Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations
2008 Polymer chemistry effects on protein release and immune activation Senja Katarina Lopac Iowa State University
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Polymer chemistry effects on protein release and immune activation
by
Senja Katarina Lopac
A thesis submitted to the graduate faculty
in partial fulfillment of the requirements for the degree of
MASTER OF SCIENCE
Major: Chemical Engineering
Program of Study Committee: Balaji Narasimhan, Major Professor Michael Wannemuehler Aaron Clapp
Iowa State University
Ames, Iowa
2008
Copyright © Senja Katarina Lopac, 2008. All rights reserved. UMI Number: 1450148
UMI Microform 1450148 Copyright 2008 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.
ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, MI 48106-1346 ii
TABLE OF CONTENTS ACKNOWLEDGEMENTS ...... vi
CHAPTER 1: INTRODUCTION...... 1
1.1 Introduction ...... 1
1.2 References ...... 4
CHAPTER 2: BACKGROUND AND LITERATURE REVIEW...... 5
2.1 Summary ...... 5
2.2 Polyanhydrides...... 5
2.2.1 History ...... 5
2.2.2 Structures...... 6
2.2.2.1 Aliphatic...... 6
2.2.2.2 Aromatic ...... 7
2.2.2.3 Novel polyanhydride...... 7
2.2.2.4 Other ...... 8
2.2.3 Synthesis...... 9
2.2.3.1 Melt polycondensation...... 9
2.2.3.2 Solution polymerization...... 10
2.2.3.3 Dehydrative coupling...... 11
2.2.3.4 Ring-opening polymerization ...... 12
2.2.4 Characterization...... 13
2.2.5 Degradation and erosion...... 15
2.2.6 Polyanhydrides for drug delivery ...... 17
2.3 Microspheres ...... 18
2.3.1 Fabrication...... 18
2.3.1.1 Hot-melt microencapsulation...... 18 iii
2.3.1.2 Solvent evaporation ...... 19
2.3.1.3 Solvent removal ...... 19
2.3.1.4 Spray drying...... 20
2.3.2 Characterization...... 21
2.3.3 Release...... 22
2.4 Mechanisms of Immune Response...... 23
2.4.1 Innate and adaptive immunity ...... 23
2.4.2 Major histocompatibility complex ...... 23
2.4.3 Dendritic cells...... 25
2.5 Adjuvants and Vaccines...... 27
2.5.1 Novel therapies using polymers ...... 28
2.6 Conclusion...... 30
2.7 References ...... 32
CHAPTER 3: RESEARCH OBJECTIVES AND ORGANIZATION ...... 38
3.1 Research Objectives ...... 38
3.2 Thesis Organization...... 39
CHAPTER 4: EFFECT OF POLYMER CHEMISTRY AND FABRICATION
METHOD ON PROTEIN RELEASE AND STABILITY FROM
POLYANHYDRIDE MICROSPHERES ...... 40
4.1 Abstract ...... 40
4.2 Introduction ...... 41
4.3 Materials and Methods ...... 44
4.3.1 Materials...... 44
4.3.2 Monomer/polymer synthesis ...... 44 iv
4.3.3 Protein...... 45
4.3.4 Contact angle measurements ...... 45
4.3.5 Microsphere fabrication methods...... 46
4.3.5.1 Solid-oil-oil emulsion ...... 46
4.3.5.2 Cryogenic atomization...... 47
4.3.6 Microsphere characterization ...... 48
4.3.7 Ova release ...... 49
4.3.8 Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).....50
4.3.9 Western blot...... 51
4.4 Results ...... 52
4.4.1 Contact angle...... 52
4.4.2 Release of ovalbumin from polyanhydride microspheres...... 53
4.4.3 SDS-PAGE...... 56
4.4.4 Western blot...... 58
4.5 Discussion ...... 59
4.6 Conclusion...... 62
4.7 Acknowledgements ...... 62
4.8 References ...... 63
CHAPTER 5: EFFECT OF POLYMER CHEMISTRY ON IMMUNE
ACTIVATION OF DENDRITIC CELLS...... 67
5.1 Introduction ...... 67
5.2 Materials and Methods ...... 70
5.2.1 Materials...... 70
5.2.2 Polyanhydride synthesis ...... 71 v
5.2.3 Microsphere fabrication and characterization ...... 72
5.2.4 Isolation and culture of dendritic cells ...... 73
5.2.5 Stimulation of dendritic cells with polyanhydrides...... 73
5.2.6 Staining of dendritic cells...... 74
5.3 Results ...... 74
5.3.1 Size distribution...... 74
5.3.2 Assessment of surface markers by flow cytometry...... 76
5.4 Discussion ...... 81
5.5 Conclusions ...... 84
5.6 References ...... 85
CHAPTER 6: CONCLUSIONS AND FUTURE WORK ...... 88
6.1 Conclusions ...... 88
6.2 Future Work ...... 90
6.2.1 Interaction of polyanhydrides and the immune system...... 90
6.2.2 Polyanhydride coated drug-eluting stents ...... 92
6.3 References ...... 95 vi
ACKNOWLEDGEMENTS
First and foremost, I would like to thank Dr. Balaji Narasimhan and Dr. Michael
Wannemuehler for their assistance, guidance, and direction on my projects. I would like to recognize Dr. Aaron Clapp for serving on my Program of Study committee as well.
I owe many thanks to the members of Dr. Narasimhan’s and Dr. Wannemuehler’s research groups, especially Maria Torres and Jenny Wilson-Welder. I cannot express how appreciative I am of the hours they have helped me, between explaining protocols, to spending numerous hours staining/stimulating/feeding dendritic cells, to giving me valuable comments on my chapters and presentations. Their optimistic attitude and upbeat spirits made those extra hours in the lab much more bearable.
I would also like to recognize the undergraduate students for working in our laboratories, in particular Kristina Staley, Mary Byron, Chelsea Sackett, and Kathleen Ross; the hours spent taking release samples, conducing BCA assays (over and over!), synthesizing diacids/pre-polymers/polymers, feeding cells, and helping with stimulating were staining dendritic cells were much appreciated.
I would like to thank Warren Straszheim and Jerry Amenson for their helping in training and imaging on the scanning electron microscopy; and Guy Sander for his numerous frustrating hours spent helping me with HPLC.
vii
I would like to acknowledge my sources for funding, including AraVasc, ORI-MURI, and the Iowa State University Institute of Combinatorial Discovery.
I would like to thank my fellow graduate students, who made working much more enjoyable. Though I won’t miss the long hours spent in Sweeney, I will definitely miss having so many friends in such a close proximity.
