Microsphere size, precipitation kinetics and drug distribution control drug release from...

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Journal of Controlled Release 94 (2004) 129–141

Microsphere size, precipitation kinetics and drug distribution

control drug release from biodegradable

polyanhydride microspheres

Cory Berklanda, Matt J. Kipperb, Balaji Narasimhanb,Kyekyoon (Kevin) Kimc, Daniel W. Packa,*

aDepartment of Chemical and Biomolecular Engineering, University of Illinois, Box C-3, 600 S. Matthews Ave., Urbana, IL 61801, USAbDepartment of Chemical Engineering, Iowa State University, Ames, IA 50011, USA

cDepartment of Electrical and Computer Engineering, University of Illinois, Urbana, IL 61801, USA

Received 24 September 2003; accepted 27 September 2003

Abstract

A thorough understanding of the factors affecting drug release mechanisms from surface-erodible polymer devices is critical

to the design of optimal delivery systems. Poly(sebacic anhydride) (PSA) microspheres were loaded with three model drug

compounds (rhodamine B, p-nitroaniline and piroxicam) with a range of polarities (water solubilities). The drug release profiles

from monodisperse particles of three different sizes were compared to release from polydisperse microspheres. Each of the

model drugs exhibited different release mechanisms. Drug distribution within the polymer was investigated by laser scanning

confocal microscopy and scanning electron microscopy. Rhodamine, the most hydrophilic compound investigated, was

localized strongly toward the microsphere surface, while the much more hydrophobic compound, piroxicam, distributed more

evenly. Furthermore, all three compounds were most uniformly distributed in the smallest microspheres, most likely due to the

competing effects of drug diffusion out of the nascent polymer droplets and the precipitation of polymer upon solvent

extraction, which effectively ‘‘traps’’ the drug in the polymer matrix. The differing drug distributions were manifested in the

drug release profiles. Rhodamine was released very quickly independent of microsphere size. Thus, extended release profiles

may not be obtainable if the drug strongly redistributes in the microspheres. The release of p-nitroaniline was more prolonged,

but still showed little dependence on microsphere size. Hence, when water-soluble drugs are encapsulated with hydrophobic

polymers, it may be difficult to tailor release profiles by controlling microsphere size. The piroxicam-loaded microspheres

exhibit the most interesting release profiles, showing that release duration can be increased by decreasing microsphere size,

resulting in a more uniform drug distribution.

D 2003 Elsevier B.V. All rights reserved.

Keywords: Controlled release; Uniform microspheres; Polyanhydrides

0168-3659/$ - see front matter D 2003 Elsevier B.V. All rights reserved.

doi:10.1016/j.jconrel.2003.09.011

* Corresponding author. Tel.: +1-217-244-2816; fax: +1-217-

333-5052.

E-mail address: [email protected] (D.W. Pack).

1. Introduction

Biodegradable, microsphere-based controlled re-

lease systems are widely studied drug delivery devi-

ces. By delivering therapeutic at a controlled rate

C. Berkland et al. / Journal of Controlled Release 94 (2004) 129–141130

over a prolonged time, such devices can maintain

optimal drug concentrations [1–4], protect and sta-

bilize the drug [5] and aid patient compliance by

reducing the frequency of administration. In addition,

microspheres are easily administered by injection,

and they do not require surgical removal. Drug

release rates can be controlled by manipulation of

the particle size [6–9], the polymer degradation and/

or erosion rates [9,10], and polymer erosion mecha-

nism (bulk vs. surface erosion), among other factors.

Surface eroding polymers, such as polyanhydrides,

may simplify the drug release kinetics because water

penetration into the microsphere interior is mini-

mized, and the drug release rate becomes dependent

predominantly on the polymer erosion rate [11].

However, in practice the situation is often not so

simple (see Ref. [12] for examples). Efficient, repro-

ducible design of controlled release delivery systems

will require increased understanding of the release

mechanisms.

