Separate and simultaneous binding of tamoxifen and

Journal of Molecular Liquids 249 (2018) 1083–1096

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Journal of Molecular Liquids journal homepage: www.elsevier.com/locate/molliq

Separate and simultaneous binding of tamoxifen and estradiol to human serum albumin: Spectroscopic and molecular modeling investigations Nastaran Moradi a,b, Mohammad Reza Ashrafi-Kooshk a,⁎, Jamshidkhan Chamani b,⁎⁎, Dareuosh Shackebaei a, Fatemeh Norouzi c a b c

Medical Biology Research Center, Kermanshah University of Medical Sciences, Kermanshah, Iran Department of Biochemistry and Biophysics, Faculty of Sciences, Mashhad Branch, Islamic Azad University, Mashhad, Iran Department of Genetics, Reproductive Biomedicine Research Center, Royan Institute for Reproductive Biomedicine, ACECR, Tehran, Iran

a r t i c l e

i n f o

Article history: Received 31 May 2017 Received in revised form 1 September 2017 Accepted 7 November 2017 Available online 10 November 2017 Keywords: Human serum albumin Tamoxifen Estradiol Binding Spectroscopy Molecular modeling

a b s t r a c t In the recent years, tamoxifen has been often co-administrated with estradiol to treatment of breast cancer. Hence, we examined the interaction of two anti-breast cancer drugs, tamoxifen and estradiol, with human serum albumin (HSA) using spectroscopic and molecular modeling techniques. The quenching of intrinsic fluorescence and docking were used to calculate binding affinity and location, while the RLS, synchronous fluorescence and CD spectroscopy were done to investigate the effect of these drugs on protein conformational changes, both separately and simultaneously. While the photo-stability of drugs increased in the presence HSA, the enhancement of RLS intensity was attributed to the formation of a new complex between the two drugs and the protein. Both drugs showed a strong ability to quench the fluorescence of HSA via static mechanism. The fluorescence quenching action at different temperatures and enhancement of RLS intensity were much stronger when the two drugs co-existed, whereas the binding affinity of both drugs were partially affected by each other, which demonstrate that two drugs bind to distinct/different location on HSA via hydrophobic interactions. On the other hand, appearance of new peak at 405 nm accompanying with quenching of HSA fluorescence at 340 nm by tamoxifen revealed that the drug binds near the protein Trp residue in subdomain IIA, which confirmed the docking results. Furthermore, synchronous fluorescence and CD data clearly indicate that the conformation of HSA was changed upon addition of the drugs which can affect the physiological functions of HSA and binding of other drugs. © 2017 Elsevier B.V. All rights reserved.

1. Introduction Proteins are important chemical substances in our life and the main target of many drugs in different organisms. Human serum albumin (HSA), lipoprotein, glycoprotein, α, β, and γ globulins are the main drug carrier proteins [1]. From a biopharmaceutical point of view drugs-serum proteins interaction are particularly important, because the free form of the drugs or the pharmacologically active form directly relate to their affinities to serum proteins. Binding of a drug to albumin, results in an increased drug solubility in plasma, decreased toxicity, and protection against oxidation of the bound drug [2]. In some pathogenic states of albumins, weaker interactions lead to an increase of drug concentration in the blood serum, which in turn can cause toxic poisoning,

⁎ Correspondence to: M. R. Ashrafi-Kooshk, Medical Biology Research Center, P. O. Box 67155-1616, Kermanshah University of Medical Sciences, Kermanshah, Iran. ⁎⁎ Corresponding author. E-mail addresses: mrashrafi@kums.ac.ir (M.R. Ashrafi-Kooshk), [email protected] (J. Chamani).

https://doi.org/10.1016/j.molliq.2017.11.056 0167-7322/© 2017 Elsevier B.V. All rights reserved.

or even death. Therefore, knowledge of a drug's binding to the albumin is very useful in therapeutic drug monitoring. Most of the protein–ligand binding research is primarily focused on investigating the interactions of various natural and synthetic chemicals with HSA, their binding site mapping and characterization [3,4]. Investigating the interaction of drugs with HSA can elucidate the properties of drug–protein complexes, as it may provide useful information of the structural features that determines the therapeutic effectiveness of drugs [5]. Serum albumins, which are abundant in plasma, are the most widely studied proteins in the circulatory system of a wide variety of organisms. Structural aspects and properties of these transport proteins have been well explored. HSA is a globular protein consisting of a single polypeptide chain with 585 amino acids, but only one tryptophan residue (Trp214) [6]. The main binding sites of aromatic and heterocyclic ligands on HSA are within two hydrophobic pockets in subdomains IIA and IIIA, namely Sudlow's sites I and II [7]. Sudlow's site I is major importance, as it contains the Trp214 residue [8,9]. This protein is also responsible for the regulation of colloidal osmotic pressure, maintenance of blood pH, and is a possible source of amino acids for different tissues

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N. Moradi et al. / Journal of Molecular Liquids 249 (2018) 1083–1096

[10]. HSA binding can extend the metabolic half-life of a compound, providing a convenient way in which to extend the duration of activity. One of the most important biological functions of serum albumin is its ability to reversibly bind to a large variety of endogenous and exogenous ligands, such as fatty acids, drugs, and metal ions in the blood stream [1,6,11–16]. Estradiol (EST) is a sex hormone (estra-1, 3,5,(10)-triene-3, 17- βdiol) which has two hydroxyl groups in its molecular structure (Scheme 1A). Estradiol is using for the treatment of hypoestrogenism, advanced androgen-dependent carcinoma of the prostate, moderate to severe vasomotor symptoms associated with menopause and is also used to prevent osteoporosis [17]. In plasma, EST is largely bound to sex hormone-binding globulin, also to albumin. Its cellular functions were initiated by binding to estrogen receptor. It has been shown long-term consumption of estradiol can raise risk of breast cancer. One of the several antagonists that use for treatment of hormone-positive breast cancers is tamoxifen (TMX). Tamoxifen (2-[4-[(Z)-1,2di(phenyl)but-1- enyl]phenoxy]-N,N-dimethylethanamine) is a nonsteroidal anti-estrogen drug, which for the past 20 years has been widely used to prevent and treat breast, liver, pancreas and brain cancers (Scheme 1B) [18]. TMX is an antagonist of the estrogen receptor in breast tissue. It has been the endocrine therapy for hormone-positive early breast cancer in post-menopausal women, although aromatase inhibitors have been proposed [19]. Lum et al. observed that the longterm tamoxifen therapy can be associated with increased serum levels of estrogen hormones especially EST [20]. A decade later, Gjerde et al. indicated that there are the significant correlations between the serum concentrations of tamoxifen, N-dedimethyltamoxifen, and tamoxifen-N-oxide and estrogens (P b 0.05) [21]. They proposed that the genotype predicted CYP2C19 activity influenced the levels of both tamoxifen metabolites and EST [22]. However, since similar to EST, TMX could bind to HSA, be effectively transported and eliminated in body, the effect of competition between TMX and EST for one drug binding site on HSA, which influences their serum concentrations, has not been studied. Thus, we analyzed of the interaction between TMX and EST with HSA as binary and ternary systems constitutes the continuation of a study of HSA–drug binding in multidrug therapy. In this study, spectroscopic evidences regarding the drug-binding mode, drug binding constant and protein structural change of both anticancer drugs has been investigated combined with molecular modeling. 2. Experimental 2.1. Materials Essentially free fatty acid HSA (fraction V), TMX, and EST were all purchased from Sigma-Aldrich (St. Louis, MO, USA) and used without further purifications. Double-distilled water with very low conductivity was used throughout the experiments. Due to sensitivity of TMX and EST to the light, all solutions were stored in a refrigerator a 4 °C in the

dark (bottles). Due to poor solubility of EST and TMX in water, the stock solutions of TMX and EST (3 × 10−3 M) were prepared in DMSO and diluted to 3 × 10−4 M (1:10) in a phosphate buffer 50 mM pH 7.4 prior to use. The final concentration of DMSO in all experiments are below 0.5% that has any significant alteration in HSA structure/conformation and function. Also, 10 mg of HSA was gently dissolved in 1 mL of phosphate buffer 50 mM pH 7.4 (~1.5 × 10−4 M). The exact concentration of HSA in a stock solution was determined spectrophotometrically using molar extinction coefficient of 35,700 M−1·cm−1, and the molecular weight of 66.4 kDa [23]. The adequate amount of HSA stock (~ 13.5 μl) was added to a 1.0 cm quartz cell to make up 2 mL, 1 μM, HSA solution and the range of the drug solution was progressively titrated physically into the cell using a micro-injector for the binary and ternary systems. In all titration experiments, the dilution factor of the ligand titration was corrected. 2.2. Methods 2.2.1. Drug stability experiments Experiments were performed with a self-designed and self-made photochemical reactor equipped with one UV lamp with an intensity of 200 μW/cm2. A temperature of 25 ± 1 °C was maintained for reactions. The aqueous solutions of EST, TMX, their complexes with HSA and the protein solution as a control were simultaneously placed in the reactor for 120 h (5 days). The concentrations of EST, TMX and HSA were 30 μM. Aliquots from solutions were collected every 6 h for the first day and then every 24 h. The aliquots were centrifuged and their absorbance at 275 nm were recorded to follow photodegradation of drugs in the absence and the presence of the protein [24–27]. Also, because a part of decrement in absorption is due to degradation or precipitation of the protein, the absorption difference of the control was added to the absorbance of HSA-drug complexes. 2.2.2. Resonance light scattering (RLS) Fluorescence measurements were carried out on a spectrofluorometer Model F-2500 (Hitachi, Japing) linked to a personal computer and equipped with a 150-W Xenon lamp, a 1.0 cm quartz cell and a thermostat bath. For RLS measurement, the excitation and emission monochromators were scanned simultaneously, with Δλ = 0 nm from 220 nm to 600 nm with slit widths of 5 nm for the excitation and emission with the same device used for fluorescence measurements. A 2 mL solution containing 1 μM HSA was injected successively by 40 aliquots of drug solutions to obtain a final concentration of drugs ranging from 0 to 5.4 μM. For the ternary systems, an identical procedure was carried out with pre-incubation of HSA with 1 μM interferer drug. 2.2.3. Circular dichroism (CD) spectroscopy Far-UV CD spectra in the binary and ternary systems were carried out on a Jasco-815 spectropolarimeter (Jasco, Tokyo, Japan) equipped with a Jasco 2-syringe titrator under constant N2 flush at room