And finally, I owe a huge thanks to my family and close friends for keeping my spirits up and giving me the support I needed to finish my studies in graduate school. 1
CHAPTER 1
INTRODUCTION
1.1 Introduction
Vaccines have prevented at least two million deaths worldwide in 2003 alone, yet the
World Health Organization has estimated that at least 27 millions infants go without proper immunizations, making them susceptible to diseases easily prevented 1. Current alum-based adjuvants (which are molecules that enhance immune response) have been shown to be weak to poor at inducing an immune response; in addition, often more than one dose is needed for protection. The use of Freund’s complete adjuvant and the addition of lipopolysaccharide have both received much attention as inclusions in carriers, yet both have been proven to be toxic in humans 2,3. Clearly there is a need to
develop new adjuvants for vaccine delivery that are easy to manufacture and non-toxic.
Biodegradable polymeric systems, in particular poly(D,L-lactide-co-glycolide) (PLGA)
and polyanhydrides, have received much attention as potential vaccine carriers. These
polymers have the ability to degrade by hydrolysis into their non-toxic monomeric units,
lactic and glycolic acids and dicarboxylic acids, respectively. Parenteral delivery of these
polymers can be achieved by fabricating microspheres, which allows the drug to be
released over a desired period of time; due to the degradation of the polymers, there is no
need to remove the device once the drug has been delivered. However, the relatively
acidity (due to dissolution of the lactic and glycolic acid monomers) and the bulk eroding
properties of PLGA may be detrimental to the stability of encapsulated proteins, causing
denaturation and even irreversible aggregation 4,5. On the other hand, recent studies with polyanhydrides 6,7 have shown that the surface erodible feature of these polymers can be 2
combined with less hydrophobic monomers to create a new class of amphiphilic materials that are preferable for a wide range of protein and vaccine delivery applications.
Polyanhydrides are currently being studied as possible alternatives to alum-based adjuvants 8. In addition, polyanhydrides are surface eroding, which allows the antigen to be released over a predictable period of time. This eliminates the need to administer subsequent doses of antigen in the case of vaccines 9. Because degradation varies
depending on the chemistry and structure, a tailored release rate can be achieved 10. The surface-eroding properties of polyanhydrides, combined with their relative hydrophobicity, implies that proteins are less likely to undergo moisture-induced aggregation, thereby retaining their stability 11,12. Polyanhydrides have been proven to be
compatible in vivo, as shown by the polyanhydride-comprised Gliadel® wafer.
Approved by the FDA in 1996, Gliadel® is used to deliver bis-chloroethylnitrosourea to
treat a form of brain cancer. In addition, an in vivo study demonstrated the
immunomodulatory properties of tetanus toxoid-loaded polyanhydride microspheres; by
8 changing the chemistry, a TH0 (balanced) immune response was attained . Protection
from diseases such as cancer require induction of TH1 (cell-mediated) immune responses;
13 traditional alum-based vaccines enhance TH2 (humoral) responses .
The overall objective of the research therein was to demonstrate the effect of polymer chemistry on release kinetics, protein stability, and immune activation. It is anticipated
that application of the results of these studies will lead to rational design of carriers for
specific therapeutic modalities. Polyanhydrides of various chemistries based on the 3
monomers of sebacic acid (SA), 1,6-bis(p-carboxyphenoxy)hexane (CPH), and 1,8-bis(p- carboxyphenoxy)-3,6-dioxaoctane (CPTEG) were investigated in course of this research; these polymers are shown in Figure 1.1. Chapter 2 summarizes literature regarding polyanhydride synthesis and characterization, microsphere fabrication and characterization, mechanisms of immune response, adjuvants and vaccines, and novel therapies using polymers. Chapter 3 lists the goals of the research project. Chapter 4 details the release kinetics of protein-loaded polyanhydride microspheres and analyzes the protein’s primary structure and activity once released from the microspheres. Chapter
5 describes the effects of polyanhydride chemistry on the activation of bone marrow derived dendritic cells as measured by studying changes in cell surface marker expression. Finally, Chapter 6 summarizes the work of the previous two chapters, and gives a direction as to where the work conducted will lead.
O O O O O O O n O n
O O O O O O O n
Figure 1.1 – Chemical structures of polymers used, from top, left to right: poly (sebacic acid), poly (1,6-bis(p-carboxyphenoxy)hexane), and poly (1,8-bis(p-carboxyphenoxy)- 3,6-dioxaoctane), where n represents the number of repeating monomer units. 4
1.2 References
1. Global Immunization Vision and Strategy: 2006-2015. Geneva, Switzerland: World Health Organization Department of Immunizations, Vaccines, and Biologicals and UNICEF Programme Division, Health Section; 2005. 2. Stuart-Harris CH. Adjuvant influenza vaccines. Bull World Health Organ 1969;41(3):617-21. 3. Johnson AG, Gaines S, Landy M. Studies on the O antigen of Salmonella typhosa. V. Enhancement of antibody response to protein antigens by the purified lipopolysaccharide. J Exp Med 1956;103(2):225-46. 4. Zhu G, Schwendeman SP. Stabilization of proteins encapsulated in cylindrical poly(lactide-co-glycolide) implants: mechanism of stabilization by basic additives. Pharm Res 2000;17(3):351-7. 5. Jaenicke R. Stability and stabilization of globular proteins in solution. J Biotechnol 2000;79(3):193-203. 6. Torres MP, Determan AS, Anderson GL, Mallapragada SK, Narasimhan B. Amphiphilic polyanhydrides for protein stabilization and release. Biomaterials 2007;28(1):108-16. 7. Torres MP, Vogel BM, Narasimhan B, Mallapragada SK. Synthesis and characterization of novel polyanhydrides with tailored erosion mechanisms. J Biomed Mater Res A 2006;76(1):102-10. 8. Kipper MJ, Wilson JH, Wannemuehler MJ, Narasimhan B. Single dose vaccine based on biodegradable polyanhydride microspheres can modulate immune response mechanism. J Biomed Mater Res A 2006;76(4):798-810. 9. Determan AS, Trewyn BG, Lin VS, Nilsen-Hamilton M, Narasimhan B. Encapsulation, stabilization, and release of BSA-FITC from polyanhydride microspheres. J Control Release 2004;100(1):97-109. 10. Eldridge JH, Staas JK, Meulbroek JA, McGhee JR, Tice TR, Gilley RM. Biodegradable microspheres as a vaccine delivery system. Mol Immunol 1991;28(3):287-94. 11. Ron E, Turek T, Mathiowitz E, Chasin M, Hageman M, Langer R. Controlled release of polypeptides from polyanhydrides. Proc Natl Acad Sci USA 1993;90:4176-4180. 12. Tabata Y, Gutta S, Langer R. Controlled delivery systems for proteins using polyanhydride microspheres. Pharm Res 1993;10(4):487-96. 13. Singh M, O'Hagan DT. Recent advances in veterinary vaccine adjuvants. Int J Parasitol 2003;33(5-6):469-78. 5
CHAPTER 2
BACKGROUND AND LITERATURE REVIEW
2.1 Summary
This chapter discusses the literature that is relevant for the polyanhydride work described
in this thesis. Section 2.2 provides an in-depth discussion of polyanhydrides, focusing on
the chemistry, synthesis, characterization, and what sets polyanhydrides from other
biodegradable polymers used in drug delivery. Section 2.3 details microsphere
fabrication and characterization. A brief introduction to immunology, mainly directed at
dendritic cells, innate versus adaptive immunity, and the major histocompatibility
complex, is given in Section 2.4. Adjuvants and vaccines are discussed in Section 2.5.
