Macromolecule release from monodisperse PLG microspheres: Control of release rates and investigation...

Macromolecule Release from Monodisperse PLGMicrospheres: Control of Release Rates and Investigationof Release Mechanism

CORY BERKLAND,1 EMILY POLLAUF,1 CHANDRASHEKAR RAMAN,1 ROSHELLE SILVERMAN,1

KYEKYOON (KEVIN) KIM,2,3 DANIEL W. PACK1,3

1Department of Chemical and Biomolecular Engineering, University of Illinois, Urbana, Illinois 61801

2Department of Electrical and Computer Engineering, University of Illinois, Urbana, Illinois 61801

3Department of Bioengineering, University of Illinois, Urbana, Illinois 61801

Received 25 May 2006; revised 28 November 2006; accepted 29 November 2006

Published online in Wiley InterScience (www.interscience.wiley.com). DOI 10.1002/jps.20948

ABSTRACT: Novel macromolecular therapeutics such as peptides, proteins, and DNAare advancing rapidly toward the clinic. Because of typically low oral bioavailability,macromolecule delivery requires invasive methods such as frequently repeatedinjections. Parenteral depots including biodegradable polymer microspheres offer thepossibility of reduced dosing frequency but are limited by the inability to adequatelycontrol delivery rates. To control release and investigate release mechanisms, we haveencapsulated model macromolecules in monodisperse poly(D,L-lactide-co-glycolide)(PLG) microspheres using a double-emulsion method in combination with the precisionparticle fabrication technique. We encapsulated fluorescein-dextran (F-Dex) andsulforhodamine B-labeled bovine serum albumin (R-BSA) into PLG microspheres ofthree different sizes: 31, 44, and 80 mm and 34, 47, and 85 mm diameter for F-Dex andR-BSA, respectively. The in vitro release profiles of both compounds showed negligibleinitial burst. During degradation and release, the microspheres hollowed and swelled atcritical time points dependant uponmicrosphere size. The rate of these events increasedwithmicrosphere size resulting in the largestmicrospheres exhibiting the fastest overallrelease rate. Monodisperse microspheres may represent a new delivery system fortherapeutic proteins and DNA and provide enhanced control of delivery rates usingsimple injectable depot formulations. � 2007 Wiley-Liss, Inc. and the American Pharmacists

Association J Pharm Sci 96:1176–1191, 2007

Keywords: controlled release; uniform microspheres; poly(D,L-lactide-co-glycolide);bovine serum albumin; dextran

INTRODUCTION

Macromolecules such as proteins, poly-saccharides, and DNA are rapidly developing asspecific and potent therapeutics. However, macro-

molecules present many new challenges for theirformulation and delivery. Often, the deliverymethod can impact efficacy—and certainly com-mercialization—as much as the nature ofthe drug itself. Although there has been somesuccess in pulmonary, transdermal, and evenoral delivery of macromolecular drugs, mostcurrent protein-based products require frequentlyrepeated injections.1–5 In addition, many macro-molecular therapeutics require stringent controlof the in vivo concentration and localization,6,7

1176 JOURNAL OF PHARMACEUTICAL SCIENCES, VOL. 96, NO. 5, MAY 2007

Cory Berkland’s present address is 2030 Becker Dr.,Lawrence, KS 66045.

Correspondence to: Cory Berkland (Telephone: 785-864-1455; Fax: 785-864-1454; E-mail: [email protected])

Journal of Pharmaceutical Sciences, Vol. 96, 1176–1191 (2007)� 2007 Wiley-Liss, Inc. and the American Pharmacists Association

sometimes necessitating the use of invasivemechanical devices such as implantable pumps.In response, research in advanced drug deliverysystems seeks to provide safe, effective, andsimple methods for administration with spatialand temporal control of drug concentrations.

Controlled release systems, especially biode-gradable polymermicrospheres, are a popular andpromising drug delivery method for small-mole-cule and macromolecular drugs. Polymer micro-spheres are relatively easy to fabricate, can beadministered via a simple injection, and canrelease drug for weeks to months.8 Poly(D,L-lactide-co-glycolide) (PLG) is perhaps the mostcommonly used polymer due to its relatively highbiocompatibility and the availability of numerouspolymers with varying monomer ratios and mole-cular weights. Nonetheless, PLG microspherespresent their own formulation challenges. Pro-teins and nucleic acids are often easily deactivatedduring fabrication or after administration due toadsorption to polymer surfaces; chemical changes,denaturation, or aggregation caused by harshenvironmental conditions such as the presenceof organic solvents or acidic pH9,10; or in vivoenzymatic activity.8,11,12 Such problems oftennecessitate modified processing techniques,13 theinclusion of excipients,14,15 or the synthesis ofentirely new biomaterials, any of whichmay affectthe subsequent drug release profile.

A number of reports have attempted to explainthe mechanisms of macromolecule release fromPLG microspheres (e.g., see16–19). Nevertheless,design of PLG microsphere systems for macro-molecule delivery is hindered by the complexity ofthe problem and incomplete understanding of themechanisms governing polymer degradation, ero-sion and drug release.20 It is often believed thatprotein release rates can be controlled by choosinga polymer with appropriate degradation rates, forexample, by varying lactic/glycolic acid ratios orpolymer molecular weight. However, the polymerdegradation alone does not always control drugrelease. Degradation, erosion, particle porosity,protein aggregation, protein/polymer interactions,etc. all impact release mechanisms to varyingdegrees. In addition, the size, solubility, andcharge of the macromolecule can further confoundprediction of release rates.

