Effect of cucurbit[7]uril on DPPC-containing liposomes: Interactions with the lipid bilayer

Abstract

Liposomes, which are bilayer lipidic nanocarriers, have been utilized in many pharmaceutical applications to enhance the solubility and therapeutic index of drugs. Liposomes have also been used as carriers for smaller drug carriers, such as cucurbiturils, to achieve a more controlled release of the drug into the targeted site in the body. In this study, we investigated the effects of cucurbit[7]uril, a macrocyclic organic compound, on the integrity of liposome lipid membranes. The average liposome size, measured by dynamic light scattering, increased with increasing concentrations of cucurbit[7]uril. In addition, fluorescence spectroscopy was used to calculate an association constant (K_a) between cucurbit[7]uril and cholesterol of 3 $\times {10}^6/\mathrm{M}$ . This high K_a value demonstrated the ability of cucurbit[7]uril to reduce liposome stability by extracting cholesterol molecules from the lipid bilayer. Thermogravimetric analysis demonstrated the localization of cucurbit[7]uril molecules on the surface of the liposomes. As the concentration of cucurbit[7]uril increased, the thermal stability increased, i.e. the mass loss of the liposomal suspension decreased. The biocompatibility of cucurbit[7]uril was also investigated using a hemolysis test on human red blood cells. In conclusion, the current study is the first to explain the relationship between lipid membranes and cucurbit[7]uril. The results of this study can be used to develop a new drug delivery system comprising liposomes and cucurbit[7]uril.

Introduction

Recent decades witnessed many advances in drug delivery systems particularly through the use of nanoparticles, cucurbiturils (CBs), and cyclodextrins (CDs). These systems have advanced the delivery of pharmaceutical agents to the disease-affected sites, offering improved therapeutic outcomes and reduced side effects.^1–4 Among these, liposomes have emerged as a main platform due to their ability to encapsulate both hydrophobic and hydrophilic molecules. Encapsulation of active drugs in liposomes enables specific targeting and drug localization. For example, doxorubicin encapsulated in PEGylated liposomes became the first FDA-approved liposomal formulation with anticancer activity. ⁵

Liposomal drug nanocarriers are favored over other nanoformulations due to higher biocompatibility, the ability to enhance drug solubility, and targeted delivery while reducing toxicity.^3,6 These sophisticated properties of liposomes aid in improving the therapeutic index, establishing liposomes as a cornerstone in modern chemotherapy.^7,8 Furthermore, recent innovations in developing multifunctional liposomes, which incorporate combined targeting mechanisms, represent advanced strategies for targeted treatments.^9,10

CBs are a class of macrocyclic compounds synthesized by acid-catalyzed condensation reaction between glycoluril and formaldehyde. ¹¹ CBs are versatile for use in drug delivery and other research areas because of their ability to form stable binary and ternary host–guest complexes. ¹² For example, cucurbit[7]uril (CB[7]) has been shown to selectively bind to insulin and β-amyloid, offering potential applications in Alzheimer's and diabetes research by inhibiting protein aggregation. ¹¹ Additionally, CBs have been explored for developing stable drug-in-CB-in-liposome formulations to improve drug delivery and control lipophilic drugs’ rapid release from liposomes. ¹³

Despite these advances in the use CBs and liposomes for drug delivery, the interaction of CBs with liposomes remains unclear and needs further investigation. To address this gap, this study aims to investigate the effects of CB[7] on the integrity and stability of liposomes. Conventional liposomes composed of 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and cholesterol (Chol) were prepared using the thin-film hydration technique. The prepared liposomes were characterized in terms of their average size, zeta potential (ξ), morphology, and thermal behavior. The ability of CB[7] to act as a host for Chol and DPPC was also explored using fluorescence spectroscopy. Furthermore, the permeability of the liposomal membrane was assessed by monitoring the release of the fluorescent dye (calcein) in the presence of different concentrations of CB[7]. Finally, the biocompatibility of CB[7] was evaluated using a hemolysis assay with human red blood cells (RBCs). Elucidating the mechanisms underlying the CBs–liposomes interactions can contribute to the development of stable and efficient drug delivery systems, while also providing deeper insights into the interaction of CBs with cell membranes.

Materials and methods

Materials

CB[7] was supplied by Dr Anthony Day, University of New South Wales (Australia), and used as received. Deuterium oxide (D₂O; 99 at% D) and deuterated chloroform (CDCl₃) were purchased from Sigma-Aldrich, and deionized water was prepared using an Osmo Aries device (CA, USA). High-performance liquid chromatography (HPLC) isocratic-grade chloroform stabilized with ethanol was purchased from CARLO ERBA reagents (Sabadell, Spain). Pierce^® radio immunoprecipitation assay (RIPA) buffer was obtained from Thermo Scientific (Waltham, MA, USA). Chol and DPPC were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL, USA). Phosphate-buffered saline (PBS; pH 7.4, 1×) was obtained from Gibco Thermofisher (Paisley, UK). Calcein was purchased from Sigma-Aldrich, Inc. (Baden-Württemberg, Germany). Absolute ethanol (HPLC-grade, 99.9%) was obtained from Carbon Group (Cork, Ireland).

Preparation of liposomes

Liposomes were prepared using the thin-film hydration–extrusion method. ¹⁴ Briefly, 4.77 $\pm 0.01$  mg DPPC and 1.35 $\pm 0.01$  mg Chol (molar ratio = 65:35) were weighed (Cubis^® Sartorius laboratory balance, Göttingen, Germany) and dissolved in 3–4 mL of HPLC-grade chloroform. The organic solvent was then evaporated under a reduced pressure of 100 mbar (IKA RV 05 Basic Rotary Evaporator attached to a BÜCHI Vacuum Pump V-700, Essen, Germany) at 20 r/minute in a 30 °C water bath until a transparent and homogeneous dry thin film was formed. The thin film was then hydrated with 2 mL PBS (pH = 7.4) at a temperature of 50 °C for 30 minutes with robust vortexing every 3–4 minutes. ¹⁵ Then, the vesicle suspension was extruded 13 times in succession through a polycarbonate membrane (Nuclepore, Track-Etch Membrane, Filtration products, 100 nm, Whatman^®, Cleves, OH, USA) using a mini-extruder (Avanti Polar Lipids, Inc., Alabaster, AL, USA) at 25 °C. The nanovesicles were passed through the same size 13 times to obtain a final liposomal suspension of the desired size with a low polydispersity index (PDI). The liposome suspension was stored at 4 °C for further analysis.

