The influence of manufacturing procedure on the degradation of poly(lactide-co-glycolide) 85:15 and 50:50 implants

Journal of Controlled Release 43 (1997) 161–173

The influence of manufacturing procedure on the degradation of poly(lactide-co-glycolide) 85:15 and 50:50 implants a, b c Monica Ramchandani *, Marvin Pankaskie , Dennis Robinson a

b

Integra LifeSciences Corporation, 105 Morgan Lane, Plainsboro, NJ 08536, USA Drug Information and Epideimology Center, 137 Victoria Hall, University of Pittsburgh Medical Center, Pittsburgh, PA 15261, USA c Department of Pharmaceutical Sciences, College of Pharmacy, University of Nebraska Medical Center, 600 South 42 nd Street, Omaha, NE 68105, USA Received 22 September 1995; revised 4 April 1996; accepted 23 April 1996

Abstract Poly(lactides-co-glycolides) (PLGA) are widely investigated biodegradable polymers and are extensively used in several biomaterials applications as well as drug delivery systems. The PLGA polymers degrade by bulk hydrolysis of ester bonds and breakdown into their constituent monomers, lactic and glycolic acids which are excreted from the body. The purpose of this investigation was to study the effect of manufacturing procedure on the in vitro degradation of two PLGA copolymers, 85:15 and 50:50, which were fabricated as implants. Implants were compressed from microcapsules prepared by nonsolvent induced phase separation using two solvent-nonsolvent systems, viz., methylene chloride-hexane (non-polar) and acetonephosphate buffer (polar). Studies were performed at pH 4.5, 7.4 and 9.4 and polymer degradation was monitored by measuring the decrease in number- and weight-average molecular weights (M n and M w ), mass loss, formation of lactic and glycolic acids and pH of the degradation medium. Representative scanning electron micrographs (SEMs) were also obtained to study the changes in surface morphology of the implants. Results of these studies indicated that both PLGA 85:15 and 50:50 implants prepared by the non-polar procedure degraded faster than the implants prepared by the polar procedure. The decrease in polymer M n and M w followed pseudo first-order kinetics. Changes in M n and M w occurred before the onset of mass loss, after which implant mass loss was described by pseudo first order kinetics. The appearance of lactic and glycolic acids corresponded to the initiation of mass loss and also resulted in decrease in pH of the bulk degradation medium. The SEMs indicated that water uptake was faster in implants prepared by the non-polar method resulting in a more porous matrix which degrades more rapidly. Finally, the average degradation time for PLGA 85:15 was ¯26 weeks and that for PLGA 50:50 was 6–8 weeks. Keywords: Poly(lactides-co-glycolides); Microcapsules; Nonsolvent phase separation; Polymer degradation; Surface morphology; Degradation time

1. Introduction Poly(lactide-co-glycolides) (PLGA) are widely investigated biodegradable polymers and copoly*Corresponding author.

mers. The biocompatibility of these polymers is well established and they have been extensively used in biomaterial applications such as tracheal replacement, ligament reconstruction, surgical dressings and dental repairs. The potential of these polymers as carriers in controlled-release drug delivery systems

0168-3659 / 97 / $17.00  1997 Elsevier Science Ireland Ltd. All rights reserved PII S0168-3659( 96 )01481-2

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was evidenced by some of the first reports for the delivery of narcotic antagonists, antimalarials and contraceptive hormones [1]. The early efforts were directed towards the homopolymer, polylactic acid, rather than the copolymers. Since then, numerous research teams have used various copolymers of PLA and PGA in drug delivery formulations. The advantages of using copolymeric forms are that they are amorphous and also degrade faster than the homopolymers. PLGA polymers degrade by bulk hydrolysis of ester bonds, and breakdown to their constituent monomers, lactic and glycolic acids, which are eliminated from the body through Kreb’s cycle. The hydrolytic cleavage involves reaction with one molecule of water to form acidic and alcoholic end groups (Scheme 1), and can be catalyzed by acid, alkali, or enzymes [2]. Although several investigators have studied the in vitro and in vivo degradation of these polymers, most studies primarily focus on local tissue reactions to these polymers rather than attempt to elucidate either the mechanism or kinetics of degradation [2–4]. A few recent literature reports, however, have addressed the correlations between the physicochemical properties of the PLGA polymers and degradation rates [5–7]. The investigations of Li et al., [8,9] revealed that degradation of P(D)LA and PLGA copolymers proceeded more rapidly in the center than at the surface of the devices. The degradation of PLGA polymers is reported to be influenced by several physical and chemical factors, such as initial pH, ionic strength and temperature of the external bulk medium, molecular weight, crystallinity, exposure to gamma radiation