Most often PLG microspheres release macro-molecules with a tri-phasic shape consisting of aninitial ‘‘burst,’’ a lag period of very slow release,and finally a phase of accelerated release.16,18,19

The initial burst of protein therapeutics has been

attributed to their tendency to partition tothe microsphere surface or adsorb to the surfaceduring the encapsulation process.7 Multipleapproaches have attempted to alleviate the burstby varying the polymer chemistry,20–23 addingexcipients to the polymer phase,24,25 utilizingnovel polymers,26–30 encapsulating particulateforms of the drugs,31 or exchanging the aqueousnonsolvent phase used in the fabrication processwith nonpolar oils or alcohols to reduce the affinityof the encapsulated drugs for the bulk phase.13,32

The burst is usually followed by a lag period wherediffusion is limited and little release of themacromolecule occurs. The duration and flatnessof this phase is determined by the polymerdegradation kinetics, particle size, and micro-sphere porosity.16 In addition, the drug size,charge, and any potential interactions of the drugwith the polymer can influence the lag phase.33

During this time, however, polymer degradation(hydrolytic cleavage, decrease in molecularweight) and erosion (loss of polymer mass due torelease of soluble degradation products less thanMW �1100)34 both occur. Finally, the lag phase isfollowed by a period of steady release typicallycontrolled by the polymer degradation rate.

Initially, polymer degradation and erosionwerethought to occur homogeneously throughout PLGmicrospheres. However, increasing evidencesupports a heterogeneous process. Ester cleavagegenerates acid end groups that can contribute to alocal acidification inside the polymer matrix.Diffusional limitation of buffer componentsbetween the external aqueous phase and theinterior of the microsphere will induce a radialpH gradient with pH decreasing toward the centerof the particle.9,10 Further, diffusion of solubleoligomers and perhaps monomers (also acids) outof the microspheres may be slower than that of thebuffer components (phosphate ions). Accumula-tion of these degradation products would alsocontribute to the pH gradient in themicrospheres.Degradation of PLG is acid catalyzed.16,35,36 Thus,the core of degrading microspheres becomes moreacidic than the surface,9,10 and one should expectfaster degradation inside the microspheres thannear the periphery. This phenomenon is known asautocatalytic degradation. Nevertheless, degrada-tion and erosion ultimately lead to formation ofinterconnecting, water-filled pores. Diffusion ofmacromolecules through this pore system resultsin the final, relatively fast protein release phase.These parameters can be further confounded byrecent reports of an increase in polymer density or

MACROMOLECULE RELEASE FROM MONODISPERSE PLG MICROSPHERES 1177

DOI 10.1002/jps JOURNAL OF PHARMACEUTICAL SCIENCES, VOL. 96, NO. 5, MAY 2007

pore sealing at the surface of PLGA microspheresover the first several hours of incubation.37

The microsphere size is a critical parameterimpacting many of the factors that can controldrug release, especially drug diffusion, polymerdegradation, and erosion. In particular, the impactof autocatalytic degradation is clearly expected toincrease with device size. This complex system isdifficult to untangle in part due to the broadsize distribution inherent in most conventionalmicrosphere fabrication methods.

Precision particle fabrication (PPF) technologycan produce monodisperse PLG microspheresencapsulating small molecules and macromole-cules.38,39 Here we report monodisperse PLGmicrospheres with diameters from 30 to 85 mmencapsulating fluorescein-labeled dextran (F-Dex)and sulforhodamine B-labeled bovine serumalbumin (R-BSA). The microsphere morphologyand intraparticle drug distribution were investi-gated by scanning electron microscopy and laser-scanning confocal microscopy, respectively. Invitro release of both drugs was investigated forthe three different microsphere sizes. We alsoinvestigated the mechanisms of degradation andF-Dex andR-BSA release fromPLGmicrospheres.This study is greatly facilitated by the uniform sizeof particles generated with PPF. Light, confocalfluorescence, and scanning electron micrographsshow the progression of intraparticle drugdistribution and interior microsphere morphologyduring the in vitro release experiment.Correlationof the release data with physical changes inthe microspheres provides insight into themechanisms controlling the macromoleculerelease rates. In particular, this article shows thatautocatalytic degradation of the microspheres isan important phenomenon and depends dramati-cally on the microsphere size, even given therelatively small size and narrow size rangestudied.

MATERIALS AND METHODS

Materials

PLG copolymer (50:50 lactic acid/glycolic acid;i.v.¼ 0.17 dL/g corresponding to Mw �15,000according to the manufacturer) was obtainedfrom Birmingham Polymers (Pelham, AL).Fraction V BSA (Mw �67,000), fluorescein iso-thiocyanate-dextran (F-Dex, Mw �70,000) andsulforhodamine B acid chloride were purchased

from Sigma (St. Louis, MO). Polysciences, Inc.(Warrington, PA), provided 88% hydrolyzed poly(vinyl alcohol) (PVA). Reagent grade dichloro-methane (DCM) from Fisher (Hampton, NH) wasutilized as the polymer solvent.

