Exploring Hydrophobic Subdomain IIA of the Protein Bovine Serum Albumin in the Native, Intermediate,...

Exploring Hydrophobic Subdomain IIA of the Protein Bovine Serum Albumin in theNative, Intermediate, Unfolded, and Refolded States by a Small Fluorescence MolecularReporter

Bijan Kumar Paul, Anuva Samanta, and Nikhil Guchhait*Department of Chemistry, UniVersity of Calcutta, 92 A. P. C. Road, Calcutta 700009, India

ReceiVed: January 1, 2010; ReVised Manuscript ReceiVed: March 23, 2010

A simple intramolecular charge transfer (ICT) compound, 5-(4-dimethylamino-phenyl)-penta-2,4-dienoic acidmethyl ester (DPDAME), has been documented to be a potential molecular reporter for probing microhet-erogeneous environments of a model transport protein bovine serum albumin (BSA) using spectroscopictechniques. Meteoric modifications to the emission profile of DPDAME upon addition of BSA come out tobe a result of its binding to hydrophobic subdomain IIA. The highly polarity-sensitive ICT emission ofDPDAME is found to be a proficient extrinsic molecular reporter for efficient mapping of native, intermediate,unfolded, and refolded states of the protein. Experimental data coupled with a reinforcing support fromtheoretical simulation using CHARMM22 software confirm the binding site of the probe to be the subdomainIIA of BSA, while FRET study reveals a remarkably close approach of our extrinsic molecular reporter toTrp-212 (in domain IIA): the distance between DPDAME and Trp-212 is 1.437 nm. The caliber of DPDAMEas an external fluorescence marker also extends to the depiction of protein-surfactant (BSA-SDS) interactionto commendable fruition. Additionally, the protective action of small amounts of SDS on urea-denaturedprotein is documented by polarity-sensitive ICT emission of the probe. The present study also reflects theenhancement of the stability of BSA with respect to chemically induced denaturation by urea as a result ofbinding to the probe DPDAME.

1. Introduction

Serum albumins are abundantly found in blood plasma andare often considered as transport proteins, and this class ofproteins belongs to the most widely studied category. Theyfunction as carriers for numerous exogenous and endogenouscompounds in the body. The primary structure is composed of583 amino acid residues and is characterized by low tryptophancontent along with a high content of cystine, stabilizing a seriesof nine loops. The secondary structure of serum albumins has67% of helix of six turns and 17 disulfide bridges.1-3 The tertiarystructure is composed of three domains I, II, and III, and eachdomain is constituted of two subdomains A and B.1,2 Becauseof the common interface between domains II and III, bindingof a probe to domain III associates conformational changes ofdomain II and hence its binding affinities. Bovine serum albumin(BSA) displays approximately 80% sequence homology and arepeating pattern of disulfides, which are strictly conserved. Themolecule BSA contains two tryptophan residues, Trp-134 andTrp-212, of which the former is located in hydrophilic subdo-main IB, and it is proposed to be located near the surface ofthe albumin molecule in the second helix of the first domain.2

The protein BSA is known to exhibit a very high conformationaladaptability to a large variety of ligands.4-6 On the basis ofstudies using absorption, fluorescence, and circular dichroismspectroscopy,7-9 information on the binding process of manyexogenous ligands like long-chain fatty acids, amino acids,metals, drug, bilirubin, etc., has been furnished at the molecularlevel. There are also reports where such binding has been found

to enhance the solubility of the ligands6 and the toxicity of someligands like bilirubin diminished on binding to albumins.9

On another extreme, the past few years in the field ofphotochemistry and photobiology have witnessed an awesomeevolution of research surrounding the photophysical studies onorganic molecules of donor (D)-acceptor (A) pattern. Thepioneering work of Lippert et al.10 to observe the anomalousdual fluorescence from the model compound N,N-dimethylaminobenzonitrile (DMABN) envisaged a new arena of research inthe realm of photochemistry through intramolecular chargetransfer (ICT) reaction. Ever since its first report, the elatingphenomenon of ICT reaction has continued to grow on capturingattention of researchers because of its tremendous potential toform a splendid avenue for a massive lexicon of applicativeresearch.10-25 These include applications as electroopticalswitches, chemical sensors, fluorescence probes, and so forth.26-29

In a previous work,30 we have reported the photophysicalproperties of DPDAME (Scheme 1) by a detailed spectroscopicstudy in combination with quantum chemical calculations. Thepresent program aims at utilizing DPDAME as a molecularreporter for the investigation of the microenvironment of

* To whom correspondence should be addressed. Phone: 91-33-2350-8386. Fax: +91-33-2351-9755. E-mail: [email protected].

SCHEME 1: Schematic of the Structure of DPDAME

J. Phys. Chem. B 2010, 114, 6183–6196 6183

10.1021/jp100004t 2010 American Chemical SocietyPublished on Web 04/16/2010

proteinous medium. The remarkable sensitivity of the emissionspectral properties of DPDAME toward medium polaritythrough the operation of ICT reaction paved the way forimplementing DPDAME as an extrinsic fluorescent reporter forthe study of proteinous microenvironment, since the process ofincorporation of the probe into protein cavity will be reflectedthrough the significant and interesting modifications on theemission profile of the former. Additionally, steady-state ani-sotropy, red-edge excitation shift (REES), acrylamide-inducedfluorescence quenching, and fluorescence resonance energytransfer (FRET) studies have been exploited to extract valuableinformation regarding the nature and mechanism of the bindingphenomenon through deciphering the modified photophysics ofBSA-bound DPDAME. Thermal and chemical denaturationof the protein and also protective influence of small amountsof SDS on urea-induced denaturation have been manifestedsimply through modulation of the emission spectral behaviorof DPDAME. Very recently, a nice report by Abou-Zied andAl-Shihi3a explored the native, unfolded, and refolded states ofhuman serum albumin (HSA) mainly by using intrinsic tryp-tophanyl fluorescence of HSA and fluorescence lifetime tech-niques. The present study advocates the documentation ofDPDAME as a new and efficient extrinsic molecular reporterfor microheterogeneous environments of the protein BSA.

2. Experimental Section

2.1. Materials. The molecule DPDAME was synthesized andpurified following a literature procedure and is describedelsewhere.30 The purity of the sample was established on a TLCplate before use. Tris buffer was purchased from SRL (India),and 0.01 M Tris-HCl buffer of pH 7.4 was prepared. Bovineserum albumin (BSA) and sodium dodecyl sulfate (SDS) werepurchased from SRL (India) and used as received. Analyticalgrade urea from SRL (India) was used after recrystallizing twicefrom MeOH (AR grade, Spectrochem, India). Triple distilledwater was used for the preparation of all solutions. The solventappeared visually transparent, and its purity was also tested byrunning the fluorescence spectra in the studied wavelength range.

2.2. Instrumentations and Methods. The absorption andemission spectra were acquired on a Hitachi UV-vis U-3501spectrophotometer and Perkin-Elmer LS-50B fluorimeter, re-spectively. In all measurements, the sample concentration wasmaintained in the range 10-6-10-7 M in order to avoidaggregation and reabsorption effects. Experiments have beencarried out at an ambient temperature of 25 °C, unless otherwisespecified. Only freshly prepared air-equilibrated solutions wereused for spectroscopic measurements. For spectral backgroundcorrections, a similar set of solutions in increasing BSAconcentration was prepared except that the probe was omitted.During protein denaturation study by urea, spectral backgroundcorrections have been ensured with a set of solutions without aprobe but containing the protein BSA (30 µM) and respectiveconcentrations of urea as required in each set of solutions underexperiment.

Fluorescence quantum yield (Φf) was determined usingrecrystallized �-naphthol as the secondary standard (Φf ) 0.23in methylcyclohexane) using the following equation:18-21,31

where A terms denote the fluorescence area under the curve,“Abs” denotes absorbance, n is the refractive index of the

medium, and Φ is the fluorescence quantum yield and thesubscripts “S” and “R” stand in recognition of respectiveparameters for the studied sample and reference, respectively.

Steady-state anisotropy measurements were carried out usinga HORIBA JOBIN YVON Fluoromax-4 spectrofluorimeter. Thesteady-state anisotropy is defined as3,6,7,31

in which IVV and IVH are the emission intensities when theexcitation polarizer is vertically oriented and the emissionpolarizer is oriented vertically and horizontally, respectively.G is the correction factor.

Fluorescence lifetimes were measured by a time-correlatedsingle photon counting (TCSPC) spectrometer using nanoLED-07 (IBH, U.K.) as the light source at 370 nm to trigger thefluorescence of DPDAME, and the signals were collected at amagic angle of 54.7°. The observed fluorescence intensities werefitted by using a nonlinear least-squares fitting procedure to afunction (X(t) ) ∫0

t E(t′)R(t - t′) dt) comprising the convolutionof the IRF (E(t)) with a sum of exponentials (R(t) ) A +∑i)1

N Biet/τi) with pre-exponential factors (Bi), characteristiclifetime (τi), and a background (A). The relative contributionof each component was obtained from a biexponential fitting,expressed by the following equation:

The mean (average) fluorescence lifetimes for the decay curveswere calculated from the decay times and the relative contribu-tion of the components using the following equation:31

The excellence of the fits was judged by �2 criteria and visualinspection of the residuals of the fitted function to the data.

Circular dichroism (CD) spectra were recorded on a JASCOJ-815 spectropolarimeter, using a cylindrical cuvette of 5 mmpath length. The reported CD profiles are an average of foursuccessive scans obtained at a 20 nm/min scan rate withappropriately corrected baseline. BSA concentration for CDmeasurements was 1.25 µM.

