Exploring Hydrophobic Subdomain IIA of the Protein Bovine Serum

J. Phys. Chem. B 2010, 114, 6183–6196

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Exploring Hydrophobic Subdomain IIA of the Protein Bovine Serum Albumin in the Native, Intermediate, Unfolded, and Refolded States by a Small Fluorescence Molecular Reporter 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 acid methyl ester (DPDAME), has been documented to be a potential molecular reporter for probing microheterogeneous environments of a model transport protein bovine serum albumin (BSA) using spectroscopic techniques. Meteoric modifications to the emission profile of DPDAME upon addition of BSA come out to be a result of its binding to hydrophobic subdomain IIA. The highly polarity-sensitive ICT emission of DPDAME 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 from theoretical simulation using CHARMM22 software confirm the binding site of the probe to be the subdomain IIA of BSA, while FRET study reveals a remarkably close approach of our extrinsic molecular reporter to Trp-212 (in domain IIA): the distance between DPDAME and Trp-212 is 1.437 nm. The caliber of DPDAME as an external fluorescence marker also extends to the depiction of protein-surfactant (BSA-SDS) interaction to commendable fruition. Additionally, the protective action of small amounts of SDS on urea-denatured protein is documented by polarity-sensitive ICT emission of the probe. The present study also reflects the enhancement of the stability of BSA with respect to chemically induced denaturation by urea as a result of binding to the probe DPDAME. 1. Introduction

SCHEME 1: Schematic of the Structure of DPDAME

Serum albumins are abundantly found in blood plasma and are often considered as transport proteins, and this class of proteins belongs to the most widely studied category. They function as carriers for numerous exogenous and endogenous compounds in the body. The primary structure is composed of 583 amino acid residues and is characterized by low tryptophan content along with a high content of cystine, stabilizing a series of nine loops. The secondary structure of serum albumins has 67% of helix of six turns and 17 disulfide bridges.1-3 The tertiary structure is composed of three domains I, II, and III, and each domain is constituted of two subdomains A and B.1,2 Because of the common interface between domains II and III, binding of a probe to domain III associates conformational changes of domain II and hence its binding affinities. Bovine serum albumin (BSA) displays approximately 80% sequence homology and a repeating pattern of disulfides, which are strictly conserved. The molecule BSA contains two tryptophan residues, Trp-134 and Trp-212, of which the former is located in hydrophilic subdomain IB, and it is proposed to be located near the surface of the albumin molecule in the second helix of the first domain.2 The protein BSA is known to exhibit a very high conformational adaptability to a large variety of ligands.4-6 On the basis of studies using absorption, fluorescence, and circular dichroism spectroscopy,7-9 information on the binding process of many exogenous ligands like long-chain fatty acids, amino acids, metals, drug, bilirubin, etc., has been furnished at the molecular level. There are also reports where such binding has been found * To whom correspondence should be addressed. Phone: 91-33-23508386. Fax: +91-33-2351-9755. E-mail: [email protected].

to enhance the solubility of the ligands6 and the toxicity of some ligands like bilirubin diminished on binding to albumins.9 On another extreme, the past few years in the field of photochemistry and photobiology have witnessed an awesome evolution of research surrounding the photophysical studies on organic molecules of donor (D)-acceptor (A) pattern. The pioneering work of Lippert et al.10 to observe the anomalous dual fluorescence from the model compound N,N-dimethylamino benzonitrile (DMABN) envisaged a new arena of research in the realm of photochemistry through intramolecular charge transfer (ICT) reaction. Ever since its first report, the elating phenomenon of ICT reaction has continued to grow on capturing attention of researchers because of its tremendous potential to form a splendid avenue for a massive lexicon of applicative research.10-25 These include applications as electrooptical switches, chemical sensors, fluorescence probes, and so forth.26-29 In a previous work,30 we have reported the photophysical properties of DPDAME (Scheme 1) by a detailed spectroscopic study in combination with quantum chemical calculations. The present program aims at utilizing DPDAME as a molecular reporter for the investigation of the microenvironment of

10.1021/jp100004t  2010 American Chemical Society Published on Web 04/16/2010

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Paul et al.

proteinous medium. The remarkable sensitivity of the emission spectral properties of DPDAME toward medium polarity through the operation of ICT reaction paved the way for implementing DPDAME as an extrinsic fluorescent reporter for the study of proteinous microenvironment, since the process of incorporation of the probe into protein cavity will be reflected through the significant and interesting modifications on the emission profile of the former. Additionally, steady-state anisotropy, red-edge excitation shift (REES), acrylamide-induced fluorescence quenching, and fluorescence resonance energy transfer (FRET) studies have been exploited to extract valuable information regarding the nature and mechanism of the binding phenomenon through deciphering the modified photophysics of BSA-bound DPDAME. Thermal and chemical denaturation of the protein and also protective influence of small amounts of SDS on urea-induced denaturation have been manifested simply through modulation of the emission spectral behavior of DPDAME. Very recently, a nice report by Abou-Zied and Al-Shihi3a explored the native, unfolded, and refolded states of human serum albumin (HSA) mainly by using intrinsic tryptophanyl fluorescence of HSA and fluorescence lifetime techniques. The present study advocates the documentation of DPDAME as a new and efficient extrinsic molecular reporter for microheterogeneous environments of the protein BSA. 2. Experimental Section 2.1. Materials. The molecule DPDAME was synthesized and purified following a literature procedure and is described elsewhere.30 The purity of the sample was established on a TLC plate before use. Tris buffer was purchased from SRL (India), and 0.01 M Tris-HCl buffer of pH 7.4 was prepared. Bovine serum albumin (BSA) and sodium dodecyl sulfate (SDS) were purchased from SRL (India) and used as received. Analytical grade urea from SRL (India) was used after recrystallizing twice from MeOH (AR grade, Spectrochem, India). Triple distilled water was used for the preparation of all solutions. The solvent appeared visually transparent, and its purity was also tested by running the fluorescence spectra in the studied wavelength range. 2.2. Instrumentations and Methods. The absorption and emission spectra were acquired on a Hitachi UV-vis U-3501 spectrophotometer and Perkin-Elmer LS-50B fluorimeter, respectively. In all measurements, the sample concentration was maintained in the range 10-6-10-7 M in order to avoid aggregation and reabsorption effects. Experiments have been carried out at an ambient temperature of 25 °C, unless otherwise specified. Only freshly prepared air-equilibrated solutions were used for spectroscopic measurements. For spectral background corrections, a similar set of solutions in increasing BSA concentration was prepared except that the probe was omitted. During protein denaturation study by urea, spectral background corrections have been ensured with a set of solutions without a probe but containing the protein BSA (30 µM) and respective concentrations of urea as required in each set of solutions under experiment. Fluorescence quantum yield (Φf) was determined using recrystallized β-naphthol as the secondary standard (Φf ) 0.23 in methylcyclohexane) using the following equation:18-21,31

ΦS AS (Abs)R nS2 ) × × ΦR AR (Abs)S n 2 R

(1)

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 the subscripts “S” and “R” stand in recognition of respective parameters for the studied sample and reference, respectively. Steady-state anisotropy measurements were carried out using a HORIBA JOBIN YVON Fluoromax-4 spectrofluorimeter. The steady-state anisotropy is defined as3,6,7,31

r)

(IVV - G · IVH) (IVV + 2G · IVH)

(2)

IHV IHH

(3)