Section 2.6 summarizes the current state-of-the-art and raises unsolved issues in this area,
some of which are the subject of this thesis.
2.2 Polyanhydrides
2.2.1 History
In 1909, Bucher and Slade documented the first synthesis of polyanhydrides upon heating
isophthalic and terephthalic acid with acetic anhydride 1. Their work led to the
investigation of polyanhydrides as textiles by Hill and Carothers in the 1930s; they soon
learned that this class of polymers was not suited for such an application due to the
polymers’ capabilities to hydrolytically degrade. Hill and Carothers also discovered new
properties of polyanhydrides, such as their instability at high temperatures, which causes
them to transform into cyclic dimers and polymeric rings 2-4. It was not until the 1980s 6
that a connection was made between polyanhydrides, due to their degradation properties and biomedical applications.
2.2.2 Structures
Polyanhydrides can be grouped into different classes, based on the properties of their
constituent monomers. Varying the chemistry of the polyanhydride backbone can
considerably change the properties of the resulting polymer, such as solubility, mechanical strength, crystallinity, and degradation rate. While there are literally
hundreds of polyanhydride structures synthesized, two chemistries are predominantly
used in the medical field: aliphatic and aromatic.
2.2.2.1 Aliphatic
Poly(sebacic anhydride) was the first aliphatic polyanhydride synthesized by Hill and
Carothers in 1932 3,4. Aliphatic polyanhydrides are crystalline, brittle, typically melt at
temperatures below 100°C, and are soluble in chlorinated hydrocarbons. These
polyanhydrides are known to be sensitive to hydrolysis, and degrade and are eliminated
from the body within weeks 5. To fabricate a slower degrading aliphatic polyanhydride,
the alkyl chain length of the monomer can be increased, thereby increasing the
hydrophobicity of the polymer and lengthening the degradation time of the polymer 5.
Aliphatic polyanhydrides are usually prepared by melt condensation reactions, as their
diacids are typically not affected by heat 6. Typical diacids used for aliphatic
polyanhydrides include adipic acid, dodecanoic acid, fumaric acid, sebacic acid, and fatty
acid dimers 7. 7
2.2.2.2 Aromatic
Aromatic polyanhydrides of isophthalic and terephthalic acid were first synthesized by
Bucher and Slade in 1909 1. Typical diacids used include phthalic acid and several types of carboxyphenoxyalkanes 7. Aromatic polyanhydrides are less soluble in common organic solvents and characteristically more hydrophobic than aliphatic, melt at temperatures above 200°C, degrade over time spans of years, and possess fairly high strength and stability 8,9. These properties make pure aromatics difficult to process into films or microspheres, since these techniques usually require lower temperatures to melt at or for the polymer to be easily solubilized. In order to overcome these barriers, aromatics are typically copolymerized with other polyanhydrides, such as aliphatic diacids; the copolymers tend to take on the characteristic properties of their aliphatic
portion 10.
2.2.2.3 Novel polyanhydrides
The hydrophobic chemistry of polyanhydrides help to prevent water-induced covalent
aggregation of proteins since water penetration into the bulk is negligible; however, when
incorporated with a protein, noncovalent aggregation may result due to hydrophobic interactions 11,12. By incorporating a hydrophilic oligomeric ethylene glycol unit, such as
triethylene glycol, into the backbone of a hydrophobic aromatic polymer, both covalent
and noncovalent aggregation can be controlled 13. The resulting amphiphilic polymer is
also closer to bulk-eroding, a novel characteristic for the normally surface-eroding
polyanhydrides. However, upon copolymerizing with an aromatic polyanhydride, the 8
surface-eroding properties can be regained, providing a controlled release profile characteristic 13.
2.2.2.4 Other
Many other polyanhydride structures exist, each with their own characteristics. The
crystalline unsaturated polyanhydride homopolymers are insoluble in common organic
solvents; to solubilize in chlorinated hydrocarbons, unsaturated polyanhydrides are
copolymerized with aliphatic diacids, which also makes them less crystalline 14.
Crosslinked polyanhydrides are used for situations where a greater mechanical strength is
required, as most other polyanhydrides are rather brittle and cannot be used in load-
bearing applications 14. The copolymerization with imides can enhance the tensile
strength and thermal resistance of polyanhydrides, making them ideal for applications
such as sutures 15. Fatty acid-based polyanhydrides based on oleic acid and eurucic acid
are viscous liquids; typically, these polymers are copolymerized with an aliphatic
polyanhydride such as sebacic acid to solidify 16 .
9
O O
STRUCTURE R group from: R O n EXAMPLES x = 4 Poly(adipic acid) Aliphatic polyanhydrides CH x = 8 Poly(sebacic acid) 2 x x = 10 Poly(dodecanedioic acid)
para- Poly(terephthalic acid) meta- Poly(isophthalic acid) Aromatic polyanhydrides
x = 1 Poly[1-bis(p-carboxyphenoxy)methane] x = 3 Poly[1,3-bis(p-carboxyphenoxy)propane] O CH2 xO x = 6 Poly[1,6-bis(p-carboxyphenoxy)hexane]
O O Poly[1,8-bis(p-carboxyphenoxy)-3,6-dioxaoctane] Novel polyanhydrides O O
Figure 2.2 – Common structures of polyanhydrides used in drug delivery applications
2.2.3 Synthesis
Polyanhydrides can be prepared through a variety of techniques, including melt condensation in conjunction with acetic anhydride, solution polymerization, dehydrative coupling, interfacial condensation, and ring-opening polymerization.