Scheme 1. The chemical structures of estradiol (A) and tamoxifen (B).


temperature. Dry nitrogen gas was used to purge the testing environment before and during the measurements. Also using a quartz cuvette having bandwidth of 1 nm was utilized and the scanning speed was 100 nm·min−1. The instrument was calibrated with ammonium d-10camphor sulfuric acid. The induced ellipticity was achieved by subtracting the ellipticity of the drug from that of the drug-HSA mixture at the same wavelength. The results are expressed as the mean residue ellipticity [θ], which is defined as [θ] = 100 × θobsd/(LC), where θobsd is the observed ellipticity in degrees, C is the concentration in residue mol·cm−3, and L is the length of the light path in cm. The samples for CD analysis were prepared with a fixed concentration of HSA (1.5 μM) and varying drug concentrations (0, 1.5, 3 and 6 μM) resulting in equal volumes. For the ternary systems, HSA solution was pre-incubated with 1.5 μM interferer drug. 2.2.4. Synchronous fluorescence spectroscopy Information on the molecular environment in the proximity of the Trp and Tyr fluorophores of HSA in the case of binary and ternary complexes was acquired by this technique, which suggests the assists of spectral simplification and bandwidth reduction. Synchronous fluorescence spectra were acquired by concurrently scanning the excitation and emission monochromators by spectrofluorometer. Synchronous fluorescence spectra are characteristics of the Tyr and Trp residues of HSA when the wavelength interval (Δλ) is 15 nm and 60 nm, relatively [28]. A 2 mL solution containing 1 μM HSA was injected successively by 40 aliquots of drug solutions to obtain a final concentration of drugs ranging from 0 to 8.9 μM. For the ternary systems, an identical procedure was carried out with pre-incubation of HSA with 1 μM interferer drug. 2.2.5. Fluorescence quenching measurements The fluorescence intensity of a protein can be made weak by a variety of molecular interactions, which is called fluorescence quenching. Fluorescence measurements were carried out on the Hitachi spectrofluorometer equipped with a 1.0 cm quartz cell and a thermostat bath. The widths of excitation and emission slit were set at 5 nm. The protein concentration was in all our experiments kept constant 1 μM and the concentrations of the quenchers were different (ranging from 0 to 8.5 μM). For the ternary systems, HSA solution was pre-incubated with 1 μM interferer drug. The fluorescence spectra were measured with an excitation wavelength at 280 nm and 295 nm, and an emission wavelength of 300–600 nm at 280, 290, 300 and 310 K. To decline the inner filter effect, the fluorescence intensities used in the present study were corrected for absorption of the excitation light and re-absorption of emitted light using the formula: Fcor = Fobs × e(Aex + Aem)/2 where Fcor and Fobs are the corrected fluorescence intensity and observed fluorescence intensity, relatively, and Aex and Aem are the absorption of the system at the excitation wavelength and at the emission wavelength, respectively [29]. To analyses of quenching mechanism and accessibility of Trp residues in proteins, the Stern–Volmer (Eq. 1) and modified Stern–Volmer (Eq. 2) equations were used respectively [13,29,30]. F0 ¼ 1 þ K SV ½Q ¼ 1 þ kq τ0 ½Q F

ð1Þ

where F0 and F are the fluorescence intensities of HSA in the absence and the presence of quencher, respectively. kq, Ksv, τ0 and [Q] are the quenching rate constant of the biomolecule, the Stern–Volmer dynamic quenching constant, average life-time of the biomolecule without quencher (10−8 s−1) and the concentration of the quencher, respectively [8,31]. F0 1 1 ¼ þ F 0 − F f a K Q ½Q f a

ð2Þ

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where KQ is the effective quenching constant and fa is the fraction of initial fluorescence that is accessible to the quencher (fa = Fa/(Fa + Fb)); a and b refer to the accessible and inaccessible or buried fluorophore, respectively [30]. 2.2.6. Binding analyses The analysis of the dependence of fluorescence intensity of HSA on the EST/TMX concentration can be carried out according to the earlier reports [32,33] to determine of ligand binding affinity. Knowing n value from previous reports [34,35], the fluorescence quenching data were fitted to Eq. 3 to obtain the dissociation constant of ligand binding site (Kdiss) [33]. Eq. 3 is the most generally valid equation to analyze fluorescence changes upon formation of a 1:1 complex [36]. qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ffi 2 F F lim n½P0 þ ½L0 þ K diss − n½P0 þ ½L0 þ K diss −4n½P0 ½L0 ¼ 1− 1− F0 F0 2n½P0

ð3Þ

2.2.7. Thermodynamic analysis of the binding process In order to explain the nature of the interaction between drugs and HSA in the binary and ternary systems, the dissociation constant of HSA-drugs complexes was determined at four different temperatures. The thermodynamic parameters of binding, entropy change (ΔS°) and enthalpy change (ΔH°), were obtained from van't Hoff equation: ln

1

K diss

¼

−ΔH∘ ΔS∘ þ RT R

ð4Þ

where Kdiss is the dissociation constant at the given temperature and R is the universal gas constant (8.314 J·mol−1·K−1) [37]. The values of ΔS° and ΔH° can be obtained from the intercept and slope of the van't Hoff plot, respectively, and the free energy change (ΔG°) can be estimated by using the Gibbs equation [8]: ΔG∘ ¼ ΔH ∘ −TΔS∘

ð5Þ

2.2.8. Molecular docking simulation The AutoDock Vina 1.1.2 docking package was used for ligand flexible docking simulations using the default settings [38]. The structures of estradiol and tamoxifen were generated by HyperChem Professional 8.0 [39]. Then, their configurations were imposed to semi-empirical method AM1 with Polak-Ribiere algorithm to maintain their energy minimized structures. Prior to docking, the ligands were optimized with Gasteiger-Hückel charges [40] by the package of MGLTools [41,42]. Also, the three-dimensional (3D) coordinate of the HSA (PDB ID: 1AO6) was downloaded from the Protein Data Bank (http://www. rcsb.org/pdb) [43]. The pdbqt files of ligands and receptor were generated using the MGLTools. The pdbqt file is a modified protein data bank file format containing the information of atomic charges, atom type definitions and, for ligands, topological information. The size of the docking grid was kept as 126 Å × 126 Å × 126 Å and the grid spacing set at 0.6 Å. The docking studies were performed by AutoDock Vina using a genetic algorithm [44]. Default docking parameters were used. AutoDock Vina has reported high accuracy in predicting binding free energies by setting the receptor rigid while appraising flexible ligands, with a comparatively low standard error [38]. Therefore, HSA conformational flexibility was neglected by rigid receptor docking. The interactions of the ligands with protein were visualized in VMD 1.9.2 (www.ks.uiuc.edu) and LIGPLOT+ v 1.4 (www.ebi.ac.uk) [45,46]. The binding energy was obtained for each ligand. The lowest binding energy conformation out of ten generated binding modes was considered as the most favourable docking pose.

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3. Results and discussion 3.1. Drug photostability measurements Estrogens and estrogen modifiers have photosensitive phenolic structures and can be efficiently reduced by photolysis [24,47]. Liu et al. note that UVC can rapidly degrade E1 by direct photolysis, in which the estrogen molecules absorb photon energy, transit to a high level, and then undergo irreversible reactions that result in molecular bond fractures, etc. [48]. Also, Mboula et al. illustrate that EST under photocatalytic degradation formed several intermediates with estrogenic activity which the phenol group of EST is still present in their structures but with OH groups in different positions [26]. On the other hand, photodegradation of TMX was studied by Salamoun et al. and DellaGreca et al. [24,49]. Salamoun et al. reported that during the UV irradiation of TMX, isomerization of the trans to the cis isomer takes place and consequently, corresponding highly fluorescent phenanthrene derivatives are formed [49]. The structures of photoproducts were identified by HPLC, GC–MS, 1H NMR spectroscopy and LC-MS. DellaGreca et al. studied the photodegradation of TMX in water by prolonged exposure to sunlight irradiation [24]. The main photoproducts, have been identified by spectroscopic means, where photoisomerization, photocyclization and, to a lesser extent, photooxygenation appear to be involved in the drug degradation [27]. Since EST and TMX have a strong UV bands at 250–300 nm, the degradation process can be analyzed by UV–Vis spectrophotometer at 275 nm [24,25,27,50]. Fig. 1A and B show the remained intact drug in the solution in the absence and the presence of HSA during the 120 h period. Under UV radiation with or without HSA, the degradation of drugs occurs at different rates. When HSA was absent, EST and TMX were efficiently photolyzed by UV light, whereas in the presence of protein, the degradation process is slowed down. Fig. 1A and B indicate the fast degradation of TMX and EST in the absence of HSA with a conversion of 66% and 42% after 120 h of radiation, respectively; however, only 19% and 25% of TMX and EST were photodegraded in the presence of the protein, respectively. The faster degradation of TMX can be explained by its much stronger absorption of photons in the UVC region of the electromagnetic spectrum. These results may suggest that the albumin can be used as a stabilizing agent in the formulation of injectable forms of EST and TMX. 3.2. Resonance light scattering (RLS) measurements During recent years light-scattering has been widely utilized to study of the size, shape and aggregation of biological and chemical

