A C S

S Y M P O S I U M

S E R I E S

Formulation and Deliveryof Proteins and Peptides

Jeffrey L. Cleland, EDITOR
Genentech, Inc.

Robert Langer, EDITOR
Massachusetts Institute of Technology

Developed from a symposium sponsoredby the Division of Biochemical Technology
at the 205th National Meetingof the American Chemical Society,Denver, Colorado,
March 2 8 - A p r i l 2,1993

A m e r i c a n C h e m i c a l S o c i e t y , W a s h i n g t o n , D C 1994

567

Chapter 15

Drug Delivery from Bioerodible Polymers
Systemic and Intravenous Administration
1 2

1 3

1

Achim Gopferich ' , Ruxandra Gref ' , Yoshiharu Minamitake ,Lisa Shieh , Maria Jose Alonso , Yashuhiko Tabata , andRobert Langer
1

14

15

1

department of Chemical Engineering, Massachusetts Instituteof Technology, Building E25, Room 342, Cambridge, MA 02139
Lehrstuhl fur Pharmazeutische Technologie, Universitat
Erlangen-Niirnberg, Cauerstrasse 4, 91058 Erlangen, Germany
Laboratoire de Chimie Physique Macromoleculaire, Ecole Nationale
Superieure des Industries Chimiques, 1 rue Grandville, 54001 Nancy
Cedex, France
Laboratorio de Farmacia Galenica, Facultad de Farmacia, Universidade
de Santiago de Compostella, 15706 Santiago de Compostella, Spain
Research Center for Biomedical Engineering, Kyoto University, 53
Kawahara-cho, Shogoin, Sakyo-ku, Kyoto 606, Japan
2

3

4

5

The recent progress i n understanding the erosion of
biodegradable polymers and the manufacturing of controlled
release devices for proteins and peptides is reported. The
erosion mechanism of poly(anhydrides) was investigated as
an example of biodegradable polymers and the erosion behavior is modeled mathematically. The results provide useful
information on the microstructure and chemical environment
inside these polymers during erosion. It is shown how theymight affect the stability and the release of proteins.
Concomitantly proteins were processed in controlled releasedevices. Special attention was paid to the stability of thebiomolecules during the manufacturing of dosage forms.
Furthermore, the development of a new type of biodegradable
nanosphere as a future dosage form is shown.
Progress in the field of biology and biochemistry has led to the discoveryof numerous bioactive peptides and proteins in the last few decades. Manyof them, like insulin, as a classical example, are very useful for medical
therapy. The rapid development of biotechnology and progress in peptide
and protein chemistry allowed the mass production of many compounds
and made their broad introduction into medical therapy possible(7). The
0097-6156/94/0567-0242$10.88/0© 1994 American Chemical Societyuse of such biomolecules poses, however, severe problems. Some of thesecompounds have very short half lives in body fluids (under a minute, insome cases(2)) and due to degradation in the gastrointestinal tract the majority of them cannot be administered orally. These limitations motivatedthe development of controlled release dosage forms that improve protein
and peptide stability as well as prolong drug activity after application.
Processing these substances into dosage forms is not always easily
achieved. Many of them have limited chemical and physical stability.
Common instabilities are irreversible aggregation(J), oxidation(4) orconformational changes(5) all of which may affect activity. The preparation of controlled release devices for such proteins and peptides has, therefore, become one of the major challenges in the field of controlled release.
Controlled release devices have been prepared for various routes ofadministration such as oral, parenteral, nasal, rectal, buccal, vaginal andtransdermal(6-7). They all control the release of peptides and proteins by a
few basic principles. Classical systems control release by diffusion
through a polymer matrix, in which the drug might be dissolved or suspended^). Closely related to diffusion controlled release systems arethose in which swelling controlled release occurs(9). The swelling of thepolymer, usually an ionic network, increases the permeability of the matrix and allows drugs to be released by diffusion. Parenterally applied
dosage forms are the most often used at present as many proteins and peptides are unstable in the gastrointestinal tract resulting in poor bioavailability. The introduction of degradable drug carrier materials brought new
progress into this field of controlled release research. Biodegradable
polymers as drug carriers control the release of the drugs through diffusionand erosion and have the advantage of dissolving after the application.
Instead of simple diffusion-controlled release, drug is release by an erosion-controlled mechanism, which provides the advantage of decreasing
the release rate of the proteins and peptides. The search for new
biodegradable polymers for controlled release stimulated research in several areas, such as polymer synthesis, polymer processing and the formulation of devices. In the area of polymer synthesis, there have been major
efforts to 'design' polymers for the purpose of controlled release.
Appropriate examples are poly(ortho esters)(70) and poly (anhydrides)(77). In providing the biodegradable polymer raw material for controlled
release devices for peptides and proteins two fundamental problems arose:
1. In order to efficiently design a controlled release device from biodegradable polymers, the polymer properties and its erosion mechanism should be known.
2. The technology for manufacturing devices ranging from the centimeterto the nanometer scale must be developed. The requirements of stability for such sensitive compounds as peptides and proteins must,
thereby, be met.
The successful design of release systems will heavily depend on
detailed knowledge in both areas. Unfortunately, a unique theory does notexist for either set of issues. There is not, for example, a basic theory that
could predict the erosion of all biodegradable polymers or the stability ofproteins and peptides from their structural formulas alone. Due to the expanded research in these areas, however, there is progress towards a
broader understanding of polymeric properties and the development of
stable protein formulations compatible manufacturing technology. This
article intends to show some of the progress that has been made in understanding the release of peptides and proteins from polymers. The focus
will be on two major areas:
1. Understanding the erosion mechanism of biodegradable polymers.
2. Providing some examples to illustrate recent advances in the development of controlled release systems. We cover the release of proteins frompolymers as well as the development of future dosage forms.
The Erosion of Biodegradable Polymers
Investigating erosion mechanisms of biodegradable polymers might lookrather academic and not that beneficial at first glance. The
microenvironment of a biodegradable polymer during erosion, however,provides the medium in which peptides and proteins are embedded andfrom which they have to be released. Factors like pH or the amount ofdissolved monomers not only influence release rates, but also affect thestability of proteins and peptides substantially(7 2) and thus become very
important parameters for the stability of a controlled release dosage form.
The design of manufacturing systems for the controlled release of proteinsand peptides from polymers requires, therefore, a fundamental knowledge
the erosion mechanism of a biodegradable polymer.
Diffusion controlled protein release from non-degradable polymers
is well-understood(73). The release of such compounds from biodegradable polymers in contrast depends heavily on the erosion of the polymer, acomplex process whose mechanism is not entirely understood. The erosion of biodegradable polymers follows mechanisms specific for a certain
type of polymer and is influenced by a number of parameters. Most
important is the chemical degradation of bonds in the polymer chain. The
rate at which the bonds are cleaved depends on the type of bonds between
the monomers(74), the diffusivity of water in the polymer(75), and thepolymer crystallinity(76). The pH of the degradation medium exhibits acatalytic effect on the hydrolysis of bonds(77), influences the solubility of
degradation products, and finally regulates their release. Degradation
products such as Monomers and oligomers carry functional groups thatchange p H inside cracks and pores in the eroded polymer and these groups
can have feedback on the erosion process(7S). It can be concluded that thepolymer erosion is a composite process that can be very complex and isspecific for each polymer. The erosion mechanisms of poly(lactic-co glycolic acid) and poly(ortho esters), for example, follow completely different mechanisms(79-20).