G)

in which IVV and IVH are the emission intensities when the excitation polarizer is vertically oriented and the emission polarizer is oriented vertically and horizontally, respectively. G is the correction factor. Fluorescence lifetimes were measured by a time-correlated single photon counting (TCSPC) spectrometer using nanoLED07 (IBH, U.K.) as the light source at 370 nm to trigger the fluorescence of DPDAME, and the signals were collected at a magic angle of 54.7°. The observed fluorescence intensities were fitted by using a nonlinear least-squares fitting procedure to a function (X(t) ) ∫t0 E(t′)R(t - t′) dt) comprising the convolution of the IRF (E(t)) with a sum of exponentials (R(t) ) A + N Biet/τi) with pre-exponential factors (Bi), characteristic ∑i)1 lifetime (τi), and a background (A). The relative contribution of each component was obtained from a biexponential fitting, expressed by the following equation:

an )

Bn N

∑ Bi i)1

The mean (average) fluorescence lifetimes for the decay curves were calculated from the decay times and the relative contribution of the components using the following equation:31

〈τf〉 )

∑ aiτi2 i

∑ aiτi

(4)

i

The excellence of the fits was judged by χ2 criteria and visual inspection of the residuals of the fitted function to the data. Circular dichroism (CD) spectra were recorded on a JASCO J-815 spectropolarimeter, using a cylindrical cuvette of 5 mm path length. The reported CD profiles are an average of four successive scans obtained at a 20 nm/min scan rate with appropriately corrected baseline. BSA concentration for CD measurements was 1.25 µM. 2.3. Theoretical Simulation Protocol. The native structure of HSA was taken from the Protein Data Bank having PDB ID 1AO6. BSA was generated from it by performing necessary additions at the N-terminal as well as some mutations in the required regions, as no PDB is available for BSA. The probe of our interest was also created by using CHARMM.32a Then, BSA was solvated in a water box and any water that comes within 2.6 Å from any protein atom was deleted. Then, three different coordinates were prepared: (1) where the probe was

Exploring Hydrophobic Subdomain IIA of BSA

Figure 1. Effect of increasing concentration of BSA (curves 1 f 14 correspond 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 the absorption profile of DPDAME on the lower energy absorption band region.

near the W212 (Trp-212) residue, (2) where the probe was near the third domain of BSA, and (3) where the probe was at the center geometry of BSA. The final simulation box contained 21918, 21927, and 21911 water molecules, respectively, for the above three arrangements. Each of the systems was energy minimized by using the ABNR method until the difference in kinetic energy of the two successive steps was less than 0.0001 kcal/mol. Then, each of the minimized structures was equilibrated for 65 ns and at 298 K. Coordinates after equilibration were generated by using the PyMOL software package32b which shows that the stable interacting conformation in all three cases is the one stated in arrangement 1. The observed conformation might be attributed to the hydrophobic interaction between the probe and Trp-212 residue. SHAKE was applied to maintain the bond lengths and bond angles of water molecules. PME was applied to measure the electrostatic interaction with a 9 Å cutoff. All of the simulations were performed using the CHARMM22 force field and parameters. 3. Results and Discussion 3.1. Solvatochromism and ICT Process in DPDAME. The molecule DPDAME shows two broad and structureless absorption bands at ∼370 and ∼260 nm in water which are ascribed to S1 r S0 and Sn r S0 (S2 r S0 or higher) transitions, respectively.30 Emission spectra of DPDAME exhibit characteristic dual fluorescence in polar solvents of which a solvent polarity insensitive higher energy local emission band is centered at ∼425 nm and a lower energy but relatively intense charge transfer band is located at ∼540 nm (in water). This large Stokes shifted (∆ν ) 8508 cm-1 in water) emission band arises from the CT state of DPDAME and thus experiences remarkable solvent polarity dependency. Steady-state absorption and fluorescence spectroscopic studies of DPDAME in solvents of different polarity clearly establish the phenomenon of ICT reaction in DPDAME.21,30 3.2. Probe-Protein Complexation Equilibrium. The complexation reaction between the probe DPDAME and protein BSA has been monitored by following the spectral changes of the probe upon binding to BSA. As seen in Figure 1, gradual addition of BSA to a solution of DPDAME in aqueous buffer (Tris buffer, 0.01 M, pH 7.4) is associated with a red shift of the absorption maxima of DPDAME (from ∼370 to ∼395 nm for the lower energy band and from ∼260 to ∼278 nm for the higher energy band) with simultaneous increase in absorbance. This might be the first indication toward the occurrence of

J. Phys. Chem. B, Vol. 114, No. 18, 2010 6185 interaction between the two concerned parties, although nothing concrete can be derived from such a response because of overlapping regions of BSA-tryptophan absorption and the higher energy band of the probe. The increment of absorbance coupled with the red shift on the longer wavelength region of DPDAME might be taken into account, as there is negligible absorption of tryptophan at this wavelength region. The protein BSA-induced spectral changes in Figure 1 suggest strong interaction between the two parties involved and can be connected to the modulation of local polarity surrounding the fluorophore (DPDAME) which subsequently modulates the stabilization of its different energy levels. With lowering of local polarity in the immediate vicinity of the fluorophore in protein environments, the energy gap between the HOMO (highest occupied molecular orbital) and LUMO (lowest unoccupied molecular orbital) decreases to give rise to the bathochromic shift. Such interpretation of our findings finds reasonable consistency with literature reports.33 Furthermore, our interpretation is substantiated by a relative red shift of absorption maxima of DPDAME upon moving from aqueous solution to a nonpolar medium like n-hexane or methylcyclohexane (λabsmax ∼ 370 nm in water vs λabsmax ∼ 375 nm in n-hexane/methylcyclohexane).30 On the other extreme, the result of interaction of DPDAME with BSA is found to produce dramatic modifications on the emission profile in the form of a large blue shift of emission maxima (Figure 2a, from ∼540 nm in aqueous buffer to ∼509 nm in 150 µM BSA) together with remarkable intensity enhancement. (However, this blue shift on the emission profile is not to be confused with the observed red shift on the absorption spectral profile because of the enormously differential natures of the potential energy surfaces (PESs) of the ground and excited states.31 The ICT phenomenon is exclusively an excited state affair, and the CT state is generated through LE state excitation; i.e., the CT state does not put its signature on the ground state PES.30,31 Hence, the interpretations have been fabricated accordingly.) Usually, the neutral and hydrophobic nature of the probe molecule favors solubilizing in the hydrophobic cavities of the protein BSA and such a large shift of the emission maxima toward the blue end of the spectrum is actually a reflection of the high degree of sensitivity of the probe toward changes in the polarity of its surrounding microenvironment.3,6 In fact, the emission maximum of DPDAME in nonpolar solvent is blue-shifted with respect to that in water (from ∼540 nm in water to ∼425 nm in methylcyclohexane).30 The blue shift in the aforementioned case is thus argued on the basis of the idea of encapsulation of the probe in the hydrophobic interior of the protein backbone which thereby offers a reduced polarity in the immediate vicinity of the probe inside proteinous medium compared to that in aqueous buffer phase. Solvent water is wellknown to act as a quencher for the emission of the CT state due to specific interactions like intermolecular hydrogen bonding,10-12,21 and as the probe molecules are less exposed to water when present inside the hydrophobic cavity of BSA, the deactivation of nonradiative decay channels comes into operation.6,7,12,31 The lower polarity of the interior of the protein cavity results in enhancement of the energy gap between the CT state and the triplet/ground states, which according to the energy gap law leads to a reduction in the radiationless decay accounting for the increment of the CT emission intensity with addition of BSA.12,31 Increased quantum yield with increasing concentration of BSA is also consistent with the same idea. Figure 2b portrays the variation of CT emission intensity of DPDAME with increasing BSA concentration. An initial steep rise of intensity is found to be followed by attainment of a

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Paul et al. TABLE 1: Different Parameters Obtained from DPDAME-BSA Binding medium

Ka (M-1)

BSA

(12.55 ( 0.4) × 103

∆Ga KSVb kETc (kJ · mol-1) (M-1) Ec (%) (ns-1) -23.38

0.913

86.96

1.216

a

K is the binding constant, and ∆G is the free energy change as obtained for DPDAME-BSA complexation from analysis of the emission data on the Benesi-Hildebrand equation. b Stern-Volmer quenching constant for acrylamide-induced quenching of BSAbound DPDAME. c E is the energy transfer efficiency, and kET is the rate of energy transfer from Trp-212 of BSA to DPDAME during FRET.

