Identification of modes of interactions between 9-aminoacridine hydrochloride hydrate and serum proteins by low and high resolution spectroscopy and molecular modeling

Abstract

Photophysical studies on binding interactions of a therapeutically important drug, 9-aminoacridine hydrochloride hydrate (9AA-HCl) with serum proteins, bovine serum albumin (BSA) and human serum albumin (HSA), have been performed using several low and high resolution spectroscopic techniques in conjunction with molecular modeling, which disclose some salient features of such interactions in ground and excited sates. The studies reveal that although 9AA-HCl forms ground state complexes with both BSA and HSA, their individual modes of binding interaction are quite different. Its resonance energy transfer efficiency is 79% and 72% with BSA and HSA respectively. It undergoes photoinduced electron transfer (PET) with BSA, but both PET and excited state proton transfer simultaneously with HSA, which are confirmed further by laser flash photolysis studies coupled with a magnetic field. Thermodynamic analyses indicate that the binding of 9AA-HCl with BSA and HSA is controlled primarily by changes in enthalpy and entropy, respectively, with corresponding binding stoichiometries of 1 : 1 and 1 : 2, respectively. Circular dichroism spectra depict some structural changes of both the serum proteins while interacting with 9AA-HCl. Docking analyses unveil the crucial role of the disparity between the cavity sizes of the proteins which might be the foremost reason behind the differential behavior of the drug towards BSA and HSA. The binding site pocket of HSA to be docked with 9AA-HCl is 1.7 times larger in dimensions than that of BSA, which facilitates HSA to bind with 9AA-HCl with higher stoichiometry compared to BSA. These differences in binding modes as well as affinities have been further confirmed by saturation transfer difference (STD) NMR experiments which show the ligand 9AA-HCl binds to BSA with higher affinity compared to HSA. In addition, unlike BSA, HSA can accommodate more than one ligand, which corroborates well with docking and fluorescence studies.

---

1. Introduction

The most extensively studied serum albumins are identified as major transport proteins in blood plasma for many compounds such as fatty acids, which are otherwise insoluble in plasma, hormones and bilirubin. Furthermore, albumins are effective in increasing the solubility of hydrophobic drugs in plasma and modulate their delivery to cells in vivo and in vitro. However, depending on the molecular and physical properties of the interacting ligands, the binding site and stoichiometry may differ, since the overall distribution, metabolism and efficacy of many drugs in the body are correlated with their affinities towards serum albumin.¹ The interaction of drugs and proteins has been playing a very important role in drug pharmacology and pharmacokinetics for a long time.^2–13 The proteins, bovine serum albumin (BSA) and human serum albumin (HSA) are characterized by their high homology (80%) and similar conformations, containing 17 disulfide bridges and a series of nine loops, assembled in three domains (I, II, III), each containing two subdomains, A and B. From the spectroscopic point of view, one of the main differences between the two proteins is that BSA has two tryptophan (Trp) residues whereas HSA has a single Trp residue.^14–18

Acridine and its derivatives have attracted much attention due to their immense pharmacological importance and extensive uses in biological fields owing to their strong interaction with biomolecules such as DNA and proteins.^19–28 Moreover, these acridine-based drugs are well-known for their antifungal, antibacterial and antimalarial properties as well as these dyes also act as model photosensitizers in photodynamic therapy (PDT).^29–31 Among them, 9-aminoacridine hydrochloride hydrate (9AA-HCl) is used as an antibacterial, mutagenic and antitumor cytotoxic drug. An additional interesting feature about 9AA-HCl is that it belongs to those rare classes of compounds which readily form excimer^32–38 and exists in neutral, singly protonated and doubly protonated forms.³⁵ A good number of biochemical and molecular biological investigations have already been made earlier using 9AA-HCl. One of the most interesting biological applications of 9AA-HCl is its ability to interact with DNA as an intercalant agent and with enzymes as actomyosin ATPase, α-chimotrypsin, and guanidinobenzoatase (GB) inhibiting their enzymatic activity.³⁶ The spectroscopic and photophysical properties of 9AA-HCl in homogeneous and micro heterogeneous media are very much helpful for a better understanding of the nature of binding and biodistribution of this kind of dye inside the living cells. In our earlier work we explored the photophysical and photochemical properties of 9AA-HCl in different micellar environment from which it is evident that in photoinduced electron transfer (PET) reactions 9AA-HCl can act as both donor and acceptor.^39,40 We also investigated the conjugation of this biologically important acridine derivative with gold nanoparticles (AuNPs) for better targeted drug delivery.⁴¹

From the viewpoint of future biophysical applications, it seems interesting to study the interaction of 9AA-HCl with different serum proteins. In this present article, we have exploited both steady-state and time-resolved studies including laser flash photolysis techniques coupled with magnetic field to understand the mode of interaction of 9AA-HCl with the proteins and to determine the binding affinity constants as well as to resolve the location of the probe molecule in the microenvironment. Although it is a challenge to identify the exact location of a small probe molecule in a complex protein environment, the binding parameters and fluorescence resonance energy transfer (FRET) along with theoretical docking studies have thrown some light on this aspect. Further, saturation transfer difference (STD) NMR experiments have also been employed to support the results obtained from spectroscopic and theoretical docking studies as well as to have deeper insight of the drug–protein interaction.

2. Experimental section

2.1. Materials

Fatty acid and globulin-free BSA and HSA were obtained from Sigma. 9AA-HCl was also purchased from Sigma. All the solutions were prepared in a phosphate buffer of pH 7.4. D₂O was purchased from Cambridge Isotope Laboratories (CIS).

2.2. Methods

2.2.1. Absorption and fluorescence measurements. The absorption spectra were recorded on a Jasco V-650 absorption spectrophotometer at 298 K. Steady-state fluorescence excitation spectra were recorded in a Spex Fluoromax-3 spectrofluorimeter using 1 cm path length quartz cuvettes. The fluorescence data was corrected for inner filter effect using the following equation,where F_corr and F_obs are the corrected and observed fluorescence intensities, respectively, and OD_ex and OD_em are the absorbances at excitation and emission wavelengths, respectively.

Fluorescence lifetime in singlet state was measured using a diode laser-based JobinYvon Horiba picosecond-resolved time-correlated-single-photon-counting (TCSPC) spectrometer with excitation wavelength at 280 nm. The data were fitted to multiexponential functions after deconvolution of the IRF by an iterative reconvolution technique using IBH DAS 6.2 data analysis software. Analysis of the fluorescence decay data I(t) was done using the following equation:

where, B_i and τ_i are the pre-exponential factor and the fluorescence lifetime, respectively. The values of reduced χ² and residuals serve as the parameters for goodness of the fit.

2.2.2. Transient absorption measurement. A nanosecond flash photolysis setup (Applied Photophysics) containing a Nd:YAG (Lab series, Model Lab150, Spectra Physics) laser was used for the measurement of transient absorption spectra. The sample was excited at 355 nm (FWHM = 8 ns) laser light. Transients were monitored through absorption of light from a pulsed xenon lamp (150 W). The photomultiplier (R928) output was fed into an Agilent Infiniium oscilloscope (DSO8064A, 600 MHz, 4Gs/s), and the data were transformed to a computer using the IYONIX software. The software origin 8.0 was used for curve fitting. The solid curves were obtained by connecting the points using B-spline option. The samples were deaerated by passing pure argon gas for 20 min prior to each experiment. No degradation of the samples was observed during the experiments.

