J. Phys. Chem. B 1998, 102, 6107-6113
6107
Water-Soluble Hollow Nanospheres as Potential Drug Carriers Jianfu Ding and Guojun Liu* Department of Chemistry, The UniVersity of Calgary, 2500 UniVersity DriVe, NW, Alberta, Canada, T2N 1N4 ReceiVed: December 17, 1997; In Final Form: May 13, 1998
A polyisoprene-block-poly(2-cinnamoylethyl methacrylate) (PI-b-PCEMA) sample with 88 units of isoprene and 2.3 × 102 units of CEMA formed vesicles in THF/hexanes with hexanes volume fractions between 50% and 95%. The vesicles had PCEMA as the shell and PI chains stretching into the solution phase from both the inner and outer surfaces of the PCEMA shell. The cavity diameter of the vesicles is ∼38 nm, and the volume fraction of PI chains there is low. After the PCEMA shell was cross-linked, the PI chains were converted to water-soluble poly(2,3-dihydroxyl-2-methyl-butane) chains. The water-soluble hollow nanospheres uptook a large amount of rhodamine B in methanol and released the compound into water at a tunable rate depending on the amount of ethanol added to the aqueous medium. This study demostrates the potential of hollow nanospheres, prepared from tailor-made biocompataible and degradable polymers, as drug carriers in controlled drug release applications.
I. Introduction A diblock copolymer consists of two linear polymer chains joined together in a head-to-tail fashion. Polyisoprene-blockpoly(2-cinnamoylethyl methacrylate), PI-b-PCEMA, is a diblock copolymer. A diblock copolymer may form micelles in a block-
selective solvent, which solubilizes one but not the other block of the copolymer. In a micelle, the soluble block stretches into the solvent phase and the insoluble block of different chains aggregates to form a polymer-rich phase to minimize polymer and solvent contact. The shape of the micelles formed ranges from spheres1,2 to vesicles,3-6 cylinders,7,8 and donuts.6 In the case of spheres or cylinders, the cores made up of the insoluble block would be spherical or cylindrical with chains of the soluble block on their surfaces. In a vesicular micelle, the insoluble block forms an essentially solvent-free shell with chains of the soluble block stretching into the solvent phase from both the outer and inner surfaces. A donut micelle is an enclosed “hairy” cylinder. Donut micelles form because the end caps of the cylindrical micelles are not stable energetically. Micelles of different shapes are formed to minimize the free energy of a system. In general, diblocks with a long soluble and short insoluble block tend to form spherical micelles3-4,6 because the curvature of the interface between the core and corona is positive in this case and a chain in the corona can occupy more lateral space than a chain in the core, which reduces chain-chain repulsion in the corona. As the length of the soluble block decreases relative to that of the insoluble block, less space is required to accommodate the soluble chains, the curvature of the interface between the two blocks decreases,
and the micelle shape changes from spheres to cylinders or donuts and then to vesicles. In a cylinder, the interfacial curvature along the cylinder axis direction is zero. In the case of vesicles, the curvature of the interface between the soluble and insoluble block on the cavity side is negative. This micelle morphological transitional trend is similar to that observed for small-molecule surfactants in water.9 We recently established that a PI-b-PCEMA sample with 88 units of isoprene and 2.3 × 102 units of CEMA formed vesicles in THF/hexanes mixtures with hexanes (HX) volume fractions between 50% and 95%.6,10-11 In such solvents, the PCEMA block is insoluble and makes up the shell. Transmission electron microscopic studies revealed that these vesicles had a cavity diameter of ∼38 nm. Such cavities should hold a large amount of material. In this paper, we demonstrate the preparation of water-soluble PCEMA-cross-linked vesicles from PI-b-PCEMA by modifying the PI chains chemically. Also shown is the ability of these water-soluble cross-linked vesicles or hollow nanospheres to uptake rhodamine B (RB), a model compound for a drug, in an organic solvent, and to release it from the hollow nanospheres in water. If the polymeric hollow nanospheres are prepared from tailormade diblocks that are biocompatible and degradable, they may serve as an alternative to small-molecule surfactant vesicles or liposomes12,13as drug delivery vehicles. Small-molecule vesicles or liposomes are unstable in biological milieu, and the stability and thus the circulation time of these species in human bodies12,14 are recently drastically increased by attaching watersoluble polymer chains to the vesicle or liposome surfaces. Since small-molecule vesicles or liposomes have to be modified by polymers for use in controlled drug delivery, the direct production of polymeric vesicles for such a purpose may be more costeffective. Furthermore, polymeric vesicles may be made to possess a greater range of cavity sizes to enable the delivery of large species such as proteins and genes. Polymeric vesicles may also be advantageous over diblock spherical micelles15-19 as drug carriers because the cavities of the vesicles should enable the loading of larger and more species.