Dynamic light scattering measurements

The mean hydrodynamic diameter, PDI, and zeta potential (ξ) of the liposomes were measured via dynamic light scattering (DLS)^16,17 using a Nano ZS (Malvern Instruments, UK) before and after incubation with CB[7] for 2 and 24 hours at room temperature. ¹⁶ The liposomes were diluted with PBS to obtain a suitable count rate of between 200 and 300 K. The measurements were performed in triplicate at 25 °C for each sample. The standard deviation (SD) was calculated to estimate the repeatability of the measurements. ¹⁷

Fluorescence measurements

The fluorescence intensity of calcein dye released from the liposomes was measured using a fluorometer (Promega, Glama^®-Multi Detection System, Madison, WI, USA) to evaluate the integrity of the liposome bilayer in response to CB[7] addition. Briefly, 80 mM calcein in PBS (pH = 12.00 ± 0.01) was added to liposomes during the hydration step. The liposomal suspension was then extruded below the transition temperature of liposomes ( $<$ 41 °C) ¹⁸ to enhance the flow of the suspension through the membrane. The unloaded calcein was removed via ultrafiltration using Amicon filtration tubes (Amicon Ultra-0.5 Centrifugal Filter Unit, Merck Millipore Ltd, Darmstadt, Germany) with a molecular weight cutoff of 10 kDa at 4 °C and flow rate of 13,000 L/minute. The sample was washed and centrifuged six times with PBS (pH 7.4) to ensure that all untrapped calcein molecules were removed, and the solution was adjusted to a neutral pH. The fluorescence intensity of calcein was measured at different time intervals ( $I_t$ ) over 21 hours for each liposomal sample with CB[7] concentrations ranging from 0.02866 to 1.433 mM. Dilution using 1% RIPA buffer along with the PBS was performed to prevent saturation of the fluorescence on the fluorometer. The amount of calcein released was then calculated using the below equation:

$$\mathrm{RF}(\text{\%})=100\times (I_t-I_0)(I_{\max }-I_0),$$

(1)

where $I_t$ , $I_0$ , and $I_{\max }$ are the fluorescence intensities measured at time t, at the beginning of the experiment, and after the addition of 1% RIPA buffer, respectively. ¹⁹

Benesi–Hildebrand (B–H) linear regression analysis of the fluorescence spectra was used to determine the association constant ( $K_{\mathrm{a}}$ ) between CB[7] and Chol, or between CB[7] and DPPC. This analysis can provide reliable information on the stoichiometry of the complex. The B–H analysis for a 1:1 stoichiometry yields:

$${\displaystyle \frac{1}{\mathrm{\Delta }{I}_t}={\displaystyle \frac{1}{K_{\mathrm{a}}(I_1-I_0){[\mathrm{G}]}_0}+{\displaystyle \frac{1}{(I_1-I_0)},}}}$$

(2)

where Δ $I_t$ is the difference between the observed fluorescence intensity of the guest and guest–host mixture and the fluorescence intensity of free guest, $I_1$ is the intensity of the guest–host complex, $I_0$ is the initial fluorescence intensity of free guest, and $[\mathrm{G}{]}_0$ is the initial concentration of the guest. ²⁰ The measured data satisfied equation (2), yielding a linear fit. Therefore, plotting 1/Δ $I_t$ versus 1/ $[\mathrm{G}{]}_0$ produced a linear B–H plot, which indicates the formation of a 1:1 inclusion complex between CB[7] and Chol or DPPC. ²¹ The $K_{\mathrm{a}}$ value was calculated from the ratio between the intercept and slope of the plot. ²² The B–H method was applied as follows: calcein and CB[7] were dissolved in PBS at pH 12 and 7.4, respectively. Then, the high pH value of 12 was adjusted to pH $\cong$ 7 by continuous dilution resulting from the addition of different solutions of CB[7], Chol, and/or DPPC (as described below). Subsequently, a CB[7] solution with a concentration of 0.002441 mM (X) was mixed with the calcein solution (also at a concentration equal to X) at a 1:1 volume ratio. Furthermore, DPPC and Chol solutions were prepared by dissolving different amounts of Chol and/or DPPC in absolute ethanol (99.9%) to obtain solutions with final concentrations of 2X, X, 0.5X, and 0.25X. The latter solutions were pipetted using a 200 μL pipette (PIPETMAN Classic P200, GILSON^®, France) into a plate reader (96-well black plate reader, Vision Plate™ 96 Well, USA) that contained a fixed volume of CB[7] and calcein solution (1:1 volume ratio). All of the samples in the plate reader were of a fixed volume (160 μL), and the fluorescence intensities of the different solutions were measured and compared with the fluorescence intensity of CB[7] with calcein solution.

Titration of liposomes with different concentrations of CB[7] using nuclear magnetic resonance

Nuclear magnetic resonance (NMR) spectroscopy was performed in deuterium oxide (D₂O; 99.0 at% D, Sigma-Aldrich, USA) using a Bruker Biospin AG Magnet system, 500MHz/54 mm instrument (Bruker BioSpin, Switzerland) with a PA BBO 500S1 BBF-H-D-05 Z SP probe. The temperature was controlled at a constant 298 K using a variable temperature unit. The effect of CB[7] on the liposomes was investigated by adding different concentrations of CB[7] (0.3189–1.085 mM) and measuring the resulting ¹H-NMR spectra. The results were then compared with the ¹H-NMR spectra of the liposomes and CB[7] alone.

Thermogravimetric analysis

Thermogravimetric analysis (TGA) was used to investigate the thermal stability and mass loss of the liposomal suspensions after CB[7] addition. TGA is an empirical technique in which the mass of a sample is measured as a function of the sample temperature or time to obtain a thermogram or thermal decomposition curve. ²³ Liposome suspensions were mixed with CB[7] solutions of different concentrations in a 1:1 volume ratio in alumina crucibles, and the wt% versus temperature for the different solutions was measured using a thermogravimetric analyzer (Mettler Toledo, TGA/DSC2 STAR^® System, Greifensee, Switzerland) and compared with the thermogram of blank liposomes.