Polyanhydrides are a promising class of materials

for controlled release devices [1–4,13–17]. These

polymers are relatively hydrophobic, yet hydrolysis

of the anhydride bond is typically fast, resulting in

predominantly surface-eroding devices [4,11]. Varia-

tion of monomer chemistries and co-monomer ratios

provides erosion durations ranging from weeks to

months [4]. In addition, they are relatively non-toxic

and biocompatible [16,17]. Polyanhydride-based wa-

fers, employing poly(1,3-bis-p-carboxyphenoxy pro-

pane-co-sebacic acid) (20:80 CPP:SA), that deliver

carmustine for treatment of brain cancer have been

approved by the FDA.

A variety of factors affect drug release rates from

polyanhydride microspheres. Because the polymer is

surface eroding, the polymer degradation/erosion

often dominates release. However, drug release and

polymer erosion rates are sometimes not correlated,

depending on the copolymer composition, the rela-

tive hydrophobicity of the co-monomers and the

miscibility of the drug with the polymer [18,19].

Previous studies have shown that copolymers rich in

one component may microphase separate forming

two amorphous phases with different compositions

[12,20]. Thus, encapsulated compounds may parti-

tion within the microsphere to the more thermody-

namically favorable phase, resulting in drug release

profiles controlled by the more complex erosion

kinetics of the microphase separated device. Clearly,

the interactions between the polymer and drug

should be considered in understanding and manipu-

lating the mechanisms of release from polyanhydride

microspheres.

Microsphere size is another critical factor deter-

mining drug release rates. In many cases, smaller

microspheres release faster due to increased surface

area/volume ratio. However, the effects of micro-

sphere size can be more complex. For example,

particle size can affect the relative importance of

polymer degradation in controlling release rates.

Small (< 20-Am diameter), uniform poly(lactide-co-

glycolide) (PLG) microspheres exhibit diffusion-con-

trolled release of small-molecule compounds such as

rhodamine and piroxicam, whereas for uniform par-

ticles larger than 50 Am, polymer degradation plays

an increasingly important role [21]. In addition, we

have recently found that microsphere size can strong-

ly affect the spatial distribution of encapsulated

compounds within PLG particles [22]. Highly wa-

ter-soluble compounds tended to migrate toward the

microsphere surface during the production process.

Yet, we observed that small microspheres, which

harden quickly, appear to kinetically trap the drug

in the microsphere interior, resulting in a relatively

uniform drug distribution in the particle. Thus, the

relative hydrophilicity/hydrophobicity of the drug

and polymer, the microsphere size and the precipi-

tation kinetics all may play key roles in determining

drug distribution within and drug release mecha-

nism(s) from surface erodible polyanhydrides.

This work examines the role of all three of these

factors on the mechanisms of drug release from

poly(sebacic anhydride) (PSA) microspheres. Our

ability to fabricate monodisperse particles [23] allows

us to investigate these effects without interference

from the broad size distributions typical in previous

studies. We chose three model compounds, rhodamine,

p-nitroaniline and piroxicam, with varying water sol-

ubilities (Table 1) to investigate the role of different

‘‘compatibility’’ with the relatively non-polar PSA on

the drug release mechanism. However, the water-

solubilities of the drugs were not sufficient to interpret

the release profiles. Hence, we have examined the

effect of particle size on drug distribution within and

release from uniform and non-uniform microspheres.

These studies provide new insights into the factors

Table 1

Microsphere size, theoretical drug loading and encapsulation efficiency of various microsphere samples

Drug Water

solubility

(mg/ml)

Mean diameter

F S.D. (Am)

Drug

released

(Ag/mg)

Theoretical

loading

(Ag/mg)

Encapsulation

efficiency

Rhodamine B 8 18F 7 6.2 30 21%

52F 8 6.6 30 18%

f 88F 37 (non-uniform) 9.4 30 31%

p-Nitroaniline 1 18F 7 14 100 14%

52F 7 16 100 16%

f 120F n/a 21 100 21%

f 124F 27 (non-uniform) 34 100 34%

Piroxicam 0.1 20F 6 34 100 34%

54F 6 13 100 13%

f 112F n/a 15 100 15%

f 111F 37 (non-uniform) 50 100 50%

C. Berkland et al. / Journal of Controlled Release 94 (2004) 129–141 131

affecting the drug release kinetics from surface eroding

polymer microspheres.