species [51,52]. RLS studies are using to monitor molecular gather and characterize the extended aggregates of chromophores. Pasternack et al. first developed the RLS technique for studying the aggregation of porphyrins on the surface of nucleic acids with a common fluorimeter [53]. RLS can be improved and has confirmed to be able to examine the aggregation of small molecules and also the long-range gathering of drugs on biological patterns. This technique is ready for utilize to prepare for in advance some insight into processes that are certain for the formation of complexes. When interaction happens between two molecules, the structural firmness depends on the solubility in suitable solvents, the interacting molecules' sizes, their symmetry and their charges. Since the manufacture of RLS is associated with the formation of specific aggregates, the RLS intensity is mainly dominated by the particle dimensions of the formed aggregate in solution. The RLS spectra of the protein were studied in the absence and presence of increasing concentrations of quenchers (EST and TMX). The results for the binary and ternary systems of EST and TMX are indicated in Fig. 2. As can be seen, the intensity of the RLS spectra were clearly enhanced with the increase of the drug concentration for the binary and ternary systems. It is known that RLS enhancement could be the result of enlargement of the molecular volume of the complexes. So, it was suggested that aggregation is principally operated by an interaction between HSA and drugs. The similar results were obtained by Danesh et al. and Pourgonabadi et al. for the interaction of EST and TMX with HSA, respectively [54,55]. Furthermore, Fig. 3 compares the values of ΔIRLS plotted against various concentrations of the drugs in the binary and ternary systems. It was found that the increase of the RLS intensity differed for various concentrations of the HSA–drug solution and there was a non-linear relationship between the enhanced intensity and the concentration of the drug. The RLS intensity of the systems progressively increased with increasing drug concentrations in all systems and acceleration happen in the solutions that included high concentrations of drug. Also, it can be seen from Fig. 3 that the RLS intensity in both ternary systems was much higher as compared to their binary complement, which exhibit the formation of a larger complex in the three-component systems form complexes and aggregates on the HSA surface. This suggests that the two drugs can bind to HSA simultaneously. The intersection of a horizontal and a slanting line at high concentrations of drugs yields the critical induced-aggregation concentration of drug-induced protein aggregation (CCIAC value) [56]. Under corresponding experimental conditions, a smaller CCIAC value signified a smaller concentration of drug-induced protein aggregation. It is important to note that, smaller CCIAC values testified to a higher affinity to create aggregates. Due to a greater interaction between the drug and HSA in the

Fig. 1. Degradation kinetics of EST (A) and TMX (B) in the absence (●) and the presence of HSA (○) under UV radiation during 120 h. The degradation process monitored by decrement in absorbance at 275 nm that normalized to initial absorbance. The concentration of EST, TMX and HSA in aqueous solution are 30 μM.


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Fig. 2. RLS spectra of HSA in the HSA–EST (A), [HSA–TMX]–EST (B), HSA–TMX (C) and [HSA–EST]–TMX systems (D). The concentration of drugs were progressively increased from 0 to 5.3 μM. For the ternary systems, 1 μM HSA solution was pre-incubated with 1 μM interferer drug.

presence of the other drug, the CCIAC values for both ternary systems were smaller than for the binary ones. So that the CCIAC values for EST and TMX were decreased from 1.4 and 1.2 to 0.3 and 0.4, respectively. 3.3. CD spectroscopy analysis It is known that binding of ligand to a globular protein can alter its secondary structure, which results in changes in the protein's conformation. These alterations are reflected in their circular dichroism (CD) spectra [57]. To get a clear image of how the structure of HSA was

affected by drugs binding, CD measurements on free HSA, as well as binary and ternary systems were performed. Changes in CD spectra can provide information on ligand–protein interactions [58]. Secondary structural elements, such as α-helix, β-sheets, β-turns and random coil structures, all induce bands of distinctive shapes and magnitudes in the far ultraviolet region [59]. To ascertain the possible influence of the two anti-cancer drugs binding on the secondary structure of HSA, we performed far-UV CD studies in the presence of different concentrations of drugs at pH 7.4 and room temperature. As shown in Fig. 4, the CD spectra of HSA had two negative bands in the far-UV region at 208

Fig. 3. RLS enhancement of the (A) HSA–EST (●) and [HSA–TMX]–EST (○) and (B) HSA-TMX (●) and [HSA-EST]-TMX (○) systems as a function of drug concentration. Points are the experimental data.

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amino acid residues of the main polypeptide chain of HSA and that the interaction destroyed the original hydrogen-bonding networks. This conclusion is consistent with the results of the synchronous fluorescence spectroscopic study pointing at the microenvironment and secondary structure undergoing some changes but to a limited extent. A notable point was that the structural changes of the α-helixes, βsheets, turns and unordered coils were more pronounced for the ternary systems as opposed to for their binary counterparts (from 53.97% to 49.66% in the [HSA–TMX]–EST system, and to 51.43% in the [HSA– EST]–TMX system). Therefore, the presence of EST or TMX in the ternary systems led to a more significant decrease in the secondary structure stabilization of HSA. To sum up, the results revealed that both anti-cancer drugs had an effect on the physiological function of HSA by altering its conformation and destabilization of the protein's secondary structure. The alteration of HSA secondary structure upon drugs binding is not as great as one was reported by Bourassa et al. (2011), Bourassa et al. (2011) and Bourassa et al. (2016) which may be due to significant difference in the ratio of TMX to HSA concentration [60–62]. In these papers the concentration of TMX is 10-fold higher than the protein concentration, whereas in the present study and the other reports such as Pourgonabadi et al., Sharif-Barfeh et al. and Amani et al. with the similar CD results, the ratio of HSA to drug is close to 1 [55,63,64]. However according to Imoto (1997) and Robic (2010), the negative changes in the Gibbs free energy upon drugs binding accompanying with the minor changes in the protein secondary structure might imply that the binding of the drugs stabilize the protein conformation [65,66]. Fig. 4. Far-UV CD spectra of HSA in the absence and presence of different concentrations of EST (A) and TMX (B) in the binary system. 1.5 μM of HSA was incubated for 30 min with 0 (1), 1.5 (2), 3 (3) and 6 μM drugs (4). Conditions: T = 298 K, pH = 7.4.

and 222 nm, which are typical of α-helix structure in the protein [58, 59]. If the α-helixes were to change, the spectrum would change accordingly. The addition of drugs to HSA in both the two- and the three component systems caused a decrease in the negative ellipticity in the region of far-UV CD in a concentration-dependent manner, without any significant shift of the peaks. This was believed to be the result of the formation of complexes between HSA and the drugs. However, the similarity between the shapes of the CD spectra pertaining to HSA in the presence and the absence of the drugs in all interacting systems suggested that the structure of HSA was still predominantly α-helical. Table 1 lists the fractions of α-helix, β-sheet, turn and unordered coil, and as can be seen, the pure HSA comprised 53.97% α-helix, 18.31% β-sheet, 13.47% turn and 14.25% unordered coil. The calculated results exhibited a reduction of the α-helical content from 53.97% to 51.48% in the HSA–EST system and to 51.8% in the HSA–TMX system, whereas those of unordered coil increased, thereby leading to a more random structure in both the binary and the ternary systems. Furthermore, the content of the β-sheet and turn slightly decreased on the addition of larger amounts of drug. Also, the percentage of α-helix structure in HSA decreased, indicating that both drugs bound to the

Table 1 Analysis of secondary structure of HSA and the HSA-drugs binary and ternary systems from CD data. System

Helix %

Sheet %

Turn %

Random coil %

HSA HSA-EST (HSA-TMX)-EST HSA-TMX (HSA-EST)-TMX

53.97 51.48 49.66 51.80 51.43

18.31 18.02 19.76 17.65 17.24

13.47 13.45 13.40 13.15 13.11

14.25 17.05 20.04 17.40 18.22

3.4. Synchronous fluorescence spectra The synchronous fluorescence spectra are frequently utilized to characterize the interaction between small molecules and proteins since it can provide the information about the molecular microenvironment in a proximity of the chromospheres. The synchronous fluorescence spectroscopy was introduced by Lloyd in 1971, who initially applied it in the field of forensic science where it was used to characterize complex mixtures providing fingerprints of intricate samples [67]. It is a useful method for studying the environment of amino acid residues by measuring the possible shift in position of the wavelength emission maximum corresponding to the changes in polarity around the chromophore molecule [30,68,69]. Moreover, the synchronous fluorescence of HSA at Δλ = 15 nm and 60 nm is characteristic of Tyr and Trp residues, respectively [28]. The Δλ value, representing the difference between excitation and emission wavelengths, is significant operating parameter. This technique concludes the conformational changes of the protein due to its binding reaction. The principal characteristics of synchronous fluorescence spectroscopy are a narrowing of the spectral band, a simplification of the emission spectra, sensitivity, the contraction of the spectral range and the possibility to avoid different alarming effects. Fig. 5A and B present a synchronous fluorescence spectra of the binary and ternary systems at Δλ = 15 and 60 nm with fixed the unchanged concentration of HSA. Fig. 5A demonstrate that the emission maximum of tyrosine residues has a little red shift (2 nm) which shows that the polarity around the Tyr residues has enhanced due to conformational changes of HSA. Also, it can be observed that the Tyr fluorescence emission of the systems was gradually decreased with enhancement of drugs. On the other hand, Fig. 5B indicate that the maximum emission wavelength has only a weak blue shift (about 3 nm) at the investigated concentration range when Δλ = 60 nm which indicate that the hydrophobicity around the Trp residue was enhanced. This could be an important hint to suggest conformational changes to has, which correspond to alter in polarity around the chromophore molecule during the formation of the complexes [28,68]. In agreement with these results, Amani et al. and Pourgonabadi et al. reported that the interaction of EST and TMX cause the blue shift in HSA spectra at Δλ = 60 nm [55,63].


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Fig. 6A and B demonstrate that the slope of the curves of the ternary systems at Δλ = 15 and 60 nm decreased in comparison to the binary systems. Although differences in degree of fluorescence quenching between binary and ternary systems at Δλ = 60 nm are not significant (Fig. 6B), but indicate that the access of drugs to protein fluorophores is partially limited by pre-binding of second drug. However, the slope was similar in binary and ternary systems at Δλ = 15 nm (Fig. 6A), indicating that the opportunity of EST and TMX to approach the Trp and Tyr residues was more or less the same in both systems. This fact signified that the presence of the first drug did not affect the quenching mechanism of HSA by the second drug in the ternary system. To sum up, the two anti-breast cancer drugs bound to the HSA molecules through different interactions.