Investigation of Erosion Mechanisms. We were interested in the erosion
mechanism of a class of related poly(anhydrides) consisting of sebacic
acid (SA), l,3-bis(p-carboxyphenoxy)propane (CPP) and a fatty aciddimer ( F A D ) monomers which are shown in Figure 1. The techniques thatwere used for these studies may be applied to any class of polymer and theresults that we obtained provide a clear outline for the important fundamental characteristics of polymer erosion.
The Structure of Polymers. Prior to the investigation of erodedpolymer discs, the polymers were investigated for their internal structure.
Figure 2 shows the appearance of various polymers that were taken fromthin melted polymer films under polarized light. The Maltese Crosses indicate that these polymers are partially crystalline and consist of
spherulites. The crystallinity of the polymers depends on the copolymer
composition. In the case of poly(l,3,-bis(p-carboxyphenoxy)propane-cosebacic acid) abbreviated p(CPP-SA) changing the content of either
monomer to a 50:50 composition lowers the crystallinity from more than
60% to almost 10% (27). Concomitantly, the glass transition temperaturedrops substantially. p(FAD) homopolymer is amorphous(22), so that the
crystallinity of p(FAD-SA) copolymers depends only on the S A monomer
content. A s shown in Figure 3, increasing the content of F A D monomer
above 80% results in amorphous polymers. The effect of microstructure
and crystallinity on erosion is observed in investigating eroded polymer
discs that were produced from the polymer by melt molding(77). Figure 4
a shows a scanning electron photograph of a cross section through a discof p(CPP-SA) 20:80 that has been eroded in phosphate buffer p H 7.4 at
37°C for 3 days. Eroded and non-eroded polymer sharply separated by theerosion front are readily distinguishable. Observation of this front withtime confirms that the erosion of these polymers starts on the surface andmoves towards the center of the discs which has been referred to as surface erosion(23). Figure 5 shows the kinetics of front movement in p(CPPSA) 20:80 and p(CPP-SA) 50:50 as measured from S E M pictures. The
way erosion affects the polymer can be seen from S E M pictures showing
the eroded parts of the polymer in more detail. A s shown in Figure 4b theamorphous parts of the spherulites have been eroded, while the large partsof the crystalline skeleton remain in place(24). A s erosion progresses, ahighly porous erosion zone is created. Once the erosion front has reacheda certain spot in the matrix, the release of drug begins from that point. The
release of compounds is then determined by the diffusion through poresand cracks.
pH Changes during Erosion. When these polymers erode, they
release oligomers and monomers that carry carboxylic groups (25). We
suspected that the p H in the vicinity of the polymer surface or inside thecracks and pores of the polymer is not determined by the pH of the buffer
medium, but instead by the released monomers. To support this hypothe-HOOC-(CH ) -COOH
2

8

sebacic acid (SA)H O O C — O — (CH )3— O — { ^ ) ~ - COOH
2

1,3-bis-(p-carboxyphenoxy)propane (CPP)

CH -(CH )32(CH2) -COOH

7

8

CH

/
HOOC-(CH )2\
7(CH2) -CH
8

3

fatty acid dimer (FAD)

Figure 1. Monomers contained in the investigated poly (anhydrides).

Figure 2. Polarized light microscopic pictures of poly(anhydride)films: a) p S A (250x), b) p-(FAD-SA) 20:80 (400x), c) p - ( F A D - S A )50:50 (400x).

20

40

60

80

100

FAD content (%w/w)

Figure 3. Crystallinity of p(FAD-SA) poly(anhydrides) depending on
F A D content.

