Modern Physical Methods in Biochemistry, Part B (New Comprehensive Biochemistry)

MODERN PHYSICAL METHODS IN BIOCHEMISTRY, PART B

New Comprehensive Biochemistry

Volume 11B

General Editors

A. NEUBERGER London

L.L.M. van DEENEN Utrecht

ELSEVIER AMSTERDAM. NEW YORK . OXFORD

Modern Physical Methods in Biochemistry Part B

Editors

A. NEUBERGER and L.L.M. VAN DEENEN London and Utrecht

1988 ELSEVIER AMSTERDAM * NEW YORK . OXFORD

0 1988, Elsevier Science Publishers B.V. (Biomedical Division)

All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior written permission of the Publisher, Elsevier Science Publishers B.V. (Biomedical Division), P.O. Box 1527, loo0 BM Amsterdam, The Netherlands.

No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, neghgence or otherwise, or from any use or operation of any methods; products, instructions or ideas contained in the material herein. Because of the rapid advances in the medical sciences, the Publisher recommends that independent verification of diagnoses and drug dosages should be made. Special regulations for readers in the USA. This publication has been registered with the Copyright Clearance Center, Inc. (CCC), Salem, Massachusetts. Information can be obtained from the CCC about conditions under which the photocopying of parts of this publication may be made in the USA. All other copyright questions, including photocopying outside of the USA, should be referred to the Publisher.

ISBN 0-444-80968-6(volume) ISBN 0-444-80303-3 (series) Published by: Elsevier Science Publishers B.V. (Biomedical Division) P.O. Box 211 loo0 AE Amsterdam The Netherlands Sole distributors for the USA and Canada: Elsevier Science Publishing Company, Inc. 52 Vanderbilt Avenue New York, NY 10017 USA

Library of Congress Cataloging-in-PublicationData (Revised for volume 11 B) Modem physical methods in biochemistry. (New comprehensive biochemistry; v. 11 A, B) Includes bibliographies and index. 1. Spectrum analysis. 2. Biochemistry-Technique. I. Neuberger, Albert. 11. Deenen, Laurens L.M. van. QD415.N48 vol. 11 A, etc. 574.19’2 s [574.19’283] 85-4402 [QP519.9S6] ISBN 0-444-80649-0 (v. 11 A) 0-444-80968-6 (v. 11 B) Acknowledgment Many illustrations and diagrams in this volume have been obtained from other publications. In all cases reference is made to the original publication. ThejuN source can be found in the reference list. Permission for the reproduction of this material is gratefully acknowledged.

Printed in The Netherlands

Preface

In the former series of Comprehensive Biochemistry the contributions of physical methods to biochemistry were considered in volumes 1-4, a section which was devoted to the physicochemical and organic aspects of biochemistry. In 1962 the series editors M. Florkin and E.H. Stotz emphasized the importance of these basic sciences for the future progress in the life sciences. Since that time, the application of physical methods to biological problems has solved many questions and opened new avenues of research. Volume 11,part A, of the present series contained chapters on protein crystallography, nuclear magnetic resonance spectroscopy, electron spin resonance, mass spectroscopy, circular dichroism and optical rotatory dispersion. In this volume the range of spectroscopic techniques is extended to chapters on fluorescence and Raman spectroscopy. One chapter deals extensively with neutron and X-ray solution scattering techniques, and a choice of rapid reaction methods is discussed in a further chapter. The use of electron microscopy has been another very important development in the biological sciences and the results are illustrated by a chapter with emphasis on biomembranes. The New Comprehensive Biochemistry series contains a volume (8) devoted to separation methods. This area is now supplemented by a chapter in the present volume on high performance liquid chromatography of nucleic acids and a chapter on reversed phase HPLC of peptides and proteins. The editors hope that the publication of this volume may serve the needs of many biochemists and thus contribute to further research in the biological sciences.

A. Neuberger L.L.M. van Deenen

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Contents

Preface

V

Chapter I Fluorescence spectroscopy; principles and application to biological macromolecules J.R. Lakowicz (Baltimore, MD, USA)

1

1. The phenomenon of fluorescence 2. Factors affecting the fluorescence emission 2.1. Solvent polarity and viscosity 2.2. Emission spectra of melittin 2.3. Quenching of fluorescence 2.4. Fluorescence energy transfer 2.5. Fluorescence anisotropy 3. Time-resolved fluorescence spectroscopy 3.1. Resolution of the emission spectrum of liver alcohol dehydrogenase 3.2. Pulsed lasers for time-resolved fluorescence 3.3. Frequency-domain resolution of protein fluorescence 3.4. Anisotropy decays of protein fluorescence 4. Harmonic-content frequency-domain fluorometry 5. summary

1 4 4 6 7 9 11 13 15 18 19 21 23 25

Acknowledgements References

25 26

Chapter 2 Raman and resonance Raman spectroscopy P.R. Carey (Ottawa, Ont., Canada)

27

1. 2. 3. 4. 5. 6. 7.

Introduction The units used in Raman spectroscopy A model for Raman scattering based on classical physics Raman and resonance Raman scattering: a quantum mechanical interpretation Polarisation properties of Raman scattering Basic experimental aspects Raman studies on biological materials 7.1. Proteins 7.1.1. Amide I and amide I11 features 7.1.2. Side chain contributions to the Raman spectrum 7.1.3. Applications 7.1.4. UV excited resonance Raman spectra of proteins

27 29 31 34 37 38 40 40 40 42 43 43

...

Vlll

7.2. Proteins containing a natural, visible chromophore 7.3. Resonance Raman labels 7.4. Nucleic acids 7.4.1. The purine and pyrimidine bases 7.4.2. Conformation of the (deoxy)ribose-phosphate backbone 7.4.3. Resonance Raman studies of nucleic acids 7.5. Viruses 7.6. Lipids and membranes 7.6.1. The C-C stretching region between 1050 and 1150 cm-' 7.6.2. The C-H stretching region between 2800 and 3000 cm-' 7.6.3. Deuterated lipids as selective probes 7.6.4. Lipid protein interactions and natural membranes

44 48 50 50 52 53 54 56 57 57 58 59

References

61

Chapter 3 Rapid reaction methods in biochemistry Quentin H. Gibson (Ithaca, NY, USA)

65

1. Introduction 2. Continuous flow 3. Stopped flow 3.1. Miscellaneous stopped-flow devices 3.2. Relaxation methods 3.2.1. Flash sources 3.2.2. Observation light sources 3.2.3. Light detectors 4. Combinations of flash photolysis with other techniques 5. Temperature jump 6. Miscellaneous methods 6.1. Time-resolved resonance Raman spectroscopy 6.2. Competitive methods 7. Data reduction

65 65 69 71 72 73 75 75 76 76 77 77 78 78

References

83

Chapter 4 High performance liquid chromatography of nucleic acids M. Colpan and D. Riesner (Dusseldorf, FRG)

85

1. Introduction 2. Techniques 2.1. Size exclusion chromatography 2.2. Anion-exchange chromatography 2.3. Reversed phase and hydrophobic interaction chromatography 2.4. RPC-5 and other mixed mode chromatography 2.5. Sample preparation and recovery 3. Applications 3.1. Oligonucleotides 3.2. Natural RNA 3.3. DNA fragments 3.4. Plasmids 4. Concluding remarks

85 86 86 88 91 94 95 96 96 98 99 101 102

Acknowledgements References

Chapter 5 Reversed phase high per,,rmance liquid chromatography of peptides and proteins M.T.W. Hearn and M.I. Aguilar (Clayton, Vic., Australia)

103 103

107

1. Introduction 2. Retention relationships of peptides in RP-HPLC 3. The relationship between peptide retention behaviour and hydrophobicity coefficients 4. Bandwidtb relationships of peptides in RP-HPLC 5. Dynamic models for interconverting systems 6. Conclusion

107 111 120 126 131 139

Acknowledgements References

139 140

Chapter 6 X-ray and neutron solution scattering S.J. Perkins (London, UK)

143

1. Introduction Part A: Theoretical and Practical Aspects 2. Theory of X-ray and neutron scattering 2.1. Scattering phenomena and their angular ranges 2.1.1. X-ray scattering 2.1.2. Neutron scattering 2.1.3. Scattering angles, vectors and resolution 2.2. The scattering event and the Debye equation 2.3. Scattering densities and allowance for solvent 2.3.1. Concept of scattering densities 2.3.2. Scattering densities and volumes 2.3.3. The contrast difference A p 2.3.4. Mean macromolecular scattering densities p 2.3.5. Scattering density fluctuations pF(r) 2.4. The Guinier plot: Z ( 0 ) and R , 2.4.1. The innermost scattering curve 2.4.2. Cross-sectional and thickness Guinier analyses 2.5. Analyses of I ( 0 ) values 2.6. Analyses of R, values 2.7. Non-uniform scattering densities and contrast variation 2.7.1. The Stuhrmann plot 2.7.2. Solvent penetration and exchange effects 2.7.3. Isomorphous replacement 2.7.4. Matchpoints of multicomponent systems 2.8. Label triangulation 2.9. Wide-angle scattering and modelling strategies 2.9.1. Spheres and ellipsoids 2.9.2. Scattering curves at large Q 2.9.3. Independent parameters from scattering 2.9.4. Debye curve simulations 2.9.5. Interparticle interference 2.10. Distance distribution functions

143 144 144 144 144 145 146 147 149 149 150 152 154 154 160 160 162 163 165 167 167 170 172 173 173 175 175 177 178 178 180 180

X

3. Experimental practice and instrumentation 3.1. Sample preparation and measurement 3.1.1. Sample monodispersity and concentrations 3.1.2. Sample assays 3.1.3. Sample backgrounds 3.1.4. Sample holders 3.1.5. Instrumental calibration 3.2. Labelling techniques and deuteration 3.3. Sources of X-rays and neutrons 3.3.1. Anode sources 3.3.2. Synchrotron radiation 3.3.3. Reactor neutron sources 3.3.4. Spallation neutron sources 3.4. Scattering instrumentation 3.4.1. X-ray cameras 3.4.2. Neutron cameras 3.5. Data reduction Part B: Biochemical Applications to Proteins, Carbohydrates, Lipids and Nucleic Acids 4. Applications of X-ray and neutron scattering 4.1. Introduction 4.2. X-ray studies on globular proteins 4.2.1. Relationship between R , and M , 4.2.2. Comparison of crystal and solution structures 4.2.3. Conformational changes and ligand binding 4.2.4. AUostericism 4.2.5. Molecular modelling of proteins 4.2.6. Associative systems and time-resolved synchrotron radiation studies 4.2.7. Interparticle interference 4.2.8. X-ray contrast variation and anomalous scattering 4.2.9. Label triangulation of heavy metal probes 4.3. Neutron studies on globular proteins 4.3.1. Contrast variation studies 4.3.2. Label triangulation and deuteration 4.4. X-ray and neutron studies on glycoproteins 4.4.1. Plasma glycoproteins, proteoglycans and polysaccharides 4.4.2. Immunoglobulins 4.4.3. Components of complement 4.5. Lipids, detergents, membrane proteins and lipoproteins 4.5.1. Lipid vesicles and complexes with proteins 4.5.2. Detergent micelles and complexes with proteins 4.5.3. Lipoproteins 4.6. Nucleic acids and nucleoproteins 4.6.1. DNA studies by X-ray scattering 4.6.2. X-ray and neutron studies on transfer RNA 4.6.3. Protein-nucleic acid interactions by neutron scattering 4.6.4. Chromatin and chromosomes by X-rays and neutrons 4.6.5. Ribosomes and their constituents 4.6.6. Viruses 5. Conclusions Acknowledgements References

182 182 183 183 184 184 185 186 187 187 187 189 190 190 190 191 193 194 194 194 194 194 196 196 198 199 201 203 204 207 208 208 21 1 213 21 3 218 219 221 221 224 226 230 230 231 234 236 239 244 249 25 1 251

Chapter 7 Electron microscopy W.F. Voorhout and A.J. Verkleij (Utrecht, The Netherlands)

267

1. Introduction 2. Negative staining and metal shadowing 2.1. Negative staining 2.2. Metal shadowing 3. Thin sectioning 4. Low-temperature techniques 4.1. Cryofmation 4.2. Freeze-fracturing 4.2.1. The freeze-fracture technique 4.2.2. Biological membranes 4.2.3. Lipid phase transitions and lipid polymorphism as visualized by freeze-fracturing 4.3. Localization studies 4.3.1. Introduction 4.3.2. Immunocytochemistry 4.3.3. Marker system 4.3.4. Cryo-ultramicrotomy 4.3.5. Cryo-fractures 4.3.6. Label-efficiency 5. Conclusions

261 268 268 269 210 212 212 214 214 215 219 286 286 281 288 289 290 293 295

Acknowledgements References

295 295

Subject index

301

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A. Neuberger and L.L.M. Van Deenen (Eds.) Modern Physical Methoak in Biochemimy. Part B 0 1988 Elsevier Science Publishers B.V. (Biomedical Division)

1

CHAPTER 1

Fluorescence spectroscopy; principles and application to biological macromolecules * JOSEPH R.LAKOWICZ University of Maryland at Baltimore School of Medicine, Department of Biological Chemistry, 660 West Redwood Street, Baltimore, M D 21201, USA

1. The phenomenon of fluorescence Luminescence is the emission of photons from electronically excited states. Luminescence is divided into two types, fluorescence and phosphorescence. In phosphorescence, the emission is from an excited triplet state to a ground state singlet. Since this transition is forbidden the rate of return to the ground state is slow, which means the decay times are long (msec to sec). Fluorescence is the emission from excited singlet states, also yielding a ground state singlet. These allowed transitions to occur rapidly, with rates near lo8 sec-'. Consequently, the decay times for fluorescence are typically near lo-' sec or 10 nsec. In this chapter we will discuss primarily fluorescence, but the concepts are also applicable to events on a slower timescale if the phosphorescence is observed. The nanosecond timescale of fluorescence provides much of its usefulness in biophysical chemistry. In solutions near room temperature, a variety of molecular events can occur within 10 nsec and alter the emission. These events include rotational diffusion, collisions with quenchers, solvent reorientation, and energy transfer. These events alter one or more of the spectral observables, and can thus be detected by analysis of the emission. Substances which display fluorescence are generally delocalized aromatic systems with or without polar substituents (Fig. 1).It is difficult to predict which molecules will be fluorescent or non-fluorescent because exceptions can usually be found. However, several general rules are generally true. Rigid molecules are usually more fluorescent, or at least their fluorescence more predictable, than molecules with the possibility of internal rotation. Hence, perylene and anthracene fluoresce with high efficiencies, whereas stilbene can be much less efficient. In viscous solvents, in which rotational reorientation to cis-stilbene cannot occur, trans-stilbene is highly fluorescent. In non-viscous solution stilbene is only weakly fluorescent. This illustrates an important aspect of fluorescence, which is that the excited states are involved, * Dedicated to Professor Gregorio Weber on the occasion of his seventieth birthday.