3.5. Red edge excitation shift measurements

Fig. 5. Synchronous fluorescence spectra of binary systems of HSA with (A) EST at Δλ = 15 nm and (B) TMX at Δλ = 60 nm. The insets are the corresponding ternary systems under identical conditions except the protein (1 μM) was pre-incubated with 1 μM interferer drug. For the sake of clarity, the down black arrows show the fluorescence quenching process and the left or right blue arrows indicate the shift of spectra maximum peak. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Red edge excitation shift (REES) is a shift in the emission maximum toward a higher wavelength caused by a shift in the excitation wavelength toward the red edge of the absorption band [70]. REES is especially useful in monitoring motions surrounding the Trp residues in protein studies [71]. Various red-edge-excitation effects are manufacture by a redistribution of the energy levels both in the ground and the exited states of the chromophore aroused by alters of external conditions [72]. Calculation of the fluorescence emission and red edge excitation shift (REES) of HSA upon interaction with TMX and EST in all interacting systems cause to be it possible to compare the environmental and mobility features of the Trp residue in the HSA–drugs complexes. This mean that, in the presence of TMX or/and EST, the values of REES increased. For the present experiments, the choice was made to excite the Trp at both 295 nm and 305 nm to investigate the REES effect, and the results are listed in Table 2. In this Table, the difference of the emission maximum between that excited at 295 nm and at 305 nm is shown as Δλem,max. The restricted mobility around the Trp residue in the binary and ternary systems can be compared by the magnitude of REES. As shown, similar to the report of Danesh et al. the REES of HSA-drug binary and ternary complexes, especially in the case of TMX, presents higher values compared to the HSA alone indicating that the Trp residue in the complexes is in a slightly motionally restricted environment [54]. Also, the different values of the Δλem,max observed for the systems presumably reflected the different Trp environments in the binary and ternary complexes. In other words, the Trp microenvironment in the case of the ternary system appeared to be less polar while the binary complex offered a more polar environment.

Fig. 6. Comparison of curves of F/F0 versus EST and TMX concentration for the binary (●) and ternary systems (○) at (A) Δλ = 15 nm and (B) Δλ = 60 nm. Points are the experimental data.

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Table 2 Red edge excitation shift effects for HSA and HSA-drugs in the binary and ternary complexes at λex = 305 nm and 295 nm. System

λex: 295 nm

λex: 305 nm

Δλem−max

HSA HSA-EST (HSA-TMX)-EST HSA-TMX (HSA-EST)-TMX

341 338 339 337 335

344 344 345 345 345

3 6 6 8 10

3.6. Fluorescence quenching of HSA by TMX and EST in binary and ternary systems The fluorescence technique can be utilized to investigate the binding of small molecules to proteins involving the binding mechanism, binding mode, binding constant, binding forces, and intermolecular distances [8,30,33,42,69,73]. Among biopolymers, proteins are unique in displaying useful intrinsic fluorescence. On the other hand, the fluorescence manner of acting can provide information regarding the molecular environment in the proximity of chromophore groups [8,28,30,42, 69]. The fluorescence of HSA originates Trp, Tyr and Phe residues, but in fact, the main donor to the intrinsic fluorescence of HSA is the Trp residue by oneself [29,74]. When small molecule bind to HSA, alters of the intrinsic fluorescence intensity of HSA are induced by the micro-environment of the Trp residue [29,30,69]. Here the technique was employed to study the interaction of HSA with TMX and EST. The results

pointed at HSA having a strong fluorescence that presence of TMX and EST causes the quenching fluorescence of protein. Fig. 7A, B and their insets show the fluorescence spectra of HSA-EST and (HSA-TMX)-EST at the excitation wavelengths 280 nm and 295 nm, (pH = 7.4), respectively. When a fixed concentration of HSA was titrated with varying amounts of EST, a remarkable fluorescence decrease of HSA was observed. Such a decrease in intensity is known as quenching and this quenching was concentration-dependent. The fluorescence intensity of HSA decreased regularly and a slight blue shift was observed for the emission wavelength when increasing the EST concentration in the HSA–EST binary and ternary systems. The slight blue shift of the maximum emission wavelength and decline in fluorescence intensity exhibit a decreased polarity of the solvent, a change in the micro environment of the Trp and Tyr residues and a less hydrophilic nature of the fluorophore micro environment. The results demonstrated in this work show that EST bound to human serum albumin with high affinity and efficiently quenched the intrinsic fluorescence of HSA. The strong quenching of HSA fluorescence clearly indicates that it interacted with the drug, that conformational changes took place within HSA and that an inter-molecular energy transfer occurred between the drug and protein. A comparison of fluorescence quenching of the protein excited at 280 nm and 295 nm allows us to estimate the participation of the Trp and Tyr groups in the complex [75]. Fig. 7C, D and their insets show the same results of the Fig. 7A and B that present the fluorescence spectra of HSA-TMX and (HSA-EST)-TMX at the excitation wavelengths 280 nm and 295 nm. The results pointed

Fig. 7. Fluorescence emission spectra of the protein–drug complexes at 310 K in the presence of various concentrations of drugs (0–8.9 μM for the binary and 0–8.3 μM for the ternary systems) with the excitation at 295 nm; for the HSA-EST (A), [HSA-TMX]-EST (B), HSA-TMX and [HSA-EST]-TMX systems. The inset of figures are the same system with excitation at 280 nm. For the sake of clarity, the black arrows show the direction of fluorescence intensity change (enhancement or quenching) and the blue arrows indicate the shift of spectra maximum peak. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)


at HSA having a strong fluorescence emission at 340 nm after being excited with wavelength of 280 nm and 295 nm. In addition of varying concentrations of TMX caused a noticeable decrease in the fluorescence intensity of HSA. Similar to EST, the blue shift of the maximum emission wavelength indicate the polarity of micro environment of the protein fluorophores was decreased upon TMX binding which in good agreement with REES results. On the other hand, in the case of TMX along with fluorescence quenching at 340 nm, the enhancement of intensity was observed at 405 nm is due to fluorescence resonance energy transfer (FRET) which implies that TMX was bind near Trp214 (Fig. 7B, C and D). However, the insignificant part of intensity at 405 nm is related to the intrinsic fluorescence of TMX, which was corrected by measuring the fluorescence intensity of drug separately. Also, the occurrence of an isosbestic point at about 375 nm indicated the existence of bound and free tamoxifen in equilibrium [76]. Similar FRET was observed for TMX-human holo-transferrin interaction which quenching of protein tryptophan fluorescence at 330 nm accompanied with appearance/increase of new peak intensity at 405 nm [76].

3.7. Fluorescence quenching mechanism Quenching of fluorescence emission of a macromolecule induced by a quencher can be classified either as dynamic or static quenching. In dynamic quenching, increasing the temperature results in faster diffusion and hence larger amounts of collision which causes rising of the Stern–Volmer dynamic quenching constants (Ksv). In the other hand, in static one, increasing the temperature weakens the stability of the complexes that formed mainly by H-bounds/van der Waals interactions and hence reduces their quenching constant [77–79]. Contrary, the quenching constant of complex formed by hydrophobic interactions increases with increasing temperature because the hydrophobic effect been strengthened [80]. The mechanism of quenching can also be distinguished by the quenching rate constant (kq); when the values of kq (with assuming τ0 = 10−8 s) were much greater than the maximum diffusion rate constant of biomolecules (2 × 1010 M−1·s−1) indicated that the fluorescence quenching of protein was initiated by complex formation between protein and ligand [37,42,78,79]. In order to speculate the fluorescence quenching mechanism for the binary and ternary systems, the quenching experiments were carried out at 280, 290, 300 and 310 K. The fluorescence quenching data are analyzed by the Stern–Volmer equation (Eq. 1) and the results for the binary and ternary systems of EST as representative are depicted in Fig. 8A and B, respectively. The linearity in the plots of F0/F against [Q] confirmed one-to-one interaction between drugs and HSA up to 1 μM drug concentration (HSA

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to drug ratio is 1:1). As can be seen, the slope of curves, that are equal to Ksv values, increased with increasing temperature for all systems which indicates the involved binding forces are mainly hydrophobic interactions, because endothermic apolar interactions are strengthened with increasing temperature. The calculated values of Ksv at 310 K for the binary and ternary systems are also listed in Table 3. It can be found from the Table, the values of Ksv were higher for HSA–TMX than for the HSA– EST system. This means that TMX had a higher affinity to HSA than EST. In addition, the Ksv for EST and TMX in the binary system was higher than ternary system, which suggest that the quenching ability of EST and TMX to HSA was altered in the presence of the each other. This also implies that binding of each drug caused conformational change in HSA, thereby slightly reducing the affinity of binding of other drug in the ternary system. Also, the values of kq for all systems are greater than maximum diffusion rate constant of biomolecules, which indicated that the fluorescence of HSA was quenched by static mechanism in the presence of drugs [37,42,78,79]. In other words, the fluorescence quenching of HSA resulting from complex formation is predominant, while that from dynamic collision could be negligible. The modified Stern-Volmer analysis is useful in the estimation of the accessibility of Trp residues in proteins to the quencher molecules (Eq. 2). In addition of the Ksv and kq values, the fa values of interaction between EST and TMX to HSA are given in Table 3. As can be seen, the presence of the drugs affected the fa values in the same way as has been reported earlier [81,82]. When the fa value was equal to 1, all the Trp residues were accessible to the quencher. Consequently, a change in the value of fa caused the fraction of fluorescent components accessible to the quencher to become altered. As can be seen in Table 3, the value of fa was decreased for ternary systems compared to binary complexes because some of protein fluorophores have been out of TMX reach by EST and vice versa.