Figure 4. S E M pictures of cross sections through eroded p(CPP-SA)
20:80 dies: a) erosion front separating eroded(right) from non-eroded(left) polymer, b) eroded polymer.

sis, we measured the pH on the surface of p(CPP-SA) 20:80 during erosion using confocal microscopy(26). Figure 6 shows the value for the p H
as a function of the distance from the surface of the polymer matrix disc.
The p H clearly drops when approaching the surface of the polymer. Thiseffect is due to the high concentration of released monomers upon degradation. Keeping in mind the highly porous structure of the eroded zone ofthe polymer it can be hypothesized that the p H inside this network is determined by the monomers and is even lower than the p H at the surface(26). This issue is very important with respect to the stability of pH
sensitive drugs such as some proteins or peptides.
Changes in Crystallinity. From the structural changes during erosion, changes in polymer crystallinity could be suspected. They were investigated by differential scanning calorimetry and x-ray diffraction. In
the case of the p-SA homopolymer, it was found that the crystalline regions are more resistant towards degradation than the amorphous regions.
These results agree with the S E M pictures obtained from the degraded partof polymer discs. The monomers S A and CPP were found to crystallize torelatively high extents in all polymers qualitatively shown by wide-angle
x-ray diffraction and quantitatively by DSC. Figure 7 shows the calculated
amount of crystallized monomers in p(CPP-SA) 20:80, as determined by
D S C according to (26). The presence of crystallized material proves indirectly that the pores of eroded polymer are filled with a saturated solutionof monomers. F A D monomer is liquid at room temperature and therefore
not able to crystallize upon release. Due to its low solubility in aqueous
buffer solution, however, it sticks to the eroding discs as a film.
The Release of Monomers during Erosion. The release of acidic
monomers indicates the importance of p H during erosion. A s an example,
the release profile of S A and C P P monomer during the erosion of p(CPPSA) 20:80 is shown in Figure 8. The release profile of S A is similar to therelease of substances from monolithic devices. The release profile of C P P
is discontinuous as the release rate increases after S A has completely leftthe eroded polymer matrix. This spontaneous increase can be explained by
the higher solubility of SA compared to CPP. The dissolved S A , therefore,
controls the p H inside the pores of the eroded zone limiting, thereby, thesolubility of CPP. After S A has been completely released, the p H inside
the pores starts to rise, which increases the solubility of CPP(26). This
process finally increases the release rate of C P P from the eroded polymer
matrix.
Modeling Polymer Erosion. The modeling of polymer erosion is the firststep towards the theoretical design of controlled release systems with an apriori adjusted release rate. The lack of knowledge about erosion, however, hindered the development of models substantially. The first attempts

time [days]

Figure 5. Relative position of the erosion front in p(CPP-SA) 20:80
depending on time.

7.00

-200

-150

-100

-50

0

distance from surface |pm]

Figure 6. pH in the degradation medium as a function of distance tothe surface of a p(CPP-SA) 20:80 polymer disc after 2 days of erosion
in daily changed 0.1 M phosphate buffer pH 7.4 at 37°C.
(Reproduced with permission from ref. 26. Copyright 1993 Wiley &Sons.)

50

time [days]

Figure 7. Content of crystallized monomers in a p(CPP-SA) 20:80
matrix during erosion: o CPP, • SA.
(Reproduced with permission from ref. 26. Copyright 1993 Wiley &Sons.)

120

21

time [days]

Figure 8. Release of monomers during the erosion of a p(CPP-SA)
20:80 polymer matrix disc: o CPP, • SA.
(Reproduced with permission from ref. 26. Copyright 1993 Wiley &Sons.):

to describe the erosion of biodegradable polymers stem from the description of the dissolution of erodible tablets(27). It was assumed that erosion
could be described by surface dissolution kinetics. Later, diffusion processes were taken into account and modeled as a pseudo-steady state process, where erosion was modeled assuming first order kinetics or a constantly moving erosion front(2S-29). Introducing non-steady state analysis,
erosion was described as a moving front problem with constant velocity(JO). A l l these models describe erosion in one dimension and do notallow descriptions of the microstructural changes upon erosion. Inspiredby the two-dimensional description of the dissolution of matrix tablets(3732), a two-dimensional model able to describe the structural changes of
our investigated polymers upon erosion was developed. Rather than taking
individual factors of polymer erosion into account, we combined them byregarding the erosion of small parts of polymer matrix as a random
event(JJ).
The Basic Concept. A polymer matrix was represented by a twodimensional computational grid, in which each grid point represents either
an amorphous or crystalline part of the polymer matrix. The erosion ofsuch a 'polymer pixel' was assumed to depend on two features: the contactof the polymer with the degradation medium, and the crystalline or amorphous nature of a matrix part which is represented by a pixel. Polymers
not in contact with water will not erode. Pixels on the surface of the polymer matrix or next to an eroded neighbor have contact to water. The erosion of crystalline matrix parts occurs at a slower rate than amorphousparts, a characteristic which was observed in the structural studies(26).
Besides these fundamental assumptions, a general algorithm must be defined to develop a working model: the matrix must be represented by acomputational grid divided into individual pixels. A distribution must be
chosen for crystallites and amorphous parts. After contact with an eroded
neighbor, each pixel is assigned an individual life expectancy after which
it is regarded eroded. The life expectancies are distributed according to afirst order Erlang distribution 34) from which they are chosen randomly.
To account for the crystalline or amorphous nature, different constants arechosen for use in the Erlang distribution.
The Representation of a Polymer Matrix. To represent a polymer
matrix prior to erosion, it is sufficient to represent only a cutout of thematrix, if the erosion problem is symmetrical. Figure 9 shows such acutout for a cross section through a cylindrical matrix.
To account for the crystallinity, some of the pixels have to be designatedto represent crystalline polymer matrix parts. In the easiest case, a random
distribution of them can be assumed:

1-Xc(xij)= 1 X0for:l^i^n ,

Xij=0

xy=l
all other valuesx(1)