and presence of drugs and other agents in the matrices. However, data regarding the effects of these factors on polymer decomposition are controversial, for example, some researchers have shown no influence of pH on degradation [3,5], whereas others have observed an appreciable effect [4,10,11]. An important consideration regarding degradation of PLGA matrices is the influence of manufacturing procedure on polymer decomposition. It has been demonstrated that drug release from PLGA matrices is influenced by polymer degradation [12,13], and that the method of manufacture of a delivery system influences drug release [14], however, there are no reports regarding the effect of manufacturing procedure on the degradation of PLGA matrices. The purpose of this investigation, therefore, was to study the effect of manufacturing procedure and bulk medium pH on the in vitro degradation of two PLGA copolymers, 85:15 and 50:50, which were fabricated as implants. These implants were compressed from microcapsules prepared by two new microencapsulation procedures.

2. Materials and methods

2.1. Materials DL-lactic acid, glycolic acid, sodium citrate and sodium borate were purchased from Sigma Chemicals Co., St. Louis, MO. The polymers, PLGA 85:15 and 50:50 were obtained from Medisorb Technologies International L.P., Cincinnati, OH. The polystyrene standards for gel permeation chromatography (GPC) were purchased from Polysciences,

Scheme 1.

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Waarington, PA. Methylene chloride, acetone, hexanes, hydrochloric acid, disodium hydrogen phosphate, potassium dihydrogen phosphate and potassium hydroxide were obtained from Fisher Scientific, Fairlawn, NJ.

2.2. Methods 2.2.1. Preparation of PLGA 85:15 and 50:50 microcapsules and implants 2.2.1.1. Preparation of microcapsules. Microcapsules were prepared by nonsolvent-induced, phaseseparation using two solvent-nonsolvent systems, namely, methylene chloride-hexane (nonpolar) and acetone-Sorenson’s phosphate buffer (polar). The precise procedures for both microencapsulation methods used are described below. Non-polar procedure. A 2.5% w / v solution of the polymer in methylene chloride was added drop-wise to a 20-fold excess of the nonsolvent, hexane, while stirring the system with a magnetic stirrer at 700– 800 rpm. The rate of addition of the polymer solution was #2.5 ml / min for PLGA 85:15 and #5 ml / min for PLGA 50:50. PLGA 85:15 microcapsules were equilibrated to harden for 5.5 h, whereas PLGA 50:50 microcapsules were hardened for 2.5 h only. The microcapsules were then filtered over vacuum, air-dried at ambient room temperature and sieved to remove any aggregates. Polar procedure. A 2.5% w / v solution of the polymer in acetone was added drop-wise to a 20-fold excess of the nonsolvent, Sorenson’s phosphate buffer, pH 7.4. Microcapsules of PLGA 85:15 and 50:50 were prepared by a procedure identical to that described for the organic method above, equilibrated to harden for 3–4 h, filtered over vacuum, dried in a vacuum oven at room temperature for 3–5 h, and sieved to remove any aggregates. 2.2.1.2. Preparation of implants. Implants (0.45 3 0.51 cm) were prepared by compression of 100 mg of microcapsules, prepared either by polar or nonpolar procedure, in a Carver hydraulic press at a pressure of one metric ton, using a die-and-punch apparatus. The mean weight of implants was greater than 98 mg, the small batch-to-batch coefficient of

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variation (,3%) was indicative of the reproducibility of the method.

2.2.2. In vitro degradation of PLGA 85:15 and 50:50 implants Degradation studies were performed at pH 7.4 (Sorenson’s phosphate buffer), pH 4.5 (citrate buffer) and pH 9.4 (borate buffer). Each implant was immersed in 10 ml buffer maintained at 378C in a shaking water bath. Polymer degradation was measured by monitoring the decrease in number- and weight-average molecular weight (M n and M w ), changes in polydispersity, mass loss, formation of lactic and glycolic acid monomers and measuring the pH of the degradation medium. All studies were performed in triplicate.