Figure 2. (a) Effect of increasing concentration of BSA (curves 1 f 15 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 of DPDAME (λex ) 370 nm and ([DPDAME] ) 2.0 µM)). (b) Variation of intensity at emission maxima and λemmax (nm) of DPDAME as a function 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).

plateau region marking the saturation of the interaction being at [BSA] ≈ 60 µM. Though the modulations of the spectral properties of DPDAME in the presence of BSA provide indications of interaction between the probe and the protein, a quantitative estimate or the extent of binding of DPDAME to the hydrophobic cavity of protein is obtained from an examination of the emission data on the Benesi-Hildebrand relation.26,34 A detailed discussion on the Benesi-Hildebrand equation is avoided here, since it is routine and profusely available in the literature6a,b,26,34 (a little elaboration is given in the Supporting Information). We thus start with the equation

I)

I0 + I1K[BSA] 1 + K[BSA]

(5)

which upon rearrangement gives

1 1 1 ) + (I - I0) (I1 - I0) (I1 - I0)K[BSA]

(6)

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

of BSA. Thus, an analysis of the fluorescence data on eq 6 paves the way for simplistic mapping of the spectroscopic modulations on a quantitative scale through estimation of the binding parameters and stoichiometry of the DPDAME:BSA complex. As seen in Figure 2c, the plot of 1/[I - I0] vs 1/[BSA] produces a straight line indicating the formation of a 1:1 complex between DPDAME and BSA. The excellence of this linear fit can be justified from the correlation factor R ) 0.9953 and standard deviation (SD) of 8.6 × 10-4. A quantitative estimate of the extent of binding, i.e., the binding constant (K), is determined from the intercept to slope ratio of the Benesi-Hildebrand plot, and the computed value is K ) (12.55 ( 0.4) × 103 M-1. Using this value of K, the free energy change for this process of complexation is determined to be ∆G ) -23.38 kJ-mol-1 (Table 1). A high K value indicates strong binding between probe and protein BSA, and the favorable process of complexation is dictated by the negative free energy change. The values obtained for K and ∆G are found to bear good consistency with literature reports.3,6,7,31,35-41 3.3. Polarity of the Microenvironment around the Fluorophore. Over the past few years, the determination of the microscopic polarity of biological systems applying fluorescent probes has been recognized as an efficient technique.42-46 The polarity determined through different photophysical parameters of the probe gives a relative measure of the polarity of the microenvironments. In the present report, we have made an attempt to estimate the micropolarity of the proteinous environments around the fluorophore. With the proteinous medium being heterogeneous, the probe molecules experience varied polarity depending on their precise location inside the protein environment.3,6,31,36,46 As mentioned earlier, the blue shift of the CT emission band of DPDAME in protein solution (in Tris buffer) reflects the lower polarity of the probe-binding site in the protein backbone. To find out the polarity experienced by our probe molecules inside the protein solution, we have employed the high sensitivity of its CT emission band toward the polarity of the medium, by taking the probe in different percentages of water and dioxane mixtures. The standard ET(30) values of the dioxane-water mixtures were determined on the basis of the intramolecular charge transfer transition of betaine dye 2,6-diphenyl-4(2,4,6-triphenyl-1-pyridino)phenolate.45 The polarity, in terms of ET(30), of the proteinous microenvironment surrounding the fluorophore can hence be determined on the basis of the position of the emission maxima of the CT band in these environments. As found from Figure 3, the micropolarity of BSA is determined to be 48.44 on the ET(30) scale, which is considerably low compared to that of bulk water (ET(30) ) 63.145) and hence accounts for the blue shift of the CT emission band of DPDAME. 3.4. Steady-State Fluorescence Anisotropy Measurements: Microviscosity of the Environment around the Fluorophore. Steady-state fluorescence anisotropy measurement occupies a commanding position in biochemical and biophysical research because of its delicate sensitivity toward sensing of any factor


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

Figure 4. (a) Variation of steady-state fluorescence anisotropy (λex ) 370 nm and λmonitored ) λemmax) of DPDAME with increasing concentration 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 an average of 10 individual measurements.

that influences the size, shape, or segmental flexibility of the fluorophore.3b,31,46 The degree of restrictions imparted by the microenvironments on the dynamical properties of the probe is directly manifested through fluorescence anisotropy values31 whence it can fruitfully be exploited in assessing the probable location of the probe in the confined medium. The environment of the probe molecule is governed by its precise location in such a complex molecular assembly. The inferences drawn from the steady-state fluorescence measurements need to be complemented with the anisotropy values in order to provide another supportive platform to our findings and interpretations. Figure 4a depicts a marked increase in the fluorescence anisotropy with increasing concentration of BSA. Such gradual enhancement of anisotropy indicates interaction between the probe and BSA. At the same time, the trend of change of anisotropy with increasing concentration of BSA reflects that the rotational diffusion of the probe is constrained significantly in the proteinous medium. Attainment of the plateau in Figure 4a implies saturation in the binding interaction between the two parties. The high anisotropy value of ∼0.30 in 50 µM BSA

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

indicates strong binding between DPDAME and BSA, complementing the values of complexation constant (K) and free energy change (∆G). Fluorescence anisotropy is very sensitive to the viscosity of the environment around the fluorophore. The microviscosity at a given temperature is often estimated by comparing the fluorescence anisotropy of a fluorophore in an environment with those of the probe in solvents of known viscosity.46a,47 In order to determine the microviscosity of the proteinous environment, the fluorescence anisotropy of DPDAME in glycerol-water mixtures of different composition has been measured and a calibration curve has been constructed by plotting the anisotropy values against weight percent of glycerol (Figure 4b). The anisotropy value suggests that the average environment around the probe molecule upon incorporation into the protein cavity corresponds to approximately 90% glycerol-water mixture (weight percentage) composition (ranis ≈ 0.30 in 90% glycerol-water mixture vs ranis ≈ 0.31 in 110 µM BSA; this concentration of BSA ensures saturation of interaction with DPDAME, as discussed in section 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 of acrylamide to a solution of DPDAME in 60 µM BSA results in quenching of the fluorescence intensity of the emission band with a slight red shift of the emission maxima (from ∼511 to ∼513 nm in 2 M acrylamide). Acrylamide does not bind to the protein but is known for its activity as an efficient static quencher of tryptophanyl fluorescence of serum albumins.31 It releases the probe molecules from the hydrophobic sites by approaching close to the site6,31 when release of the probe molecules from the hydrophobic protein environment to the aqueous phase is necessarily accompanied with quenching of the CT emission along with a shift of the emission maxima to the red end of the spectrum. Thus, the extent of perturbation of BSA-bound fluorescence of DPDAME (here in terms of quenching) by an external agent like acrylamide can be exploited as an indirect but fruitful strategy to assess the strength of binding of the probe to the protein. A careful scrutiny of the emission spectral changes imparted by the presence of acrylamide will also throw light on the probable binding site of the extrinsic fluorescence reporter.6,31 The inset of Figure 5 shows that the Stern-Volmer plot for acrylamide quenching of fluorescence of BSA-bound DPDAME leads to a Stern-Volmer constant of KSV ) slope of the plot ) 0.913 M-1. Such low magnitude of KSV obtained for quenching of fluorescence of BSA-bound DPDAME along with a little red shift conforms to only an insignificant perturbation of emission properties of DPDAME when bound to BSA, signaling toward the possibility that the probe molecules are deeply embedded in the hydrophobic pocket of the protein

