Indian Journal of Biochemistry & Biophysics Vol. 40, June 2003, pp. 147-159

Minireview

Fluorescence spectroscopy in molecular description of biological processes G Krishnamoorthy

Dept. of Chemical Sciences, Tata Institute of Fundamental Research, Homi Bhabha Road, Mumbai 400 005, India

Received 20 January 2003; revised & accepted 2 May 2003

The power of fluorescence spectroscopy in enhancing our ability to obtain molecular description of complex biological processes is discussed. A brief account of sensitivity, selectivity and observable parameters of the various fluorescence methods and some typical applications which are mostly based on work in our laboratory are provided. The applications cover a wide range of areas such as protein dynamics, protein folding dynamics, protein-DNA interactions, membrane dy-namics and intracellular dynamics. Interesting information revealed by fluorescence-based methods in each of these applica-tions is highlighted.

Key words: 2-Aminopurine, cell membrane raft, DNA dynamics, Time-resolved fluorescence, fluorescence spectroscopy, fluorescence probes, Maximum entropy method, membrane heterogeneity, protein dynamics, protein folding.

Introduction Fluorescence spectroscopy is one of the most

widely used forms of molecular spectroscopy in bio-chemical research and is surpassed only by electronic absorption spectroscopy. Its early use was as an ana-lytical tool for determining the concentration of vari-ous molecular species. When the molecular species (analyte) is fluorescent, its concentration can be de-termined directly from the intensity of fluorescence. This is of immense use especially in complex physio-logical fluids having a multitude of components. Thus the need to separate the component of interest could be avoided. Based on this principle, assay systems for a large number of enzyme reactions and various bio-molecules had been developed and effectively used. Besides, a variety of new and quite exotic (in some cases) applications of fluorescence spectroscopy in biological research problems are encountered in re-cent times. The main thrust behind these new applica-tions is two fold: (i) technical developments in laser systems and optical detection methods and (ii) devel-opment and maturation of novel optical methods and data analysis. These developments are rapidly and

significantly enhancing our capability of gaining deeper understanding of the complex biological sys-tems. In this article, I shall give a few glimpses of novel information generated by the application of state-of-the-art techniques of fluorescence spectros-copy to biological research. The applications which are mostly based on work in our laboratory cover a wide range of areas such as protein dynamics, protein folding dynamics, protein-DNA interactions, mem-brane dynamics and intracellular dynamics. For a more exhaustive coverage, the reader is referred to the recent monographs and reviews1-4. What are the observable parameters in fluores-cence?

Although fluorescence intensity has been the most widely used observable parameter in many early use of fluorescence spectroscopy, there are several other observable parameters which also could be put to ef-fective use. These and the typical information they provide are listed in Table 1. They are classified into two broad categories, namely (i) steady-state and (ii) time-resolved parameters. Steady-state measure-ment of fluorescence properties is marked by their relative simplicity and robustness of measurement. Fluorescence intensity could be used either as inte-grated total intensity under the fluorescence emission spectrum or as shape of emission spectrum. Both the total intensity, which is proportional to the fluores-cence quantum yield and the emission spectrum, pro-vide information on the environment of fluorescent

—————— E-mail: gk@tifr.res.in Abbreviations used: 2-AP, 2-aminopurine; DCCD, N, N’-dicyclo-hexylcarbodiimide; DODCI, 3, 3’-diethyloxadicarbocyanine io-dide; FCS, fluorescence correlation spectroscopy;Trp, tryptophan; FRET, fluorescence resonance energy transfer; GFP, green fluo-rescent protein; GPI, glyscosyl phosphatidylinositol; MEM, maximum entropy method; TCSPC, time-correlated single photon counting; TNB, trinitrobenzoate.

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Table 1—Observable parameters in fluorescence

Parameter Information offered Comments Steady-state parameters Total intensity Molecular concen-

tration, molecular environment

Interference in complex media

Emission spectra Molecular envi-ronment (polarity etc.)

-do-

Fluorescence ani-sotropy (or polari-zation)

Molecular flexibil-ity, microviscosity of medium

Robust measure-ment but interfer-ence likely in com-plex media

Time-resolved parameters

Fluorescence fluc-tuation

Translational dy-namics, chemical kinetics

Requires expensive instruments

Fluorescence life-time

Molecular envi-ronment, structural information, popu-lation distribution, translational diffu-sion

Very sensitive, but requires expensive instruments

Fluorescence ani-sotropy decay time (or rotational corre-lation time)

Molecular motions, microviscosity, structural informa-tion

-do-

Time-resolved emission spectra

Solvation dynam-ics, excited state reactions, popula-tion heterogeneity

-do-

molecule. High sensitivity of intensity and emission spectrum on the environment has been efficiently used to derive useful information. Another useful pa-rameter is fluorescence anisotropy, r (or polarization, p which is similar to r. r = (I|| - I┴)/((I|| + 2I┴) and p = (I|| - I┴)/(I|| + I┴), where I|| and I┴ are the intensities of emitted light having polarizations parallel and per-pendicular to that of excitation light). When excited by a stream of linearly polarized photons the resultant fluorescence emission could either be polarized (r >0) or depolarized (r = 0) depending mainly on rotational motion of fluorophore in its environment. This phe-nomenon could be very effectively used in many seemingly intractable situations. Several of the exam-ples given in subsequent sections will amply clarify this claim.

All the three observable parameters, namely inten-sity, emission spectrum and fluorescence anisotropy could also be monitored with time as extra dimension. Such time-resolved parameters (Table 1) offer us

added advantages enabling deeper understanding of the system being probed. Time as the extra dimension is associated with a quantum leap in information re-trieval. Time resolution of fluorescence can be thought of in two ways: (i) change in fluorescence parameters (mainly the intensity) originating from time-dependent variation in the concentration of the fluorescent species. Change in concentration could be either due to some chemical or physical (such as transport across membrane) process or to fluctuations in concentration of the fluorescent species, brought about by brownian motion. While the former type of time-dependence forms the fluorescence based assay for many chemical and biochemical reactions, the lat-ter (fluctuation) process forms the core of recent de-velopments such as ‘Fluorescence Correlation Spec-troscopy’ (FCS)5-7. Examples of applications of FCS in biological problems are given in a later section; (ii) time dependence of fluorescence arising out of spec-troscopic processes such as transition from excited electronic level to the ground level of fluorescent molecule. Direct observation of this time-dependence requires capability to excite the molecules with short pulses from an ultrafast laser and to detect the emitted fluorescence with time resolutions in the range of femtoseconds to nanoseconds. All the three parame-ters, that is intensity, spectra and polarization (or ani-sotropy) can be monitored with such time-resolution. Time-resolved parameters significantly enhance the quality of information which could be extracted from complex systems. Fluorescence lifetime (Table 1) gives information on the environment around the fluo-rescent molecule and fluorescence anisotropy decay time offers a window to observe the motional dynam-ics.