2.2.2.1. Average molecular weight and polydispersity. The decrease in number and weight average molecular weight (M n and M w ), and the changes in polydispersity of the implants were determined by gel permeation chromatography (GPC) after calibrating the system using polystyrene standards (600– 600 000 g / mol). Calibration curves were obtained daily to account for the interday variability, and the correlation coefficients exceeded 0.996 in the M w range used. The molecular weights of all samples were determined directly from the polystyrene standard curve and not from PLGA standards, and therefore, represent approximate values. Because the purpose of these investigations was to compare the degradation of implants under different experimental conditions, the determination of absolute molecular weights was not essential. The polymer samples, retrieved at various time intervals, were dissolved in methylene chloride (1% w / v) and eluted, with methylene chloride as mobile phase at a flow rate of 1 ml / min using a solvent delivery module, Model 110B (Beckman Instruments Inc., Fullerton, CA). Two Ultrastyragel columns, ˚ and 10 4 A ˚ 7.8 3 300 mm, with a pore size of 10 3 A (Waters, Milford, MA) maintained at 338C in a column heater (Biorad, Knauer, Germany) were used to separate degraded polymer fractions as detected using a differential refractometer, Model 1750 (Biorad, Knauer, Germany). The average molecular weights and polydispersity were calculated using a

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GPC 6000 software package (Jones Chromatography, CO).

2.2.2.2. pH. After retrieving the sample, pH of the dissolution medium was recorded at each sampling interval using a Beckmann pH-meter, Model F 40. 2.2.2.3. Mass loss. The initial (t 5 0), and both wet and dry final weights of the implants at each sampling interval were recorded. At appropriate intervals, implants were retrieved from the medium and the wet weight was recorded after drying off excess water using Kimwipes. The implants were then dried for 18–24 h over phosphorus pentoxide in a vacuum desiccator at ambient room temperature, after which the final dry weight was recorded. The percentage mass loss was calculated from the following equation: % mass loss 5 (initial weight 2 final dry weight) / initial weight (1)

2.2.2.4. Formation of lactic and glycolic acids. To monitor the formation of lactic and glycolic acid monomers, a reverse-phase high pressure liquid chromatographic (HPLC) assay was established by modifying the procedure developed by Owen et al. [15]. Two Beckmann Ultrasphere 5-m m ODS columns, 4.6 3 250 mm (Alltech, Deerfield, Ill) connected in series were used to achieve simultaneous separation of both monomers. The mobile phase, phosphate buffer, pH 2.25, was maintained at a flow rate of 1 ml / min using a Shimadzu liquid chromatographic pump, Model LC-6A. The samples were automatically injected onto the columns using a Shimadzu Autoinjector, Model, SIL 9A. Each acid was detected spectrophotometrically at 215 nm using a UV-VIS spectrphotometric detector, Model SPD6AV (Shimadzu, Colombia, MO) and the concentration was determined from calibration curves in the range of 0.5–5 mg / ml using Shimadzu chromatopac data processors Models CR 60 and C-R3A. 2.2.3. Scanning electron microscopy studies Changes in the surface morphology of implants before and during in vitro implant degradation were evaluated by scanning electron microscopy (SEM).

The implants were sputter coated with gold under vacuum using an electron beam (10 kV). The implant surface was viewed under low (310.6) and high (3342) magnifications and representative photomicrographs obtained.

3. Results and discussion

3.1. Preparation of microcapsules 3.1.1. Non-polar procedure The conventional nonsolvent-induced phase separation procedure for the preparation of microcapsules involves the drop-wise addition of equivalent quantities of the nonsolvent to the polymeric solution maintained under constant agitation conditions. Although this procedure using methylene chloride-hexane system had been successfully used with P(L)LA by Sampath et al. [16], for PLGA 85:15, it resulted in the formation of a gel-like mass. The conventional methylene chloride-hexane method was then modified such that the polymeric solution of PLGA 85:15 (2.5% w / v) was now added drop-wise to a large excess of the nonsolvent, hexane. Studies performed to establish the volume excess required for this procedure indicated that a 20-fold excess of nonsolvent was necessary to produce flowable, discrete microcapsules with approximately 90% yield. The microcapsules were required to be equilibrated for about 4–6 h in the presence of nonsolvent before they were filtered under vacuum, washed with hexane, and dried at ambient room temperature. A slower rate of addition of the polymer solution and a longer equilibration time were necessary to obtain discrete microcapsules with PLGA 85:15 as compared to PLGA 50:50. These effects were attributed to the increased lactic acid content of PLGA 85:15, and hence increased solubility in methylene chloride, in which it dissolved relatively rapidly compared to the more hydrophilic PLGA 50:50. Therefore, the rate of desolvation of PLGA 85:15 in methylene chloride may be slower, requiring a slower rate of addition and increased equilibration time to obtain discrete, free-flowing microcapsules. 3.1.2. Polar procedure This method is innovative and novel in that it involved the use of an aqueous nonsolvent, thereby

