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Feb 15, 2017 - 2 mV whereas HSA-MPNPs complex showed negative surface charge of around −20 ± 1 mV. The increase in the zeta potential of HSA−MPNPs ...

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Complexation With Human Serum Albumin Facilitates Sustained Release of Morin From Polylactic-Co-Glycolic Acid Nanoparticles Pooja Ghosh, Jayita Patwari, and Swagata Dasgupta* Department of Chemistry, Indian Institute of Technology Kharagpur, Kharagpur 721302, India S Supporting Information *

ABSTRACT: Understanding the interaction of proteins with nanoparticles has become an important area of research in biomedical and pharmaceutical fields. Morin is a flavonol which shows several properties including antioxidant, anticancer, and anti-inflammatory activities. However, the major limitation is its poor aqueous solubility. Therefore, morin-loaded polylacticco-glycolic acid (PLGA) nanoparticles (MPNPs) were prepared to improve the solubility of morin. The resulting MPNPs were characterized by spectroscopic and microscopic techniques. The nanoparticles were spherical with an average size of 237 ± 17 nm. UV−visible, fluorescence, and circular dichroism (CD) spectroscopy were employed to study the interaction of the MPNPs with human serum albumin (HSA). Our study revealed that a static fluorescence quenching mechanism was involved in the interaction between HSA and MPNPs. Hydrophobic interactions also play an important role in stabilizing the HSA-MPNP complex. CD results suggest that there is an alteration of the secondary structure of HSA in the presence of MPNPs. MPNPs exhibit antioxidant properties which are supported by the DPPH assay. We have further checked the effect of HSA on the antioxidant property of morin and MPNPs. HSA binding with MPNPs was also found to influence the in vitro release property of morin from MPNPs wherein a delayed release response is observed.

1. INTRODUCTION Over the past decade, polymeric nanoparticles (PNPs) have gained popularity in biomedical and pharmaceutical applications due to their increasing use in the field of drug delivery. PNPs, not only have the ability to deliver small bioactive molecules to the target site, but can also enhance its transport, increase its stability and bioavailability, control its release rate, therapeutic efficiency, and decrease its side effects. Morin (3,5,7,2′,4′-pentahydroxyflavone) is a polyphenolic compound (Figure 1a) which belongs to the flavonol group. It is widely present in red wine, herbs, and fruits.1−3 It possesses several activities including antioxidant, antimutagenesis, anti-inflammatory,4−6 antineoplastic, cardioprotective activities7,8 in addition to anticancer,9 and cell proliferation inhibition,10 etc. Despite its pharmacological and biological activities, its poor aqueous solubility is its main limitation. We have prepared morin-loaded poly lactic-co-glycolic acid (PLGA) NPs (MPNPs) to increase the solubility of morin. Poly lactic-co-glycolic acid (PLGA) has been chosen as a carrier in the preparation of nanoparticles as drug delivery systems in several biomedical applications. The advantages of choosing PLGA are that it is biodegradable, biocompatible,11 and nontoxic.12 Also, PLGA has been accepted by the US Food and Drug Administration (FDA) and European Medicine Agency (EMA) for human use.13 Hydrolysis of PLGA results in the formation of natural biodegradable metabolites (lactic acid and glycolic acid) (Figure 1b) that are metabolized in the body via the Krebs cycle and eliminated as carbon dioxide and water. For the © XXXX American Chemical Society

effective use of any therapeutic drug molecule or nanoparticles in the human body, it is important to understand their role in the bloodstream. The biodistribution, availability, and metabolism of therapeutic drugs are greatly affected by their interaction with proteins in blood. Human serum albumin (HSA), (Figure 2) is the major soluble protein in blood plasma. It is well-known for its versatile nature as it plays an important role in the transportation and distribution of several compounds, such as drugs, hormones, amino acids, and fatty acids.14 It is the most abundant protein in blood plasma and also maintains the colloidal osmotic pressure in blood. HSA is a heart shaped globular monomeric protein with a molecular weight of 66.5 kDa. It has a single polypeptide chain, consisting of 585 amino acid residues. Crystal structure analysis has revealed that HSA has three specific domains, I, II, and III, each containing two subdomains (A and B).15,16 There are two main drug binding sites in HSA referred to Sudlow site 1 which is the warfarin binding site is located in subdomain IIA. Sudlow site 2, located in subdomain IIIA is referred to as the benzodiazepine binding site.17 Drugs like ibuprofen and diazepam are selective probes for this site. The single tryptophan residue (Trp 214) of HSA is present in subdoman IIA. HSA is also used as a biomarker in various diseases as its nontoxic and anti-immunogenic property makes Received: August 24, 2016 Revised: February 15, 2017 Published: February 15, 2017 A

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investigate drug-loaded NPs in the body for clinical applications, the effect of HSA binding on drug release must be examined. In the current study, we have prepared morin-loaded PLGA NPs that have been characterized by spectroscopic and microscopic analyses. In addition the interaction of MPNPs with HSA has been investigated to explore the influence on the release efficiency of morin. Several biophysical techniques, such as UV−visible, fluorescence, and circular dichroism (CD) spectroscopy have been used to study the interaction of the prepared NPs with HSA. The in vitro release behavior of morin from MPNPs and HSA bound MPNPs has been evaluated using a dialysis technique. To elucidate the role of HSA on the release of morin from MPNPs, we have further performed an in vitro release study to observe the release property of morin from HSA bound MPNPs. The HSA binding influenced the drug release property of NPs, which makes it significant for these drug carriers to be further explored for in vivo efficacy.