3.8. Determination of binding affinity Assuming that the quenching of protein fluorescence caused by ligand binding (the mechanism of static quenching is dominant), the dependence of protein fluorescence intensity on the ligands (EST and TMX) concentration can be analyzed according to previously published papers [32,33]. Given that the protein fluorescence intensity change is reached to plateau in the 1:1 ratio of protein-ligand concentration and also previous reports [34,35], the value of n was assumed one. Thus, the Eq. 3 was used for calculation of the lignads dissociation constant in the binary and ternary systems without preliminary estimation of the Flim value (Fig. 9A, B and their insets). As can be seen in Table 3,

Fig. 8. Stern-Volmer plots for quenching of HSA fluorescence (λex: 280 nm, λem: 340 nm) by EST in the binary (A) and ternary (B) systems at 280 (●), 290 (○), 300 (▲) and 310 K (△). For the ternary systems, 1 μM HSA solution was pre-incubated with 1 μM interferer drug. Points are the experimental data. Solid line was calculated from Eq. 1.

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Table 3 Ksv, kq, fa, KQ and Kdiss values of EST and TMX upon interaction with HSA at 310 K in the binary and ternary systems with excitation of the protein at 280 and 295 nm. System

HSA-EST (HSA-TMX)-EST HSA-TMX (HSA-EST)-TMX

Ksv × 10−4 (M−1)

kq × 10−12 (M−1·s−1)

fa

280

295

280

280

295

280

295

280

295

98.38 90.96 105.31 88.26

73.88 27.84 40.70 22.02

9.84 9.10 10.53 8.83

0.77 0.94 0.79 0.70

0.61 0.49 0.49 0.36

2.82 1.04 2.61 4.81

4.60 1.07 3.19 2.17

2.12 1.96 1.63 1.98

1.02 4.68 4.51 8.21

when the protein was excited at 295 nm, the Kdiss values of EST and TMX in the binary complex are smaller than ternary complex while by exciting the protein at 280 nm, the binding affinity of binary and ternary systems have no significant difference. This implies that this two drug may bind to different sites on serum albumin. The value of Kdiss is important to understand the distribution of the drug in plasma since the weak binding can lead to a short life time or poor distribution, while strong binding can decrease the concentration of free drug in plasma [8,42]. The value of Kdiss illustrated that EST and TMX bind to HSA strongly similar common HSA ligands such as heme, bilirubin and fatty acids [83– 85]. It also implies that EST and TMX can be tightly stored and carried by HSA in the body. It was found that the simultaneous administration of EST and TMX might affect their ADME (Absorption, Distribution, Metabolism and Elimination). The obtained values of dissociation constant for EST-HSA and TMX-HSA complexes are smaller than the binding affinity values reported by Pearlman et al., Moll et al., Sharif-Barfeh et al., and Danesh et al. for EST and Pourgonabadi et al., Bourassa et al.

KQ (M−1)

Kdiss × 107 (M)

(2011), Bourassa et al. (2016), and Maciazek-Jurczyk et al. for TMX which are between 10−4 and 10−6 [54,55,60,61,64,86–88]. This is can be attributed to differences in experimental techniques, models used for calculation of binding affinity and the use of fatty acid free (FA free) HSA in our experiment. However, Zsila and, Chen and Hage, who used FA free HSA in the binding experiments, obtained similar Kdiss value for TMX (10−7) [89,90]. Surprisingly, although the binding affinity of TMX in the absence and the presence of EST has a little change, the enhancement of fluorescence intensity at 405 nm is almost 10-fold amplified in the presence of EST. In addition, calculation of Kdiss based on enhancement at 405 nm indicate that the binding affinity of TMX to HSA in the presence of EST is 20-fold less than the absence of EST (Kdiss of TMX-HSA is 0.7 versus 35.1 μM for [EST-HSA]-TMX). These implies that the prior binding of EST to HSA causes the structural changes which decrease the distance between Trp residue and TMX without alteration of drug binding site which in agreement with REES results. This is in accordance with the enhancement of intensity at 405 nm of [TMX-HSA]-EST system in the presence of increasing concentration of EST which confirmed the neighboring of protein Trp to TMX in the presence of EST (Fig. 7C and D). 3.9. Forces involved in the binding process Considering the dependence of dissociation constant on temperature, a thermodynamic process was considered to be responsible for the formation of a complex. Therefore, the thermodynamic parameters dependent on temperatures were analyzed. The acting forces between a small molecule and macromolecule mainly include hydrogen bonds, van der Waals forces, electrostatic forces, and hydrophobic interaction forces [42,77,78,80]. In order to obtain the information about the forces persisting in the present binding processes, the thermodynamic parameters of binding were obtained from van't Hoff plot followed by the Gibbs equation (data not shown). Table 4 shows the values of ΔH°, ΔS° and ΔG° obtained for the binary and ternary systems from the slopes and ordinates at the origin of the fitted lines. From Table 4, it can be seen that the negative sign for free energy (ΔG°) means that Table 4 Thermodynamic parameters (ΔH°, ΔS° and ΔG°) for the interaction of HSA with EST and TMX in the binary and ternary systems with excitation of the protein at 280 nm. System HSA-EST

Fig. 9. The dependence of HSA relative fluorescence intensity (F/F0) at 340 nm on total concentrations of (A) EST and (B) TMX in the binary (●) and the ternary complexes (○) with excitation at 295 nm. The inset of figures are the same system with excitation at 280 nm. Points are the experimental data. Solid line was calculated from Eq. 3.

Temp. (K)

280 290 300 310 (HSA-TMX)-EST 280 290 300 310 HSA-TMX 280 290 300 310 (HSA-EST)-TMX 280 290 300 310

Kdiss × 107 ΔH° ΔS° ΔG° (M) (kJ·mol−1) (J·mol−1·K−1) (kJ·mol−1) 3.46 2.93 2.53 2.12 3.21 2.86 2.33 1.96 2.70 2.28 1.96 1.63 3.33 2.80 2.40 1.98

11.66

90.24

12.15

89.33

12.01

91.30

12.31

88.73

−13.61 −14.51 −15.41 −16.32 −12.86 −13.75 −14.64 −15.54 −13.56 −14.47 −15.38 −16.30 −12.53 −13.42 −14.31 −15.19


the interaction process is spontaneous [42,80]. The positive enthalpy (ΔH°) and entropy (ΔS°) values for all systems indicate that the binding processes are mainly entropy-driven and the hydrophobic forces play a major role in the interactions [31,37,42,78,80]. This result is in accordance with Bourassa et al. (2011), Bourassa et al. (2016), Pourgonabadi et al., Danesh et al. and Ling et al. which suggested the hydrophobic interactions are the predominant intermolecular forces stabilizing the EST-HSA and TMX-HSA complexes [54,55,60,61,76,91]. 3.10. Molecular modeling In order to provide a considerable insight into the interactions of HSA with EST and TMX, molecular modeling has been employed which locates the binding sites for these drugs. The 3D structure of HSA was obtained from Proteins Data Bank (1AO6). The possible conformations of the ligand-protein complex were calculated using Autodock program. Among 100 conformers obtained from 200 runs, the conformer with the lowest binding free energy was used for further analyses. The summary of the ligands docking results into the whole protein (all known protein binding sites set to rigid) are shown in Table 5. As can be seen, EST shows less affinity to the protein than TMX in terms of docking score and total binding energy which are consistent with fluorescence quenching data. While the main acting force between both of drugs is hydrophobic interaction, but hydrogen bond and hydrogen bond/salt bridge (electrostatic) are present in EST- and TMX-HSA systems, respectively. The best/most plausible docking poses for EST and TMX are shown in Fig. 10A. As can be seen, the binding sites for EST and TMX did not overlap and two drugs did not compete for the same site in HSA. The principal regions where the ligands tends to bind to HSA are located in hydrophobic pockets in subdomains IIA and IIIA, and tryptophan residue located in subdomain IIA is also a primary binding region. These two subdomains are positioned in the two major binding sites, i.e. Sudlow's site I and II, respectively [8,10]. From Fig. 10A, it can be seen that EST binds to the sixth fatty acid binding site (FA6) between two α-helix regions (H2-IIA and H2-IIB) and properly fits the binding pocket. Then EST can be bound to HSA with the effect on two α-helixes which was in agreement with the far-UV CD curves. On the other hand, TMX formed a salt bridge with Glu153 (in H3-IB) and located in hydrophobic pocket of Sudlow site I close to Trp residue (3.42 nm). Binding of EST to the sixth fatty acid binding site was more confirmed by two reports: (i) Bruning and Bonfror report that a physiological in vivo or in vitro increase of plasma free fatly acid (such as linoleate, linolenate, and arachidonate) concentration or lipase activity was found to be accompanied by a rise of the available non-protein bound estradiol fraction [92]; (ii) Westphal (1986) and Brown (1982) suggested that the BSA subdomain IIC (amino acids 307–385 of BSA), which was isolated from BSA by Pearlman and Fong (1972) after limited peptic hydrolysis, may be the primary steroid binding site of serum albumins [93–95]. The subdomain IIC of BSA is consistent with subdomain IIB of HSA which the FA6 is between its H2-IIB and H2IIA. In the case of TMX, while mostly of the papers (such as Buttar et al., Shah and Parsons), point out that the main binding site of TMX is in subdomain IB (drug binding site III) or IIIA (drug binding site II) with Kdiss between 10−4 and 10−6, there are some reports (such as Sjöholm