2.2.3.1 Melt polycondensation
The conversion of dicarboxylic acids into polyanhydrides takes place in two steps, the first being the reaction of the monomers with excess acetic anhydride by reflux. By heating the aforementioned prepolymer under vacuum at a high temperature to eliminate the acetic anhydride, the subsequent polyanhydride is formed, with a molecular weight of
100 to 1000 16,17. Further studies indicated that polymerization at 180°C for 90 minutes, with a 10-4 mmHg vacuum and dry ice/acetone trap, provided the optimal molecular weight. Additionally, it was found that increasing the reaction time or reaction 10
temperature did not increase the molecular weight of the polymer 18. The use of
coordination catalysts, such as cadmium acetate, earth metal oxides, orZnEt2·H2O, could be employed to achieve even higher molecular weights with less reaction time 19. The resulting polyanhydride is dissolved in an organic solvent, and precipitated in hexane.
O O O O + OH R OH O
REFLUX
O O O O
O R O m
180 C, 90 min 10-4 mm Hg
O O O O
O R O n
Figure 2.3 – Schematic of melt polycondensation reaction
2.2.3.2 Solution polymerization
Solution polymerization employs a Schotten-Baumann condensation reaction to
synthesize polyanhydrides. It is advantageous to use this method over melt condensation
when synthesizing heat-sensitive monomers, due to possible charring at the high reaction temperature or the high melting point of the subsequent polymer. However, the diacid halide monomer needs to be of utmost purity in order to use this technique 20. Yoda and
Mikaye were the first to develop this technique in 1959; Subramanyam and Pinkus
further developed this method with the inclusion of an acid acceptor 21,22. Briefly, a
dehydrochlorination reaction occurs when a diacid halide is added to an ice-cold solution
of a dicarboxylic acid, in a dry nitrogen environment and in the presence of an organic
solvent, such as benzene, ethyl ether, or chloroform. Polycondensation was only 11
facilitated in the presence of an acid acceptor, such as triethylamine. The key to obtaining high molecular weight polymers is to add the diacid dropwise to the diacid halide solution to allow full contact, and not the reverse; otherwise an ionic salt is formed as a result of the triethylamine reacting with the terephthaloyl chloride complexes 21.
High molecular weight polymers are desirable to allow for control over the degradation process of polyanhydrides.
O O O O + OH R OH Cl R' Cl
BASE
O O O O * + BASE * HCl R O R' O n *
Figure 2.4 – Schematic of solution polymerization reaction
2.2.3.3 Dehydrative coupling
Dehydrative coupling, like solution polymerization, is an alternative to melt-
polycondensation reactions for monomers that are thermolabile 9. This method utilizes a dehydrative coupling agent, such as N’N-bis(2-oxo-3-oxazolidinyl)phosphonic chloride, in conjunction with a dicarboxylic acid monomer to produce the polymer 23. Other coupling agents include phosgene and diphosgene; the use of an acid acceptor such as poly(4-vinyl pyridine) has been shown to yield higher molecular weight polymers 24.
Dehydrative coupling is considered the simplest polymerization technique, due to its one step protocol and no need for an intermediate, prepolymer forming step. However, many drawbacks exist, such as the need for fresh catalyst for each reaction and the possible contamination of the polymer with catalyst by-products. In order to rid the polymer of the 12
catalyst derivatives, a washing step with a protic solvent is needed, which can potentially lead to hydrolysis of the polymer 23.
2.2.3.4 Ring-opening polymerization
Hill and Carothers first studied the ring-opening polymerization mechanism with adipic
acid and acetic anhydride under a reaction similar to melt polycondensation 2. Upon noting the low molecular weight of the polymer, they decided to subject this polymer (α- anhydride) to elevated temperatures to see if it would improve the molecular weight.
This resulted in a distillate of cyclic monomers and dimers (β-anhydride), and a much higher molecular weight polymer remaining in the still (ω-anhydride). Upon standing, the cyclic monomers and dimers converted into cyclic polymers (γ-anhydride), with a molecular weight of 5000; the schematic of this process is shown in Figure 2-4. They later extended their studies to include a larger number of diacids, which included HOOC-
3,4 (CH2)n-COOH, with n being 4-12 or 16 .
O O α-anhydride
OH CH OH (linear polymer) 2 n
β-anhydride + ω-anhydride (cyclic dimer) (residue)
γ-anhydride (cyclic)
Figure 2.5 – Schematic of ring opening polymerization reaction
Albertsson and Lundmark 25 demonstrated the formation of poly(adipic anhydride) from oxepane-2,7-dione in the presence of stannous 2-ethylhexanoate. The preparation of this 13
cycle polyanhydride was completed in two steps, the first being the reflux of adipic acid with acetic anhydride and, consequently, the removal of acetic anhydride with a zinc chloride hydrate catalyst to form oxepane-2,7-dione monomer. The second step involved melt polymerization with a stannous 2 ethylhexanoate inhibitor; the pair also tried other
- + inhibitors such as cationic (AlCl3 or BF3·(C2H5)2O) and anionic (CH3COO K or NaH) categories, as well as other coordination-type inhibitors (dibutylinoxide) 25,26. First attempts to polymerize (80°C for 5 hours) resulted in a low molecular weight polymer.
Later efforts produced a polyanhydride with a slightly higher molecular weight after improving upon the monomer-initiator ratio and the optimal polymerization time (two hours) 26.
2.2.4 Characterization
Many characterization techniques are employed to analyze polyanhydrides and confirm
their purity, based on chemical structure, crystallinity, molecular weight, and thermal
properties. These techniques include Fourier transform infrared spectroscopy (FTIR),
Fourier transform Raman spectroscopy (FTRS), proton nuclear magnetic resonance
spectroscopy (1H-NMR), gel permeation chromatography (GPC), differential scanning
calorimetry (DSC), and X-ray diffraction.
Fourier transform infrared spectroscopy (FTIR) has been used to characterize
polyanhydrides by identification of a carboxylic anhydride doublet located between 1670
and 1800 cm-1 9. Aromatics normally show peaks at 1720 and 1780 cm-1, aliphatics at
1740 and 1810 cm-1; degradation of the polymer can be monitored by the ratio of the 14
peaks of 1810 and 1700 cm-1. Fourier transform Raman spectroscopy (FTRS) is used to
determine how the monomer composition changes when copolymerized with another
monomer by monitoring the methylene bands. These bands can be altered due to
deformation, stretching, rocking, and twisting. The location of carbonyl bands, present in all polyanhydrides, can also be an indication of the monomer change. Aromatic polyanhydrides show the carbonyl band pairs at 1764 and 1712 cm-1; for aliphatic
polyanhydrides, the bands are displayed at 1804 and 1739 cm-1 27.