Biocompatibility of CB[7]:hemolysis test

Hemoglobin loss or hemolysis is an indicator of the stability of the cell membrane of RBCs. ²⁴ A hemolysis assay using CB[7] was conducted by isolating fresh RBCs from serum through centrifugation at 2000 r/minute for 5 minutes. The supernatant containing plasma and platelets was discarded, and the remaining suspension was washed continuously with PBS (pH 7.4) to obtain a clear supernatant. The resultant RBC suspension (200 μL or more) was diluted in 9.8 mL of PBS, producing a stock RBC suspension containing approximately 2% RBCs. RBCs incubated with deionized water and 3% acetic acid, and RBCs incubated with PBS were used as the positive and negative controls, respectively. In these experiments, 50 μL of the 2% RBC suspensions were incubated with 450 μL CB[7] samples for at least 2 hours at 37 °C and 100 r/minute. The final concentrations of the resulting solutions ranged from 0.16 to 3.1 mg/mL. The absorbance of the supernatant was measured at 541–543 nm (Evolution™ 220 UV–Visible Spectrophotometer, Thermo Scientific™, Waltham, MA, USA) to evaluate the release of hemoglobin. All measurements were performed in triplicate, and the hemolysis rate was calculated using the below equation:

$$\mathrm{Hemolysis}(\text{\%})={\displaystyle \frac{(A_{\mathrm{test}}-A_{\mathrm{negative}})}{(A_{\mathrm{positive}}-A_{\mathrm{negative}})}\times 100,}$$

(3)

where A_test is the absorbance of the test sample, and $A_{\mathrm{negative}}$ and $A_{\mathrm{positive}}$ are the absorbances of the negative and positive controls, respectively.^25,26 The institutional review board (IRB) and ethical issues committee from Al-Ahliyya Amman University examined and authorized the hemolysis assay (IRB:AAU/1/7/2021-2022).

Transmission electron microscopy

The structure and morphology of the liposomes with and without added CB[7] were analyzed using transmission electron microscopy (TEM). TEM analysis of the liposome samples was performed using a negative staining method. First, 200-mesh formvar copper grids (SPI Supplies, USA) were equally coated with carbon using a low-vacuum Leica EM ACE200 glow discharge coating system (Leica, Austria). The carbon-coated grids were then coated with 1.5% vinylec K solution in chloroform. A drop of PBS-diluted liposome suspension was placed on the carbon-coated copper grids and left to air-dry overnight to evaluate the stability of the liposome samples over extended periods according to Odeh et al. and Alshaer et al.^3,16 Then, the loaded grids were stained using a 3% (v/v) aqueous solution of uranyl acetate dye for 20 minutes at room temperature. The grids were then rinsed with distilled water and air-dried at room temperature. The assembled grids were examined using a Versa 3D TEM system (FEI, Netherlands) with an acceleration voltage of 30 kV. ²⁷

Statistical analysis

The reported results were the averages of three or four complete and independent experiments. Data were reported as the mean ± SE (standard error of measurement) or $\pm$ SD. One-way analysis of variance was used to determine the differences in the means using OriginPro 2018 (OriginPro 2018 64 bit SR1 b9.5.1.195 for Windows, serial number: GF3S4-9089-7991320 Statistical, Northampton, MA, USA). Statistical significance was set at P $<0.05$ . ¹⁹

Results and discussion

Effect of CB[7] on liposome properties

This study investigated the effect of CB[7] on the stability of a DPPC:liposome (65:35) formulation. Previous research identified DPPC ratios of 60–40% and 70–30% as the most stable, with a 2:1 lipid ratio being optimal for characterization. ²⁸ Liposomes lacking electrosteric stabilization tend to aggregate because of electrostatic interactions between vesicles. Nevertheless, the liposomal formulation consisting of a 65:35 molar ratio of DPPC, lipid, and Chol was considered stable, with a surface potential sufficient to reduce the rate of aggregation and fusion. ²⁹ Table 1 presents the DLS measurements of conventional liposomes at different CB[7] concentrations. The results reveal a significant increase in the average size of liposomes at the highest CB[7] concentration (1.548 mM) compared to liposomes without CB[7]. Liposome size increased with higher CB[7] concentrations during both 2- and 24-hours incubation periods (Table 1). Particle size is an essential property of lipidic nanocarriers that influences the encapsulation efficiency, stability, bio-distribution, cellular uptake, mucoadhesion, and drug release profile. ³⁰ The sizes of liposomes used for clinical applications range from 50 to 450 nm. ³¹ Table 1 indicates that the sizes of the blank liposomes (L1) and those with CB[7] were smaller than 450 nm, except for L7 (with the highest CB[7] concentration), which reached 507.1 ± 173.7 nm after 2 hours. The PDI value was below 0.1 for L1 and increased with higher CB[7] concentrations, unaffected by the incubation time. The PDI value of L1 confirmed the existence of a monodisperse distribution. The TEM image of the blank liposomes (Figure 1(a)) shows they had a homogenous size and spherical shape. ¹⁶ Upon CB[7] addition, the liposomes were disrupted into either small micelles or aggregates of lipid–CB[7] complexes, as shown in Figure 1(b). ³²

Figure 1.

Representative DLS (size measurement) and TEM plots of (a) blank liposomes and (b) liposomes with 0.1698 mM CB[7] 24 hours after preparation. DLS measurements were performed at 25 °C, and the results are reported as triplicate averages ± SD.

Table 1.

DLS measurements of conventional liposomes (formulas) at different CB[7] concentrations after 2 and 24 hours of treatment at 25.0 ±  $0.1{}^{\circ }\mathrm{C}$ .

Formula	CB[7] (mM)	Time of incubation after treatment (hours)	Mean hydrodynamic diameter (nm $\pm$ SD)	PDI ± SD	ξ -Potential (mV ± SD)
L1	0	0	125.2 ± 4.277	0.096 ± 0.019	−3.71 ± 0.47
L2	0.02150	24	148.3 ± 5.367	0.15 ± 0.03	−4.29 ± 0.30
L3	0.1698		205.7 ± 10.42	0.34 ± 0.02	−1.98 ± 0.23
L4	1.548		426.1 ± 115.5	0.99 ± 0.01	8.29 ± 0.31
L5	0.03869	2	128.3 ± 2.401	0.11 ± 0.01	−3.73 ± 1.3
L6	0.1591		246.6 ± 24.36	0.46 ± 0.16	−10.9 ± 0.6
L7	1.548		507.1 ± 173.7	1 ± 0.002	7.32 ± 0.2

DLS: dynamic light scattering; CB[7]: cucurbit[7]uril; PDI: polydispersity index.