2.2.3. Study of energy transfer. The Förster resonance energy transfer (FRET) was used to evaluate the distance between the fluorophore, using 9AA-HCl as accepter and the Trp fluorescence of the serum proteins as donor with equimolar concentrations.⁴² The efficiency of energy transfer (E) was determined by using the given equation:where r is the distance between the donor and acceptor and R₀ is the critical energy transfer distance, at which 50% of the excitation energy is transferred to the acceptor and is given by the following equation:

R₀⁶ = 8.79 × 10⁻²⁵k²n⁻⁴Jϕ (4)

where, k² is the spatial orientation factor describing the relative orientation in space of the transition dipoles of the donor and acceptor, n is the refractive index of the medium, ϕ is the fluorescence quantum yield of the donor in the absence of the acceptor, and J is the overlap integral of the fluorescence emission spectrum of the donor and the absorption spectrum of the acceptor. Under experimental conditions, k² = 2/3, n = 1.336 ⁴² and ϕ of BSA and HSA = 0.11 and 0.15 respectively.⁴³ The value of J is given by the equationwhere F(λ) is the fluorescence intensity of serum proteins (donor) at wavelength λ and ε(λ) is the molar extinction coefficient at wavelength λ. Values of E and J were obtained experimentally. Rate of energy transfer (k_ET) depends on the lifetime of the donor molecule and is given by⁴²where τ_D is the lifetime of the donor molecule in the absence of the acceptor moiety.

2.2.4. Estimation of binding constant and binding stoichiometry. The binding affinity constant (K_a) was determined from the steady state fluorescence data, with variation of the 9AA-HCl concentration. The binding affinity constant (K_a) for the association of a drug to a protein was determined according to the following equation^44–46where ΔF = F₀ − F and F₀ and F denotes the protein fluorescence in the absence and presence of the ligand (L), respectively. ΔF_max represents the maximum quenched fluorescence intensity obtained from the intercept of the linear plot of 1/ΔF against 1/L and the slope gives the value of binding affinity constant (K_a).

The method of analysis is based on the following Scatchard's equation from which we can calculate the association binding constant and number of binding sites for drug–protein interactive system by measuring the change in intrinsic fluorescence of the tryptophan in serum proteins on successive addition of drug^47–49

where r (r = ΔF/F₀) is the moles of drug bound per mole of protein and C_f is the molar concentration of free drug. The binding site n is determined from the intercept and slope of the plot of r/C_f vs. r. The binding affinity constant (K_a) is calculated from the slope.

For cooperative interaction, binding parameters were calculated from fluorescence quenching data using the procedure of Hill plot.⁵⁰ This method was based on the general equation:

where, θ = fraction of occupied sites where the ligand can bind to the active site of the receptor, protein, [L] = free (unbound) ligand concentration, K_d = apparent dissociation constant derived from the law of mass action (equilibrium constant for the dissociation), n = Hill coefficient, describing cooperativity. Positively cooperative reaction i.e., once one ligand molecule is bound to enzyme, its affinity for other ligand molecules increases, shows n > 1, negatively cooperative reaction i.e., once one ligand molecule is bound to the enzyme, its affinity for other ligand molecule decreases, gives n < 1 and noncooperative reaction i.e., the affinity of the enzyme for a ligand molecule is not dependent on whether or not other ligand molecules are already bound, has n = 1.

2.2.5. Analysis of thermodynamic parameters. The van't Hoff and Gibbs–Helmholtz equations were used for determination of the thermodynamic parameters:

ΔG⁰ = ΔH⁰ − TΔS⁰ (11)

In the above equations K_a is the binding affinity constant at corresponding temperature T, and R is the gas constant. Standard enthalpy change (ΔH⁰) and standard entropy change (ΔS⁰) were obtained from the slope and intercept respectively of the plot of ln K_a vs. 1/T and then the values of free energy change (ΔG⁰) at different temperature were obtained using eqn (10).

2.2.6. Measurements of circular dichroism (CD). The Far-UV and near CD spectra measurements were made on a Jasco-720 automatic recording spectrophotometer. An accumulation of three scans with a scan speed of 20 nm per minute and response time of 2 s was performed to collect data. The CD results were analyzed in terms of mean residue ellipticity (MRE) according to the following equation⁵¹where C_p is the concentrations of protein, n is the number of amino acid residues, and l is the path length. The α-helical content of free and bound protein was determined from the MRE value at 208 nm using the following formula⁵¹where, MRE₂₀₈ is the observed MRE value at 208 nm, 4000 is the MRE of β-form and random coil conformation at 208 nm, and 33 000 is the MRE value of a pure α-helix at 208 nm.

2.2.7. Theoretical docking studies. Structural information for serum albumins were obtained from Protein Data Bank (PDB IDs: 4L8U and 3V03 for HSA and BSA respectively, UniProt numbering convention was followed, BSA (UniProt KB: P02769) has two tryptophan residues at position 158 and 237, respectively. HSA (UniProt KB: P02768) has a single tryptophan at position 238). Model 4L8U for HSA has a resolution of 2.01 Å and it is the best model so far according to the validation report available at wwPDB.⁵² Only four models for BSA are available and the best model 3V03 with resolution 2.7 Å was chosen. The structure with missing side chain atoms (3V03) was modeled at PDB_Hydro server.⁵³ Chemical structure of 9AA-HCl was drawn and geometrically optimized using the B3LYP/6-311+G(2d,p) level of density functional theory as done earlier by our group.³⁸ Binding pockets of the serum albumins were analyzed by POCASA program.⁵⁴ Volumes of the binding pockets were calculated at 3V server.⁵⁵ Molecular Docking was performed using AutoDock Vina⁵⁶ following the previously published protocol.^9,12 All the hetero atoms and water and the extra subunits were removed from the PDB structures. Polar hydrogen atoms and Gasteiger charges were added to the proteins and the ligand. All the rotatable bonds in the ligand were set free. The whole protein was placed in the center of a simulation box. The box dimension was 87 × 66 × 80 cubic angstroms for BSA and 87 × 66 × 73 cubic angstroms for HSA. The grid point spacing of 1 Å was used for the docking. Distance constrain as obtained by FRET experiment was used to filter the docking results. For HSA one molecule of ligand was docked at first and then with the best complex, another ligand molecule was docked following the same procedure described above. Docking results were rendered in PyMOL molecular graphics program. Solvent accessible surface area (SASA) of the proteins, ligand and lowest energy protein–ligand complexes were calculated using 1.4 Å probe size, with Mark Gerstein's calc-surface program⁵⁷ on Helix Systems server at NIH (http://www.helixweb.nih.gov/structbio/). Ligand interaction diagrams were produced in Schrödinger Maestro.

2.2.8. Saturation transfer difference (STD) NMR studies. We have performed STD NMR experiments to investigate the binding behaviour of 9AA-HCl with BSA and HSA. The serum proteins as well as 9AA-HCl have been prepared in deuterated 10 mM phosphate buffer (pH 7.4). One dimensional STD experiments have been performed with STD pulse sequence containing WATERGATE scheme for water suppression as described previously.^58,59 On-resonance irradiation frequency of the receptor proteins have been set to 1.0 ppm for selective saturation of protein resonances. Similarly, off resonance frequency has been set to 40 ppm where neither the protein nor the ligand signals was detected. A cascade of 40 selective Gaussian-shaped soft pulses (49 ms each) in a delay of 1 ms has been applied ensuing in total saturation transfer of 2 s to the ligand from the macromolecule. Internal subtraction of the two spectra (on resonance-off resonance) by phase cycling generates the difference spectrum that contains ligand signals attenuated via saturation transfer from protein. It is to be noted that, the STD experiments have been performed over several saturation times (t_sat) i.e., 0.5, 1, 2, 3 and 5 s to overcome spin lattice (T₁) relaxation bias in group epitope mapping (GEM) (data not shown).⁶⁰ However, the reported data correspond to 2 s saturation time, which falls, in the slope region of build-up curves of saturation time. The total number of scans is 128 and 64 with 16 dummy scans using 14 ppm spectral widths for the STD and reference NMR spectra, respectively. Complete assignment of the molecule has been performed using two-dimensional ¹³C–¹H HSQC, ¹H–¹H COSY experiments as well as one-dimensional ¹³C DEPT experiments using standard Bruker pulse programs. All NMR experiments have been executed on a Bruker AVANCE III 500 MHz NMR spectrometer furnished with a 5 mm SMART probe operating at a proton frequency of 500.13 MHz at 298 K. Data acquisition and data processing have been carried out using Topspin™v3.1 software suite (Bruker Biospin, Switzerland).