2.3. Theoretical Simulation Protocol. The native structureof HSA was taken from the Protein Data Bank having PDB ID1AO6. BSA was generated from it by performing necessaryadditions at the N-terminal as well as some mutations in therequired regions, as no PDB is available for BSA. The probeof our interest was also created by using CHARMM.32a Then,BSA was solvated in a water box and any water that comeswithin 2.6 Å from any protein atom was deleted. Then, threedifferent coordinates were prepared: (1) where the probe was

ΦS

ΦR)

AS

AR×

(Abs)R

(Abs)S×

nS2

nR2

(1)

r )(IVV - G · IVH)

(IVV + 2G · IVH)(2)

G )IHV

IHH(3)

an )Bn

∑i)1

N

Bi

⟨τf⟩ )∑

i

aiτi2

∑i

aiτi

(4)

6184 J. Phys. Chem. B, Vol. 114, No. 18, 2010 Paul et al.

near the W212 (Trp-212) residue, (2) where the probe was nearthe third domain of BSA, and (3) where the probe was at thecenter geometry of BSA. The final simulation box contained21918, 21927, and 21911 water molecules, respectively, for theabove three arrangements. Each of the systems was energyminimized by using the ABNR method until the difference inkinetic energy of the two successive steps was less than 0.0001kcal/mol. Then, each of the minimized structures was equili-brated for 65 ns and at 298 K. Coordinates after equilibrationwere generated by using the PyMOL software package32b whichshows that the stable interacting conformation in all three casesis the one stated in arrangement 1. The observed conformationmight be attributed to the hydrophobic interaction between theprobe and Trp-212 residue. SHAKE was applied to maintainthe bond lengths and bond angles of water molecules. PMEwas applied to measure the electrostatic interaction with a 9 Åcutoff. All of the simulations were performed using theCHARMM22 force field and parameters.

3. Results and Discussion

3.1. Solvatochromism and ICT Process in DPDAME. Themolecule DPDAME shows two broad and structureless absorp-tion bands at ∼370 and ∼260 nm in water which are ascribedto S1 r S0 and Sn r S0 (S2 r S0 or higher) transitions,respectively.30 Emission spectra of DPDAME exhibit charac-teristic dual fluorescence in polar solvents of which a solventpolarity insensitive higher energy local emission band is centeredat ∼425 nm and a lower energy but relatively intense chargetransfer band is located at ∼540 nm (in water). This large Stokesshifted (∆ν ) 8508 cm-1 in water) emission band arises fromthe CT state of DPDAME and thus experiences remarkablesolvent polarity dependency. Steady-state absorption and fluo-rescence spectroscopic studies of DPDAME in solvents ofdifferent polarity clearly establish the phenomenon of ICTreaction in DPDAME.21,30

3.2. Probe-Protein Complexation Equilibrium. The com-plexation reaction between the probe DPDAME and proteinBSA has been monitored by following the spectral changes ofthe probe upon binding to BSA. As seen in Figure 1, gradualaddition of BSA to a solution of DPDAME in aqueous buffer(Tris buffer, 0.01 M, pH 7.4) is associated with a red shift ofthe absorption maxima of DPDAME (from ∼370 to ∼395 nmfor the lower energy band and from ∼260 to ∼278 nm for thehigher energy band) with simultaneous increase in absorbance.This might be the first indication toward the occurrence of

interaction between the two concerned parties, although nothingconcrete can be derived from such a response because ofoverlapping regions of BSA-tryptophan absorption and thehigher energy band of the probe. The increment of absorbancecoupled with the red shift on the longer wavelength region ofDPDAME might be taken into account, as there is negligibleabsorption of tryptophan at this wavelength region. The proteinBSA-induced spectral changes in Figure 1 suggest stronginteraction between the two parties involved and can beconnected to the modulation of local polarity surrounding thefluorophore (DPDAME) which subsequently modulates thestabilization of its different energy levels. With lowering of localpolarity in the immediate vicinity of the fluorophore in proteinenvironments, the energy gap between the HOMO (highestoccupied molecular orbital) and LUMO (lowest unoccupiedmolecular orbital) decreases to give rise to the bathochromicshift. Such interpretation of our findings finds reasonableconsistency with literature reports.33 Furthermore, our interpreta-tion is substantiated by a relative red shift of absorption maximaof DPDAME upon moving from aqueous solution to a nonpolarmedium like n-hexane or methylcyclohexane (λabs

max ∼ 370 nmin water vs λabs

max ∼ 375 nm in n-hexane/methylcyclohexane).30

On the other extreme, the result of interaction of DPDAMEwith BSA is found to produce dramatic modifications on theemission profile in the form of a large blue shift of emissionmaxima (Figure 2a, from ∼540 nm in aqueous buffer to ∼509nm in 150 µM BSA) together with remarkable intensityenhancement. (However, this blue shift on the emission profileis not to be confused with the observed red shift on theabsorption spectral profile because of the enormously differentialnatures of the potential energy surfaces (PESs) of the groundand excited states.31 The ICT phenomenon is exclusively anexcited state affair, and the CT state is generated through LEstate excitation; i.e., the CT state does not put its signature onthe ground state PES.30,31 Hence, the interpretations have beenfabricated accordingly.) Usually, the neutral and hydrophobicnature of the probe molecule favors solubilizing in the hydro-phobic cavities of the protein BSA and such a large shift of theemission maxima toward the blue end of the spectrum is actuallya reflection of the high degree of sensitivity of the probe towardchanges in the polarity of its surrounding microenvironment.3,6

In fact, the emission maximum of DPDAME in nonpolar solventis blue-shifted with respect to that in water (from ∼540 nm inwater to ∼425 nm in methylcyclohexane).30 The blue shift inthe aforementioned case is thus argued on the basis of the ideaof encapsulation of the probe in the hydrophobic interior of theprotein backbone which thereby offers a reduced polarity inthe immediate vicinity of the probe inside proteinous mediumcompared to that in aqueous buffer phase. Solvent water is well-known to act as a quencher for the emission of the CT statedue to specific interactions like intermolecular hydrogenbonding,10-12,21 and as the probe molecules are less exposed towater when present inside the hydrophobic cavity of BSA, thedeactivation of nonradiative decay channels comes intooperation.6,7,12,31 The lower polarity of the interior of the proteincavity results in enhancement of the energy gap between theCT state and the triplet/ground states, which according to theenergy gap law leads to a reduction in the radiationless decayaccounting for the increment of the CT emission intensity withaddition of BSA.12,31 Increased quantum yield with increasingconcentration of BSA is also consistent with the same idea.Figure 2b portrays the variation of CT emission intensity ofDPDAME with increasing BSA concentration. An initial steeprise of intensity is found to be followed by attainment of a

Figure 1. Effect of increasing concentration of BSA (curves 1 f 14correspond to [BSA] (in µM) ) 0, 10, 15, 20, 25, 30, 35, 40, 45, 50,60, 70, 80, 90) on the absorption spectra of DPDAME ([DPDAME] )2.0 µM). The inset shows a magnified view of the changes in theabsorption profile of DPDAME on the lower energy absorption bandregion.

Exploring Hydrophobic Subdomain IIA of BSA J. Phys. Chem. B, Vol. 114, No. 18, 2010 6185

plateau region marking the saturation of the interaction beingat [BSA] ≈ 60 µM.

Though the modulations of the spectral properties of DPDAMEin the presence of BSA provide indications of interaction betweenthe probe and the protein, a quantitative estimate or the extent ofbinding of DPDAME to the hydrophobic cavity of protein isobtained from an examination of the emission data on theBenesi-Hildebrand relation.26,34 A detailed discussion on theBenesi-Hildebrand equation is avoided here, since it is routine andprofusely available in the literature6a,b,26,34 (a little elaboration is givenin the Supporting Information). We thus start with the equation

which upon rearrangement gives

in which I0, I, and I1 are the emission intensities, respectively, in theabsence of, at intermediate concentration, and at infinite concentration

of BSA. Thus, an analysis of the fluorescence data on eq 6 paves theway for simplistic mapping of the spectroscopic modulations on aquantitative scale through estimation of the binding parameters andstoichiometry of the DPDAME:BSA complex. As seen in Figure 2c,the plot of 1/[I - I0] vs 1/[BSA] produces a straight line indicatingthe formation of a 1:1 complex between DPDAME and BSA. Theexcellence of this linear fit can be justified from the correlation factorR) 0.9953 and standard deviation (SD) of 8.6 × 10-4. A quantitativeestimate of the extent of binding, i.e., the binding constant (K), isdetermined from the intercept to slope ratio of the Benesi-Hildebrandplot, and the computed value is K) (12.55( 0.4) × 103 M-1. Usingthis value of K, the free energy change for this process of complexationis determined to be ∆G)-23.38 kJ-mol-1 (Table 1). A high K valueindicates strong binding between probe and protein BSA, and thefavorable process of complexation is dictated by the negative freeenergy change. The values obtained for K and ∆G are found to beargood consistency with literature reports.3,6,7,31,35-41

3.3. Polarity of the Microenvironment around the Fluo-rophore. Over the past few years, the determination of themicroscopic polarity of biological systems applying fluorescentprobes has been recognized as an efficient technique.42-46 Thepolarity determined through different photophysical parametersof the probe gives a relative measure of the polarity of themicroenvironments. In the present report, we have made anattempt to estimate the micropolarity of the proteinous environ-ments around the fluorophore. With the proteinous mediumbeing heterogeneous, the probe molecules experience variedpolarity depending on their precise location inside the proteinenvironment.3,6,31,36,46 As mentioned earlier, the blue shift of theCT emission band of DPDAME in protein solution (in Trisbuffer) reflects the lower polarity of the probe-binding site inthe protein backbone. To find out the polarity experienced byour probe molecules inside the protein solution, we haveemployed the high sensitivity of its CT emission band towardthe polarity of the medium, by taking the probe in differentpercentages of water and dioxane mixtures. The standard ET(30)values of the dioxane-water mixtures were determined on thebasis of the intramolecular charge transfer transition of betainedye 2,6-diphenyl-4(2,4,6-triphenyl-1-pyridino)phenolate.45 Thepolarity, in terms of ET(30), of the proteinous microenvironmentsurrounding the fluorophore can hence be determined on thebasis of the position of the emission maxima of the CT band inthese environments. As found from Figure 3, the micropolarityof BSA is determined to be 48.44 on the ET(30) scale, which isconsiderably low compared to that of bulk water (ET(30) )63.145) and hence accounts for the blue shift of the CT emissionband of DPDAME.