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Figure 6. (a) Overlap of emission spectrum of BSA (-9-) and absorption 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 correspond to [DPDAME] (in µM) ) 0, 3.79, 7.59, 11.38, 15.18, 22.77, 26.57) (λex ) 280 nm).

BSA, which in turn ascertains a feeble approach of the quencher to the fluorophore leading to only insignificant static quenching (acrylamide is a well-known static quencher of intrinsic tryptophanyl fluorescence of serum albumins).3,6,31 3.6. Fluorescence Resonance Energy Transfer. Ever since its discovery, fluorescence resonance energy transfer (FRET)6,31,35,48 continues to be exploited as a powerful tool for measuring the distance (in nanometer scale) between donor fluorophore and acceptor fluorophore in vitro and in vivo. FRET is an electrodynamic phenomenon that occurs between a donor (D) molecule in the excited state and an acceptor molecule in the ground state. The donor molecules typically emit at shorter wavelengths that overlap with the absorption spectrum of the acceptor.31,48 Energy transfer in the process occurs without the appearance of a photon and is the result of long-range dipole-dipole interactions between the donor and acceptor. The FRET efficiency is found to depend on three parameters:31 (i) the distance between donor and acceptor must be within the specified Fo¨rster distance of 2-8 nm; (ii) there must be appreciable overlap between donor fluorescence and acceptor absorption bands (Figure 6a); (iii) proper orientation of the transition dipole of the donor and acceptor fluorophores. During FRET study, the donor emission intensity is found to be depleted with concomitant increment of acceptor emission intensity as the acceptor concentration increases. Figure 6b depicts the intrinsic fluorescence spectra of tryptophan of BSA with increasing concentration of the probe DPDAME. Increasing the concentration of DPDAME results in a diminution of tryptophanyl fluorescence of BSA with simultaneous emergence of a new band at ∼504 nm which is the CT emission of DPDAME molecule originating through the operation of FRET (with a characteristic isoemissive point at 464 nm). At the same time, the emission maximum of the tryptophan residue of BSA is found to undergo a slight hypsochromic shift in DPDAME:BSA complex (λemmax ∼ 340 nm) compared to that in free BSA (λemmax ∼ 349 nm). This observation can be rationalized on the basis of the idea of some

Paul et al. modification of the local environments of the Trp residue imparted by the presence of the probe. With a view to the neutral and hydrophobic character of DPDAME, it is not unlikely to assume a somewhat more hydrophobic environment of the emitting Trp residue in the presence of the external probe leading to a little blue shift in the emission maxima.31 Also, this observation is consistent with a recent literature report.48e At the same time, Figure 14 illustrates the location of Trp-212 and DPDAME over a hydrophobic surface (see section 3.14). Using the required spectroscopic parameters, we have calculated the FRET parameters for the presently investigated system (according to Fo¨rster’s theory) as follows: a remarkably high 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 Supporting Information; here, J ) overlap integral between donor emission and acceptor absorption spectra, R0 ) Fo¨rster’s distance, 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 energytransfer efficiency. This also indicates that the probe can measure the proteinous microenvironment at a distance of 1.437 ( 0.1 nm close to the tryptophan of protein; i.e., it can be used as a nanometric ruler. As far as our knowledge goes, it seems to be the first documentation of an extrinsic molecular reporter which is capable of approaching deep inside the protein cavity to this extent. That the ratio r/R0 amounts to 0.712, i.e., within the range 0.5-2.0, rationalizes the practical reliability of the FRET process to measure the distance between donor and acceptor chromophores 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 remarkably faster than the donor decay rate (τD ) 6.3 ns31). This in turn advocates for the observed high efficiency of energy transfer.31 However, it is ethical to mention here that the value of the orientation factor κ2 used in the calculation is 2/3, which is not precisely correct as it is the value for donor and acceptor that randomize by rotational diffusion prior to energy transfer.31 The reasons behind using the value κ2 ) 2/3 have been ornately argued in the Supporting Information, and we advocate that with κ2 ) 2/3 no significant error is included.3,6,31,46,48 Indeed, our results are well assisted by available literature that describe the use of κ2 ) 2/3 during FRET measurements in various confined environments.3,6,31,35,46,48 Confirmation for the occurrence of FRET between the present choices of pair has been derived from blank experiments which produced 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 considerably low compared to that in the presence of the latter. However, predictions about the location of the probe during FRET are not that easy, since the situation in BSA is pretty complicated by the presence of two tryptophan moieties (Trp212 and Trp-134). Previous studies reveal that Trp-134 is more exposed to the aqueous phase and its fluorescence properties are quite different.49 Our findings in the following sections, however, tend to support the occurrence of FRET from Trp212 of BSA to DPDAME. This proposition is indeed reinforced from spectral blue shift on the emission profile of DPDAME in the presence of BSA (section 3.2, Figure 2), which corroborates to binding of the extrinsic probe to the hydrophobic domain of BSA, since Trp-212 and Trp-134 are, respectively, located at hydrophobic subdomain IIA and hydrophilic subdomain IB.


Figure 7. Effect of changing excitation wavelength on the emission maxima of DPDAME:BSA complex under different experimental conditions.

3.7. Wavelength-Sensitive Fluorescence Parameters. A wavelength-sensitive tool for directly monitoring the environment and dynamics around a fluorophore in a complex biological system and the solvation dynamics in an organized medium is the “red-edge excitation shift” or REES, i.e., the shifting of the emission maxima to the red end of the spectrum upon shifting of the excitation wavelength to the red end of the absorption spectra of the fluorophore.49-51 Here, we have monitored the dependence of emission maxima of DPDAME on excitation wavelength under various conditions of binding to BSA. The occurrence of excitation-wavelength dependence is connected to the presence of an ensemble of molecules in the ground state differing in their solvation sites and hence energies.49-51 However, this condition alone does not ensure the occurrence of REES because of rapid relaxation (in the form of rapid solvation of the fluorescent state or energy transfer between energetically different excited states or conformers) of the excited state. Thus, apart from the condition of selective excitation of energetically different species, REES is also subject to slow (and hence incomplete) relaxation of the excited state. Precisely, the operation of REES is subject to the following conditions: (a) The molecule should be polar with the dipole moment higher in the excited state than that in the ground state. In fact, the extent of inhomogeneous broadening of the absorption spectra allowing the provision of site photoselection of energetically different species is dependent on the change of dipole moment (∆µ) upon photoexcitation through the relation ∆υ ) A∆µF-3/2(kT)1/2 according to the Onsager sphere approximation50 (here, A is a constant that depends on the dielectric constant of the medium, F is the Onsager cavity radius, and k is Boltzmann’s constant). For DPDAME, ∆µ ) 16.42 D30 (at the DFT/B3LYP/6-31G(d,p) level of calculation and using the Lippert-Mataga equation31). However, additional broadening, which can play even a greater role in inhomogeneous broadening of absorption spectra, may be induced by specific interactions of the sort of hydrogen bonding, electrostatic interactions, and so forth.49-51 (b) The solvent molecules around the fluorophore must be polar, and the solvent reorientation time (〈τsolvent〉) should be slower or comparable to the fluorescence lifetime (τf) of the fluorophore so that unrelaxed fluorescence can give rise to excitation-wavelength-dependent emission behavior. Figure 7 furnishes the information obtained from REES measurements of DPDAME under different conditions. It is seen that the shift of excitation wavelength from 370 to 460 nm (i.e., during REES measurements, scrupulous care has been devoted to the selection of excitation wavelength to ensure that the variation of λex involves shifting to the red end of the absorption spectra and not merely the absorption maxima; this is a very crucial criterion for REES monitoring50,51) results in shift of