Where to place the fluorescent probe? This question is not obviously relevant in situations

where the fluorescence arises naturally from the par-ticular molecule of interest. However it becomes ex-tremely relevant when either an intrinsic fluorophore such as tryptophan is engineered in a protein or an extrinsic fluorophore is covalently attached to a spe-cific location in a macromolecular system. In fact, the success of using fluorescence-based approaches to extract information on the molecular mechanism de-pends largely on the selection of fluorophore location. Intuition could guide us in selecting the location ap-propriately in some situations. For example, placing the fluorescence label at a location close to the site of action such as enzyme active site8, ligand binding site,

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protein-protein interaction site could result in success-ful information retrieval. Furthermore, selection of fluorophore is also important. The guiding principles involved in selection of site and the type of fluoro-phore depend on the problem being addressed and is best illustrated in the typical examples which are given below.

Fig. 1 shows some of the fluorophores which are useful in biological research. The probes are classified by their spectral properties, molar extinction coeffi-cient, fluorescence quantum yield, size, environment sensitivity, pH dependence, metal ion dependence, photostability, etc. Apart from organic molecules, the list contains some exotic species such as quantum dots. These semiconductor nanoparticles such as ZnS, CdSe are new entrants coming from an unrelated area of science and they have remarkable luminescence properties9-12. Their emission spectra are quite narrow. The emission wavelength depends on the size of quantum dot (1-10 nm) and spans the complete visible region9,11. However, their excitation spectra are largely similar. This property is expected to be of tremendous use in cell biology, wherein probes of different emission colour could be localized at differ-ent regions in the same cell and excited by a single laser source. Other advantage of quantum dots include higher level of photostability compared to organic fluorophores12. However, the lack of newer develop-ments in the technique of coupling them to biological macromolecules has restricted the efficient use of quantum dots. Most of the organic fluorophores cur-rently in use could be found in the catalog of Molecu-lar Probes Inc (www.probes.com). However, some of the probes could also be obtained at lower prices from other manufacturers. Commercial availability of quantum dots can be explored at www.evidentech.com. Instruments required for fluorescence experiments

The level of complexity of intsrumentation re-quired for fluorescence-based experiments depends on the type of information being sought. Measurement of steady-state properties such as emission spectra and fluorescence anisotropy can be carried out by using commercially available instruments. Spectrofluorime-ters based on photon counting have higher sensitivity, compared to photocurrent measuring fluorimeters. Most of them are designed in L-format, where the fluorescence is detected at a single direction perpen-dicular to excitation beam. However, precision meas-

urement of fluorescence anisotropy (or polarization) in time-evolving samples requires T-format configuration, where fluorescence is de-tected simultaneously in two directions perpendicular to the excitation beam. Thus, the fluorescence inten-sity at the two polarizations (parallel and perpendicu-lar to that of excitation) required for estimating fluo-rescence anisotropy, could be estimated simultane-ously13.

Measurement of time-resolved parameters such as fluorescence lifetime or fluorescence anisotropy de-cay time (i.e. rotational correlation time) requires sig-nificantly more complex instruments. Basically they are measured two ways, viz. time-domain and fre-quency-domain. Although both have similar capabil-ity, the results from time-domain method are easier to visualize and comprehend2-4. However, time-domain methods requires expensive ultrafast lasers and detec-tors based on techniques such as Time-Correlated-Single-Photon-Counting, (TCSPC)2,3,14, fluorescence up-conversion15 and stroboscopic methods16. In con-trast, frequency-domain instruments are less expen-sive and hence are more popular among biologists. They also have wider wavelength range available for excitation. In time-domain experiments, the fluoro-phore is excited by a narrow (fs to ns) pulse of light and the decay of fluorescence intensity is monitored directly as a function of time by a variety of tech-niques. Thus, observable parameters such as fluores-cence lifetime and anisotropy decay time could be visualized directly in the time-domain. In frequency-domain measurements the fluorophore is excited con-tinuously by sinusoidally modulated light. The de-tected fluorescence signal also will have the sinusoi-dal modulation. However, due to finite value of ex-cited state lifetime, the sinusoidal nature of fluores-cence signal will differ from that of excitation in two ways: (i) the phase shift, ϕ and (ii) modulation ratio, M. Both ϕ and the decrease in M could then be used to retrieve the time-resolved parameters2-4.

Time-domain measurements of fluorescence where temporal changes originate from fluctuations about a mean level require instrumentation and analysis, which fall in the realm of Fluorescence Correlation Spectroscopy (FCS). The reader is referred to some of the recent reviews in this field5-7. The need to observe very small sample volumes having an average number of molecules less than ~10 requires the use of fluores-cence microscope coupled with laser source and hardware correlators of the intensity monitored by

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Fig. 1—Typical and widely used fluorescence probes [1, Tryptophan (natural amino acid); 2, Aladan, an unnatural amino acid used site-specifically to study protein electrostatics and dynamics (ref. 87); 3, IANBD, an environment-sensitive probe; 4, Kiton Red, a cytoplas-mic probe (ref. 80); 5, SNARF-1, dual wavelength pH probe (ref. 47); 6, Pyranine, membrane impermeant pH probe (ref. 88); 7, Nile Red, polarity-sensitive membrane probe (ref. 65, 70); 8, Badan, a membrane probe; 9, DPH, a membrane probe; 10, fluorescein; 11, rho-damine; 12, erythrosine (ref. 8); 13, BODIPY, a high quantum yield environment insensitive probe (ref. 75); 14, Pyrene methyl iodoace-tate, a long lifetime probe; 15, Fura, a Ca2+ sensor and 16, YOYO-1, a DNA dynamics probe (ref. 89)]