X is the crystallinity of the polymer. The status Xjj (1 for crystalline and 0
for amorphous) of all pixels is assessed from consecutive Bernoulli tria l s ^ ) . c(Xij) is the probability that a pixel at location x=i and y=j on thegrid represents an amorphous or crystalline part of the matrix. Figure 10ashows the theoretical representation of a polymer matrix prior to erosion.
Dark pixels represent crystalline, whereas white pixels represent amorphous parts of the polymer.
The Simulation of Erosion. After having set up the grid, the erosion algorithm can be applied. A first order Erlang distribution34) wasused for the calculation of the life expectancy of a pixel after its contact
with water
e(t)=

(2)

t3

X*e- ^

e(t) is the probability that a pixel degrades after a time t. Random variable
t is the time that elapsed between the first contact with water and the erosion of the pixels. X can be considered as a rate constant and is different
for amorphous and crystalline pixels. To achieve results independent fromthe grid size, the grid size n has to be taken into account in equation 2:
1

e (t)= X+n+e- *^*
n

11

(3)

Using Monte Carlo sampling techniques( 36), the life expectancy of an individual pixel can now be calculated from equation 3. Erosion begins at
the pixels that represent the surface of the polymer matrix. Their lifetime
is calculated and the pixel with the shortest lifetime is determined. Erosion
proceeds with the removal of eroded pixels from the grid, now considered
to represent aqueous pores. They then initiate the erosion process of noneroded neighbors, the life expectancies of which are calculated(ii). The
next pixel is eroded, and so on. A s an example, the erosion of the gridshown in Figure 10a can be seen in Figure 10b. In contrast to Figure 10adark pixels represent non-eroded polymer and white pixels eroded polymer. The appearance of the cross sections of eroded discs shown in Figure
4a agrees with the appearance of the lattice during erosion. In both cases,
the erosion front is clearly visible.

polymermatrixcenter of matrix

computationalgridsurface of matrix
i=l

i=n

x-axisFigure 9. Representation of a polymer disc by a computational grid.
(Reproduced with permission from ref. 33. Copyright 1993 A C S . )Figure 10a. Theoretical representation of a polymer matrix: prior toerosion (dark pixels = crystalline areas, white pixels = amorphous areas).
(Reproduced with permission from ref. 33. Copyright 1993 ACS.)

t* Xa = 0.042

t* Xa = 0.074

t*Xa = 0.339

t**a = 0.396

Figure 10b. Theoretical representation of a polymer matrix: changes during
erosion (dark pixels = non-degraded areas, white pixels = degraded areas).
(Reproduced with permission from ref. 33. Copyright 1993 ACS.)The Quantitative Evaluation of Experimental Results. The unknown parameters in this model are the erosion rate constants for amorphous and crystalline pixels. Once these parameters are known the model
becomes a tool to estimate the erosion behavior of polymer matrices andto predict when the last portion of drug will be released. The computational grid can, thereby, be fit to any desired shape.
In order to determine the erosion rate constants, system parametersmust be defined to develop a model to fit the experimental data. Therefore
two functions were chosen:if(o = -

n(4)

2 fid)
i=0

i n n

(5)

i=lj=lEquation 4 defines the position of the erosion front f(t) at time t as an average value of the position fj(t) of the foremost eroded pixel in each column of the grid. Experimentally, the position of the erosion front was determined by S E M . Equation 5 calculates the remaining mass of the matrix
m(t) at time t, s(xij) being equal to 1 for non-eroded pixels and 0 for
eroded pixels. Experimentally, the remaining mass was determined byweighing dried eroded polymer matrix discs. Using equations 4 and 5, themodel was fit to experimental data of p(CPP-SA) 20:80 and p(CPP-SA)
50:50, minimizing the squared distances between experimental points andsimulated data calculated from equation 4 and 5 (37-38). A s shown in
Figures 1 1 a and b, the movement of the erosion front is almost linear inboth polymers. The larger deviation between experimental and predicteddata for p(CPP-SA) 50:50 (cf. Fig. l i b ) is mainly due to problems in thedetermination of erosion fronts in this polymer by light microscopy. The
loss of mass after an induction period also follows linear kinetics. Fromthe good fit to this parameter it can be assumed that the effect of the error
in the determination of the erosion front movement has only moderate effect on the determination of the rate constants. The erosion rate constants
obtained are shown in Table I.
A s expected from the structural information on eroded polymer matrices,
the erosion rate constants for crystalline pixels are substantially lower

Figure 11. Fit of theoretical functions for mass loss and erosion front
movement to experimental data: a) p(CPP-SA) 20:80, b) p(CPP-SA)
20:80.
(Reproduced with permission from ref. 33. Copyright 1993 A C S . )Table I. Erosion rate constants for crystalline and amor
phous areas in two poly(anhydrides)
X [s-1]p(CPP-SA) 20:80
7.32*10-7
8.75*10-9p(CPP-SA) 50:50
2.7*10-7
3.85*10-11
Source: Reprinted with permission from ref. 33. Copyright1993.
acompared to the rate constants for amorphous pixels. The obtained microconstants allow the simulation of erosion of any kind of polymer matrix
cross section under various boundary conditions. A good example is thesimulation of erosion of devices partially coated with a water impermeable
coating. Such devices could be very useful for the modification of releaseprofiles for slowing down the overall erosion velocity of a biodegradable
device.
The Consequences for the Release of Proteins from Polymers. The erosion of the poly(anhydrides) follows a well defined pattern. A l l polymers
degrade from the surface of the polymer matrix discs towards their center.
A moving front sharply separates eroded from non-eroded polymer. It
could be shown by scanning electron microscopy that crystalline areas ofthe polymers are more resistant to erosion than amorphous areas. Due tothe spherulitic microstructure of the polymers (cf. Figure 2), erosion creates highly porous devices consisting of the crystalline skeleton of eroded
spherulites. Due to the high degradation rates, many oligomers and finallymonomers are released. Reaching the limit of solubility, these substances
start to precipitate out of solution, which was observed by wide angle xray diffraction and differential scanning calorimetry. Substantially highamounts of monomers were found to be in a crystalline state(26).
Confocal fluorescence microscopy showed that the p H on the surface ofthe polymers during erosion was lower than in the buffer medium. Fromthe presence of crystalline monomers inside the erosion zone, it can beconcluded that the aqueous phase inside the polymers is saturated withmonomers. A s S A has the highest solubility of all monomers, this compound determines the pH. From the p K a which is about 4.5 and the solubility which is 0.011M, the theoretical p H minimum is about 3.5(59). Dueto the diffusion of hydroxide ions from the degradation medium into theerosion zone, however, some of the free acids are neutralized, whichbuffers the p H probably around the pKa. Buffer salts like HPO4 " are apparently not able to raise the p H inside the porous zones of the eroded2