2. Materials and methods

2.1. Materials

PSA was synthesized by melt polycondensation of

acetylated prepolymer under vacuum as previously

described [20]. The polymer was characterized by

GPC (Mw = 17000, PDI = 3.7), DSC (Tm = 81 jC,X = 61%; DSC7, Perkin Elmer, Shelton CT) and 1H

NMR (in deuterated chloroform; VXR 300, Varian

Palo Alto, CA). GPC was conducted in chloroform

with PL Gel columns (Polymer Laboratories, Amherst,

MA) and calibrated to poly(methyl methacrylate)

standards (Fluka, Milwaukee, WI). Poly(vinyl alcohol)

(PVA; 88% hydrolyzed) was supplied by Polysciences

(Warrington, PA). Rhodamine B chloride and p-nitro-

aniline were obtained from Sigma. Piroxicamwas a gift

fromDongwha Pharmaceuticals (Seoul, Korea). HPLC

grade dichloromethane (DCM) was purchased from

Fisher Scientific.

2.2. Preparation of microspheres

Precision particle fabrication technology, originally

developed by K.K. [24–29] and previously reported

[23,30], was used to fabricate uniform PSA micro-

spheres. Briefly, solutions of 20% w/v PSA in DCM

containing 3% w/w rhodamine B, 10% w/w p-nitro-

aniline or 10% w/w piroxicam were pumped through

a small-gauge hypodermic needle at various flow

rates. An ultrasonic transducer (Branson Ultrasonics)

controlled by a frequency generator (Hewlett Packard

model 3325A) acoustically excited the emerging

polymer jet, disrupting the stream into a train of

uniform droplets. A carrier stream (1% w/v PVA in

deionized water) flowed around the DCM/PSA/drug

stream to thin the jet to the desired diameter. The

emerging droplets were collected inf 1 l of 1% PVA.

Microspheres were hardened with stirring for 3 h, then

filtered and rinsed with deionized water. Microspheres

were lyophilized for a minimum of 48 h and stored at

� 20 jC under desiccant.

Non-uniform PSA microspheres were fabricated

using a traditional stirred emulsion method wherein

2 ml of polymer solution containing the same polymer

and drug concentrations to those used for uniform

microsphere production were added to 50 ml of 1%

PVA and stirred vigorously for 30 s. To facilitate

solvent extraction, an additional 50 ml of PVA solu-

tion were then added. Non-uniform PSA microspheres

were hardened, filtered and rinsed analogously to

uniform microsphere preparations.

2.3. In vitro drug release

To determine drug release, f 5 mg of each

microsphere formulation were suspended in 1.3 ml

of phosphate buffered saline (PBS, pH 7.4) contain-

ing 0.5% Tween 20. Microsphere suspensions were

continuously agitated by inversion (at f 10 rpm) in a


37 jC incubator. At certain time points, the samples

were centrifuged, 1 ml of supernatant was removed

and the medium was replaced with fresh PBS. Tubes

were briefly vortexed to resuspend the microspheres.

Concentration of rhodamine B in the supernatant was

determined by measuring the absorbance at 550 nm

in a multi-well plate spectrophotometer (Molecular

Devices Spectra Max 340PC). p-Nitroaniline and

piroxicam release were subjected to the same release

Fig. 1. Scanning electron micrographs depicting the uniformity of unlo

morphology. Scale bar = 10 Am.

conditions. However, the concentrations of p-nitro-

aniline and piroxicam in the supernatant were

obtained by measuring the absorbance at 387 and

276 nm, respectively. The microsphere loading is

reported as the total amount of drug obtained at the

conclusion of the release experiments, and these

values are within 5–10% of loading measured by

assaying for concentration of drug recovered from

fresh microspheres (by accelerated degradation of

aded 20-Am PSA microspheres often exhibiting a rough surface


microspheres in 0.2 N NaOH at 37 jC for 48 h).

After the release experiment, no residual, unreleased

drug was detectable by fluorescence microscopy or

spectrophotometric analysis of the residual samples

dissolved/degraded in 0.2 N NaOH. Absorbance

spectra of drug, obtained from fresh solutions and

supernatant sampled at various times during the

release experiment, were identical in shape indicating

that there was no degradation of drug during the

release period.