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et al. and, Chen and Hage) which suggested that TMX binds to subdomain IIA (drug binding site I) with Kdiss b 10− 6 [89,96–98]. It seems that the difference in the proposed TMX site is due to the prebinding of fatty acids to HSA; so that the reports that have used the serum HSA (non\\FA free) suggested the subdomain IB and IIIA are the TMX binding site, and on the other hand, the studies which have utilized the FA free HSA proposed the subdomain IIA. Also, the proposed binding sites for TMX on HSA and BSA by Bourassa et al., who have used ArgusLab 4.0.1 software, are different with our docking results [60,62]. This may be due to use of different setting/software for molecular modeling. However, Bourassa et al. suggested TMX binding site are our third modeled pose in terms of binding energy (data not shown). Moreover, Chen and Hage reported that when warfarin was used in competition with TMX, this had a positive allosteric effect on TMX-HSA binding, whereas using TMX as a competitor indicated that TMX had a negative allosteric effect on warfarin-HSA binding [89]. Given that warfarin binds to the bottom of drug binding site I (subdomain IIA) and our docking results indicate that TMX binds to entrance of drug binding site I in front of warfarin binding site, it seems that the binding of warfarin to the bottom of site I results in conformational changes that facilitates/improves the interaction of TMX, whereas contrary to this, the binding of TMX leads to the structural alterations that causing the binding of warfarin be more difficult [1,5]. There are six hydrophobic residues (Ala213, Val216, Val235, Leu327, Leu331 and Ala350), two acidic residue (Asp324 and Glu354), three basic residue (Arg209, Lys212 and Lys351), and one glycine residue (Gly328) nearby EST (Fig. 10B). In addition, one hydrogen bond was observed between side chain of Glu354 (located in H3-IIB) and EST with energy −2.1 kcal·mol−1 in distance 3.17 nm (Table 5). In agreement with the results, Danesh et al. by ITC experiments showed that both van der Waals forces and hydrogen bonding are predominant in ESTHSA interaction [54]. There were large hydrophobic cavities in HSA of which one of the largest was in the sub-domain IIA where the drugs can bind (Sudlow's site I) [99]. The proposed binding site for TMX overlaps with the seventh fatty acid binding site (FA7) of HSA. TMX was accommodated into a pocket surrounded by hydrophobic residues (Phe157, Trp214, Leu219, Leu238, Leu260, Ile290 and Ala291) which indicates that the nature of interaction between HSA and EST is mainly hydrophobic. Also, the quenching of protein fluorescence accompany with efficient FRET confirms the docking results. In the binding site, TMX is surrounded by polar/basic Tyr50, Lys195, Gln196, Lys199, Arg218, Arg222, His242, Arg257, His288, residues. Fig. 10B indicates that there are three glutamate residues (Glu153, 3.93 nm; Glu188, 5.48 nm; Glu292, 5.59) near the NH+ of TMX (at pH 7) which reveal that the ionic interaction is involved in the interaction between protein and ligand. The potency of TMX for the salt bridge formation in the interaction with proteins has already been reported by Sarzehi et al. [76]. Moreover, TMX was able to form hydrogen bonds with Ser192 and Glu292 with energy −5.8 kcal·mol−1 (Table 5). In accordance, Bourassa (2011), Bourassa (2016) and Pourgonabadi have suggested that TMX bounds HSA via both hydrophobic and hydrophilic interactions with extended H-bonding network [55,60,61]. The average distance of Trp residue of HSA from EST and TMX is 12.17 and 3.42 nm, respectively. Due to efficient FRET

Table 5 Molecular docking results of EST and TMX against HSA showing binding energies, docking score and residues involved in the drug-protein interaction. Ligand Residues involved in the interaction

Etotala

EST TMX

−108.7 0.0 −145.5 −7.6

a

Arg209, Lys212, Ala213, Val216, Asp324, Val235, Leu327, Gly328, Leu331, Ala350, Lys351, Glu354 Tyr50, Glu153, Phe157, Glu188, Ser192, Lys195, Gln196, Lys199, Trp214, Arg218, Leu219, Arg222, Leu238, His242, Arg257, Leu260, His288, Ile290, Ala291, Glu292

The unit of energy is kcal/mol.

Eelectrostatic EH-Bond EVdW −2.1 −5.8

−31.1 −47.9

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Fig. 10. (A) Docking poses of EST and TMX into HSA (PDB ID 1AO6). Target protein is illustrated as cartoon with secondary structure colour-coded. The two primary drug binding sites are indicated as Sudlow's site I and Sudlow's site II. The drug molecules are depicted in surface model and coloured according to lipophilicity; Red: polar and Green: hydrophobic. (B and C) Two dimensional representation of HSA binding site interactions with EST and TMX, respectively. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

was typically occurred in the range of 1–10 nm, the molecular modeling results are in total agreement with results of fluorescence quenching experiments. Also, the exploration of different/distinct binding sites for ligands confirms all experimental data.

we would especially like to thank Dr. R. Khodarahmi, the head of our lab. As our teacher and mentor, he has taught us more than we could ever give him credit for here. He has shown us, by his example, what a good scientist and person should be.

4. Conclusions

References

The performed binding study of two anti-breast cancer drugs (TMX and EST) with HSA should exhibit significant importance in pharmacy, pharmacology and biochemistry for explaining the transport and storage of drugs during cancer treatment. Combining the results obtained with intrinsic fluorescence, RLS and molecular modeling demonstrated that TMX and EST may bind to distinct sites on HSA. The transfer of fluorescence resonance energy from protein to TMX illustrated that this drug bind near the Trp residue in domain IIA, which confirmed the docking results. Also, amplification of fluorescence enhancement at 405 nm in the presence of EST implies that the prior binding of EST to HSA causes the structural changes which decrease the distance between Trp residue and TMX without alteration of drug binding site. The CD and synchronous fluorescence data revealed that TMX and EST induced conformational changes which can modify the physiological functions of HSA and binding of other drugs. The present study on the interaction between TMX and EST with HSA should open new vistas for what is reasonable regarding drug design. Declaration of interest The authors certify that no actual or potential conflict of interest in relation to this article exists. Acknowledgments The research and technology council of Kermanshah University of Medical Sciences and Islamic Azad University-Mashhad Branch were gratefully acknowledged for financial supports (Grant No. 94517). Also,

[1] I. Petitpas, A.A. Bhattacharya, S. Twine, M. East, S. Curry, Crystal structure analysis of warfarin binding to human serum albumin anatomy of drug site I, J. Biol. Chem. 276 (2001) 22804–22809. [2] J. Hodgson, ADMET—turning chemicals into drugs, Nat. Biotechnol. 19 (2001) 722–726. [3] X.M. He, D.C. Carter, Atomic structure and chemistry of human serum albumin, Nature 358 (1992) 209–215. [4] R.L. Rich, Y.S.N. Day, T.A. Morton, D.G. Myszka, High-resolution and high-throughput protocols for measuring drug/human serum albumin interactions using BIACORE, Anal. Biochem. 296 (2001) 197–207. [5] J. Ghuman, P.A. Zunszain, I. Petitpas, A.A. Bhattacharya, M. Otagiri, S. Curry, Structural basis of the drug-binding specificity of human serum albumin, J. Mol. Biol. 353 (2005) 38–52. [6] J. Tang, S. Qi, X. Chen, Spectroscopic studies of the interaction of anti-coagulant rodenticide diphacinone with human serum albumin, J. Mol. Struct. 779 (2005) 87–95. [7] A. Kober, I. Sjoholm, The binding sites on human serum albumin for some nonsteroidal antiinflammatory drugs, Mol. Pharmacol. 18 (1980) 421–426. [8] R. Khodarahmi, S.A. Karimi, M.R.A. Kooshk, S.A. Ghadami, S. Ghobadi, M. Amani, Comparative spectroscopic studies on drug binding characteristics and protein surface hydrophobicity of native and modified forms of bovine serum albumin: possible relevance to change in protein structure/function upon non-enzymatic glycation, Spectrochim. Acta A Mol. Biomol. Spectrosc. 89 (2012) 177–186. [9] P.A. Schwartz, C.T. Rhodes, D.S. Greene, Effect of free fatty acid concentration on furosemide binding to human serum albumin, Pharmacology 22 (1981) 364–370. [10] T. Peters Jr., All About Albumin: Biochemistry, Genetics, and Medical Applications, Academic press, San Diego, 1996. [11] C. Bertucci, G. Ascoli, G. Uccello-Barretta, L. Di Bari, P. Salvadori, The binding of 5fluorouracil to native and modified human serum albumin: UV, CD, and 1H and 19F NMR investigation, J. Pharm. Biomed. Anal. 13 (1995) 1087–1093. [12] T.E. Bull, B. Halle, B. Lindman, Internal motion at the chloride binding sites of human serum albumin by NMR relaxation studies, FEBS Lett. 86 (1978) 25–28. [13] C.Q. Jiang, M.X. Gao, X.Z. Meng, Study of the interaction between daunorubicin and human serum albumin, and the determination of daunorubicin in blood serum samples, Spectrochim. Acta A Mol. Biomol. Spectrosc. 59 (2003) 1605–1610.