The remarkable increase in the average size of liposomes along with the fluctuating PDI values with increasing CB[7] concentration was attributed to the high affinity of CB[7] toward the lipid molecules forming the synthetic liposome bilayer, particularly Chol. Consequently, CB[7] may disrupt liposomes by solubilizing the lipids forming the liposomes. ³³ The tendency of CB[7] to migrate to the suspension surface reduces the packing density of the lipid membrane. ³⁴ Low negative values of the ξ-potential of the liposomal formulations indicated that phosphate ions in the phosphate buffer did not adsorb to the liposome head groups in large quantities. In addition, the ξ-potential depends on the ionic strength of the phosphate buffer used (I = 0.165 M). As noted by Chibowski and Szcześ, ³⁵ at high ionic strengths, the choline groups are located above the phosphate group in the polar heads of DPPC lipids. Additionally, compression of the diffuse part of the electric double-layer around the liposomes at high ionic strength results in low negative ξ-potential values. ³⁵ However, liposomal formulations with high CB[7] concentrations (L4, L7) exhibited more positive values of the ξ-potential than the liposomes without any CB[7] added (Table 1). For example, the ξ-potential value for L1 was −3.71 mV, whereas those for L4 and L7 were 8.29 and 7.32 mV, respectively. The sign and magnitude of the ξ-potential is evaluated based on the net charge accumulated on the liposome surface. ³⁵ The higher ξ-potential values for L4 and L7 were attributed to the binding of metal cations (Na⁺ from the PBS) to the portals of CB[7] via the ion-dipole effect. ¹¹ According to Tovani et al., macrocycles such as CB[7] have an affinity for membrane lipid molecules, particularly Chol. ³³ CB[7] molecules bind to the liposome surface, thus resulting in positive ξ-potential values for L4 and L7 owing to the metal cations on their rims.

Calcein release assay and liposome stability

Liposomes can form leaky structures when the phospholipids are hydrated under specific conditions. ³⁶ Calcein release from liposomes depends on the bilayer rigidity, diffusion coefficient calcein, and its solubility in the release medium. ¹⁹ A calcein release assay was used in this study to evaluate the changes in the liposome stability and permeability with varying CB[7] concentrations. Self-quenching has been used for quantitative leakage assays of lipid formulations. Initially, a high-concentration fluorophore is encapsulated inside the membrane to enhance self-quenching (Supplemental 1). An increase in the volume or permeability of the membrane decreases the concentration of fluorophore inside the liposomes. This generates a fluorescence signal of the fluorophore due to the decrease in self-quenching. ³⁷ Changes in the fluorescence intensity ( $I_t$ ) of liposomes with CB[7] concentrations from 0.02866 to 1.433 mM were investigated and plotted (Figure 2) over different time intervals ranging from 4 minutes to 21.33 hours. Calcein molecules exit liposomes via diffusion across the vesicle membranes. This physical mechanism allows for tuning of the release rate by modifying factors such as membrane thickness. The results show that the calcein permeability through the lipid bilayers changed significantly based on ¹⁹ the CB[7] concentration and exposure time to CB[7] (t). Different fluorescence responses were observed for low and high CB[7] concentrations and varying times. At low and medium CB[7] concentrations (from 0.02866 to 0.6879 mM), the fluorescence intensities reached their maxima at the beginning of the measurements (t  $=$  0 minutes); the fluorescence intensities then started to decrease and reached their minimum values after approximately 21 hours of exposure to CB[7] (Figure 2). The high fluorescence intensity of calcein released from the liposomes with low and medium CB[7] concentrations at t = 0 minutes was attributed to the difference in osmolarity, which caused calcein to diffuse across the membrane under osmotic pressure. ¹⁹ The lower fluorescence intensities after 21 hours were attributed to the quenched fluorescence of calcein through a mechanism by which the electronic excitation of the fluorophore is resonantly transferred to overtones and combined transitions of high-frequency vibrational stretching modes (of OH groups) of the solvent through space. ³⁸ However, the fluorescence intensities for higher CB[7] concentrations (1.146 and 1.433 mM) increased until reaching maximum values after 21.33 hours of exposure to CB[7] (for 0.8598 mM CB[7], the maximum fluorescence intensity occurred at t = 4 minutes) (Figure 2). In addition, the fluorescence intensity of liposomes with higher CB[7] concentrations was lower than that of liposomes with lower CB[7] concentrations. The high affinity of CB[7] molecules to lipid molecules in the liposome membrane, particularly Chol molecules, ³³ enabled CB[7] to deposit on the liposomes, resulting in stabilization of the liposomes. ³⁹ The release of calcein from the liposomes increased with time at CB[7] concentrations ranging from 0 to 0.1433 mM, reaching a maximum RF _t % after 80 minutes for concentrations between 0.02866 and 0.1433 mM (Figure 2). The delayed fluorescence intensity and higher release were linked to the effect of CB[7] on reducing the lipid packing density in the membrane. ³⁴ Liposomes with higher concentrations of CB[7], ranging from 0.2292 to 1.433 mM, exhibited lower values of RF _t %, reaching their maximum values after 21.33 hours of exposure to CB[7]. For example, liposomes with a CB[7] concentration of 1.433 mM exhibited a maximum RF _t % value of 3.5989% (Figure 2). Notably, the graph of RF _t % versus t for liposomes containing calcein obtained in this study was similar to the trend of RF _t % versus t for DPPC liposomes in Figure 2 of the study by Maherani et al. ¹⁹ These results can be explained by the high affinity of the CB[7] molecules for the lipid molecules in the liposome membrane. ³³ The accumulation of CB[7] molecules on the surface of the liposomes decreased calcein release from the vesicles. ³⁹ Accordingly, the amount of calcein released from liposomes with higher CB[7] concentrations was lower than that released from liposomes treated with lower CB[7] concentrations. Lin et al. ³⁹ demonstrated that liposomal formulations with a higher concentration of (2-hydroxypropyl)-β-cyclodextrin (CD) formed a more compact bilayer structure than that of the “liposome only” sample. Moreover, the integration of CD molecules into the outer surface of the liposomal sphere resulted in stabilization of the liposome. ³⁹ These changes in fluorescence intensity indicate that CB[7] affected the permeability of calcein through the lipid membrane. ¹⁹

Figure 2.

Fluorescence intensities ( $I_t$ ) and time-history of calcein release from liposomes with different CB[7] concentrations over 21.33 hours at 25 °C and different CB[7] concentrations; the results are the averages $\pm \mathrm{SE}$ (standard error of measurement; SE is 0.3 arbitrary units (a.u.)) of four independent experiments with statistical significance of P < 0.05. The relative standard deviation (RSD%) is ≤ 0.52% for all time intervals.

Behavior of calcein release from liposomes

Neutral vesicles have a net surface potential of zero, whereas calcein is a polyanion with a negative surface potential. The calcein molecule has approximately 3.5 negative charges at pH 7.4, as confirmed by acid–base titration (Figure 3). Consequently, a significant increase in the ionic strength inside the liposome produces a large difference in the osmolarity between the external and internal parts of the liposome. Considering the very small distance between the two lipid layers (nearly 4 nm), the external surface potential is influenced by the internal surface potential. Osmotic pressure inside the liposome forces calcein to diffuse through the lipid bilayer. Consequently, the release of calcein follows zero-order kinetics with a constant release rate. However, the release rate decreased over time because of the desorption/migration of calcein molecules from the surface to the outer aqueous sections of the liposomes, producing a non-zero or first-order release (Table 2, Supplemental 2). ¹⁹

Figure 3.