S1089-5647(98)00551-3 CCC: $15.00 © 1998 American Chemical Society Published on Web 07/11/1998
6108 J. Phys. Chem. B, Vol. 102, No. 31, 1998
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Figure 1. FTIR spectra of RB (top), PHI-b-PCEMA nanospheres (bottom), and RB-loaded PHI-b-PCEMA nanospheres (RB loading density ) 10%, middle). The peaks marked by arrows are used to determine RB loading densities.
TABLE 1: Characteristics of the PI-b-PCEMA Sample n/m from NMR
M h w/M hn from GPC
10-4M hw (g/mol) GPC
dnr/dc a (mL/g)
hw 10-4M (g/mol) LS
10-2 n
10-2 m
0.38
1.19
2.7
0.150
6.6
0.88
2.3
a
Specific refractive index increment.
II. Experimental Section Polymer Synthesis and Characterization. PI-b-PCEMA was prepared by reacting PI-b-PHEMA, where PHEMA denotes poly(2-hydroxylethyl methacrylate), with excess cinnamoyl chloride in pyridine at room temperature. The precursor to PIb-PHEMA was PI-b-P(HEMA-TMS), where P(HEMA-TMS) denotes poly[2-(trimethylsiloxyl)ethyl methacrylate)]. PI-bP(HEMA-TMS) was prepared by anionic polymerization.7,10 PIb-PCEMA was characterized by NMR, gel permeation chromatography (GPC), and light scattering as described previously,10 and the characterization results are shown in Table 1. Instrumentation and Techniques. Light scattering experiments were performed using a Brookhaven model 9025 instrument equipped with an argon-ion laser operated at 488 nm. FTIR measurements were performed on a Mattson model 4030 instrument. Transmission electron microscopic (TEM) images
were obtained using a Hitachi-7000 electron microscope operated at 100 kV. The fluorometer used was Photon Technology International Alpha Scan system equipped with a 75 W xenon lamp. Vesicle Preparation. PI-b-PCEMA, 400 mg, was dissolved in 65 mL of THF. A quantity of 35 mL of hexanes was added. The solution was then slowly added into an equal volume of THF/HX with 85% HX to induce vesicle formation. The vesicle solution in THF/HX with 60% HX was stirred for 2 weeks before an equal volume of HX was added to yield a vesicle solution at a concentration of 1.0 mg/mL in THF/HX with 80% HX. The solution was immediately irradiated to obtain a PCEMA conversion of 40%. The vesicles were initially equilibrated in THF/HX with 60% HX, since the vesicles were found to have the narrowest size distribution at this solvent composition. More HX was added just before irradiation because it was supposed to increase the rigidity of the PCEMA shell (due to the reduced PCEMA swelling by THF) and reduce the degree of intervesicle fusion during the cross-linking process. The irradiated mixture was concentrated by rotor-evaporation to 12 mL. Hydroxylation of the PI Chains. A literature method20-22 was followed to add vicinal hydroxyl groups across a double bond in PI. This should convert PI to poly(2,3-dihydroxyl-2methylbutane) or hydroxylated PI (PHI). To 8 mL of 90% formic acid (0.188 mol) at 10 °C were added 2 mL of acetic anhydride (0.021 mol), 2 mL of 30% hydrogen peroxide (0.019 mol), and 0.06 mL of concentrated sulfuric acid.20 The mixture was stirred for 5 min before 6 mL of it was taken and mixed with 3 mL of the concentrated cross-linked vesicle solution (1.3 × 10-4 mol of isoprene units). The new mixture was stirred for 8 h at room temperature and then dropped into 100 mL of water. The polymer was precipitated after centrifugation and rinsed with water before it was stirred in 10 mL of 1 N NaOH solution at 60 °C for 1 h. After this, the polymer was once again precipitated by centrifugation, redispersed in fresh water, and neutralized by adding 0.10 N hydrochloric acid. The precipitation and dispersion in the fresh
Figure 2. TEM image of PI-b-PCEMA vesicles. The sample was sprayed in THF/hexanes with 80% hexanes.