2. MATERIALS AND METHODS Materials. PLGA (poly lactic coglycolic acid), poly(vinyl alcohol) (PVA), morin hydrate, human serum albumin (HSA), and 8-anilino-1-naphthalene sulfonic acid (ANS) were purchased from Sigma Chemical Co. (St. Louis, USA). All organic solvents were of HPLC grade. All the chemicals were of analytical grade and used as received. Milli-Q grade water was used for the preparation and characterization of NPs. The concentration of HSA was determined spectrophotometrically (UV-1800 Shimadzu) using a molar extinction coefficient of 35 500 M−1 cm−1 at 280 nm. Phosphate buffer (20 mM) pH 7.0 was used to study the interaction of HSA and MPNPs. Methods. Preparation of PVA Solution. 0.5% PVA solution was prepared by heating at 85 °C on a magnetic stirrer until the PVA dissolved. Preparation of Morin-Loaded PLGANPs (MPNPs). Morinloaded PLGA NPs were prepared using a solid in oil in water (S/O/W) emulsification technique33 with slight modifications. In the first step 50 mg of PLGA was dissolved in 1.5 mL ethyl acetate which was then incubated at room temperature for 2 h and 10 mg of morin added to the PLGA solution. The mixture was sonicated for 2 min in a bath sonicator (Oscar Ultrasonic Cleaner, Microclean-101). The organic phase (morin and PLGA in ethyl acetate) was then added dropwise to 3 mL of PVA solution (aqueous phase) by continuous stirring. The stirring was continued for 24 h. The prepared NPs were then centrifuged and washed at least three times with Milli-Q water. PLGA nanoparticles were also prepared using a similar technique. Characterization of NPs. UV−vis Spectrophotometric Study. PLGA, morin, and morin-loaded PLGA NPs were characterized by UV−vis spectroscopy (UV-1800, Shimadzu). UV−vis studies were performed at 25 °C in the range of 200− 600 nm. Fourier Transform Infrared Spectroscopy (FTIR) Study. FTIR spectra of morin and morin-loaded PLGA NPs were obtained from a Spectrum BX FTIR (PerkinElmer) equipped with a Lithium Tantalate (LiTaO3) detector and a KBr beam splitter at room temperature. The resolution used was 4 cm−1 and the scanning range was from 4000 to 400 cm−1. Field Emission Scanning Electron Microscopy (FESEM). Field emission scanning electron microscopy (FESEM) was used to determine the morphology of the nanoparticles. A drop

Figure 1. (a) Structure of morin hydrate. (b) Hydrolysis of PLGA to glycolic acids and lactic acids.

Figure 2. Cartoon representation of HSA (PDB id: 1AO6), Trp 214 is shown in red.

it a suitable candidate for drug binding.18,19 Interactions of bovine serum albumins with nanoparticles have been explored to observe possible applications of nanoparticles in drug delivery.20−25 Nanoparticles (NPs) can be administered into the body via different routes, such as inhalation, oral contact, and intravenous injection. The NPs enter the bloodstream and first interact with several biomacromolecules like serum proteins (primarily HSA), lipids, and nucleic acids. A conformational change of protein occurs due to the interaction of proteins with NPs which in turn can also affect the transport properties of albumins.26 HSA binding also affects the uptake and intercellular trafficking of NPs and it has a great impact on particle biodistribution, biocompatibility, and therapeutic efficiency.27,28 HSA is often coated on the surfaces of many nanoparticles for in vivo treatment29,30 which changes the release property of drug, affects the targeted tissue and in turn results in the altered nanodrug efficacy.31,32 To further B