Chemical structure of calcein.

Table 2.

Apparent rate constants (k) and permeability coefficients $(P_m$ ) for calcein release from liposomes with different concentrations of CB[7].

Concentration of CB[7] (mM)	Reaction order	k (1/second)	Mean hydrodynamic diameter (ZAvg) (nm $\pm$ SD)	$r$ (nm)	$P_m$ (nm/second)
0	1	4.100 × 10−3	162.0 ± 1.0	80.99	0.1107
0.02866	1	7.717 × 10−4	369.6 ± 12.0	184.8	0.04754
0.08598	1	1.160 × 10−3	384.6 ± 2.7	192.3	0.07434
0.1433	1	2.820 × 10−3	369.9 ± 14.3	185.0	0.1739
0.2292	1	2.250 × 10−3	347.5 ± 7.9	173.7	0.1303
0.4585	1	1.520 × 10−3	238.1 ± 6.0	119.0	0.06031
0.6879	1	2.740 × 10−3	369.2 ± 20.6	184.6	0.1520
0.8598	1	1.740 × 10−3	508.6 ± 7.8	254.4	0.1475
1.146	1	9.848 × 10−4	810.3 ± 24.6	405.1	0.1330
1.433	1	8.889 × 10−4	1817 ± 37	908.6	0.2692

CB[7]: cucurbit[7]uril.

Data with standard errors were obtained from at least three independent experiments. All experiments were performed at 25 °C.

Considering the time-dependent release of calcein, the permeability coefficient ( $P_m$ ) values of the liposomes with different CB[7] concentrations were evaluated according to an analysis of the kinetics of calcein fluorescence using the below equation:

$${\mathrm{RF}}_t(\text{\%})={\mathrm{RF}}_{\max }(\text{\%})(1-e^{-kt}).$$

(4)

Finally, the permeability coefficient of calcein was calculated using the below equation:

$$P_m=\left({\displaystyle \frac{r}{3}}\right)k$$

(5)

where r is the radius of the liposome. This equation considers the role of the liposome radius and evaluates the apparent rate constant (k) for calcein release. This model was consistent with the calcein release process and showed effective parameters for the release behavior, such as the size of the liposome. ¹⁹

The values of ln ( $({\mathrm{RF}}_{\max }(\text{\%})-{\mathrm{RF}}_t(\text{\%}))$ / $({\mathrm{RF}}_{\max }(\text{\%})$ ) were plotted versus time (t), and the linear relationship indicated that the reaction was first order; k was then calculated from the slope (Supplemental 2). The permeability coefficients $(P_m$ ) for calcein release from liposomes with different concentrations of CB[7] were calculated as presented in Table 2. ¹⁹

Liposome size is an essential factor for calcein release, as indicated in Table 2. The permeability of hydrophilic molecules increases with increasing size of liposomes having the same unilamellarity. ¹⁹ As indicated in Table 2 and Supplemental 3, the permeability of calcein from the liposomes increased with the liposome size and reached a maximum (0.2692 nm/second) for the liposomal sample with the highest CB[7] concentration (1.433 mM).

The results showed that increasing concentrations of CB[7] (from 0.02866 to 1.433 mM) caused the size of large unilamellar vesicles (over 150 nm) to increase, which also increased their permeability (Table 2, Supplemental 3). CB[7] molecules, which strongly bind to lipids such as Chol, can solubilize Chol and DPPC from the lipid bilayer. ³³ Moreover, the packing density was reduced, which in turn disturbed the barrier to calcein permeation and resulted in increased $P_m$ values (Table 2). ¹⁹ The calcein release parameters were also calculated by Maherani et al. ¹⁹ following first-order kinetics. The results obtained in this study were higher owing to the addition of different concentrations of CB[7], which was expected to increase calcein release and permeability with time. Additionally, the presence of Chol in the liposomes affects their size, with higher Chol concentrations leading to increased liposome size. ²⁹

Stoichiometry and association constant determination

The stoichiometry and association constants $(K_{\mathrm{a}})$ for different complex formations were calculated using the B–H method. ²¹ In this study, this approach was employed to determine the stoichiometry and $K_{\mathrm{a}}$ of CB[7] with Chol and/or DPPC complexes (if present) by measuring the calcein fluorescence. It is important to investigate the different types of interactions between DPPC and/or Chol molecules and CB[7]. By comparing these interactions and identifying the stronger ones, the affinity of CB[7] toward lipid molecules in liposomes was confirmed. Anderson et al. and Zidovetzki and Levitan^40,41 reported a newly developed model of the interactions between a chosen macrocyclic molecule and a lipid molecule that incorporates the characteristics of two previous models. The mechanism involves the formation of an initial inclusion complex between the bilayer-phase lipid and a single macrocyclic molecule, followed by gradual attachment of three more macrocyclic molecules. The first step requires the removal of a phospholipid molecule from the bilayer environment into the aqueous phase before, or simultaneously with, binding to a macrocyclic molecule; the following three steps involve only the binding of additional macrocyclic molecules to the emergent complex. ⁴¹ The results showed that CB[7] interacted with DPPC in a similar manner (Supplemental 7(b)). However, ¹H-NMR spectra of liposomes titrated with varying CB[7] concentrations revealed no new peaks corresponding to a lipid–CB[7] complex. Instead, the chemical shifts were altered: an upfield shift was observed for the protons of the liposomes, and a downfield shift was observed for the protons of CB[7]. This indicates that CB[7] interacted only with the outer polar head group of DPPC and the –OH group of Chol, as shown in Supplemental 7(a) and supported by other studies.^33,39,42

The difference in fluorescence intensity ( $\mathrm{\Delta }{I}_t$ ) between the [CB[7]]:[Calcein] solution with a 1:1 volume ratio and the same solution containing different concentrations of Chol and/or DPPC were measured. Calcein and CB[7] had the same concentration of 0.002441 mM (X). The concentrations of DPPC and/or Chol were equal to 2X, X, 0.5X, and 0.25X. Calcein was used as a fluorescent dye, and its fluorescence was measured at an excitation wavelength of 490 nm and emission wavelength of 520 nm. ⁴³ A weak interaction between CB[7] and calcein was observed. Mohanty and Nau ⁴⁴ reported that CB[7] forms inclusion complexes with fluorescent dyes such as rhodamine 6G (Rh6G). CB[7] forms a 1:1 host–guest complex with Rh6G, as demonstrated by Job's plot. However, the size of Rh6G prevents the dye from being fully inserted into the cavity of the CB[7]. ⁴⁴ Calcein and Rh6G are both xanthene dyes; therefore, the size of calcein can be compared to that of Rh6G.^44,45 Additionally, calcein contains two attached methyleneiminodiacetic acid groups. ⁴⁵ Consequently, it is also difficult for calcein to fully insert into the CB[7] cavity.