Nanospheres as Potential Drug Carriers
Figure 3. FTIR spectra of PI-b-PCEMA (top), PI-b-PCEMA vesicles at a CEMA conversion of 40% (second from the top), cross-linked PI-b-PCEMA vesicles reacted with performic acid for 8 h and hydrolyzed in 1.0 M NaOH at 60 °C for 1 h (third from the top) and in 2.0-M NaOH at 70 °C for 2 h (bottom). The marked peaks at 828, 1635, and 3514 cm-1 are derived from absorption characteristic of the hydrogen atom attached to the double bond of PI, the PCEMA double bond, and the hydroxyl groups.
water step was repeated another three times before the precipitate from water was dried in vacuo to obtain solid, water-soluble, hollow nanospheres. Loading Rhodamine B. RB (50 mg), PHI-b-PCEMA nanospheres (10 mg), and methanol (0.12 mL) were mixed in a 1.5 mL polyethylene capsule. The mixture was stirred at room temperature for 5 days before 0.80 mL water was added, and the mixture was centrifuged to remove the supernatant. The
Figure 4. TEM image of PHI-b-PCEMA nanospheres.
J. Phys. Chem. B, Vol. 102, No. 31, 1998 6109 vesicles were then redispersed in ∼1 mL of water and centrifuged out at 14 × 103 rpm. This rinsing step was repeated four times with vesicle loss at every step due to the difficult removal of the vesicles by centrifugation. The RB-loaded vesicles were then dried for RB content analysis. RB Content Analysis. Compared in Figure 1 are the IR absorption spectra of RB, PHI-b-PCEMA nanospheres, and RBloaded PHI-b-PCEMA nanospheres between 1500 and 1800 cm-1. Since RB and the nanospheres have distinct absorption peaks at 1590 and 1730 cm-1, respectively, the ratio in the intensities of these two peaks was used to evaluate the relative RB and nanosphere contents in solid RB-loaded nanosphere samples. This was done by calibrating the instrument with a series of mixtures with known RB to PHI-b-PCEMA nanosphere mass ratios first. RB Loading Kinetics. A quantity of 20 mg of the nanospheres, 100 mg of RB, and 0.25 mL of methanol were mixed in a 1.5 mL polyethylene capsule and stirred. At different times, four drops of the mixture were dropped into 1.5 mL of water. The aqueous mixture was shaken and centrifuged to remove the supernatant. The solid material was subsequently rinsed four times in this fashion before it was dried in vacuo and used for FTIR measurements to evaluate the RB content. TEM Analysis. Nanospheres in a solvent were sprayed on carbon-coated copper grids using a home-built device, as described previously.6 The solvent used for dispersing the PIb-PCEMA hollow nanosphere was THF/hexanes with 20% THF by volume and that for PHI-b-PCEMA was methanol. RBloaded PHI-b-PCEMA nanospheres were sprayed from water. While the PI-b-PCEMA and PHI-b-PCEMA nanospheres were stained with OsO4 vapor overnight before TEM analysis, the RB-loaded nanospheres were stained by stirring them in ∼1% CsOH solution overnight. Fluorescence Measurements. Fluorescence emission spectra were measured by exciting at 550 nm. For RB release kinetic studies, the increase in the fluorescence intensity at 576 nm was monitored as a function of time.