1H-NMR is often used to determine the copolymer composition, the average length of the
sequence, the frequency of copolymer sequences, and the chain structure, such as block,
alternating, or random (given by degree of randomness) 28. Protons in aromatic
polyanhydrides, due to being closer to electronegative groups, exhibit chemical shifts at
6.5 to 8.5 ppm; aliphatic protons exhibit higher chemical shifts of 1 to 2 ppm 29.
For determining molecular weight, GPC is the most common method to use.
Characteristic weight average molecular weight (Mw) of polyanhydrides vary from 5,000
28 to 300,000; polydispersities range from 2 to 15, and typically rise with increasing Mw .
Viscosity measurements can also be used for verifying molecular weight; the relationship between viscosity and molecular weight can be established using the Mark-Houwink equation. For example, for Poly(CPP-SA), the relationship was found to be
o [η]23 C ×= 1088.3 − M 658.07 CHCl3 w
Here, η is the intrinsic viscosity 28. 15
Differential scanning calorimetry (DSC) can be used to verify thermal properties, such as melting and glass transition temperatures, as well as heats of fusion. Typically, the glass
30 transition temperature, Tg, decreases with increasing chain length of the polymer . For
processing polyanhydrides into drug delivery devices, Tm is needed to dictate the lowest temperature needed for melt pressing or molding, while Tg determines the lowest
31 temperature for compression molding . A wider range of values between Tg and Tm is observed with Poly(CPH) (32°C and 140°C, respectively) 32 than with Poly(SA) (62° and
79°C, respectively) 30; copolymers exhibit temperatures between the two homopolymers .
Wide angle X-ray diffraction, a combination of differential scanning calorimetry (DSC)
and X-ray diffraction, and 1H-NMR can be used to determine the crystallinity of
polyanhydrides 33,34. In general, most aromatic and aliphatic homopolymers were found
to be semi-crystalline. A copolymer rich in either monomer exhibited a slight decrease in
crystallinity; while copolymerizing at equal mole ratios, the resulting polymers became
amorphous 33.
2.2.5 Degradation and erosion
Polymer degradation can occur by either enzymatic or hydrolytic degradation; the latter is much preferred for drug delivery applications, because of large amount of water naturally present in the body 35. Polyanhydrides degrade by the hydrolysis of their anhydride
linkages, in the presence of water, to form dicarboxylic acids, and further more to form
carbon dioxide and water. While monomeric anhydrides degrade in the presence of both
acids and bases, hydrolysis of polyanhydrides are base catalyzed 5. To impede the 16
degradation rate, methylene groups can be introduced into the core of the polymer.
Degradation can be affected by many factors, such as pH of the medium and hydrophobicity and crystallinity of the polymer. As expected, the higher the hydrophobicity of the polymer, the slower the degradation rate, as the rate at which water penetrates into the bulk slows in correlation 35. Polyanhydride degradation is base
catalyzed; an increase in pH of the medium increases the rate of degradation; acidic
media tends to impede and can even halt degradation in some cases 9. An increase in
crystallinity of a polyanhydride can also lead to a slower degradation rate 36.
Two types of polymer erosion mechanisms exist: bulk erosion and surface erosion 35.
Considering polyanhydrides are primarily hydrophobic, they are characterized as surface eroding, meaning they degrade layer by layer, and material is only lost from the surface.
This is opposed to bulk erodible polymers, which degrade as a whole, since water penetrates into the bulk. Figure 2.5 gives a schematic of bulk eroding versus surface eroding mechanisms.
Figure 2.6 – Bulk eroding versus surface eroding mechanisms 17
Biodegradation is the process by which the polymers degrade into products that are either metabolites or could be easily metabolized by a living organism. Degradation varies drastically depending on the monomer used. Poly(sebacic acid) and Poly(1,6-bis-(p- carboxyphenoxy)hexane, for example, degrade in 54 days and 1 year, respectively, in saline37. By combining different ratios of polyanhydride monomers during the
fabrication of copolymers, degradation rates can be tailored for a specific application.
O O O O O O O O " + " " " R1 OH OH R2 O n * R1 O R2 n * *
Figure 2.7 – Hydrolysis reaction of polyanhydrides
2.2.6 Polyanhydrides for drug delivery
Polyanhydrides were not researched for drug delivery applications until 1983, when
Langer and co-workers reported their potential for controlled drug delivery, based on
their biodegradable properties 38. There are several reasons why polyanhydrides are
suitable candidates for use in drug delivery applications. Due to their ability to surface
erode, polyanhydrides exhibit a predictable release rate; this can be modified simply by
changing the chemistry or the molecular weight. As mentioned previously,
polyanhydrides are quick and easy to produce, and can be manufactured in a variety of
ways, depending on the properties of the preceding monomers. Finally, polyanhydrides
are non-toxic and non-mutagenic, and degrade into carboxylic acids that are further
catabolized intocarbon dioxide and water. For use in medical applications, they can be
sterilized by terminal-γ-irradiation without affecting molecular weight or structure 39-41.
18
One of the most prominent uses of polyanhydrides is with the Gliadel® wafer, comprised
a copolymer of 1,3-bis(p-carboxyphenoxy)propane and sebacic acid. Approved by the
Food and Drug Administration (FDA) in 1996, Gliadel® is used to deliver bis-
chloroethlylnitrosourea (BCNU) to treat a form of brain cancer known as glioblastoma
multiforme 42.
2.3 Microspheres
Biodegradable polymers are preferred for parenteral drug delivery systems as there is no
need to remove any device post implantation. A size of less than 25 μm is normally preferred for such applications; this size also allows for delivery into the tissue by the use of a syringe 43.
2.3.1 Fabrication
The choice of which fabrication method to use in order to produce the appropriate microspheres depends on the properties of both the desired polymer and the drug to be encapsulated, including hydrophobicity, crystallinity, thermal properties, and stability.
Four of the most common ways to fabricate microspheres include hot-melt microencapsulation, solvent evaporation, solvent removal, and spray drying.
2.3.1.1 Hot-melt microencapsulation
Hot-melt microencapsulation involves adding a mixture of melted polymer and solidified
drug particles to a polymer immiscible-solvent at a temperature slightly above the
melting point of the polymer. The spherical structure forms when the solution is cooled; 19
the resulting microspheres are rinsed with a nonsolvent, such as petroleum ether.
Adjusting the stirring speed can change the size of the uniform microspheres. While the microspheres formed have a smooth surface, only thermolabile polymers can be fabricated by this process due to the high temperatures used 44.