2

Perylene

Anfhracene

I ndole

P PO

trans- Stilbene

2-Naphthol

Fig. 1. Typical fluorescent molecules.

and these states have a different electronic distribution which may alter their chemical properties. In the excited state trans-stilbene (Fig. 1) isomerizes to the non-fluorescent cis-stilbene. The altered electronic distribution can also alter chemical reactivity. For instance, the pKa of the hydroxyl group on naphthol decreases from 9 to 2 upon excitation, presumably as the result of transfer of electron density from the oxygen into the aromatic ring. The emission from an aqueous solution of naphthol can be due to unionized naphthol, naphtholate, or both, depending upon pH and the concentration of basic species available to accept the dissociated proton. The presence of substitutes for carbon in the aromatic system generally alters the emission from the aromatic nucleus. Insertion of oxygen or nitrogen into the ring system often results in good fluorescence. Hence, indole, fluorescein, PPO, the rhodamines and similar substances are fluorescent (Fig. 1). The presence of sulfur, nitro groups or heavy atoms like iodide generally result in quenching of fluorescence. Biological systems contain a variety of intrinsic (natural) fluorophores (Fig. 2). In proteins, tryptophan is the most highly fluorescent amino acid, accounting for 90% of the emission from most proteins. Emission from tyrosine residues is also observed, especially in proteins lacking tryptophan, in denatured proteins, or in those with a high ratio of tyrosine to tryptophan. Tyrosine is highly fluorescent in solution, but its emission is often quenched in native proteins, due either to the quenching effects of hydrogen bonding to the hydroxyl group or because of energy transfer from tyrosine to tryptophan. The emission of phenylalanine from proteins is less studied. The nucleotides and nucleic acids are generally non-fluorescent. However, some notable exceptions are known. Phenylalanine transfer RNA from yeast (tRNAPhe) contains a single highly fluorescent base, called the Y-base, whch has an emission maximum near 470 nm. The presence of this intrinsic fluorophore has resulted in numerous studies of tRNAPheby fluorescence spectroscopy. Regarding the “nonfluorescent” nucleic acids, it should be noted that they do fluoresce, but with very low yields and with short decay times.

3

CH

/I

CH

I

CH

II

CH

I

CH

II

ANS

DNS-CI

DPH

-ATP

Ethidium Bromide

Acridine Orange

Fig. 2. Intrinsic biological fluorophores. For NADH and FAD we only showed the fluorescent part of the molecule.

Other natural fluorophores include NADH and FAD, whose fluorescent moieties are shown in Fig. 2. In both cases the amount of fluorescence depends upon their local environments. For instance, the emission of NADH is usually increased about three-fold upon binding to proteins, whereas the emission of FAD is usually quenched. C02H

C02H I

a;H2 6 H2N-CHI

H

Tryptophan

Y,- b a s e

H2N-CH

I

C02H

I

H 2N-CH

6 I

\

OH

Tyrosine

NADH

Phenylalanine

FAD

Fig. 3. Typical extrinsic fluorophores used to label macromolecules.

4

In instances where nature has not provided an appropriate fluorophore, one can often add an extrinsic label. The earliest probes include dansyl chloride [l]and ANS (Fig. 3). Dansyl chloride can be covalently attached to macromolecules by reaction with amino groups. ANS often binds spontaneously but non-covalently to proteins and membranes, probably by hydrophobic and electrostatic interactions. The emission of both molecules is sensitive to the polarity of the surrounding environment. ANS is nearly non-fluorescent in water, but fluoresces strongly upon association with serum albumin, immunoglobulins and other proteins. A wide variety of covalent and non-covalent probes are available [2,3]. Studies of cell membranes by fluorescence depends almost exclusively upon the use of extrinsic probes. This is because most lipids are not fluorescent, and the emission from membrane-bound proteins is too heterogeneous for interpretation of the data. The probe DPH (Fig. 3) is typical of membrane probes, as is perylene (Fig. 1).These non-polar molecules partition spontaneously into membranes. And finally, Fig. 3 shows several extrinsic probes for nucleic acids. Addition of the etheno bridge to ATP results in a highly fluorescent residue. Unfortunately, this modification also disrupts the base pairing of the nucleotide. Alternatively, nucleic acids can be labeled by spontaneous binding of planar cations such as ethidium bromide and acridine orange (Fig. 3). Depending upon the structure, the fluorescence of the probe may be quenched or enhanced upon intercalation into DNA, and the emission may depend upon whether the intercalation site is adjacent to A-T or G-C pairs. For instance, the fluorescence yield of ethidium bromide is enhanced about 30-fold upon intercalation into DNA. Other intercalating dyes such as proflavin and 9-aminoacridine are quenched by their interactions with DNA [4].

2. Factors affecting the fluorescence emission 2. I . Solvent polarity and viscosity The variety of factors which can affect the fluorescence emission are illustrated by the modified Jablonski diagram [5] shown in Fig. 4. In this diagram we emphasize fluorescence emission and quenching, and hence we have not included the higher electronic states or the triplet states. Upon absorption of light the fluorophore arrives instantaneously in the first singlet state (Sl), usually with some excess vibrational energy. This excess energy is usually dissipated quickly in lo-'' sec by interaction with the solvent, resulting in a molecule in the lowest vibrational level of S,. The fluorophores remain at this level for the mean duration of the excited state, which is typically 10 nsec. Any process or interaction which occurs during this interval can alter the fluorescence emission. These processes and interactions are the origin of much of the information available from fluorescence spectroscopy. Fluorophores with polar groups are often sensitive to solvent polarity. Interaction between the excited fluorophore and surrounding polar groups lowers the energy of the excited state, which shifts the emission to longer wavelengths. The relative amounts of emission from the relaxed and the unrelaxed states depend upon the

5

\j

S L a R e l a Y Donor Emission

k,

Energy Transfer Acceptor absorption

Xkil

h

a

so

Fig. 4. Jablonski diagram for fluorescence emission and quenching.

relative rates of depopulation of the excited state ( r+ Cki) and that of solvent relaxation ( k R ) .The rate of emission is r and the rate of return to the ground state exclusive of emission is Cki. In fluid solvents near room temperature the rate of solvent reorientation is near 10-1'-10-12 sec. Hence, this process is mostly complete prior to emission, so the observed emission is that of the relaxed state. If the solution is cold or viscous, or if the probe is bound to a rigid site on a macromolecule, then the rates of relaxation and emission can be comparable, so emission is seen from both the relaxed and the unrelaxed states. If the solution is very viscous then solvent relaxation does not occur during the lifetime of the excited state, and the observed emission is from the higher energy (shorter wavelength) unrelaxed state. The effects of solvent polarity are best understood by specific examples. To model the fluorescence emission of proteins we examine spectra for N-acetyl-Ltryptophanamide (NATA). This molecule is analogous to tryptophan in proteins. It is a neutral molecule, and its emission is more homogeneous than that of tryptophan itself. In solvents of increasing polarity the emission spectra shift towards longer wavelengths (Fig. 5). The emission maxima of NATA in dioxane, ethanol and water are 333, 344 and 357, respectively. These solvents are non-viscous, so the emission is dominantly from the relaxed state (Fig. 4). The spectral shifts can be used to calculate the change in dipole moment which occurs upon excitation [6]. More typically, the emission spectrum for a sample is compared with that found for the same fluorophore in various solvents, and the environment judged as polar or non-polar. While this approach is qualitative, it is simple and reliable, and does not involve the use of theoretical models or complex calculations. The timescale of the relaxation process also affects the emission. This effect is illustrated for NATA in propylene glycol (Fig. 6). At room temperature the relaxation is mostly complete, and a red shifted spectrum is observed (348 nm). Lowering the temperature results in a progressive shift of the emission to shorter wavelengths, with an emission maximum of 329 nm at - 60 O C . As the temperature is decreasing the relaxation rate ( k R )becomes slow relative to that of the decay rate ( r+ Cki). Hence, an increasing proportion of the emission is from the unrelaxed and the intermediate states (Fig. 4), which have higher energies and shorter emission

6

WAVELENGTH ,(nanometers)

Fig. 5 . Emission spectra of N-acetyl-L-tryptophanamidein various solvents of different polarities.

wavelengths (Fig. 6). In proteins it is probable that the emission maxima are affected by both the average environment of the tryptophan residues [7] and by the relaxation rates [8]. Rather detailed data and analysis are needed for an unambiguous separation of these effects, but the average environment of the tryptophan residues seems to be the dominant determinant of the emission maxima. 2.2. Emission spectra of melittin Melittin is an amphipathic peptide component of bee venom which associates with cell membranes, enhances the phospholipase activity of venom and participates in the disruption of cell membranes. This protein has been studied extensively by

WAVELENGTH [nanometers)

Fig. 6. Emission spectra of N-acetyl-L-tryptophanamidein propylene glycol at 25 and

- 60

C

WAVELENGTH (nanometers) Fig. 7. Emission spectra of melittin in the absence (data are courtesy of N. Joshi.

) and presence (-

- -)

of 2 M NaCl. The

fluorescence and other physical methods [9,10], and its X-ray structure is known [ll]. In solution melittin exists as either a monomer or a self-associated tetramer. The self-association is driven by high salt concentration, which apparently shields the positive charges on the monomer from each other and allows the hydrophobic interactions to cause association. The monomeric form of melittin is thought to be largely random coil with a high degree of segmental mobility. In the tetrameric state the monomeric units are mostly a-helical. Melittin is an ideal protein to illustrate the effects of structure on the fluorescence spectral properties. Each monomer contains a singly tryptophan residue and no tyrosine residues. The X-ray structure of the tetrameric form shows that the tryptophan residues are buried in a non-polar pocket and are not directly exposed to the aqueous phase. The emission spectra of melittin illustrate the effects of solvent exposure on the tryptophan emission (Fig. 7). In the absence of salt, the emission maximum of 360 nm is comparable to that found for NATA in water. In the presence of 2 M NaCl the emission maximum is blue shifted by 12 nm to 348 nm. This shift is a result of shielding of the indole ring from the aqueous phase. Hence, solvent relaxation proceeds to a lesser extent because there is less solvent available for interaction with the fluorophore. 2.3. Quenching of fluorescence

Collisional quenching of fluorescence requires contact between the fluorophore and the quencher. For quenching to occur the quencher must diffuse to and collide with the fluorophore in the excited state. If this occurs the fluorophore returns to the ground state without emission of a photon (Fig. 4). Many small molecules act as collisional quenchers of fluorescence [6,12]. These include iodide, acrylamide, halogenated hydrocarbons and occasionally amines and metal ions. The excited state lifetimes provide ample opportunity for quenching. For instance, acrylamide is known to be an efficient quencher of tryptophan fluorescence [12,13]. Su.ppose its

8

I P

Melittin 7.01 2 5 O C , p H - 7

Monomer K =

7,2 M-'

k = 2.1x I 0 ' M-'s-'

5.0-

-FF.

0.6

0.4

0.2

0

0.8

[Acrylamidel ,M

Fig. 8. Stern-Volmer plot for acrylamide quenching of melittin monomer and tetramer. From M. Eftink, University of Mississippi, Chemistry Department, unpublished observations. The lifetimes are from [35]. The broken lines are the initial slopes, corresponding to the values on the figure.

diffusion coefficient is lop5cm2/sec. In 10 nsec an acrylamide molecule can diffuse a distance of 44 A, as calculated using A x 2 = 2 0 7 , where A x is the distance, D is the diffusion coefficient and 7 is the fluorescence lifetime. This distance is comparable to the diameter of many proteins. Hence, we expect quenching to occur to a measurable degree and the extent of quenching to be sensitive to the average degree of exposure of the tryptophan to the aqueous phase. Once again melittin illustrates the effect of protein structure on the fluorescence emission. Acrylamide quenching data for melittin monomer and tetramer are shown in Fig. 8. Stern-Volmer plots are often used to present quenchmg data. The Stern-Volmer equation is FO = 1+ k.,[Q]

F

=

1

+K [ Q ]

where Fo and F are the fluorescence intensities in the absence and presence of quenching, respectively, T~ is the fluorescence lifetime in the absence of quenchmg, [ Q ]is the concentration of quencher, k is the bimolecular quenching constant, and K is the Stern-Volmer quenching constant. The lifetime T,, is the reciprocal of the rates which depopulate the excited state. From Fig. 4,

If every collisional event results in quenching, the bimolecular rate constant can be estimated using the diffusion constants of the fluorophore ( D F )and quencher ( DQ) and the radius expected for contact ( R ) , k

(3)

= 4~rRDN/1000

where N is Avogadro's number and D

= D,

+ DQ. If the fluorophore is exposed to

9

the solvent we expect k to be near 0.5 X lo1' M-' sec-'. If the residue is shielded from collisional encounters this rate will be smaller. This comparison is the basis for estimating the extent of exposure from quenching data. The quenching constant measured for a protein is compared with that expected for a completely exposed fluorophore. Typically, model compounds with no possibilities for shielding are studied to account for lack of precise knowledge of diffusion coefficients, and the possibility that the quenching encounter is not 100% efficient. The quenching data for both the monomeric and tetrameric forms of melittin indicate the tryptophan residues are accessible to acrylamide with the accessibility being greater in the monomeric state. This conclusion is reached by comparison with acrylamide quenching data for NATA. At 25°C in water the acrylamide quenching constant for NATA is 0.58 X 10" M-' sec-' [47]. For the monomer the quenching constant is about one-third of this value, which is indicative of a rather fully exposed residue [13]. The value of k for the tetramer is less, indicating shielding of the tryptophan residue from the aqueous phase. It should be noted that the relative shielding is only 40%, which probably indicates considerable penetration of the tetramer by acrylamide. In other more extensive studies Eftink and Ghiron showed that acrylamide quenching reflects the average degree of tryptophan exposure to the aqueous phase [13]. The penetration of proteins by quenchers has been known for some time [14,15]. For melittin tetramer the penetration by acrylamide is not unexpected since acrylamide is neutral and the tryptophans are located in a loosely packed non-polar region of the protein [ll]. 2.4. Fluorescence energy transfer

Another process which can occur during the excited state is fluorescence energy transfer, which is the transfer of the excited state energy from a donor (D) to an acceptor (A) (Fig. 4). The transfer is called radiation-less because it occurs without the appearance of a photon. This process is strongly dependent upon distance because it is the result of dipole-dipole coupling between the donor and the acceptor [16]. A requirement for energy transfer is that the emission spectrum of the donor overlaps with the absorption spectrum of the acceptor. The rate of transfer ( kT) is gven by (4) where R , is the distance at which 50% of the energy is transferred, and r is the donor-to-acceptor distance. The value of R o can be calcuated from the spectral properties of donor and acceptor [6,16]. The efficiency of energy transfer is given by the ratio of the rates of transfer to the total rate of depopulation of the donor. Hence,

10

Usually, both the transfer efficiency ( E ) and R , are determined experimentally. Then, the donor-to-acceptor distance is calculated using

This method is widely used to measure the distance between sites on a macromolecule, and has been the subject of considerable experimentation and discussion [17- 191. Energy transfer has been used to measure the self-association of melittin. The melittin was labeled with a N-methylanthraniloyl (NMA) residue on one of the lysine residues. This fluorophore serves as the energy acceptor for the single tryptophan residue. Only a small fraction (5%) of the melittin monomers was labeled with NMA. In the monomer there is only one tryptophan residue near the acceptor, whereas four such residues are present in the tetramer. Hence, the extent of tryptophan to NMA energy transfer should be sensitive to and increased by melittin self-association. In this experiment the intention is not to determine a distance, but rather to use the association-dependent energy transfer to determine the extent of self-association [20].

300

350

400

450

500

WAVELENGTH (nanometers)

Fig. 9. Emission spectra of N-methylanthraniloyl-labeled melittin. Spectra are shown for the monomer (0 M NaC1) and for the tetramer (2 M NaCI). From [20]. The broken lines are the emission spectra of the unlabeled melittin.

11

250

300

350

400

WAVELENGTH (nanometers 1

Fig. 10. Excitation spectra of NMA-labeled melittin. From [20].