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Fluorescence Spectroscopy. The interaction between HSA and MPNPs was monitored by a fluorimetric titration. Fluorescence experiments were performed in a Horiba Jobin Yvon spectrofluorometer (Fluoromax 4) using a 1 cm quartz cell, using an excitation wavelength of 295 nm. The emission spectra were recorded from 305 to 500 nm. The bandwidth of excitation and emission slits were 5 nm with an integration time of 0.3 s. A 2.5 mL solution of 2 μM HSA was titrated with successive addition of morin-loaded PLGA NPs. Each spectrum was subtracted from the corresponding blank. ANS Displacement Study. HSA (2 μM) was saturated with 6 μM ANS35 in 20 mM phosphate buffer of pH 7.0. The mixture was then incubated for 1 h in the dark. After 1 h incubation, the mixture was titrated with successive addition of morin-loaded PLGA NPs. The concentration of ANS was determined spectrophotometrically using its molar absorption coefficient of 4950 M−1 cm−1 at 350 nm.36 The excitation wavelength for ANS-bound HSA was set at 375 nm and emission was observed at 474 nm. Circular Dichroism. Far-UV CD measurements were carried out on a Jasco-810 automatic recording spectrophotometer, using a 0.1 cm path length cell at 25 °C. The concentration of HSA used for the experiment was 2 μM in 20 mM phosphate buffer of pH 7.0. Four different sets of solutions were prepared containing HSA and MPNPs in the range 1:0 to 1:3. The spectra were recorded in the region 190−240 nm with a response time of 4 s and scan speed of 50 nm/min. At least three scans were accumulated for each spectrum. The secondary structure was analyzed using DICHROWEB, an online server for protein secondary structure analyses from CD spectroscopic data. In Vitro Release Study of Morin from MPNPs and HSABound MPNPs. An in vitro release study of morin from MPNPs and HSA-bound MPNPs were determined by a dialysis technique. Briefly, MPNPs were dispersed in 500 μL of phosphate buffer saline (PBS) (pH 7.4) and taken in a dialysis bag with a molecular weight cut off of 12.6 kDa. The dialysis bag was then tied and kept inside 5 mL PBS (pH 7.4). At fixed time intervals 1 mL aliquots were withdrawn from the solution in which the dialysis bag was kept and the same amount of buffer added. Concentration of the release media was determined by measuring the absorbance at 351 nm which is the maximum absorption wavelength of morin (ε351 = 13 100 M−1 cm −1) using UV−vis spectroscopy (UV-1800, Shimadzu). A mixture of HSA and MPNPs were added to the dialysis bag and the release of morin from HSA-bound MPNPs was determined as described above. DPPH (2,2-Diphenyl-1-picrylhydrazyl) Assay. DPPH assay37 was carried out in order to find out the antioxidant activity of the MPNPs. DPPH solution was prepared in methanol and was protected from light using aluminum foil. Morin at various concentrations (300 μL), MPNPs as well as the standard compound (ascorbic acid) were prepared in methanol. DPPH solution (300 μL) was then added to each of them. DPPH in methanol was treated as the control for our study. The samples were incubated for 30 min in the dark. Absorbance was taken at 517 nm (absorption maxima of DPPH) on a UV−vis spectrophotometer (UV-1800, Shimadzu). HSA was then added to morin and MPNPs and DPPH assay was carried out as described above to observe the effect of HSA on the antioxidant property of morin and MPNPs. The percentage of scavenging which is a measure of the antioxidant activity was calculated using the following expression

of sample was mounted on a glass slide, air-dried, and then scanned in a NOVA NANOSEM 450 operating at 10 kV. Atomic Force Microscopy (AFM). The morphology of the NPs was also monitored by atomic force microscopy (AFM) using Agilent Technologies, Model 5500. A drop of sample was deposited on freshly cleaved mica foil. The mica foil was then allowed to dry in air and then scanned. The images were taken in tapping mode using a silicon probe cantilever of 215−235 μm length, at a resonance frequency of 146−236 kHz, and a force constant of 21−98 N/m. Dynamic Light Scattering (DLS). The size of the NPs was determined using DLS. NPs were dispersed in water and the average size of the NPs measured using a Malvern Nano ZS instrument employing a 4 mW He−Ne laser (λ = 632 nm), with a scattering angle of 173°. Encapsulation Efficiency. The morin content of the prepared NPs was determined by UV−vis spectroscopy (UV1800, Shimadzu). The NPs were centrifuged and the amount of morin (w) present in the clear supernatant calculated by taking the absorbance of the supernatant at 351 nm. A standard calibration curve of concentration versus absorbance was plotted for this purpose. The amount of morin in the supernatant (w) was then subtracted from the total amount of morin added initially (W). Effectively, (W − w) will give the amount of morin entrapped within the NPs. The percentage of encapsulation is given by,

W−w × 100 W

(1)

Hemolytic Assay. A hemolytic assay was performed to find out the toxicity of the MPNPs. Fresh blood was collected and centrifuged at 3600 rpm for 10 min. The red blood cells (RBC) were separated from plasma and buffy coat. PBS (pH 7.4) was used to wash it five times. The morin-loaded PLGA NPs was then incubated with RBC suspensions (1% hematocrit) at 37 °C for 40 min. The reaction mixtures were again centrifuged at 3600 rpm for 10 min and the absorbance of the supernatant was taken at 540 nm which is the absorption maxima of hemoglobin. The percentage of hemolysis was calculated using the following formula, 34

A sample − A negative control A positive control − A negative control

× 100 (2)

where, Asample is the absorbance of the sample, Anegative control is the absorbance of the negative control (PBS), and Apositive control is the absorbance of the positive control (RBC in water). HSA and Morin-Loaded PLGA NPs (MPNPs) Interaction Studies. Zeta Potential Measurements. The surface charge of MPNPs and HSA-MPNPs complex were measured by means of zeta potential measurement. The measurement was carried out using Malvern ZetaSizer Nano ZS instrument. The analysis was performed at a scattering angle of 173° at 25 °C. Each reported value is the measurements of 20 runs and the data here represents the average of three independent readings. Diluted samples were taken in a cuvette and zeta potential measured using instrument software. UV−vis Spectroscopy. The UV−vis absorption spectra were recorded at room temperature on a UV spectrophotometer (UV-1800 Shimadzu) equipped with 1.0 cm quartz cells. The scanning range was 250−500 nm. A 1 mL of 5 μM of HSA in 20 mM phosphate buffer (pH 7.0) was titrated with successive addition of MPNPs. C