discs. The major reason is the long diffusion pathway into the disc along
which they get steadily protonated by dissolved monomers and lose theirpotential to raise p H . The monomer saturated solution inside polymer
pores with a pH of approximately 4.5 is, therefore, the environment that
proteins and peptides will be in contact prior to release out of the matrix.
Once the stability data of such a biomolecule is known, it can readily be
decided whether these types of polymers will make suitable drug carriers.
The well defined erosion behavior of the investigated polymers allows thedevelopment of mathematical models that describe changes in polymer
microstructure during erosion(JJ). Once fit to experimental data, thesemodels are a useful tool to predict changes in polymer microstructure.
They describe important parameters like porosity, loss of weight, erodedpolymer area and may, therefore, become an important tool to predict therelease kinetics of proteins and peptides from such polymers in the future.
Controlled Release Devices for the Delivery of Proteins and Peptides
from Polymers
For the delivery of proteins and peptides, non-degradable andbiodegradable polymers have been used in controlled release. Due to thehigh molecular weight of proteins, direct diffusion through a nondegradable polymer matrix is not possible(40). By the introduction of a
network of pores in manufacturing, however, the release of such large
compounds does occur(47). Another possibility is the creation of hydrophilic pathways using swellable polymers, or embedding the compounds into gels. In the case of biodegradable polymers, pores are createdupon erosion of the polymer matrix enabling the release of proteins fromthe dosage form. A l l of these options have certain advantages and disadvantages. Embedding suspended compounds into a non-degradable matrix
prevents some of the protein from being released(42), or might causesome instabilities due to the intense contact with organic solvents(43).
Their disadvantage with respect to parenteral application, however, is theneed for removing such systems after therapy. By using gels as a carrier,the protein may be released very quickly if not combined with some othersort of material(44-45). Degradable polymers change their properties substantially during erosion, a characteristic which may or may not be beneficial for the stability of proteins and peptides. In general, decisions about
the suitability of a release device for specific proteins or peptides and appropriate manufacturing technology must be made on a case by case basis.
In the following section, we provide a number of examples from theprogress we made in the delivery of proteins from polymers.
Biodegradable Polymers for Immunization. The idea of using polymers
as antigen releasing carriers for the stimulation of immune responses

emerged in the seventies (46). Due to their ease of application and theprogress in their development, microspheres emerged as a potentiallyefficient carriers to enhance the immune response (47). They have so far
been used for a number of vaccines (48-50). These microspheres weremade from polymers based on poly-lactic acid ( P L A ) and its copolymers
with glycolic acid ( P L G A ) . For these applications, loading a polymer witha high molecular weight protein requires adjusting the desired release rate
and preventing the protein's loss of activity.
In an attempt to develop a controlled release system for vaccination
against tetanus, we investigated the design of microspheres with a desired
release rate and methods to maintain the imunogenicity of the processed
tetanus toxoid(57). For the manufacturing of microspheres, the solvent
evaporation and solvent extraction methods were applied to the double
emulsion technique. The solvent evaporation method has been used successfully for the encapsulation of hydrophilic drugs and peptides(52-5J)
and was recently applied to the encapsulation of proteins(54). According
to the protocol,of saline tetanus toxoid solution (3600 Lf/mL) was
emulsified by vortex mixing in l m L organic solution (methylene chloride
or ethyl acetate) containing 200 mg of polymer. The emulsion was homogenized by sonication and dispersed into l m L of a 1% aqueous P V A
solution by vortex mixing. The final double emulsion was diluted byadding it to 100 m L of a 0.3% P V A solution while stirring with a magnetic stirrer. The organic solvent was then allowed to evaporate for 3
hours (solvent evaporation) or extracted by adding 200 m L of 2%(V/V)
aqueous isopropanol solution (solvent extraction). Both techniques produced microspheres with a high loading capacity (>80%). Critical points
in the encapsulation procedure are the protein stability upon contact withmoisture(55) and organic solvent (43) during the formation of the emulsion as well as the subsequent freeze drying.
Protein stability under the manufacturing conditions was investigated varying three parameters. We varied the type of solvent for the dissolution of polymer using methylene chloride and ethyl acetate and investigated the effect of three different types of stabilizers: Pluronic F68, P E G
4,600 and sodium glutamate. To mimic the conditions that prevail during
the manufacturing of microspheres, emulsions were prepared from 100/^Laqueous tetanus toxoid solution and dispersed it in l m L of organic solvent. These emulsions were freeze dried and investigated for aggregation
by HPLC(56). Table II shows the results from this study.
Table II gives a good survey on the impact of manufacturing conditions on
the stability of the protein. Sample one shows that moisture alone causes