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minimizing the use of conventional organic solvents in the preparation of PLGA microcapsules. Similar to the non-polar procedure, phase separation was induced by drop-wise addition of the polymeric solution of either PLGA 85:15 or 50:50 in acetone (2.5% w / v) into a 20-fold excess of Sorenson’s phosphate buffer to obtain discrete microcapsules (yield ¯ 90%). After equilibration for 3–4 h, the microcapsules were filtered under vacuum, washed with water, and dried in a vacuum oven at ambient room temperature. In this method the rates of addition of the polymer solution (4–5 ml / min) into the nonsolvent and equilibration times required for the microcapsules to harden (3–4 h) were similar for both PLGA 85:15 and 50:50.

3.2. In vitro degradation studies 3.2.1. Effect of manufacturing procedure on the degradation of PLGA 85:15 and 50:50 implants 3.2.1.1. Changes in molecular weight and polydispersity. The implants compressed from microcapsules prepared by the nonpolar procedure degraded more rapidly than those made by the polar procedure. For nonpolar procedure, the M n of PLGA 85:15 implants decreased from 41 642 g / mol to 2369 g / mol in 12 weeks, compared to a decrease from 57 081 g / mol to 28 756 g / mol observed for implants prepared by the polar procedure during the same period (Fig. 1A). Similarly, PLGA 50:50 implants manufactured by the nonpolar procedure also degraded more rapidly (Fig. 1B). In 3.5 weeks, the M n decreased from 53 565 g / mol to 3523 g / mol for implants made by nonpolar method as compared to a decrease from 51 437 to 20 845 g / mol for implants made by polar method. Due to the more hydrophilic nature of PLGA 50:50, almost complete degradation of implants was obtained in 8 weeks as compared to almost 30 weeks for PLGA 85:15. The differences in M w during degradation of both PLGA 85:15 and 50:50 implants were similar to those observed for M n (Table 1). The decrease in average molecular weight of PLGA 85:15 and 50:50 implants (M n and M w ) was biphasic with no change in molecular weight being observed initially. This lag time was followed by a pseudo-first-order decrease (r 2 . 0.93) in M n and M w for both PLGA 85:15 and 50:50 implants made

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Table 1 Weight-average molecular weight of PLGA 85:15 and 50:50 implants during degradation Polymer PLGA 85:15 0 weeks 12 weeks PLGA 50:50 0 weeks 3.5 weeks

Polar procedure

Non-polar procedure

90 801 53 552

73 031 21 120

81 852 42 811

86 176 8185

by the two procedures (Fig. 1A,B and Fig. 2A,B). Sanders et al., [17] observed a similar degradation profile for PLGA 50:50, and suggested that the lag time represents the time taken for the aqueous degradation medium to penetrate the polymeric matrix. Biphasic profiles for PLGA 50:50 matrices, due to an initial slow degradation, followed by a faster decomposition phase, have also been reported by Hutchinson et al. [18] and Sampath [16]. Changes in polydispersity of PLGA 85:15 and 50:50 associated with the decomposition of implants prepared by both polar and non-polar procedures are shown in Fig. 2A,B. In both cases, polydispersity increased with time indicating that there was an increase in the formation of lower molecular weight fractions. This was followed by a decrease in polydispersity, which indicated that most of the higher molecular weight fractions had degraded. This confirmed that significant changes in polydispersity correlate to the time when the monomeric carboxylic acids are detected in the medium. Consistent with the more rapid decrease in molecular weight, the changes in polydispersity occurred at earlier times for both PLGA 85:15 and 50:50 implants prepared by the non-polar procedure. For example in the case PLGA 85:15 implants, changes in polydispersity were observed between 9 and 20 weeks for implants made by organic procedure as compared to 20–26 weeks for implants prepared by the aqueous procedure. A bimodal distribution of average molecular weights was observed for PLGA 85:15 implants prepared by the non-polar method between 12 and 18 weeks after which the distribution became unimodal again (Fig. 3A). Similarly, PLGA 50:50 implants prepared by the non-polar method exhibited a bimodal distribution between 6–7 weeks of degradation. Several investigators have reported that a bimodal

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Fig. 2. Changes in the polydispersity of PLGA 85:15 (A) and PLGA 50:50 (B) during degradation of implants. Fig. 1. Decrease in number-average molecular weight of PLGA 85:15 (A) and PLGA 50:50 (B) implants during degradation.