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× 100

1458, 1306, and 972 cm−1. Several researchers have used FTIR spectroscopy to study the encapsulation of bioactive molecules into NPs.38,39 We have also observed that the specific functional groups of PLGA in the prepared NPs have almost the same characteristic peaks of pure PLGA. Our FTIR data also suggests that the peaks in the NPs result from the superposition of PLGA and morin, from which, we can conclude that morin is entrapped within the PLGA matrix and the chemical structure of morin does not change after the encapsulation. This is in agreement with a previous result from this laboratory, where fisetin-loaded human serum albumin (HSA) nanoparticles40 and epicatechin/morin-loaded HSA nanoparticles41 exhibited the same characteristics as fisetin, epicatechin, and morin as expected on account of encapsulation. Particle size, size distribution, and morphology play a key role in nanoparticle systems that can affect drug loading, drug release, and stability of the drug-loaded NPs. The morphology of the prepared NPs was monitored by field emission scanning electron microscopy (FESEM). It was observed that the particles were homogeneous, smooth, and spherical in shape which is further confirmed by atomic force microscopy (AFM). The topography of the particles was visualized by AFM. The FESEM and AFM images are shown in Figure 5. The smallest capillaries in the human body are ∼5−6 μm in diameter, thus particles distributed into the bloodstream must be much smaller than 5 μm. In addition they should not form aggregates. In this regard, nanoparticles provide a large number of advantages over microparticles as the sizes of the particles are of importance. NPs have already been used to deliver drugs to target sites for cancer therapeutics42 since NPs can easily enter into the body and penetrate tissues and target tumors. Dynamic light scattering (DLS) was used to determine the sizes of the NPs. It was observed that the average size of PLGA NPs is 131 ± 9 nm and there is a ∼ 81% increase in size of the morin-loaded PLGA NPs to 237 ± 17 nm. Figure 6 shows the DLS profile of the prepared NPs. The sizes of the NPs are further confirmed by FESEM and AFM images. From FESEM images the sizes obtained were almost similar to that obtained from DLS measurements. Figure 5 shows the AFM height profile of nanoparticles from which we observe that the sizes of the nanoparticles are ∼230 nm. We have checked the stability of the nanoparticles after 6 months of storage using field emission scanning electron microscopy (FESEM) and dynamic light scattering (DLS) studies. Figure 7 shows the FESEM images and DLS measurements of morin-loaded PLGA NPs after 6 months of storage to check the stability. From DLS measurements, we have observed that the sizes of the nanoparticles are almost similar even after 6 months of storage. There is only 5% decrease in the size of the nanoparticles. FESEM images showed that no distinct change occurs in the shape of the nanoparticles which indicates that the nanoparticles are stable. Encapsulation Efficiency. The encapsulation efficiency depends on various factors, such as the nature of the polymer used, encapsulating compounds, medium of NP synthesis, etc.38 UV−vis spectroscopy was used to calculate the encapsulation efficiency and the amount of morin present in the supernatant was calculated with the help of a calibration curve. Our study showed ∼97% encapsulation efficiency of morin-loaded PLGA NPs. Hemolytic Assay. Hemolysis causes damages to RBCs (Figure 8). The toxicity of the NPs must be evaluated to ensure

(3)

Statistical Analysis. The data presented in our study have been expressed as mean ± standard deviation. All experiments have been conducted at least three times.

3. RESULTS AND DISCUSSION Preparation and Characterization of NPs. PLGA NPs and morin-loaded PLGA NPs were prepared by a modified solvent emulsification method. In this method the organic phase solution containing morin and PLGA serves as the oil phase (O) and PVA aqueous solution serves as the water phase (W) in an O/W emulsion. Physical sonication induces reduction of the size of nanodroplets in this emulsion. UV− vis spectroscopy was used to characterize morin-loaded PLGA NPs. Figure 3 shows the absorption spectra of morin in ethanol

Figure 3. UV−vis absorption spectra of morin in ethanol and morin, morin-loaded PLGA NPs, and PLGA NPs in buffer.

and morin, morin-loaded PLGA NPs, and PLGA NPs in buffer. Morin in ethanol shows two major absorption peaks at 351 and 264 nm. The peak at 351 nm is due to the cinnamoyl moiety and the peak at 264 nm is due to the benzoyl moiety. However, in the presence of buffer the peak at 351 nm of morin is shifted to 390 nm. After the formation of NPs, the absorbance of the two peaks of morin decrease, but the appearance of both the peaks of morin in the prepared NPs implies that the chemical structure of morin does not change after encapsulation. In this experiment, we have used the buffer as the control since we want to observe the change in the spectra after the formation of NPs. Each spectra has been corrected with respect to the corresponding buffer solution (control). Figure 4 shows the FTIR spectra of PLGA, morin, and MPNPs. For PLGA, the CO stretching frequency appears at 1757 cm−1 and the characteristic peaks at 1044 cm−1and 1248 cm−1 are due to the C−O stretching frequency. For morin, a band of strong intensity appears at 3374 cm−1 due to the − OH group. The bands at 1613, 1509, and 1461 cm−1 are due to C C stretching vibration in the aromatic ring, the −C−OH deformation vibration appears at 1307 cm−1. The bands at 1250 and 973 cm−1 involve the −C−OH stretching vibration. The spectra of morin-loaded PLGA NPs shows the characteristic peaks of morin with almost negligible shifts at 1621, 1510, D

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Figure 4. FTIR spectra of PLGA, morin, and morin-loaded PLGA NPs.