Table II. manufacturing conditions and their effect on protein aggregation
Set of conditions
Stabilizer
Aggregation
Solvent
(%)
14
1
none
none
none
Pluronic F68
02
P E G 4,600
3.9
3
none
Sodium4.4
4
none
Glutamatenone
Methylene
13.11
5
chloride
Pluronic F68
9.6
6
Methylene
chloride9
P E G 4,600
7
Methylene
chloride
Sodium
3.4
8
Methylene
Glutamatechloridenone
5.4
Ethyl acetate
9
the protein to aggregate substantially. The impact of even small amountsof moisture on the structure of lyophilized proteins can be tremendous(55). The aggregation of the protein in methylene chloride (sample 5)
is higher than in ethyl acetate (sample 9) which is unfortunately the poorerof both solvents for many biodegradable polymers. The use of stabilizers
like Pluronic F68 might be benefical in preventing some of the proteinsand peptides from aggregating or from adsorbing to hydrophobic surfaces
i.e. polymer forming the microspheres. This has been reported for a number of surfactants(57).
The goal for the release of tetanus toxoid from microspheres wascontinuous release over a period of weeks. For that purpose two types ofpolymers were used: L - P L A and D , L - P L G A 1 : 1 . The P L G A polymers areamorphous whereas L - P L A are crystalline. Depending on their molecularweight they erode in weeks or months(5#). To investigate the effect oferosion velocity the polymers were chosen with a molecular weight of either 100,000 or 3,000 Da. The release was studied by incubation of the
microspheres in phosphate buffer p H 7.4 at 37°C. Due to the slow degradation of the polymers it was sufficient to replace the buffer at least every
5 days. The released protein was determined by using a microBCA protein
assay. The results for the two types of polymers are shown in Figure 12a
and b. The release rate was clearly affected by polymer composition withP L G A faster than from P L A at the same molecular weight. In both cases,

Figure 12. In vitro release of tetanus toxoid from different types ofpolymer:a) • P L A (Mw 100,000); • P L G A (Mw 100,000)b) a P L A (Mw 3,000); a P L G A (Mw 3,000)(Reproduced with permission from ref. 56. Copyright 1993
Butterworth-Heinemann Ltd.)the release from the low molecular weight polymers is faster. This result isdue to the smaller size of microspheres prepared from these polymers
since the organic polymer solution had a lower viscosity. Other reasons
could be their faster degradation due to differences in molecular weight.
A l l preparations release over a period of weeks, as was intended. With the
exception of low molecular P L G A , there is no substantial burst effect asreported for other vaccine preparations (59).
A general disadvantage of methods that use chemical reactions or
physical interactions like chromatographic methods for the determinationof protein concentrations is the lack of structural information. We, therefore, tested the in vivo response to the encapsulated tetanus toxoid by injecting the microspheres subcutaneously into mice. The sera was assayed
for the antitoxin units using the toxin neutralization test(60-67) as well as
an E L I S A test. Figure 13a and b show the results for all microspheres aswell as a control injection of fluid toxoid and aluminum adsorbed toxoid.
The use of microspheres increases the immune response compared to thefluid toxoid. The use of aluminum adsorbed toxoid produces initiallyhigher antitoxin titers which decrease, however, more rapidly compared tothe encapsulated toxoid. The levels resulting from microsphere injection
are considerably higher than the estimated minimum protective level
which is about 0.01 A U / m L ( 6 2 ) . The antibodies determined by theneutralization test and the IgG antibodies reach their maximum at different
times. This could be explained by the induction of progressive affinity
maturation tetanus antibodies induced by prolonged exposure to low
concentrations of tetanus toxoid.
Protein Delivery from Poly(anhydride) Microspheres. A major
problem associated with many dosage forms for drug release is the releaseof large amounts of the drug during initial stages of release known as theinitial burst-effect. For microspheres, this burst effect tends to be enhanced since microspheres have a larger surface area to volume ratio thanmore sizable slabs. In addition, the change of biological activity of proteins can occur during microsphere formulation because of their sensitivity
to environmental alternation. Our goal in this study was to preparepoly(anhydride) microspheres which permitted the controlled release of
biologically active proteins without any initial burst in release. We selected two enzymes, trypsin (Mw24,000) and heparinase (Mw43,000) in
addition to B S A ( M w 62,000) which has been used as a model protein.
The measurement of enzymatic activity allowed us to investigate the effect of microsphere formulation on protein activity.
We encapsulated proteins into poly(anhydride) microspheres withthe solvent evaporation method by using a double emulsion (54).
However, this technique comprises of a number of formulation processes
which may affect protein activity: contact of aqueous protein solution with

•

Fluid toxoid

H

PLAMW 3,000

9PLAMW 100,000m PLGAMW 3,000•

PLGA MW 100,000•

Alum-adsorbed toxoid•

Fluid toxoid

B

PLAMW3,000•

PLAMW 100,000

0

PLGAMW 3,000•

PLGAMW 100,000•

Alum-adsorbed toxoid

time [weeks]

4

8

13

time [weeks]
Figure 13. Results from the in vivo test of tetanus toxoid loaded m i crospheres:a) In vivo neutralizing antibody response (antitoxin units/ml serum),b) IgG antibody response by E L I S A .
(Reproduced with permission from ref. 56. Copyright 1993
Butterworth-Heinemann Ltd.)the organic phase, vortex mixing, exposure to ultrasound and freeze drying.
The activity loss of proteins was measured based on the specific activityof proteins at each stage of the process (63). Table III shows the effect of
the formulation process on the enzymatic activity of trypsin.
Table III. Activity loss of trypsin after eachstep of microsphere preparation
remaining activity (%)
Preparation step100
Trypsin solution
vortex mixing
100
59
sonication for 30s
56
3 h solvent evaporation
freeze drying
59

The sonication process which was performed in presence of the organic
solvent to prepare a primary emulsion, causes the main activity loss ofprotein, while freeze-drying did not have any addition effect. It is known
that ultrasound has an effect on the biological properties of proteins inaqueous solution due to a number of factors such as temperature, mixing
and cavitation(64). Table IV shows the effect of ultrasound exposure period on the activity loss of proteins during microsphere preparation together with the additional effect of some protein stabilizers(65).