Emission spectra of the labeled melittin are shown in Fig. 9. Recall that only a small fraction of the melittin contains a NMA label. Hence, the emission spectra are mostly characteristic of tryptophan, with shoulders at 430 nm due to the NMA emission. In the presence of 2 M NaCl the NMA emission is enhanced, reflecting increased energy transfer from the additional tryptophan residues. Excitation spectra are often used to study energy transfer. This is because energy transfer can be detected by enhanced emission from the acceptor when the excitation is centered at the donor absorption. The effects of melittin self-association are evident from the excitation spectra (Fig. 10). For these spectra the emission monochromator is centered on the NMA emission (430 nm) and the intensity recorded as the excitation monochromator is scanned through the absorption bands of the NMA label (350 nm) and the tryptophan absorption (280 nm). Increasing salt concentrations result in increased intensity of the tryptophan excitation band (280 nm). This increase in energy transfer is due to the close proximity of the three additional donors to the NMA acceptor.

2.5. Fluorescence anisotropy The timescale of fluorescence emission is comparable to that of rotational diffusion of proteins and the timescale of segmental motions of protein domains or individual amino acid residues. The polarization or anisotropy of the emission provides a measure of these processes. Suppose a sample is excited with vertically polarized light (Fig. ll),and that the sample is viscous so that the fluorophores do not rotate during the lifetime of the excited state. Then the emission is polarized, usually also in the vertical direction. This polarization occurs because the polarized excitation selectively excites those fluorophores in the isotropic solution whose absorption

12

LIGHT

SOURCE-

m+ f/

DETEC~OR Fig. 11. Measurement of fluorescence anisotropies.

moments are aligned vertically. The extent of polarization is most conveniently defined by the anisotropy [6,40].

where I refers to the intensities, and the subscripts indicate the parallel (11) or component. perpendicular (I) A number of processes can result in the loss of anisotropy, the most common being rotational diffusion. Melittin is expected to have rotational correlation times near 2 and 8 nsec in the monomeric and tetrameric states, respectively. The effect of rotational diffusion on the anisotropy is described by the Perrin equation,

where r, is the anisotropy in the absence of rotational diffusion, r is the anisotropy, is the lifetime and 6 is the correlation time. The value of r, is usually measured in a separate low-temperature experiment. Its value depends upon the excitation wavelength, and is typically in the range of 0.1 to 0.4. The r, value is a measure of the angle between the absorption and emission transition moments of the fluorophore. For tryptophan the value of ro on the long wavelength side of the absorption is near 0.32. Values of r, which are less than 0.1 are usually not useful because the difference between 11,and I* will be small, and the precision of the measurements will be decreased. When 7, and 8 are of similar magnitude then the measured anisotropy is dependent upon the correlation time. Self-association of melittin is expected to increase its correlation time about four-fold. Since the lifetime of melittin fluoresence is near 3 nsec we expect self-association to have a dramatic effect on the 7,

13

old0

'

0'4

'

0.8 '

'

I.2 '

'

1'.6

'

2.0 I

'

[NaCII. M

Fig. 12. Fluorescence anisotropies of melittin [20]. Anisotropies are shown both for the intrinsic tryptophan emission (A), and for that of the NMA label (0). Also shown is the effect of salt concentration on the extent of energy transfer (0).

anisotropy. Anisotropy data for melittin at various salt concentrations are shown in Fig. 12. As the salt concentration is increased the anisotropy values increase and reach a plateau whch is characteristic of the tetramer. Also shown in Fig. 12 are the anisotropy values of the NMA label. These also increase with salt concentration, and reach a constant value at 1 M NaC1. The NMA anisotropy values are lower than for tryptophan because the decay time of the NMA label is longer, near 8 nsec. For comparison this figure also shows the extent of energy transfer. All three measurements reflect the monomer to tetramer transition. Anisotropy measurements are generally useful for measuring any process which increases or decreases the rate or extent of rotational diffusion. These processes include domain motions of immunoglobulins [21], denaturation of proteins [22] and the association of proteins with membranes [lo]. Additionally, there are numerous applications of anisotropy measurements to membranes, in which the phase state and apparent fluidity are estimated from the anisotropy of probes which are bound to the membranes [23,24].

3. Time-resolved fluorescence spectroscopy The previous discussion and examples emphasized the use of steady-state fluorescence data. Steady-state data are measured with constant illumination of the sample. The timescale of these measurements is slow relative to the fluorescence decay times. Hence, the effects of the time-dependent processes are averaged to yield the average emission spectra, anisotropies, or extents of energy transfer. Each measured steady-state quantity is the average of the time-dependent values of that quantity averaged over the time-dependent decay of the sample. For instance, the

14

steady-state anisotropy is determined by its time-resolved decay ( r (t )) and the time-dependent decay of the emission ( Z ( t ) )

If we assume that both r ( t ) and I ( t ) decay as single exponentials with time constants of 1/8 and 1/~,,, respectively, then application of equation 9 yields the steady-state form, whch is the Perrin equation (8). At present, there is widespread interest in directly measuring the time-dependent processes. This is because considerably more information is available from the time-dependent data. For example, the time-dependent decays of protein fluorescence can occasionally be used to recover the emission spectra of individual tryptophan residues in a protein. The time-resolved anisotropies can reveal the shape of the protein and/or the extent of residue mobility within the protein. The time-resolved energy transfer can reveal not only the distance between the donor and acceptor, but also the distribution of these distances. The acquisition of such detailed information requires high resolution instrumentation and careful data acquisition and analysis. There are presently two methods of obtaining the time-resolved data. These are by direct measurements in the time-domain [25,26] and by less direct measurements

TIME

I

0

TIME

I

20

1

I

40

I

1

60

TIME (nanoseconds)

Fig. 13. Measurement of time-resolved fluorescence in the time-domain (top) and in the frequency-domain (bottom).

15

in the frequency-domain [27,28]. For time-domain measurements the sample is excited with a brief pulse of light (Fig. 13). The time-dependent fluorescence intensities are used to estimate the decay time(s) of the sample. In the frequency-domain the sample is excited with intensity modulated light. The frequency response (phase and modulation) of the sample are used to estimate the decay time(s). Both methods are rapidly evolving to take advantage of the increased time resolution obtainable using picosecond pulse lasers and faster detectors [29,30]. The complex equipment and analyses necessary for time-resolved measurements has been the subject of numerous publications and monographs [25,26]. In this article we will not describe the instrumentation, but will rather describe the results and their interpretation. The objective of either time or frequency-domain fluorometry is to determine the decay law of the sample. For example, consider protein containing two tryptophan residues, and assume further that each residue has a single decay time. The impulse response of the sample is the decay which would be observed with an ideal instrument following excitation with a &function light pulse. For our hypothetical protein we expect a doubly exponential decay of intensity, 2

I( t ) =

C aie-'/'~ i=l

In this expression the 7; values are the decay times of the individual residues and ai values are the preexponential factors. The contribution of each residue to the emission is

Suppose the data are measured at a number of wavelengths across the emission spectrum. Then the data are described by 2

I(X, t)=

C a,(X)e-f'Ti i=l

where X indicates the emission wavelength. Under favorable conditions, such data allow the spectrum of each residue to be recovered. This can be seen by recognizing that the values of h(X) represent the fraction of the total emission due to each residue. This illustrates how the time-resolved data provide information not obtainable from the steady-state spectra. 3.1. Resolution of the emission spectrum of liver alcohol dehydrogenase

The resolution and understanding of the emission from proteins is a difficult task. This is because most proteins contain two or more tryptophan residues, and even

16

10'

10'

b double

I oJ

I oJ

in

in

tZ

t-

z

:

3

u

u

3

;I 0 2

102

0

0

1

_I

10'

10'

1

100

0

1

1

1

I

100

200

300

400

I oo

i

100

0

500

200

30C

400

500

400

500

CHAFJNELS

CHANNELS

aooo

aooo

6000

I'

A

'I

6000

m

! I

54 0 0 0

5400C

t-

+

0

0

u

0

200[

2000

in0

200

300

400

so0

CHANNELS

0

100

200

300

CHANNELS

Fig. 14. Time-resolved fluorescence intensity of liver alcohol dehydrogenase. Left: single exponential fit, x i = 1.14. The faster decaying curves are the lamp profiles found using scattered light. The upper panels are on a semilogarithmic scale, and the lower are on a linear scale. The lower panels show the deviations, and the autocorrelation of the deviations as insets.

x i = 3.02; right: double exponential fit,

the emission from proteins containing a single tryptophan is complex or multi-exponential. One of the most detailed studies is for liver alcohol dehydrogenase (LADH) by Brand and co-workers [7]. LADH consists of two identical subunits (dimeric) with a molecular weight of 80,000 daltons. Each subunit contains two tryptophan residues, trp-15 and trp-314. One residue (trp-15) is exposed to the solvent, while t h s other (trp-314) is in a hydrophobic pocket at the dimer interface. Time-resolved data for LADH are shown in Fig. 14. These data for a single emission wavelength, were obtained by the method of time-correlated single photon

17

counting [25,26]. The light source is a flash lamp which fires repetitively at rates near 20 kHz. The time-resolved decay is obtained by measuring the time interval between the lamp pulse and the first emitted photon. The rate of detecting the emitted photons is kept near 2% of the lamp rate. In this way the first photon represents a sampling of the entire time-resolved decay. If many photons reach the detector, and only the first one is counted, then the measured decay does not represent the true decay. The electronic circuits (in current use) which detect the arriving photons cannot accept and process multi-photon events, which limits the rate of data acquisition. A second difficulty is the width of the lamp pulse. Flash lamps are inexpensive, but the pulse widths are near 2 nsec, whch is not much shorter than the decay times of proteins. Consequently, mathematical procedures, known as deconvolution or reconvolution, are used to account for the lamp pulse width. Nonetheless, the necessary procedures are highly refined and reliable. The data for LADH (Fig. 14) illustrate how the time-resolved data are analyzed. The analysis starts by assuming the decay is a single exponential (equation 1 with i = l),and calculation of the best fit. This fitting procedure involved use of the lamp profile and a guess of the decay time to predict the data. The numerical value of the single decay time is varied until the best fit is found. Frequently, a single decay time model is not adequate, as was found for LADH. The inadequacy of the fit is revealed by systematic deviations between the measured and calculated data (Fig. 14, lower left), and an unacceptably high value for the goodness-of-fit parameter xi = 3.02. For an acceptable fit xi should be near unity. Greater values of xi can be the result of either an inadequate model or systematic errors in the data. The acceptable values of xi are determined, in principle, by statistical criteria [41], but individual judgements are necessary. The next step in the analysis is to fit the data using a more complex model. The best fit for LADH for two decay times yields an improved match (Fig. 14, right). The calculated and measured values are now in agreement, the deviations are small and random, and xi = 1.14 is acceptably close to unity. The parameters (a,and 7,) which yield this match are taken as the decay law of the sample. The decay times (3.8 and 7.2 nsec) were taken to be due to tryptophan 314 and 15, respectively. It must be emphasized that this result does not prove the decay is a double exponential, but only shows that the double exponential model is adequate to explain the data. If data were available with higher time resolution or statistical accuracy (more photons) then it may be necessary to use a more complex three-exponential decay model to explain the data. To resolve the emission spectra of each residue similar data were collected at closely spaced wavelengths across the emission. The assumption was made that the decay time of each tryptophan residue was independent of emission wavelength, and constant across the emission. The measured decay law was used to calculate the individual spectra using

18 I

I

I

1

m

F 5

w

LL4

T, = 3 8 0 n s T2 = 7.22 ns

3 2 0 3 5 0 380

410

WAVELENGTH (nm)

0 WAVELENGTH

(NMi

Fig. 15. Lifetimes, amplitudes and emission spectra of the two tryptophan residues in liver alcohol dehydrogenase 171.

These spectra are shown in Fig. 15. The shorter-lived trp-314 (3.8 nsec) showed a shorter-wavelength emission, whereas the longer-lived trp-15 emitted at longer wavelengths. Hence, the emission spectrum and decay time of each tryptophan residue in LADH were resolved. 3.2. Pulsed lasers for time-resolved fluorescence

The time-resolved emission of LADH was found to be complex due to the presence of two tryptophans. It is highly probable that the results in Fig. 15 are only approximate, and that the decay of each tryptophan is multi-exponential. Such complexity is hidden from view by the limited resolution of the available instrumentation. The resolution of more complex decays for LADH or any protein requires higher resolution. This is being accomplished by the use of pulsed laser sources. One of the most popular sources is shown in Fig. 16. The main pump is shown as an Ar-ion laser, but the Nd-YAG lasers are used in a similar manner. Normally, these lasers yield continuous output. In this case the argon laser is mode-locked, which yields a repetitive train of pulses. These pulses are spaced at the time interval for light to travel the length of the laser cavity, which is 83 MHz. The argon laser output is then used to pump a dye laser, whose cavity length is identical to that of the Argon laser. This results in a 83 MHz train of pulses in the dye laser cavity which has pulse widths near 5 psec. These pulses are extracted at a slower rate using a cavity dumper. The cavity dumper is a small piece of quartz into which a time R F pulse is launched via a piezoelectric device. This RF pulse sets up a diffraction grating which deflects the desired laser pulse. The extracted pulse train is then

19

514nm

I

1

n

I

I

U

1

570-610 nm

u

0.1 (0 4 MHz

u 83 MHz

I

I

I

D Y E LASER

CAVITY D U M P E R

-

285 305 nm DOUBLER

5ps 4MHz

Fourier Transform

-

b 0

a

Time Frequency (GHz)

Fig. 16. Pulsed laser source for time-resolved fluorescence. The lower panel shows the pulse train from the laser source and its Fourier transform.

frequency-doubled to yield the UV necessary to excite protein fluorescence. Additional details are available in other books on this topic [42,43]. There are several significant advantages to the laser source over the more conventional flash lamp. The laser source is more intense. Its pulse width is near 5 psec, as compared for 2000 psec for a flash lamp. And finally, the repetition rate can be much higher, typically 800 kHz to 4 MHz. Because of all these factors it is possible to rapidly acquire data to a much higher level of statistical accuracy than with a flash lamp. For example, a recent paper by Small and co-workers describes a multi-component resolution of a histone, which contains a single tyrosine residue [31]. Because of the substantial increases in resolution, the laser sources are becoming more widely used in the biochemical applications of fluorescence, as illustrated by recent studies of the tryptophan emission from phospholipase A, [44] and hemoglobin [45]. 3.3. Frequency-domain resolution of protein fluorescence

Multi-exponential decays of fluorescence can also be recovered by measurements in the frequency-domain. This has only become practical within the past four years [27,28]. The resolution of multi-exponential decays requires measurements over a wide range of light modulation frequencies. Earlier instrumentation could operate at only one to three frequencies, and these limited data were not adequate to determine the four parameters in equation 10 (two a, and two T,). The new instruments

20

2.57

1.0

0.15

0.04

5.52

0.33

2 1 ~ 5 l O l

(I. 25

! 2

5

1.03

10

20

50

100

200

FREQUENCY (MHz) Fig. 17. Frequency-domain data for the intrinsic fluorescence of S, Nuclease and melittin.