Labeling of Bovine Serum Albumin

Approximately 500 mg of BSA was added to100 mL of phosphate-buffered saline solution(PBS, pH¼ 7.4) that contained 840.39 mg ofsodium bicarbonate resulting in a solution pH of8.95. Nine milligrams of sulforhodamine B acidchloride was mixed with 200 mL of DMF and thisdye solution was added to the protein solution,which was mixed for 2 h at 378C. The reaction wasstopped by the addition of 1.5 M hydroxylamineand incubating for an additional hour. To removeunbound dye, the solution was placed in anultrafiltration column (Millipore Centriplus YM-10, Billerica, MA, molecular weight cut-off100 kDa) and centrifuged at 3,000g for four30-min periods, refreshing the column andretained R-BSAwith nanopure water each period.The resulting concentrated, labeled, and salt-freeprotein solution was lyophilized and stored at�208C in the dark, under desiccant.

Microsphere Preparation

PPF technology produced monodisperse PLGmicrospheres using methods similar to thosepreviously reported.39,40 Macromolecules wereencapsulated via a double emulsion technique.A primary aqueous solution of 100 mg F-Dex orR-BSA/mL deionized water was emulsified into anoil phase of PLG (40% w/v) dissolved in DCM at a1:10 v/v ratio using a sonicator (Heat Systems-Ultrasonics, Inc., model W-220, Farmingdale, NY)fitted with a microtip operating near the microtiplimit continuously for 30 s. A small-gauge hypo-dermic needle was utilized to spray the primaryemulsion. The liquid jet emerging from the nozzlewas acoustically excited using an ultrasonictransducer (Branson Ultrasonics, Danbury, CT)controlled by an amplified signal (Trek modelPZD700) from a frequency generator (Agilentmodel 33120A). The vibration yielded regular jetinstabilities and disrupted the stream into mono-disperse polymer/drug/solvent droplets. A carrierstream (0.5% w/v PVA in distilled water) sur-rounded the emerging PLG jet. The dropletsflowed into a beaker containing approximately900 mL of 0.5% PVA. PLG/water/drug drops were

1178 BERKLAND ET AL.

JOURNAL OF PHARMACEUTICAL SCIENCES, VOL. 96, NO. 5, MAY 2007 DOI 10.1002/jps

stirred �3 h then were filtered and rinsed with anequal volume of distilled water to remove residualPVA. Lastly, microspheres were freeze-dried(Labconco benchtop model) for 2 days and storedat �208C under desiccant.

Drug Loading

F-Dex loading was determined by dissolving 2–5 mg of spheres in 250 mL of DCM. Afterdissolution of F-Dex and PLG, the solution wassplit between two microcentrifuge tubes with125 mL in each, and 1 mL of PBS containing0.005% v/v Tween-20 was added to each tube. Thetubes were incubated overnight at 378C withshaking at 225 rpm to extract the FITC-dextranfrom the organic to the aqueous phase. F-Dexconcentration was determined by fluorescencespectrometry of the isolated aqueous phase withan excitation wavelength of 491 nm and emissionwavelength of 513 nm (Varian Cary EclipseFluorescence Spectrophotometer). Concentrationstandard curves were determined at the sameconditions.

To determine the loading of R-BSA, 2–5 mg ofeachmicrosphere formulationwereweighed out intriplicate, dissolved in 1 mL of acetonitrile, thenvortexed for 30 s to completely dissolve thepolymer phase. The samples were centrifuged at12,000 rpm for 15 min to pellet undissolved BSA.The supernatant was removed and the pellet wasdissolved in 1 mL of PBS containing 0.005% v/vTween-20. The concentration of R-BSA in thesupernatant was determined by fluorescencespectrometry using excitation and emission wave-lengths of 560 and 578 nm, respectively.

In Vitro Drug Release

Approximately 15 mg of microspheres wereaccurately weighed into glass vials and 1.3 mLof PBS containing 0.005% Tween-20 were addedto each vial. The samples were incubated at 378Cwhile shaking at 225 revolutions per minute. Ateach time point, 1 mL of supernatant wasremoved and replaced with fresh buffer. Theconcentration of R-BSA and F-Dex in the super-natant for each time point was determined asdescribed for drug loading measurements. Therelease data were summed for each time point,and the total divided by the calculated drugloading (experimental loading times mass ofmicrospheres), to arrive at the ‘‘cumulativepercent released.’’

Microsphere Degradation Study

Microsphere samples for tracking PLG molecularweight were prepared to mimic in vitrorelease conditions by adding �15–20 mg of eachmicrosphere formulation to 1.3 mL of PBScontaining 0.005% Tween-20 while shaking at225 revolutions per minute in a 378C incubator.At regular time intervals, supernatant wasrefreshed and a small sample (�1 mg) of micro-spheres was removed and placed in a �208Cfreezer. At the culmination of the study, micro-spheres were centrifuged, the supernatantwas discarded, and samples were freeze-dried(Labconco benchtop model).

Confocal Microscopy

The intraparticle distribution of R-BSA andF-Dex could be tracked over time via laserscanning confocal microscopy. A small amount(<1 mg) of each formulation was suspended indistilled water and placed on glass microscopeslides. Microspheres were imaged using an Olym-pus Fluoview FV300 Laser Scanning BiologicalMicroscope to determine the distribution ofeach molecule at the midline of the microsphere.R-BSA was excited with a krypton laser (565 nm)and emission detected at 586 nm. F-Dex wasexcited with an argon laser (495 nm) and emissiondetected at 535 nm. Care was taken to ensurethe microscope settings (i.e., gain, aperture size,number of scans, etc.) were the same for allimages.