According to the kinetic indicator displacement assay (kinIDA) reported by Sinn et al., ⁴⁶ the addition of a guest solution (Chol or DPPC) to a pre-equilibrated host–dye pair ([CB[7]]:[Calcein] solution) results in the replacement of the dye molecules with guest molecules. Therefore, the fluorescence intensity of calcein changed because DPPC or Chol replaced the dye molecules inside the CB[7] cavity. The latter phenomenon was attributed to the higher affinity of CB[7] toward lipid molecules, particularly Chol. ³³ To further explain this observation, ¹H-NMR spectra were recorded for Chol and 1:1 [CB[7]]:[Chol] in D₂O (Supplemental 7(a)). As reported by He et al., ⁴⁷ the hollow cavity of CB[7] shields protons, causing guest molecules to exhibit higher field signals in ¹H-NMR. The protons inside the cavity shift upfield, whereas those near the carbonyl oxygen atoms at the portals shift downfield. This makes ¹H-NMR an effective method for studying host–guest interactions. ⁴⁷ The formation of hydrogen bonds causes a loss of electron density around the hydrogen nucleus, resulting in a downfield shift. ⁴⁸ According to Supplemental 7(a), H₃ and H₁₉ shifted downfield and upfield, respectively. The downfield shift for H₃ was due to the formation of a hydrogen bond between the hydroxyl group of Chol and the highly electronegative carbonyl-fringed portals of CB[7]^11,49 (Supplemental 5 and 7(a)). However, the upfield shift for H₁₉ arose from the encapsulation of ring B of the Chol molecule (Supplemental 5) inside the CB[7] cavity. In conclusion, rings A and B of the nearly planar Chol molecule (Supplemental 5) were included in a single CB[7] cavity. ⁴⁹ To verify this result, the association constant was determined using equation (6) by plotting the inverse of the fluorescence differences against the inverse of the concentration of DPPC or Chol (Figure 4(b)):

$$K_{\mathrm{a}}={\displaystyle \frac{\mathrm{Intercept}}{\mathrm{slope}}}$$

(6)

Figure 4.

Formation of CB[7]–Chol inclusion complexes: (a) fluorescence intensity over 13 minutes as a function of Chol concentration; (b) Benesi–Hildebrand method based on fluorescence spectroscopy by titration of the CB[7] host solution with a Chol guest solution at different ratios. The temperature was 25 °C. The measurements represent averages of five independent trials with SE and RSD% values of ≤ 0.083 a.u. and ≤ 0.033%, respectively.

The association constant between CB[7] and Chol was strong: $3.08\times {10}^3/\mathrm{mM}$ or 3.08 $\times {10}^6/\mathrm{M}$ with 1:1 stoichiometry (Figure 4(b)). In addition, no linear relationship between 1/ $\mathrm{\Delta }{I}_t$ and 1/[Chol]² was observed, which eliminates the possibility of 1:2 or 1:3 complexation. ¹⁶ Our results are consistent with the association constant reported by Sinn et al. utilizing the B–H method based on fluorescence spectroscopy. ⁴⁶ In addition, as shown in Figure 4(a), at the beginning of the measurements (t = 0 minutes), the fluorescence intensity at a Chol concentration of 0 mM (not shown in Figure 4(a)) decreased from 5.6250 $\times {10}^4$ to 5.2924 $\times {10}^4$ a.u. when the Chol concentration was equal to 0.0006104 mM. The high fluorescence intensity at 0 mM Chol was attributed to the fluorescence enhancement typically observed when CB[7] is added to calcein solutions. ⁵⁰ For Chol concentrations of 0.25X and 0.5X, the quenching of the [CB[7]]:[Calcein] solution was higher than that at Chol concentrations of X and 2X. As demonstrated by Maillard et al., ³⁸ the OH groups in Chol cause three times stronger fluorescence quenching of calcein compared with DPPC. This is due to energy transfer from the excited dye to the vibrational modes of the solvent. However, the fluorescence intensity of calcein increased and reached a maximum at a Chol concentration of 0.002441 mM or at a 1:1 molar ratio of Chol to CB[7]. Hwang et al. ⁵¹ reported that water cluster molecules close to the electronegative portals of CB are anchored to carbonyls by hydrogen bonds. The stretching vibrations of the OH group are shifted to lower frequencies with stronger hydrogen bonds. ⁵¹ Encapsulation of the hydrophobic part of the Chol molecule inside the CB[7] cavity ³³ enabled the formation of hydrogen bonds between the OH group of the Chol molecule and the carbonyl groups of CB[7]. ⁵¹ Therefore, the OH group of the Chol molecule was shielded by CB[7], and the quenching caused by this group was less at a 1:1 molar ratio of Chol to CB[7]. ⁵²

This result can be attributed to the ability of DPPC to form micelles starting from extremely low concentrations.^53,54 As shown by Smith and Tanford, ⁵³ the critical micelle concentration of DPPC decreased rapidly as the methanol concentration in the solvent was reduced, reaching a value of 0.46 nM in water. To investigate the stoichiometry of the interaction between DPPC and CB[7], the condition of the lipid in solution must be known. Therefore, the amounts of lipids existing as monomers and micelles must be accounted for. ⁵³ However, the interaction between DPPC monomers or vesicles and CB[7] did not occur or was minimal. ⁵⁵ The addition of ethanol to DPPC vesicles in the gel phase caused structural changes, leading to hydrocarbon interdigitation. This resulted from the interaction of ethanol with the polar region of the lipid bilayer, which enhanced the head group solvation. ⁵⁶