Table IV. Activity loss of trypsin during microsphere preparetion (66)Period ofType ofAmount ofremainingstabilizer (%activitysonication [s] stabilizer
w/w)
80.5
10
none
none
67.3
20
none
58.9
30
5
58.6
30
BSA
10
B
S
A
58.1
30
4
Glycine30
58.7*
30
Glycine
7.5
59.8*
30
Glycine8
56.6*
Glycine1530
57.2** Differences within the error margin of the test

The extent of activity loss becomes smaller with a decrease in the exposure period of sonication. In addition, the decreased period did not reduce
the size of the primary emulsion. These findings demonstrated that the observed activity loss results from the sonication probe and not from the increase in the interfacial area between aqueous and organic phase. Little effect of the stabilizers used here on activity loss was observed, indicating
the poor potential of these substances to protect the investigated proteins
against ultrasound exposure. Trypsin and heparinase were encapsulated to
study the enzymatic activity of proteins in poly(anhydride) microspheres.
Figure 14 shows the comparison of the remaining activity between theproteins encapsulated in microspheres and the proteins in a soluble form
after incubation in phosphate buffer pH 7.4 at 37°C. A t different stages ofthe experiment, the remaining % activity of proteins in microspheres wasassessed by determining the activity of protein that could be extracted
from these systems (66) These results clearly indicate that encapsulationof proteins inside poly(anhydrides) may stabilize their activity for a shortperiod of time (l-2days).
Figure 15 shows release profiles of B S A from p(FAD-SA) 25:75 atvarious protein loadings. No initial burst is observed irrespective of theprotein loading. The protein is released for up to three weeks at a nearconstant rate.
The release rate of protein depends on the monomer composition ofpoly (anhydrides) used as can be seen from Figure 16. Release periods ofseveral days to a couple of weeks are possible for poly(anhydride) microspheres by changing their monomer composition(66).
Biodegradable Injectable Nanospheres Composed of Diblock
Poly(ethyleneglycol)-Poly(lactic-co-glycolic acid) Copolymers A major
challenge in the area of parenteral administration of drugs is the development of a drug carrier that is small enough for intravenous application andwhich has a long circulation half-life. There are numerous potential applications for such a system: the suppression of toxic side effects which canoccur when the drug is injected in the form of a solution, or the protectionof sensitive compounds against degradation in the plasma. Consequently
there have been many approaches to develop such systems (67-68). The
major obstacle in the use of injectable systems is the rapid clearance fromthe blood stream by the macrophages of the reticuloendothelial system(RES). Polystyrene particles as small as 60 nm in diameter, for example,
are cleared from the blood within 2 to 3 min(69). B y coating these particles with block copolymers based on polyethylene glycol and polypropylene glycol, their half-life was significantly increased (70). Polystyrene
particles are, however, not biodegradable and are not therapeutically useful. For this reason, only liposome systems have been developed for intravenous administration (71-76). Due to their small size they were expectedtime [h]Figure 14. Stability of Heparinase and Trypsin with and without
polymer encapsulation in phosphate buffer p H 7.4: o Heparinase
solution, • Heparinase encapsulated, • Trypsin solution, • Trypsin
encapsulated.
(Reproduced with permission from ref. 66. Copyright 1993 Plenum
Press.)
120

0

7

14

21

28

time [days]

Figure 15. Release of B S A from poly (anhydride) microspheres depending on loading: o 1% B S A , • 2% B S A , • 4% B S A .
(Reproduced with permission from ref. 66. Copyright 1993 Plenum
Press.)

to freely circulate in the blood. It has been found, however, that they arecleared from the blood by uptake through the reticuloendothelial system(RES). The coating of liposomes with poly(ethyleneglycol) (PEG) increased their half life substantially (71-77). The presence of the flexible
and relatively hydrophilic P E G chains induces a steric effect at the liposome's surface, thus reducing protein adsorption and RES uptake (77).
Rather than using liposomes, we designed PEG-coated biodegradable nanospheres that consist of polymer only(7&). These are among thefirst degradable polymer-based systems with potential use for prolonged
intravenous administration or drug targeting.
The 'ideal' nanosphere (Figure 17) is biodegradable, biocompatible,
has a size of less than 200 nm, and has a rigid biodegradable core. The
PEG-coated surface theoretically provides a prolonged half-life in the
blood by masking it from the R E S . W e chose the F D A approved
polyesters poly(lactic-co-glycolic acid) ( P L G A ) to form the core of theparticles. These polymers allow us to adjust erosion rates and therefore
drug release by changing the monomer ratio. We synthesized a series of
diblock copolymers P L G A - P E G , starting with P E G of various chain
lengths and progressively increased the chain length of P L G A . Stannous
octoate was used as a catalyst for the ring-opening polymerization of lactide and glycolide in the presence of monomethoxy polyethylene glycol( M P E G ; M w 5,000; 12,000 and 20,000). The polymers obtained werecharacterized by nuclear magnetic resonance spectroscopy ( N M R ) , gelpermeation chromatography (GPC), infrared spectroscopy (IR), differential scanning calorimetry (DSC) and X-ray diffraction. The polymerization
reaction was followed by G P C from which molecular weight and polydispersity were determined. The consumption of lactide and glycolide can beobserved from a decrease of peak D (Figure 18). The shifting of the peakP towards lower retention times (higher molecular weights) indicates thatan addition reaction takes place at the hydroxyl end group of the P E G
chains. The polydispersity remains low during the polymerization( M n / M w less than 1.1). Both the crystallinity and the glass transition temperature (Tg) were determined by using D S C . The diblock copolymers
show a single Tg, which shifts to higher temperatures with an increase of
the P L G A chain length inside the P L G A - P E G (Figure 19). A complete
discussion of the D S C results can be found elsewhere(79). Random lactic
acid-ethylene oxide copolymers show two distinct Tg's (80), suggesting aphase separation inside the polymer. We suppose that the single T g we observed is due to an entanglement effect of long P E G and P L G A chains in
the polymers, which cannot easily phase-separate. The exact chemicalcomposition of the polymers was determined by C N M R spectroscopy.
Sterically stabilized particles were prepared from diblock P L G A P E G copolymers or from blends of P L G A and P L G A - P E G . These polymers were dissolved in a common solvent (ethyl acetate or methylene
i 3