2.3. Results and discussion

2.3.1. Absorption and fluorescence measurements. 9AA-HCl exists in three different forms i.e., neutral (9AA), protonated (9AAH⁺) and doubly protonated forms depending on the pH of the medium (Fig. S1†).³⁴ The absorption peaks and the fluorescence peaks of 9AA at pH = 11.0 are slightly red shifted compared to that of 9AAH⁺ at pH = 4.0 and pH = 7.0. At pH = 7, 9AAH⁺ predominates^39,40 which can give rise its deprotonated form by donating proton to surrounding environment both in ground as well as excited states since there is an equilibrium between 9AAH⁺ ↔ 9AA. Absorption spectroscopy is used to explore the possibility of ground state interaction between drug–protein. Therefore, the interactions of the acridine derivative with the serum proteins have been inferred from the changes in absorption spectra of serum proteins with gradual addition of drug at 298 K. In case of BSA–9AA-HCl system isosbestic point appear at 440 nm and in case of HSA–9AA-HCl system it generates at 438 nm as shown in Fig. 1A and B respectively. Since the appearance of isosbestic points suggests that an equilibrium is established between two species and reflect the formation of complexes between them, therefore, these observations signify that the ground state complexes are formed between respective serum proteins and 9AA-HCl.

Fig. 1 (A) Absorption spectra showing the formation of ground-state complex between BSA and 9AA-HCl in phosphate buffer (pH = 7.4) where [BSA] = 5.0 × 10⁻⁵ M and [9AA-HCl] ranges from 2.0 × 10⁻⁵ M to 4.0 × 10⁻⁵ M. (B) Absorption spectra showing the formation of ground-state complex between HSA and 9AA-HCl in phosphate buffer (pH = 7.4) where [HSA] = 5.0 × 10⁻⁵ M and [9AA-HCl] ranges from 1.60 × 10⁻⁵ M to 4.50 × 10⁻⁵ M.

However, the observations obtained from absorption studies are not sufficient to study the interactions in detail. Therefore other excited state spectroscopic techniques, e.g., fluorescence spectroscopy, laser flash photolysis etc., have been preferred for studying binding mode of interactions. The fluorescence of BSA comes from two Trp residues, one of them is located on the surface and the other resides in the hydrophobic pocket of the protein molecule.⁶¹ However, HSA bears only one Trp residue residing in subdomain IIA.^14,17 Fig. 2A and B depict that on progressive addition of various concentration of acridine derivative the fluorescence intensity of the Trp residues of respective serum proteins decreases along with a slight blue shift of maximum wavelength of Trp. The shift of maximum wavelength signifies hydrophobic effect on Trp residue due to complex formation between drug and respective proteins. The data of fluorescence quenching are analyzed using the following Stern–Volmer equation⁴²

where F₀ and F are the relative fluorescence intensities in the absence and presence of quencher, respectively, K_SV is the Stern–Volmer constant and [Q] is the concentration of the quencher. The plot of F₀/F vs. concentration of quencher is linear for BSA–9AA-HCl system (Fig. 3A). However, the fluorescence lifetime of BSA does not change with the gradual addition of 9AA-HCl which rules out the possibility of any dynamic quenching for BSA–9AA-HCl (Fig. S2A†). Therefore for BSA–9AA-HCl system the quenching mechanism is solely static. However, for HSA the plot is non-linear (Fig. 3B). The non-linearity with upward bending might be either due to simultaneous occurrence of static (ground state) and dynamic (excited state) quenching or for the presence of charges on interacting molecules or through the interaction of more than one fluorophore with the quencher. The static quenching has already been proven by ground state complex formation. On the other hand, fluorescence lifetime of HSA decreases on simultaneous addition of acridine derivative which indicates the occurrence of dynamic quenching (Table 1 and Fig. S2B†). Since the decrease in fluorescence lifetime in presence of quencher accounts only for dynamic quenching, the corresponding Stern–Volmer plot is expected to be linear (Fig. S3†). Therefore the positive deviation of Stern–Volmer plot for HSA–9AA-HCl system is attributed for the simultaneous occurrence of static and dynamic quenching.

Fig. 2 (A) Fluorescence spectra showing quenching of intrinsic fluorescence of BSA with increase in 9AA-HCl concentration in phosphate buffer (pH = 7.4) where [BSA] = 2.0 × 10⁻⁶ M and [9AA-HCl] ranges from 0.0 to 1.15 × 10⁻⁴ M; λ_ex = 280 nm. (B) Fluorescence spectra showing quenching of intrinsic fluorescence of HSA with increase in 9AA-HCl concentration in phosphate buffer (pH = 7.4) where [HSA] = 2.13 × 10⁻⁶ M and [9AA-HCl] ranges from 0.0 to 2.64 × 10⁻⁴ M; λ_ex = 280 nm.

Fig. 3 Stern–Volmer plot of quenching (A) of BSA by 9AA-HCl where [BSA] = 2.0 × 10⁻⁶ M and [9AA-HCl] ranges from 0.0 to 1.15 × 10⁻⁴ M; (B) of HSA by 9AA-HCl where [HSA] = 2.13 × 10⁻⁶ M and [9AA-HCl] ranges from 0.0 to 2.64 × 10⁻⁴ M.

Table 1 Variation of lifetime (τ₁ and τ₂) of HSA with increase in concentration of 9AA-HCl in phosphate buffer (pH = 7.4); (λ_em = 350 nm)^a

HSA (M)	9AA-HCl (M)	Relative amplitude = B1	Relative amplitude = B2	τ1^b (ns)	τ2^b (ns)	χ^2
a λ = wavelength, χ^2 is the goodness of fit.b ±5%.
1.5 × 10^−5	0	34.59	65.41	4.14	7.51	1.14
	2.02 × 10^−5	25.28	74.72	3.55	7.16	1.19
	2.02 × 10^−4	24.86	75.14	2.74	6.72	1.23
	4.03 × 10^−4	26.84	73.16	2.03	6.39	1.26
	6.30 × 10^−4	24.49	75.51	1.53	5.79	1.49

---

2.3.2. Determination of binding constant and binding stoichiometry. The binding affinity constant (K_a) is evaluated by making use of the steady-state fluorescence data obtained with the variation of 9AA-HCl concentration in serum proteins. For BSA–9AA-HCl system the K_a value is 1.13 × 10⁵ M⁻¹, whereas for HSA–9AA-HCl system that becomes 2.82 × 10³ M⁻¹ at 298 K, which is almost forty times smaller compared to the previous one (Tables 2 and 3). Moreover, with increase in temperature the changes in the K_a values contradict each other in case of two serum proteins. The K_a value for BSA–9AA-HCl system decreases with increasing temperature, whereas that obtained for HSA–9AA-HCl system increases with temperature. From the absorption and fluorescence quenching studies it has already been confirmed that ground state complex is formed between BSA and 9AA-HCl. The increased temperature decreases the stability of the complex which is formed on addition of 9AA-HCl to BSA; thus it decreases the static quenching constant, hence K_a. However, it is well reported in the literature that HSA is much more flexible compared to BSA.¹² In this case, HSA undergoes both static and dynamic quenching with 9AA-HCl. It is pertinent to mention here that 9AA-HCl somehow distorts the structure of the protein. Therefore, 9AA-HCl became much more accessible towards Trp which is reflected in the enhancement of K_a values with increase in temperature.