2.3.1.2 Solvent evaporation
An oil-in-water emulsion is a simple method for making microspheres; since heat is not
involved, this method can be used to produce microspheres from heat-sensitive polymers.
The polymer is dissolved in an organic solvent, such as methylene chloride, and stirred
into an aqueous solution of surfactant, such as poly(vinyl acetate). The resulting solution
is stirred for two hours, which is sufficient time to allow the organic solvent to evaporate
completely, and leaves behind the hardened microspheres 45-49
For hydrophilic polymers, a double-emulsion, consisting of water-oil-water, can be used.
The aforementioned oil-in-water method is altered slightly by adding a small amount of
the aqueous phase to the organic solution prior to dispersing 50. However, the problem
exists that the process is aqueous and increases the possibility that the polymer can begin
to degrade during the fabrication of the microspheres 51-54.
2.3.1.3 Solvent removal
A variation of this technique is sometimes referred to as solvent removal, though it is a
similar process. This is also referred to as solid-oil-oil emulsion. The advantage of this 20
technique is that it prevents hydrolysis of the polyanhydride by eliminating water from the process 50.
Polymers and excipients are dissolved in an organic solvent, such as methylene chloride.
An organic oil nonsolvent, such as silicon or paraffin oil, is slowly added to the solution with stirring; a surfactant such a Span 85 is slowly introduced. The emulsion is added to an immiscible nonsolvent, such as petroleum ether or hexane, and stirred for a couple hours; during this time, the oil slowly extracts the organic solvent, and microspheres are formed 55,56.
2.3.1.4 Spray drying
Mathiowitz developed a spray-drying method for producing microspheres; the
polyanhydrides and drugs were dissolved into methylene chloride and sprayed through a
0.5 mm diameter atomizing nozzle. The microspheres are dried by a flow of nitrogen gas
as they descend to the base of the spray dryer. Lower crystallinity polymers, such as
aromatics and copolymers of aromatics and aliphatics, produced spherical, smooth
microspheres. Aliphatic polymers tended to have a rutted surface; despite this, the
method is conducive as it is fast and results in the production of uniform microspheres, 1-
5 μm in diameter, and can easily be scaled up 31,57. This method is preferred for making
microspheres as it can be easily scaled up for industrial applications and also can be used
for heat-sensitive drugs.
21
A modified spray drying technique, known as cryogenic atomization, was first described by Johnson and Cleland with polylactic-co-glycolic acid 58,59. The polymer and any
protein or other excipients are dissolved in a solvent such as methylene chloride, acetone,
or ethyl acetate, and sprayed over frozen ethanol topped with liquid nitrogen. The liquid
nitrogen acts to hold the microsphere structure intact by freezing and gradually evaporate;
the ethanol removes the residual solvents. The solution is filtered, and microspheres
were found to be between 50 and 60 µm.
2.3.2 Characterization
Due to the small physical size of microspheres, microscopy is generally used to
determine the relative size and shape of the microsphere. Light microscopy is a simple method for imaging, as it requires no special sample preparation. For much higher
resolution, scanning electron microscopy (SEM) can be used; however, since polymers
are nonconducting materials, they are coated with a thin layer of gold to allow for
imaging 60. SEM can also be used to view cross sections of cut microspheres, due to the ability to tilt the sample holder upon imaging 61; this ensures the microsphere is a solid object.
For drug-loaded microspheres, confocal fluorescence microscopy is a useful method for
detecting the distribution of the drug within the microspheres. By staining the drug with
a dye, such as carboxyfluorescein or Nile red, and imaging at different wavelengths (550
nm or 600 nm, respectively), the surface of the microspheres can be analyzed for
dispersal of the drug 60,61. 22
2.3.3 Release
Due to the surface erosion characteristics of polyanhydrides, microspheres of these polymers have predictable, zero-order release kinetics. Polyanhydride microspheres have been shown to provide a sustained release of BSA-FITC, and with a differing ratio of copolymers, can offer a differing release profile 62. Manipulation of the ratio of
monomers used in copolymers is the simplest way to alter the release kinetics 63. Other
factors not as easily controlled can change the way the polyanhydrides release the drug.
Since polyanhydrides are base-catalyzed, a higher pH of the surrounding medium can
lead to a faster release rate of drug. A microsphere with a more porous surface has been
shown to degrade faster than non-porous surfaces. Size and shape of the microsphere can
also alter the release kinetics; in general, smaller microspheres have faster release
kinetics due to an increased surface area to volume ratio. However, drug solubility in
water and fabrication method can affect this trend 64. Drug hydrophobicity also can play a
role in release kinetics; the higher the hydrophobicity, the slower the release rate.
When a drug is incorporated into a microsphere, some of the particles are adsorbed onto
the polymer’s surface rather than fully incorporated into the microsphere. Therefore,
when immersed into a solution, the adsorbed drug immediately dissolves into solution,
resulting in a large, instantaneous release of the drug; this is known as the burst effect.
This phenomenon typically occurs upon incorporation of a hydrophilic drug. An increase
in drug size results in a larger burst 65.
23
2.4 Mechanisms of Immune Response
2.4.1 Innate and adaptive immunity
An immune response can be categorized as one of two types: innate or adaptive. An innate immune response is non-specific, meaning it reacts to a pathogen, regardless of type, in a standard way. It is active and functional at an early age, and will not change nor develop as one ages. In contrast, an adaptive immune response is quite specific, and will develop only after an initial innate response occurs. It will develop over the course of an individual’s life. Memory cells develop as part of the adaptive immune response; therefore, upon re-exposure to prior infectious agent, the adaptive immune response will rapidly recall pathogen-specific effector mechanisms, thereby inducing a stronger reaction than the initial primary immune response. Adaptive immunity occurs over the course of an individual’s life, and becomes stronger upon each subsequent encounter of the antigen 66.
2.4.2 Major histocompatibility complex
The major histocompatibility complex (MHC) is comprised of the molecules responsible for the recognition of foreign antigens. There are two main types of MHC molecules,
class I and class II; their differences lie not only in their structure, but also on the source
of the antigen and the type of cells they provoke.
All nucleated cells express class I MHC proteins; this presentation is known as the
cellular response, and allows for recognition of non-self. When nucleated cells become
infected with a virus or bacteria, the protein of the pathogen is transferred to the cytosol. 24
Both the pathogen’s protein and self proteins are degraded by the proteasome, and the fragments are transferred to the cell’s endoplasmic reticulum. There, the fragments bind to the peptide binding clefts of the class I MHC complex, and are translocated to the plasma membrane where they can now react with the specific T cell receptor.. Since viral proteins are made in the cells cytoplasm, MHC I present these foreign peptides to
CD8+ cytootoxic T cells. Once activated, these cells, form synapses between their TCR
and the peptide MHC I complex and subsequently release the cytotoxins perforin and
granzymes. Perforins form pores within the target cell’s plasma, which allows
granzymes to migrate into the plasma membrane, resulting in cell death 66.