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high quantum efficiency detectors such as avalanche photo diode (APD). What can we learn from fluorescence experi-ments? (Some examples)

In this section, we shall see some typical applica-tions of fluorescence spectroscopy in biological sci-ences. Although these might cover both steady-state and time-resolved experiments, emphasis will be higher on the latter, due to higher level of information obtainable from time-resolved parameters. The exam-ples have been categorized into various types of bio-molecular systems and interactions among them. (i)Protein dynamics

The notion that dynamics along with their struc-tures form the basis of activity of biomolecular sys-tems is undergoing experimental tests in recent years17,18. Proteins would be the most suitable area to test this notion. Of several experimental and simula-tion methods available for studying macromolecular dynamics fluorescence-based methods have the fol-lowing advantages: (i) dynamics of a loca-tion/segment of a massive macromolecular system can be specifically monitored without the interference from the rest of the system; (ii) the timescale of dy-namics covers a wide range of femtosecond to sec-onds; and (iii) experiments can be performed with very sparse levels of molecules.

The naturally occurring fluorescent amino acid tryptophan is an excellent probe of protein dynamics in spite of its complex fluorescence decay kinetics19,20. Engineering tryptophan in various locations enables us to learn about local and global dynamics of pro-teins. Local dynamics informs us about the flexibility of the protein around the tryptophan residue, while global dynamics the overall hydrodynamic radius and the multimer/aggregation behaviour of the protein. Information on local dynamics could then be corre-lated with the function. Rotational correlation time is the most suitable method for monitoring local and global dynamics. Fluorescence lifetime and the effect of fluorescence quenchers such as iodide, acrylamide also provides information on the accessibility of tryp-tophan and hence the flexibility of the environ-ment13,21,22. Since iodide and acrylamide are polar and non-polar respectively, differential information on the location of fluorophores (a structural information) could be obtained.

Activity of proteins depends on specific interaction with its partner such as ligand, other proteins, DNA

sites, etc. Hence, dynamics of side chains at the sur-face of proteins is of great concern and is possibly controlled by the layer of water bound to the protein surface. This bound water (‘biological water’23) has different properties compared to bulk water. This problem has been approached primarily in two ways: (i) relaxation of bound water has been monitored with fs time resolution24; Monitoring time-resolved fluo-rescence emission spectrum of surface-located trypto-phan residue using the technique of fluorescence up-conversion15 showed that the relaxation of ‘biological water’ is significantly slower than that of bulk water similar to observations in heterogeneous media25; and (ii) by monitoring the local dynamics of surface-exposed tryptophan side chains in several proteins through their rotational dynamics in ps timescales26. In latter case, monitoring the effect of increasing bulk solvent viscosity on the surface dynamics in folded and unfolded proteins26 showed that the flexibility (or dynamics) of surface groups is controlled by the bound water and not by bulk water in folded proteins. Furthermore, no such difference was observed in un-folded proteins indicating the absence of bound water layer in unfolded proteins (Fig. 2). Thus, our studies suggest that interaction involving proteins could get

Fig. 2—A schematic diagram showing that the dynamics of side chain is controlled by bound water layer in native protein (top) and by bulk water in denatured protein (bottom) [Spheres, dark ellipsoid and rectangles represent bulk water, ordered water and glycerol respectively. Surface-labeled fluorescence probe is repre-sented by an ellipsoid linked by a cone. For more details see ref. 26]

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modulated by the properties of the layer of ‘biological water’. In all these examples, the intrinsic fluorophore tryptophan was used as probe, since it was possible to have a single tryptophan in these small proteins. Thus, it is better to use intrinsic probes, wherever possible in order to ensure that the observed dynamics origi-nates from the system rather than induced by the probe.

A direct correlation between local dynamics and function was observed in the electron transfer-driven proton pump, NADH-CoQ oxidoreductase of mito-chondria8. In this case, rotational dynamics of the ex-trinsic probe, DODCI bound to quinone- binding site was used to infer the segmental dynamics. The qui-none-binding site has been implicated as the proton-pumping site and hence its dynamics is expected to be correlated with the efficiency of the pump. The seg-mental dynamics as reported by the fluorescence ani-sotropy decay kinetics of DODCI was damped when the proton pump was inhibited by the well character-ized inhibitor DCCD27, thus supporting the identifica-tion of quinone-binding site as the site of pumping. Thus, dynamics information could even be used for identifying functional domain in a complex system. (ii) Protein folding dynamics

Mechanism of protein folding is one of the areas which has benefited significantly from the power of fluorescence dynamics studies. One of the main rea-sons is that the process of protein folding from an ex-tended random polypeptide chain goes through sev-eral intermediate forms, which are likely to have par-tial structures. Such an ensemble of unfolded and par-tially folded forms is not amenable to many physical techniques, such as X-ray crystallography and NMR. Fluorescence-based methods are thus ideally suited for studying such distributed ensemble structures.

Barstar, a small protein (89 amino acids) and a nau-ral inhibitor of the ribonuclease barnase28 having no disulfide bonds has been used extensively in protein folding studies as a model protein13,21,29-32,. Of the three tryptophan residues one of them is located at the core of the protein (Fig. 3). This side chain (Trp53) is found to be an ideal marker for studying dynamics of various structural forms encountered in the folding process. Hence, a mutant protein having only Trp53 was made. It helps us to monitor the core dynamics specifically. Firstly, it was observed that Trp53 was associated with a single lifetime in the native state of the protein13. This unusual observation was inter-

Fig. 3—Rasmol representation of the structure of barstar used as a model protein in folding dynamics studies (refs. 13, 21, 29-32) [The core tryptophan (W53) which is used as the probe and the sites of fluorescence acceptors (refs 31, 32, 37, 38) are marked]

preted as due to rigid core freezing the conformational freedom of Trp53. It was also supported by direct ob-servation of its rotational dynamics with the help of fluorescence anisotropy decay, which showed the ab-sence of any local motion, but the presence of only global motion due to tumbling of the protein13. In un-folded state, conformational freedom of Trp53 was relaxed and was seen from multiple fluorescence life-times and the presence of unrestricted local dynam-ics13,31. These effects are anticipated. However, the most interesting observation was made during transi-tion from native to unfolded form, aided by denatur-ants. Presence of partially folded structure(s) was in-ferred from the fluorescence anisotropy decay kinet-ics13,31. The kinetics were distinctly different from that of the native (N) or denatured (U) forms, indicating the inadequacy of two states (N = U) model, which has been assumed to represent the folding process of single domain proteins such as barstar. Thus, the presence of intermediate structural form (I) was re-vealed from its characteristic motional dynamics (Fig. 4).