Table 2 Determination of binding affinity constants and thermodynamic parameters of BSA–9AA-HCl system at different temperatures^a

Temperature (K)	Binding affinity constant (Ka) M^−1	ΔH^0 (kJ mol^−1)	ΔS^0 (J mol^−1 K^−1)	ΔG^0 (kJ mol^−1)
a ΔH^0 = standard enthalpy change, ΔS^0 = standard entropy change, ΔG^0 = Gibbs free energy change.
298	1.13 × 10^5	−310.42	−944.44	−28.97
303	1.71 × 10^4			−24.25
308	1.93 × 10^3			−19.53

---

Table 3 Determination of binding affinity constants and thermodynamic parameters of HSA–9AA-HCl system at different temperatures^a

Temperature (K)	Binding affinity constant (Ka) M^−1	ΔH^0 (kJ mol^−1)	ΔS^0 (J mol^−1 K^−1)	ΔG^0 (kJ mol^−1)
a ΔH^0 = standard enthalpy change, ΔS^0 = standard entropy change, ΔG^0 = Gibbs free energy change.
298	2.82 × 10^3	175.37	654.36	−19.63
303	8.43 × 10^3			−22.90
308	2.81 × 10^4			−26.17

---

In the earlier report it had been depicted that either 1 : 1 or 1 : 2 complexes are formed between proteins and drug moieties.^12,13 To ascertain its viability in the present complexes i.e., between 9AA-HCl and serum proteins, the Scatchard plot and the Hill plots are used. The Scatchard plot for BSA–9AA-HCl system, give the binding stoichiometry is 1 : 1 (Fig. 4). However, the Hill plot evaluates the presence of cooperative binding that means the binding of a ligand to a macromolecule which is often enhanced if there are other ligands present previously at the same molecule. Fig. 5 illustrates the Hill plot with the slope i.e., the Hill coefficient n_H, as 1.8 (∼2) which signifies positive cooperativity (n_H > 1) that means when one 9AA-HCl binds to HSA, it helps another drug moiety to interact with protein. The indication of formation of 1 : 2 complex between HSA and drug is also supported by the docking as well as STD NMR studies as discussed later.

Fig. 4 Scatchard plot for BSA–9AA-HCl system in phosphate buffer; [BSA] = 10.0 × 10⁻⁶ M and [9AA-HCl] ranges from 5.22 × 10⁻⁵ to 1.15 × 10⁻⁴ M.

Fig. 5 Hill plot for HSA–9AA-HCl system in phosphate buffer where [HSA] = 10.0 × 10⁻⁶ M and [9AA-HCl] ranges from 1.60 × 10⁻⁵ to 2.64 × 10⁻⁴ M.

2.3.3. Thermodynamic parameters. It is necessary to determine the nature of interacting forces between drug and protein which can be predicted by measuring the thermodynamic parameters of binding, i.e., changes in standard enthalpy (ΔH⁰), entropy (ΔS⁰) and Gibb's free energy (ΔG⁰) using van't Hoff isotherm. Usually four types of forces play a part in drug–protein interaction, like electrostatic forces, hydrophobic forces, van der Waal's interactions and hydrogen bonding. From the signs of the thermodynamic parameters the nature of forces involved in the drug–protein interaction may be predicted. Tables 2 and 3 depict two sets of values on thermodynamic parameters, one is for BSA–9AA-HCl system and the other is for HSA–9AA-HCl system respectively. The nature of van't Hoff plot (Fig. 6A and B) for two serum proteins appreciably differs from each other and as well as the signs of thermodynamic parameters (ΔH⁰ and ΔS⁰) for two interactive systems differ. These kinds of discrepancy support the fact that forces involved in the BSA and HSA interactions are of different type.

Fig. 6 Van't Hoff plot for the binding of (A) BSA (2.0 × 10⁻⁶ M) with 9AA-HCl (0.0 to 1.15 × 10⁻⁴ M) and (B) of HSA (2.13 × 10⁻⁶ M) with 9AA-HCl (0.0 to 2.64 × 10⁻⁴ M) in the phosphate buffer at three different temperatures, 298 K, 303 K and 308 K.

In case of BSA–9AA-HCl system, the negative change in both enthalpy and entropy values clearly suggest that primarily the key binding forces are hydrogen bonding and van der Waal's. Unlike BSA–9AA-HCl system, the positive values of entropy and enthalpy are calculated for HSA system. The positive thermodynamic parameters indicate that the binding of drug with HSA is entropy driven process where ionic and hydrophobic interaction are the key forces and 9AA-HCl binds to a more hydrophobic environment in HSA compared to that of BSA. The overall negative changes in free energy depict the spontaneity of the binding interaction between drug and serum proteins.

2.3.4. Energy transfer between 9AA-HCl and both serum proteins. It has been observed that the absorption spectrum of 9AA-HCl and the fluorescence spectra of both serum proteins overlap with each other to some extent which is the primary requirement for the occurrence of FRET. The serum proteins which act as donor emit in the shorter wavelength region (maximum at 340 nm) and that overlaps with the corresponding absorption spectrum of 9AA-HCl which acts as an acceptor (Fig. 7A and B). Now, Förster's theory of nonradiative energy transfer is employed to calculate the efficiency of energy transfer (E) and the distance between 9AA-HCl and serum proteins which show that for smaller donor–acceptor distance the value of energy transfer efficiency become higher. Here the efficiency of energy transfer signifies the fraction of photons absorbed by the donor moiety which transferred to acceptor.⁴² The values of the efficiency of energy transfer (E), the critical energy transfer distance (R₀) and the distance between the donor and the acceptor (r) are evaluated by using the required equations for both the systems and their values are depicted in Table 4. The values of r are 1.82 nm and 1.83 nm for BSA–9AA-HCl and HSA–9AA-HCl respectively. In case of average distance r < 8 nm, that follows the condition, 0.5R₀ < r < 1.5R₀, the energy transfer from the proteins to 9AA-HCl occurs with highest probability.⁶²

Fig. 7 Spectral overlap between the absorption spectrum of 9AA-HCl (acceptor) and the fluorescence spectrum (A) of BSA (donor) where [BSA] = [9AA-HCl] = 5.0 × 10⁻⁵ M; (B) of HSA (donor) where [HSA] = [9AA-HCl] = 5.0 × 10⁻⁵ M.

Table 4 The values obtained from the FRET analyses (E, R₀ and r) for 9AA-HCl and serum proteins systems

System	Efficiency of energy transfer (E) in %	Critical energy transfer distance (R0) in nm	Distance between the donor and the acceptor (r) in nm
9AA-HCl–BSA	79	2.27	1.82
9AA-HCl–HSA	72	2.15	1.83

---

2.3.5. Circular dichroism studies. The circular dichroism (CD) is a sensitive technique to monitor the conformational changes in serum proteins upon interaction with acridine derivative. Native serum proteins show two characteristic negative bands in the far-UV region at ∼209 nm and ∼222 nm which depict a typical characteristic of an α-helical protein.^63–65 The α-helix contents of native BSA and HSA in the presence of 9AA-HCl have been calculated which are summarized in Table 5 and depicted in Fig. 8A and B respectively. It is observed that binding of 9AA-HCl to serum proteins causes a reduction of α-helix of the proteins signifying that secondary structure of protein changes in presence of acridine derivative. Besides that, moderate change in spectrum is observed for both serum proteins in near-UV-CD region (250–350 nm) in presence of 9AA-HCl as depicted in Fig. 9A and B respectively, which indicates that 9AA-HCl perturbs the tertiary structure of the protein to some extent.