surface charge of around −20 ± 1 mV. The increase in the zeta potential of HSA−MPNPs complex is due to the adsorption of HSA on the surface of the MPNPs. Instantaneous protein adsorption leads to a slight increase in the particle size and alters the zeta potential of NPs toward the positive direction. HSA molecules have an average size of ∼36 nm.44 There is a ∼17% increase in the size of the HSA−MPNPs complex compared to MPNPs. The nanoparticle size increase is also an indication of the HSA adsorption onto the nanoparticle surface which is in agreement with an earlier report.45 UV−vis Spectroscopy. UV−vis spectroscopy was used to observe the structural changes in the protein. The spectra were recorded by continuous addition of morin-loaded PLGA NPs into HSA keeping the concentration of HSA constant. Figure 9 corresponds to the absorption spectra of HSA in the presence of morin-loaded PLGA NPs at different concentrations. As shown in Figure 9, HSA shows a strong absorption band at

the minimum destruction of red blood cells. There should not be any interaction between the NPs and blood constituents during administration of NPs into the body. Therefore, it is essential to perform a hemolytic assay. According to ISO/TR 7405−1984(f),43 the samples showing less than 5% of hemolysis are nonhemolytic, 5−10% slightly hemolytic, and values greater than 10% highly hemolytic. In our present study the hemolytic percentage of morin-loaded PLGA NPs is ∼2.8% which suggests that the NPs showed no hemolytic activity, i.e., NPs do not damage RBCs. Effect of MPNPs after Interacting with HSA. Zeta Potential Measurements. HSA has a pI of 4.9 which means that at the experimental pH of 7.4, HSA possesses a negative charge. At pH 7.4, the − OH group (pKa 5.2) located at 2′ position in the B ring of morin is ionized. Morin-loaded PLGA NPs (MPNPs) showed negative surface charge of around −27 ± 2 mV whereas HSA-MPNPs complex showed negative E

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Figure 5. (a) FESEM and (b) AFM images with height profile of PLGA NPs and morin-loaded PLGA NPs.

already shown that there is an appearance of the peak of morin (at 390 nm) in the prepared NPs which implies that the chemical structure of morin does not change after encapsulation. The addition of NPs also resulted in the formation of a new peak between 300 and 400 nm (∼390 nm) which indicate that there is a ground state complex formation between NPs and HSA, i.e., there is a strong interaction between the HSA and NPs through a ground state complex formation, which is caused by the partial adsorption of HSA on the surface of the nanoparticles. Our result is also consistent with the results obtained from previous reports.46,48 Fluorescence Spectroscopy. The effect of morin-loaded PLGA NPs on quenching of HSA in 20 mM phosphate buffer (pH 7.0) is shown in Figure 10. HSA exhibits strong fluorescence emission at 343 nm upon excitation at 295 nm because of its single tryptophan residue (Trp 214). The

Figure 6. DLS measurements of (a) PLGA NPs and (b) morin-loaded PLGA NPs.

∼280 nm and the absorbance at 280 nm gradually increased with a significant blue shift to 270 nm with increase in the concentration of morin-loaded PLGA NPs.46−48 We have F

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fluorescence maximum intensity of HSA decreases progressively with consecutive addition of NPs which indicates that NPs are able to quench the fluorescence of HSA. We have also observed a significant blue shift from 343 to 338 nm in the emission maxima of HSA on addition of NPs which suggests that the fluorophore of HSA (Trp 214) shifts to a more nonpolar environment. This blue shift is a result of hydrophobic interactions with the NPs. This study suggests that the NPs bind near Trp 214 (site 1, subdomain IIA) of the HSA. Depending on the type of interaction between the quencher and fluorophore, fluorescence quenching occurs by two mechanisms: static quenching and dynamic quenching. Static quenching involves ground state complex formation between the quencher and the fluorophore. Dynamic quenching is otherwise referred to as collision quenching and occurs when there is a contact between the quencher and fluorophore in the excited state. The quenching mechanism was analyzed by using the following Stern−Volmer equation49

Figure 7. (a) FESEM images and (b) DLS measurements of morinloaded PLGA NPs after 6 months of storage to check the stability.

F0 = 1 + Kqτ0[Q ] = 1 + K sv[Q ] F

(4)

where F0 and F denote the relative fluorescence intensities in absence and presence of the quencher respectively, Kq is the bimolecular quenching constant, τ0 is the lifetime of the fluorophore in absence of quencher, [Q] is the quencher concentration, and Ksv is Stern- Volmer quenching constant. The Stern−Volmer plot of the quenching of HSA fluorescence by NPs is shown in Figure 10. The result showed that the plot of F0/F versus [NPs] is linear. The calculated Ksv and kq values were found to be 0.015 × 103 L g−1 and 0.015 × 1011 L g−1 s−1, respectively. The lifetime of HSA is 10−8 s.50 The linear nature of the Stern−Volmer plot (Figure 10b) is in agreement with the formation of a ground state complex (confirmed from the UV−vis studies) and indicates that the quenching mechanism is static in nature. The number of binding sites and binding constant of the interaction of NPs with HSA can be determined using the following equation51 log