distribution of molecular weights, which is characterized by two maxima, is due to the differences in degradation rates of the crystalline and amorphous microdomains of the polymer [2,3]. However, for intrinsically, amorphous polymers such as PLGA 85:15 and 50:50, the hypothesis of Li et al. [8,9], seems to be more appropriate. These investigators showed that degradation of P(DL)LA and PLGA copolymers proceeded more rapidly in the center than at the surface of the devices. The authors related this observation to the formation of an outer layer of

slowly degrading polymer that entraps the degrading macromolecules in the interior of the matrix, allowing only relatively low molecular weight oligomers to diffuse and dissolve in the surrounding media. As a result, a bimodal distribution is observed because the rate of ester bond cleavage in the inner part of the degrading specimens is accelerated due to autocatalysis because of an increase in the number of carboxylic acid groups present in this region relative to the surface. Results similar to those of Li et al. [8,9] were obtained in these studies when the surface and core of the implant were analyzed separately, i.e., a unimodal distribution was obtained, with the

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Fig. 3. GPC chromatograms of PLGA 85:15 implants prepared by non-polar (A) and polar (B) methods during degradation.

surface portion exhibiting a shorter retention time indicative of larger molecular weights (Fig. 4). No bimodal distribution was observed for implants made by polar method (Fig. 3B). A possible explanation for this observation may be the relatively lower matrix porosity for implants prepared by this procedure. As a result of this lower porosity, the free

carboxylic acid groups formed due to polymer degradation cannot penetrate out into the bulk medium.

3.2.1.2. Changes in implant mass and medium pH. Fig. 5A,B shows the implant mass loss profile and the corresponding changes in the pH of the medium

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Fig. 4. GPC chromatogram PLGA 50:50 implants showing differences in degradation of surface and interior of the devices.

for PLGA 85:15 and 50:50 implants. These data are typical of these polymers in that there was a characteristic onset time before which no mass loss occurred, followed by a rapid mass loss phase, described by pseudo-first-order kinetics (r 2 . 0.92). Similar results have been reported for PLGA 50:50, 85:15 and 70:30 [6–8]. Table 2 lists times of onset for mass loss, the associated pseudo first-order degradation rate constants and the correlation coefficients for PLGA 85:15 and 50:50 implants prepared using the two procedures. The onset time was 9 weeks for PLGA 85:15 and 4 weeks for PLGA 50:50 implants made by non-polar method, as compared to 23 weeks for PLGA 85:15 and 6.5 weeks for PLGA 50:50 implants made by the polar method. The decrease in pH corresponded to the formation of acidic monomers. Some investigators have reported only slight changes in pH of the bulk medium during degradation [8,9]. In the studies of Li et al. [8,9], the initial weight of samples or buffer capacity of the phosphate buffer was not reported, and, it is possible that due to smaller initial sample weights and / or higher buffer capacity, the pH of the buffer was unaffected by the presence of acidic monomers.

3.2.1.3. Formation of lactic and glycolic acids. The formation of monomers, lactic and glycolic acids,

during implant degradation was monitored by HPLC. The retention time for glycolic acid was 5.5 min and lactic acid was 7 min. To account for any betweenday variability, calibration curves were run concurrently with all samples. The calibration curves were linear, the correlation coefficients ranging between 0.97 and 0.99 for lactic and glycolic acid concentrations of 0.5–5 mg / ml. The coefficients of variation were less than 5% at all times. Fig. 6A,B illustrates the cumulative amounts of lactic and glycolic acids produced due to degradation of PLGA 85:15 implants made by both polar and nonpolar methods. The appearance of monomers corresponded to the initiation of implant mass loss due to formation of these water-soluble species. Further, the production of monomers occurred earlier for implants made by non-polar method, consistent with the earlier findings that these implants degraded faster than those manufactured by aqueous procedure. Fig. 7A,B shows the cumulative amounts of lactic and glycolic acids produced as a result of degradation of PLGA 50:50 implants. The amount of lactic acid produced is greater than glycolic acid, and it is also detected later in the medium. Lactic acid has a bulkier methyl group attached to the a -carbon atom, and therefore, compared to glycolic acid, is

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169

Fig. 5. Changes in mass and medium pH during degradation of PLGA 85:15 (A) and PLGA 50:50 (B) implants.

Fig. 6. Cumulative amounts of lactic (A) and glycolic (B) acids produced during degradation of PLGA 85:15 implants.