In order to get structural information on intermedi-ate forms, we used fluorescence resonance energy transfer (FRET) between Trp53 and TNB group cova-lently attached to the single Cys (Cys82) side chain in a mutant form of barstar32. Picosecond time-resolved decay of fluorescence intensity of Trp53 depends upon the rate of energy transfer to Cys82-TNB and this, in turn is a function of the distance between

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Fig. 4—Rotational dynamics of W53 (labeled F) of barstar reveals the presence of partially folded intermediate (I) states (ref. 13) [Value 12 ns for the rotational correlation time τr is taken as the evidence for I]

Trp53 and Cys82. Kinetics of intensity decay, ana-lyzed by the model-free Maximum Entropy Method (MEM)33,34 provides information on distribution of distance which arises due to ensemble distribution of protein structural forms32. Such vital information on population distribution is a unique outcome of the combined techniques of time-resolved FRET and MEM. This method when applied to the folding tran-sition showed that transition between N and U occurs in a continuous manner, rather than either as a two-state process or as a process involving a few discrete forms of I (Fig. 5). The N state expands progressively to reach the dimension of a molten globule-like state before transformation to the U form. It should be mentioned that this result is not inconsistent with the information derived from rotational dynamics (Fig. 4). While rotational dynamics indicated the presence of intermediate structure (Fig. 4), it did not preclude the existence of ensemble of intermediate structures. FRET studies (Fig. 5) showed clearly that there is a continuum of expanded structures in be-tween N and the transition state. The ensemble of such structural form was revealed collectively from their rotational dynamics (Fig. 4) whereas, FRET brought out the continuous nature of this ensemble. Thus, these time-resolved experiments (Figs 4 and 5) have shown that the transition between folded (N) and unfolded (U) states of even a single domain small pro-tein, such as barstar, is not a two-state process, but is associated with a continuum of intermediate states. In contrast, steady-state spectroscopic measurements have been unable to bring out these information since they all could be fitted to the minimal model of two-state process.

The above observations were made on equilibrium situations. Similar observations made during the ki-netic process of protein folding would provide infor-

Fig. 5—Continuum of states (N, N’, N” …..) during the unfolding of barstar as inferred from t-FRET-MEM analysis of folding tran-sition (ref. 32) [The continuum is separated from the unfolded state U by an energy barrier]

mation on the energy landscape of folding process. Steady-state fluorescence observations on time evolu-tion of folding are limited only by the time resolution of initiation of folding. (For example, time-resolution in steady state observations in stopped-flow and flash photolysis experiments is ~1-2 ms and ~5 ns, respec-tively). In contrast, time-resolution of folding kinetics monitored by time-resolved fluorescence is limited by the time needed to make a single observation with sufficient signal to noise ratio rather than the time needed to initiate the folding reaction. Methods based on TCSPC provide a time resolution of folding kinet-ics in the range of tens of millisecond35 and those based on streak camera enable slightly better time resolution36. The reader should note the difference between time-resolution of dynamic fluorescence sig-nal, which is spectroscopic and molecular dynamics in origin and the time resolution of protein folding kinetics which depends on reaction rate constants. The latter process is much slower than the former. Such double kinetics were applied to the slow folding process of barstar31. Monitoring rotational dynamics of the core Trp53 during progress of folding process revealed that the protein collapses initially and core gets rigidified prior to formation of secondary struc-ture. The transformation of initially collapsed form to the final compact native state in a time scale similar to that of secondary structure formation was also re-vealed from the dynamics of Trp5331.

Double kinetics experiments performed by moni-toring FRET between Trp53 and Cys-82-TNB pro-vided information on distance and distance distribu-tion between the two locations in the protein37,38. Thus, it helps in acquiring structural information on the intermediates along the folding pathway and their population distribution. By engineering location of fluorescence acceptor group TNB at various places in the protein, it was possible to estimate several in-tramolecular distances during the folding process38. Results from these studies revealed the heterogeneous

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nature of folding process, supporting the rugged land-scape model of folding39. (iii) Protein-ligand interactions

Fluorescence-based methods have been extensively used in this area and its reach next only to fluores-cence-based assay of biochemical reactions. Determi-nation of binding affinity of ligand is the main aim in this level of usage. Fluorescence intensity from either protein or the ligand and changes in the quantum yield on binding has been the main observables. Mostly, binding results in changes in quantum yield and hence, estimation of binding constants becomes rou-tine albeit its own complexities40. However, situations where binding does not lead to significant change in the quantum yield, fluorescence anisotropy of fluores-cent ligand has been effectively used to visualize the binding isotherm41. It becomes necessary to label the ligand rather than the protein. Fluorescence anisot-ropy of free ligand is significantly small when com-pared to the protein-ligand complex, due to larger size of the complex. Since global rotational correlation time (τrg) scales linearly with the molecular volume, V (τrg = ηV/kT, where η is the microviscosity, k is Boltzman constant and T is absolute temperature) the stead-state anisotropy (rss = r0/(1+τf/τrg) where r0 is the limiting anisotropy and τf is fluorescence lifetime) increases on binding. Due to its simplicity and robust-ness of measurement of steady-state anisotropy, it has become one of the methods of choice in high throughput screening of drugs, etc.