2.4. Scanning electron microscopy

Microsphere surface and interior morphology was

observed by scanning electron microscopy (Hitachi S-

4700). Several droplets containing different formula-

tions of microspheres suspended in water were placed

onto a SEM sample stage. The samples were lyophi-

Fig. 2. Size distributions of p-nitroaniline- (A and C) an

lized overnight. Cross-sections were obtained by freez-

ing lyophilized microspheres on the SEM stage with

liquid nitrogen and chopping with a razor blade.

Sectioned and whole microsphere samples were sputter

coated with gold prior to imaging.

2.5. Confocal fluorescence microscopy

Drug distribution within PSA microspheres was

determined by confocal fluorescence microscopy prior

to in vitro release. A small amount (f 1 mg) of each

sample was suspended in distilled water, placed on

Petri dishes and dried overnight. Microspheres were

then imaged using an Olympus Fluoview FV300 Laser

Scanning Biological Microscope. Rhodamine was

excited with a krypton laser, while p-nitroaniline and

piroxicam were excited with a helium/neon laser.

Optical cross-sections were taken at various depths

d piroxicam-loaded (B and D) PSA microspheres.


for each microsphere size and drug type to determine

drug distribution at the microsphere midline.

2.6. Particle size distribution

A Coulter Multisizer 3 (Beckman Coulter) deter-

mined the size distribution of the various sphere

preparations. The lyophilized particles were resus-

pended in Isoton electrolyte with a type II-A disper-

sant to prevent microsphere aggregation. A minimum

of 5000 microspheres was analyzed for each sample.

3. Results

3.1. Production of uniform poly(sebacic anhydride)

microspheres

Monodisperse PSA microspheres were produced

using precision particle fabrication technology as

reported previously for PLG microspheres [23,30].

PSA microspheres exhibited a non-spherical surface

morphology (Fig. 1) that has been observed previ-

ously [31]. ‘‘Folds’’ in the surface often penetrate to

near the microsphere center. Typically, over 90% of

microspheres are within 1.5 Am of the average

diameter. To our knowledge, this is the first study

reporting the production of uniform polyanhydride

microspheres. Size distributions for p-nitroaniline-

loaded and piroxicam-loaded microspheres show

Fig. 3. In vitro, normalized release of rhodamine (A), p-nitroaniline (B), an

the relative uniformity of precision particle fabrica-

tion (Fig. 2A and B) versus typical emulsion micro-

sphere production methods (Fig. 2C and D). Bimodal

distributions (e.g. 20-Am piroxicam microspheres,

Fig. 2B) are the result of fusion of nascent PSA

droplets.

3.2. Drug loading and in vitro drug release

Three model drugs exhibiting various degrees of

water solubility were encapsulated in PSA micro-

spheres (Table 1). Piroxicam, having the lowest solu-

bility in water (f 100 Ag/ml), showed encapsulation

efficiency in the range 13–34%. The p-nitroaniline

solubility in water isf 1 mg/ml, and the encapsulation

efficiency ranged from 14% to 21%, depending on the

particle size. The encapsulation efficiency of rhoda-

mine (21–31%) was higher than that for p-nitroaniline,

which is contradictory to expectations based on drug

solubility. However, the theoretical loading of rhoda-

mine was only 30 Ag/mg (drug/polymer) compared to

100 Ag/mg for p-nitroaniline and piroxicam. Lower

loading typically results in higher encapsulation effi-

ciency due to smaller concentration gradients driving

the drug out of the polymer/solvent droplets. Thus, the

rhodamine encapsulation efficiency cannot be directly

compared.

In general, previous studies with other degradable

polymers (e.g., PLG) have shown that smaller,f 10–

20-Am, microspheres have higher drug encapsulation

d piroxicam (C) from uniform and non-uniform PSA microspheres.


efficiency than larger microspheres, probably due to

rapid precipitation of these small microspheres [22].