In addition to MHC I, antigen presenting cells, such as dendritic cells or macrophages,
also express class II MHC proteins. Induction of an immune response by this process is
referred to as humoral or antibody response. This process starts with the pathogen being
engulfed by the antigen presenting cell; following proteolytic cleavage, the peptides are retained within the endosome. At this point, the peptides are bound by the class II MHC
molecule that is then translocated to the cell surface. It presents the foreign antigens to
the CD4+ cells. Cytokines released from these activated CD4 helper T cells then either
activate B cells, phagocytes, or other T cells such as CD8+ T cells 66. 25
Figure 2.7 – Major Histocompatibility Complex and Mechanisms of Immune Response
2.4.3 Dendritic cells
The term “dendritic cell” was first documented in 1973, by Steinman and Cohn 67, and are termed because of their branches, or “dendrites” that extend from the cell body’s exterior. Found in the lymph, spleen, skin, and bloodstream, they are a type of antigen presenting cell, meaning they display the foreign antigen on the surface. Immature dendritic cells (DCs) constantly sample the surroundings for pathogens such as viruses and bacteria. Once they have come into contact with such pathogens, they engulf it through phagocytosis, migrate to the lymph nodes, and become mature. Upon maturation, dendritic cells present the protein fragments, which are bound to class II
MHC molecules, on their surface; this leads to a surge of class II molecule production.
As antigen presenting cells, dendritic cells assign directives to both T and B cells. 26
Consequent to antigen presentation, CD4+ T cells recognize this peptide complex, and
either activate B cells to differentiate into antibody secreting plasma cells or stimulate other T cells, such as CD8+ cytotoxic T cells 66,68-71.
Unfortunately, naïve T cells are not activated by MHC molecules alone; they also need a
co-stimulatory signal expressed by an antigen presenting cell 66. Co-stimulatory
molecules are upregulated on antigen presenting cells following stimulation with a
pathogen or pathogen mimicking molecules, and are not specific towards a certain
antigen. Co-stimulatory molecules commonly contributing to T cell activation include
CD80, CD86, and CD40, which bind to CD28 and the C40 ligand on T cells,
respectively. The binding of CD40 to the CD40 ligand is not only important for
activation of the T cell, but is also a necessary signal for T cell proliferation 66,70,71.
Another important molecule recognized by DCs is the intercellular adhesion molecule
(ICAM). ICAM-3, in particular, is found on the surface of naïve T-cells, and has a strong attraction for the DC specific mannose C-type lectin of DC-SIGN (dendritic cell specific
ICAM-3 grabbing non-integrin), also known as CD209 66,72. Found in high levels among
mature dendritic cells, DC-SIGN is hypothesized to be responsible for recruiting and
trafficking resting T cells through binding of ICAM-2 and promoting T cell proliferation
72,73.
Dendritic cells, when activated, also secrete cytokines, which are proteins that affect the
vigor and/or bias of the immune response. As a consequence of IL-12 secretion from
+ DCs, T cells are induced to become TH1 cells. These CD4 T cells typically produce 27
IFN-γ, the cytokine responsible for initially activating macrophages, and TNF-β, which, in addition to activating macrophages, hinders B ells and can prove toxic to some cells.
The release of IL-10 from DCs will induce CD4+ T cells to become TH2 cells that secrete
IL-4, IL-5, IL-6, IL-13, as well as IL-10. The TH2 cytokines are critical for the
differentiation of B cells to inhibit the activation of macrophages, and attenuate
inflammation. In the absence of DC activation (i.e., no co-stimulatory signals), a TH0 response is induced, that is, typically characterized by the secretion of IL-2, as well as IL-
4 and low amounts of IFN-γ 66.
2.5 Adjuvants and Vaccines
Derived from the Latin word ‘adjuvare’ meaning ‘to help’, adjuvants are compounds
used conjunction with antigens to boost the body’s immunological response 74. In 1926,
aluminum-based compounds (aluminum hydroxide or phosphate, primarily) were shown
to enhance the effects of immunizing diphtheria toxoid, and these compounds are still the primary adjuvants used today and the only ones approved for use in humans by the Food
and Drug Administration in the United States 75,76. However, these adjuvants are rather poor at inducing cellular immune responses. Other adjuvants have been investigated, such as Freund’s complete adjuvant and the inclusion of lipopolysaccharides (LPS); however, because of their toxicity, they have been deemed unfit for use in humans 77,78.
A need still remains to find adjuvants with minimal toxicity that is cheap to manufacture, and has no side effects.
28
2.5.1 Novel therapies using polymers
According to the World Health Organization, an estimated 27 million infants are lacking proper immunizations, thus causing them to be more vulnerable to easily avoided diseases 79. Many immunizations require more than one dose to provide effective
immunity; one such example is the vaccine against chicken pox, proven to not be
protective when only one dose is given 80. Additionally, in less developed countries, it
may be difficult for patients to receive more than one dose of a vaccine, due to locations
of clinics or ability to reach one 81. One feasible approach would be to produce a vaccine
that, with one dose, could give controlled release over a pre-determined period of time 82;
a schematic of this is shown in Figure 2.8.
The first controlled release of an antigen from a polymer, poly(ethylene-co-vinyl acetate)
(EVA), was demonstrated in the late 1970s 83,84; however, EVA is not a biodegradable
polymer, meaning it needs to be removed after the antigen has fully released.
Biodegradable systems, involving polymers such as poly(lactic-co-glycolic acid) (PLGA)
and polyanhydrides, have been proven as an effective alternative to alum-based adjuvants
85 and are able to provide a continual and controlled release over a given period of time.
In addition, microsphere formulations are preferred, as there is no surgery required for
implantation; a size of 25 μm in diameter is preferential for delivery through a syringe43.
Ideally, microspheres should be less than 10 μm, which can lead to efficient phagocytosis by dendritic cells, and thus effective antigen presentation to immune cells 86. 29
Figure 2.8 – Conventional release versus controlled release
A single injection of microspheres comprised of PLGA have been used successfully to deliver antigens such as tetanus toxoid 87, diphtheria toxoid 88, influenza virus 89 to induce an equivalent antibody response to that observed following repeated doses of conventional vaccines. The most prominent example of PLGA biodegradable microspheres approved by the FDA, in 1989, and currently on the market, is the Lupron
Depot®. It provides palliative treatment for prostate cancer through release of leuprolide
acetate from the microspheres 90.