ΔF = n log[Q ] + log Kb F

(5)

where ΔF = F0 − F, F0 and F stand for the fluorescence intensities of protein in absence and presence of quencher, n is the number of binding sites, and Kb the equilibrium binding constant. The plot of log(ΔF/F) versus log[Q] is linear (Figure 10c) and the values of “n” and “Kb” were obtained from the slope and intercept, respectively. From the plot, the binding constant of the NPs to HSA was evaluated to be 0.026 × 103 L g−1 and “n” is 0.852. ANS Displacement Study. 8-Anilino-1-naphthalenesulfonic acid (ANS) is a fluorescent hydrophobic probe which binds to the nonpolar region of proteins. Figure 11 shows that the fluorescence intensity of ANS bound HSA decreases on increasing the concentration of morin-loaded PLGA NPs. The displacement of ANS by morin-loaded PLGA NPs indicates that ANS and the NPs bind to a common site which is the hydrophobic pocket of HSA. This suggests that there is formation of a complex and that hydrophobic interactions play an important role in the process. CD Spectroscopy. Circular dichroism (CD) is a powerful technique to investigate the interaction of proteins with other molecules including nanoparticles as it primarily provides

Figure 8. Hemolytic effect of morin-loaded PLGA NPs.

Figure 9. Absorption spectra of human serum albumin in absence (bottom line) and presence (top lines) of morin-loaded PLGA NPs. G

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Figure 10. (a) Fluorescence quenching spectra of the HSA-MPNPs system. (b) Stern−Volmer plot for the HSA-MPNPs system. (c) Doublelogarithm plot for interaction of HSA with MPNPs in 20 mM phosphate buffer pH 7.0 at 273 K; λex = 295 nm.

HSA decreases from ∼52% to ∼46% upon binding with morinloaded PLGA NPs. There is also a slight increment in the βsheet and unordered structure. This suggests that the complexation between MPNPs and HSA induces a slight conformational change in the protein leading to a loss in its helical content. Figure 12 represents the CD spectra of the HSA−MPNPs complex. In Vitro Release of Morin from MPNPs and HSA-Bound MPNPs. The release behavior of morin of MPNPs and HSAbound MPNPs was monitored in vitro in phosphate buffer saline (PBS) for 120 h. The in vitro release profile of morin from MPNPs and the effect of HSA on its release behavior were shown in Figure 13. It was observed that there is a burst release of morin from MPNPs in the initial phase which may be due to the small fraction of morin present on the surface of the PLGA matrix. There is a sustained and controlled release of morin from 24 to 120 h. Diffusion of morin entrapped within the matrix resulted in the slow and sustained release of morin. This kind of biphasic release is consistent with previous studies.39 The experiment was continued until 120 h and the total release at the end of 120 h was found to be ∼78% for MPNPs. HSAbound MPNPs showed comparatively slower release of morin compared to that of the MPNPs. The major reason behind the slower release of morin from HSA-bound MPNPs is attributed to a steric hindrance effect. Figure 14 shows a schematic representation of the effect of HSA complexation on the release of morin from MPNPs. Our observation is in agreement with a previous report published by Tao et al.52

Figure 11. Displacement of bound ANS from HSA-ANS complex by MPNPs. [HSA] = 2 μM; [ANS] = 6 μM; λex = 375 nm. Arrows indicate the increase in concentration of MPNPs.

information about the secondary structure changes on interaction. CD spectra of HSA shows two negative bands at 208 nm (∏-∏*) and 222 nm (n-∏*) that are characteristic of the α-helical structure of the protein. In the analyses of secondary structural content of the protein, we have used DICHROWEB, an online server for protein secondary structure analyses using the SELCON3 analysis program. Since we have carried out far-UV CD measurements, the reference set used for this online server was Set 4 (optimized for 190−240 nm). It was observed that the α-helical content of H

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by Xie et al.53 that morin shows specific binding to HSA. We have also studied the interaction of morin with HSA by biophysical techniques including UV−vis spectroscopy, fluorescence spectroscopy and circular dichroism. From the UV− vis studies we have observed a significant red shift from 390 to 398 nm in the UV absorption band of morin (due to the cinnamoyl moiety) upon binding with HSA which indicates that there is a complex formation between HSA and morin. Our results are also in agreement with the observation reported by Xie et al.53 The fluorescence maximum intensity of HSA decreases gradually with successive addition of morin indicating that morin can quench the fluorescence of HSA. The addition of morin also results in a significant blue shift in the emission maxima of HSA which indicates that Trp 214 is shifted to a more nonpolar environment. CD results show that there is a slight conformational change in the protein after interaction with morin which results in a loss in its helical content. Figure 15 shows the UV−vis, fluorescence and CD spectra of HSA−morin complex. Thus, morin present on the surface of MPNPs is initially bound to the HSA present and the HSA−morin complex is in turn bound to the surface of the NPs. Instantaneous adsorption of HSA on MPNPs was also confirmed through zeta potential measurements where we have observed a distinct increase in the zeta potential of the HSA−MPNP complex. This protein adsorption causes a slight increase in the particle size and also alters the zeta potential of NPs toward the positive direction.45 Moreover a slight increase in the size of HSA−MPNPs complex compared to MPNPs also confirms the adsorption of HSA onto the nanoparticle surface. On the other hand, it is possible that the morin entrapped within the matrix of MPNPs can also be adsorbed by the complexed HSA on the surface. This causes a steric effect on the surface which restricts the release of morin from MPNPs that subsequently results in a comparatively slower release of morin. A similar observation was found for the release of mitoxantrone from pullulan nanoparticles in the presence of protein.52 To study the drug release rate, a number of mathematical models have been developed. The drug release kinetic profiles are generally studied by fitting the data into five different mathematical model equations which are (i) zero-order (cumulative percentage of drug released versus time), (ii) first-order (log cumulative percentage of drug remaining versus time), (iii) Higuchi (cumulative percentage of drug released versus square root of time), (iv) Korsmeyer−Peppas (log cumulative percentage of drug released versus log time), and (v) Hixson−Crowell (cube root of cumulative percentage of drug remaining versus time).54−58 In our case, the data were fitted according to zero-order, firstorder, Higuchi, and Korsmeyer−Peppas model equations to analyze the release profile of morin from MPNPs and HSAbound MPNPs. We have obtained a linear relationship (Figure S1) with high regression value (0.992 for MPNPs and 0.993 for HSA bound MPNPs) from Higuchi square root model equation (shown in Supporting Information) which implies that release occurs through a diffusion mechanism. According to Korsmeyer−Peppas model (log cumulative percentage of drug released versus log time) equation: F = K4tn (F represents the cumulative percentage of drug released in time t, K4 is the release constant, and n is the diffusional exponent) when n ≤ 0.45, the release mechanism follows a Fickian diffusion mechanism (Case I diffusion).59 When, the n