120

time [days]

Figure 16. Release of B S A from poly(anhydride) microspheres depending on polymer type: o p(SA), • p(FAD-SA) 8:92, • p ( F A D SA) 25:75, • p(FAD-SA) 44:56.
(Reproduced with permission from ref. 66. Copyright 1993 Plenum
Press.)

Figure 17. Schematic representation of an 'ideal' nanosphere having abiodegradable drug containing core and a coating of Polyethylene
glycol chains.

"i

1

8

1

9
10
time (minutes)

1—

11

Figure 18. A T i m e course of the polymerization reaction between
P E G and lactide/glycolide followed by G P C : Peak P: polymer P E G P L G A , peak D: starting material (glycolide+lactide).

Figure 19. D S C thermograms of P L G A - P E G diblock copolymers(heating rate 10 °C/min, M W PEG=5000, for increasing chain lengthof P L G A ) .

chloride). A n 0 / W emulsion was formed by vortex mixing and sonicating
for one minute. The organic solvent was then slowly evaporated, at room
temperature, under gentle stirring for two hours. Slow removal of the solvent allows the reorganization of the polymer chains inside and on the surface of the droplets. B y optimizing the viscosity of the organic phase, thevolume ratio of the two phases and the sonication parameters, nanospheresof a mean size of about 150 nm were obtained. After removal of the organic solvent, the nanospheres were isolated from the aqueous phase by
centrifugation. They could later be readily redispersed in water, probably
due to the presence of P E G chains on their surface.
The nanospheres were characterized by various methods. The surface composition of lyophilized particles was determined by X-ray photoelectron spectroscopy (XPS). Figure 20 shows the carbon Is envelopesthat were observed by analyzing the nanosphere powder. One main carbon
environment is observed for P E G . It corresponds to the ether carbon C - O(81). The spectrum of P L G A polymer (curve a) shows three main carbon
environments. In the case of P E G - P L G A nanospheres, the peaks are stillpresent. A s the information obtained by X P S corresponds to the first layers on the nanosphere's surface (about 5 nm(S2)), this spectrum indicatesthat P E G is present on the surface of the nanosphere powder. Moreover,
we verified that P E G remains on the surface after incubation in distilledwater for 24 hours at 37 °C.
To investigate the in vivo half life of the systems, the nanospheres
were labeled with ^Indium, which has been used for gamma-scintigraphy studies in humans (83). Hn was attached directly to the polymer
chain by complex formation following a method that has already beenused for liposomes (83). The incubation at 37 ° C in P B S or horse serum
for more than 24 hours showed that there was less than 4 % label loss.
This indicated that the label is bound tightly enough to prevent loss duringin vivo study. Subsequently biodistribution studies were performed by i n jecting
In-labeled P L G A and PEG-coated nanospheres into the tail veinof Balb/c mice (18-20g). Five minutes after injection of uncoated P L G A
nanospheres, 40% of nanosphere-associatedl n radioactivity was foundin the liver and approximately 15% in the blood. In the case of P E G coated nanospheres, the results were reversed (15% of injected radioactivity in the liver, 60% i n the blood). After four hours, 30% of thenanospheres were still circulating i n the blood, whereas the P L G A
nanospheres had disappeared completely from the blood.
1

ll

1 1 1

U 1

These results are very encouraging and we hope that thesenanospheres might be used as drug carriers for intravenous administrationin the future. Studies are now underway to achieve the encapsulation ofdifferent hydrophobic drugs and small peptides into the core of thesenanospheres. W e are also determining the effect of P E G molecular weight

6e+3
5e+3.
4e+3.
(0
c
3

3e+3.
2e+3.
1e+3.
Oe+0
283

285

287

289

291

293

binding energy [eV]

Figure 20. Surface analysis by X P S of P L G A powder: a) P L G A
powder, b) P E G crystals, c) P L G A - P E G nanospheres.

and density on the R E S uptake and on drug encapsulation efficiency andrelease.
Summary
Progress in various fields of protein delivery has been illustrated. Usingpoly(anhydrides) as a model we showed that it is possible to describe
erosion mechanisms and to develop mathematical models of the complex
processes involved in polymer degradation. Based on such models the design of delivery systems using computers might become possible in the
future. Concerning manufacturing technologies for protein delivery, we
showed how existing systems, like microspheres, can be improved andprovided the development of vaccines as an example for future applicationof such systems. Finally the recent progress in the development of new
dosage forms was illustrated with the example of biodegradable
nanospheres with an increased circulation half-life. A l l these projects aim
at a better understanding and an improved design of protein delivery systems.
Acknowledgments
The authors want to thank Dr. R . K . Gupta and Dr. G.R. Sieber for theircontribution to the protein stability study and in the vivo immune response
studies. Thanks are due to the Deutsche Forschungsgemeinschaft who
sponsored the polymer degradation studies with grant N o G O 565/1-1 andto the National Institutes of health who sponsored the same project with
grant No. G M 26698. We thank Scios Nova for supplying us with polymers.

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RECEIVED April 19,1994