operate over a wide range, typically 1 to 200 MHz, and some instruments now operate to 2000 MHz [37]. Typical frequency-domain data for two proteins are shown in Fig. 17. The data consist of the phase angles and modulation, each measured over the widest possible range of frequencies. This requirement illustrates the transform relationship between the time and the frequency-domain measurements. In the time-domain, the most desirable excitation profile is the shortest obtainable pulse. The Fourier transform of a &function consists of all frequencies. Hence, the experimental requirements are similar, short pulses or wide range frequencies. For each protein (Fig. 17) the phase angle increases and the modulation decreases as the frequency increases. The data are analyzed in a manner analogous to the time-domain data. That is, a decay law is assumed and the parameters varied until the best possible match is obtained - - -) values. The adequacy of between the measured (0)and calculated (-, the fit is judged by the value of the xi, which is a weighted sum of the squared deviations between the measured and the calculated values. If the value of is significantly larger than unity then the model can be rejected. Both melittin and S, Nuclease contain a single tryptophan residue. The data illustrated the point that the emission from such simple proteins can be multiexponential. Even though only a single residue is responsible for the emission, it was not possible to fit the data using a single decay time. This is shown by the failure of the single decay time model (- - -) to explain the data. The decays are referred to as being heterogeneous. The decays of both nuclease and melittin are signifi-

xi

21

cantly heterogeneous. The decay of melittin is more heterogeneous as seen by the greater deviations of the data from the one decay time model, and the larger value of xk. For both proteins the data can be explained by more complex decay laws. Two decay times are needed to fit the data for nuclease, and three are needed for melittin. It is important to exercise clear thinking when fluorescence data in either domain are fitted to various models. A poor fit can be used to reject a model. The poor fit can be due to either an inadequate model or due to systematic errors in the data not known to the researcher. If systematic errors occur a more complex model could be accepted to account for the errors, not because the model is appropriate for the sample. Secondly, a good fit does not prove the model which yields the good fit is correct. A good fit only shows that the model is adequate to explain the data. Alternatively stated, the data which yield the good fit are not adequate to support a more complex model. What is the origin of the multi-exponential decays found for single tryptophan proteins? Surprisingly, this is an unanswered question which is the focus of current research. Clearly, the protein matrix provides a unique but asymmetric environment for each tryptophan residue. The spectral properties (emission maximum, lifetimes, anisotropy and yield) are determined by this environment. The protein environments can now be conveniently examined using the X-ray coordinates and modem computer graphics, Additionally, it is becoming increasingly easy to replace individual amino acid residues using the techniques of molecular biology. These capabilities, and the increased resolution available from state-of-the-art instrumentation, should allow the linkage to be established between structural data and fluorescence spectral parameters. 3.4. Anisotropy decays of protein fluorescence

There is considerable interest in measuring the rotational diffusion and the dynamic properties of proteins. The rates of rotation diffusion can reveal the size and shape of the protein. Also, proteins are known to undergo structural fluctuations, a topic which has been broadly studied by both experimentation and computer simulations [32-361. The time-resolved experiments are often directed towards a comparison of the measurable dynamics of proteins with the calculated dynamics. One promising approach is to use anisotropy data from intrinsic protein fluorescence. If such data are available with picosecond resolution then such a comparison should be possible. In the time-domain the anisotropy decay is obtained from the time-resolved decays of the parallel and perpendicular polarized components of the emission. More specifically, one measures the time-resolved decays of the parallel ( II) and the perpendicular ( I) components of the emission, and calculates the time-resolved anisotrop y,

22

Generally, the anisotropy decay is multi-exponential

In this expression r( t ) is the time-dependent anisotropy, Bi the correlation times and g, the fraction of the total anisotropy ( r o )whch decays with this correlation time. In general we expect one component (8,) due to rotational diffusion of the protein, and one due to torsional motions of the tryptophan residue, if such motions are significant. In proteins which contain more than a single fluorescent residue there can be energy transfer among the residues, whch can appear as a component in the anisotropy decay. The timescale of energy transfer depends upon the distance and orientation between the residues, but there is little information on the timescale of energy transfer between intrinsic fluorophores in proteins. The measurements are different in the frequency-domain. In this case we measure the phase shift between the parallel and perpendicular components of the emission, and a frequency-dependent anisotropy, which is analogous to the steady state anisotropy. These two types of data are used to determine the decay law for the anisotropy (equation 15). Melittin and S, Nuclease illustrate how the anisotropy decay is reflected in the frequency-domain data. From earlier studies it was known that the single tryptophan residue in S, Nuclease was mostly rigid [36], so that its anisotropy decay should display a single correlation time for rotational diffusion near 11 nsec. In contrast, melittin monomer is thought to be disordered in aqueous solution, so that a rapid anisotropy decay is expected due to local tryptophan motions. The frequency-domain anisotropy data for nuclease and melittin are shown in Fig. 18. The data for nuclease are nearly Lorentzian and centered near 30 MHz, which is expected for a single correlation time near 11 nsec. In contrast, the differential phase data for melittin show no such maximum, and the phase angles increase up to the 200 MHz limit. This is characteristic of a subnanosecond anisotropy decay. For both proteins a multi-exponential anisotropy decay was necessary to explain the data, and in both cases a short correlation time ( < 1 nsec) was indicated by the data. In the case of nuclease only 12% of the anisotropy decays by this rapid process, indicating that the torsional motions have a limited amplitude. In contrast, 75% of the melittin anisotropy decays by the rapid process, which indicates considerable free motion of the tryptophan residue. To illustrate the nature of the anisotropy decays the equivalent time-dependent anisotropies are shown as an insert. These were calculated from the frequency-domain data. For S, Nuclease the plot of log r ( t ) versus time is mostly linear with a slope of (12 nsec)-'. This is the portion of the anisotropy decay due to overall rotational diffusion of the protein. The rapid component in the nuclease anisotropy decay is seen only near the t = 0 origin. The anisotropy decay of melittin is much more rapid, which reflects the greater motional freedom of the tryptophan residue in this disordered polypeptide. Because of the segmental motions which depolarize the

23

O

2

5

10

20

FREQUENCY

50

100

200 f

(MHz)

Fig. 18. Frequency-domain resolution of the anisotropy decay of S, Nuclease and melittin monomer. Melittin: r( t ) = 0.24 exp (- t/0.26) + 0.08 exp (- r/3.04). Nuclease: r( t ) = 0.04 exp (- r/0.20) + 0.28 exp ( - r/12.18).

emission the rotational diffusion of melittin is barely evident in the anisotropy decay.

4. Harmonic-content frequency -domain j7uorometry The frequency-domain data for melittin (Fig. 18) revealed the need for still higher modulation frequencies. Resolution of the anisotropy decay parameters is decreased if the phase angle maximum is not reached. This is perhaps analogous to the data obtained with flash lamps (Fig. 14), in which the width of the pulse was comparable to the decay times of the emission. Very recently, this laboratory developed a hybrid instrument which uses components typical of both time-domain and frequency-domain fluorometers [37]. The instrument uses a 4 MHz train of 5 psec pulses from a cavity-dumped dye laser, whch is the same source as is used for time-correlated single photon counting (Fig. 16). However, the pulses are not used to perform time-domain measurements. The pulse train possesses intrinsic modulation to many GHz, which is shown by the Fourier transform in Fig. 16. This source can be used directly as the modulated light source, an idea proposed originally for pulsed laser excitation by Merkelo et al. [46] and expanded to use the higher harmonics of pulsed synchrotron radiation by Gratton and Degado [48,49]. When used with a fast detector the frequency-domain measurements now extend to 2 GHz [37].

24

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.. .. ....... ._.-.-a_..

.

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. .. .,

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... . ... ..

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FREQUENCY (MHz) Fig. 19. Frequency-domain intensity decay of oxytocin at 20 O C. The symbols (0)represent the data, the solid line the best three-exponential fit, and the dashed line the best one-exponential fit. The lower panels show the deviations between the data and the calculated values for the one (A) and three (0) decay time models. The values of xk are 377, 5.9 and 2.1 for the 1, 2 and 3 exponential fits, respectively.

6

I

0

/'

I

20

I 1 1 1 1 1 1

50

100

I

200

I

I

I

I I , , ,

500

I

1000 20'0

FREQUENCY (MHz) Fig. 20. Anisotropy decay of oxytocin tyrosine fluorescence. The data (0) could not be fitted using a ). The single correlation time (- - -), but were adequately fitted using two correlation times (values of x i are 292, 3.3 and 3.4 for the 1, 2 and 3 correlation time fits, respectively.

25

This development is recent, and harmonic-content data are not yet available for the proteins described above. However, the potential of the 2 GHz measurements is illustrated by the data for oxytocin, which is a cyclic nona-peptide containing a single tyrosine residue. The frequency-response for the intensity decay is shown in Fig. 19. The mean decay time for oxytocin is near 0.7 nsec. Even with this short decay time the entire frequency response was measured, as seen by phase angles which extend to 70" and modulations which decrease to 20%. These data are adequate to support a three exponential analysis. The apparent decay times are 80, 359 and 927 psec. Once again, we find that even the decay of a single tyrosine residue can be complex. The 2 GHz data provide resolution of complex anisotropy decays on the picosecond timescale. Data for the anisotropy decay of oxytocin are shown in Fig. 20. It was not possible to fit the data using a single correlation time (- - -, xi = 292). In contrast, a two-correlation time model provides a good fit, which is not improved by the use of a third correlation time. We believe the correlation times of 29 and 454 psec reflect local tyrosyl motions and overall rotational diffusion, respectively. It is important to note that the measurements to 2 GHz provide considerable information beyond the data to the previous 200 MHz limit. Data to 200 MHz would not display the shoulder seen at 600 MHz, which represents the transition from rotational diffusion to segmental motions.

5. Summary The phenomenon of fluorescence can provide information about the physical properties of proteins and other macromolecules. The information content results from the sensitivity of the spectral properties to the average and dynamic properties of the environment surrounding the fluorescent residues. In general, more detailed information is obtainable from time-resolved data than from steady-state measurements. However, the steady-state measurements are considerably easier to perform. At present, the ability to recover time-resolved spectral data is rapidly improving, primarily because of advances in instrument design. The newer instruments may possess resolution adequate to correlate experimental data with the structural or dynamic properties of macromolecules.

Acknowledgements This work was supported by grant DAAG29-85-G-0017 from the Army Research Office, grants DMB-8511065 and DMB-08502835 from the National Science Foundation and grants GM-39617 and GM-29318 from the National Institutes of Health. The author wishes to especially acknowledge the National Science Foundation for providing support to construct the frequency-domain fluorometer, at a time when there were doubts about their design and usefulness.

26

References 1 Weber, G. (1952) Biochem. J. 51, 155-167. 2 Haugland, R.P. (1983) In: Excited States of Biopolymers (R.F. Steiner, ed.) pp. 29-58, Plenum Press, New York. 3 Beddard, G.S. and M.R. West, eds. (1981) Fluorescent Probes. Academic Press, New York. 4 Steiner, R.F. and Y. Kubota (1983) In: Excited States of Biopolymers (R.F. Steiner, ed.) Plenum Press, New York. 5 Jablonski, A. (1935) Z. Phys. 94, 38-46. 6 Lakowicz, J.R. (1983) Principles of Fluorescence Spectroscopy. Plenum Press, New York. 7 Ross, J.A.B., C.J. Schmidt and L. Brand (1981) Biochemistry 20, 4369-4377. 8 Lakowicz, J.R. and H. Cherek (1980) J. Biol. Chem. 255, 831-834. 9 Talbot, J.C., J. Dufourcq, J. DeBong, J.F. Faucon and C. Lussan (1979) FEBS Lett. 102, 191-193. 10 Faucon, J.F., J. Dufourcq and C. Lussan (1979) FEBS Lett. 102, 187-190. 11 Terwillinger, T.C. and D. Eisenberg (1982) J. Biol. Chem. 257, 6016-6022. 12 Eftink, M.R. and C.A. Ghiron (1981) Anal. Biochem. 144, 199-227. 13 Eftink, M.R. and C.A. Ghiron (1976) Biochemistry 15,672-680. 14 Lakowicz, J.R. and G. Weber (1973) Biochemistry 12,4161-4170. 15 Lakowicz, J.R. and G. Weber (1973) Biochemistry 12, 4171-4179. 16 Forster, Th. (1948) Ann. Phys. (Leipzig) 2, 55-77. Translated by R.S. Knox. 17 Stryer, L. (1978) Annu. Rev. Biochem. 47, 819-846. 18 Stryer, L. and R.P. Haugland (1967) Proc. Natl. Acad. Sci. USA 58, 719-726. 19 Dale, R.E., J. Eisinger and W.E. Blumberg (1979) Biophys. J. 26, 161-194. 20 Hermetter, A. and J.R. Lakowicz (1986) J. Biol. Chem. 261, 8243-8248. 21 Hanson, D.C., Y. Yguerabide and V.N. Schumaker (1981) Biochemistry 20, 6842-6852. 22 Brand, J.C. and R.H. Cagen (1977) Biochim. Biophys. Acta 497, 178-187. 23 Dale, R.E., L.A. Chen and L. Brand (1977) J. Biol. Chem. 252, 2163-2169. 24 Kawato, S., K. Kinosita and A. Ikegami (1977) Biochemistry 16, 2319-2324. 25 OConnor, D.V. and D. Phillips (1984) Time-Correlated Single Photon Counting. Academic Press, New York. 26 Demas, J.N. (1983) Excited State Lifetime Measurements. Academic Press, New York. 27 Lakowicz, J.R. and B.P. Maliwal (1985) Biophys. Chem. 21, 61-78. 28 Gratton, E. and M. Limkemann (1983) Biophys. J. 44, 315-324. 29 Visser, A.J.W.G., ed. (1985) Anal. Instrum. 14, 193-566. 30 Lakowicz, J.R., G. Laczko and I. Gryczynski (1986) Rev. Sci. Instrum. 57, 2499-2506. 31 Libertini, L.J. and E.W. Small (1985) Biophys. J. 47, 765-772. 32 Gurd, F.G. and T.M. Rothgeb (1979) Adv. Prot. Chem. 33, 73-165. 33. Karplus, M. and J.A. McCammon (1981) CRC Crit. Rev. Biochem. 9, 243-249. 34 Lakowicz, J.R. (1986) Methods in Enzymology, in press. 35 Lakowicz, J.R., G. Laczko, I. Gryczynski and H. Cherek (1986) J. Biol. Chem. 261, 2240-2245. 36 Munro, I., I. Pecht and L. Stryer (1979) Proc. Natl. Acad. Sci. USA 76, 56-60. 37 Lakowicz, J.R., G. Laczko and I. Gryczynski (1986) Rev. Sci. Instrum. 57, 2499-2506. 38 Lakowicz, J.R., G. Laczko and I. Gryczynski (1986) Biophys. Chem. 24, 97-100. 39 Lakowicz, J.R. and Balter, A. (1982) Photochem. Photobiol. 36, 125-132. 40 Jablonski, A. (1960) Bull. Acad. Pol. Sci. 8, 259-264. 41 Bevington, P.R. (1969) Data Reduction and Error Analysis for the Physical Sciences. McGraw-Hill, New York. 42 Svelto, 0. (1982) Principles of Lasers. Plenum Press, New York. 43 Visser, A.J.W.G., ed. (1985) Analytical Instrumentation 14. Marcell Dekker, New York. 44 Ludescher, R.D., Volwek, J.R., DeHaas, G.H. and Hudson, B.S. (1985) Biochemistry 24, 7240-7249. 45 Albana, J., Albert, B., Krajcarski, D.T. and Szabo, A.G. (1985) FEBS Lett. 182, 302-304. 46 Merkelo, H., Hartman, S.R., Mar, T., Singhal, G.S. and Govindjie (1969) Science 164, 301-302. 47 Eftink, M. and Ghiron, C.A. (1976) J. Phys. Chem. 80, 486-493. 48 Gratton, E. and Degado, L.R. (1980) Nuovo Cimento 56, 1110-1124. 49 Gratton, E. and Delgado, L.R. (1979) Rev. Sci. Instrum. 50, 789-790.