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The amount of RB released into water from nanospheres after RB-releasing equilibrium is established was estimated by RB fluorescence intensity measurement. For this purpose, the fluorometer was calibrated with a series of RB solutions in water with known concentrations. III. Results and Discussion Vesicle Formation. Illustrated in Figure 2 is a TEM image of the PI-b-PCEMA vesicles prepared by following the procedures described in the Experimental Section. The vesicular structure has been established on the basis of the following observations.10 First, the particles are spherical. This was established by comparing the images of the particles obtained at different sample-stage tilting angles. Second, the PI block reacted more readily with OsO4 and should appear darker in a TEM picture. Also, the PI block should be in contact with the solvent because PI is soluble and PCEMA is insoluble in the THF/HX mixture. Third, the weight fraction of the PI block is only 9.0%. The overall conclusion was that the lighter centers of the particles in Figure 2 correspond to the cavities. The dark ring separating the cavities and the shell represents the location of a collapsed PI layer. The cavities should have been originally filled with the THF/hexanes that swelled the PI chains. Further unambiguous evidence for the proposed structure was the preparation of “semishaved hollow nanospheres” by cutting the outer PI chains from the cross-linked PI-b-PCEMA vesicles selectively or the preparation of “fully shaved nanospheres” by cutting the PI chains from both the outer and inner vesicle surfaces.10 Cross-Linked PI-b-PCEMA Vesicles. The cross-linking of PCEMA has been used by our group to make various nanostructures including nanofibers23 and nanochannels in polymer thin films.24,25 CEMA dimerizes because of photoinduced cycloaddition. PCEMA cross-links because of the dimerization among many CEMA groups from different chains.26,27 The conversion of PCEMA can be seen by comparing the first and second FTIR spectra shown at the top in Figure 3. UV irradiation reduced the carbon-carbon double bond peak intensity at 1635 cm-1 substantially. CEMA conversion was also accompanied by a UV absorbance decrease at 274 nm, and the conversion estimated from this method was ∼40%. Properties of the Hairy Hollow Nanospheres. Dried crosslinked vesicles were soluble in a wide range of solvents including THF, chloroform, toluene, etc. The survival of the vesicular structure in such solvents was confirmed by our TEM results. The cross-linked vesicles, however, did aggregate with time in the solid state probably because of intervesicle linking through the PI chains. This aggregation was confirmed by an increase in the molar mass and size of the cross-linked vesicles with storage time. Light scattering studies of the cross-linked vesicles were carried out in THF/HX with 20% THF immediately after they were prepared. The weight-average molar mass, M h w, and radius of gyration, RG, of the vesicles determined this way were 2.1 × 108 g/mol and 39 nm, respectively. Assuming a cavity radius, Rc, of 19 nm and a polymer density of F ) 1.0 g/cm3, this molar mass should give a vesicle outer radius, Ro, of 42 nm as calculated from
(4/3)πNAF(Ro3 - Rc3) ) 2.1 × 108 (g/cm3)
(1)
This Ro value compares well with 39 nm obtained from Figure 2, considering the relatively wide vesicle size distribution.
The hydrodynamic radius, Rh, of the hairy hollow nanospheres was measured over the scattering angle range 30-150° in THF/ HX with 20% THF, and the average value is 53 nm. This gives Rh/RG ) 1.36, which is close to the theoretical value of 1.29 for homogeneous spheres.19 Hydroxylation of the PI Chains. Performic acid forms by reacting formic acid with hydrogen peroxide.20-22 Acetic anhydride was used to consume the water present in acetic anhydride
HCO2H + H2O2 98 HCO3H + 2CH3COOH formic acid and hydrogen peroxide. Concentrated sulfuric acid was used as a catalyst. The reaction between performic acid and the double bond of PI is supposed to yield an oxirane initially, which is not stable and may be further converted into a hydroxyformoxy derivative:
Hydrolysis of the formyl ester in 1.0 M aqueous sodium hydroxide yields PHI.