Scanning Electron Microscopy

The surface and interior morphologies of micro-spheres were imaged at various time points byscanning electron microscopy (Hitachi S-4700). Tofracture the particles, a small amount of driedmicrospheres was placed onto a glass microscopeslide and suspended immediately over liquidnitrogen. Once the microspheres were adequatelyfrozen, a razor blade was used to section theparticles, which were then rinsed with water intoa vial and allowed to settle. Approximately 20 mLof settled particles were removed and placed ontoa scanning electron microscope sample stage. Thestages were freeze dried overnight (Labconcobenchtop model) and sputter coated with gold-palladium (Emitech K-575) prior to imaging at2–10 kV.



Particle Size Distribution

Microsphere size distributions of each of sixformulations were attained by a Coulter Multi-sizer 3 (Beckman Coulter, Inc., Fullerton, CA)equipped with a 100- or 280-mm aperture. Lyo-philized particles were resuspended in Isoton IIelectrolyte and a minimum of 5,000 microsphereswere analyzed for each distribution.

Quantitation of Intraparticle FluorescenceDistribution

The images obtained from confocal microscopywere analyzed to obtain drug distributions withinthe microspheres. The images were converted tointensity plots using ImageJ software and thenconverted to radial distributions by using a simplecomputer program to average the intensity at allpoints at a given radial distance from the center ofthe microsphere.

RESULTS

Production of Uniform MicrospheresEncapsulating F-Dex and R-BSA

PPF technology38,39,41 produced uniform PLGmicrospheres encapsulating fluorescently labeledmacromolecules via a modified double emulsion/solvent extraction technique. F-Dex or R-BSA wasencapsulated at 2.5% w/w PLG resulting in highencapsulation efficiency (88–97%) for F-Dex andacceptable encapsulation efficiency (64–66%) forR-BSA (Tab. 1). PPF produced highly uniform 31,44, and 80 mm F-Dex-loaded microspheres and 34,47, and 85 mm R-BSA-loaded microspheres(Fig. 1A). Laser scanning confocal micrographsrevealed that each model drug was evenlydistributed throughout the polymer matrix forall of the formulations (Fig. 1B). F-Dex-loaded

microspheres exhibited a dense surface andinterior morphology compared to the moreporous surface and interior of the R-BSA-loadedmicrospheres (Fig. 2). The pore sizes were similarfor all three sizes of R-BSA-loaded microspheres.Although the same three sets of conditions wereutilized for fabricating F-Dex- and R-BSA-loadedmicrospheres of the three sizes, the increasedporosity resulted in R-BSA-loaded microspheresbeing slightly larger than F-Dex-loaded micro-spheres.

In Vitro Release of F-Dex and R-BSA

The in vitro release of F-Dex or R-BSA wasstudied to elucidate the effect of microsphere sizeon the release profiles of these model macromole-cules. As described above, the release of hydro-philic macromolecules often follows a tri-phasicpattern in which an initial drug burst is followedby a lag period of very slow release and a finalphase of steady, more-rapid drug release gov-erned by significantly higher effective diffusivitythrough water-filled pores.

Notably, the initial F-Dex and R-BSA releaseappears to be purely diffusion-controlled (Figs. 3Aand B and Fig. 4, discussed below). At 28 h, F-Dex-loaded microspheres had only released 1.2% of thetotal encapsulated drug for all three microspheresizes while R-BSA-loaded microspheres released1.9–4.8% depending on microsphere diameter.Rather than conventional burst and lag phases,uniformmicrospheres produced one smooth phaseof slow release. The duration of the slow releasephase varied with the microsphere diameter(Fig. 3C and D), and the rates decreased withincreasing sphere diameter (decreasing surfacearea:volume ratio).

F-Dex release profiles were very distinctivefor the three different microsphere sizes.The diffusion-controlled slow-release period was

Table 1. Characteristics of Macromolecule-Loaded Monodisperse PLGMicrospheres

AverageMicroparticleDiameter (mm)

TheoreticalLoading (w/w Polymer)

EncapsulationEfficiency

F-Dex Mw¼ 70000 31 2.5% 64� 6%44 2.5% 66� 16%80 2.5% 66� 10%

R-BSA Mw¼ 67000 34 2.5% 97� 9%47 2.5% 93� 3%85 2.5% 88� 3%



followed by a sharp increase in release rate(Fig. 3C). The time at which the releaserate increased was strongly dependent upon butinversely proportional tomicrosphere size. Thirty-one, 44, and 80 mm spheres began releasingmore quickly after �250, �75, and �25 days,respectively. Thirty-one- and 44-mmmicrospheresshowed incomplete (but still increasing) release ofF-Dex after nearly 1 year. The long duration ofrelease from microspheres of this relatively lowmolecular weight PLG is surprising, and theresults should be interpreted with some caution;however, the key finding of this article is thenotable differences inmicrosphere releasekineticsas a function of size (see Discussion).

R-BSA release profiles more smoothly transi-tioned from the lag phase to steady release(Fig. 3D). The slow phase for 34, 47, and 85 mmspheres lasted �45, �40, and �20 days, re-spectively, although the transition point for34 and 47 mm spheres was difficult to distinguish.

Again, after the slow phase, R-BSA release ratesincreased dramatically. Release of R-BSA from34- to 47-mm particles was indistinguishable from�50 to 120 days. Essentially all of the R-BSAwas released from particles of all three sizes by160 days. The final amount of R-BSA releaseappeared to be greater than 100%, most likely dueto a slight error in the loading measurementalthough experiments were repeated with care.