A. Neuberger and L.L.M. Van Deenen (Eds.) Modern Physical Methoak in Biochemistry, Part B 0

27

1988 Elsevier Science Publishers B.V. (Biomedical Division)

CHAPTER 2

Raman and resonance Raman spectroscopy * P.R. CAREY Division of Biological Sciences, National Research Council of Canada, Ottawa, Ontario, Canada KIA OR6

1. Introduction When a beam of monochromatic light is incident on, for example, a liquid a small portion of the light undergoes a change in wavelength. That portion which changes wavelength is known as the Raman spectrum after the Indian scientist C.V. Raman who with his colleague K.S. Krishnan first reported the effect [l].In the biochemical context Raman spectroscopy is used principally to monitor the vibrational motions of atoms within a molecule. Since the vibrational spectrum is sensitive to the molecule’s conformation and environment the Raman spectrum is used to probe the detailed chemistry of the biochemical molecule under study [2-41. The physical origin of Raman scattering lies in inelastic collisions between the molecules composing the liquid and photons which are the particles of light making up the light beam. An inelastic collision means that there is an exchange of energy between the photon and the molecule with a consequent change in energy, and hence wavelength, of the photon (Fig. 1). Moreover, since total energy is conserved during the scattering process the energy gained or lost by the photon must equal an energy change within the molecule. It follows that by measuring the energy gained or lost by the photon we can probe changes in molecular energy. The changes in the molecule’s energy are called transitions between molecular energy levels. As already mentioned, in biochemical studies Raman spectroscopy is concerned primarily with the molecule’s vibrational energy level transitions although the resonance Raman effect also provides detailed information on electronic energy levels. Although present-day Raman spectroscopy uses high technology-based instrumentation the experimental technique is simple in conception and can usually be depicted as in Fig. 2. A monochromatic laser beam of wavelength, A, is focussed into the sample to produce a high photon density and the resulting scattered light, which includes the Raman spectrum, is analysed for wavelength and intensity. The Raman effect is extremely weak and only a minute proportion of the incident * Published as NRCC Number 28, 789.

28

Molecule w

incident photon wavelength A, Scattered photon wavelength A,

For Raman scattering

A,

A,

Fig. 1. An inelastic collision between a photon and a molecule. From [2].

photons become Raman photons of wavelengths A,, A,, etc. The inherent weakness of the effect means that relatively high-power lasers must be used to create a high photon flux and that sophisticated optical and electronic equipment is required to detect the scattered photons. The normal Raman scattering process is improbable and gives rise to a weak Raman spectrum. However, there is a condition under which the spectrum can be greatly intensified, as a result of a phenomenon known as the resonance Raman (RR) effect. The Raman spectrum, obtained using an excitation wavelength of 488 nm, of the dithioester methyl dithioacetate is illustrated in Fig. 3. The spectrum has peaks whch contain contributions from the stretching motions of the C-H, C = S respectively. This Raman spectrum, although and C-S bonds, vC-H, vC+ and easily recorded using modern equipment, is extremely weak. However, under the resonance condition considerable intensity enhancement can occur. This condition is also illustrated in Fig. 3; the dithioester group has an intense electronic absorption transition near 302 nm and using an excitation wavelength (e.g. 324 nm) that lies under the absorption band a marked increase in Raman scattering is observed. Peaks occur in the same positions (usually denoted on a cm-' scale as in infrared spectroscopy) in both the Raman and the RR spectra, but only peaks which are

-

/ scattered light analysed forwavelength,

sample

_C

A, A2A3etc, and intensity

\ \ collecting lens focussing lens

laser beam wavelength A ,

Fig. 2. Schematic of a Raman experiment. From [2].

29

t S

H,C-C,

4

S-CH,

Resonance Ramon spectrum

1

f w=s "C-H "c-s FExciting line 488 nrn I

;

1

300

I

400

I

I

500 NANOMETERS

c

Fig. 3. The main features in the absorption, resonance Raman and normal Raman spectra of methyldithioacetate showing their relative positions on the wavelength scale. The choice of the 488 nm exciting line is arbitrary since any wavelength longer than 400 nm would yield a normal Raman spectrum. From

PI. intimately connected with the electronic absorption process are more intense under resonance conditions. For example, in the Raman spectrum shown in Fig. 3 three features are evident; however, in the RR spectrum it appears that only two remain. This is because the v,--~ and v , - = ~features are associated with the electronic transition at 302 nm and are thus intensity enhanced by approximately 2000-fold compared to the normal Raman case. However, the C-H bonds are not associated with the electronic transition and the v , - - ~feature is not intensity enhanced. This results in the relatively weak v ~ being - ~'lost' in the background noise of the RR spectrum. A major advantage of Raman spectroscopy for the analysis of biomolecules stems from the fact that water has a weak Raman spectrum. This enables us to record spectra of aqueous solutes at 10-'-10-2 M with little interference from the solvent. For a chromophore under the RR condition the concentration range becomes 10-4-10-6 M. Moreover, the intensity enhancement associated with the RR effect confers the important advantage of selectivity, allowing one to observe selectively the vibrational spectrum of a chromophore which is just one component of an extremely complex biological system. Other advantages are that Raman and RR spectra can be recorded using small amounts of material (e.g. 10 pl of a liquid) and that most materials such as liquids, gases, fibers and crystals are amenable to analysis.

2. The units used in Raman spectroscopy The units employed in Raman spectroscopy can best be explained by reference to an example. The Raman spectrum of water, the biological solvent, is shown in Fig.

30

,

I

500

NANOMETERS

525

550

575

600

> k

m

z + W

2

I

1

I

RAMAN SHIFT (cm-1)

a00

I9500

18500

I7500

I6500

ABSOLUTE (cm-1)

Fig. 4. A Raman spectrum of water.

4. The spectrum was obtained in the manner illustrated in Fig. 2 using a laser beam

of wavelength 488 nm. The most readily discernable peaks are the intense feature marked 0-H stretch and the weak feature marked H-0-H bend. These Raman peaks arise, respectively, from an exchange of energy between the incoming photons and a vibrational motion corresponding to a stretching of the 0-H bond or a bending motion of the water molecule.

In Fig. 4 the abscissa represents an energy difference between the scattered and incoming photons with the zero point energy difference being at the wavelength of excitation, marked 488 nm. It is therefore apparent that more energy is required to bring about the 0-H stretching motion than the H-0-H bending motion. It is also apparent from the intensities of the peaks in the spectrum that the exchange of energy between molecules and photons giving rise to the 0-H stretching peak is more probable than that giving rise to the H-0-H bending feature. A monochromatic light beam is characterised by its wavelength, A, its power and its polarisation. Instead of quoting the wavelength of a light beam this property is often given in terms of the equivalent units of frequency or wavenumber. These quantities are related to wavelength thus: Frequency = v

C

=

- Hertz (or cycles per second)

h

(1)

31

where c is the speed of light (2.998 X 10" cm sec-' in vacuum) and A must be expressed in the same units as c. 1 Wavenumber (often denoted w or u) = - cm-'

A

(2)

where A is expressed in centimeters. The wavelength of light is commonly expressed in Angstroms (A) or nanometers (nm) 10 A = 1 nm = 1 0 - ~meter

(3)

Finally, the wavelength of light is related to the energy, E , by using E=hv

(4)

where h is Planck's constant. The units described in Fig. 4 are now clear. The wavelength of the laser line used to excite the Raman spectrum is 488 nm which is equivalent to 20,492 cm-'. When discussing energies it is simpler to use wavenumbers since, from equation (4), energy is linear with wavenumber (and therefore proportional to 1/ wavelength). The exchange of energy between the incoming photons and the vibrational energy transitions giving rise to the H-0-H bending and 0 - H stretching features results in a loss in energy for the photons equivalent to 1640 and 3400 cm-', respectively. Thus, on the scale of absolute cm-' the H-0-H bending and 0-H stretch appear at 20,492-1640 = 18,852 cm-' and 20,492-3400 = 17,092 cm-', respectively, and this can be seen by referring to the bottom scale in Fig. 4. The usual convention in Raman spectroscopy is to quote the positions of the vibrational peaks as the difference between the absolute wavenumbers of the exciting line and the absolute wavenumbers of the scattered photon. Thus, the Raman shift scale seen in Fig. 4 is normally the only scale encountered in published Raman spectra. The equivalent wavelength scale in nanometers is shown at the top of Fig. 4 and the nonlinearity of the top scale reflects the fact that spectrum is plotted as a linear function of energy but that energy is proportional to the reciprocal of wavelength.

3. A model for Raman scattering based on classical physics A light wave is a travelling wave of electric and magnetic fields, of which only the electric component gives rise to Raman scattering. When a light wave meets a molecule consisting of electrons and nuclei the electric field will exert the same force on all electrons in the molecule and will tend to displace them from their average positions around the positively charged nuclei. Crucially, for the Raman process the displacements result in an induced dipole moment B in the molecule which is, to a good approximation, proportional to the electric field strength E . Thus Ir=(YE

(5)

32

where the proportionality factor a is called the electric polarisability of the molecule. In general the vector IT will have a different direction from that of the vector E , and therefore a is not a simple scalar quantity. In fact, a is a tensor but in the following simplified treatment we will be dealing with the relationship between just one component of the tensor, a,, and the components of IT and E in the z direction. The electric field of the light wave varies with time. If a fixed molecule is irradiated with monochromatic radiation of frequency vo, expressed in Hertz, which is plane polarised in the z direction, E, as a function of time is given by E , ( t ) = E m , cos 2.rrvot

(6)

where Em, is the value of E, at its maximum and f is the time in seconds from an arbitrary starting time. Thus, for the z component of IT .rr,(t) = aZZEm,cos 27rv0t

(7)

Since .rr, depends on a,= as well as E, the properties of the molecule can change In the present context a varies with time as a consequence of the vibrations of the molecule, since the ease with which electrons may be displaced by the electric field depends on how tightly they are bound to the nuclei, which in turn depends on the internuclear separation. The result of the time dependence of a and E on IT can be seen by considering the simple example of a single diatomic molecule. For a diatomic molecule the difference from equilibrium in the internuclear distance at time t , Ar(t) can be written rZvia a.

Ar( t ) = Arm,(2.rrv,,t

+ cp)

(8)

where vVib is the vibrational frequency of the molecule in Hertz, and Arm, is the maximum extension of the distance between the two atoms. By taking the time t to have the same starting point as the time scale for the light wave in equation (6), the phase constant cp may be equated to zero. Using the postulate that the polarisability of the diatomic molecule depends linearly on Ar, the azzcomponent of a can be written: a,,(t) =

dazz + -Ar(t) dr

The constant ( Y ~ ~is' the polarisability element of the nonvibrating molecule and da,,/dr characterizes the manner in which the polarisability changes with r .

33

Substituting the expression for a,, into equation (7) the dependence of time fluctuations of both a and E becomes:

on the

I I

Using the identity cos 6 cos + = B[cos(O + +) + cos(6-@)] equation (10) becomes:

Equation (11) demonstrates that when a light wave interacts with a vibrating diatomic molecule the induced dipole moment, in this case exemplified by n;, has three components contributing to its time dependence. The first term on the right of equation (11) is a component vibrating with the frequency of the incident light and with a magnitude determined by azi’ and Ern,. According to classical electromagnetic theory an oscillating dipole radiates energy in the form of scattered light. Thus, as a result of the first term in equation (11)light of the incident frequency v,, will be emitted and will be observable in directions which differ from that of the incoming light. This is the phenomenon of Rayleigh scattering. The second term is a component vibrating at a frequency which is the sum of the frequencies of the light and the molecular vibration. The scattered light arising from this second term is known as anti-Stokes Raman scattering. The third term is a component vibrating at a frequency given by that of the light wave minus that of the molecule and the scattered light resulting from this term is known as Stokes Raman scattering. Both these components have magnitudes depending on the field strength of the light, the amplitude of vibration and the polarisability derivative da,,/dr. The appearance of scattered radiation, e.g. at vo + VGb, in units of Hertz arising from the second term in equation (ll),or vo-vvib from the third term in that equation, means that by analysing the scattered light we can monitor the vibrations within a molecule. It is this ability to measure molecular vibrations which gives the Raman effect its importance in the study of molecules. The information about molecular vibrational frequencies provided by infrared absorption spectroscopy is of the same kind as that provided by the VGb values of the Raman lines. However, Raman and infrared spectroscopies originate from two distinct physical processes. The Raman process is a scattering effect involving an induced dipole, a,which, in turn, depends on a change in molecular polarisability during a vibration. In contrast, infrared spectroscopy is an absorption process caused by a change in the permanent molecular dipole, p , with change in bond length during a vibration. Both infrared and Raman spectra are required to provide a complete picture of molecular vibrations. But because of the complementary nature of the effects only one technique may be applicable in a given instance. For

34

example Raman spectroscopy is often favored for biochemical studies because water has a relatively weak Raman spectrum (Fig. 4). In contrast the infrared absorption of water is very hgh in most of the region of interest and this often obscures the infrared absorption of the biochemical solute.

4. Raman and resonance Raman scattering; a quantum mechanical

interpretation The quantum mechanical approach to the scattering process is quite different to the classical model: the light beam is considered to be made up of packets or quanta of light particles known as photons, the quantisation of molecular energy levels is taken into account, and a means is provided for calculating the polarisability a,and thus Raman intensities, in terms of the electronic properties of a molecule. It is a reasonable approximation, for a gaseous diatomic molecule, to write the molecular energy E m ,as a sum of terms:

where the respective subscripts refer to electronic and vibrational components of the total energy. For present purposes the contributions due to molecular rotation and translation can be ignored. Electronic energy transitions involve much larger quantities of energy than vibrational transitions, with values of 10,000-50,000 cm-' for the former and 10-4000 cm-' for the latter. This situation is depicted in Fig. 5 with a large energy spacing between the ground and excited electronic states, and smaller spacings between the vibrational levels contained within each state. Moreover, for the vibrational energy, E,,, of a diatomic molecule it is a good approximation to write

where the vibrational quantum number u has only integral values so that the vibrational energy levels in the ground state in Fig. 5 are equally spaced by the amount vvib. The energy E,,, of course, takes the units of v,b, Hertz if the latter is expressed in Hertz and so on. In the quantum mechanical model light scattering is depicted as a two-photon process. The first step in t h s process is the combination of a photon and a molecule to raise the molecule to a higher energy state of extremely short lifetime. This state is reached by the upward arrows in Fig. 5 and as shown in the figure, the higher energy state may or may not correspond to a quantised energy state of the molecule. The second step, indicated by the downward arrows in Fig. 5, involves the release of a photon after a very short time interval (< lo-" sec). The energy of this second photon is given by the length of the downward arrows in Fig. 5. For Rayleigh scattering the upward and downward transitions have the same length and have

35

Upper (excited) electronic stote

3

Lower (ground) electronic state

2 I Infrared Rayleigh Normal Roman

Preresonance Roman

Resonance Roman

Fluorescence

n "

J

Fig. 5. Some of the possible consequences of a photon-molecule interaction. The lengths of the upward-pointingarrows are proportional to the frequencies of the incoming light while the lengths of the downward-pointing arrows are proportional to the frequency of the scattered (or in the case of fluorescence, emitted) light. The vibrational quantum numbers in the upper and lower electronic states are v' and v" respectively. The energy spacing v" between the lower state vibrational levels is equal to vmb. From [2].