3.4. Steady-State Fluorescence Anisotropy Measurements:Microviscosity of the Environment around the Fluorophore.Steady-state fluorescence anisotropy measurement occupies acommanding position in biochemical and biophysical researchbecause of its delicate sensitivity toward sensing of any factor

Figure 2. (a) Effect of increasing concentration of BSA (curves 1 f15 correspond to [BSA] (in µM) ) 0, 4, 10, 15, 20, 25, 30, 35, 40, 45,50, 60, 90, 140, 150) on the fluorescence emission spectral profile ofDPDAME (λex ) 370 nm and ([DPDAME] ) 2.0 µM)). (b) Variationof intensity at emission maxima and λem

max (nm) of DPDAME as afunction of BSA concentration. (c) Benesi-Hildebrand plot of 1/[I -I0] vs 1/[BSA] (M-1) for binding of DPDAME with BSA (for [BSA]) 10, 20, 25, 30, 35, 40, 45, 50, 60, 90, 140, 150 in µM).

I )I0 + I1K[BSA]

1 + K[BSA](5)

1(I - I0)

) 1(I1 - I0)

+ 1(I1 - I0)K[BSA]

(6)

TABLE 1: Different Parameters Obtained fromDPDAME-BSA Binding

mediumKa

(M-1)∆Ga

(kJ ·mol-1)KSV

b

(M-1) Ec (%)kET

c

(ns-1)

BSA (12.55 ( 0.4) × 103 -23.38 0.913 86.96 1.216

a K is the binding constant, and ∆G is the free energy change asobtained for DPDAME-BSA complexation from analysis of theemission data on the Benesi-Hildebrand equation. b Stern-Volmerquenching constant for acrylamide-induced quenching of BSA-bound DPDAME. c E is the energy transfer efficiency, and kET is therate of energy transfer from Trp-212 of BSA to DPDAME duringFRET.


that influences the size, shape, or segmental flexibility of thefluorophore.3b,31,46 The degree of restrictions imparted by themicroenvironments on the dynamical properties of the probe isdirectly manifested through fluorescence anisotropy values31

whence it can fruitfully be exploited in assessing the probablelocation of the probe in the confined medium. The environmentof the probe molecule is governed by its precise location insuch a complex molecular assembly. The inferences drawn fromthe steady-state fluorescence measurements need to be comple-mented with the anisotropy values in order to provide anothersupportive platform to our findings and interpretations. Figure4a depicts a marked increase in the fluorescence anisotropy withincreasing concentration of BSA. Such gradual enhancementof anisotropy indicates interaction between the probe and BSA.At the same time, the trend of change of anisotropy withincreasing concentration of BSA reflects that the rotationaldiffusion of the probe is constrained significantly in theproteinous medium. Attainment of the plateau in Figure 4aimplies saturation in the binding interaction between the twoparties. The high anisotropy value of ∼0.30 in 50 µM BSA

indicates strong binding between DPDAME and BSA, comple-menting the values of complexation constant (K) and free energychange (∆G).

Fluorescence anisotropy is very sensitive to the viscosity ofthe environment around the fluorophore. The microviscosity ata given temperature is often estimated by comparing thefluorescence anisotropy of a fluorophore in an environment withthose of the probe in solvents of known viscosity.46a,47 In orderto determine the microviscosity of the proteinous environment,the fluorescence anisotropy of DPDAME in glycerol-watermixtures of different composition has been measured and acalibration curve has been constructed by plotting the anisotropyvalues against weight percent of glycerol (Figure 4b). Theanisotropy value suggests that the average environment around theprobe molecule upon incorporation into the protein cavity corre-sponds to approximately 90% glycerol-water mixture (weightpercentage) composition (ranis ≈ 0.30 in 90% glycerol-watermixture vs ranis ≈ 0.31 in 110 µM BSA; this concentration of BSAensures saturation of interaction with DPDAME, as discussed insection 3.2), i.e., a considerably hydrophobic region.

3.5. Steady-State Acrylamide Quenching of DPDAME:BSA Fluorescence. As displayed in Figure 5, the addition ofacrylamide to a solution of DPDAME in 60 µM BSA results inquenching of the fluorescence intensity of the emission bandwith a slight red shift of the emission maxima (from ∼511 to∼513 nm in 2 M acrylamide). Acrylamide does not bind to theprotein but is known for its activity as an efficient static quencherof tryptophanyl fluorescence of serum albumins.31 It releasesthe probe molecules from the hydrophobic sites by approachingclose to the site6,31 when release of the probe molecules fromthe hydrophobic protein environment to the aqueous phase isnecessarily accompanied with quenching of the CT emissionalong with a shift of the emission maxima to the red end of thespectrum. Thus, the extent of perturbation of BSA-boundfluorescence of DPDAME (here in terms of quenching) by anexternal agent like acrylamide can be exploited as an indirectbut fruitful strategy to assess the strength of binding of the probeto the protein. A careful scrutiny of the emission spectralchanges imparted by the presence of acrylamide will also throwlight on the probable binding site of the extrinsic fluorescencereporter.6,31 The inset of Figure 5 shows that the Stern-Volmerplot for acrylamide quenching of fluorescence of BSA-boundDPDAME leads to a Stern-Volmer constant of KSV ) slopeof the plot ) 0.913 M-1. Such low magnitude of KSV obtainedfor quenching of fluorescence of BSA-bound DPDAME alongwith a little red shift conforms to only an insignificantperturbation of emission properties of DPDAME when boundto BSA, signaling toward the possibility that the probe moleculesare deeply embedded in the hydrophobic pocket of the protein

Figure 3. Plot of the variation of emission maxima of DPDAME indioxane-water mixture against ET(30) values. The polarity of thebinding site of DPDAME in proteinous medium (BSA) is indicated.

Figure 4. (a) Variation of steady-state fluorescence anisotropy (λex )370 nm and λmonitored ) λem

max) of DPDAME with increasing concentra-tion of BSA. (b) Variation of steady-state fluorescence anisotropy (λex

) 370 nm and λmonitored ) λemmax) of DPDAME as a function of

composition of the glycerol-water mixture. Each data point is anaverage of 10 individual measurements.

Figure 5. Variation of emission spectra of the DPDAME-BSAcomposite system as a function of acrylamide concentration in 60 µMBSA: (inset) Stern-Volmer plot of I0/I vs [acrylamide] (M).


BSA, which in turn ascertains a feeble approach of the quencherto the fluorophore leading to only insignificant static quenching(acrylamide is a well-known static quencher of intrinsic tryp-tophanyl fluorescence of serum albumins).3,6,31

3.6. Fluorescence Resonance Energy Transfer. Ever sinceitsdiscovery,fluorescenceresonanceenergytransfer(FRET)6,31,35,48

continues to be exploited as a powerful tool for measuring thedistance (in nanometer scale) between donor fluorophore andacceptor fluorophore in vitro and in vivo. FRET is an electro-dynamic phenomenon that occurs between a donor (D) moleculein the excited state and an acceptor molecule in the ground state.The donor molecules typically emit at shorter wavelengths thatoverlap with the absorption spectrum of the acceptor.31,48 Energytransfer in the process occurs without the appearance of a photonand is the result of long-range dipole-dipole interactionsbetween the donor and acceptor. The FRET efficiency is foundto depend on three parameters:31 (i) the distance between donorand acceptor must be within the specified Forster distance of2-8 nm; (ii) there must be appreciable overlap between donorfluorescence and acceptor absorption bands (Figure 6a); (iii)proper orientation of the transition dipole of the donor andacceptor fluorophores. During FRET study, the donor emissionintensity is found to be depleted with concomitant incrementof acceptor emission intensity as the acceptor concentrationincreases. Figure 6b depicts the intrinsic fluorescence spectraof tryptophan of BSA with increasing concentration of the probeDPDAME. Increasing the concentration of DPDAME resultsin a diminution of tryptophanyl fluorescence of BSA withsimultaneous emergence of a new band at ∼504 nm which isthe CT emission of DPDAME molecule originating through theoperation of FRET (with a characteristic isoemissive point at464 nm). At the same time, the emission maximum of thetryptophan residue of BSA is found to undergo a slighthypsochromic shift in DPDAME:BSA complex (λem

max ∼ 340nm) compared to that in free BSA (λem

max ∼ 349 nm). Thisobservation can be rationalized on the basis of the idea of some

modification of the local environments of the Trp residueimparted by the presence of the probe. With a view to the neutraland hydrophobic character of DPDAME, it is not unlikely toassume a somewhat more hydrophobic environment of theemitting Trp residue in the presence of the external probe leadingto a little blue shift in the emission maxima.31 Also, thisobservation is consistent with a recent literature report.48e Atthe same time, Figure 14 illustrates the location of Trp-212 andDPDAME over a hydrophobic surface (see section 3.14).

Using the required spectroscopic parameters, we have cal-culated the FRET parameters for the presently investigatedsystem (according to Forster’s theory) as follows: a remarkablyhigh energy transfer efficiency E ) 86.96%, J ) 3.50 × 10-15

L mol-1 cm3, R0 ) 2.018 ( 0.1 nm, and r ) 1.437 ( 0.1 nm(the calculation procedures have been elaborated in the Sup-porting Information; here, J ) overlap integral between donoremission and acceptor absorption spectra, R0 ) Forster’sdistance, and r ) distance between the donor and acceptor).That the donor (tryptophan of BSA) and acceptor (DPDAME)can approach very close paves the way for very high energy-transfer efficiency. This also indicates that the probe can measurethe proteinous microenvironment at a distance of 1.437 ( 0.1nm close to the tryptophan of protein; i.e., it can be used as ananometric ruler. As far as our knowledge goes, it seems to bethe first documentation of an extrinsic molecular reporter whichis capable of approaching deep inside the protein cavity to thisextent. That the ratio r/R0 amounts to 0.712, i.e., within the range0.5-2.0, rationalizes the practical reliability of the FRET processto measure the distance between donor and acceptor chro-mophores in the present case.31 Also, the rate of energy transfer,i.e., kET ) τD

-1(R0/r)6 ) 1.216 ns-1, is found to be remarkablyfaster than the donor decay rate (τD ) 6.3 ns31). This in turnadvocates for the observed high efficiency of energy transfer.31

However, it is ethical to mention here that the value of theorientation factor κ2 used in the calculation is 2/3, which is notprecisely correct as it is the value for donor and acceptor thatrandomize by rotational diffusion prior to energy transfer.31 Thereasons behind using the value κ2 ) 2/3 have been ornatelyargued in the Supporting Information, and we advocate that withκ2 ) 2/3 no significant error is included.3,6,31,46,48 Indeed, ourresults are well assisted by available literature that describe theuse of κ2 ) 2/3 during FRET measurements in various confinedenvironments.3,6,31,35,46,48

Confirmation for the occurrence of FRET between the presentchoices of pair has been derived from blank experiments whichproduced negative results when conducted with the acceptor(DPDAME) alone;31,35 i.e., for the given excitation wavelength(280 nm), the total fluorescence coming from the acceptor(DPDAME) in the absence of the donor (BSA) is considerablylow compared to that in the presence of the latter.