It is well known that for diffusion of a substancefrom a spherical particle, plots of the amountreleased versus the square root of time are linearat early times (cumulative fraction released<0.6).Dextran and BSA are most likely released byrestricted diffusion through water-filled poresconnected to the particle surface, which can bedescribed by Fickian diffusion with an effectivediffusivity, leading to an analogous linear plot ofrelease versus t0.43. Such plots of our data suggestthat the slow phases of F-Dex and R-BSA releaseare controlled bydiffusion for all spheres sizes, anda conventional ‘‘initial burst’’—defined as a rapidrelease, faster than that accounted for by simplediffusion, in the first 1–2 days—was not observed(Fig. 4). Linear least squares fits to the lag phasesof each plot were quite good (see r2 values for eachfit in Tab. 2). In addition, fits of the initial releasedata to the well-known analytical solution fordiffusion from amicrosphere42 provided estimatesof the effective drug diffusivity in the slow phase(Tab. 2).

The release versus t0.43. plots also provide ameans to more quantitatively determine theduration of the slow phase. We define the end ofthis phase to occur at the last time point that lieswithin the linear fit to the initial release (Fig. 4).Thus, the duration of the F-Dex slow phase was243, 68, and 19 days for 31-, 44-, and 80-mmdiameter microspheres, respectively. For R-BSArelease, the slow release phase lasted 47, 40, and15 days for 34-, 47-, and 85-mm microspheres,respectively (Tab. 2).

Microsphere Morphology and Intraparticle DrugDistribution During Degradation and Drug Release

To investigate the mechanisms responsible for theinteresting shapes of the macromolecule releaseprofiles, we observed each of the formulationsimmediately after fabrication and at 19, 40, 75,and 152 days during the in vitro release experi-ment using light, confocal fluorescence, andscanning electron microscopies (Figs. 5 and 6).Further, the fluorescence distributions were

Figure 1. A: Size distributions of F-Dex- and R-BSA-loaded PLG microspheres. B: Confocal fluorescencemicrographs showing intraparticle distribution of F-Dex and R-BSA within PLG microspheres of threedifferent sizes.



Figure 3. In vitro release of (A and C) F-Dex and (B and D) R-BSA from uniform PLGmicrospheres of different sizes. InpanelsAandC,microspheres are 31mm(closed circles),44 mm (open circles), and 80 mm (triangles) in diameter. In panels B and D, microspheresare 34 mm (closed circles), 47 mm (open circles), and 85 mm (triangles) in diameter.

Figure 2. Scanning electron micrographs of the surfaces and interiors of cross-sectioned PLG microspheres loaded with (A) F-Dex and (B) R-BSA.



quantified by averaging the fluorescence intensityat each radial position using a simple imageanalysis algorithm (Fig. 7).

After 19 days of incubation, fluorescence micro-graphs and radial distributions show that both

F-Dex and R-BSA concentrations decrease nearthe particle surface. The internal morphologyof F-Dex-loaded microspheres changed littlebetween 0 and 19 days. The initially porousR-BSA-containing microspheres, however, showan increase in both the number and size of pores by19 days.

Subsequently, all six formulations underwentsimilar changes in morphology and drug distri-bution, but importantly the rate of changeincreases with microsphere diameter. In general,as degradation and release progressed, the inter-ior of themicrospheres growsmore porous, and thedrug distribution in the microspheres continuedto recede from the surface (e.g., Figs. 5A and 6A,40 days). Next, we observed times at whichthe fluorescence was redistributed toward theparticle surface and the core of the microspheresappears relatively dark (e.g., Fig. 5A, 75 days; 5B,40days; 6A, 75days; and6B40days). [Adetectablelevel of fluorescence was still observed in the coresof 31-mm F-Dex-containing particles at 152 days(Fig. 5A) and in 44-mmR-BSA-containing particlesat 75 days (Fig. 6B).] SEMsat the same time pointsshow that the microspheres became hollow. Asdegradation and release progressed, the hollowcore grew in size and the layer of fluorescence onthe periphery became thinner. The polymer shellbecame quite porous in all cases, and the micro-particle diameter increased by as much as 2.5-fold(see lightmicrographs inFigs. 5 and 6at 152days).Eventually the polymer shell became so thin thatwhen sectioned and dried the particles collapsedentirely (SEMs inFig. 5C, 40and152days;Fig. 6C,75 and 152 days). All of these changes can beobserved semi-quantitatively in the radial fluor-escence distributions (Fig. 7).

It is interesting to note that the transition fromthe slow to the faster release phase (Fig. 3) appearsto correspond with changes in the particle mor-phology. In each case, the transition occurredshortly after the fluorescence darkened in the

Figure 4. Initial in vitro release of F-Dex (A) andR-BSA (B) plotted versus t0.43. Solid lines representlinear least square fits to the experimental data.Correlation coefficients for the fits are shown in Table 2.

Table 2. Parameters Resulting from Linear Least Squares Fit of Initial Cumulative Release According to FickianDiffusion (t0.43) and the Effective Diffusivity Calculated by Fitting the Original Release Data

AverageDiameter (mm)

r2 for Linear Fit to Lag Phasein Terms of t0.43

Duration of Lag(Days)

Deff Lag Phase(cm2/s)

F-Dex 31 0.992 243 3.3� 10�16

44 0.992 68 3.3� 10�16

80 0.982 19 5.6� 10�16

R-BSA 34 0.997 47 4.2� 10�15

47 0.998 40 6.9� 10�15

85 0.990 15 5.6� 10�15



particle interior and the core appeared hollow. Atthe next time point after the transition at whichthemorphologywas investigated, the fluorescenceappeared predominantly at the particle peripheryand the microspheres appeared as hollow shells.