In recent years, fluorescence correlation spectros-copy (FCS) has become a powerful tool in studying protein-ligand interactions by registering the fluctua-tions of fluorescence intensity (see the section ‘Ob-servable parameters of fluorescence’) of ligand5-7. Fluctuation arises due to Brownian diffusion of mole-cules in and out of a very small observation volume. Autocorrelation of such intensity fluctuations are used to extract the translational diffusion coefficient and the average number of molecules in the observation volume. Thus, the latter parameter gives the concen-tration of the free ligand which can be used to con-struct the binding isotherm. The main advantage of FCS is its capability to probe in extremely sparse conditions such as situations inside a single cell. Di-rect real time measurement of association and disso-ciation kinetics in protein-ligand interactions has been demonstrated from FCS measurements42. However, in spite of these advantages, FCS is yet to catch up as a

routine analytical tool due to expensive instrumenta-tion and very strict sample conditions such as low background signals. (iv) Sensing of ions

A large number of fluorescent molecules have been developed for sensing ions such as H+, Ca2+, K+ , etc. and newer probes are being discovered at a significant rate43. The main advantages of fluorescence-based sensors44 are the simplicity of measurement, the high sensitivity of fluorescence and the ability to monitor with high spatial resolution in living cells. The limita-tions often encountered are the selectivity for a par-ticular ion in a medium composed of several species and the uncertainty associated with the probe volume especially in microscopic spaces. The observation volume is essential for estimation of concentration from fluorescence intensity and this is a nontrivial problem in microscopy of cellular structures. To overcome this problem, ratiometric probes have been developed. They are associated with dual emission spectra and an isoemissive point at which the fluores-cence intensity is insensitive to concentration of the species being sensed. Hence, ratio of emission inten-sity at emission maximum and at the isoemissive point is independent of the observation volume and the concentration of the probe45. In situations, where such biemission ratiometric probes are unavailable, covalent coupling of the sensor fluorophore with an-other fluorophore which is insensitive to the analyte may be used; the two fluorophores could be designed to have mutually exclusive emission band positions46.

Binding of fluorescence probes to molecules other than the species being sought after is often encoun-tered in cell biology. It could be overcome by moni-toring fluorescence lifetime instead of steady-state intensity, which is more popular due to its simplicity. Time resolution of the fluorescence signal, which also enables us to separate the population of probes bound to non-specific locations. This technique was used in the case of estimation of intracellular pH with the aid of fluorescent pH sensor SNARF-147. It also demon-strated the need to correct for probe binding for ob-taining reliable estimates of intracellular pH. (v) Protein-Nucleic acid interaction

Molecular level description of cellular processes such as growth, differentiation and development in-volves specific interaction between nucleic acid and proteins. The plethora of molecular components and variety of mechanisms that are being brought out in

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this area of research is quite breathtaking. Popular techniques such as gel shift assay and filter binding assay for studying the interaction have following seri-ous limitations. These techniques separate out the components involved in interaction and hence do not assess equilibrium status of interaction. Furthermore, they are unsuited for weaker binding between protein and nucleic acid segment48. Fluorescence-based methods are ideal for such situations49. They allow us to observe under equilibrium conditions in a wide range of environments that are close to physiological situations. As before, fluorescence label could be lo-cated either on protein or on the nucleic acid. How-ever, in many situations, it has been found that label-ing the nucleic acid is more effective for the following reasons: (i) delineating the binding isotherm of a small fragment of nucleic acid by the method of steady-state anisotropy (see section iv) requires fluo-rophore to be located in smaller of the two interacting partners, viz. the nucleic acid fragment; (ii) fluores-cence labeling of nucleic acid could be more conven-iently performed at the terminal residue50 and labeling at a specific location in the nucleotide sequence could be achieved by using fluorescent analog of adenine, 2-aminopurine51-53; and (iii) specific labeling of the much larger component, the protein is complicated by lack of information on the suitable site and also the increased level of technical difficulty involved. How-ever, the above information could be used only as general guidelines and several exceptions to the above are regularly encountered. Also, FRET studies which reveal valuable structural information on the complex requires that appropriate (intrinsic or extrinsic) labels are inserted into both the interacting partners.

Estimation of binding affinity between the nucleic acid and the protein is the most commonly encoun-tered application of fluorescence. Since there could be many sites for the protein on a single molecule of DNA and also protein binding could be cooperative, evaluation of binding isotherms requires special ana-lytical treatments of the binding data54,55.

Several other interesting aspects of protein-nucleic acid interactions could also be addressed by fluores-cence-based methods. Structure and dynamics of the nucleic acid component in the complex is one of them. Furthermore, novel and complex architectures of multistranded DNA have been elucidated by FRET based methods56,57. Population distribution in such exotic structures and changes in population distribu-tion caused by changes in the dynamics of these struc-

tures could also be revealed by techniques such as FRET and fluorescence lifetime distribution56,57. One of the unique applications of fluorescence in such massive complexes is the ability to address the role of dynamics in eliciting physiological function.

The answer to the question why apparently similar structures do not elicit similar activities could lie in differences in their dynamics. This paradigm was tested recently in RecA-mediated recombination be-tween two double-stranded DNA molecules which have homologous regions in them58. One of the ele-mentary steps in the overall process is formation of a three-stranded complex from RecA-coated single strand and a homologous double strand. Strand ex-change involving long stretches of homology tracts requires hydrolysis of ATP by RecA, which has coated one of the strands. The non-hydrolyzable ana-log of ATP, ATPγS is unable to effect strand ex-change, although a three stranded structure (similar to that formed in the presence of ATP) is formed in the presence of ATPγS. With the aim of monitoring dy-namics of DNA-RecA filament DNA was labeled at several single locations with 2-aminopurine (2-AP) which is a fluorescent analog of adenine and forms Watson-Crick base pairs similar to that of adenine51-53. The extent of base-pairing is shown by the amplitude of the shortest fluorescence lifetime of 2-AP59. The dynamics of DNA was monitored through fluores-cence depolarization kinetics of 2-AP60. Results from all these observations showed that ATP hydrolysis induces an unique segmental dynamics in DNA-RecA filament which was absent in the presence of ATPγS60. It suggests that the segmental dynamics set-up by ATP hydrolysis is an obligatory requirement for exchange of homologous tracts. It would be interest-ing to see whether such dynamics-induced steps are common features of many control processes in mo-lecular interactions in biology. (vi) Membrane structure and dynamics