However, predictions about the location of the probe duringFRET are not that easy, since the situation in BSA is prettycomplicated by the presence of two tryptophan moieties (Trp-212 and Trp-134). Previous studies reveal that Trp-134 is moreexposed to the aqueous phase and its fluorescence propertiesare quite different.49 Our findings in the following sections,however, tend to support the occurrence of FRET from Trp-212 of BSA to DPDAME. This proposition is indeed reinforcedfrom spectral blue shift on the emission profile of DPDAMEin the presence of BSA (section 3.2, Figure 2), which cor-roborates to binding of the extrinsic probe to the hydrophobicdomain of BSA, since Trp-212 and Trp-134 are, respectively,located at hydrophobic subdomain IIA and hydrophilic subdo-main IB.

Figure 6. (a) Overlap of emission spectrum of BSA (-9-) andabsorption spectrum of DPDAME (-0-) in Tris-HCl buffer (pH 7.4)and (b) fluorescence emission spectra of tryptophan in BSA (60 µM)with increasing concentration of DPDAME (curves 1 f 7 correspondto [DPDAME] (in µM) ) 0, 3.79, 7.59, 11.38, 15.18, 22.77, 26.57)(λex ) 280 nm).


3.7. Wavelength-Sensitive Fluorescence Parameters. Awavelength-sensitive tool for directly monitoring the environ-ment and dynamics around a fluorophore in a complex biologicalsystem and the solvation dynamics in an organized medium isthe “red-edge excitation shift” or REES, i.e., the shifting of theemission maxima to the red end of the spectrum upon shiftingof the excitation wavelength to the red end of the absorptionspectra of the fluorophore.49-51 Here, we have monitored thedependence of emission maxima of DPDAME on excitationwavelength under various conditions of binding to BSA. Theoccurrence of excitation-wavelength dependence is connectedto the presence of an ensemble of molecules in the ground statediffering in their solvation sites and hence energies.49-51

However, this condition alone does not ensure the occurrenceof REES because of rapid relaxation (in the form of rapidsolvation of the fluorescent state or energy transfer betweenenergetically different excited states or conformers) of theexcited state. Thus, apart from the condition of selectiveexcitation of energetically different species, REES is also subjectto slow (and hence incomplete) relaxation of the excited state.Precisely, the operation of REES is subject to the followingconditions: (a) The molecule should be polar with the dipolemoment higher in the excited state than that in the ground state.In fact, the extent of inhomogeneous broadening of theabsorption spectra allowing the provision of site photoselectionof energetically different species is dependent on the change ofdipole moment (∆µ) upon photoexcitation through the relation∆υ ) A∆µF-3/2(kT)1/2 according to the Onsager sphere ap-proximation50 (here, A is a constant that depends on the dielectricconstant of the medium, F is the Onsager cavity radius, and kis Boltzmann’s constant). For DPDAME, ∆µ ) 16.42 D30 (atthe DFT/B3LYP/6-31G(d,p) level of calculation and using theLippert-Mataga equation31). However, additional broadening,which can play even a greater role in inhomogeneous broadeningof absorption spectra, may be induced by specific interactionsof the sort of hydrogen bonding, electrostatic interactions, andso forth.49-51 (b) The solvent molecules around the fluorophoremust be polar, and the solvent reorientation time (⟨τsolvent⟩) shouldbe slower or comparable to the fluorescence lifetime (τf) of thefluorophore so that unrelaxed fluorescence can give rise toexcitation-wavelength-dependent emission behavior.

Figure 7 furnishes the information obtained from REESmeasurements of DPDAME under different conditions. It is seenthat the shift of excitation wavelength from 370 to 460 nm (i.e.,during REES measurements, scrupulous care has been devotedto the selection of excitation wavelength to ensure that thevariation of λex involves shifting to the red end of the absorptionspectra and not merely the absorption maxima; this is a verycrucial criterion for REES monitoring50,51) results in shift of

the emission maxima of the probe from 499 to 509 nm; i.e.,REES of 10 nm is observed in 80 µM BSA. This observation,therefore, suggests that binding of the probe to BSA offersrestriction to the rotation of solvent dipoles around the excitedfluorophore. Furthermore, given the complex structural archi-tecture of BSA, additional broadening of absorption spectra (asmentioned above) is not unlikely to contribute to the operationof REES in the present case. However, for the same change ofexcitation wavelength in the presence of 4.5 M urea (unfoldingof BSA by urea is believed to proceed through a two-statetransition process with the intermediate being formed at 4.5-5M concentration of urea52), the REES value is found to diminishto 5 nm, and keeping track of this, REES trims down to zero inthe presence of 8.0 M urea. Denaturation of BSA with increasingconcentration of urea results in greater exposure of the probemolecules to the aqueous buffer environment from the confinedenvironment inside the protein backbone, a direct consequenceof which is manifested through minimization of the REES effectwith gradual increase of urea concentration (i.e., the extent ofdenaturation). At a urea concentration as high as 8.0 M,complete denaturation of BSA takes place when the microen-vironment around the fluorophore is likely to be qualitativelycomparable with that in the bulk aqueous phase, as supportedby the absence of REES. This proposition is also supported bythe absence of REES in 80 µM BSA at 80 °C at which thermaldenaturation of BSA takes place, resulting in greater exposureof the probe molecules to aqueous buffer phase compared toits bound state in rigid protein surroundings.

3.8. Urea-Induced Protein Unfolding Studies. Steady-statefluorescence measurements that dictate the changes in thetertiary structure of proteins are complementary pathways toexplore the environmental stability of globular proteins.1,6 Theunfolding process of serum albumins with urea is quite wellstudied.6 Denaturation of BSA with urea takes place via a two-state transition through intermediate state (Int) at 4.5-5.0 Murea.52,53 After finding the binding interaction between the probeand BSA, we intended to see the effect of denaturation of proteinon its binding activity and on the overall photophysics ofDPDAME. Here, the urea-induced modifications to spectralcharacteristics of protein-bound DPDAME have been followedby steady-state fluorescence measurements. Figure 8a displaysthe changes in the emission spectra of BSA-bound DPDAMEwith increasing concentration of urea. As seen in Figure 8a,gradual addition of urea to a solution of DPDAME in 30 µMBSA results in a decrement of intensity of the CT emissionband with simultaneous shift of the emission maxima to thered; i.e., the pattern observed is quite reversed with respect tothat in Figure 2a. Addition of urea leads to weakening of theprobe-protein binding and the probe molecules are therebyexposed more to the bulk aqueous buffer phase compared toits bound condition in the native state of the protein. It isbelieved that urea displaces some of the water moleculesadjacent to the probe in the protein environment with theconsequent denaturation of the protein.54,55 The resultingdestabilization of the complex should be associated with agreater exposure of the probe to the aqueous buffer phasecompared to that in its bound state in the native conformationof the protein,54 and hence, the CT emission band intensitydecreases along with a red shift of the emission maxima. It isimportant to note that, at a substantial concentration of urea (9M), the emission wavelength (Figure 8b) and steady-stateanisotropy value (inset of Figure 8b) tend to correspond to thevalues in aqueous buffer solution. These observations are inline with the idea that the probe DPDAME binds to the protein

Figure 7. Effect of changing excitation wavelength on the emissionmaxima of DPDAME:BSA complex under different experimentalconditions.


BSA in its native conformation, and denaturation of the latterleads to a greater exposure of the probe from the proteinbackbone to the bulk aqueous buffer phase. Figure 8b depictsthe variation of emission maxima of the CT band withconcentration of urea. As seen in this figure, up to a ureaconcentration of 3 M, λem

max increases slowly and then the riseis quite sharp followed by another more or less steady changegetting off from a urea concentration of 7 M. This observationis consistent with reports available in the literature.3,6,46 However,we think it is ethical to point out here that the appearance ofemission spectra in Figure 8a exhibits some noticeable broaden-ing particularly toward the region of high concentration of urea.Appropriate spectral background correction negates the pos-sibility of interference from any unusual instrumental response(signal-to-noise ratio), while the use of low emission andexcitation slit widths (3.0 nm) and conservation of all otherexperimental conditions during the entire experiment confirmthe minimization of instrumental artifacts. Under such circum-stances, we presume that such noticeable broadening of theemission profile of DPDAME shows signs of extensivesolute-solvent interactions. The solution under experiment isquite complicated by composition containing three individualspecies, viz., BSA (30 µM), DPDAME (2 µM), and urea. Thus,it is not unlikely on the part of the probe to encounter complexand extensive solute-solvent interaction (contributing to spectralbroadening31) with denaturation of the protein. In fact, the extentof broadening is found to be more prominent with the progressof the process of denaturation with increasing denaturantconcentration.

It seems crucial at this stage to make an attempt to decipherwhether the probe is completely free to move or is in asomewhat restricted environment in the denatured state of theprotein. For this purpose, a meticulous perusal of the steady-state fluorescence anisotropy values has been undertaken andit appears that, despite the lowering of the anisotropy of theprotein-bound probe with increasing urea concentration (insetof Figure 8b), the anisotropy value at a reasonable ureaconcentration remains still somewhat higher than the value inthe absence of the protein: anisotropy, ranis ≈ 0.013 for the probein buffer (Figure 4a) vs ranis ≈ 0.125 for the probe in thepresence of 9.5 M urea (inset of Figure 8b).