Table 5 Variation in α-helix% of BSA and HSA with increase in 9AA-HCl concentration

[BSA] M	[9AA-HCl] M	α-Helix%	[HSA] M	[9AA-HCl] M	α-Helix%
1.25 × 10^−6	0	61.25	1.35 × 10^−6	0	67.42
	6.30 × 10^−6	53.53		6.30 × 10^−7	56.79
	9.45 × 10^−6	45.66		3.15 × 10^−6	46.02
	2.52 × 10^−5	36.95		6.30 × 10^−6	36.50

---

Fig. 8 Far-UV CD spectra (A) of BSA (1.25 × 10⁻⁶ M) on addition of 9AA-HCl where [9AA-HCl] are (i) 0 M, (ii) 6.30 × 10⁻⁶ M, (iii) 9.45 × 10⁻⁶ M, (iv) 2.52 × 10⁻⁵ M; (B) of HSA (1.35 × 10⁻⁶ M) on addition of 9AA-HCl where [9AA-HCl] are (i) 0 M, (ii) 6.30 × 10⁻⁷ M, (iii) 3.15 × 10⁻⁶ M, (iv) 6.30 × 10⁻⁶ M in phosphate buffer (pH = 7.4).

Fig. 9 Near-UV CD spectra (A) of BSA (1.25 × 10⁻⁶ M) where [9AA-HCl] are (i) 0 M, (ii) 2.52 × 10⁻⁶ M, (iii) 3.78 × 10⁻⁶ M; (B) of HSA (1.35 × 10⁻⁶ M) where [9AA-HCl] (i) 0 M (ii) 6.30 × 10⁻⁶ M in phosphate buffer (pH = 7.4).

2.3.6. GuHCl-induced denaturation of BSA and HSA: studied by 9AA-HCl. Now, the conformational stability of globular protein can be explored by two complementary pathways such as steady-state fluorescence measurements and changes in the tertiary structures of proteins. The unfolding processes of serum albumins on increasing concentration of denaturating agent have been well studied.^12,66 After finding the binding interaction between serum proteins and drug, we intend to see the denaturing effect of the protein on its binding activity and on the overall photophysics of the drug. In this present case, GuHCl-induced modification of the protein bound drug is studied by means of steady-state fluorescence measurements. For both BSA–9AA-HCl and HSA–9AA-HCl systems, the fluorescence intensities in the presence of GuHCl (F) to that in absence (F₀), are plotted against added concentration of GuHCl. Fig. 10A and B depict the variation patterns which appear sigmoidal in nature. From the transition curves, the values of GuHCl concentration are measured where one-half of the native state of protein is denatured indicated as [den]_1/2 and it is determined from the midpoints of these transition curves. In presence of GuHCl, [den]_1/2 moves from 1.29 for BSA to 1.49 in the presence of 9AA-HCl and from 1.74 to 1.40 for HSA in the presence of the same amount of 9AA-HCl. In case of BSA, the serum protein is stabilized by 9AA-HCl, therefore [den]_1/2 increases in presence of the acridine derivative. However, in case of HSA, the scenario is different. The ionic nature of 9AA-HCl increases overall ionic strength of the medium because of the presence of unbound or free 9AA-HCl associated with ionic GuHCl which makes denaturation much more facile for HSA systems.

Fig. 10 Denaturation plot for (A) BSA–9AA-HCl system where [BSA] = 4.24 × 10⁻⁶ M and [9AA-HCl] = 0 M to 1.35 × 10⁻⁵ M and (B) HSA–9AA-HCl system where [HSA] = 7.50 × 10⁻⁶ M and [9AA-HCl] = 0 M to 1.11 × 10⁻⁴ M; λ_ex = 280 nm and λ_em = 350 nm.

2.3.7. Laser flash photolysis study. Laser flash photolysis study coupled with an external magnetic field (MF = 0.08 T) is employed to investigate the interaction of 9AA-HCl with two serum proteins in the excited state. From the literature it is found that the TrpH˙⁺ and Trp˙ show maxima absorbance around 350 nm, 560–570 nm, and at 520 nm respectively.^11,47,67,68 Earlier we have observed the transient absorption spectrum of the transients, 9AAH˙ and 9AA˙⁻ have maxima absorbance at 380 nm and 570 nm respectively.^39,40

Fig. 11A depicts the transient absorption spectra of the acridine derivative after 0.6 μs time delay of the laser flash at 355 nm with BSA in phosphate buffer. In absence of serum protein, the spectrum shows two strong characteristic absorption maxima around 440 nm and 470 nm and a small peak around 530–540 nm. The two proximal peaks at 440 nm and 470 nm have been attributed to 9AAH⁺ and the red shifted one at 530–550 nm to 9AA.^39,40 However, in the presence of BSA, the transient spectrum shows overall quenching around 440 nm, 470 nm and 530–540 nm accompanied by an increase in absorbance around 380 nm and 570 nm along with the generation of an isosbestic point at 408 nm. Quenching of the absorbance in the region characteristic to 9AAH⁺ and 9AA gives an indication of PET occurring from serum protein to acridine derivative. Previous reports show that the peak at 380 nm and 570 nm can be accounted for the presence of TrpH˙⁺.¹¹ However, the spectral signatures of 9AAH˙ and 9AA˙⁻ have also been assigned by the peak at 380 nm and 570 nm respectively. Therefore the peaks around 380 nm, 570 nm are unambiguously assigned for TrpH˙⁺, 9AAH˙ and 9AA˙⁻ respectively. These radical moieties may be formed via PET from the tryptophan of BSA to both the 9AAH⁺ and 9AA. In absence and presence of BSA, the lifetimes obtained from the decay profile at 380 nm are 1.37 μs and 2.16 μs respectively (inset of Fig. 11A). The increase in lifetime in presence BSA corroborates the fact that radical ions are formed due to the course of PET reactions.

Fig. 11 (A) Transient absorption spectra of 9AA-HCl (1.81 × 10⁻⁵ M) in the (i) absence and (ii) presence of BSA (6.0 × 10⁻⁴ M) in phosphate buffer after the laser flash at 355 nm at a time lag of 0.6 μs. Inset shows the decay profiles at 380 nm of 9AA-HCl (1.81 × 10⁻⁵ M) in the (i) absence and (ii) presence of BSA (6.0 × 10⁻⁴ M) in phosphate buffer at 0.4 μs after the laser flash at 355 nm. (B) Transient absorption spectra of 9AA-HCl (1.81 × 10⁻⁵ M) and BSA (6.0 × 10⁻⁴ M) in absence and in presence of magnetic field at 1 μs after the laser flash at 355 nm in phosphate buffer.

Fig. 11B shows the transient absorption spectra of 9AA-HCl (1.81 × 10⁻⁵ M) and BSA (6.00 × 10⁻⁴ M) in the presence and absence of MF in phosphate buffer at a delay of 1 μs after the laser flash. On application of an external MF in BSA–9AA-HCl system, there is significant enhancement in absorbance in the wavelength regions around 370–380 nm and 570 nm which correspond to TrpH˙⁺ as well as for 9AAH˙ and 9AA˙⁻ respectively (Fig. 11B). MFE at the above wavelengths arises out of the efficient coupling of spin and diffusion dynamics of the spin correlated geminate pairs proving that these radicals are triplet-born and PET primarily occurs in the triplet state.

The scenario is somewhat different in case of HSA because of its mobile nature compared to BSA. Fig. 12A shows the transient absorption spectra of 9AA-HCl and HSA after 0.4 μs time delay of the laser flash at 355 nm in phosphate buffer. It is observed that a peak develops at 530 nm and 570 nm accompanied by formation of an isosbestic point at 520 nm as well as quenching of absorbance around 440 nm and 470 nm. In this case, increase in absorbance at 530 nm and 570 nm implies the formation of Trp˙ and TrpH˙⁺respectively. The peak at 570 nm can also be assigned for 9AA˙⁻. These changes in transient spectra imply that both PET and proton transfer are present in the HSA–9AA-HCl system. The quenching at 440 nm and 470 nm are not much significant as in BSA. That may be due to the occurring of excited state proton transfer (ESPT) from TrpH˙⁺to 9AA which results in the formation of 9AAH⁺ leading to an increase in absorbance in that region. Similar cases of proton transfer from TrpH˙⁺ to other species have been reported earlier.¹¹ In absence and presence of HSA, the lifetimes obtained from the decay profile at 530 nm are 1.95 μs and 2.12 μs respectively (inset of Fig. 12A), supporting the formation of radical ions through PET.