DISCUSSION

PPF technology has been used to producemonodisperse polymer microspheres encapsulat-ing therapeutic agents.39,41,43 Previously, PPF-

fabricated particles were utilized to better controland understand the release profiles of smallmolecule drugs by comparing a range of particlesizes encapsulating drugs of various solubility inwater.41,44,45 For all formulations studied to date,an initial burst of drug was absent, although forsmall microspheres encapsulating a model drugwith high water solubility (>8 mg/mL), rapiddiffusion and complete release after only a fewdays were often noted. These uniform particleformulations revealed that for small-molecules,�10–30 mm diameter microspheres would tend to

Figure 5. Morphologies of F-Dex-containing micro-spheres during in vitro degradation and release. (A) 31mm; (B) 44mm;and (C) 80mmdiameter. Ineachpanel, thethree rows of images are light micrographs (top), laserscanning confocal micrographs (middle), and scanningelectron micrographs (bottom).

Figure 6. Morphologies of R-BSA-containing micro-spheres during in vitro degradation and release. (A) 31mm; (B) 44mm;and (C) 80mmdiameter. In eachpanel, thethree rows of images are light micrographs (top), laserscanning confocal micrographs (middle), and scanningelectron micrographs (bottom).



release in a concave downward, simple diffusion-controlled manner, but as microsphere diameterincreased to �50–100 mm, the release profilesteadily shifted toward a sigmoidal shape due tothe effect of polymer degradation.41,46 Overall, therelease of small molecules from uniform micro-spheres exhibits an increased rate and decreasedduration of release as microsphere size decreases.A similar trend is typically expected for delivery ofmacromolecules from PLG microspheres.16 As

shown in our work with small molecules, theuniformity of the particles may provide theopportunity to control delivery rates and toinvestigate the fundamental mechanisms govern-ing release. Here we have reported that PPFtechnology can also be used to encapsulatemacromolecules in uniform PLG microspheresusing a water-in-oil emulsion.

The very long duration of particle erosion anddrug release is somewhat surprising given the

Figure 7. Radial fluorescence distributionprofiles obtained fromCLSMimages shownin Figures 5 and 6. r/R is the normalized radial position from the core to the surface of themicrospheres. (A) 31-mm F-Dex; (B) 44-mm F-Dex; (C) 80-mm F-Dex; (D) 34-mm R-BSA;(E) 47-mm R-BSA; (F) 85-mm R-BSA.



relatively low initial PLG molecular weight.PLG 50:50 degradation rates reported in theliterature16,17,47–49 are �0.02–0.08 day�1. Basedon images of microspheres obtained at 152 days(Figs. 5 and 6), as well as preliminary degradationstudies, the F-Dex- and R-BSA-loaded PLGmicro-spheres reported here appear to degrade moreslowly (�0.012–0.018 day�1 estimated from asingle sample of each microsphere size). Althoughthe discrepancy is not fully understood, there areseveral possible explanations for our observations.First, the encapsulated Dex and BSA may adsorbto or otherwise interact with the polymer, hinder-ing degradation. Another possible explanation forthe slower degradation is the unique emulsifica-tion mechanism of the PPF process, which couldaffect the polymer matrix, especially near themicrosphere surface, leading to differences inwater penetration, for example. The majority ofthe experimental parameters (e.g., solvents, tem-peratures, polymer concentrations, volumes, etc.)and processes (e.g., solvent extraction, washing,particle drying, etc.) are identical to conventionalmethods, however, and any effect should be due tothe emulsification step only, making this anunlikely hypothesis. The uniformity of the parti-cles may impact polymer degradation, though aprobable mechanism for such an effect is notknown. Microspheres showed minimal clumpingduring incubation, which may have increased thelocal renewal of microsphere surface with freshbuffer, thus, decreasing the local build-up ofdegradation products. A potential mechanismcausing the decreased degradation rate observedhere is the difference in agitation of the samplesduring release studies. Shaking the release sam-ples produces an undisturbed ‘‘pile’’ of micro-spheres at the bottom of the sample vial, which iscontrasted to typical methods of stirring ortumbling samples during dissolution studies.Indeed, researchers at Merck have demonstratedthat rapid stirring released hepatitis B antigenover a few days as compared to three months ofrelease observed by gentle tumbling, a strikingdifference.50

Another possible complication stems frompossible instability of the encapsulated F-Dexand R-BSA in an acidic microclimate, such asthemicrosphere interior,9,10 over prolonged times.SDS–PAGE of R-BSA released after 50 and200 days verified that the protein partiallyaggregates but is not hydrolyzed or otherwisedegraded (not shown). The fluorescent dyes,furthermore, remained covalently attached to

BSA and Dex over the entire course of the study,as no low-molecular weight (<3 kDa) fluorescentspecies were detected in the supernatants (notshown).

Given the long duration of the release ex-periments reported here and the possible insta-bility of the F-Dex and R-BSA after prolongedincubation in degrading microspheres, it is never-theless prudent to use caution in interpretation ofthe release data, especially at later time points.Release data after 100–150 days, times at whichwe have evidence that the F-Dex and R-BSA areintact and at which microspheres were imaged,were not relied on for the following discussion northe conclusions.