J. Phys. Chem. B, Vol. 114, No. 18, 2010 6189 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 offers restriction to the rotation of solvent dipoles around the excited fluorophore. Furthermore, given the complex structural architecture of BSA, additional broadening of absorption spectra (as mentioned above) is not unlikely to contribute to the operation of REES in the present case. However, for the same change of excitation wavelength in the presence of 4.5 M urea (unfolding of BSA by urea is believed to proceed through a two-state transition process with the intermediate being formed at 4.5-5 M concentration of urea52), the REES value is found to diminish to 5 nm, and keeping track of this, REES trims down to zero in the presence of 8.0 M urea. Denaturation of BSA with increasing concentration of urea results in greater exposure of the probe molecules to the aqueous buffer environment from the confined environment inside the protein backbone, a direct consequence of which is manifested through minimization of the REES effect with gradual increase of urea concentration (i.e., the extent of denaturation). At a urea concentration as high as 8.0 M, complete denaturation of BSA takes place when the microenvironment around the fluorophore is likely to be qualitatively comparable with that in the bulk aqueous phase, as supported by the absence of REES. This proposition is also supported by the absence of REES in 80 µM BSA at 80 °C at which thermal denaturation of BSA takes place, resulting in greater exposure of the probe molecules to aqueous buffer phase compared to its bound state in rigid protein surroundings. 3.8. Urea-Induced Protein Unfolding Studies. Steady-state fluorescence measurements that dictate the changes in the tertiary structure of proteins are complementary pathways to explore the environmental stability of globular proteins.1,6 The unfolding process of serum albumins with urea is quite well studied.6 Denaturation of BSA with urea takes place via a twostate transition through intermediate state (Int) at 4.5-5.0 M urea.52,53 After finding the binding interaction between the probe and BSA, we intended to see the effect of denaturation of protein on its binding activity and on the overall photophysics of DPDAME. Here, the urea-induced modifications to spectral characteristics of protein-bound DPDAME have been followed by steady-state fluorescence measurements. Figure 8a displays the changes in the emission spectra of BSA-bound DPDAME with increasing concentration of urea. As seen in Figure 8a, gradual addition of urea to a solution of DPDAME in 30 µM BSA results in a decrement of intensity of the CT emission band with simultaneous shift of the emission maxima to the red; i.e., the pattern observed is quite reversed with respect to that in Figure 2a. Addition of urea leads to weakening of the probe-protein binding and the probe molecules are thereby exposed more to the bulk aqueous buffer phase compared to its bound condition in the native state of the protein. It is believed that urea displaces some of the water molecules adjacent to the probe in the protein environment with the consequent denaturation of the protein.54,55 The resulting destabilization of the complex should be associated with a greater exposure of the probe to the aqueous buffer phase compared to that in its bound state in the native conformation of the protein,54 and hence, the CT emission band intensity decreases along with a red shift of the emission maxima. It is important to note that, at a substantial concentration of urea (9 M), the emission wavelength (Figure 8b) and steady-state anisotropy value (inset of Figure 8b) tend to correspond to the values in aqueous buffer solution. These observations are in line with the idea that the probe DPDAME binds to the protein

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Figure 8. Chemically (urea) induced denaturation of BSA as sensed by the polarity-sensitive ICT emission of DPDAME. (a) Representative spectra showing the effect of increasing concentration of urea (curves 1 f 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) Plot of emission maxima of DPDAME:BSA complex as a function of urea concentration (0 f 9.5 M). Inset shows the decrease in the steadystate fluorescence anisotropy of BSA-bound probe against increasing urea concentration (each data point is an average of 10 individual measurements).

BSA in its native conformation, and denaturation of the latter leads to a greater exposure of the probe from the protein backbone to the bulk aqueous buffer phase. Figure 8b depicts the variation of emission maxima of the CT band with concentration of urea. As seen in this figure, up to a urea concentration of 3 M, λemmax increases slowly and then the rise is quite sharp followed by another more or less steady change getting off from a urea concentration of 7 M. This observation is consistent with reports available in the literature.3,6,46 However, we think it is ethical to point out here that the appearance of emission spectra in Figure 8a exhibits some noticeable broadening particularly toward the region of high concentration of urea. Appropriate spectral background correction negates the possibility of interference from any unusual instrumental response (signal-to-noise ratio), while the use of low emission and excitation slit widths (3.0 nm) and conservation of all other experimental conditions during the entire experiment confirm the minimization of instrumental artifacts. Under such circumstances, we presume that such noticeable broadening of the emission profile of DPDAME shows signs of extensive solute-solvent interactions. The solution under experiment is quite complicated by composition containing three individual species, viz., BSA (30 µM), DPDAME (2 µM), and urea. Thus, it is not unlikely on the part of the probe to encounter complex and extensive solute-solvent interaction (contributing to spectral broadening31) with denaturation of the protein. In fact, the extent of broadening is found to be more prominent with the progress of the process of denaturation with increasing denaturant concentration.