Also, larger f 80- to 120-Am microspheres tend to

Fig. 4. Scanning laser confocal fluorescence micrographs revealing the

microspheres (A and B), 18-, 52- and 120-Am p-nitroaniline-loaded m

microspheres (F–H). Scale bar = 50 Am (same magnification in all image

encapsulate drugmore efficiently than ‘‘medium’’-size,

f 50-Am microspheres. The dependence of encapsu-

lation efficiency on particle size is most likely the result

distribution of drug in uniform 18- and 52-Am rhodamine-loaded

icrospheres (C–E), and 20-, 54- and 112-Am piroxicam-loaded

s).

Fig. 5. Scanning electron micrographs of uniform 18- and 52-Am rhodamine-loaded microspheres (A and B, respectively), 18- and 52-Am p-

nitroaniline-loaded microspheres (C and D, respectively), and 20- and 54-Am piroxicam-loaded microspheres (E and F, respectively). Scale

bar = 10 Am (scale bar in E applies to A and C; scale bar in F applies to B and D).



of competition between (1) decreased diffusion of drug

out of the particle as size increases, which tends to

increase encapsulation efficiency, and (2) decreased

rate of entrapment of drug due to slower ‘‘hardening’’

of the particles as size increases, which tends to

decrease encapsulation efficiency. Thus, there is a

minimum in encapsulation efficiency for particles near

50-Am in diameter where the rate of hardening is

significantly slower than in 10-Am particles, allowing

time for drug diffusion out of the particles, and the rate

of diffusional loss is still faster than in the larger

particles. The results reported here, as well as previous

Fig. 6. Scanning electron micrographs of the surface structure of unif

respectively), 18- and 52-Am p-nitroaniline-loaded microspheres (C and D

(E and F, respectively). Arrows indicate drug crystals. Scale bar = 3 Am

studies [30], are consistent with these hypotheses. Non-

uniform microspheres exhibited the highest drug en-

capsulation efficiency, possibly due to the presence of

many small, < 10-Am diameter, microspheres and

faster polymer precipitation due to rapid stirring rates

in the non-solvent hardening bath.

The effect of microsphere size on drug release was

very different for the three compounds (Fig. 3).

Rhodamine release was rapid and complete within

3 days. The profiles exhibited a smooth concave

downward shape that was independent of micro-

sphere size. p-Nitroaniline release profiles were sim-

orm 18- and 52-Am rhodamine-loaded microspheres (A and B,

, respectively), and 20- and 54-Am piroxicam-loaded microspheres

(same magnification in all panels).

C. Berkland et al. / Journal of Contro138

ilar to rhodamine with a longer duration of release.

Again, no distinction could be made between micro-

spheres of different diameter. However, p-nitroaniline

release appeared to be biphasic, exhibiting a drug

burst followed by a period of slower release. Pirox-

icam release, in contrast, varied significantly with

microsphere size. Large, uniform (f 112 Am) and

non-uniform (average = 111 Am) microspheres re-

leased drug rapidly, similar to drug-release profiles

described for rhodamine and p-nitroaniline. Interme-

diate, 54-Am microspheres showed a burst of pirox-

icam followed by a 5-day ‘‘lag’’ during which little

drug was released and a more rapid release phase

between days 5 and 8. Finally, the 20-Am micro-

spheres exhibited a much smaller burst followed by a

shorter lag and a 5- to 6-day period of near zero-order

release. It is clear from these data that microsphere

size alone does not determine the drug release profile.

Fig. 7. Scanning electron micrographs of the interior structure of unifo

piroxicam-loaded microspheres (C and D). Arrows indicate drug crystals. S

applies to D).

3.3. Distribution of drugs within the polymer matrix

The distribution of drugs within the microspheres

was examined via laser scanning confocal microsco-

py (Fig. 4). Rhodamine appears to be distributed

preferentially to the surface of the 18- and 52-Ammicrospheres. p-Nitroaniline distributed more evenly

throughout the smallest microspheres, but was local-

ized increasingly toward the microsphere surface as

the size increased. Similarly, piroxicam distributed to

the microsphere surface in large (f 112-Am) micro-

spheres and was rather evenly distributed in the 20-Ammicrospheres. The grainy appearance of the fluores-

cence may be indicative of the larger drug crystals

present in p-nitroaniline and piroxicam-loaded micro-

spheres (vide infra). The ‘‘lines’’ of drug apparently

running through the center of 52-Am p-nitroaniline-

loaded and 54-Am piroxicam-loaded microspheres are

lled Release 94 (2004) 129–141

rm 18-Am rhodamine-loaded microspheres (A and B) and 20-Amcale bar = 3 Am (scale bar in A also applies to C; scale bar in B also


the result of folds in the microsphere surface (see Fig.