Also, we have monitored the effect of increasing concentra-tion of the probe on the intrinsic fluorescence properties of BSAunder denatured conditions (30 µM BSA + 6 M urea) to followwhether the probe can induce any significant perturbation tothe intrinsic fluorescence properties of the protein. The conceptbehind the experiment was as follows. If the probe bindsstrongly to BSA in its native state and is released upon uncoilingof the protein, the external addition of the probe to denaturedprotein should have entailed no significant modulation to theintrinsic fluorescence of BSA. However, the fruitfulness of theexperiment was little crumpled by the observation reported inFigure 6a and section 3.6; i.e., an appreciable overlap betweentryptophanyl emission and absorption spectra of DPDAME ledto the operation of FRET with consequent quenching of intrinsic(tryptophanyl) fluorescence intensity of BSA. However, anoticeable reduction in energy transfer efficiency (E) to 64.2%in the denatured state (30 µM BSA + 6 M urea) of BSA (ca.E ) 86.96% in the native state of BSA) well substantiates theproposition of binding of the probe to the native conformationof the protein and its subsequent greater extent of exposuretoward the aqueous buffer phase upon denaturation of the same,whereby rendering the donor (Trp-212)-acceptor (DPDAME)distance (r) to increase as reflected in sizable depletion of FRETefficiency.48 However, with a view to fathom deeper into theresults, we have calculated the donor (Trp-212)-acceptor(DPDAME) distance in the presence of 6 M urea (see also theSupporting Information) and the result comes out to be r )1.656 ( 0.13 nm, which is not too large with respect to thevalue obtained in the case of native BSA and is thus harmonizingwith the anisotropy data quite satisfactorily, as mentioned above.Also, these data appear to corroborate with our discussions insection 3.5.

3.9. Structural Stability of BSA toward Chemically In-duced Denaturation by Urea upon Interaction withDPDAME. While carrying out a study on the application ofthe simple charge transfer probe DPDAME to study themicroenvironments of proteinous media, it is pertinent toestablish whether binding of the probe to the protein leads tostabilization or destabilization of the latter. The effect of bindingof DPDAME on the stability of BSA has been investigated byurea-induced unfolding of the protein by monitoring the steady-state fluorescence. For this purpose, we have excited BSA at280 nm (λabs

max of tryptophan residue in BSA) in the presenceand absence of the probe DPDAME and recorded the fluores-cence of tryptophan residue at 350 nm in the presence of varyingconcentrations of urea. Tryptophanyl fluorescence of BSA isfound to gradually diminish due to the denaturing action of urea.A transition curve is constructed by plotting I/I0 against theconcentration of urea, where I and I0 are the fluorescenceintensities, respectively, in the presence and absence of urea.The transition curves appear sigmoidal (Figure 9a). The valuesof urea concentration required for half completion of the

Figure 8. Chemically (urea) induced denaturation of BSA as sensedby the polarity-sensitive ICT emission of DPDAME. (a) Representativespectra showing the effect of increasing concentration of urea (curves1f 8 correspond to [urea] (in M) ) 0, 1.0, 2.0, 3.0, 4.0, 5.0, 6.0, 7.0)on the fluorescence of BSA-bound DPDAME (λex ) 370 nm). (b) Plotof emission maxima of DPDAME:BSA complex as a function of ureaconcentration (0 f 9.5 M). Inset shows the decrease in the steady-state fluorescence anisotropy of BSA-bound probe against increasingurea concentration (each data point is an average of 10 individualmeasurements).


transition (i.e., when one-half of the native state of protein isdenatured) are determined from the midpoints of the transitioncurves, and the values are found to be 3.48 and 4.15 M for thebare protein BSA and BSA in the presence of DPDAME,respectively. The requirement of a greater amount of denaturant(urea) in the presence of DPDAME justifies significant stabiliza-tion of BSA in the presence of the probe, with respect todenaturation by urea (it is not a concern over the absolutestability of the protein, however). These observations are in trackwith previous reports available in the literature.3

The influence of binding of the probe on the secondary structureof the protein has been ascertained through far UV circulardichroism studies. The CD spectra for BSA (Figure 9b) monitoredin the range 250-200 nm reveal the presence of two bands at ∼209and ∼222 nm, as consistent with the literature.3b,31,56 However,Figure 9b also reveals that, for the BSA:DPDAME compositesystem, the appearance of the CD spectra is exactly similar to thatof BSA alone, only differing in a slight decrement of ellipticity.Such very similar (almost superimposed) CD spectra of the proteinin the presence and absence of the probe evidence no detectablestructural change of the protein upon binding with the probe, asconsistent with literature reports.3,7b

3.10. Thermal Denaturation of BSA: ICT Emission ofDPDAME as an Extrinsic Probe. Unfolding of proteins isusually marked by a change in the secondary and globularstructure of the protein. Just like chemical denaturation of BSA,unfolding of a protein can be induced by rise of temperature.Here, we have followed the unfolding of BSA imparted by riseof temperature through changes in the spectral characteristicsof the probe when bound to BSA. Figure 10 represents theemission spectra of DPDAME in the presence of 80 µM BSA(this concentration of BSA ensures saturation of bindinginteraction between BSA and the probe) at different tempera-tures. As evident from the figure, rise of temperature inducesnoticeable diminution in the CT emission intensity of DPDAMEtogether with a red shift of the emission maxima (inset of Figure

10). The unfolding of the protein BSA with rise of temperatureis necessarily to induce a greater exposure of the probemolecules to the bulk aqueous phase compared to that in itsBSA-bound state in the native conformation of the protein. Thus,the study of thermal denaturation of BSA adds another supportto the inference that DPDAME prefers to bind to BSA in itsnative conformation and, due to its neutral and hydrophobicnature, DPDAME is favorably solubilized in the hydrophobicpocket of BSA.

3.11. Variation of Fluorescence Emission with Changesin Hydrophobicity of the Microenvironment of DPDAME.Figure 11a shows that the CT band maxima of DPDAME isblue-shifted from ∼540 nm in bulk aqueous buffer phase to∼528 nm in 20 mM SDS. In the presence of 60 µM BSA, theblue shift is even higher, to ∼510 nm. This observation pointsto the fact that the polarity in SDS micelle is much higher

Figure 9. (a) Plot of the relative fluorescence (I/I0) of tryptophan ofBSA against the concentration of urea in the absence and presence ofDPDAME (λex ) 280 nm, λem ) 350 nm). (b) Far UV CD spectraof BSA in the absence and presence of DPDAME ([BSA] ) 1.25 µMand [DPDAME] ) 0.25 µM).

Figure 10. Effect of increasing temperature on the emission spectralprofile of the DPDAME-BSA complex ([BSA] ) 80 µM; λex ) 370nm). Curves 1 f 8 correspond to T ) 273, 283, 293, 303, 313, 323,333, and 353 K.

Figure 11. (a) Normalized emission spectra of DPDAME in water,60 µM BSA, and 20 mM SDS medium. (b) Plot of fluorescenceintensity of DPDAME at λem ) 528 nm as a function of SDSconcentration in aqueous medium.


compared to that in the proteinous media. However, thepolarities of binding hydrophobic regions of both SDS and BSAare much less than that of pure water (buffer). The micropolarityof the surroundings of the probe in terms of ET(30) in 20 mMSDS is determined to be 54.7 (Figure 3b), which is much higherthan that of BSA (ET(30) ) 48.4). Here, the concentration ofthe probe was kept the same in all media. The emission intensityof the blue-shifted band of the probe in SDS as well as in BSAis found to increase manifold compared to that in aqueous buffersolution. As already mentioned earlier, depletion of the radia-tionless decay routes due to restrictions imposed on the mobilityof the probe molecules contributes to such intensity enhancement.

In fact, the CT emission intensity of DPDAME when plottedas a function of SDS concentration (Figure 11b) allows one todetermine the critical micellar concentration (CMC) of SDS (7.10mM which is in excellent agreement with the literature value54,55).This advocates for the efficiency of DPDAME for probingmicroheterogeneous environments of micellar environments.

3.12. Surfactant (SDS)-Induced Unfolding of BSA. Severalstudies have been reported by different spectroscopic techniqueson the binding of BSA with small molecules, particularly fattyacids and surfactants.1,9,57-61,64 Binding of these molecules to aglobular protein leads to alterations in the intramolecular forcesresponsible for maintaining the secondary structure of the proteinand thereby producing conformational changes.62 In fact,characterization of these changes at atomic resolution has alsobeen possible.61 Here, we discuss the binding of the anionicsurfactant SDS with the globular protein BSA based on thefluorescence approach. The binding of SDS with BSA is knownto display four characteristic regions with increasing concentra-tion of SDS: (A) specific binding region; (B) noncooperativebinding region; (C) co-operative binding region; (D) saturationbinding region.9,62 Addition of SDS results in decrement of theintrinsic fluorescence intensity of Trp-212 of BSA due to greaterexposure of the tryptophan moiety to the polar environment,resulting from SDS-induced uncoiling of BSA. The bindingisotherms for SDS-BSA interaction can be obtained from aplot of R vs total SDS concentration (Figure 12a) whichdistinctly produces the four characteristic zones, viz., A, B, C,and D. The term R, i.e., the fraction of a protein bound tosurfactant, is defined as

where Iobs, Ifree, and Imin are, respectively, the observed emissionintensity values of Trp-212 at any concentration of SDS, in theabsence of SDS, and under saturation conditions.63 As is evidentfrom the figure, an initial fast rising region A (ranging from[SDS] ≈ 0.0 to 0.2 mM) is followed by a slow rising zone Bup to ∼0.5 mM of SDS. Here, region A represents specificbinding of SDS to some high energy binding sites of BSA, whileregion B is representative of the noncooperative binding wherethe rise is quite slow or almost flat. The next zone, i.e., zone C,is the most important one which observes a steep rise in R valueup to ∼7 mM SDS representing massive co-operative bindingof SDS, resulting in uncoiling of the BSA native conformation.After region C, saturation binding comes into play (region D)where no significant change in R occurs, indicating no furtherbinding.9