The strongest evidence for the conversion of PI to PHI is the water solubility of the hollow nanospheres after reacting the nanospheres with performic acid for 8 h at room temperature and hydrolyzing the sample in 1.0 M NaOH at 60 °C for 1 h. Then the characteristic broad absorption of the hydroxyl groups at 3514 cm-1 in the IR region appeared (third curve from the top in Figure 3) at the expense of a peak at 828 cm-1, which derives from the wagging motion of the hydrogen atom next to the PI double bond. Unfortunately, this peak is very weak and is visible only in the cross-linked PI-b-PCEMA vesicle sample because of the reduced PCEMA absorption at 864 cm-1 (insert of Figure 3). The quantification of the degree of isoprene double bond conversion is difficult because the PI double bond absorption peaks are all over-shadowed by those of PCEMA. We also attempted solid-state NMR and Raman experiments. Again, PCEMA peaks over-shadowed those of PI. This is understandable because the weight fraction of PI in the polymer is only 9.0%. Reactions of PCEMA. Only ∼40% of the double bonds of PCEMA were consumed in the UV cross-linking step. The partial CEMA conversion is evident from the second curve from the top in Figure 3 because the 1635 cm-1 peak remains strong after UV photolysis of the vesicles. The residual double bonds of PCEMA can also be converted to hydroxyl groups owing to performic acid and hydrolysis treatments. The reaction of CEMA double bonds with performic acid should, however, be much slower than that between the isoprene double bonds and performic acid because 2-methyl-2-butene, an excellent model compound for an isoprene unit, reacted ∼1 × 105 times faster with performic acid than alkyl cinnamates.22 It is thus not
Nanospheres as Potential Drug Carriers
J. Phys. Chem. B, Vol. 102, No. 31, 1998 6111 TABLE 3: Change in Nanosphere Loading Density as a Function of the Mass of RB Useda run
mass of RB (mg)
mass of nanospheres (mg)
loading density (g/g)
1 2 3
49.8 25.9 12.6
10.8 11.1 10.7
0.10 0.04 0.02
a
Figure 5. Increase in RB loading density as a function of RB and nanosphere equilibration time in methanol.
TABLE 2: Properties of the PI-b-PCEMA and PHI-b-PCEMA Hollow Nanospheres sample (nanospheres)
solvent
PI-b-PCEMA THF/HX (2/8) PHI-b-PCEMA H2O
h w*/(g/mol) Rh/nm RG*/nm dnr/dc b M by LS by LS (mL/g) by LS 53a 59a
39
0.180
1.81 × 108
a Value determined at the scattering angle of 90°. b Specific refractive index increment.
surprising to see the survival of some PCEMA double bonds (third curve from the top) after the performic acid and mild hydrolysis treatments. The lowest curve in Figure 3 was obtained after the performicacid-treated sample was hydrolyzed in a 1.0 M aqueous NaOH solution at 70 °C for 2 h. The PCEMA peak at 1635 cm-1 disappeared completely. This is most likely caused by the removal of cinnamoyl groups due to the hydrolysis of the ester groups of PCEMA. Our nanospheres were prepared utilizing the milder hydrolysis conditions, i.e., in NaOH solution at 60 °C for 1 h, to ensure the integrity of the PCEMA shell. Properties of the PHI-b-PCEMA Hollow Nanospheres. When first prepared in water containing NaCl salt, the PHI-bPCEMA hollow nanospheres precipitated readily by centrifugation at a spinning rate of 1.5 × 103 rpm. As the salt was removed by repeated rinsing, the vesicles became more stable in water. A spinning rate of 1.4 × 104 rpm was required to precipitate out the vesicles. The freshly prepared wet hollow nanospheres were found by dynamic light scattering to disperse well in water. Dry hollow nanospheres did disperse in water, but the dispersion contained some nanosphere aggregates. The particles in Figure 4 appear to have the same shape as those in Figure 2. This suggests the integrity of the hollow nanosphere after the hydroxylation step. The fact that the size of the vesicles did not change very much further suggests the integrity of the hollow nanospheres. The cross-linked PI-bPCEMA vesicles in THF/HX with 80% HX had a hydrodynamic radius of 53 nm, which changed to 59 nm for the PHI-b-PCEMA nanospheres in water. Kinetics of RB Loading. We loaded the hollow nanospheres with RB by equilibrating the nanospheres with a concentrated RB solution in methanol. We followed the kinetics of RB incorporation into the nanosphere by analyzing RB loading densities, the amount of RB incorporated into each gram of nanospheres, at different times, and the results are illustrated in Figure 5. The majority of RB is incorporated into the nanospheres in less than 1 h. Thus, the RB loading rate is high. Equilibrium RB Loading Densities. Shown in Table 3 are the loading densities obtained after the hollow nanospheres were equilibrated with RB solutions at different concentrations for 5 days. The loading densities increase linearly with the RB concentration in methanol. This is expected because the cavities of the nanospheres are supposed to be eventually filled, except for the presence of some PHI chains, with a RB solution in
In all cases, 0.12 mL methanol was used as the solvent.