To investigate the interactions of DPPC with CB[7], the ¹H-NMR spectrum of DPPC and 1:1 [CB[7]]:[DPPC] in D₂O was recorded; the shielded and deshielded protons are shown in Supplemental 7(b). As illustrated in Supplemental 7(b), most of the protons at the sn-1 and sn-3 positions of the glycerol backbone shifted upfield. In addition, the methyl protons of the tail groups were also shifted upfield. The peak at 3.2444 ppm corresponds to the nine hydrogens of the three methyl groups of the quaternary amine, and it was shifted downfield to 3.2469 ppm. The peak at 2.3020–2.2304 ppm is representative of the four β-hydrogens of DPPC on C1 of the tail groups. ⁵⁷ Most of the latter peaks were also shifted downfield. A schematic structure of the DPPC:CB[7] complex^32,49 (Supplemental 7(b), right) was used to identify the shielded and deshielded protons. Nagle and Tristram-Nagle ⁵⁸ reported that CB[7] formed very stable complexes with several cationic cholines. Regarding the phosphocholine group, the phosphate P exhibits a small downfield shift because the anionic groups are positioned outside the CB[7] cavity. The stabilization of the complex is achieved by ion–dipole and dipole–dipole interactions of the protonated and polar head groups of the guests with carbonyl-laced portals. In addition, the quadrupolar cavity of CB[7] stabilizes the oxygen in the carboxylate through dipolar–quadrupolar interactions with the trimethylammonium group, which is located near the carbonyl groups at the portals of the cavity. ⁵⁸ The peaks of the ¹H-NMR spectrum for the 1:1[CB[7]]:[DPPC] sample are consistent with a downfield shift of the protons of N(CH₃)₃⁺ and C1 of the tail groups (both are located at or outside of the portal). For the CB[7] molecules residing at the end of the alkyl chain of the fatty acids of DPPC, He et al. ⁴⁷ showed that the alkyl chains stabilize the CB[7] host when it bonds to external sites, and their hydrophobic character prevents the CB[7] host from sliding. The ¹H-NMR spectrum in Supplemental 7(b) clearly shows that the protons of the fatty acid tails were shifted upfield from 1.3130–1.2340 ppm to 1.2971–1.1783 ppm, except for the C1 and C2 protons. Anderson et al. and Zidovetzki and Levitan^40,41 considered the relative dimensions of the macrocycle cavity (e.g. the height) and phospholipid acyl chains. The phospholipid used in the latter study contained one acyl chain, which is the same as that used in this study (palmitoyl group). Accordingly, the length of the previous acyl chain was about 2 nm or 20 Å. The height of CB[7] is equal to 9.1 Å; thus, according to the previous study, ⁴¹ each acyl chain requires two CB[7] molecules to form an inclusion complex. As presented in Supplemental 7(b), the C1 and C2 protons were shifted downfield, and thus two C–C bonds with lengths of 1.54 Å ⁵⁹ were excluded from the total length of the acyl chain available for complexation with CB[7]. Consequently, the remaining length was (20 Å − (2 × 1.54 Å)) = 16.92 Å; thus, each acyl chain had one CB[7] molecule to complex with. Simultaneously, the peak of the free alkyl chain was considerably induced, which may have been caused by the inclusion of two tail chains in CB[7] separately. ⁴⁷

NMR titration of liposome with CB[7]

The stability of the liposomes toward CB[7] was investigated by measuring the chemical shifts of protons in both the liposomes and CB[7] (Supplemental 4) after the addition of CB[7] to the liposomes (Figure 5). The resulting spectra were compared to ¹H-NMR spectra for 5.00 mM liposomes and 5 mM CB[7] solutions. The ¹H-NMR spectrum of liposomes appeared as a high-resolution spectrum (Figure 5), as if the sample was in a liquid state (fast isotropic tumbling of the molecules canceled out the anisotropy); ⁶⁰ however, the formation of the liposomes usually caused broadening in the NMR signals. ³² In solids and soft matter, molecular motions do not exist or, when present, are slow and/or anisotropic, producing substantial line broadening, which completely prevents the separation of different peaks in the spectrum in some circumstances. ⁶⁰ A Chol concentration of 35 mol% increased the membrane fluidity, which explains the distinct peaks observed in the NMR spectrum of the liposomes. ⁶¹

Figure 5.

Stacked ¹H-NMR spectra of liposomes titrated with CB[7]. The proton peak of D₂O is distinguished by an asterisk. D₂O, 25 °C, 500 MHz.

The differences in the chemical shift values of the protons ( $\mathrm{\Delta }\delta$ ) of both liposomes and CB[7] were not significant when the two were mixed together (Supplemental 8). A slight upfield shift and broadening of the proton peaks of the liposomes was observed; however, the protons of CB[7] were shifted downfield, as shown in Figure 5 and Supplemental 8. In general, if the protons of a guest molecule are located inside the hydrophobic cavity of CB[7], the protons will be shifted upfield. However, if they reside near or around the negatively polarized carbonyl rims of CB[7], a downfield shift will be observed. ⁶² The upfield shift observed for the liposome protons was due to shielding of these protons by the CB[7] molecules located on the surface of the liposomal suspension. ³⁴ However, the protons of CB[7] exhibited a downfield shift, particularly the ${\mathrm{H}}_{\mathrm{b}}$ and ${\mathrm{H}}_{\mathrm{c}}$ protons. The downfield shift of the CB[7] protons was due to hydrogen bonding between the carbonyl groups of CB[7] and the positively charged choline groups in the polar head of DPPC. This bonding reduces the electron density around the protons in the CB[7] portals.^11,63

A comparison of the proton peaks of Chol in CDCl₃ and the proton peaks of liposomes in D₂O (Supplemental 6(b) and (c)) shows that the liposome protons appeared at approximately the same frequency as Chol protons H₃ and H₁₉ (Supplemental 5, Figure 5). The Chol proton peaks were not obvious in D₂O because of the weak solubility of Chol in D₂O (spectrum not presented). However, the solubility of Chol in D₂O was enhanced by the addition of CB[7] at a 1:1 molar ratio (Supplemental 6(a)). The purpose of this comparison was to identify the liposome proton peaks using the same solvent (D₂O); however, Chol was not soluble in D₂O even after heating. Consequently, the addition of CB[7] to liposomes mainly affected the Chol proton peaks by shielding them, and they were also slightly broadened. Therefore, the Chol molecules were encapsulated inside the CB[7] cavity at a fast-to-intermediate exchange rate. ⁶⁴ CB[7] can also interact with the DPPC molecule, but it has a higher affinity toward Chol because it can solubilize one Chol molecule for every five DPPC molecules, as demonstrated by Tovani et al. ³³ The comparison between the spectra did not exclude the presence of DPPC proton peaks in the ¹H-NMR spectrum of liposomes. However, the DPPC proton peaks possibly overlapped with those of Chol. For example, methyl and methylenic protons in the lipid chains resonate in the region of 0.85–1.50 ppm, but the methylenic protons of choline were assigned at 3.6 ppm. ⁶⁵