We have attempted to explore this binding phenomenonspectroscopically utilizing DPDAME as an extrinsic reporter.As seen from Figure 12b, the intensity of the CT band of

DPDAME solubilized in 30 µM BSA exhibits specific variationupon addition of SDS, reflecting the binding isotherm forSDS-BSA interaction. The CT band shows a red shift (from∼506 to ∼520 nm) upon increasing concentration of SDS,which is related to the exposure of the probe to the bulk aqueousbuffer phase due to uncoiling of BSA in the presence of SDS.The sudden reduction of CT emission intensity at ∼0.1 mMSDS is probably due to expulsion of the probe molecules onaddition of SDS. In the region of specific binding A (rangingfrom [SDS] ≈ 0.0 to 0.2 mM), initial uncoiling of BSA takesplace and increase of the number of exposed binding sites leadsto increment of the intensity of the CT band due to competitivebinding of some of the expelled probe molecules to thehydrophobic binding sites of BSA. This region is followed bya small rising region B (ranging from [SDS] ≈ 0.2 to 0.5 mM)representing the noncooperative binding of SDS to BSA. Themost important zone of cooperative binding, i.e., zone C, showsits onset in the SDS concentration range 0.6-0.8 mM. In thiszone, maximum uncoiling or denaturation of BSA makes thedeeply buried probe molecules inside the protein hydrophobiccavity expose more to the aqueous buffer phase, leading todecrease of the intensity of CT fluorescence. This region is foundto move up to around 8-10 mM of total [SDS], beyond whichfurther change in intensity is minimized, reflecting saturationof binding past this concentration of SDS and the probemolecules being hydrophobic remains solubilized in SDSmicelles (CMC of SDS ) 7.1 mM as determined usingDPDAME as the probe).54 In fact, solubilization of the probemolecules in SDS micelles is also supported by the final positionof the emission maxima, ∼528 nm as against ∼540 nm in pureaqueous buffer. The stability of the native globular conforma-tions of proteins is, indeed, an outcome of a delicate balance

R )Iobs - Ifree

Imin - Ifree(7)

Figure 12. (a) Binding isotherm for protein-surfactant (BSA-SDS)interaction presented in the form of R vs [SDS] curve. (b) Variation ofemission intensity at 508 nm of the DPDAME-BSA complex withincreasing concentration of SDS ([BSA] ) 30 µM, λex ) 370 nm, λem

) 508 nm).


between various interactions and can be affected by severalfactors of which the impact of surfactants on the stability ofproteins has long been a subject of much extensive and criticalstudy. Herein, the excellent harmony of our results with availableliterature reports9,54 advocates for the commendable efficiencyof the polarity-sensitive ICT emission of DPDAME in constru-ing an important phenomenon, protein-surfactant interactions.

3.13. Refolding of Urea Denatured BSA with SDS. Thepresence of SDS results in unfolding of the native conformationof the protein since the gain in free energy due to binding ofSDS to the hydrophobic sites of BSA can surpass the loss infree energy due to loss of the native conformation.3-7,9,31,45

Hence, the destructive action of SDS occurs. However, SDSplays an interesting dual role as a destabilizer for the nativeconformation of BSA and a stabilizer for urea-denatured BSA.It is well-known that some of the helices in BSA are disruptedin the presence of SDS (which binds to hydrophobic sites ofBSA) but most of them are drastically destroyed in the presenceof urea (whose denaturing action is based on breaking of waterstructures responsible for stabilization of the native form ofBSA). Thus, interestingly, the coexistence of SDS and urea wasfound to protect the protein conformation and this phenomenonwas first observed by Duggan and Luck65 as rise in the viscosityof BSA-urea solution was prevented upon addition of certainorganic anions. Later on, the problem has been addressed byseveral groups.57-59,66 Disulfide bridges play quite an importantrole in stabilizing the native conformation of BSA, the reductionof which accompanies a huge disruption of the helices. Thepresence of urea results in denaturation of BSA when the probemolecules are more exposed to the bulk aqueous phase whichis reflected through the red shift of the polarity-sensitive ICTemission of the probe and thereby making way for followingthe protective action of SDS on urea-denatured BSA bymonitoring the shift of the CT band upon addition of SDS inBSA-urea solution. In the range of 4-6 M urea, addition ofSDS is found to result in an initial blue shift of the CT bandmaxima of DPDAME, indicating recoiling of urea-denaturedBSA, but further increment of SDS concentration accompaniesred shift of the CT band, reflecting greater exposure of the probeto the bulk aqueous phase; i.e., further addition of SDS inducesuncoiling of BSA (Figure 13). Thus, the ICT probe DPDAMEcan successfully be utilized in probing the conformationalchanges induced upon addition of SDS to urea-denatured BSAsolution.

3.14. Binding Site of the Probe. The efficiency of DPDAMEto function as a molecular reporter for probing proteinous (BSA)

and micellar (SDS) microheterogeneous environments has beendemonstrated in the foregoing sections. Now, it is pertinent atthis point to explore the binding site of the probe in the proteinbackbone for understanding the efficacy of the probe as amolecular reporter to furnish information about the microen-vironment of a protein molecule. In order to assess the bindingsite of DPDAME in BSA, we have intertwined the results ofthe urea-induced denaturation study, micropolarity measure-ments, and the fluorescence resonance energy transfer (FRET)study. The micropolarity around the probe has been determinedin different states of the protein, i.e., native (N), intermediate(Int), and unfolded (U) states. Literature reports reveal that theintermediate state (in the presence of 4.5-5.0 M urea52) ischaracterized by the unfolding of domain III along with a partialloss of the native form of domain I.1,31,56,66-68 The unfolded state(U) is, however, characterized mainly by unfolding of domainII.1,31,67,68 The measured micropolarity values around the probeat different states of the protein are summarized in Table 3 whichreveals that the Int to U transition of BSA (involving domainII) entails a marked difference of micropolarity (∆ET(30) (kcal/mol) ) 7.18) as against that in the case of N to Int transition(involving domains I and III) for which ∆ET(30) (kcal/mol) isonly 2.92. This result implies a greater possibility for DPDAMEto reside in domain II relative to domain I or III. In addition,the principal hydrophobic binding sites of BSA are located indomains II and III, while domain I, characterized by a strongnegative charge, can serve as a better binding site for cationicprobes.2,3b With an eye to the neutral and hydrophobic natureof the probe in our case, it is reasonable to exclude domain I asone of its probable binding sites.

The occurrence of FRET with a very high energy transferefficiency (E ) 86.96%; section 3.6) strongly suggests thelocation of the probe to be in near vicinity of the tryptophanresidue in BSA. However, predictions are quite difficult here,since the situation is pretty complicated by the presence of twotryptophan residues in BSA. From the micropolarity measure-ments in different states of the protein, it can be inferred thatthe most probable binding location of the probe in BSA isdomain II. On top of this, Trp-134 is located in domain IB(hydrophilic region) which has been excluded as a probablebinding site of the probe and domain III (hydrophobic region)contains no tryptophan residue which infers that the locationof the probe in domain III would have led to inoperativenessof FRET from Trp to DPDAME as contrary to the presentfindings.

Our assessment receives additional support on a theoreticalmilieu which reveals that out of various possibilities the stableinteracting conformation is the one having DPDAME in nearvicinity of Trp-212 (Figure 14; computations done accordingto the simulation protocol described in section 2.1). Themagnified segment of Figure 14 illustrates the location of theprobe and Trp-212 on a background of electrostatic potentialsurface on the protein (as generated using PyMOL software31),revealing that the aromatic nuclei of both DPDAME and Trp-212 are located on a hydrophobic region (white surface) whichfurther reinforces our previous assignments.

3.15. Time-Resolved Measurement. Because of its inherentsensitivity toward excited state interactions, fluorescence lifetimemeasurements have long been recognized as a sensitive indicatorof the local environment of a fluorophore, apart from beingapplied in monitoring sensitive issues like differential degreesof solvent relaxation around a fluorophore, the presence of morethan one chemical entity in a solution, and so forth.3,6,7,17-21,31,46

Time-resolved studies of the intrinsic fluorophore tryptophan

Figure 13. Plot of emission maxima of DPDAME in 30 µM BSAand (a) 4 M urea, (b) 5 M urea, and (c) 6 M urea in Tris-HCl bufferof pH 7.4 vs total SDS concentration in mM.


are complex, and nothing concrete can be concluded from thefluorescence decays. This is because most proteins contain morethan one tryptophan which lies in different environments and,second, protein folding and unfolding are very complex andfast processes and collection of suitable data ranges in hours.Under such circumstances, the fluorescence lifetimes of probesbound to proteins throw light on the microenvironment sur-rounding the probe molecule in its excited state and a greatdeal can be inferred about the nature of protein-probe bindingas well as conformational changes of proteins under variouscircumstances. The fluorescence decay curves for such bindingprocesses in heterogeneous media are generally multiexponen-tial. For DPDAME bound to BSA, the fluorescence decay curveswere best fitted to a biexponential function with acceptablevalues of �2 (Table 2) and exhibit an increase in the lifetime ofDPDAME with increasing BSA concentration (Figure 15 andTable 2). In analogy to other such studies,3,6,7,17-21,31 the increasein lifetime is inferred to be due to depletion in nonradiativedecay channels as a result of encapsulation of DPDAME in ahydrophobic cavity of the protein.