Figure 12. CD spectra of HSA-MPNPs complex obtained in 20 mM phosphate buffer pH 7.0 at room temperature. (a) represents the CD spectra of HSA and (b)−(d) represent the CD spectra of HSAMPNPs complex (1:1, 1:2, and 1:3 molar ratio).

Figure 13. In vitro release study of morin from morin-loaded PLGA NPs and HSA-bound morin-loaded PLGA NPs for 120 h.

Figure 14. Schematic representation of the effect of HSA complexation on the release of morin from MPNPs.

The comparative studies carried out included the in vitro release experiment by taking MPNPs into the dialysis bag in one case and adding HSA to the dialysis bag containing MPNPs in the second case. In the first case the initial burst release of morin from MPNPs observed is most likely due to the small amount of morin present on the surface of the matrix. In the second case, a relatively slower release of morin occurred. This indicates that the morin present on the surface of MPNPs is initially bound to the HSA present. It has been already reported I

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Figure 15. (a) UV−vis absorption spectra of morin in the presence of HSA. (b) Fluorescence quenching spectra of the HSA-morin complex. (c) CD spectra of HSA−morin complex obtained in 20 mM phosphate buffer pH 7.0 at room temperature. (i) represents the CD spectra of HSA and (ii)− (iv) represent the CD spectra of HSA−morin complex (1:1, 1:2, and 1:3 molar ratio).

value is between 0.45 to 0.89, it indicates a combination of both diffusion and erosion drug release mechanisms which are basically non-Fickian type of release (anomalous transport). On the other hand, when n ≥ 0.89, the release involves an erosion mechanism (Case II Transport). In case of MPNPs, we have obtained an n value less than 0.45 which confirms the diffusion mechanism (Fickian diffusion). However, in case of HSA bound MPNPs the n value is between 0.45 to 0.89 which indicates a combination of both diffusion and erosion release mechanism. Thus, we can conclude that the release of morin from MPNPs is mainly controlled by a diffusion mechanism whereas a combination of both diffusion and erosion release mechanisms are involved in the release of morin from HSA bound MPNPs. A detailed discussion has been given in Supporting Information. DPPH Assay. The DPPH assay is a stable free radical method and is an easy, rapid, and sensitive way to determine the antioxidant activity of specific compounds. DPPH contains one odd electron and gives a strong absorbance at 517 nm (purple color). The principle of the DPPH method is based on the

reduction of DPPH in the presence of antioxidant compounds, such as a hydrogen donor. As a result the odd electron of DPPH is paired off and the absorbance decreases. Morin is an antioxidant compound and the sample compounds reduce the color of DPPH due to its hydrogen donating ability. The DPPH scavenging potential of morin and MPNPs in the presence of HSA has also been investigated in this study. Our result shows that morin exhibits strong DPPH scavenging activity whereas HSA significantly masks the DPPH scavenging potential of morin. However, the DPPH scavenging potential of HSA bound MPNPs is higher compared to that of the HSA morin complex. Figure 16 shows the result of the DPPH assay where the DPPH radical scavenging power was found to be 83.5 ± 1.3% for MPNPs (100 μg/mL), 94 ± 1.5% for morin (100 μg/mL) alone, 68.3 ± 1.4% for HSA-bound morin, and 75.8 ± 3.5% for HSA bound MPNPs. MPNPs showed lower DPPH radical scavenging activity than morin alone which may be due to the slow release of morin from MPNPs during its incubation. J