therefore, apart from a change in sign, the same energies. Thus, in the Rayleigh process, no change in frequency of the photon occurs. The various kinds of Raman processes may now be outlined. If the downward arrow stops on a vibrational energy level that is higher than the starting level a Stokes process has occurred. In this, the second photon has a frequency vo-vvib corresponding to the third term in equation (11) of the discussion of the classical model for the Raman effect. Conversely, an anti-Stokes process results from the transition terminating at a lower vibrational energy level compared to the starting level. In the anti-Stokes process the second photon has an energy vo + v,+ giving the same result as the second term in equation (11). Of course, in both processes total energy is conserved so for Stokes scattering the molecule gains a quantum of energy vvib while for anti-Stokes scattering the reverse is true. For both Stokes and anti-Stokes processes a selection rule can be derived from equation (13) which says that v", in Fig. 5, can only change by k l . Thus equation (11) derived from the classical model agrees with the results obtained by considering quantised energy levels in that both models predict the difference in frequency between the incident and scattered light, vo-( vo-vvib), corresponds directly to the molecular vibrational frequency vvib. The quantum mechanical model also illustrates an important generalisation namely: that the position of Raman peaks is a property solely of the electronic ground state. This follows from the fact that vvib is a vibrational transition within the lower or ground electronic state in Fig. 5. In the classical model, equation (11) indicates no difference in the expected intensities of Stokes and anti-Stokes transitions, since the coefficients of the two

36

terms in the equation are the same. However, the model of quantised energy levels depicted in Fig. 5 shows that for anti-Stokes transitions to take place the molecule must be in a hgher (u” > 0) vibrational state within the electronic ground state. Since the population of these higher states is governed by a Boltzmann distribution only a small percentage of molecules are in higher vibrational states. The ratio of the number of molecules in the u” = 1 and u” = 0 vibrational states in the ground electronic state (Fig. 5) which we will denote Nl and No respectively is given by Nl = exp( - hv,,/kT) -

NO

where h is Planck’s constant, T is the absolute temperature and k is the Boltzmann constant. When T = 300 O K and the vibrational frequency is 480 cm-’, Nl is 0.1 No. As a result of the exponential nature of equation (14) an anti-Stokes line at 3 X 480 cm-’, or 1440 cm-I, would be 0.001 times as strong as the corresponding Stokes line. In practice this means that the feeble anti-Stokes scattering is usually ignored in conventional Raman spectroscopy and only the Stokes spectrum is recorded. Having outlined normal Raman scattering in which the energy of the incident light is considerably less than that needed to reach the higher energy electronic state in Fig. 5 the result of the light beam energy approaching that of the energy gap between the lower and higher electronic states can now be considered. In Fig. 5 the transition labelled ‘preresonance Raman’ is due to a light frequency that has almost enough energy to produce direct electronic absorption by the molecule. Under this condition the intensity of Raman scattering shows a marked increase. For normal or non-resonant conditions Raman intensities are proportional to the fourth power of the scattered light frequency, vs. However, as preresonance Raman conditions are approached the intensity of scattering goes up much more rapidly than u,”. A slight increase in the energy of the exciting radiation over that for the preresonance case will place the upward transition in Fig. 5 within the higher electronic state. Absorption of a photon can now occur and by the prompt re-emission of a second photon can give rise to the RR process. Band intensities in resonance Raman spectra can be orders of magnitude greater than those in normal Raman spectra. The reason for this can be seen by considering the quantum mechanical expression for Raman intensities. In the quantum mechanical treatment for randomly oriented molecules, the total intensity of the scattered light resulting from a molecular transition between states m and n is:

where I a,,, I m n is the transition polarisability tensor. This quantity can be derived from second-order perturbation theory to give

37

In these expressions the molecule is considered to be in the molecular state m. It is perturbed by an electromagnetic wave of frequency vo and intensity I , causing the transition to a state n and scattering light of frequency v, f vmn.The sum over index r covers all of the quantum mechanical eigenstates of the molecule, h is Planck’s constant, and r,. is a damping constant which takes into account the finite lifetime of each molecular state. The ( n I p,, I r ) , etc., are the amplitudes of the electric dipole transition moments where p,, is the electric dipole moment operator along direction p. Immediately, we see from equation (16) that as vo approaches the energy of an allowed molecular transition vrm, (vrm - vo ir,)becomes small and consequently one term in the sum becomes very large. This is the resonance condition. The intensities of various features in the RR spectrum are sometimes measured as a function of the excitation wavelength (usually in reference to an internal intensity standard). The intensity variation of a Raman mode with excitation wavelength is known as the excitation profile and can reveal a considerable amount of information regarding the electronic absorption transitions giving rise to RR intensity enhancement [5,6]. An important practical outcome of the resonance Raman effect is that the accompanying intensity enhancement allows one to obtain the Raman spectrum of molecules with suitable electronic absorption bands at low concentration, e.g. in the l op5 M range in solution. At the same time the resonance Raman effect enables the spectroscopist to selectively and specifically ‘pick out’ the Raman spectrum of an absorbing molecule in a complex environment which only contributes a weak normal Raman spectrum. This property is used to great effect in biologcal Raman studies since the chromophore is often found at the site of biological activity.

+

5. Polarisation properties of Raman scattering Most natural light is unpolarised, which in a simplified form means the electric ( E ) vector performs linear oscillations of constant amplitude in a plane perpendicular to the light path but that the orientation of the E vector within that plane is completely random. Light from lasers has the special property of being plane polarised; that is the terminus of the electric vector varies periodically within a single plane through the light path. Thus, for the plane polarised beam shown in Fig. 6 the E vector remains in the X Z plane but varies in magnitude along the light path according to the hatched lines. In Fig. 6 the laser beam is depicted as a vertical line starting at the bottom of the diagram. Information on the scattering process and on the assignments of Raman bands can be gained by analysing the scattered light parallel and perpendicular to the incoming E vector. The depolarisation ratio, p, of a feature in the Raman spectrum is defined as

where I, and Il arelthe intensities of Raman radiation of a given frequency that is polarised, respectively, perpendicular and parallel to the plane normal to the

38

Ill

Fig. 6 . The orientations of the E vectors of the incoming ( E ) and scattered light parallel (Ill) and perpendicular (IL) to the incident E vector. The depolarization ratio p = I , /Il,.From [2].

incident beam. These relationships are illustrated in Fig. 6. Changes in the polarisation of the incident light upon scattering result from the tensorial nature of the interaction denoted in equation (5). In fact for a single crystal the individual For fluids, however, elements of the scattering tensor a may be related to I , and I,,. the molecules are randomly oriented with respect to the laboratory-fixed coordinate system used to define a and when an average over molecular orientations is made, I , and Illare found to be related to certain combinations of the components of the tensor a. Detailed calculations show that for normal vibrations which do not preserve molecular symmetry during the motion of the nuclei (non-totally symmetric modes) p = 3/4. For modes which do preserve molecular symmetry p 5 3/4 and, in fact, p is often found to be substantially less than 3/4 for totally symmetric vibrations. Under resonance or near resonance Raman conditions in certain rare cases it is possible to observe ‘anomalously’ polarised bands with p > 3/4. T h s phenomenon is discussed in the literature [7].

6. Basic experimental aspects In a present-day Raman experiment intense monochromatic radiation provided by a laser is focussed onto or into the sample. In Fig. 7 the laser beam enters the sample in the vertical plane with the direction of its E vector shown by double-headed

39 x

MIRROR

ENTRANCE SLIT ~

SOLID ANGLE

Y

SAMPLE CELL

I

LENS

\

FOCUSING ~-

Fig. 7. The optics about the sample for a conventional Raman experiment. From [2].

arrows, Some of the resulting scattered light is gathered by collecting optics (seen as a single lens in Fig. 7) and directed to a dispersing system which is usually a double monochromator. The function of the spectrometer is to spatially separate the scattered light on the basis of frequency. At the exit port of the spectrometer the Raman spectrum forms an image in the form of a series of very faint lines. These are detected and recorded either sequentially by a single photomultiplier used with a scanning monochromator or simultaneously by a multi-channel detector which is the modern electronic equivalent of a photographic plate. The two options are shown schematically in Fig. 8. The basis of the dispersing process is depicted in Fig. 8 by a single monochromator although in most Raman experiments the dispersing process is repeated by linking two single monochromators to form a double monochromator. A double monochromator is often necessary to separate the Raman photons from the overwhelming number of Rayleigh photons. As shown in Fig. 8 there are two main ways in which the line spectrum across the exit port may be detected. The first and most commonly used method is to use a scanning spectrometer and to place a narrow exit slit over the exit port, followed by a photomultiplier tube. By slowly turning the grating, using the accurate drive of the spectrometer, the lines of the spectrum move in succession across the slit and are detected and recorded as outlined in Fig. 8. In the second method a multichannel detector is placed at the exit port. This detector is akin to having several hundred minute photomultiplier tubes across the port. All the Raman lines are then registered on different elements of the detector all the

40

Collection optics and sample

Slll

€7.11

SPECTROMETER

I

II

I

MEASUREMENT

1

Grating /

Image of spectrum1

\Photoactive surface

\

DETECTOR

Fig. 8. Schematic of a Raman spectrometer showing the options of single-channel (scanning) and multi-channel detection. From [2].

time. Thus, it is possible to observe the entire Raman spectrum on a TV screen or an oscilloscope in real time. When using multichannel detection the grating is turned only to change the spectral region across the detector.

7. Raman studies on biological materials Both the Raman and resonance Raman effects have been used extensively in biochemical studies [2-41 and it would be possible to divide the discussion under these two main headings. However, a more useful approach is to treat each class of biological material in turn and to show how Raman and resonance Raman experiments can provide different and often complementary information on its chemical properties. The three main classes of biological molecules to be considered are proteins, nucleic acids and lipids and membranes. 7.1. Proteins 7.1.1. Amide I and amide III features As can be seen in Fig. 9, the observed Raman spectrum of a protein recorded under non-resonance conditions consists of contributions from various amino acid side chain modes together with modes originating from the peptide backbone. Among the latter the amide I and amide I11 modes are widely used to characterise the secondary structure of the peptide backbone. The amide I modes have a high degree of C=O stretchmg character while the amide I11 modes have a large contribution from N-H in-plane bending. The use of the amide I and I11 regions in the Raman spectrum to characterise the secondary structure of a protein depends, in part, on determining characteristic frequencies for helical, P-sheet, p-turn and random protein conformations. Initially, the most widely used means of doing this was to use polypeptide models and proteins of known conformation [8]. The ranges within

41 T.0

Lysozyme Crystals

-

cm-' Fig. 9. A Raman spectrum of a protein, viz. the enzyme lysozyme in the crystalline phase. From [81].

which the amide I and amide I11 Raman bands occur for a-helical, P-sheet and unordered structures are given in Table 1. More recently the assessment of protein secondary structure from the amide I and I11 regions of the Raman spectrum has been made by a number of different approaches. In one of these, PCzolet and co-workers [9] have proposed a means of estimating the amount of P-sheet structure by using the relative intensity of the peaks at 1240 and 1450 cm-', due to the amide 111 and CH, bending modes (from the many CH, groups in the protein) respectively. The method is only applicable when there is a well-defined amide 111 band at 1240 3 cm-'. The 1450 cm-' feature is used, in Pkzolet's approach, as an internal intensity standard and this idea is common to the second method developed by Lippert et al. [lo]. In their method the relative intensities at 1240, at 1632 and at 1660 cm-' are taken to be a linear combination of contributions from a-helix, P-sheet and random coil conformations. The percentages of these conformations are derived from a set of four linear equations involving the intensity measurements as input data. The third method proposed by Williams [ l l ] for delineating the secondary structure of proteins analyzes the amide I band of a protein in terms of a linear TABLE 1 Approximate positions (cm-') of the most intense amide I and 111 bands in Raman spectra for various polypeptide conformations

a-Helix P-Sheet Unordered a

Amide I

Amide I11

1645-1 660 1665-1 680 1660-1 670

1265-1 300 1230-1 240 1240-1 260

Weak and may be confused with side-chain modes.

a

42

combination of amide I bands of proteins whose secondary structure is known. For 14 proteins analyzed by removing each one from the reference sets and analysing its structure in terms of the remaining proteins, the average correlation coefficients between the Raman and X-ray diffraction estimates of helix, P-strand, p-turn and undefined were 0.98, 0.98, 0.82 and 0.35, respectively. 7.1.2. Side chain contributions to the Raman spectrum 7.1.2.1. The disuljide group and -SH groups A relatively intense Raman line near 500-550 cm-’ is observed from the stretching mode, v ~ - of~ ,a disulfide linkage, CCS-SCC; and this feature can be seen in the spectrum of lysozyme shown in Fig. 9. The vs-s band can be used to follow changes in conformation about the SS linkage [2-41 (and see [12] for a recent discussion) and also, to follow the formation or breakdown of S-S bonds in proteins. Both cysteine (disulfide) and methionine groups in proteins give rise to C-S stretching vibrations in the 600-750 cm- regions. Hence, correlations proposed between vc-s and conformation within the -C-C-Smoiety in disulfide and methionine are easier to apply when only one group is present. The C-S stretching frequency has been suggested, on the basis of studies on dialkyldisulfides, to lie near 630-670 cm-’ when the site (X), trans to the sulfur atom, in

’

H

x-c- c -ss H

is occupied by a hydrogen, and 700-745 cm-’ when X is a carbon atom [13,14]. Contributions from C-S bonds are identified near 663, 696 and 721 cm-’ in the Raman spectrum of lysozyme (Fig. 9). The SH stretching band of cysteine residues occurs for proteins in the 2550-2585 cm-’ range. In one interesting application the intensity of the -SH feature has been used to follow the conversion of -SH to S-S linkages during the aging process of the lens from the eye of a rat [15]. 7.1.2.2. Tyrosine uibrations Tyrosine residues contribute peaks to the Raman spectra of proteins at 1617, 1210, 1180, 850 and 830 (the latter two form the so-called tyrosine doublet) and 645 cm-’; some of these features can be seen in Fig. 9. It was suggested by Yu et al. [16] that the ratio of the intensities of the tyrosine ring vibrations at 850 and 830 cm-’ (850/830 = R T y r )reflects ‘buried’ and ‘exposed’ tyrosines. Subsequently, it was shown that the doublet arises from Fermi resonance between a ring-breathing vibration and the overtone of an out-of-plane ring-bending vibration [17] and that the intensity ratio is sensitive to the nature of hydrogen bonding or state of ionization of the phenolic hydroxyl group. If the OH group is strongly bound to a negative acceptor RTyr is near 0.3. If the OH forms moderately strong hydrogen bonds to H,O, RTyr is approximately 1.25 and if the OH participates as a strong hydrogen bond acceptor RTy, is close to 2.5. For the -0- form of the phenol side chain RTyr is near 1.5. The doublet is seen in the spectrum of lysozyme at 836 and 856 cm-’ (Fig.9).

43

7.1.2.3. Ttyptophan modes Tryptophan’s 3-substituted indole ring can give rise to features near 1622, 1578, 1555, 1361, 1342, 1016, 880 and 762 cm-’ in the visible light excited Raman spectra of proteins and some of these features can be seen in Fig. 9. A sharp line at 1361 cm-’ has been suggested as an indicator of buried tryptophan residues [18]. Thus, the presence or absence of a line in this region in the spectrum of proteins containing tryptophan suggests buried or exposed tryptophans. In human carbonic anhydrase B whose four tryptophan residues are known to be buried, the Raman spectrum does indeed show a sharp line at 1363 cm-’ [19]. A line at 1386 cm-’ from deuterated tryptophan can be used to follow the kinetics of H/D exchange for tryptophan residues in proteins exposed to D,O. The rate of exposure of the tryptophan to the solvent due to the ‘opening and closing’ of the protein structure can thereby be calculated [20]. 7.1.2.4.Histidine vibrations The imidazole side chain of histidine often plays an important role in protein function. Unfortunately characteristic imidazole modes are rarely seen in protein Raman spectra. An extensive discussion of imidazole modes is given by Harada and Takeuch [12]. 7.1.2.5.Acid groups Although the -COOH moiety has a group frequency in the 1700-1750 cm-’ region, features in this part of the spectrum are rarely reported in the Raman spectra of proteins. However, a mode due to the ionised side chains, -COO-, is sometimes seen near 1417 cm-’ in the Raman spectra of proteins containing a hgh percentage of acidic amino acids. 7.1.3. Applications In recent years Raman spectroscopy has been used extensively to study the protein components found in the lens of an eye and the protein (and nucleic acid) components of virus particles. The reader is referred to the work of Yu et al. [15] and Ozaki et al. [21] concerning lens proteins and to section 7.5 for a discussion of the work of Thomas and associates [22] on viruses.