In the present case, however, we prefer to avoid placing toomuch emphasis on the magnitude of the individual decayconstants of the multiexponential decays; rather, the mean(average) fluorescence lifetimes (as defined by eq 4) areemployed as an important parameter for exploiting the behaviorof DPDAME when bound to the protein. (It is, indeed, not easyand desired to specifically assign the two components obtainedin time-resolved fluorescence decay patterns of DPDAME underthe present experimental conditions, since the situation is pretty

complicated with the extrinsic fluorescence probe bound to theprotein backbone.3,6,7,31,46,54 However, with a view to the normalobservations with ICT chromophores, the two decay componentscan be argued to be attributable to two excited state species,namely, the LE and CT species.12,69 In fact, a reasonably steadyconsistency in the relative abundances of the two species tendto corroborate with our steady-state spectral findings in the sensethat the more abundant species conform to the higher intensityof the longer wavelength species (ICT) while, conversely, theless populated one conforms to the lower intensity of the shorterwavelength species (LE).12,69 These results are further substanti-ated by the theoretical calculations of the potential energysurface across the ICT reaction coordinate for DPDAME.30

However, the environment is enormously modified in thepresence of the protein BSA and solvent reorganization can alsoput a signature in the observed decay patterns,69 which is whythe mean fluorescence lifetime has been emphasized in furtherdiscussion.) The time-resolved emission decay profiles ofDPDAME in pure aqueous phase and in the presence of twodifferent concentrations of BSA are presented in Figure 15, andthe relevant parameters are summarized in Table 2. From thevalues of mean fluorescence lifetime (⟨τf⟩) and quantum yield(Φf) of DPDAME in different environments, the radiative andnonradiative rate constants have been calculated using thefollowing equations:31

The calculated values are tabulated in Table 2. Scrutiny of thedata in Table 2 evidences that in protein environment thenonradiative decay constants knr are reasonably reduced fromthose in aqueous buffer medium so that the enhanced lifetime

Figure 14. Final conformation obtained after energy minimization followed by equilibration of the DPDAME-BSA composite system. The picturehas been prepared using PyMOL software.32b

TABLE 2: Fluorescence Lifetimes of DPDAME with Increasing Concentration of BSA

environment a1 a2 τ1 (ns) τ2 (ns) �2 ⟨τf⟩ (ns) quantum yield (Φf) kr (s-1) × 10-9 knr (s-1) × 10-9

Aq. buffer 0.934 0.065 0.508 1.696 0.98 0.732 0.034 0.046 1.32BSA (8 µM) 0.916 0.839 0.564 2.159 1.04 1.805 0.121 0.067 0.487BSA (60 µM) 0.681 0.318 1.385 3.835 1.14 2.766 0.243 0.088 0.274

TABLE 3: Micropolarity Values in Terms of ET(30)(kcal/mol) at Different States of BSA

different states of protein ET(30)

native (N) 48.44intermediate (Int) 51.36unfolded (U) 58.54

kr ) Φf/⟨τf⟩ (8)

1/⟨τf⟩ ) kr + knr (9)


of the fluorophore in protein environment seems attributable tothe diminution of radiationless decay.

Fluorescence anisotropy is a property that depends onrotational diffusion of the fluorophore as well as the fluorescencelifetime.31 Thus, it is necessary to ensure that the observedchange in steady-state fluorescence anisotropy of DPDAME inthe presence of BSA (section 3.4) is not an outcome of anychange in fluorescence lifetime. For this purpose, the averagerotational correlation times have been calculated using Perrin’sequation31 for DPDAME in BSA. Perrin’s equation is given as

in which r0, r, and ⟨τf⟩ are the fundamental anisotropy, steady-state anisotropy, and mean fluorescence lifetime of DPDAME,respectively. Although ideally Perrin’s equation is not applicablein a microheterogeneous environment, it can be used to a gooddegree of approximation considering the mean fluorescencelifetime of the system.31,46a Now, taking r0 ) 0.383,31,46a andusing eq 10, we have calculated τc for DPDAME in the proteinenvironments and it is seen to be appreciably enhanced uponbinding to the protein (from τc ) 0.026 ns in aqueous buffer toτc ) 4.297 ns in 8 µM BSA to τc ) 11.082 ns in 60 µM BSA).Significant increase of rotational correlation time in BSAenvironments dictates that the observed change in steady-stateanisotropy of the probe (section 3.4 and Figure 4) is not a resultof any lifetime-induced phenomena and thereby reinforces ourearlier assignment that binding to the protein imparts anenhanced rotational restriction on the probe.

4. Summary and Conclusion

In the present work, the binding phenomenon of our simplecharge transfer fluorescence probe DPDAME with BSA is

presented using spectroscopic techniques. The photophysics ofthe ICT probe is remarkably modified upon binding with thetransport protein BSA, as compared to those in pure aqueousphase. Modulations of ICT photophysics of DPDAME in aprotein cavity have been shaped into a tool to explore thehydrophobic subdomain IIA of BSA through determination ofbinding efficiency, the nature of the microenvironment aroundthe probe, and the micropolarity and microviscosity at thebinding site. Steady-state anisotropy and REES measurementshave been exploited to complement the efficient binding of theprobe into the protein cavity. DPDAME is also shown to be apotent molecular reporter for probing chemical and thermaldenaturation of the protein. The specific binding phenomenonof SDS with BSA has also been adroitly followed by exploringthe polarity-sensitive ICT emission of the probe. The renaturingaction of low concentrations of SDS on urea-denatured proteinis demonstrated simply through variation of emission maximaof DPDAME. A conjugate analysis of urea denaturation study,FRET study, and micropolarity measurements leads to assessingthe probable binding site of DPDAME inside the proteinbackbone to be subdomain IIA, which is further reinforced bya theoretical simulation. Urea denaturation study also reveals asubstantial enhancement of stability of the protein as a resultof binding with the probe. Overall, we have demonstratedDPDAME to be an efficient molecular reporter for probingproteineous and micellar microheterogeneous environments andalso the action of urea, SDS, and temperature on the stabilityof the protein are well reflected in the course of modificationsof the photophysics of the probe.

Acknowledgment. N.G. acknowledges DST, India (ProjectNo.SR/S1/PC/26/2008),andCSIR, India (ProjectNo.01(2161)07/EMR-II), for financial support. B.K.P. and A.S. gratefullyacknowledge CSIR, New Delhi, India, for research fellowships.The authors convey their special thanks to Mr. Atanu Das andProfessor Dr. Chaitali Mukhopadhyay of the Department ofChemistry of our university regarding the theoretical simulationand their continuous encouragement throughout the work. Wegreatly appreciate the cooperation received from Ms. DeboleenaSarkar and Professor Dr. Nitin Chattopadhyay of JadavpurUniversity, Calcutta, India, for fluorescence lifetime and ani-sotropy measurements. Dr. Gautam Basu of Bose Institute,Calcutta, India, is gratefully acknowledged for allowing us torecord the CD spectra. We sincerely thank the reviewers fortheir meticulous inspection of our manuscript and constructivesuggestions.

Supporting Information Available: Information on theBenesi-Hildebrand equation and fluorescence resonance energytransfer. This material is available free of charge via the Internetat http://pubs.acs.org.

References and Notes

(1) (a) He, X. M.; Carter, D. C. Nature 1992, 358, 209–215. (b)Chakraborty, T.; Chakraborty, I.; Moulik, S. P.; Ghosh, S. Langmuir 2009,25, 3062–3074.

(2) Peters, T. Serum albumin. AdVances in Protein Chemistry; Aca-demic Press: New York, 1985; Vol. 37, pp 161-245.

(3) (a) Abou-Zied, O. K.; Al-Shihi, O. I. K. J. Am. Chem. Soc. 2008,130, 10793–10801. (b) Chakrabarty, A.; Mallick, A.; Haldar, B.; Das, P.;Chattopadhyay, N. Biomacromolecules 2007, 8, 920–927. (c) Abou-Zied,O. K. J. Phys. Chem. B 2007, 111, 9879–9885.

(4) Brown, J. R. Albumin Structure, Function and Uses; Rosenoer,V. M., Oratz, M., Rothschild, M. A., Eds.; Pergamon Press: Oxford, U.K.,1977; Vol. 27.

(5) (a) Carter, D. C.; Ho, J. X. AdV. Protein Chem. 1994, 45, 153–159. (b) He, X. M.; Carter, D. C. Nature 1996, 358, 209–216.

Figure 15. Typical time-resolved fluorescence decay profiles ofDPDAME with increasing concentration of BSA (λex ) 370 nm,λmonitored ) λem

max).

τc )⟨τf⟩r

r0 - r(10)


(6) (a) Mahanta, S.; Singh, R. B.; Guchhait, N. J. Fluoresc. 2009, 19,291–302, and references therein. (b) Singh, R. B.; Mahanta, S.; Bagchi,A.; Guchhait, N. Photochem. Photobiol. Sci. 2009, 8, 101–110, andreferences therein. (c) Singh, R. B.; Mahanta, S.; Guchhait, N. Spectrochim.Acta, Part A 2009, 72, 1103–1111, and references therein. (d) Wu, F.-Y.;Ji, Z.-J.; Wu, Y.-M.; Wan, X.-F. Chem. Phys. Lett. 2006, 424, 387–393,and references therein.

(7) (a) Das, R.; Mitra, S.; Nath, D.; Mukherjee, S. J. Phys. Chem. 1996,100, 14514–14519. (b) Banerjee, P.; Pramanik, S.; Sarkar, A.; Bhattacharya,S. C. J. Phys. Chem. B 2009, 113, 11429–11436.

(8) Zhong, D.; Douhal, A.; Zewail, A. H. Proc. Natl. Acad. Sci. U.S.A.2000, 97, 14056–14061.

(9) (a) Gelamo, E. L.; Silva, C. H.; Imasato, H.; Tabak, M. Biochim.Biophys. Acta 2002, 1594, 84–99. (b) Gelamo, E. L.; Tabak, M. Biochim.Biophys. Acta 2000, 56, 2255–2271.

(10) Lippert, E.; Luder, W.; Boss, H. In AdVances in MolecularSpectroscopy; Mangini, A., Ed.; Pergamon Press: Oxford, U.K., 1962; p443.

(11) Ishikawa, H.; Sugiyama, M.; Baba, I.; Setaka, W.; Kira, M.;Mikami, N. J. Phys. Chem. A 2005, 109, 8959–8961.

(12) Grabowski, Z.; Rotkiewicz, K.; Rettig, W. Chem. ReV. 2003, 103,3899–4032.

(13) Sobolewski, A. L.; Domcke, W. J. Photochem. Photobiol., A 1997,105, 325–328.

(14) Murali, S.; Rettig, W. J. Phys. Chem. A 2006, 110, 28–37.(15) Rotkiewicz, K.; Grellmann, K. H.; Grabowski, Z. R. Chem. Phys.