methanol at approximately the same concentration as that outside the nanospheres. The higher the RB solution concentration, the more RB the nanospheres uptake. Methanol was used as the solvent because RB dissolves well in it. Other solvents such as THF, chloroform, acetone, and water were also used. In all cases, the loading densities were less than ∼4%. Location of RB. Figure 6 shows a TEM image of RB-loaded nanospheres. This nanosphere sample was equilibrated with a RB methanol solution for 5 days and has a RB loading density of 10%. The sample was then stained by CsOH, which reacted supposedly selectively with the carboxylic acid groups of RB. The observation of a dark phase on the inner wall of the nanospheres suggests that the majority of RB was incorporated into the cavities of the nanospheres. The nanospheres in Figure 6 seem to be aggregated, and some nanospheres are deformed. When stained with OsO4, the nanospheres appeared normal but the RB component cannot be seen because of lack of contrast. We do not know the exact reason for nanosphere aggregation and deformation when CsOH was used as the staining agent. Kinetics of RB Release. Illustrated in Figure 7 are the fluorescence emission and excitation spectra of RB. RB release kinetics was monitored by following the increase in the fluorescence intensity of RB at 576 nm as a function time after a RB-loaded hollow nanosphere sample was mixed with water as shown in Figure 8. Released RB has a higher fluorescence quantum yield, because RB concentrations in the cavities are high and the fluorescence is quenched due to concentration quenching there. Similar intensity increase was observed for calcein as calcein was released from the phosphatidylcholine liposomes.28 The kinetics of RB release from the hollow nanospheres should be analogous to that of reagent release from the traditional vesicles or liposomes. We assume that all of the RB molecules are initially dissolved in the aqueous phase in the cavities of the hollow nanospheres and that the volume fraction of the hollow nanospheres is low. Under these conditions, the result of Johnson and Bangham,30 who modeled reagent release from single-compartment liposomes contained in a dialysis bag, can be modified to describe our system:
( )
ln 1 -
nw ) -(p/Vc)t n0
(2a)
or
nw ) n0(1 - e-(p/Vc)t)
(2b)
In eq 2, n0 is the total number of RB molecules, nw is the number of RB molecules in the bulk aqueous phase at time t, and Vc and p denote the total volume and permeability, measured in cm3/s, of the hollow nanospheres. The fluorescence intensity, I(t), of a RB-nanosphere sample in water should be
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Figure 6. TEM image of RB-loaded PHI-b-PCEMA nanospheres. The sample was stained with CsOH.
where I∞ is the fluorescence intensity of RB after it is completely released into the aqueous phase and
I∞ ) φwn0
Figure 7. Excitation (left) and emission (right) spectra of RB in water. The excitation spectrum was obtained at the emission wavelength of 576 nm. The excitation wavelength used was 550 nm for the emission spectrum.
Illustrated in Figure 8 is a set of RB release kinetic data together with the best fit by eq 4. Unfortunately, the agreement between eq 4 and experimental data is poor. Such poor agreement between experimental data and eq 4 has been observed previously as well29,30 and has been attributed to some unrealistic assumptions made in the Johnson and Bangham model. These unrealistic assumptions included the uniform size distribution of the vesicles and the thinness of the shell layer so that all the trapped reagent was initially in the cavities. A further assumption made about this system is the complete dissolution of RB in the aqueous phase in the cavities. The sample with its kinetic data shown in Figure 8 has a RB loading density of 10%, and the solubility of RB in water at room temperature is ∼9% (w/w) as established by us experimentally. Thus, RB cannot be fully dissolved in the aqueous phase in the cavities of the vesicles. Since eq 4 does not fit experimental data well, I(t) has been fitted by the empirical relation
I(t) ) a0 - a1 exp(-t/τ1) - a2 exp(-t/τ2) Figure 8. Increase in RB fluorescence intensity (scatters) as a function of time after 0.488 mg of RB-loaded nanospheres (loading density ) 10%) was added into 5.00 mL of water. The solid lines represent the best fit to the experimental data by eqs 4 (poorer fit) and 6 (better fit with the line hidden mostly in the data points).