Thermogravimetric analysis

TGA was used to detect the change in the mass percent of liposomes after the addition of different concentrations of CB[7] as a function of the sample temperature. ²³ The thermal stability of the sample and the presence of residual components in the sample were evaluated using TGA curves. ⁶⁶ The weight of blank liposomes decreased between $\cong$ 85 °C and $\cong$ 109 °C (Figure 6). However, with increasing concentrations of CB[7], the mass loss of the corresponding liposomal samples was less than that of the sample containing only liposomes (Figure 6). In TGA, the weight change below 100 °C can be attributed to the water adsorbed and bound to pure compounds or polymers. Below 100 °C, the blank liposomes exhibited a mass loss of approximately 63% ⁶⁷ (Figure 6) because the liposome suspension was mostly water. ⁶⁶ However, when CB[7] molecules were deposited on the liposomes or the hydrophobic cavity of CB[7] was incorporated with the long hydrophobic chains of the liposomes, ³⁹ CB[7] acted as a protective layer, which could reduce the water loss of the liposomal samples treated with CB[7]. Consequently, we observed a mass loss below 100 °C of only approximately 27% in liposomes with 1.146 mM CB[7]. ⁶⁷ Notably, CBs are thermally stable and do not decompose up to 673 K or 400 °C. ⁶⁸ As shown in the thermogram of CB[7] (Figure 6), water loss started at approximately 42 °C and was complete in one step by nearly 109 °C. ⁶⁹ The mass remaining after heating above 110 °C was the residual weight of CB[7]. Xu et al. also reported that the addition of (2-hydroxypropyl)-β-cyclodextrin (HP-β-CD) to liposomes increased their thermal stability compared with the stability of the liposomes without HP-β-CD. Consequently, the increase in thermal stabilization of the liposomal samples in this study was consistent with the observations reported in previous studies. ⁶⁶

Figure 6.

TGA thermograms representing the mass losses (%) of CB[7] and liposomes with different concentrations of CB[7] versus temperature. Heating rate: 10 °C/minute.

Hemolysis

Erythrocytes or RBCs were used to evaluate the hemolysis rate. One method of toxicity evaluation relies on measuring the release of hemoglobin from suspended RBCs. To investigate the concentration–response hemolysis of CB[7], a suspension of RBCs (2% RBCs) was incubated with solutions containing different concentrations of CB[7] for 2 hours or more at 37 °C and 100 r/minute. To determine the hemolysis rate (%), the amount of hemoglobin in the supernatant was detected using a UV–Vis spectrophotometer at wavelengths of 541–543 nm. The concentration of hemoglobin released in the supernatant was directly related to the number of cells that had undergone lysis. The level of in vitro cytotoxicity of hemolytic action is assessed by the observed mortality rates, as follows: 0–9% is classified as non-toxic materials, 10–49% are slightly toxic, 50–89% are toxic, and 90–100% are highly toxic materials. ²⁴ The hemolysis results showed that 2.580 mM of CB[7] resulted in 56.88% hemolysis (Figure 7). However, CB[7] at concentrations 0.05159 to 0.8254 mM yielded hemolysis rates of less than 1%, as shown in Figure 7. Moreover, Fink et al. ²⁵ reported that incubation with CB[5]–CB[7] did not cause hemolysis. The concentration of the CB samples ranged from 0.3955 to 25.80 mM. The hemolytic ratio was below 1% for all of the CB samples tested, which was comparable to the negative control (negative values were caused by statistical deviations of the medium blank). ²⁵ The concentrations of CBs used in the previous study were much higher than those used in recent studies and yet did not cause hemolysis. Many factors can increase the hemolysis rate, such as inappropriate handling during processing of blood, inappropriate storage conditions, bacterial hemolysins, antibodies that cause complementary lysis, defects in the RBC membrane, or an abnormality in the blood donor. ⁷⁰ In summary, as demonstrated by the mortality rates of RBCs in a previous study, ²⁴ it is advisable to use concentrations of CB[7] in the range of 0.05159–0.8254 mM, which yield hemolysis rates below 1% . One possible explanation for the induction of hemolysis at CB[7] concentrations equal to or greater than 1 mM is the extraction of Chol molecules from the cell membrane of RBCs by CB[7]. ²⁵

Figure 7.

Hemolysis rate of RBCs under the effect of different CB[7] concentrations. The temperature was 37 °C, and the data are averages of three independent experiments with RSD% ≤ 1%.

Conclusion

In this study, the effects of CB[7] on the lipid bilayer of conventional liposomes were investigated. The CB[7] molecules were mainly localized on the surface of the liposomes and at low concentrations ( $<0.800\mathrm{mM}$ ) they had slight adverse effects on the stability of the lipid membrane. However, at CB[7] concentration ≥1 mM, the liposomal suspension suffered from elevated levels of instability. Fluorescence measurements of calcein released from liposomes at different concentrations of CB[7] also provided evidence for the placement of CB[7] molecules near or on the surface of the liposomes. The ability of CB[7] macromolecules to extract Chol and DPPC into their hydrophobic cavities is the main factor that disrupts lipid membranes. The results of the B–H method using fluorescence spectroscopy confirmed that CB[7] had more complexation affinity toward Chol than toward DPPC. Consequently, the most probable mechanism by which CB[7] affected the stability of the liposomes and the integrity of the lipid membrane was the extraction of Chol in a time- and dose-dependent manner. Furthermore, TGA thermograms confirmed the ability of CB[7] to localize on the surface of the liposomes and negatively affect the integrity of the lipid membrane (especially at high concentrations) by incorporating the long chains of lipids into their hydrophobic cavities. Moreover, the results showed that CB[7] concentrations below 0.825 mM produced insignificant hemolysis rates after a short incubation time of approximately 2 hours. Finally, our results demonstrated that CB[7] did not significantly affect the stability of the lipid membrane of conventional liposomes when used at low concentrations. Therefore, CB[7] can be used safely as a new drug delivery system that can enhance the stability and therapeutic index of potentially active hydrophobic drugs.

Supplemental Material

sj-docx-1-sci-10.1177_00368504251334687 - Supplemental material for Effect of cucurbit[7]uril on DPPC-containing liposomes: Interactions with the lipid bilayer

Supplemental material, sj-docx-1-sci-10.1177_00368504251334687 for Effect of cucurbit[7]uril on DPPC-containing liposomes: Interactions with the lipid bilayer by Summay Altaher, Walhan Alshaer, Fedaa Al Adaileh, Hamdi Nsairat, Shrouq Alsotari, Majdoleen H Rababah, Abeer Al Bawab and Fadwa Odeh in Science Progress