7.1.4. UV excited resonance Raman spectra of proteins After pioneering work using 250-350 nm excitation (see [2], pp. 96-98, for a summary), the mid 1980’s has seen a surge of interest in the 200-230 nm UV range excited RR spectra of proteins. Pulsed laser light in this regon is usually generated by nonlinear frequency doubling and mixing of a dye laser output or doubled output with the 1.06-pm YAG fundamental; with a H, or D, Raman shifter being used to reach the lower wavelengths. The present high activity in the 200-230 nm UV excited RR spectra of protein presages many new findings. At the present time the following generalisations are emerging: 1. The excitation range below 230 nm is better than the 230-300 nm region from the point of view of having minimal interference from fluorescence. 2. Tryptophan and tyrosine chromophores can be selectively excited by 218 or 200 nm irradiation, respectively. 3. Amide I, 11, and I11 features are seen in the UV excited RR spectra, unlike the normal Raman case where amide I1 bands are usually too weak to be observed.

44

4. Care has to be taken to control unwanted photochemistry caused by the short-lived, intense UV excitation pulses [23]. Up to mid-1985, the UV-laser excited Raman spectra have been reported for peptide backbone model compounds acetamide and N-methylacetamide [24], insulin and a-lactalbumin (218 and 200 nm; [25]), myoglobin (230 nm; [26]), hemoglobin (218 and 200 nm; [27]), and cytochrome c (218 and 200 nm; [28]). 7.2. Proteins containing a natural, visible chromophore

Most protein-bound chromophores absorbing light in the 350-700 nm region have been the subject of RR study. There is extensive literature concerning RR investigation of proteins containing hemes, visual pigments and their bacterial analog bacteriorhodopsin, metal ions, flavins, chlorophylls, and carotenoids. Table 2 summarises the kind of information obtainable for each class of chromophore. In every case the RR spectrum contains detailed chemical information on the chromophore and usually the RR spectrum is uncluttered by contributions from the protein matrix or other species present. In a sense the RR spectrum can be regarded as a vibrational (and highly informational) counterpart of an electronic absorption spectrum and the concentrations used to obtain RR and absorption spectra are normally in the same range as those used to obtain routine absorption spectra, typically 10-4-10-6 M in chromophore. The discussion in the rest of this section will focus on some aspects of heme protein-RR spectroscopy. Hemes are by far the most widely studied chromophore and can be used to illustrate the degree of detail which may be elicited from RR spectra. A typical heme absorption spectrum is shown in Fig. 10. RR spectra can be generated by exciting into the Soret region near 400 nm or into the a-p bands in the 500-570 nm wavelength range. The structure of RR spectrum depends markedly on the excitation region employed. Excitation into the a-/3 region produces intense RR bands which are either depolarised ( p = 0.75) or anomalously polarised ( p > 0.75). Some early high quality RR spectra, excited in or near the /3 electronic transition, of a cytochrome and a hemoglobin are shown in Fig. 11. The spectra are dominated by peaks in the 1100-1650 cm-' region, because these peaks are due to the in-plane stretching of C-C and C-N bonds and the bending of C-H bonds which are effective in vibronically mixing the in-plane a and Soret electronic transitions to create the j3 band. Several peaks in Fig. 11have high utility as marker bands. For example the so-called band I, which occurs between 1358 and 1377 cm-' can be used as an oxidation state marker for the Fe atom. Band IV, which occurs between 1552 and 1590 cm-' is easily recognised by its high intensity and anomalous polarisation (Fig. 11). Band IV has great value in that its position correlates with the Fe-porphyrin core size as defined by the Fe-N (porphyrin ligand) distance [29]. These and other marker bands have been used to follow the details of chemical changes in a wide variety of heme proteins under a wide variety of conditions [30]. Excitation into the Soret band often enables one to detect RR bands in the lower frequency range, 100-800 cm- which are associated with Fe-axial ligand motions.

',

45

In one study Kitagawa and colleagues [31] identified the Fe-histidine stretching peak in the 214-225 cm-' region for various mutant deoxyhemoglobins. Separate oxygen binding experiments enabled them to obtain K,, the 0, equilibrium binding TABLE 2 A summary of resonance Raman studies on natural chromophores in the visible spectral region Biochemical class

Examples

Chromophore

Typical information

Hemes

Hemoglobin a Myoglobin Cytochrome c a,c Cytochrome P-450 Horseradish peroxidase Cytochrome oxidase

Usually porphyrin moiety, occasionally charge transfer transition involving porphyrinbound metal may contribute

Fe spin and oxidation state 8 Fe-N (from tetrapyrrole) core size g Fe-axial ligand chemistry 8 Protein and poryphyrin relaxation following photolysis of Fe-CO Quaternary structure transitions '

Visual pigments and bacteriorhodopsin

Rhodopsin Bacteriorhodopsin Halorhodopsin

Retinal, covalently linked to protein as Schiffs base

For each intermediate in photocycle: conformation of polyene and state of protonation of Schiffs base

Metalloproteins

Hemerythrin m.n Hemocyanin Tyrosinase Ribonucleotide reductase O Ferridoxin Andrenodoxin Rubredoxin 'Blue' copper Transferrins and other Fe(II1)-tyrosinate proteins "

Metal ligand(s) chargetransfer transition

Identification and coordination geometry of metal ligands

Flavoproteins

Yeast fatty acyl-CoA oxidase ' Porcine liver fatty acylCoA dehydrogenase Yeast glutathione reductase ' Egg-white flavoprotein " Old yellow enzyme " D-aminO acid oxidase NADPH-cytochrome P450 reductase '

Flavin

Type of bonding between flavin and protein. Chemistry of charge-transfer complexes involving flavin and second ligand (e.g. phenol " or amino-acid derivative ")

J

'

Table 2 continued on next page.

Ir

46

TABLE 2 (continued) Biochemical class

Examples

Chromophore

Typical information

Chlorophylls

Chlorophylls Chloroplasts Algae Reaction centers

Tetrapyrrole

Selectively obtain RR spectra of e.g. chlorophyll a and b in situ. Nature of binding site in protein. Mg coordination number

Carotenoids

Many in vivo situations bb e.g. photosynthetic membranes '' lobster shells

Polyene

Polyene conformation; polyene-polyene exciton interactions; membrane potential; triplet state properties ee

Phytochrome

Isolated bile pigments

Biliverdin dimethyl ester

Conformation

f f

For references see pp. 63-64.

constant for each mutant hemoglobin. Kitagawa et al. were then able to show that v ~ correlates ~ remarkably ~ ~well with ~ K, (Fig. 12). The variation in v ~ was ~ attributed to strain imposed on the Fe-histidine bond by the globin. Measurements such as these provide key evidence on the mechanics of hemoglobin function. In another study involving excitation into the Soret band Yu and his co-workers [32] have challenged the detailed X-ray crystallographic analysis of a heme protein. A fragment of the 406.7 nm excited RR spectrum of CO bound to the monomeric

Y

A (nm)

Fig. 10. Absorption spectrum of cytochrome c in the reduced form.

-

-

0-

h

1.

Y

h

INTENSITY

0 v

41

B 6'

5a

E. *

Fig. 11. Resonance Raman spectra of (a) ferrocytochrome c , 0.5 mM obtained with 514.5 nm excitation and (b) oxyhemoglobin, 0.5 mM, obtained with 568.2 nm excitation. From [7b].

hemoglobin I11 from Chironomzrs thummi thummi is shown in Fig. 13. The isotopic substitutions identify the Fe(I1)-CO stretching mode at 500 cm-' and the Fe(I1)-C-0 bending mode at 574 cm-'. Similarly in the complex involving CN-, Fe(II1)-C-N- stretch and Fe(JI1)-C-N- bend were assigned at 453 cm-' and 412 cm-', respectively. The RR data, in conjunction with those obtained from heme model complexes with known Fe-C bond distances, suggested strongly that the

48

K,/(mmHg)

Fig. 12. Correlation between the Fe-His stretching frequency, mutant deoxyhemoglobins. From [31].

Y,

and oxygen affinity K , for various

A)

Fe(II1)-CN- bond ( - 1.91 is longer (hence weaker) than the Fe(I1)-CO bond This result is at variance with the X-ray crystallographic findings of 2.2 and 2.4 A for the Fe-CN and Fe-CO bonds, respectively; leading Yu to suggest that there may have been errors in the X-ray crystallographic refinement. ( - 1.80

A).

7.3. Resonance Raman labels The advantages of the RR effect are that it provides specificity and selectivity in very complex biological systems at chromophore concentrations in the 10-3-10-6

*

Hb CTT 111. CO

407

489

I

I

300

400

cm-'

I

I

500

600

Fig. 13. CO isotope effects on the resonance Raman spectrum of a heme protein with a CO molecule bound to the porphyrin Fe atom. From [32].

49

M range. However, many sites of high biological interest do not have suitable chromophores to act as RR probes. For these, the RR labelling technique was developed [33]. A RR label, usually mimicking a natural component, is introduced into the system as a reporter group and there has been success in using chromophoric ligands to study antibody-hapten, enzyme-inhibitor (and drug), DNA-drug and cell-dye interactions [34]. The great majority of studies have involved protein-chromophoric ligand interactions. Recently, the labelling technique has been used extensively to monitor the chemical details of enzyme-substrate complexes. One approach is to use substrates which are themselves chromophoric [33,35], in another approach the chromophore is formed at the point of the transient covalent linkage between the substrate and the enzyme. The latter method relies on a reaction involving the hydrolysis of thionoesters (which may be based on a single amino acid or a multipeptide complex) by cysteine proteases such as papain. The hydrolysis of the thionoester substrate

N

U

-

O N

cch.

0

II

Ph - C - N H

S -CH2-

II

C -S-C-PAP

Fig. 14. Monitoring the group (and its neighbors) undergoing transformation in an enzyme’s active site. The resonance Raman spectrum shown is of the enzyme substrate transient PhC( = O)NHCH,C( = S)SPapain, 324 nm excitation; data acquisition time is 10 sec. From [82].

50 TABLE 3 Information

Method

Number and types of substrate conformers in active site

Identified using a library of structure-Raman spectra correlations

Search for geometric strain in bound substrate

By comparing RR spectra of enzyme-substrate intermediate with those for model compounds in standard state

Molecular information on individual kinetic rate constants

By combining RR and kinetic data

Conservation of catalytic mechanism during evolution

By comparing RR data for plant and mammalian cysteine proteases

Effects of enzyme-contacts far from points of catalyses

By investigating the effects of substrates of increasing sue on the RR spectrum of the enzyme-substrate linkage

Different species on the reaction pathway

Rapid mixing-rapid flow system using RR detection

Enzyme-substrate dynamics

Study intermediates under cryoenzymological conditions

RC(=O)NHCHR’C(=S)OCH, occurs via the formation of a transient dithioester RC(=O)NHCHR’C(=S)-S-papain, where the thiol sulfur is the S of cysteine 25 in papain’s active site. The dithioester is chromophoric absorbing near 315 nm and gives rise to a label just at the time and place of catalytic transformation. A natural intermediate in the hydrolysis of an ester or amide linkage is a thiol ester -C(=O)S, so the RR label involves substituting a C=S for a C=O group. The 324 nm excited RR spectrum in Fig. 14 shows how different spectral features monitor conformation in different parts of the enzyme-substrate complex near the scissile bond. The type of information gained by studying such complexes is summarised in Table 3 [ 36- 381. 7.4. Nucleic acids 7.4.1. The purine and pyrimidine bases The normal Raman spectrum of a polynucleotide contains approximately 30 detectable bands. Nearly all of these are due to purine or pyrimidine ring modes and the prominence of the base ring modes has led to detailed analysis of their vibrational spectra. The ring modes can be used to gain a semi-quantitative estimate of base content in a heteropolynucleotide, to monitor C-H C-D exchange on the rings [39] and to follow protonation of the bases. Protonation can even be detected for a small percentage of the total bases in RNA encapsulated in a virus [40]. In addition to the base modes, two important features originating from the phosphate group have been identified near 800 and 1100 cm-’ and are dealt with in the following section. The ribose or deoxyribose groups are poor Raman scatters and do not contribute any

51

cm"

Fig. 15. Raman spectra of H,O and D,O solutions of poly(rA). poly(rU) at 32 and 85°C. From [43].

intense or moderately intense features to the spectrum. However, in the case of guanosine it has been demonstrated recently that the guanine ring-breathing frequency near 650 cm-' is sensitive to conformational events in the sugar and the glycosidic linkage [41]. In both DNA and RNA certain purine and pyrimidine ring modes are sensitive to base-pairing and base-stachng interactions. Some of these features, and the general appearance of polynucleotide spectra, are illustrated by the spectrum shown in Fig. 15 of poly(rA)poly(rU), a double helix in which the adenine residues of poly(rA) hydrogen bond to the uracil residues of poly(rU). The system was initially investigated by Small and Peticolas [42] and independently by Lafleur et al. [43]. As Fig. 15 shows, marked spectral changes occur upon raising the temperature from 32 to 85 " C. The spectral changes result from the thermal disruption of the helix with, at high temperature, the almost total loss of the base-pairing and base-stacking interactions. Key spectral changes in Fig. 15 are intensity variations of certain ring modes and the radical changes in the carbonyl profiles between 1650 and 1700 cm-'. In the latter region in D,O at 85 C the carbonyl groups of U give strong lines at 1661 and 1696 cm-' while at 32°C a single line occurs at 1688 cm-'. Several lines, assigned to A and U ring vibrations, show appreciable intensity loss upon helix formation at32°C. This is called Raman hypochromism [44,45]. The phenomenon has a direct correspondence with the hypochromism observed in the UV adsorption spectrum and is believed to arise from the electronic interactions between vertically stacked bases. A few lines, however, show modest intensity gain in the 32" C spectrum (e.g. the mode near 1570 cm-' increases in intensity with increasing structural order) and this effect is termed Raman hyperchromism. O

52

Similar effects are seen in the spectrum of the poly(rC)poly(rG) complex. Again there are several hypochromicities and a notable hyperchromic band at 670 cm-' due to a G ring mode.In poly(rC)poly(rG), however, the intensities of lines in the carbonyl region are weaker, by a factor of 6-8, compared to the carbonyl modes in the A.U. helix. It is tempting to make the assumption that in a RNA containing a heterogeneous sequence of bases the carbonyl region may be used to quantitate-base pairing and the purine and pyrimidine ring modes may be used to form a semi-quantitative picture of base stacking. However, difficulties arise from the fact that the profile in the carbonyl region and the degree of hypochromism are both independent upon base sequence. Moreover, some hyperchromic or hypochromic lines may reflect base pairing as well as stacking interactions [46]. 7.4.2. Conformation of the (deoxy)ribose-phosphate backbone The phosphate groups:

in the (deoxy)ribose-phosphate backbone of RNA (and DNA) give rise to two characteristic Raman bands near 800 and 1100 cm-' whch are very useful probes of backbone conformation. The 800 cm-' feature is assigned to the -0-P-Osymmetrical stretching vibration [47] while the 1100 cm-' band originates from the PO; symmetric stretch motion. Conformational utility stems from the fact that the frequency and intensity of the PO, feature is insensitive to backbone geometry, and thus provides an internal standard, while the 800 cm-' is highly sensitive to conformation of the -C-0-P-0-Cgroup and possibly the ribose ring [48,49]. The Raman spectrum of an ordered, single-stranded or double-helical RNA shows an intense sharp band at 810-815 cm-' which shifts to near 795 cm-' upon disordering. This transition is clearly seen in the spectrum of poly(rA)poly(rU) in Fig. 15 while the same spectrum demonstrates the insensitivity of the 1100 cm-' feature to disruption of secondary structure. The intensity of the PO; Raman mode remains constant at constant ionic strength. For RNA, at low ionic strength, the ratios of intensities of the bands at 851 and 1100 cm-' can be used to monitor the amount of secondary structure. Z(815)/( I(1100) = 1.66 in completely ordered ribopolymers (double-helical or single-stranded) and 0.0 in completely disordered ribopolymers [50]. The phosphate modes for DNA resemble those for RNA; the DNA 1100 cm-' band is insensitive to changes in secondary structure and, as in the case of RNA, the features near 800 cm-' in the Raman spectra of DNA are sensitive to conformation. For DNA in solution or cast as a fiber the 800 cm-' band can be used to characterise the A, B [51] and Z [52]forms of DNA and to follow fluctuations in the sugar conformations [53].