1) and actually represent surface localized drug.

Scanning electron micrographs confirm the distri-

bution of drug in microspheres and provide a compar-

ison of microsphere morphology amongst the different

drug formulations. Rhodamine-loaded microspheres

exhibit a folded and somewhat porous microsphere

surface for both 18-Am (Fig. 5A) and 52-Am micro-

spheres (Fig. 5B) as well as a large amount of surface

localized drug crystals. The size and density of rhoda-

mine crystals on the surface appear to be similar for

both microsphere sizes (Fig. 6A and B). Folds are less

apparent in 18-Am (Fig. 5C) and 52-Am (Fig. 5D) p-

nitroaniline-loaded microspheres. However, larger

drug crystals are evident, especially in 52-Am micro-

spheres (compare Fig. 6C and D). Piroxicam-loaded

microspheres also exhibit larger crystals and a crack-

like pore structure for 20-Am (Fig. 5E) and 54-Am (Fig.

5F) microspheres. The density of piroxicam crystals on

the microsphere surface is lower compared to the p-

nitroaniline or rhodamine crystals (compare Fig. 6E

and F to A, B, C and D).

Scanning electron micrographs of fractured micro-

spheres reveal dense interior structures, and the

presence or absence of drug crystals can be visual-

ized. A cross-section of an 18-Am rhodamine-loaded

microsphere shows no evidence of rhodamine inside

the microsphere (Fig. 7A and B). However, a cross-

section of a 20-Am piroxicam-loaded microsphere

reveals large, f 1- to 3-Am drug crystals in the

microsphere interior confirming the presence of drug

throughout the microsphere matrix (Fig. 7C and D).

4. Discussion

Drug delivery applications utilizing biodegradable

polymeric microspheres are becoming an important

means of delivering therapeutic agents since micro-

spheres offer facile, noninvasive administration via

injection, protection of encapsulated drug and sus-

tained drug release over periods from hours to months

or even years [32,33]. In the past, many factors have

been adjusted to produce a desired drug release profile

including microsphere fabrication conditions, which

usually affect the resulting microsphere size distribu-

tion [6,34], and polymer chemistry (e.g., polymer

composition, co-monomer ratios, molecular weight,

etc.) [9,35]. Defining a set of these formulation param-

eters offers a degree of reproducibility, but the precise

mechanisms affecting drug release kinetics can be

difficult to determine. Here, we have utilized a tech-

nology for precision particle fabrication to more accu-

rately investigate factors influencing drug release. By

examining uniform microspheres over a size range

suitable for subcutaneous or intramuscular drug depots

(f 10–120 Am), we are able to see differences in

encapsulation efficiency, drug distribution within the

microsphere matrix and, most importantly, drug release

kinetics, which would be difficult to ascertain from a

non-uniform microsphere population.

Several different drug types exemplifying various

degrees of solubility in water were encapsulated in

uniform and non-uniform PSA microspheres. The

encapsulation efficiency of these drugs appeared to

increase slightly as drug solubility in water de-

creased. Polyanhydrides are relatively hydrophobic

polymers [4,11]. Therefore, improved incorporation

of drugs having low water solubility may be ex-

pected. Additionally, the relative hydrophobicity of

the drug determined the distribution of drug within

the microspheres. Water-soluble rhodamine (f 8 mg/

ml) exhibited a strong preference toward the micro-

sphere surface while the relatively water insoluble

piroxicam (f 100 Ag/ml) remained more evenly

distributed within the hydrophobic microsphere ma-

trix, especially for small (20-Am), rapidly precipitat-

ing microspheres (Fig. 4).