Lett. 1973, 19, 315–318.(16) Sumalekshmy, S.; Gopidas, K. R. J. Phys. Chem. B 2004, 108,

3705–3712.(17) Bangal, P. R.; Panja, S.; Chakravorti, S. J. Photochem. Photobiol.,

A 2001, 139, 5–16.(18) Singh, R. B.; Mahanta, S.; Kar, S.; Guchhait, N. Chem. Phys. 2007,

342, 33–42.(19) Chakraborty, A.; Kar, S.; Nath, D. N.; Guchhait, N. J. Chem. Sci.

2007, 119, 195–204.(20) Chakraborty, A.; Kar, S.; Guchhait, N. Chem. Phys. 2006, 320,

75–83.(21) Chakraborty, A.; Kar, S.; Nath, D. N.; Guchhait, N. J. Phys. Chem.

A 2006, 110, 12089–12095.(22) Rettig, W. J. Phys. Chem. 1982, 86, 1970–1976.(23) Zachariasse, K. A.; Druzhinin, S. I.; Bosch, W.; Machinek, R. J. Am.

Chem. Soc. 2004, 126, 1705–1715.(24) Yang, J.-S.; Liau, K.-L.; Wang, C.-M.; Hwang, C.-Y. J. Am. Chem.

Soc. 2004, 126, 12325–12335.(25) Hazra, P.; Chakrabarty, D.; Sarkar, N. Langmuir 2002, 18, 7872–

7879.(26) Kim, Y. H.; Cho, D. W.; Yoon, M.; Kim, D. J. Phys. Chem. 1996,

100, 15670–15676.(27) Jodicke, C. J.; Luthi, H. P. J. Chem. Phys. 2002, 117, 4146–4156.(28) Jodicke, C. J.; Luthi, H. P. J. Chem. Phys. 2003, 119, 12852–12865.(29) Ishikawa, H.; Shimanuki, Y.; Sugiyama, M.; Tajima, Y.; Kira, M.;

Mikami, N. J. Am. Chem. Soc. 2002, 124, 6220–6230.(30) Paul, B. K.; Samanta, A.; Kar, S.; Guchhait, N. J. Lumin. DOI:

10.1016/j.jlumin.2010.02.036.(31) Lakowicz; J. R. Principles of fluorescence spectroscopy; Plenum:

New York, 1999.(32) (a) Brooks, B. R.; Bruccoleri, R. E.; Olafson, B. D.; States, D. J.;

Swaminathan, S.; Karplus, M. J. Comput. Chem. 1983, 4, 187–217. (b) DeLano, W. L. The PyMOL Molecular Graphics System; De Lano Scientific:San Carlos, CA, 2002.

(33) (a) Modukuru, N. K.; Snow, K. J.; Perrin, B. S.; Kumar, C. V. J.Phys. Chem. B 2005, 109, 11810–11818. (b) Chuan, D.; Yu-xia, W.; Yan-li, W. J. Photochem. Photobiol., A 2005, 174, 15–22.

(34) Benesi, H. A.; Hildebrand, J. H. J. Am. Chem. Soc. 1949, 71, 2703–2707.

(35) Charaborty, B.; Basu, S. J. Lumin. 2009, 129, 34–39.(36) Krishnakumar, S. S.; Panda, D. Biochemistry 2002, 41, 7443–7452.(37) Guo, M.; Zou, J.-W.; Yi, P.-G.; Shang, Z.-C.; Hu, G.-H.; Yu, Q.-

S. Anal. Sci. 2004, 20, 465–470.(38) Mukherjee, S.; Chattopadhyay, A. J. Fluoresc. 1995, 5, 237–246.(39) Ahmad, B.; Khan, M. K. A.; Haq, S. K.; Khan, R. H. Biophys.

Biochem. Res. Commun. 2004, 314, 166–173.

(40) Gonzalez-Jimenez, J.; Cortijo, M. J. Protein Chem. 2002, 21, 75–79.

(41) Wallevik, K. J. Biol. Chem. 1973, 248, 2650–2655.(42) Macgregor, R. B.; Weber, G. Nature 1986, 319, 70–73.(43) Bismuto, E.; Jameson, D. M.; Gratton, E. J. Am. Chem. Soc. 1987,

109, 2354–2357.(44) Kosower, E. M.; Kantey, H. J. Am. Chem. Soc. 1983, 105, 6236–

6243.(45) (a) Richardt, C. In Molecular Interaction; Ratajazak, H., Orville-

Thomas, W. J., Eds.; Wiley: New York, 1982; Vol. 3, p 255. (b) Muzammil,S.; Kumar, Y.; Tayyab, S. Proteins 2000, 40, 29–38.

(46) (a) Mallick, A.; Haldar, B.; Maiti, S.; Bera, S. C.; Chattopadhyay,N. J. Phys. Chem. B 2005, 109, 14675–14682. (b) Sengupta, B.; Sengupta,P. K. Biochim. Biophys. Res. Commun. 2002, 299, 400–403.

(47) Draxler, S.; Lippitsch, M. E. J. Phys. Chem. 1993, 97, 11493–11496.

(48) (a) Seth, D.; Chakrabarty, D.; Chakraborty, A.; Sarkar, N. Chem.Phys. Lett. 2005, 401, 546–552. (b) Sahu, K.; Ghosh, S.; Mondal, S. K.;Ghosh, B. C.; Sen, P.; Roy, D.; Bhattacharyya, K. J. Chem. Phys. 2006,125, 44714–44718. (c) Barbero, N.; Barni, E.; Barolo, C.; Quagliotto, P.;Viscardi, G.; Napione, L.; Pavan, S.; Bussolino, F. Dyes Pigm. 2009, 80,307–313. (d) Selvin, P. R. Nat. Struct. Biol. 2000, 7, 730–734. (e) Sardar,P. S.; Samanta, S.; Maity, S. S.; Dasgupta, S.; Ghosh, S. J. Phys. Chem. B2008, 112, 3451–3461. (f) Ghosh, S.; Dey, S.; Adhikari, A.; Mandal, U.;Bhattacharyya, K. J. Phys. Chem. B 2007, 111, 7085–7091. (g) Adhikari,A.; Das, D. K.; Sasmal, D. K.; Bhattacharyya, K. J. Phys. Chem. A 2009,113, 3737–3743.

(49) Mattjus, P.; Molotkovsky, J. G.; Smaby, J. M.; Brown, R. E. Anal.Biochem. 1999, 268, 297–304.

(50) Demchenko, A. P. Luminescence 2002, 17, 19–42.(51) (a) Mukherjee, S.; Chattopadhyay, A. Langmuir 2005, 21, 287–

293. (b) Samanta, A. J. Phys. Chem. B 2006, 110, 13704–13716, andreferences therein. (c) Adhikari, R. M.; Neckers, D. C. J. Phys. Chem. A2009, 113, 417–422.

(52) (a) Tayyab, S.; Ahmad, B.; Kumar, Y.; Khan, M. M. Int. J. Biol.Macromol. 2002, 30, 17–22. (b) Sulkowska, A.; Bojko, B.; Rownicka, J.;Pentak, D.; Sulkowska, W. J. Mol. Struct. 2003, 651, 237–243.

(53) Santra, M. K.; Banerjee, A.; Rahaman, O.; Panda, D. Int. J. Biol.Macromol. 2005, 37, 200–204.

(54) (a) Singh, R. B.; Mahanta, S.; Guchhait, N. Chem. Phys. Lett. 2008,463, 183–188, and references therein. (b) Jones, M. N. In BiochemicalThermodynamics, 1st ed.; Jones, M. N., Ed.; Elsevier: Amsterdam, TheNetherlands, 1985; Chapter 5, p 112. (c) Jones, M. N.; Brass, A. In FoodPolymers, Gels and Colloids; Dickinson, E, Ed.; Royal Society of Chemistry:Cambridge, 1991; pp 65-80. (d) D’alfonso, L.; Collini, M.; Baldini, G.Biochemistry 2002, 41, 326–333. (e) Campbell, I. D.; Dwek, R. A. BiologicalSpectroscopy; Benjamin/Cummings: Menlo Park, CA, 1984.

(55) Belletete, M.; Lachapelle, M.; Durocher, G. J. Phys. Chem. 1990,94, 7642–7648.

(56) Ahmad, B.; Parveen, S.; Khan, R. H. Biomacromolecules 2006, 7,1350–1356.

(57) Moriyama, Y.; Takeda, K. Langmuir 1999, 15, 2003–2008.(58) Moriyama, Y.; Takeda, K. Langmuir 2005, 21, 5524–5528.(59) Markus, G.; Karush, F. J. Am. Chem. Soc. 1957, 79, 3264–3269.(60) Deep, S.; Ahluwalia, J. C. Phys. Chem. Chem. Phys. 2001, 3, 4583–

4591.(61) Yuan, H.; Antholine, W. E.; Subczynski, W. K.; Green, M. A.

J. Inorg. Biochem. 1996, 61, 251–259.(62) Parkar, W.; Somg, P. S. Biophys. J. 1992, 61, 1435–1439.(63) Kaldas, M. I.; Walle, U. K.; van der Woude, H.; McMillan, J. M.;

Walle, T. J. Agric. Food Chem. 2005, 53, 4194–4197.(64) Wang, F.; Yang, J.; Wu, X.; Liu, S. Spectrochim. Acta, A 2005,

61, 2650–2656.(65) Duggan, E. L.; Luck, F. M. J. Biol. Chem. 1948, 172, 205–220.(66) Moriyama, Y.; Watanabe, E.; Kobayashi, K.; Haranao, H.; Inui,

E.; Takeda, K. J. Phys. Chem. B 2008, 112, 16585–16589.(67) Tayyab, S.; Sharma, N.; Khan, M. M. Biochem. Biophys. Res.

Commun. 2000, 277, 83–87.(68) Hua, L.; Zhou, R.; Thirumalai, D.; Berne, B. J. Proc. Natl. Acad.

Sci. U.S.A. 2008, 105, 16928–16933.(69) Misra, R.; Mandal, A.; Mukhopadhyay, M.; Maity, D. K.; Bhat-

tacharyya, S. P. J. Phys. Chem. B 2009, 113, 10779–10791.

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