53

7.4.3. Resonance Raman studies of nucleic acids The nucleic acid bases have a rich assortment of electronic transitions in the 200-300 nm range. These absorption transitions are being increasingly exploited to provide RR spectra. In turn the RR spectra, via their excitation profiles, can help identify some of the often controversial features in the electronic absorption spectra [54,55]. The most detailed analysis of the RR spectra of nucleic acids appears in the review by Nishimura et al. [56], where these authors discuss a normal coordinate treatment of nucleic acids together with RR theory and experimental results. Several approaches have been used to generate UV laser light. Early studies used 257.3 nm obtained by frequency doubling the 514.5 nm cw argon laser line [56], sources in the 267-305 nm were then accessed using a frequency doubled pulsed dye laser [54,55]. More recently harmonics of the YAG laser output and their H, or D, Raman shifted frequencies have been used in the 200-266 nm range [57,58].

N

dAMP

W

0

2Wnm x 0.33

200nm

1

600

1

1

800

1

1

1

1

1000 1200

1

1

1400

1

1

1600

1

1

I800

A cm-' Fig. 16. Resonance Raman spectra of aqueous, 5 mM, deoxyadenine 5'-monophosphate. The different excitation wavelengths used for each spectrum are indicated. From [SS].

54

The excitation wavelength dependence of the RR spectrum of deoxyadenine 5’-monophosphate is illustrated in Fig. 16. Quite pronounced changes in the appearance of the RR spectrum occur and these can be understood on the basis of the five calculated T-T * electronic transitions of adenine in the 200-266 nm region [58]. The strong wavelength dependence seen for adenine is also found for the other bases and this behavior holds promise for selectively exciting the RR spectrum of a given class of base in a complex polynucleotide structure. The enhancement variations also assist the spectroscopist in resolving overlapping components and in making vibrational assignments. The substitution of a bromine atom on a nucleic acid base (e.g. 8Br-adenosine) provides the base with significant absorption in the 300 nm region. The RR spectrum of the brominated species can then be specifically excited by 300 nm irradiation [59]. The brominated bases are in effect RR labels but there are rare naturally occurring bases with significant absorption in the near UV, these are sulfur substituted analogs absorbing from 300-360 nm. For example Nishimura, Tsuboi and coworkers [60] obtained RR spectra of the single 4-thiouridine which occurs in a number of tRNAs. 7.5 Viruses

Raman spectroscopy has yielded detailed molecular information on several viruses and other DNA and RNA complexes (see [2] pp. 198-201). Here, one recent example, the RNA-based cowpea chlorotic mottle virus [61], will be discussed since it illustrates the wealth of detail which may be gained from the Raman spectrum. At the same time the study provides examples of the conformationally sensitive RNA and protein features mentioned in the foregoing sections. Cowpea chlorotic mottle virus is an icosahedral RNA virus made up of three nonidentical ribonucleoprotein particles. The Raman spectra of the intact virus and its individual component protein shell (capsid) and RNA parts are shown in Fig. 17. The base and backbone contributions are identified in the spectrum of the RNA while the amide, CH, and CH, contributions are indicated in the spectrum of the protein. Most of the other features in the capsid spectrum can be assigned to aromatic side chain modes. By analysing the small spectral differences observed as a function of pH and by comparing the spectrum of the intact virus by difference spectroscopy with spectra of capsids, subunit dimers and protein-free RNA, Thomas and his coworkers [61] have demonstrated that a good deal of detailed information may be obtained: a. For the RNA in the virus, the carbonyl stretching region and the purine and pyrimidine ring modes indicate that the bases are paired and stacked. The relative intensity of the -0-P-0- and PO; modes shows that the backbone of encapsulated RNA is in the conventional C3’-endo geometry. b. The proteins’ amide I and 111 profiles show that the predominant secondary structure in the virion is /3-sheet. c. Two features are seen at 2551 and 2571 cm-’ in the -SH stretching region, with the former band being broad and assigned to -SH groups in a H-bonding

55

Virus

Capsid

I

C

"I

R NA

Fig. 17. Comparison of the Raman spectrum of an intact virus (top) and its protein (capsid) and RNA components. From [61].

d.

e.

f.

g. h.

environment. The 2571 cm-' band is ascribed to the remaining cysteine chains in a hydrophobic environment. The tyrosine doublet indicates that the tryosine side chains exist on average with the p-hydroxy groups in contact with hydrogen-bonding donor and acceptor groups. Differential tryptophan and phenylalanine ring intensities suggest that these residues are in different environments in the virion and capsid states and, thus, that these aromatics are involved in interactions with the RNA. Changes in bands from C-C, -CH,- and -CH, groups in the 1130 and 1340 cm-' regions show that configurations in the aliphatic side chains are altered upon release of the RNA from the virion. There are changes in both protein and RNA backbone conformations upon dissociating the RNA from the capsid. Swelling of the virion by increasing the pH from 5.0 to 7.5 has no effect on aspartate and glutamate -COOH features and thus does not titrate these groups. However, small numbers of Ade+ and Cyt residues become deprotonated. The protonation of bases in the virion at pH 5.0 is limited to adenines and cytosines. +

i.

56

7.6. Lipids and membranes

Both Raman and infrared spectroscopy provide a wealth of molecular detail on the conformational and dynamical properties of model and real biological membranes. The infrared and Raman methods are complementary and often mutually supportive, but in keeping with the theme of this chapter just the Raman technique will be discussed here. Raman spectroscopy has the capability of monitoring both inter- and intramolecular interactions in membrane assemblies and for identifying the part of the lipid molecule undergoing a particular perturbation. Fig. 18 shows many of the conformationally sensitive features. The headgroup region can be probed via the symmetric C-N and PO, modes near 717 and 1243 cm-', respectively, and the carbonyl environments by their modes near 1720 and 1740 cm-'. The Raman skeletal C-C stretching vibration region between 1050 and 1150 cm-' enables one to monitor the truns/guuche conformational ratio, within the acyl chains, while intermolecular chain packing properties can be followed by intensity changes in the C-H stretching region from 2800-3100 cm- The methylene deformation region near 1450 cm-' can also be used to follow changes in intermolecular order. The C-C and C-H stretching regions are the most often used as probes of membrane structure and they will be discussed in more detail.

'.

Liquid crystalline;

1

I

1

520

1

1

840

h

40°C

1

1

1160

cm-'

1

1

I480

I

I

1800

Fig. 18. Raman spectra of dimyristoyl phosphatidylcholine in the gel and liquid crystalline states. From ~31.

7.6.1. The C-C stretching region between 1050 and 1150 cm-' This region contains vibrations in which alternate carbon atoms move in opposite directions along the chain length. There are at least three bands which exhibit marked changes upon disordering of the hydrocarbon chain. These can be seen in Fig. 18 for dimyristoyl phosphatidylcholine going through the gel-to-liquid crystalline transition. The intensities of the two strong lines near 1128 and 1064 cm-', assigned to skeletal optical modes of all trans conformers, decrease abruptly as the melting temperature ( T m )is approached. Plots of intensity of the 1128 cm-' feature (normalised with respect to a temperature invariant band) vs. temperature for many systems reveal that the type of transition seen in Fig. 18 is highly cooperative. The temperature dependence of the spectra, in the 1000-1200 cm-' region, of a variety of phospholipids have been studied by a number of groups and there have been attempts to extract quantitative information on lipid behaviour using the feature near 1130 cm-' [62]. However, as Levin has pointed out [63] the difficulties in extracting quantitative information are formidable due to such factors as coupling of the 1130 cm-' mode to the terminal methyl rocking mode at 892 cm-', underlying features in the 1100 cm-' region and problems in normalising the various parameters involved. Even with these difficulties temperature profiles constructed from the intensity ratios of the 1085-1090 and 1130 cm-' features, provide a convenient method for following gel-to-liquid crystalline phase transitions. An increase in the 1090,4130 cm-' ratio denotes an increase in chain disorder through the introduction of gauche bonds. 7.6.2. The C-H stretching region between 2800 and 3000 cm-' This spectral range contains C-H stretching modes from the phospholipids' methylene and methyl groups. The relative abundance of the methylene group results in -CH,- modes dominating the C-H stretching region. This is evidenced by the Raman spectrum, between 2800 and 300 cm- ',of diarachidoyl phosphatidylcholine shown in Fig. 19, where the intense peaks near 2847 and 2883 cm-' are assigned to the methylene symmetric and asymmetric stretching modes, respectively. The weaker band near 2936 cm-' is assigned, in part, to the chain terminal CH, symmetric stretch. Although these assignments establish the origin of the 'sharp' features seen in the C-H stretching region there has been considerable effort recently to elucidate the origins of the broad band or bands underlying the resolved peaks. An understanding of these broad bands is essential to the correlation of spectral with conformational and environmental change 1631. The sensitivity of the C-H region to conformational change is demonstrated in Fig. 19 which compares the Raman spectra of dispersions in the gel and liquid crystal states. As the temperature is raised and the gel state bilayer disorders and reaches the liquid crystalline state there is a marked change in the feature near 2880 cm-I. It decreases in intensity, broadens and shfts by + 10 to 12 cm-I. Although a number of spectral changes may be used, many authors have used the peak height to follow gel-to-liquid crystalline phase intensity ratio Z2ss5/I,,5, (or 12885/12935) transitions. In general, increases in peak intensity of the 2935 (or 2850) and 2880 cm- bands reflect intermolecular chain disorder and order, respectively.

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;

Liquid crystalline

1

I

I

I

I

I

I

I

2800 2842 2885 2928 2971 3014 3057 3100 cm-’

Fig. 19. Comparison of the Raman spectra, in the C-H stretching region, of diarachidoyl phosphatidylcholine in the gel and liquid crystalline states. From [63].

7.6.3. Deuterated lipids as selective probes In multicomponent systems difficulties arise from the overlapping of C-H features due to chemically different lipids, or in lipid-protein arrays, from protein C-H stretching bands. In these systems it becomes impossible to monitor a single lipid component. Mendelsohn et al. [64] showed how this problem can be overcome by the use of deuterated components. They inserted a completely deuterated fatty acid into a model membrane system and followed the C-D stretching vibrations in the spectral window 2000-2220 cm-’ which is uncluttered by modes from other Components. As the membrane passed through a gel-liquid crystal transition the line-width of the C-D stretching vibrations of the bound fatty acid was found to be a sensitive probe of membrane polymethylene chain order. Selective deuteration provides information on band assignment [65,66] and a probe of conformation within a single phospholipid. B a n d et al. [67] studied the temperature dependence of the Raman spectra of 1,2-dimyristoyl-sn-glycero-3-phosphocholines specifically deuterated in the 2 chain at positions 3, 4, 6, 10, 12 and 14. These authors showed that the frequencies of the CD, stretching modes depend on the position of the CD, label, being maximum at position 3 and decreasing until they become constant beyond position 6. The degree of the observed increase in the width of the C-D bands at T, is also position dependent. An interesting example of selective deuteration is the work of Gaber et al. [68] which involves deuterating the 1 chain or the 2 chain of dipalmitoyl phosphatidylcholine. Differences in the spectral characteristic of the compounds containing the perdeuterated 1 chain or the

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perdeuterated 2 chain at a certain temperature were noted. These differences were attributed to nonequivalent conformations of the fatty acid chains at positions 1 and 2; below the pretransition chain 2 appears to depart slightly more from the all-trans structure than chain 1. 7.6.4. Lipid protein interactions and natural membranes The interaction of lipids with small entities such as cholesterol [69] and metal ions I701 has been studied extensively by Raman spectroscopy. The effect of the perturbants has been followed via the lipids' C-C and C-H stretching regions. Recently, more attention has been focussed on lipid protein interaction, again the C-C and C-H regions are crucial to following changes in lipid properties, but now protein features, such as the amide I and I11 modes, may be recognized in the Raman spectrum [71] and analysed according to the precepts set out in the discussion of protein spectra above. In one study Taraschi and Mendelsohn [72] formed lipid-protein complexes from dipalmitoyl phosphatidylcholine (and its perdeuterated analog) and glycophorin, a protein isolated from erythrocyte membranes. The temperature dependence of the C-D stretching region of the protein-lipid complex showed that at a lipid protein mole ratio of 125 : 1, a broad melting event occurs whose midpoint is about 15°C lower than that for the pure lipid. Using their earlier postulates concerning C-D linewidth and gauche population [73] the authors concluded that the same number of gauche isomers form in the phospholipid hydrobcarbon chains during the melting process as in the phase transition of the pure lipid. There is special interest in the nature of interactions at the lipid-protein interface and Taraschi and Mendelsohn point out that the conformation of the lipid at the interface is markedly changed by the interaction. Moreover, the Raman results enable them to infer for the 125: 1 sample that the perturbation extends well beyond one shell of lipid molecules and must be at least four to five layers deep. Lavialle et al. 1741 have also obtained information on the lipid-protein boundary. These authors studied the interaction of melittin, a polypeptide consisting of 26 amino acid residues, with dimyristoyl phosphatidylcholine. The results illustrated in Fig. 20 show that for a lipid-melittin molar ratio of 14: 1 two order-disorder transitions are observed, one above (at 29°C) an$ one below (at 17°C) the transition for the pure lipid (at 22.5 " C). The low temperature transition is associated with a depression of the main lipid phase transition while the 29 " C transition is associated with the melting behavior of approximately seven immobilized boundary lipids which surround the hydrophobic portion of the melittin. The application of Raman spectroscopy to naturally occurring membranes is hindered by problems such as background luminescence, or the sheer complexity of the system. However, several groups have been able to obtain Raman spectra from natural membranes. The most extensively studied system is the membrane from human erythrocytes, commonly known as red blood cells [75-771. Lippert et al. [75] examined the membranes from human red blood cells, which had been repeatedly washed to remove traces of fluorescent material. The Raman signatures of both protein and lipid components were observed and from the amide I' (in D,O) and

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