The current view of cell membranes portrays them as heterogeneous mixtures of various types of lipids, proteins and carbohydrates61. It differs from that of classical fluid-mosaic model62 , according to which the lipid component is assumed to be homogeneous and an inert barrier. Heterogeneity in cell membranes could be both in its structural and dynamic aspects. Structural heterogeneity arises due to non-ideal mix-ing of lipid mixtures which are chemically heteroge-neous. It could result in domains (1-10 μm dia.) and

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may be visualized directly by microscopic tech-niques63. On the other hand, lipid clusters formed from a small number (~5-50) of molecules are harder to visualize by direct methods, due to their sub-micron size and their dynamic nature64. Such clusters, named ‘rafts’61, are implicated in cellular processes such as endocytosis and membrane trafficking, etc. They also could act as ‘hot spots’ of intracellular sig-naling61. Heterogeneity in temporal domain (i.e. dy-namic heterogeneity) is expected to control transport of species across membranes65. Fluorescence-based methods are ideally suited in studying both types of domains.

Quantitative information on the level of heteroge-neity (structural and dynamic) may be obtained in several ways by using fluorescence probes in mem-branes. Simultaneous use of two specific probes, hav-ing differential preference for partitioning into ‘gel’ and ‘liquid crystalline’ domains co-existing in a membrane and monitoring FRET efficiency between them has been successfully used in several systems67. Extracting quantitative information from these studies requires extensive modeling and analysis68.

Heterogeneity in the fluorescence lifetime of probes incorporated into membranes is another pow-erful approach65,69-71. Several factors contribute to life-time heterogeneity: (i) heterogeneity in the composi-tion of lipid molecules surrounding the probe; here, the fluorescence lifetime is controlled by the extent of non-radiative decay of excited state induced by nearby lipid; (ii) gel-liquid crystal phase state being different locally around each probe molecule; this in turn could control again the non-radiative rate; and (iii) difference in the orientation of flourophore with respect to the membrane plane; variation in lifetime in this case arises from the dependence of radiative rate on the molecular orientation72,73. It is easy to visualize that structural and dynamical heterogeneity would contribute to heterogeneity in fluorescence lifetime.

The polarity-sensitive probe, Nile Red74 was used for reporting heterogeneity and dynamics of lipid bi-layer vesicles65,69, supported membranes70 and cell69,75 membranes. Decay of fluorescence intensity analyzed by MEM provides an unbiased estimate of the popu-lation heterogeneity13,34,69. We found that the width of lifetime distribution was ~3 fold higher in the gel state as compared to liquid crystalline state of the mem-brane69. It could be interpreted as due to slower reori-entation of Nile Red in the gel state, as compared to its fluorescence lifetime. Apart from physical state of

membrane, the width of lifetime distribution was de-pendent on lipid composition, also. Specifically, the width increased significantly with cholesterol content of the membrane. This approach could be used to map the structure and dynamics of membranes in live sin-gle cells75.

Information on structure and dynamics of mem-branes becomes more interesting when they could be correlated with their functional activity. Transport of protons across lipid bilayers is a case in point due to its anomalously high value when compared to other monovalent cations76. We used fluorescence decay kinetics of Nile Red in membranes of various compo-sitions to arrive at the conclusion that the transport efficiency depends upon water content of lipid mem-branes and thus providing strong support to the ‘wa-ter-wire’ hypothesis of proton transport (Fig. 6).

Visualization and structural information on the cell surface architecture is best obtained from fluores-cence techniques. Perhaps fluorescence is the only method available for visualizing molecular processes occurring in single live cells! Changes in steady-state anisotropy caused by homo-FRET has been effec-tively used to show the presence of glycosyl-phophotidylinositol (GPI)-anchor-induced ‘rafts’ in cell surface64. In these experiments, fluorescence la-beled ligand for GPI anchored membrane bound re-ceptors was used as the probe. Information on the dis-tance between probes bound on adjacent receptor pro-tein was obtained from time-resolved anisotropy de-cay measurements77. Estimates of the distance (<4 nm) indicated a closely-packed structure for these clusters77. Interpretation of steady-state and time-resolved anisotropy changes as caused by homo-FRET would require exclusion of rotational dynamics as the cause of depolarization. This is feasible when

Fig. 6—H-bonded water wire spanning the membrane facilitating transport of protons [This model gets support from fluorescence dynamics of Nile Red and its correlation with transport rates (refs 65, 75)]

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rigid probes such as green fluorescent protein (GFP) is used rather than a fast rotating small molecular probe such as fluorescein77. (vii) Probing the interior of cells

Monitoring cellular processes or intracellular envi-ronment in a single living cell requires extremely sen-sitive techniques and fluorescence methods have the capability to meet the challenge. The applicability of in vitro studies to understand cellular processes re-quires information on intracellular environment. The physical state of cytoplasm is one such parameter and has been debated extensively78,79. These are based on the possibility that the high concentration of proteins (often exceeding 50 mg/mL) present in cytoplasm could result in alterations in the property of the aque-ous phase. We69,80 and others81 estimated the fluid phase viscosity of cytoplasm by monitoring rotational diffusion of hydrophilic fluorescent molecules dis-persed in the cytoplasm. Rotational dynamics is con-trolled by microviscosity felt by the probe. Hence, microviscosity could be estimated inside single living cells with a spatial resolution of ~1 μm80. The most striking conclusion from these studies is that cyto-plasmic viscosity is very nearly similar to that of wa-ter and is higher only by a factor of ~1.3 on an aver-age. It dispelled the myth that the water inside cells could have properties significantly different, com-pared to bulk water. Further, these studies also re-vealed the presence of small, but significant variation in cytoplasmic viscosity dependent upon the cell type and spatial location within a single cell80. The rele-vance of these findings may lie in functional com-partmentalization of cytoplasm which could enhance functional efficiency. It would be interesting to map the entire cell using parameters such as fluorescence lifetime82 and fluorescence depolarization time83.