Paul et al. It seems crucial at this stage to make an attempt to decipher whether the probe is completely free to move or is in a somewhat restricted environment in the denatured state of the protein. For this purpose, a meticulous perusal of the steadystate fluorescence anisotropy values has been undertaken and it appears that, despite the lowering of the anisotropy of the protein-bound probe with increasing urea concentration (inset of Figure 8b), the anisotropy value at a reasonable urea concentration remains still somewhat higher than the value in the absence of the protein: anisotropy, ranis ≈ 0.013 for the probe in buffer (Figure 4a) vs ranis ≈ 0.125 for the probe in the presence of 9.5 M urea (inset of Figure 8b). Also, we have monitored the effect of increasing concentration of the probe on the intrinsic fluorescence properties of BSA under denatured conditions (30 µM BSA + 6 M urea) to follow whether the probe can induce any significant perturbation to the intrinsic fluorescence properties of the protein. The concept behind the experiment was as follows. If the probe binds strongly to BSA in its native state and is released upon uncoiling of the protein, the external addition of the probe to denatured protein should have entailed no significant modulation to the intrinsic fluorescence of BSA. However, the fruitfulness of the experiment was little crumpled by the observation reported in Figure 6a and section 3.6; i.e., an appreciable overlap between tryptophanyl emission and absorption spectra of DPDAME led to the operation of FRET with consequent quenching of intrinsic (tryptophanyl) fluorescence intensity of BSA. However, a noticeable 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 the proposition of binding of the probe to the native conformation of the protein and its subsequent greater extent of exposure toward 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 FRET efficiency.48 However, with a view to fathom deeper into the results, we have calculated the donor (Trp-212)-acceptor (DPDAME) distance in the presence of 6 M urea (see also the Supporting Information) and the result comes out to be r ) 1.656 ( 0.13 nm, which is not too large with respect to the value obtained in the case of native BSA and is thus harmonizing with the anisotropy data quite satisfactorily, as mentioned above. Also, these data appear to corroborate with our discussions in section 3.5. 3.9. Structural Stability of BSA toward Chemically Induced Denaturation by Urea upon Interaction with DPDAME. While carrying out a study on the application of the simple charge transfer probe DPDAME to study the microenvironments of proteinous media, it is pertinent to establish whether binding of the probe to the protein leads to stabilization or destabilization of the latter. The effect of binding of DPDAME on the stability of BSA has been investigated by urea-induced unfolding of the protein by monitoring the steadystate fluorescence. For this purpose, we have excited BSA at 280 nm (λabsmax of tryptophan residue in BSA) in the presence and absence of the probe DPDAME and recorded the fluorescence of tryptophan residue at 350 nm in the presence of varying concentrations of urea. Tryptophanyl fluorescence of BSA is found to gradually diminish due to the denaturing action of urea. A transition curve is constructed by plotting I/I0 against the concentration of urea, where I and I0 are the fluorescence intensities, respectively, in the presence and absence of urea. The transition curves appear sigmoidal (Figure 9a). The values of urea concentration required for half completion of the



Figure 10. Effect of increasing temperature on the emission spectral profile of the DPDAME-BSA complex ([BSA] ) 80 µM; λex ) 370 nm). Curves 1 f 8 correspond to T ) 273, 283, 293, 303, 313, 323, 333, and 353 K. Figure 9. (a) Plot of the relative fluorescence (I/I0) of tryptophan of BSA against the concentration of urea in the absence and presence of DPDAME (λex ) 280 nm, λem ) 350 nm). (b) Far UV CD spectra of BSA in the absence and presence of DPDAME ([BSA] ) 1.25 µM and [DPDAME] ) 0.25 µM).

transition (i.e., when one-half of the native state of protein is denatured) are determined from the midpoints of the transition curves, and the values are found to be 3.48 and 4.15 M for the bare 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 stabilization of BSA in the presence of the probe, with respect to denaturation by urea (it is not a concern over the absolute stability of the protein, however). These observations are in track with previous reports available in the literature.3 The influence of binding of the probe on the secondary structure of the protein has been ascertained through far UV circular dichroism studies. The CD spectra for BSA (Figure 9b) monitored in the range 250-200 nm reveal the presence of two bands at ∼209 and ∼222 nm, as consistent with the literature.3b,31,56 However, Figure 9b also reveals that, for the BSA:DPDAME composite system, the appearance of the CD spectra is exactly similar to that of BSA alone, only differing in a slight decrement of ellipticity. Such very similar (almost superimposed) CD spectra of the protein in the presence and absence of the probe evidence no detectable structural change of the protein upon binding with the probe, as consistent with literature reports.3,7b 3.10. Thermal Denaturation of BSA: ICT Emission of DPDAME as an Extrinsic Probe. Unfolding of proteins is usually marked by a change in the secondary and globular structure 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 rise of temperature through changes in the spectral characteristics of the probe when bound to BSA. Figure 10 represents the emission spectra of DPDAME in the presence of 80 µM BSA (this concentration of BSA ensures saturation of binding interaction between BSA and the probe) at different temperatures. As evident from the figure, rise of temperature induces noticeable diminution in the CT emission intensity of DPDAME together with a red shift of the emission maxima (inset of Figure

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

10). The unfolding of the protein BSA with rise of temperature is necessarily to induce a greater exposure of the probe molecules to the bulk aqueous phase compared to that in its BSA-bound state in the native conformation of the protein. Thus, the study of thermal denaturation of BSA adds another support to the inference that DPDAME prefers to bind to BSA in its native conformation and, due to its neutral and hydrophobic nature, DPDAME is favorably solubilized in the hydrophobic pocket of BSA. 3.11. Variation of Fluorescence Emission with Changes in Hydrophobicity of the Microenvironment of DPDAME. Figure 11a shows that the CT band maxima of DPDAME is blue-shifted from ∼540 nm in bulk aqueous buffer phase to ∼528 nm in 20 mM SDS. In the presence of 60 µM BSA, the blue shift is even higher, to ∼510 nm. This observation points to the fact that the polarity in SDS micelle is much higher

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compared to that in the proteinous media. However, the polarities of binding hydrophobic regions of both SDS and BSA are much less than that of pure water (buffer). The micropolarity of the surroundings of the probe in terms of ET(30) in 20 mM SDS is determined to be 54.7 (Figure 3b), which is much higher than that of BSA (ET(30) ) 48.4). Here, the concentration of the probe was kept the same in all media. The emission intensity of the blue-shifted band of the probe in SDS as well as in BSA is found to increase manifold compared to that in aqueous buffer solution. As already mentioned earlier, depletion of the radiationless decay routes due to restrictions imposed on the mobility of the probe molecules contributes to such intensity enhancement. In fact, the CT emission intensity of DPDAME when plotted as a function of SDS concentration (Figure 11b) allows one to determine the critical micellar concentration (CMC) of SDS (7.10 mM which is in excellent agreement with the literature value54,55). This advocates for the efficiency of DPDAME for probing microheterogeneous environments of micellar environments. 3.12. Surfactant (SDS)-Induced Unfolding of BSA. Several studies have been reported by different spectroscopic techniques on the binding of BSA with small molecules, particularly fatty acids and surfactants.1,9,57-61,64 Binding of these molecules to a globular protein leads to alterations in the intramolecular forces responsible for maintaining the secondary structure of the protein and thereby producing conformational changes.62 In fact, characterization of these changes at atomic resolution has also been possible.61 Here, we discuss the binding of the anionic surfactant SDS with the globular protein BSA based on the fluorescence approach. The binding of SDS with BSA is known to display four characteristic regions with increasing concentration of SDS: (A) specific binding region; (B) noncooperative binding region; (C) co-operative binding region; (D) saturation binding region.9,62 Addition of SDS results in decrement of the intrinsic fluorescence intensity of Trp-212 of BSA due to greater exposure of the tryptophan moiety to the polar environment, resulting from SDS-induced uncoiling of BSA. The binding isotherms for SDS-BSA interaction can be obtained from a plot of R vs total SDS concentration (Figure 12a) which distinctly produces the four characteristic zones, viz., A, B, C, and D. The term R, i.e., the fraction of a protein bound to surfactant, is defined as

R)

Iobs - Ifree Imin - Ifree

(7)

where Iobs, Ifree, and Imin are, respectively, the observed emission intensity values of Trp-212 at any concentration of SDS, in the absence of SDS, and under saturation conditions.63 As is evident from the figure, an initial fast rising region A (ranging from [SDS] ≈ 0.0 to 0.2 mM) is followed by a slow rising zone B up to ∼0.5 mM of SDS. Here, region A represents specific binding of SDS to some high energy binding sites of BSA, while region B is representative of the noncooperative binding where the 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 value up to ∼7 mM SDS representing massive co-operative binding of 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 further binding.9 We have attempted to explore this binding phenomenon spectroscopically utilizing DPDAME as an extrinsic reporter. As seen from Figure 12b, the intensity of the CT band of