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concentration of MPNPs which indicates the interaction between HSA and MPNPs through a ground state complex formation. The NPs are also able to quench the fluorescence of HSA via a static quenching mechanism. Further a significant blue shift from 343 to 338 nm was observed in the emission maxima of HSA on addition of NPs from fluorescence spectroscopy. This observation leads us to conclude that the fluorophore of HSA (Trp 214) shifts to a less polar environment. This blue shift is a result of hydrophobic interactions with the NPs. Therefore, hydrophobic interactions also play a crucial role in the HSA-MPNPs complex during the binding process. In designing proper drug delivery systems, it is important to control the release properties of the drug. If too much of the drug is released at once it will be harmful, while a lower amount of drug released will limit the effectiveness of the drug. Therefore, delivery of drugs at the optimal dosage for a certain period of time will make the drugs more effective and more potent and will be beneficial for clinical purposes. A maximum cumulative drug release over time is thus preferable. HSA binding has also a great impact on the drug release properties of NPs, which is significant for these drug carriers to be further investigated for in vivo efficacy. It was observed that there is an initial burst release followed by a sustained release of morin from morin-loaded PLGA NPs (MPNPs), however a comparatively slower release of morin was observed in case of HSA bound MPNPs. Thus, HSA bound MPNPs help in enhancing the sustained release property of morin. Since sustained and controlled release of drugs at the specific target areas of the body is an important criterion of suitable drug delivery systems, we believe that this property is significant for these therapeutic molecules to be further explored for in vivo efficacy. Investigations leading to a further understanding of the interactions as well as the increased ability of sustained release of potential drug molecules are of immense therapeutic importance.

Figure 16. Histogram of DPPH radical scavenging activity of ascorbic acid, morin, morin-loaded PLGA NPs, morin in the presence of HSA, and morin-loaded PLGA NPs in the presence of HSA.

Significance of Our Study. The biodistribution, availability, and metabolism of therapeutic drugs/bioactive molecules strongly depend on their interaction with plasma proteins present in blood. Human serum albumin (HSA), one of the most studied serum proteins is well-known for its ability to bind a wide range of drugs and has been extensively used for drug− protein interaction studies as they are capable of transporting reversibly bound drugs to their specific target sites via drug− protein complex formation. For the effective therapeutic use of nanoparticles, nanoparticle and plasma protein interactions have become important topics of research. The biodistribution of the nanoparticles throughout the body is also influenced by binding with the plasma proteins. When nanoparticles enter the bloodstream the encounter with proteins present in plasma is inevitable and the protein immediately binds to the surface of the nanoparticles. Therefore, the binding interaction of NPs with HSA needs to be understood for their safe use and therapeutic applications in future. There exists a significant amount of research focusing on the interaction of nanoparticles with proteins. In some of the earlier reports, researchers have discussed about the interaction of human serum albumin with carbon nanoparticles, titanium oxide nanoparticles, ZnS nanoparticles, and silver nanoparticles, etc.60−63 All the studies relate to interactions between the NPs and HSA through spectroscopic techniques. However, not much literature is available showing the effect of HSA on the release properties of NPs.52 To further utilize the drug-loaded NPs in the body for clinical applications, it is therefore necessary to investigate the effect of HSA binding on its drug release property. The interaction between morin and HSA at the molecular level has previously been investigated using spectroscopic techniques.53 The specific binding of morin to HSA showed a clear red shift in the UV absorption band of morin upon binding to HSA which confirmed the formation of an HSA-morin complex. Further the fluorescence data indicated the presence of a specific binding site on HSA for morin and the interaction between morin and HSA also results in a distinct reduction of the protein α-helix and β-sheet structures. We have investigated the interaction of HSA with morin-loaded PLGA NPs (MPNPs) in our present work. We have observed a significant blue shift for HSA from 280 to 270 nm with increase in the

4. CONCLUSION In the present study, we have prepared morin-loaded PLGA NPs to increase the solubility and bioavailability of morin. The NPs were found to be smooth and spherical with an average diameter of 237 ± 17 nm. The hemolytic assay shows that the NPs are not toxic toward RBC. Further the interaction of the NPs with HSA has been studied by several biophysical techniques. The fluorescence intensity was found to decrease with increase in the concentration of NPs which illustrates that the NPs are able to quench fluorescence of HSA. During the binding process, hydrophobic interactions play a significant role in the HSA−MPNP complex. It was found that the fluorescence of HSA was quenched via a static mechanism. The in vitro release property of morin from MPNPs has also been facilitated after the binding of HSA with MPNPs. Therefore, our result indicates that MPNPs act as a sustained release carrier for morin and HSA-bound MPNPs help in enhancing its sustained release property.



ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcb.6b08559. The mathematical model equations explaining the possible in vitro release profiles and Higuchi model plot K

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of morin from (a) MPNPs and (b) HSA bound MPNPs Korsmeyer-Peppas model plot of morin from (c) MPNPs and (d) HSA bound MPNPs (PDF)

AUTHOR INFORMATION

Corresponding Author

*Tel.: +91-3222-283306; Fax: +91-3222-282252; E-mail: [email protected] ORCID

Swagata Dasgupta: 0000-0003-2074-1247 Notes

The authors declare no competing financial interest.



ACKNOWLEDGMENTS S.D. is grateful to Science and Engineering Research Board (SERB) for partial funding. The authors would like to acknowledge the Central Research Facility, IIT Kharagpur for providing experimental facilities. The authors are grateful to Professor Nilmoni Sarkar (Department of Chemistry, IIT Kharagpur) and his group for providing DLS facilities. The authors would like to acknowledge Professor Koel Chaudhury (School of Medical Science and Technology, IIT Kharagpur) and her group for providing zeta potential facilities. P.G. thanks CSIR, New Delhi for her fellowship.



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