I(t) ∝ φwnw + φvnv
(3)
where nv is the number of RB molecules in the nanosphere cavities with nv ) n0 - nw; φw and φv are the fluorescence quantum yields in bulk water and the nanosphere cavities, respectively. Inserting eq 2 into eq 3 and simplifying yield
I(t) ) I∞ - (φw - φv)e-(p/Vc)t
(4)
(5)
(6)
A measure of the rate of RB release is obtained from the average RB retention time:
〈τ〉 )
a1τ1 + a2τ2 a1 + a2
(7)
For the sample with its kinetic data shown in Figure 8, the RB retention time with the nanospheres is 26 h. Extent of RB Release. Figure 8 gives the relative rate of RB release but not the amount of RB released into the aqueous phase. To estimate the amount RB released, we compared the RB fluorescence intensity at the end of a RB release experiment with those of samples with known RB concentrations in water.
Nanospheres as Potential Drug Carriers
J. Phys. Chem. B, Vol. 102, No. 31, 1998 6113 IV. Conclusions
Figure 9. Effect of varying solvent composition on the rate of RB release from the nanospheres. As the ethanol volume fraction in water/ ethanol mixtures decreased from 50% (top curve) to 33% (second from the top), 16% (third from the top), and 8% (bottom), RB release rate decreased. The RB-loaded nanosphere sample was prepared by equilibrating RB and the nanospheres in methanol for 5 days. The initial RB loading density was 10%.
TABLE 4: Effect of Increasing Ethanol Content on the Time of RB Retention with the Hollow Nanospheresa solvent
τ1/h
τ1/h
〈τ〉/h
water water/ethanol ) 92/8 water/ethanol ) 84/16 water/ethanol ) 67/33 water/ethanol ) 50/50
2.0 2.3 × 10-2 1.9 × 10-2 1.6 × 10-3 5.5 × 10-4
37 0.55 0.21 0.028 0.048
26 0.49 0.099 7.7 × 10-3 8.5 × 10-3
a RB was loaded into the nanospheres by equilibrating RB with the nanospheres for 5 days. The RB loading density is 10%.
The fluorescence intensity of a RB-nanosphere sample is given by eq 3. If RB is quantitatively released into the aqueous phase, the fluorescence intensity of the RB-nanosphere sample should be φw/φv times higher than the case when no RB is released into the aqueous phase. Intensities between those of these two extreme cases should be observed for partial RB release. Since the RB fluorescence intensity of Figure 8 increased by a factor of 3.8 over time, 3.8 should be the lower bound for φw/φv. We started by assuming quantitative RB release at the end of the experiment for the sample with its data shown in Figure 8. This assumption yielded a RB mass, which gave a RB loading density of 12% for the nanospheres. This value is higher than 10% measured from FTIR. If we assume partial release of RB, the RB loading density obtained from fluorescence intensity measurement would be even higher. Thus, RB is probably released quantitatively. Tuning of RB Release Rate. Illustrated in Figure 9 is the comparison between the kinetics of RB release in water/ethanol mixtures at different ethanol volume fractions. As ethanol content increases from 0% to 33%, the RB release rate increases and the average RB retention time decreases (Table 4). No further increase in RB release rate was observed as ethanol content increased from 33% to 50%. This is probably caused by the fact that the RB release rate gets so fast at these high ethanol contents that the fast kinetics was beyond the time resolution limit of our experiment. The exact mechanism by which ethanol increases RB release rate is unknown. The two possible reasons are the increased RB solubilization and PCEMA shell swelling in water/ethanol mixtures with higher ethanol contents.