The effect of drug distribution, as controlled by

microsphere size, was manifested in the drug re-

lease profiles. Surface-localized crystals, evident in

scanning electron micrographs (Fig. 5), may be

responsible for the rapid initial release of rhoda-

mine and p-nitroaniline. The surface localization of

rhodamine and p-nitroaniline for all microsphere

sizes, and for piroxicam in large (>50-Am) micro-

spheres, further explains the rapid initial drug

release and short duration of release for these

formulations. Hydrophilic drugs, especially organic

bases, are also known to increase the rate of

polymer degradation [36], which may further ex-

plain the rapid release of rhodamine. In addition,

polymer-soluble drugs like p-nitroaniline can de-

crease the crystallinity of PSA, increasing the

polymer erosion rate [20]. This effect may also

contribute to the rapid release of p-nitroaniline.


Piroxicam release appears to be controlled primarily

by the distribution of drug within the polymer matrix

(Fig. 4). Surface-localized piroxicam was not as prev-

alent as rhodamine or p-nitroaniline (Fig. 6), and

piroxicam was present throughout the polymer matrix

(Fig. 7C and D). Laser scanning confocal microscopy

confirmed an increasingly uniform drug distribution as

microsphere size decreased, probably due to faster

precipitation of the smaller nascent PSA drops. Uni-

form drug distribution throughout the 20-Am micro-

sphere matrix resulted in nearly zero-order piroxicam

release controlled by polymer degradation. Further, the

20-Am piroxicam microsphere formulation was the

only one that sustained drug release during the life of

the microspheres [37], which supports the supposition

that drug is distributed throughout the microsphere

matrix and not just on the surface. The complex

piroxicam release profile generated by 54-Am micro-

spheres may best be explained as a combination of

release of surface localized drug, as in the f 112-Ampiroxicam-loaded microspheres, plus controlled re-

lease of drug distributed to the microsphere interior,

as in the 20-Am microspheres.

Several factors are desirable to create an improved

PSA microsphere system capable of zero-order re-

lease. Encapsulating drug as a suspension of solids

or increasing polymer solution viscosity by increas-

ing the initial polymer concentration or molecular

weight would decrease drug mobility improving

encapsulation efficiency and reducing surface local-

ization of drug. More rapid phase inversion of

nascent polymer droplets by rapid stirring or in-

creased volume of non-solvent would also minimize

redistribution of encapsulated drug. Each of these

strategies may be most efficiently investigated by

producing uniform microspheres to avoid the con-

founding effects of broad size distributions.

5. Conclusions

Precision particle fabrication technology can be

used to create monodisperse PSA microspheres of a

desired size suitable for subcutaneous or intramuscular

injection. As a direct result, different microsphere sizes

were found to exhibit various degrees of surface

localization of drug according to the relative solubility

of the drug in water and the precipitation rate of nascent

polymer droplets. Water-soluble compounds, rhoda-

mine and p-nitroaniline, localized preferentially to the

microsphere surface, resulting in rapid drug release for

only a few days. For piroxicam-loaded microspheres,

the particle size dictated the release profile, albeit in a

surprising manner. If the drug is evenly distributed in

the microspheres, the release duration should increase

as the particle size increases. However, we showed here

that the opposite trend is obtained when microsphere

precipitation kinetics play an important role in deter-

mining drug distribution. In this study, the smallest,

most rapidly precipitating microspheres (20-Am) had

the most homogeneous drug distribution and thus the

most extended release profile, while the largest par-

ticles released the fastest. This study emphasizes the

importance of the interplay between the drug water

solubility (suggestive of ‘‘compatibility’’ with the

highly non-polar polymer matrix), microsphere size

and polymer precipitation kinetics on the drug distri-

bution within and release from surface-eroding poly-

mer microspheres. A more thorough understanding of

all of the factors governing drug release kinetics is

essential for precise design of controlled-release devi-

ces based on this and similar polymer systems.

Acknowledgements

We thank Roshelle Silverman for assistance with

drug release studies. This work was supported by

grants from the Whitaker Foundation, Roy S. Carver

Trust and Iowa State University Office of Biotech-

nology to BN.

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