Fig. 12 (A) Transient absorption spectra of 9AA-HCl (1.81 × 10⁻⁵ M) in (i) absence and (ii) presence of HSA (5.0 × 10⁻⁵ M) in phosphate buffer after the laser flash at 355 nm at a time lag of 0.4 μs. Inset shows the decay profiles at 530 nm of 9AA-HCl (1.81 × 10⁻⁵ M) in (i) absence and (ii) presence of HSA (5.0 × 10⁻⁵ M) at 0.6 μs after the laser flash at 355 nm. (B) Transient absorption spectra of 9AA-HCl (1.81 × 10⁻⁵ M) and HSA (5.0 × 10⁻⁵ M) in absence and in presence of magnetic field at 0.4 μs after the laser flash at 355 nm in phosphate buffer.

Fig. 12B shows that on application of an external MF to a solution of 9AA-HCl and HSA at a time delay of 0.4 μs after the laser flash at 355 nm, there is significant enhancement in absorbance in the wavelength regions around 530 nm and 570 nm implies the formation of Trp˙ and TrpH˙⁺ respectively and the peak at 570 nm implies for 9AA˙⁻. The radical ions which are initially formed in the triplet state diffuse to an optimum distance in the medium and when an external MF is applied there is reduction of ISC leading to an enhancement in the triplet yield of the respective radical ions.

Thus, PET is the prevalent reaction mechanism in both BSA and HSA systems and initial spin state of the precursors of PET is triplet. An exceptional condition arises in this case as MFE is obtained in homogeneous medium although confined medium provides the ideal condition to obtain MFE. In reality, a pseudoconfined medium is created by the complex structure of the protein that helps to sustain the optimum conditions required to observe appreciable MFE. It may so proposed that in case of BSA–9AA-HCl system the acceptor moiety of PET is both 9AAH⁺ and 9AA whereas, in case of HSA–9AA-HCl system PET in conjunction with ESPT occur. The redox potential values of 9AA-HCl and Trp moieties are −0.781 V and −1.015 V respectively^69,70 which justify the role of the 9AA-HCl as an electron acceptor and that of Trp as an electron donor in PET reactions. Therefore laser flash photolysis supports that extent of binding with 9AA-HCl for BSA is more than HSA because the BSA–9AA–HCl complex shows only PET in the excited-state whereas HSA–9AA–HCl complex shows both PET and ESPT due to flexible nature of HSA which facilitates TrpH˙⁺, the radical ion formed through PET, to donate further its proton to 9AA.

2.3.8. Theoretical docking study. Since the experimental pH is 7.4, we performed the docking studies with the single protonated form of 9AA-HCl i.e., 9AAH⁺. Both the serum albumins show 76.6% sequence identity. Their structures are also quite similar; however there is a root mean square backbone deviation of 2.944 Å as obtained from the structural alignment between 4L8U and 3V03. Similarities in structure and function indicate their similar ligand binding properties. Moreover, FRET experiments suggest that 9AAH⁺ binds close to the tryptophan in the serum albumins. HSA and BSA both have a tryptophan (Trp238 in HSA and Trp237 in BSA) meticulously positioned near the binding pocket in domain II, which is likely to be the potential binding site for 9AA-HCl. Therefore, 9AAH⁺ might bind to the known primary drug binding site at domain IIA of serum albumins.⁷¹ BSA has another tryptophan placed near the surface in domain I, however, large solvent exposure makes it less likely to be the binding site for a small molecule ligand. The binding site pockets on domain II was further analyzed by POCASA tool. The volume, surface area, sphericity and effective radius of the binding sites are listed in Table S1.† The binding site pocket in HSA is 1.7 times larger than the pocket in BSA. This larger pocket can account for the higher stoichiometry of ligand binding in HSA. Graphical representation of the pockets and their comparisons are shown in the Fig. 13. Fig. 14 shows the docking of 9AAH⁺ into the pockets of serum proteins using AutoDock Vina. Two molecules of 9AAH⁺ were docked into HSA because of the higher stoichiometry of binding which was observed in the experiments (Fig. 14). However, BSA has only one binding site (Fig. 14). The changes in SASA of different amino acid residues in binding site pocket due to ligand binding are shown in Fig. S4.† It indicates the residues that get solvent excluded upon ligand binging. Types of interactions with different amino acids are depicted in Fig. S5.† Table 6 depicts the docking results of HSA and BSA with 9AAH⁺ obtained from AutoDock Vina respectively. Distances from the nearest tryptophan residues are also listed in the Table 6. The binding energies computed by docking correlate very well with the spectroscopic data. However, the values of the distances between the drug and the proteins obtained from fluorescence energy transfer, where dynamic interactions of the interactive molecules in solvent are considered, may differ from those obtained from docking analyses, where the presence of the solvents molecules has been totally eliminated considering all the interacting molecules to be rigid with minimum energy conformation. 9AAH⁺ has a SASA of 341 Ų, which decreases by 89% and 96% upon binding to HSA and BSA, respectively (Table 6). Entropy gain due to solvent exclusion, therefore, plays an important role in these associations. Hydrophobic, hydrogen bonding, cation–pi, pi–pi and polar interactions are the major driving forces in the 9AAH⁺ and serum albumin interactions (Fig. S5†) which also supported the spectroscopic findings.

Fig. 13 The primary drug binding pockets in two serum albumins and their comparison; (A) binding pocket near Trp238 in the domain IIA of HSA. Protein is represented in ribbon and the shape of the binding site in mesh; (B) binding pocket near Trp237 in the domain IIA of BSA; (C) comparison of the binding pockets after alignment of the proteins. Binding site of HSA and BSA are colored in green and red, respectively. Four views at 90° rotations over z axis are shown.

Fig. 14 9AA-HCl docked complexes of serum albumins. Proteins are depicted in ribbon diagrams with N–C terminal was rainbow-colored. 9AA-HCl and Trp are shown in stick model. In the close up view the shape of the binding site pockets are shown in mesh. (A) 9AA-HCl–HSA complex with the close up view of binding site. Two possible binding modes are shown. Distances between the nearest atoms of the ligand and the Trp238 are shown in Å. (B) 9AA-HCl–BSA complex with the close up view of binding site. The nearest distance of the ligand from the Trp237 is shown in Å.

Table 6 Summary of the protein–ligand docking results obtained from AutoDock Vina

Binding modes	Binding parameters	AutoDock Vina
a When two ligands are bound.b Distance between the center of mass of two residues.c Distance between the nearest atoms of two residues as shown in Fig. 14.
HSA (binding mode 1 in domain II)	ΔG^0 (kJ mol^−1)	−29.29
	SASA of ligand (Å^2)	37.93/18.73^a
	Distance of 9AA-HCl from Trp238 (Å)^b	10.72
	Nearest distance of 9AA-HCl from Trp238 (Å)^c	6.1
HSA (binding mode 2 in domain II)	ΔG^0 (kJ mol^−1)	−27.20
	SASA of ligand (Å^2)	39.68
	Distance of 9AA-HCl from Trp238 (Å)^b	12.25
	Nearest distance of 9AA-HCl from Trp238 (Å)^c	6.6
BSA (domain II)	ΔG^0 (kJ mol^−1)	−28.03
	SASA of ligand (Å^2)	14.52
	Distance of 9AA-HCl from Trp237 (Å)^b	10.74
	Nearest distance of 9AA-HCl from Trp237 (Å)^c	7.9