The overall shape of the macromolecule releaseprofiles from the uniform microspheres is compar-able to those reported for other PLG microsphereformulations,16,51,52 except a single slow phasereplaced the commonly observed burst and lag.Efforts have been made to control the duration ofthe conventional lag phase and the onset of steadymacromolecule release, for example, by controllingpolymer molecular weight or composition.53,54 Inthis work, the duration of the slow phase wasaltered by controlling the microsphere size withthe largest (80 and 85 mm) microspheres yieldingthe shortest slow-release phases of 19 and 15 daysfor F-Dex- and R-BSA-loaded microspheres,respectively.

In the formulations reported here, model com-pounds F-Dex and R-BSA were initially distrib-uted homogeneously in the PLG microspheres.Both macromolecules are insoluble in the organicphase and are expected to remain in the aqueousdroplets of the primary w/o emulsion. Fluorescentlabeling of these molecules may perturb thesolubility of native dextran or BSA to some extent,but dyes employed also exhibited high watersolubility and labeling was minimal (<1 dye permolecule). Although the fabrication conditionswere identical for the two molecules, the R-BSA-containing microspheres appeared significantlymore porous (Fig. 3). These results suggest that afiner or more stable w/o emulsion was obtainedwith the F-Dex formulation.

An initial ‘‘burst’’ of water soluble small mole-cules andmacromolecules fromPLGmicrospheresis commonly observed, in which a significantfraction of the drug is released within the first24 h.52,55,56 As reviewed in the introduction,multiple solutions have been suggested to elim-inate drug burst, but often they only work inspecial cases, necessitating reformulation for each



drug/polymer combination, or require additionalsteps such as particle coating. The burst hasoften been ascribed to desorption of drug thatmay have localized to the particle surface oradsorbed during the encapsulation process or torapid diffusion of the drug through initiallyopen pores, which quickly close as the polymergains mobility and ‘‘heals.’’7,16,25 However, wedid not observe a conventional burst phase forF-Dex or R-BSA release from monodisperse PLGmicrospheres; rather our data show a slow,diffusive release at the earliest time points.Similarly, and somewhat surprisingly, we havenot observed initial bursts for small moleculedrugs from uniform polymer microspheres,either.41,43,45

Fabrication of the microspheres reported herewas analogous to the conventional double-emul-sion/solvent extraction process.52,57 (The primarydifference, of course, is that the PPF methodgenerates a monodisperse secondary emulsion.)Thus, one might expect a similar surface localiza-tion of macromolecules in our particles, but thiswas not observed by confocal fluorescence micro-scopy (Fig. 2). It is possible that the relativelygentle process that forms the droplets, in com-parison to the strong shearing used in conven-tional processes, does not allow access to thesurface for a large proportion of the encapsulatedcompounds. An alternative explanation is that theabsence of small particles with a large surfacearea:volume ratio, which would allow rapid fluxfrom pores or channels near the surface, may beresponsible for the lack of an initial rapid release ofthe macromolecules.58,59

The first phase of F-Dex and R-BSA releaseprofiles is consistent with slow diffusion asopposed to an initial ‘‘burst.’’ Plots of the amountreleased versus t0.43 were were linear (Fig. 4).Also, the initial release rate increased withdecreasing microsphere diameter as expecteddue to increasing surface area:volume ratio.Fluorescence micrographs showed that early inthe release the drug concentration decreaseddramatically near the microsphere surface withless change in the interior (Fig. 7). These shapescompare favorably to theoretical concentrationprofiles for diffusion of a solute from a sphere.However, the drug concentration profiles are notidentical to theoretical predictions. The deviationcould possibly be due to imperfect initial distribu-tion or diffusion preferentially through aqueous-filled pores initially connected to the surface. Thisis supported by faster initial release of R-BSA

compared to F-Dex because R-BSA-containingmicrospheres are initially more porous.

In all cases, the slow release phase transitionedinto a faster phase and the time of the transitiondecreased with increasing microsphere diameter.Because the microparticle morphologies anddrug distributions were observed infrequently, itis difficult to correlate the transition directlyto specific structural changes. In most cases,the slow-release phase ended sometime after themicrospheres swelled significantly and thecore became hollow. However, in the 31-mm F-Dex-loaded microspheres, the transition betweenrelease phases did not occur until well after152 days atwhich time themicrospheres appearedas a relatively thin shell and had swelled to 1.4times their initial diameter (Fig. 5A). Also, thetransition in R-BSA release from 85-mm particlesoccurred at 15 days while the particle interiorappeared relatively dense and no swelling wasobserved even at 19 days. In general, however, thetransition appears to some extent to be caused bythe swelling of the particles and the generation of athin, porous, polymer shell with a hollow core.

F-Dex release profiles were very distinctivewhile R-BSA release varied much less over asimilar range of microsphere diameters. In addi-tion, the F-Dex release was more prolonged thanR-BSA release for comparable microsphere sizes.These differences may be due to the porosity of theR-BSA- compared to the F-Dex-containing micro-spheres since the macromolecules are believed torelease via diffusion through water-filled porescreated by polymer degradation/erosion. Theincreased initial porosity of the R-BSA-containingparticles is expected to more rapidly generateinterconnected pores that provide a route ofescape for drug in the interior of themicrospheres.The difference in macromolecule conformationmay be another factor contributing to the differ-ence between the release profiles. F-Dex is a linearmolecule whereas R-BSA is a globular protein.Although they are similar in molecular weight,R-BSA is expected to be smaller due to its foldedconformation. Thus, one may expect R-BSA todiffuse more easily through smaller and possiblymore tortuous pores. F-Dex, in contrast, mustrepeat through the same pores, which is a slowerprocess. Thus, the critical pore size, where anincrease in release rate is observed, is reachedsooner for R-BSA compared to F-Dex.