Cellular localization of a protein of interest is often carried out by the use of fluorescent proteins such as GFP and their numerous variants which are co-expressed along with the protein of interest84. Al-though this method is very powerful in spatial local-ization of the protein, it is limited in delineating the dynamics due to massive size of the fluorescent probe. However, a recent method where a small molecule probe (fluorescein linked to two arsenoxides and a hexapeptide having four cysteines, abbreviated as FlAsH85) could be specifically attached to the pro-tein of interest in living cells could overcome the limi-

tation. It may provide exciting information on in vivo dynamics of proteins and their complexes in cells.

Once the macromolecule of interest inside a living cell is fluorescence labeled, then techniques such as FCS5-7,86 and time-resolved fluorescence methods69 could be used in extracting information on in vivo dynamics. For example, experiments such as scanning the genome of a single cell for a specific sequence by using a fluorescence labeled primer and FCS could be exciting. Characteristic diffusion time (obtainable from FCS) of the primer would increase significantly on hybridization with the genomic sequence. Such experiments could also provide answers to the mechanism of search on the DNA sequence and dif-ferential rigidity of segments of chromosome. Simi-larly, time-resolved fluorescence of the macromole-cule of interest inside a living cell would offer us a view, which would be significantly closer to reality when compared with the ‘reductionist’ approach of studying cell-free systems. Such futuristic experi-ments await developments of new methodologies, for example in situ labeling of a specific macromolecule in a cell by extension of in vitro translation of proteins for specific incorporation of a fluorescent amino acid87. Conclusion

Due to its high sensitivity and selectivity, fluores-cence spectroscopy is expected to remain as one of the very revealing windows for observations on the complex biochemical reactions and physiological pro-cesses. Although a large variety of techniques and methodologies are already in use, we expect many new innovative ideas and methodologies to be devel-oped. This would require merging of ideas from areas as diverse as organic synthetic chemistry, semicon-ductor physics and molecular biology. References

1 Brand L & Johnson M L eds (1997) Methods Enzymol 278 2 Lakowicz J R (1999) Principles of Fluorescence Spectros-

copy, Plenum Press, New York 3 Lakowicz J R ed Topics in Fluorescence Spectroscopy, Ple-

num Press, New York, Vol. 1 Techniques (1991); Vol. 2 Principles (1991); Vol. 3 Biochemical Applications (1992); Vol. 4 Probe Design and Chemical Sensing (1994); Vol. 5 Non-Linear and Two Photon-Induced Fluorescence (1997); Vol. 6 Protein Fluorescence (2000)

4 Valeur B (2002) Molecular Fluorescence: Principles and Applications, Wiley-VCH, Weinheim

5 Rigler R & Elson E S, eds (2001) Fluorescence Correlation Spectroscopy: Theory and Applications, Springer-Verlag, Berlin

INDIAN J. BIOCHEM. BIOPHYS., VOL. 40, JUNE 2003

158

6 Hess S T, Huang S, Heikal A & Webb W W (2002) Bio-chemistry 41, 697-705

7 Thompson N L, Lieto A M & Allen N W (2002) Curr Opin in Struct Biol 12, 634-641

8 Ahmed I & Krishnamoorthy G (1994) Biochemistry 33, 9675-9683

9 Bruchez M, Moronne M, Gin P, Weiss S & Alivisatos A P (1998) Science 281, 2013-2016

10 Chan W C W & Nie S (1998) Science 281, 2016-2018 11 Murphy C J (2002) Anal Chem 74, 521A-526A 12 Dubertret B, Skourides P, Norris D J, Noireaux V, Brivanlou

A H & Libchaber A (2002) Science 298, 1759-1762 13 Swaminathan R, Nath U, Udgaonkar J B, Periasamy N &

Krishnamoorthy G (1996) Biochemistry 35, 9150-9157 14 O’Connor D V & Phillips D (1984) Time Correlated Single

Photon Counting, Academic Press, London 15 Mialocq J –C & Gustavsson T (2001) in New Trends in

Fluorescence Spectroscopy: Applications to Chemical and Life Sciences (Valeur B & Brochon J C, eds), pp 61-80, Springer-Verlag, Berlin

16 James D R, Siemiarczuk A & Ware W R (1992) Rev Sci In-strum 63, 1710-1716

17 Frauenfelder H, Sligar G & Wolynes P G (1991) Science 254, 1598

18 Hammes-Schiffer S (2002) Biochemistry 41, 13335-13343 19 Swaminathan R, Krishnamoorthy G & Periasamy N (1994)

Biophys J 67, 2013-2023 20 Liu B, Thalji R K, Adams P D, Fronczek F R, McLaughlin

M L & Barkley M D (2002) J Amer Chem Soc 124, 13329-13338

21 Swaminathan R, Periasamy N, Udgaonkar J B & Krish-namoorthy G (1994) J Phys Chem 98, 9270-9278

22 Eftink M R (1991) in Topics in Fluorescence Spectroscopy (Lakowicz J R, ed), Vol 2, pp 53-126, Plenum Press, New York

23 Nandi N & Bagchi B (1997) J Phys Chem B 101, 10954 24 Pal S, Peon J & Zewail A H (2002) Proc Natl Acad Sci USA

99, 1763-1768 25 Mandal D, Datta A, Pal S K & Bhattacharyya K (1998) J

Phys Chem B 102, 9070 26 Lakshmikanth G S & Krishnamoorthy G (1999) Biophys J

77, 1100-1106 27 Krishnamoorthy G & Hinkle P C (1988) J Biol Chem 263,

17566-17575 28 Hartley R W, Rogerson D L & Smeaton J R (1972) Prep

Biochem 2, 243 29 Agashe V R, Shastry M C & Udgaonkar J (1995) Nature

(London) 377, 754-757 30 Nolting B, Golbik R, Niera J L, SolerGonzalez A S, Schriebe

G & Fersht A R (1997) Proc Natl Acad Sci USA 94, 826-830 31 Sridevi K, Juneja J, Bhuyan A, Krishnamoorthy G & Ud-

gaonkar J B (2000) J Mol Biol 302, 479-495 32 Lakshmikanth G S, Sridevi K, Krishnamoorthy, G & Ud-

gaonkar J B (2001) Nature Struct Biol 8, 799-804 33 Brochon J C (1994) Methods Enzymol 240, 262-311 34 Swaminathan R & Periasamy N (1996) Proc Ind Acad Sci