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

DPDAME solubilized in 30 µM BSA exhibits specific variation upon addition of SDS, reflecting the binding isotherm for SDS-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 aqueous buffer phase due to uncoiling of BSA in the presence of SDS. The sudden reduction of CT emission intensity at ∼0.1 mM SDS is probably due to expulsion of the probe molecules on addition of SDS. In the region of specific binding A (ranging from [SDS] ≈ 0.0 to 0.2 mM), initial uncoiling of BSA takes place and increase of the number of exposed binding sites leads to increment of the intensity of the CT band due to competitive binding of some of the expelled probe molecules to the hydrophobic binding sites of BSA. This region is followed by a small rising region B (ranging from [SDS] ≈ 0.2 to 0.5 mM) representing the noncooperative binding of SDS to BSA. The most important zone of cooperative binding, i.e., zone C, shows its onset in the SDS concentration range 0.6-0.8 mM. In this zone, maximum uncoiling or denaturation of BSA makes the deeply buried probe molecules inside the protein hydrophobic cavity expose more to the aqueous buffer phase, leading to decrease of the intensity of CT fluorescence. This region is found to move up to around 8-10 mM of total [SDS], beyond which further change in intensity is minimized, reflecting saturation of binding past this concentration of SDS and the probe molecules being hydrophobic remains solubilized in SDS micelles (CMC of SDS ) 7.1 mM as determined using DPDAME as the probe).54 In fact, solubilization of the probe molecules in SDS micelles is also supported by the final position of the emission maxima, ∼528 nm as against ∼540 nm in pure aqueous buffer. The stability of the native globular conformations of proteins is, indeed, an outcome of a delicate balance


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

between various interactions and can be affected by several factors of which the impact of surfactants on the stability of proteins has long been a subject of much extensive and critical study. Herein, the excellent harmony of our results with available literature reports9,54 advocates for the commendable efficiency of the polarity-sensitive ICT emission of DPDAME in construing an important phenomenon, protein-surfactant interactions. 3.13. Refolding of Urea Denatured BSA with SDS. The presence of SDS results in unfolding of the native conformation of the protein since the gain in free energy due to binding of SDS to the hydrophobic sites of BSA can surpass the loss in free energy due to loss of the native conformation.3-7,9,31,45 Hence, the destructive action of SDS occurs. However, SDS plays an interesting dual role as a destabilizer for the native conformation of BSA and a stabilizer for urea-denatured BSA. It is well-known that some of the helices in BSA are disrupted in the presence of SDS (which binds to hydrophobic sites of BSA) but most of them are drastically destroyed in the presence of urea (whose denaturing action is based on breaking of water structures responsible for stabilization of the native form of BSA). Thus, interestingly, the coexistence of SDS and urea was found to protect the protein conformation and this phenomenon was first observed by Duggan and Luck65 as rise in the viscosity of BSA-urea solution was prevented upon addition of certain organic anions. Later on, the problem has been addressed by several groups.57-59,66 Disulfide bridges play quite an important role in stabilizing the native conformation of BSA, the reduction of which accompanies a huge disruption of the helices. The presence of urea results in denaturation of BSA when the probe molecules are more exposed to the bulk aqueous phase which is reflected through the red shift of the polarity-sensitive ICT emission of the probe and thereby making way for following the protective action of SDS on urea-denatured BSA by monitoring the shift of the CT band upon addition of SDS in BSA-urea solution. In the range of 4-6 M urea, addition of SDS is found to result in an initial blue shift of the CT band maxima of DPDAME, indicating recoiling of urea-denatured BSA, but further increment of SDS concentration accompanies red shift of the CT band, reflecting greater exposure of the probe to the bulk aqueous phase; i.e., further addition of SDS induces uncoiling of BSA (Figure 13). Thus, the ICT probe DPDAME can successfully be utilized in probing the conformational changes induced upon addition of SDS to urea-denatured BSA solution. 3.14. Binding Site of the Probe. The efficiency of DPDAME to function as a molecular reporter for probing proteinous (BSA)

J. Phys. Chem. B, Vol. 114, No. 18, 2010 6193 and micellar (SDS) microheterogeneous environments has been demonstrated in the foregoing sections. Now, it is pertinent at this point to explore the binding site of the probe in the protein backbone for understanding the efficacy of the probe as a molecular reporter to furnish information about the microenvironment of a protein molecule. In order to assess the binding site of DPDAME in BSA, we have intertwined the results of the urea-induced denaturation study, micropolarity measurements, and the fluorescence resonance energy transfer (FRET) study. The micropolarity around the probe has been determined in different states of the protein, i.e., native (N), intermediate (Int), and unfolded (U) states. Literature reports reveal that the intermediate state (in the presence of 4.5-5.0 M urea52) is characterized by the unfolding of domain III along with a partial loss of the native form of domain I.1,31,56,66-68 The unfolded state (U) is, however, characterized mainly by unfolding of domain II.1,31,67,68 The measured micropolarity values around the probe at different states of the protein are summarized in Table 3 which reveals that the Int to U transition of BSA (involving domain II) 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) is only 2.92. This result implies a greater possibility for DPDAME to reside in domain II relative to domain I or III. In addition, the principal hydrophobic binding sites of BSA are located in domains II and III, while domain I, characterized by a strong negative charge, can serve as a better binding site for cationic probes.2,3b With an eye to the neutral and hydrophobic nature of the probe in our case, it is reasonable to exclude domain I as one of its probable binding sites. The occurrence of FRET with a very high energy transfer efficiency (E ) 86.96%; section 3.6) strongly suggests the location of the probe to be in near vicinity of the tryptophan residue in BSA. However, predictions are quite difficult here, since the situation is pretty complicated by the presence of two tryptophan residues in BSA. From the micropolarity measurements in different states of the protein, it can be inferred that the most probable binding location of the probe in BSA is domain II. On top of this, Trp-134 is located in domain IB (hydrophilic region) which has been excluded as a probable binding site of the probe and domain III (hydrophobic region) contains no tryptophan residue which infers that the location of the probe in domain III would have led to inoperativeness of FRET from Trp to DPDAME as contrary to the present findings. Our assessment receives additional support on a theoretical milieu which reveals that out of various possibilities the stable interacting conformation is the one having DPDAME in near vicinity of Trp-212 (Figure 14; computations done according to the simulation protocol described in section 2.1). The magnified segment of Figure 14 illustrates the location of the probe and Trp-212 on a background of electrostatic potential surface on the protein (as generated using PyMOL software31), revealing that the aromatic nuclei of both DPDAME and Trp212 are located on a hydrophobic region (white surface) which further reinforces our previous assignments. 3.15. Time-Resolved Measurement. Because of its inherent sensitivity toward excited state interactions, fluorescence lifetime measurements have long been recognized as a sensitive indicator of the local environment of a fluorophore, apart from being applied in monitoring sensitive issues like differential degrees of solvent relaxation around a fluorophore, the presence of more than one chemical entity in a solution, and so forth.3,6,7,17-21,31,46 Time-resolved studies of the intrinsic fluorophore tryptophan

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

a1

a2

τ1 (ns)

τ2 (ns)

χ2

〈τf〉 (ns)

quantum yield (Φf)

kr (s-1) × 10-9

knr (s-1) × 10-9

Aq. buffer BSA (8 µM) BSA (60 µM)