In all formulations, a point is reached at whichthe core of the microspheres appears dark inthe fluorescence micrographs. The darkening



correlates strongly with the appearance of hollowcores in SEMs. However, the lack of fluorescencecould be due to the absence of the macromoleculesfrom the core or quenching of or damage to thefluorophores. Fluorescein is known to be pH-dependent with pKa� 6.4 between the neutraland monoionic forms. The quantum yield of themonoion of fluorescein is 0.37 compared to 0.93 forthe neutral species. Thus, fluorescein emissiondecreases with pH; emission at pH 5 is approxi-mately half of that at pH 7. If the interior ofmicrospheres becomes acidic due to accumulationof polymer degradation products (vide infra), theF-Dex signal would be strongly decreased. On theother hand, sulforhodamine B fluorescence isindependent of pH at least in the range 3–7 (datanot shown), yet R-BSA microspheres show thesame dark cores. In this case, the lack of fluores-cence in the core is not expected to be the result ofan acidic microenvironment inside the micro-spheres. The similarity between the F-Dex andR-BSA fluorescence micrographs suggests that inboth cases the macromolecules are excluded fromthe cores as the microspheres become hollow. Thecause of this effect is unknownat this timebutmaybe attributable to autocatalyzed formation ofsoluble oligomers, �1 kDa, of PLGA that candiffuse out of the particle.

The increase in degradation and erosion rateswith increasing microsphere size supports auto-catalytic degradation in these uniform micro-spheres. Because autocatalysis results fromlimited diffusion of buffer and degradation pro-ducts through the microspheres, the size of thedevice will impact the steepness of the pHgradient. Larger microspheres, with a longerdiffusion length from the core to the surface,should exhibit a lower pH and faster degradationand erosion.9 The polymer degradation studiesshowed that degradation rate increased withmicrosphere diameter. In addition, the time pointsat which the microsphere cores appeared hollowconsistently occurred earlier as microsphere dia-meter increased.

In summary, we propose the following model todescribe release of macromolecules from theseuniform PLG microspheres. Initially, drug islocated in relatively large, drug-filled ‘‘pores’’resulting from the primary emulsion. At earlytimes, drug encapsulated in interconnected poresat the microsphere surface is released by simplediffusion. The initial rate of this phase is inverselyproportional to the microsphere diameter due todifferences in the surface area:volume ratio.

During this time, polymer degrades hydrolyticallyand somemass loss occurs, increasing the pore sizeand potentially the connectivity. Once a criticalpore size and/or interconnectivity is reached, drugencapsulated deeper in the particles begins torelease by diffusion through the pores. However,because of limited diffusion of soluble degradationproducts inearly stages, autocatalytic degradationoccurs. In fact, a sufficiently acidic environmentcan be created in the interior of the particle tocause essentially complete degradation of polymerin the core. As molecular weight decreases,significant particle swelling can occur, alsoincreasing the size of the aqueous-filled pores.This autocatalytic effect is more pronounced asmicrosphere size increases. Thus, the critical stageis reached more quickly as microsphere diameterincreases. The data presented herein are consis-tent with this model. Additional experiments, forexample, measuring the pH-dependence of PLGdegradation and intraparticle pH as a function ofparticle diameter, will be needed to further testthishypothesis. Furthermore, to extend thismodelto other systems one must consider other factorssuch as the particle fabrication process, encapsu-lation method and presence of excipients, asnecessary, which can profoundly affect the releasemechanism.

This model is not dissimilar to those that haveappeared in the literature.16,19,36,60,61 In fact,Park17 described a similar mechanism for drugrelease from PLG microspheres including theprediction of a completely degraded core sur-rounded by a slowly degrading surface layer, andViswanathan et al.19 reported hollowing ofpoly(D,L-lactide) microspheres. However, to ourknowledge the hollow cores reported here have notbeen observed experimentally for microspheressmaller than 100-mm, nor has the increase inhollowing rate and subsequent swelling withincreasing microparticle size.

CONCLUSIONS

The development of PPF technology has allowedthe production of uniform, double-emulsionmicrospheres capable of efficiently encapsulatingmodel macromolecules. Confocal fluorescencemicroscopy of the fluorescently tagged macromo-lecules revealed an even distribution of drugthroughout the microsphere matrix regardless ofparticle size. An interesting observation was thelack of an initial drug burst and the ability of the



monodisperse microspheres to modulate the onsetof steady drug release with large, �80–85 mmspheres exhibiting a much reduced lag ascompared to smaller, �30–35 mm spheres. Inaddition, the overall rate of release was found toincrease with increasing microsphere size, whichat first glance may be a counterintuitive result.The interesting shapes of the release rate profilesare thought to be the result of autocatalyticdegradation and erosion of the PLG, which resultsin hollow particles, exclusion of drug from theparticle core, and particle swelling. Each of theseevents occur at critical times dependent uponmicrosphere size and all contribute to anincreased rate of macromolecule diffusion frommicrospheres.

ACKNOWLEDGMENTS

This work was supported in part by NIH GrantEB002878. Scanning electron micrographs wereobtained at the Center for Microanalysis ofMaterials, University of Illinois, which is partiallysupported by the U.S. Department of Energyunder grant DEFG02-91-ER45439.

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