Table 2 Comparative mass loss data at pH 7.4 Polymer PLGA 85:15 Polar Non-polar PLGA 50:50 Polar Non-polar

* t onset (weeks)

Mass loss (%)

k** (week 21 )

r 2 * **

23 9

0 85

0.202 0.165

0.990 0.927

0 74

0.457 0.589

0.973 0.957

6.5 4

Times of onset of implant mass loss, first-order degradation rate constants and correlation coefficients for placebo PLGA 85:15 and 50:50 implants prepared by polar and non-polar methods are shown. *Time of onset of mass loss. **First-order rate constant for mass loss. ***Correlation coefficient.

more hydrophobic and probably less susceptible to attack by the hydrolysis medium.

3.2.2. Scanning electron microscopy studies of PLGA 85:15 implants Representative scanning electron photomicrographs (SEMs), (magnification 5 3341) of PLGA 85:15 implants prepared by the non-polar and polar methods are shown in Fig. 8 and Fig. 9. These SEMs illustrate that significant differences were observed in the surface characteristics of the implants prepared by the two methods. Initially (t 5 0), the surface of implants prepared by both procedures was smooth

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3.2.3. Effect of pH on the degradation of PLGA 50:50 implants The results of the effect of medium pH on the degradation of PLGA 50:50 implants prepared by both polar and non-polar methods are summarized in Table 3. At all three pHs studied, i.e., 4.5, 7.4 and 9.4, degradation was more rapid for implants made by the non-polar method. For both manufacturing procedures, degradation was slowest at pH 4.5 and fastest at pH 9.4. These results are consistent with general, acid-base catalyzed type hydrolysis. The results of this study conform to the hypothesis of St. Pierre et al. [19], who propose that if the contribution of the buffer components to catalysis can be disregarded, the following order: KOH . KH .. KH 2 O , usually applies to the decomposition of simple esters with the pH-rate profile going through a minimum between pH 5 and 6. The increased rate of degradation at lower pH values reported by Makino et al., [11] is also consistent with this hypothesis.

4. Conclusions

Fig. 7. Cumulative amounts of lactic (A) and glycolic (B) acids produced during degradation of PLGA 50:50 implants.

(Figs. 8 and 9, top). Whereas little evidence of cracks and pores on the surface of implants prepared by polar procedure was observed after 4 weeks of degradation (Fig. 9, bottom), distinct changes were observed for implants prepared by non-polar procedure (Fig. 8, bottom). The original smooth surface was transformed to an irregular surface with the appearance of pores and channels in implants prepared by the non-polar procedure. These observations indicate that water uptake was faster in implants prepared by non-polar procedure resulting in a greater extent of swelling and faster degradation.

Two methods, using aqueous and organic nonsolvents, were developed for the preparation of PLGA 85:15 and 50:50 microcapsules by nonsolvent-induced phase separation. The polar method was innovative in that it is consistent with the current trends of minimizing the use of organic solvents in the fabrication of drug delivery systems. The SEMs indicated that water uptake was faster in implants prepared by the non-polar method resulting in a more porous matrix which degrades more rapidly. PLGA 50:50 implants prepared by both polar and non-polar procedures degraded by general, acid-base catalysis. The decrease in polymer number- and weight-average molecular weights (M n and M w ) occurred before the onset of implant mass loss, after which implant mass loss followed pseudo first-order kinetics. The monomers, lactic and glycolic acids were detectable at the time which corresponded to the initiation of implant mass loss and also resulted in decrease in pH of the bulk degradation medium. Finally, the average degradation time for PLGA 85:15 was 26 weeks and that for PLGA 50:50 was 6–8 weeks, which is consistent with literature reports [1]. These mi-

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171

Fig. 8. Scanning electron micrographs of PLGA 85:15 implants prepared by non-polar procedure at t 5 0 (top) and t 5 4 (bottom) weeks of degradation.

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Fig. 9. Scanning electron micrographs of PLGA 85:15 implants prepared by polar procedure at t 5 0 (top) and t 5 4 (bottom) weeks of degradation.

M. Ramchandani et al. / Journal of Controlled Release 43 (1997) 161 – 173 Table 3 Effect of medium pH on degradation of PLGA 50:50 implants prepared by polar and non-polar methods Factor

t *onset (weeks) % Mass loss at 6.5 weeks

Procedure

Polar Non-polar Polar Non-polar

pH 4.5

7.4

9.4

7.14 5.00 0 27

6.60 4.00 0 74

6.40 3.40 15 80

*Time of onset of implant mass loss.

crocapsules and implants can form the basis for several controlled-release drug delivery systems.

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