0.934 0.916 0.681

0.065 0.839 0.318

0.508 0.564 1.385

1.696 2.159 3.835

0.98 1.04 1.14

0.732 1.805 2.766

0.034 0.121 0.243

0.046 0.067 0.088

1.32 0.487 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) intermediate (Int) unfolded (U)

48.44 51.36 58.54

are complex, and nothing concrete can be concluded from the fluorescence decays. This is because most proteins contain more than one tryptophan which lies in different environments and, second, protein folding and unfolding are very complex and fast processes and collection of suitable data ranges in hours. Under such circumstances, the fluorescence lifetimes of probes bound to proteins throw light on the microenvironment surrounding the probe molecule in its excited state and a great deal can be inferred about the nature of protein-probe binding as well as conformational changes of proteins under various circumstances. The fluorescence decay curves for such binding processes in heterogeneous media are generally multiexponential. For DPDAME bound to BSA, the fluorescence decay curves were best fitted to a biexponential function with acceptable values of χ2 (Table 2) and exhibit an increase in the lifetime of DPDAME with increasing BSA concentration (Figure 15 and Table 2). In analogy to other such studies,3,6,7,17-21,31 the increase in lifetime is inferred to be due to depletion in nonradiative decay channels as a result of encapsulation of DPDAME in a hydrophobic cavity of the protein. In the present case, however, we prefer to avoid placing too much emphasis on the magnitude of the individual decay constants of the multiexponential decays; rather, the mean (average) fluorescence lifetimes (as defined by eq 4) are employed as an important parameter for exploiting the behavior of DPDAME when bound to the protein. (It is, indeed, not easy and desired to specifically assign the two components obtained in time-resolved fluorescence decay patterns of DPDAME under the present experimental conditions, since the situation is pretty

complicated with the extrinsic fluorescence probe bound to the protein backbone.3,6,7,31,46,54 However, with a view to the normal observations with ICT chromophores, the two decay components can be argued to be attributable to two excited state species, namely, the LE and CT species.12,69 In fact, a reasonably steady consistency in the relative abundances of the two species tend to corroborate with our steady-state spectral findings in the sense that the more abundant species conform to the higher intensity of the longer wavelength species (ICT) while, conversely, the less populated one conforms to the lower intensity of the shorter wavelength species (LE).12,69 These results are further substantiated by the theoretical calculations of the potential energy surface across the ICT reaction coordinate for DPDAME.30 However, the environment is enormously modified in the presence of the protein BSA and solvent reorganization can also put a signature in the observed decay patterns,69 which is why the mean fluorescence lifetime has been emphasized in further discussion.) The time-resolved emission decay profiles of DPDAME in pure aqueous phase and in the presence of two different concentrations of BSA are presented in Figure 15, and the relevant parameters are summarized in Table 2. From the values of mean fluorescence lifetime (〈τf〉) and quantum yield (Φf) of DPDAME in different environments, the radiative and nonradiative rate constants have been calculated using the following equations:31

kr ) Φf /〈τf〉

(8)

1/〈τf〉 ) kr + knr

(9)

The calculated values are tabulated in Table 2. Scrutiny of the data in Table 2 evidences that in protein environment the nonradiative decay constants knr are reasonably reduced from those 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 picture has been prepared using PyMOL software.32b



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

of the fluorophore in protein environment seems attributable to the diminution of radiationless decay. Fluorescence anisotropy is a property that depends on rotational diffusion of the fluorophore as well as the fluorescence lifetime.31 Thus, it is necessary to ensure that the observed change in steady-state fluorescence anisotropy of DPDAME in the presence of BSA (section 3.4) is not an outcome of any change in fluorescence lifetime. For this purpose, the average rotational correlation times have been calculated using Perrin’s equation31 for DPDAME in BSA. Perrin’s equation is given as

τc )

〈τf〉r r0 - r

(10)

in which r0, r, and 〈τf〉 are the fundamental anisotropy, steadystate anisotropy, and mean fluorescence lifetime of DPDAME, respectively. Although ideally Perrin’s equation is not applicable in a microheterogeneous environment, it can be used to a good degree of approximation considering the mean fluorescence lifetime of the system.31,46a Now, taking r0 ) 0.383,31,46a and using eq 10, we have calculated τc for DPDAME in the protein environments and it is seen to be appreciably enhanced upon binding to the protein (from τc ) 0.026 ns in aqueous buffer to τc ) 4.297 ns in 8 µM BSA to τc ) 11.082 ns in 60 µM BSA). Significant increase of rotational correlation time in BSA environments dictates that the observed change in steady-state anisotropy of the probe (section 3.4 and Figure 4) is not a result of any lifetime-induced phenomena and thereby reinforces our earlier assignment that binding to the protein imparts an enhanced rotational restriction on the probe. 4. Summary and Conclusion In the present work, the binding phenomenon of our simple charge transfer fluorescence probe DPDAME with BSA is

presented using spectroscopic techniques. The photophysics of the ICT probe is remarkably modified upon binding with the transport protein BSA, as compared to those in pure aqueous phase. Modulations of ICT photophysics of DPDAME in a protein cavity have been shaped into a tool to explore the hydrophobic subdomain IIA of BSA through determination of binding efficiency, the nature of the microenvironment around the probe, and the micropolarity and microviscosity at the binding site. Steady-state anisotropy and REES measurements have been exploited to complement the efficient binding of the probe into the protein cavity. DPDAME is also shown to be a potent molecular reporter for probing chemical and thermal denaturation of the protein. The specific binding phenomenon of SDS with BSA has also been adroitly followed by exploring the polarity-sensitive ICT emission of the probe. The renaturing action of low concentrations of SDS on urea-denatured protein is demonstrated simply through variation of emission maxima of DPDAME. A conjugate analysis of urea denaturation study, FRET study, and micropolarity measurements leads to assessing the probable binding site of DPDAME inside the protein backbone to be subdomain IIA, which is further reinforced by a theoretical simulation. Urea denaturation study also reveals a substantial enhancement of stability of the protein as a result of binding with the probe. Overall, we have demonstrated DPDAME to be an efficient molecular reporter for probing proteineous and micellar microheterogeneous environments and also the action of urea, SDS, and temperature on the stability of the protein are well reflected in the course of modifications of the photophysics of the probe. Acknowledgment. N.G. acknowledges DST, India (Project No. SR/S1/PC/26/2008), and CSIR, India (Project No. 01(2161)07/ EMR-II), for financial support. B.K.P. and A.S. gratefully acknowledge CSIR, New Delhi, India, for research fellowships. The authors convey their special thanks to Mr. Atanu Das and Professor Dr. Chaitali Mukhopadhyay of the Department of Chemistry of our university regarding the theoretical simulation and their continuous encouragement throughout the work. We greatly appreciate the cooperation received from Ms. Deboleena Sarkar and Professor Dr. Nitin Chattopadhyay of Jadavpur University, Calcutta, India, for fluorescence lifetime and anisotropy measurements. Dr. Gautam Basu of Bose Institute, Calcutta, India, is gratefully acknowledged for allowing us to record the CD spectra. We sincerely thank the reviewers for their meticulous inspection of our manuscript and constructive suggestions. Supporting Information Available: Information on the Benesi-Hildebrand equation and fluorescence resonance energy transfer. This material is available free of charge via the Internet at 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; Academic 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.

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JP100004T

Exploring Hydrophobic Subdomain IIA of the Protein Bovine Serum

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