---

2.3.9. Saturation transfer difference (STD) NMR studies. The STD NMR is an innovative screening process to reveal receptor–ligand interaction at an atomic resolution. This technique is used to recognize the orientation and mode of binding of chemical groups or residues of ligand those are in close proximity to the high molecular weight receptors, where ligands undergo a fast exchange between free and bound states at the NMR time scale.⁵⁹ Fluorescence experiments confirm that the binding of the ligand (9AA-HCl) to both proteins (HSA as well as BSA) is in the micromolar range, which motivated us to study STD NMR experiments. Fig. 15A and B depict chemical structure of 9AA-HCl and reference NMR spectrum of 9AA-HCl where all the non-exchangeable aromatic ring protons of acridine ring were clearly observed. In case of both serum proteins, BSA and HSA, the corresponding STD spectrum mimicking the reference spectrum demonstrates that the planar aromatic moiety of the molecule is well buried inside the protein cavity (Fig. 15C) and all four types of aromatic protons showed almost similar STD effects. However, differential STD effects were observed for both serum proteins. The intensity of the STD spectra in presence of BSA is slightly higher (1.4×) compared to HSA, which in turn suggests that 9AA-HCl has higher binding affinity towards BSA, supported by fluorescence experiments. In contrary, STD spectrum of the free 9AA-HCl does not show any peak which successfully probes the transfer of saturation from the protein to the ligand (Fig. 15D).

Fig. 15 (A) Chemical structure of 9AA-HCl molecule at pH 7.4 where amine group is protonated. (B) Reference ¹HNMR spectrum of 9AA-HCl showing different protons marked with different colors. (C) Overlay of STD NMR spectrum of 9AA-HCl in presence of BSA (blue) and HSA (red). The experiments have been performed at 298 K with 2 s of saturation time (on resonance = 1.0 ppm; off resonance = 40 ppm) with 1 : 450 protein: ligand ratio. (D) STD spectrum of 9AA-HCl alone (on resonance = 1.0 ppm; off resonance = 40 ppm) showing absence of any peaks.

Further we have plotted STD amplification factor (A_STD) vs. concentrations of 9AA-HCl used for both serum proteins for a better assessment of the binding phenomenon.⁶⁰ The STD amplification factor can be defined as follows

where I₀ is the intensity of the off-resonance spectrum and I_SAT is the intensity of the on-resonance spectrum. I_STD = I₀ − I_SAT is the intensity of STD NMR spectrum and L is the ligand concentration.

As discussed earlier, STD amplification factor also confirms the binding strength of the ligand 9AA-HCl to HSA or BSA are different in different serum proteins (Fig. S6A and B†).⁷² For BSA, the STD amplification factor reaches to a plateau at much earlier (∼700 μM) compared to the ligand binding to HSA (∼1200 μM). This effectively proves that the binding affinity is higher for BSA compared to HSA. Interestingly, the curve is monophasic in nature for binding of 9AA-HCl to BSA while it is biphasic for binding to HSA. It can be rationalized in terms of larger size of binding pocket for HSA (1.7 times) compared to BSA that can account more than one ligand molecule in the pocket which is also supported by fluorescence and docking analysis. Therefore, the STD NMR experiments effectively reveal not only differential binding affinity of the 9AA-HCl molecule to BSA and HSA, but also shed light on its binding modes. This result is in very good in agreement with the fluorescence as well as docking results.

3. Conclusion

In this article, the interactions of 9AA-HCl with serum proteins are explored thoroughly by using conventional spectroscopic tools as well as MF and also docking study. Although the generation of isosbestic points in absorption spectra evidences the formation of ground-state complexes between the 9AA-HCl and serum proteins, however the present study is an endeavor of the difference of specific binding modes of 9AA-HCl with serum proteins. The BSA is known to be a more rigid protein than HSA. Therefore, due to flexible nature the HSA accounts for both static and dynamic of quenching, whereas BSA shows only static quenching of Trp fluorescence in presence of 9AA-HCl. The Scatchard plot indicates that in case of BSA and 9AA-HCl the binding site is only one whereas the Hill plot shows that for HSA and 9AA-HCl more than one binding sites are present. The differences of interactions are also reflected in the thermodynamic parameters. In case of BSA–9AA-HCl system, the negative changes in both enthalpy and entropy values indicate that the key binding forces are hydrogen bonding and van der Waals. On the other hand, the positive thermodynamic parameters of HSA–9AA-HCl system decipher entropy driven nature of interaction. Laser flash photolysis studies eventually reveal that PET occurring from both the serum protein to 9AA-HCl and ESPT takes place only between HSA and 9AA-HCl. Theoretical docking studies also support the differential behavior of 9AA-HCl towards BSA and HSA. It reveals that the binding site pocket in HSA is 1.7 times larger than the pocket in BSA. This larger size of the pocket can account for the higher stoichiometry of ligand binding in HSA compared to BSA. These findings are also good in accordance with STD NMR experiments. Finally, spectroscopic studies corroborated with the docking analyses and STD NMR studies support the fact that although BSA and HSA have structural similarities, yet their mode of interactions with 9AA-HCl is quite different.

Acknowledgements

Financial assistance from the Chemical and Biophysical Approaches for Understanding Natural Processes (CBAUNP) 11-R&D-SIN-5.03-0100 and Biomolecular Assembly, Recognition and Dynamics (BARD) 12-R&D-SIN-5.04-0103 project, SINP of Department of Atomic Energy (DAE), Government of India is greatly acknowledged. P. Mitra acknowledges the Senior Research Fellowship from Council of Scientific and Industrial Research (CSIR) 09/489(0090)/2011-EMR-I, India. We would like to acknowledge our thanks to Mr Ajay Das SINP, Kolkata for his assistance in fluorescence lifetime measurements. We sincerely thank to Mrs Chitra Raha for her kind assistance and technical support.