N. Moradi et al. / Journal of Molecular Liquids 249 (2018) 1083–1096 [14] S. Nahar, R. Carpentier, H.A. Tajmir-Riahi, Interaction of trivalent Al and Ga cations with proteins of PSII. Cation binding mode and protein conformation by FTIR spectroscopy, J. Inorg. Biochem. 65 (1997) 245–250. [15] F. Zsila, Z. Bikádi, M. Simonyi, Probing the binding of the flavonoid, quercetin to human serum albumin by circular dichroism, electronic absorption spectroscopy and molecular modelling methods, Biochem. Pharmacol. 65 (2003) 447–456. [16] A.A. Bhattacharya, T. Grüne, S. Curry, Crystallographic analysis reveals common modes of binding of medium and long-chain fatty acids to human serum albumin, J. Mol. Biol. 303 (2000) 721–732. [17] C. Carani, K. Qin, M. Simoni, M. Faustini-Fustini, S. Serpente, J. Boyd, K.S. Korach, E.R. Simpson, Effect of testosterone and estradiol in a man with aromatase deficiency, N. Engl. J. Med. 337 (1997) 91–95. [18] R.M. O'Regan, V.C. Jordan, The evolution of tamoxifen therapy in breast cancer: selective oestrogen-receptor modulators and downregulators, Lancet Oncol. 3 (2002) 207–214. [19] L.S. Cook, N.S. Weiss, S.M. Schwartz, E. White, B. McKnight, D.E. Moore, J.R. Daling, Population-based study of tamoxifen therapy and subsequent ovarian, endometrial, and breast cancers, J. Natl. Cancer Inst. 87 (1995) 1359–1364. [20] S.S. Lum, E.A. Woltering, W.S. Fletcher, R.F. Pommier, Changes in serum estrogen levels in women during tamoxifen therapy, Am. J. Surg. 173 (1997) 399–402. [21] J. Gjerde, E.R. Kisanga, M. Hauglid, P.I. Holm, G. Mellgren, E.A. Lien, Identification and quantification of tamoxifen and four metabolites in serum by liquid chromatography–tandem mass spectrometry, J. Chromatogr. 1082 (2005) 6–14. [22] J. Gjerde, M. Hauglid, H. Breilid, S. Lundgren, J.E. Varhaug, E.R. Kisanga, G. Mellgren, V.M. Steen, E.A. Lien, Effects of CYP2D6 and SULT1A1 genotypes including SULT1A1 gene copy number on tamoxifen metabolism, Ann. Oncol. 19 (2008) 56–61. [23] T. Chatterjee, A. Pal, S. Dey, B.K. Chatterjee, P. Chakrabarti, Interaction of virstatin with human serum albumin: spectroscopic analysis and molecular modeling, PLoS One 7 (2012), e37468. [24] M. DellaGreca, M.R. Iesce, M. Isidori, A. Nardelli, L. Previtera, M. Rubino, Phototransformation products of tamoxifen by sunlight in water. Toxicity of the drug and its derivatives on aquatic organisms, Chemosphere 67 (2007) 1933–1939. [25] X. Ma, C. Zhang, J. Deng, Y. Song, Q. Li, Y. Guo, C. Li, Simultaneous degradation of estrone, 17β-estradiol and 17α-ethinyl estradiol in an aqueous UV/H2O2 system, Int. J. Env. Res, Public Health 12 (2015) 12016–12029. [26] V.M. Mboula, V. Héquet, Y. Andres, Y. Gru, R. Colin, J. Doña-Rodríguez, L. PastranaMartínez, A. Silva, M. Leleu, A. Tindall, Photocatalytic degradation of estradiol under simulated solar light and assessment of estrogenic activity, Appl. Catal. B 162 (2015) 437–444. [27] H.A. Merey, M.M. Galal, M.Y. Salem, E.M. Abdel-Moety, Novel stability indicating methods for the determination of certain synthetic estrogen level modifiers, Bull. Fac. Pharm., Cairo University 51 (2013) 69–79. [28] E.A. Burstein, N.S. Vedenkina, M.N. Ivkova, Fluorescence and the location of tryptophan residues in protein molecules, Photochem. Photobiol. 18 (1973) 263–279. [29] J.R. Lackowicz, Principle of Fluorescence Spectroscopy, Springer Science Business Media, New York, 2006. [30] N. Abdollahpour, A. Asoodeh, M.R. Saberi, J. Chamani, Separate and simultaneous binding effects of aspirin and amlodipine to human serum albumin based on fluorescence spectroscopic and molecular modeling characterizations: a mechanistic insight for determining usage drugs doses, J. Lumin. 131 (2011) 1885–1899. [31] N. Moradi, M.R. Ashrafi-Kooshk, S. Ghobadi, M. Shahlaei, R. Khodarahmi, Spectroscopic study of drug-binding characteristics of unmodified and pNPA-based acetylated human serum albumin: does esterase activity affect microenvironment of drug binding sites on the protein? J. Lumin. 160 (2015) 351–361. [32] M. Ghahramani, R. Yousefi, K. Khoshaman, S.S. Moghadam, B.I. Kurganov, Evaluation of structure, chaperone-like activity and protective ability of peroxynitrite modified human α-crystallin subunits against copper-mediated ascorbic acid oxidation, Int. J. Biol. Macromol. 87 (2016) 208–221. [33] B.I. Kurganov, N.P. Sugrobova, V.A. Yakovlev, Estimation of dissociation constant of enzyme-ligand complex from fluorometric data by “difference” method, FEBS Lett. 19 (1972) 308–310. [34] D.S. Hage, J. Austin, High-performance affinity chromatography and immobilized serum albumin as probes for drug-and hormone-protein binding, J. Chromatogr. B Biomed. Sci. Appl. 739 (2000) 39–54. [35] D.S. Hage, A. Sengupta, Studies of protein binding to nonpolar solutes by using zonal elution and high-performance affinity chromatography: interactions of cis-and trans-clomiphene with human serum albumin in the presence of β-cyclodextrin, Anal. Chem. 70 (1998) 4602–4609. [36] M. Van de Weert, L. Stella, Fluorescence quenching and ligand binding: a critical discussion of a popular methodology, J. Mol. Struct. 998 (2011) 144–150. [37] O.A. Chaves, B. Mathew, D. Cesarin-Sobrinho, B. Lakshminarayanan, M. Joy, G.E. Mathew, J. Suresh, J.C. Netto-Ferreira, Spectroscopic, zeta potential and molecular docking analysis on the interaction between human serum albumin and halogenated thienyl chalcones, J. Mol. Liq. 242 (2017) 1018–1026. [38] O. Trott, A.J. Olson, AutoDock Vina: improving the speed and accuracy of docking with a new scoring function, efficient optimization, and multithreading, J. Comput. Chem. 31 (2010) 455–461. [39] M. Froimowitz, HyperChem: a software package for computational chemistry and molecular modeling, BioTechniques 14 (1993) 1010–1013. [40] A. Streitwieser, Molecular Orbital Theory for Organic Chemists, Wiley, New York, 1961. [41] M.F. Sanner, Python: a programming language for software integration and development, J. Mol. Graph. Model. 17 (1999) 57–61. [42] M. Shahlaei, B. Rahimi, A. Nowroozi, M.R. Ashrafi-Kooshk, K. Sadrjavadi, R. Khodarahmi, Exploring binding properties of sertraline with human serum albumin: combination of spectroscopic and molecular modeling studies, Chem. Biol. Interact. 242 (2015) 235–246.

1095

[43] S. Sugio, A. Kashima, S. Mochizuki, M. Noda, K. Kobayashi, Crystal structure of human serum albumin at 2.5 Å resolution, Protein Eng. 12 (1999) 439–446. [44] G.M. Morris, D.S. Goodsell, R.S. Halliday, R. Huey, W.E. Hart, R.K. Belew, A.J. Olson, Automated docking using a Lamarckian genetic algorithm and an empirical binding free energy function, J. Comput. Chem. 19 (1998) 1639–1662. [45] W. Humphrey, A. Dalke, K. Schulten, VMD: visual molecular dynamics, J. Mol. Graph. 14 (1996) 33–38. [46] R.A. Laskowski, M.B. Swindells, LigPlot+: multiple ligand–protein interaction diagrams for drug discovery, J. Chem. Inf. Model. 51 (2011) 2778–2786. [47] P. Mazellier, L. Méité, J. De Laat, Photodegradation of the steroid hormones 17β-estradiol (E2) and 17α-ethinylestradiol (EE2) in dilute aqueous solution, Chemosphere 73 (2008) 1216–1223. [48] B. Liu, X. Liu, Direct photolysis of estrogens in aqueous solutions, Sci. Total Environ. 320 (2004) 269–274. [49] J. Šalamoun, M. Macka, M. Nechvátal, M. Matoušek, L. Knesel, Identification of products formed during UV irradiation of tamoxifen and their use for fluorescence detection in high-performance liquid chromatography, J. Chromatogr. 514 (1990) 179–187. [50] R.R. Chowdhury, P.A. Charpentier, M.B. Ray, Photodegradation of 17β-estradiol in aquatic solution under solar irradiation: kinetics and influencing water parameters, J. Photochem. Photobiol. A Chem. 219 (2011) 67–75. [51] J. Anglister, I.Z. Steinberg, Resonance rayleigh scattering of cyanine dyes in solution, J. Chem. Phys. 78 (1983) 5358–5368. [52] D. Gao, Y. Tian, F. Liang, D. Jin, Y. Chen, H. Zhang, A. Yu, Investigation on the pH-dependent binding of Eosin Y and bovine serum albumin by spectral methods, J. Lumin. 127 (2007) 515–522. [53] R.F. Pasternack, A. Giannetto, P. Pagano, E.J. Gibbs, Self-assembly of porphyrins on nucleic acids and polypeptides, J. Am. Chem. Soc. 113 (1991) 7799–7800. [54] N. Danesh, Z. Navaee Sedighi, S. Beigoli, A. Sharifi-Rad, M.R. Saberi, J. Chamani, Determining the binding site and binding affinity of estradiol to human serum albumin and holo-transferrin: fluorescence spectroscopic, isothermal titration calorimetry and molecular modeling approaches, J. Biomol. Struct. Dyn. (2017) 1–17, https:// doi.org/10.1080/07391102.2017.1333460. [55] S. Pourgonabadi, M. Reza Saberi, J. Khan Chamani, Investigating the antagonistic action between aspirin and tamoxifen with hsa: identification of binding sites in binary and ternary drug-protein systems by spectroscopic and molecular modeling approaches, Protein Pept. Lett. 18 (2011) 305–317. [56] X. Long, C. Zhang, J. Cheng, S. Bi, A novel method for study of the aggregation of protein induced by metal ion aluminum (III) using resonance Rayleigh scattering technique, Spectrochim. Acta A Mol. Biomol. Spectrosc. 69 (2008) 71–77. [57] P. Verdino, W. Keller, Circular dichroism analysis of allergens, Methods 32 (2004) 241–248. [58] I. Matei, M. Hillebrand, Interaction of kaempferol with human serum albumin: a fluorescence and circular dichroism study, J. Pharm. Biomed. Anal. 51 (2010) 768–773. [59] F. Ding, W. Liu, X. Zhang, L. Zhang, Y. Sun, Fluorescence and circular dichroism studies of conjugates between metsulfuron-methyl and human serum albumin, Colloids Surf. B: Biointerfaces 76 (2010) 441–448. [60] P. Bourassa, S. Dubeau, G.M. Maharvi, A.H. Fauq, T.J. Thomas, H.A. Tajmir-Riahi, Binding of antitumor tamoxifen and its metabolites 4-hydroxytamoxifen and endoxifen to human serum albumin, Biochimie 93 (2011) 1089–1101. [61] P. Bourassa, T.J. Thomas, H.A. Tajmir-Riahi, A short review on the delivery of breast anticancer drug tamoxifen and its metabolites by serum proteins, J. Nanomed. Res. 4 (2016) (00080). [62] P. Bourassa, S. Dubeau, G.M. Maharvi, A.H. Fauq, T. Thomas, H. Tajmir-Riahi, Locating the binding sites of anticancer tamoxifen and its metabolites 4-hydroxytamoxifen and endoxifen on bovine serum albumin, Eur. J. Med. Chem. 46 (2011) 4344–4353. [63] N. Amani, M. Reza Saberi, J. Khan Chamani, Investigation by fluorescence spectroscopy, resonance rayleigh scattering and zeta potential approaches of the separate and simultaneous binding effect of paclitaxel and estradiol with human serum albumin, Protein Pept. Lett. 18 (2011) 935–951. [64] Z. Sharif-Barfeh, S. Beigoli, S. Marouzi, A.S. Rad, A. Asoodeh, J. Chamani, Multi-spectroscopic and HPLC studies of the interaction between estradiol and cyclophosphamide with human serum albumin: binary and ternary systems, J. Solut. Chem. 46 (2017) 488–504. [65] T. Imoto, Stabilization of protein, Cell. Mol. Life Sci. 53 (1997) 215–223. [66] S. Robic, Mathematics, thermodynamics, and modeling to address ten common misconceptions about protein structure, folding, and stability, CBE Life Sci. Educ. 9 (2010) 189–195. [67] J.B.F. Lloyd, L.W. Evett, J.M. Dubery, Examination of petroleum products of high relative molecular mass for forensic science purposes by synchronous fluorescence spectroscopy. I1: discrimination within an arbitrary set of representative samples, J. Forensic Sci. 25 (1980) 589–602. [68] G. Zhang, Q. Que, J. Pan, J. Guo, Study of the interaction between icariin and human serum albumin by fluorescence spectroscopy, J. Mol. Struct. 881 (2008) 132–138. [69] T. Zohoorian-Abootorabi, H. Sanee, H. Iranfar, M.R. Saberi, J. Chamani, Separate and simultaneous binding effects through a non-cooperative behavior between cyclophosphamide hydrochloride and fluoxymesterone upon interaction with human serum albumin: multi-spectroscopic and molecular modeling approaches, Spectrochim. Acta A Mol. Biomol. Spectrosc. 88 (2012) 177–191. [70] A.P. Demchenko, A.S. Ladokhin, Red-edge-excitation fluorescence spectroscopy of indole and tryptophan, Eur. Biophys. J. 15 (1988) 369–379. [71] M.C. Tory, A.R. Merrill, Determination of membrane protein topology by red-edge excitation shift analysis: application to the membrane-bound colicin E1 channel peptide, Biochim. Biophys. Acta 1564 (2002) 435–448.