PI-b-PCEMA hollow nanospheres were made water-soluble by converting the PI chains to PHI chains. The occurrence of the desired reactions has been qualitatively confirmed by FTIR experiments. The hollow nanospheres survived the reactions maintaining their integrity, as demonstrated by light scattering and TEM results. The water-soluble hollow nanospheres uptook a large amount of RB in methanol. The amount of RB taken up increased with RB concentration in methanol. Once the RBloaded nanospheres were added into water or water/ethanol mixtures, the RB was quantitatively released. The RB release rate could be tuned by changing the composition of water/ ethanol mixtures. This study demonstrates the potential of hollow polymeric nanospheres as drug carriers in controlled drug delivery. Since the kinetic data of RB release could not be explained by the Johnson and Bangham model, a more complete analysis of the release process will be carried out in the future. Acknowledgment. The Natural Sciences and Engineering Research Council of Canada is thanked for financially sponsoring this research. The polymer used was synthesized by Dr. M. Yang for another project, and his contribution is cordially acknowledged. References and Notes (1) Tuzar, Z.; Kratochvil, P. Surf. Colloid Sci. 1992, 15, 1. (2) Guo, A.; Liu, G.; Tao, J. Macromolecules 1996, 29, 2487. (3) Zhang, L.; Eisenberg, A. Science 1995, 268, 1728. (4) Zhang, L.; Yu, K.; Eisenberg, A. Science 1996, 272, 1777. (5) Ding, J.; Liu, G. Macromolecules 1997, 30, 655. (6) Ding, J.; Liu, G.; Yang, M. Polymer 1997, 38, 5497. (7) Price, P. Pure Appl. Chem. 1983, 55, 1563. (8) Tao, J.; Stewart, S.; Liu, G.; Yang, M. Macromolecules 1997, 30, 2738. (9) Gelbart, W. M.; Ben-Shaul, A. J. Phys. Chem. 1996, 100, 13169 and references therein. (10) Ding, J.; Liu, G. Chem. Mater. 1998, 10, 537. (11) The same PI-b-PCEMA sample was used to obtain results in refs 5 and 9 and here. The polymer characterization results are different in ref 5 because the homo-PI impurity was not extracted at the early stage of the work. (12) Lasic, D. Chem. Ind. 1996, March 18, 210. (13) Gredoriadis, G. Liposome Technology, 2nd ed.; CRC Press: Boca Raton, FL, 1993. (14) Lasic, D. Angew. Chem., Int. Ed. Engl. 1994, 33, 1685. (15) Cammas, S.; Kataoka, K. In SolVents and Self-Organization of Polymers; Kluwer Academic Publishers: Dordrecht, The Netherlands, 1996. (16) Topp, M. D. C.; Dijkstra, P. J.; Talsma, H.; Feijen, J. Macromolecules 1997, 30, 8518. (17) Henselwood, F.; Liu, G. Macromolecules 1997, 30, 488. (18) Chu, B. Langmuir 1995, 11, 414. (19) Qin, A.; Tian, M.; Ramireddy, C.; Webber, S. E.; Munk, P.; Tuzar, Z. Macromolecules 1994, 27, 120. (20) Swern, D.; Billen, G. N.; Findley, T. W.; Scanlan, J. T. J. Am. Chem. Soc. 1945, 67, 1786. (21) Swern, D. Org. React. 1953, 7, 378. (22) Swern, D. J. Am. Chem. Soc. 1947, 69, 1692. (23) Liu, G.; Qiao, L.; Guo, A. Macromolecules 1996, 29, 5508. (24) Liu, G.; Ding, J.; Guo, A. Macromolecules 1997, 30, 1851. (25) Liu, G.; Ding, J. AdV. Mater. 1998, 10, 69. (26) Kato, M.; Ichijo, T.; Ishii, K.; Hasefawa, M. J. Polym. Sci., Part A: Polym. Chem. 1971, 9, 2109. (27) Guillet, J. E. Polymer Photophysics and PhotochemistrysAn Introduction to the Study of Photoprocesses in Macromolecules; Cambridge University Press: Cambridge, U.K., 1985. (28) Meada, M.; Kumano, A.; Tirrell, D. A. Ann. N.Y. Acad. Sci. 1991, 618, 362. (29) Johnson, S. M.; Bangham, A. D. Biochim. Biophys. Act. 1969, 193, 82. (30) Ringsdorf, H.; Schlarb, B.; Tyminski, P. N.; O’Brien, D. F. Macromolecules 1988, 21, 671.