Chem Sci 108, 39-49 35 Beechem J M (1997) Methods Enzymol 278, 24-49 36 Lyubovitsky J G, Gray H B & Winkler J R (2002) J Amer

Chem Soc 124, 5481-5485

37 Lakshmikanth G S (2001) pH D Thesis, University of Mum-bai, Mumbai

38 Sridevi K, Krishnamoorthy G & Udgaonkar J B (2003) Bio-chemistry (submitted)

39 Chan H S & Dill K A (1998) Proteins: Structure, Function and Genetics 30, 2-33

40 Eftink M R (1997) Methods Enzymol 278, 221-257 41 Terpetschnig E, Szmacinski H & Lakowicz J R (1995) Anal

Biochem 227, 140 42 Schewille P, Oelenschlager F & Walter N G (1996) Bio-

chemistry 35, 10182-10193 43 Scheller F W, Wollenberger U, Warsinke A & Lisdat F

(2001) Curr Opinion Biotech 12, 35-40 44 de Lorimer R M, Smith J J, Dwyer M A, Looger L L, Sali K

M, Paavola C D, Sadigov S, Conrad D W, Loew L & Hel-linga H W (2002) Protein Sci 11, 2655-2675

45 Kao J P Y (1994) Methods Cell Biol 40, 155-181 46 Sonawane N D, Thiagarajah J R & Verkman A S (2002) J

Biol Chem 277, 5506-5513 47 Srivastava A & Krishnamoorthy G (1997) Anal Biochem

249, 140-146 48 Sitlani A & Crothers D M (1998) Proc Natl Acad Sci USA

95, 1404-1409 49 Yang M & Millar D P (1997) Methods Enzymol 278, 417-

444 50 Morrison L E, Halder T C & Stols L M (1989) Anal Biochem

188, 231-244 51 Strivers J T (1998) Nucleic Acid Res 26, 3837-3844 52 Rachofsky E L, Osman R & Ross J B (2001) Biochemistry

40, 946-956 53 O’Niell M A & Barton J K (2002) J Amer Chem Soc 124,

13053-13066 54 McGhee J D & von Hippel P H (1974) J Mol Biol 86, 469-

489 55 Bujalowski W & Lohman T M (1987) Biochemistry 26,

3099-3106 56 Klostermeier D & Millar D P (2002) Biopolymers (Nucleic

Acid Sci) 61, 159-179 57 Parkhurst L J, Parkhurst K M, Powell R, Wu J & Williams S

(2002) Biopolymers (Nucleic Acid Sci) 61, 180-200 58 Roca A I & Cox M M (1997) Prog Nucleic Acid Res Mol

Biol 56, 129-223 59 Hochstrasser R A, Carver T E, Sowers L C & Millar D P

(1994) Biochemistry 39, 11971-11979 60 Reddy R, Sen S, Rao, B J & Krishnamoorthy G (2003) Bio-

chemistry (submitted) 61 Simons K & Ikonen E (1997) Nature (London) 387, 569-572 62 Singer S J & Nicholson G L (1972) Science 175, 720-731 63 Edindin M (1997) Curr Opin Struct Biol 7, 528-532 64 Varma R & Mayor S (1998) Nature (London) 394, 798-801 65 Ira & Krishnamoorthy G (2001) J Phys Chem B 105, 1484-

1488 66 Hao M & Maxfield F R (2002) J Fluores 11, 287-295 67 Loura L M S, de Almeida R F M & Prieto M (2001) J Fluo-

res 11, 197-210 68 Loura L M S, Fedorov A & Prieto M (2001) Biophys J 80,

776-778 69 Krishnamoorthy G & Srivastava A (1997) Curr Sci 72, 835-

845 70 Ira & Krishnamoorthy G (1998) Biochim Biophys Acta 1414,

255-259

KRISHNAMOORTHY: FLUORESCENCE SPECTROSCOPY IN MOLECULAR DESCRIPTION

159

71 Mateo C R, Acuna A U & Brochon J C (1995) Biophys J 68, 978-987

72 Toptygin D & Brand L (1993) Biophys Chem 48, 205-220 73 Krishna M M G & Periasamy N (1998) J Fluores 8, 81-91 74 Dutt G B, Doraiswamy S, Periasamy N & Venkataraman B

(1990) J Chem Phys 93, 8498-8513 75 Krishnamoorthy G & Ira (2001) J Fluores 11, 247-253 76 Paula S, Volkov A G, van Hoek A N, Haines T H & Deamer

D W (1996) Biophys J 70, 339 77 Sharma P, Varma R, Saraji R C, Ira, Krishnamoorthy G, Roa

M & Mayor S (2003) Cell (submitted) 78 Minton A P (2001) J Biol Chem 276, 10577-10580 79 Verkman A S (2002) Trends Biochem Sci 27, 27-33 80 Srivastava A & Krishnamoorthy G (1997) Arch Biochem

Biophys 340, 159-167

81 Fushimi K & Verkman A S (1991) J Cell Biol 112, 719-725 82 Cubeddu R, Comelli D, D’Andrea C, Taroni P & Valentini G

(2002) J Phys D: Applied Phys 35, R61-R76 83 Clayton A H A, Hanley Q S, Arndt-Jovin D J, Subramaniam

V & Jovin T M (2002) Biophys J 83, 1631-1649 84 Tsien R Y & Miyawaki A (1998) Science 280, 1954-1955 85 Griffin A, Adams S R, Jones J & Tsien R Y (2000) Methods

Enzymol 327, 565-578 86 Dittrich P, Malvezzi-Campeggi F & Schwille P (2001) Biol

Chem 382, 491-494 87 Cohen B E, McAnaney T B, Park E S, Jan Y N, Boxer S G &

Jan L Y (2002) Science 296, 1700-1703 88 Krishnamoorthy G (1986) Biochemistry 25, 6666-6671 89 Krishnamoorthy G, Duportail G & Mely Y (2002) Biochem-

istry 41, 15277-15287