References

1. W. R. Harris, Coord. Chem. Rev., 1996, 149, 347–365.
2. (a) P. B. Kandagal, S. Ashoka, J. Seetharamappa, S. M. T. Shaikh, Y. Jadegoud and O. B. Ijare, J. Pharm. Biomed. Anal., 2006, 41, 393–399; (b) S. M. T. Shaikh, J. Seetharamappa, S. Ashoka and P. B. Kandagal, Dyes Pigm., 2007, 73, 211–216.
3. J. L. Irvin and E. M. Irvin, J. Biol. Chem., 1952, 196, 651–668.
4. Y. J. Hu, Y. Liu, R. M. Zhao, J. X. Dong and S. S. Qu, J. Photochem. Photobiol., A, 2006, 179, 324–329.
5. Y. Q. Wang, H. M. Zhang, G. C. Zhang, W. H. Tao and S. H. Tang, J. Mol. Struct., 2007, 830, 40–45.
6. Y. Q. Wang, H. M. Zhang, G. C. Zhang, W. H. Tao, Z. H. Fei and Z. T. Liu, J. Pharm. Biomed. Anal., 2007, 43, 1869–1875.
7. A. Sułkowska, J. Mol. Struct., 2002, 614, 227–232.
8. K. Aidas, J. M. H. Olsen, J. Kongsted and H. Ågren, J. Phys. Chem. B, 2013, 117, 2069–2080.
9. A. Ray, B. Koley Seth, U. Pal and S. Basu, Spectrochim. Acta, Part A, 2012, 92, 164–174.
10. B. Chakraborty and S. Basu, J. Lumin., 2009, 129, 34–39.
11. B. Chakraborty, A. Singha Roy, S. Dasgupta and S. Basu, J. Phys. Chem. A, 2010, 114, 13313–13325.
12. M. Banerjee, U. Pal, A. Subudhhi, A. Chakrabarti and S. Basu, J. Photochem. Photobiol., B, 2012, 108, 23–33.
13. P. Mitra, M. Banerjee, S. Biswas and S. Basu, J. Photochem. Photobiol., B, 2013, 121, 46–56.
14. X. M. He and D. C. Carter, Nature, 1992, 358, 209–215.
15. D. C. Carter and X. M. He, Science, 1990, 249, 302–303.
16. D. C. Carter and J. X. Ho, Adv. Protein Chem., 1994, 45, 153–203.
17. M. Dockal, D. C. Carter and F. Ruker, J. Biol. Chem., 2000, 275, 3042–3050.
18. D. Charbonneau, M. Beauregard and H. A. Tajmir-Riahi, J. Phys. Chem. B, 2009, 113, 1777–1784.
19. M. D. Topal, Biochemistry, 1984, 23, 2367–2372.
20. D. O. Jordan and L. N. Sansom, Biopolymers, 1971, 10, 399–410.
21. C. Rehn and U. Pindur, Monatsh. Chem., 1996, 127, 645–658.
22. E. L. Loechler and J. King, Biochemistry, 1986, 25, 5858–5864.
23. T. Bentin and P. E. Nielsen, J. Am. Chem. Soc., 2003, 125, 6378–6379.
24. A. I. Kononov, J. Phys. Chem. B, 2001, 105, 535–541.
25. W. E. Lee and W. G. Galley, Biophys. J., 1988, 54, 627–635.
26. Y. F. Long, C. Z. Huang and Y. F. Li, J. Phys. Chem. B, 2007, 111, 4535–4538.
27. Y. A. Hannun and R. M. Bell, J. Biol. Chem., 1988, 263, 5124–5131.
28. G. P. Sachdev, A. D. Brownstein and J. S. Fruton, J. Biol. Chem., 1974, 249, 420–427.
29. S. I. Bae, R. Zhao and R. M. Snapka, Biochem. Pharmacol., 2008, 76, 1653–1668.
30. E. L. Loechler and J. King, Biochemistry, 1986, 25, 5858–5864.
31. H. Schmidt, A. Al-Ibrahim, U. L. B. Dietzel and L. Biker, Photochem. Photobiol., 1981, 33, 127–130.
32. S. Grzesiek, H. Otto and N. A. Dencher, Biophys. J., 1989, 55, 1101–1109.
33. P. Gangola, N. B. Joshi and D. D. Pant, Chem. Phys. Lett., 1981, 80, 418–421.
34. D. D. Pant, G. C. Joshi and H. B. Tripathi, Pramana-Journal of Physics, 1986, 27, 161–170.
35. A. Murza, S. Alvarez-Méndez, S. Sanchez-Cortes and J. V. García-Ramos, Biopolymers, 2003, 72, 174–184.
36. A. Murza, S. Sánchez-Cortés, J. V. García-Ramos, J. M. Guisan, C. Alfonso and G. Rivas, Biochemistry, 2000, 39, 10557–10565.
37. A. Murza, S. Sánchez-Cortés and J. V. García-Ramos, Biospectroscopy, 1998, 4, 327–339.
38. P. Mitra, B. Chakraborty, D. Bhattacharyya and S. Basu, J. Phys. Chem. A, 2013, 117, 1428–1438.
39. P. Mitra, B. Chakraborty and S. Basu, J. Lumin., 2014, 149, 221–230.
40. P. Mitra, B. Chakraborty and S. Basu, Chem. Phys. Lett., 2014, 610–611, 108–114.
41. P. Mitra, P. K. Chakraborty, P. Saha, P. Ray and S. Basu, Int. J. Pharm., 2014, 473, 636–643.
42. J. R. Lakowicz, Principles of Fluorescence Spectroscopy, Plenum Press, New York, 2006.
43. T. K. Maiti, K. S. Ghosh, A. Samanta and S. Dasgupta, J. Photochem. Photobiol., A, 2008, 194, 297–307.
44. T. Imoto, L. S. Forster, J. A. Rupley and F. Tanaka, Proc. Natl. Acad. Sci. U. S. A., 1971, 69, 1151–1155.
45. M. Mondal and A. Chakrabarti, FEBS Lett., 2002, 532, 396–400.
46. J. Min, X. Meng-Xia, Z. Dong, L. Yuan, L. Xiao-Yu and C. Xing, J. Mol. Struct., 2004, 692, 71–80.
47. S. Banerjee, S. Dutta Choudhury, S. Dasgupta and S. Basu, J. Lumin., 2008, 128, 437–444.
48. T. Barri, T. Trtić-Petrović, M. Karlsson and J. Åke Jönsson, J. Pharm. Biomed. Anal., 2008, 48, 49–56.
49. K. Kitamura, H. Mano, Y. Shimamoto, Y. Tadokoro, K. Tsuruta and S. Kitagawa, Fresenius. J. Anal. Chem., 1997, 358, 509–513.
50. J. A. Goodrich and J. F. Kugel, Binding and kinetics for Molecular Biologist, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 2007.
51. H. Gao, L. Lei, J. Liu, Q. Kong, X. Chen and Z. Hu, J. Photochem. Photobiol., A, 2004, 167, 213–221.
52. R. J. Read, P. D. Adams, W. B. Arendall, A. T. Brunger, P. Emsley, R. P. Joosten, G. J. Kleywegt, E. B. Krissinel, T. Lütteke, Z. Otwinowski, A. Perrakis, J. S. Richardson, W. H. Sheffler, J. L. Smith, I. J. Tickle, G. Vriend and P. H. Zwart, Structure, 2011, 19, 1395–1412.
53. C. Azuara, E. Lindahl, P. Koehl, H. Orland and M. Delarue, Nucleic Acids Res., 2006, 34, W38–W42.
54. J. Yu, Y. Zhou, I. Tanaka and M. Yao, Bioinformatics, 2010, 26, 46–52.
55. N. R. Voss and M. Gerstein, Nucleic Acids Res., 2010, 38, W555–W562.
56. O. Trott and A. J. Olson, J. Comput. Chem., 2010, 31, 455–461.
57. M. Gerstein, Acta Crystallogr., Sect. A: Found. Crystallogr., 1992, 48, 271–276.
58. M. Mayer and B. Meyer, Angew. Chem., Int. Ed., 1999, 38, 1784–1788.
59. V. Banerjee, R. K. Kar, A. Datta, K. Parthasarathi, S. Chatterjee, K. P. Das and A. Bhunia, PLoS One, 2013, 8, 72318.
60. A. Bhunia, S. Bhattacharjya and S. Chatterjee, Drug Discovery Today, 2012, 17, 505–513.
61. U. Kragh-Hansen, Pharmacol. Rev., 1981, 33, 17–33.
62. B. Valeur and J. C. Brochon, New Trends in Fluorescence Spectroscopy, Springer, Berlin, 2001.
63. G. M. Morris, D. S. Goodsell, R. S. Halliday, R. Huey, W. E. Hart, R. K. Belew and A. J. Olson, J. Comput. Chem., 1998, 19, 1639–1662.
64. M. Belletete, M. Lachapelle and G. Durocher, J. Phys. Chem., 1990, 94, 7642–7648.
65. D. Bose, D. Sarkar and N. Chattopadhyay, Photochem. Photobiol., 2010, 86, 538–544.
66. J. Gonzalez-Jimenez and M. Cortijo, J. Protein Chem., 2002, 21, 75–79.
67. D. V. Bent and E. Hayon, J. Am. Chem. Soc., 1975, 97, 2612–2619.
68. D. Creed, Photochem. Photobiol., 1984, 39, 537–562.
69. T. Uematsu, T. Waki, T. Torimoto and S. Kuwabata, J. Phys. Chem. C, 2009, 113, 21621.
70. A. Harriman, J. Phys. Chem., 1987, 91, 6102–6104.
71. A. J. Ryan, J. Ghuman, P. A. Zunszain, C.-W. Chung and S. Curry, J. Struct. Biol., 2011, 174, 84–91.
72. M. Mayer and B. Meyer, J. Am. Chem. Soc., 2001, 123, 6108–6117.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra04140j

---