1096


[72] S. Guha, S.S. Rawat, A. Chattopadhyay, B. Bhattacharyya, Tubulin conformation and dynamics: a red edge excitation shift study, Biochemistry 35 (1996) 13426–13433. [73] Z.X. Wang, N.R. Kumar, D.K. Srivastava, A novel spectroscopic titration method for determining the dissociation constant and stoichiometry of protein-ligand complex, Anal. Biochem. 206 (1992) 376–381. [74] A. Sułkowska, B. Bojko, J. Równicka, W.W. Sułkowski, Paracetamol and cytarabine binding competition in high affinity binding sites of transporting protein, J. Mol. Struct. 792 (2006) 249–256. [75] Y. Li, W. He, H. Liu, X. Yao, Z. Hu, Daidzein interaction with human serum albumin studied using optical spectroscopy and molecular modeling methods, J. Mol. Struct. 831 (2007) 144–150. [76] S. Sarzehi, J. Chamani, Investigation on the interaction between tamoxifen and human holo-transferrin: determination of the binding mechanism by fluorescence quenching, resonance light scattering and circular dichroism methods, Int. J. Biol. Macromol. 47 (2010) 558–569. [77] F. Jafari, S. Samadi, A. Nowroozi, K. Sadrjavadi, S. Moradi, M.R. Ashrafi-Kooshk, M. Shahlaei, Experimental and computational studies on the binding of diazinon to human serum albumin, J. Biomol. Struct. Dyn. (2017) 1–21, https://doi.org/10. 1080/07391102.2017.1329096. [78] X. Li, X. Cui, X. Yi, S. Zhong, Mechanistic and conformational studies on the interaction of anesthetic sevoflurane with human serum albumin by multispectroscopic methods, J. Mol. Liq. 241 (2017) 577–583. [79] M. Shahlaei, B. Rahimi, M.R. Ashrafi-Kooshk, K. Sadrjavadi, R. Khodarahmi, Probing of possible olanzapine binding site on human serum albumin: combination of spectroscopic methods and molecular dynamics simulation, J. Lumin. 158 (2015) 91–98. [80] N. Bijari, Y. Shokoohinia, M.R. Ashrafi-Kooshk, S. Ranjbar, S. Parvaneh, M. MoieniArya, R. Khodarahmi, Spectroscopic study of interaction between osthole and human serum albumin: identification of possible binding site of the compound, J. Lumin. 143 (2013) 328–336. [81] F.A. Beckford, Reaction of the anticancer organometallic ruthenium compound, [(η6p-Cymene)Ru(ATSC)Cl]PF6 with human serum albumin, Int, J. Inorg. Chem. 2010 (2010) 7. [82] R.G. Machicote, M.E. Pacheco, L. Bruzzone, Binding of several benzodiazepines to bovine serum albumin: fluorescence study, Spectrochim. Acta A Mol. Biomol. Spectrosc. 77 (2010) 466–472. [83] P. Ascenzi, M. Fasano, Serum heme-albumin: an allosteric protein, IUBMB Life 61 (2009) 1118–1122. [84] J.D. Ashbrook, A.A. Spector, E.C. Santos, J. Fletcher, Long chain fatty acid binding to human plasma albumin, J. Biol. Chem. 250 (1975) 2333–2338.

[85] R.D. Gray, S.D. Stroupe, Kinetics and mechanism of bilirubin binding to human serum albumin, J. Biol. Chem. 253 (1978) 4370–4377. [86] M. Maciążek-Jurczyk, M. Maliszewska, J. Pożycka, J. Równicka-Zubik, A. Góra, A. Sułkowska, Tamoxifen and curcumin binding to serum albumin. Spectroscopic study, J. Mol. Struct. 1044 (2013) 194–200. [87] G.W. Moll, R.L. Rosenfiel Jr., J.H. Helke, Estradiol-testosterone binding interactions and free plasma estradiol under physiological conditions, J. Clin. Endocrinol. Metab. 52 (1981) 868–874. [88] W.H. Pearlman, I.F.F. Fong, H.T. Jen-sie, A further study of a testosterone-binding component of human pregnancy serum, J. Biol. Chem. 244 (1969) 1373–1380. [89] J. Chen, D.S. Hage, Quantitative studies of allosteric effects by biointeraction chromatography: analysis of protein binding for low solubility drugs, Anal. Chem. 78 (2006) 2672–2683. [90] F. Zsila, Subdomain IB is the third major drug binding region of human serum albumin: toward the three-sites model, Mol. Pharm. 10 (2013) 1668–1682. [91] Z. Ling, L. Xiaoyan, Z. Haixia, Interaction of tamoxifen citrate with salmon sperm DNA and bovine serum albumin, Sciencepaper Online 12 (2010) 011. [92] P.F. Bruning, J.M.G. Bonfrèr, Free fatty acid concentrations correlated with the available fraction of estradiol in human plasma, Cancer Res. 46 (1986) 2606–2609. [93] U. Westphal, Serum albumin, in: F. Gross, M.M. Grumbach, A. Labhart, M.B. Lipsett, T. Mann, L.T. Samuels, J. Zander (Eds.), Steroid-Protein Interactions II. Monographs on Endocrinology, Springer, Berlin, Heidelberg 1986, pp. 8–22. [94] J.R. Brown, P. Shockley, Serum albumin: structure and characterization of its ligand binding site, in: P.C. Jost, O.H. Griffith (Eds.), Lipid-Protein Interactions, Wiley, New York 1982, pp. 25–68. [95] W.H. Pearlman, I.F.F. Fong, Steroid binding properties of some peptide fragments of bovine serum albumin obtained on peptic digestion, J. Biol. Chem. 247 (1972) 8078–8084. [96] I. Sjöholm, B. Ekman, A. Kober, I. Ljungstedt-Påhlman, B. Seiving, T. Sjödin, Binding of drugs to human serum albumin, Mol. Pharmacol. 16 (1979) 767–777. [97] D. Buttar, N. Colclough, S. Gerhardt, P.A. MacFaul, S.D. Phillips, A. Plowright, P. Whittamore, K. Tam, K. Maskos, S. Steinbacher, A combined spectroscopic and crystallographic approach to probing drug–human serum albumin interactions, Bioorg. Med. Chem. 18 (2010) 7486–7496. [98] I.G. Shah, D.L. Parsons, Human albumin binding of tamoxifen in the presence of a perfluorochemical erythrocyte substitute, J. Pharm. Pharmacol. 43 (1991) 790–793. [99] F. Yang, Y. Zhang, H. Liang, Interactive association of drugs binding to human serum albumin, Int. J. Mol. Sci. 15 (2014) 3580–3595.

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