Because, PLGA is a bulk-eroding polymer; this can potentially lead to bulk water
penetration, which can initiate water-induced covalent aggregation of the encapsulated
antigen or drug. Another problem that can affect the stability of the encapsulated
compound is the low pH generated following the degradation of PLGA into lactic and
glycolic acid 91, as well as inside the PLGA drug delivery device because of the ability of
water to penetrate the bulk eroding device 92. Such extreme pHs can lead to protein
unfolding, leading to denaturation and possibly irreversible aggregation 93.
Polyanhydrides are thus superior to PLGA in these circumstances; in addition to a higher 30
degradation pH 94, the surface-eroding polyanhydrides prevent water penetration into the
bulk. Polyanhydrides have the advantage of tailored release kinetics, since by simply
changing the chemistry of the monomer can ultimately change the rate of antigen release.
This is favored in vaccine situations, as it allows the number of immunizations to
decrease while providing tailored release kinetics for any application. While the only
application of polyanhydrides thus far is the Gliadel® wafer 42, polyanhydrides for
vaccine applications appear to be very promising. A study with polyanhydride
microspheres demonstrated therelease of tetanus toxoid in vivo with a single injection. In
addition, important immunomodulatory properties of polyanhydride delivery vehicles
were noted; by changing the chemistry of the copolymers, a balanced immune response
could be obtained 85. The ability to modulate the immune response makes the
polyanhydride delivery vehicle versatile, especially when compared to traditional alum-
based vaccines, which have only been demonstrated to provide a TH2 (humoral) response
95.
2.6 Conclusions
Biodegradable polymers have been used as carriers for the controlled delivery of drugs
and proteins for over two decades. Polyanhydrides, in particular, show tremendous
promise for use as drug delivery vehicles. A variety of synthesis techniques allows for
monomers of various structures and thermal properties to be polymerized into
polyanhydrides. Formulating the surface-eroding polyanhydrides into microspheres is
the simplest way to deliver a drug; by altering the chemistry of the backbone or
copolymerizing with other monomers, polyanhydride microspheres can exhibit a wide 31
range of predictable and controlled release kinetics. Since polyanhydrides degrade into dicarboxylic acids, there is no need to removed post-implantation, unlike other biomaterials. In order for polyanhydrides to be deemed a viable candidate for replacement of traditional vaccines, studies need to be conducted to ensure protein stability and an adequate immune response, two important factors in designing delivery devices. 32
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70. Banchereau J, Briere F, Caux C, Davoust J, Lebecque S, Liu YJ, Pulendran B, Palucka K. Immunobiology of dendritic cells. Annu Rev Immunol 2000;18:767- 811. 71. Banchereau J, Steinman RM. Dendritic cells and the control of immunity. Nature 1998;392:245-252. 72. Geijtenbeek TB, Torensma R, van Vliet SJ, van Duijnhoven GC, Adema GJ, van Kooyk Y, Figdor CG. Identification of DC-SIGN, a novel dendritic cell-specific ICAM-3 receptor that supports primary immune responses. Cell 2000;100(5):575- 85. 73. Geijtenbeek TB, Krooshoop DJ, Bleijs DA, van Vliet SJ, van Duijnhoven GC, Grabovsky V, Alon R, Figdor CG, van Kooyk Y. DC-SIGN-ICAM-2 interaction mediates dendritic cell trafficking. Nat Immunol 2000;1(4):353-7. 74. Vogel FR. Adjuvants in perspective. Dev Biol Stand 1998;92:241-8. 75. Glenny A, Pope C, Waddington H, Wallace U. The antigenic value of toxoid precipitated by potassium alum. J Pathol Bacteriol 1926;29:38-39. 76. Gupta RK, Rost BE, Relyveld E, Siber GR. Adjuvant properties of aluminum and calcium compounds. Pharm Biotechnol 1995;6:229-48. 77. Stuart-Harris CH. Adjuvant influenza vaccines. Bull World Health Organ 1969;41(3):617-21. 78. Johnson AG, Gaines S, Landy M. Studies on the O antigen of Salmonella typhosa. V. Enhancement of antibody response to protein antigens by the purified lipopolysaccharide. J Exp Med 1956;103(2):225-46. 79. Global Immunization Vision and Strategy: 2006-2015. Geneva, Switzerland: World Health Organization Department of Immunizations, Vaccines, and Biologicals and UNICEF Programme Division, Health Section; 2005. 80. Lopez AS, Guris D, Zimmerman L, Gladden L, Moore T, Haselow DT, Loparev VN, Schmid DS, Jumaan AO, Snow SL. One dose of varicella vaccine does not prevent school outbreaks: is it time for a second dose? Pediatrics 2006;117(6):e1070-7. 81. Aguado MT, Lambert PH. Controlled-release vaccines--biodegradable polylactide/polyglycolide (PL/PG) microspheres as antigen vehicles. Immunobiology 1992;184(2-3):113-25. 82. Schwendeman SP, Costantino HR, Gupta RK, Langer R. Peptide, protein, and vaccine delivery from implantable polymeric systems. In: Park K, editor. Controlled Drug Delivery: The Next Generation. Washington: The American Chemical Society; 1996. 83. Preis I, Langer RS. A single-step immunization by sustained antigen release. J Immunol Methods 1979;28(1-2):193-7. 84. Langer R. Production of antibodies. Polymers for the sustained release of macromolecules: their use in a single-step method of immunization. Methods Enzymol 1981;73(Pt B):57-75. 85. Kipper MJ, Wilson JH, Wannemuehler MJ, Narasimhan B. Single dose vaccine based on biodegradable polyanhydride microspheres can modulate immune response mechanism. J Biomed Mater Res A 2006;76(4):798-810. 37
86. Eldridge JH, Staas JK, Meulbroek JA, Tice TR, Gilley RM. Biodegradable and biocompatible poly(DL-lactide-co-glycolide) microspheres as an adjuvant for staphylococcal enterotoxin B toxoid which enhances the level of toxin- neutralizing antibodies. Infect Immun 1991;59(9):2978-86. 87. Esparza I, Kissel T. Parameters affecting the immunogenicity of microencapsulated tetanus toxoid. Vaccine 1992;10(10):714-20. 88. Singh M, Singh A, Talwar GP. Controlled delivery of diphtheria toxoid using biodegradable poly(D,L-lactide) microcapsules. Pharm Res 1991;8(7):958-61. 89. Moldoveanu Z, Novak M, Huang WQ, Gilley RM, Staas JK, Schafer D, Compans RW, Mestecky J. Oral immunization with influenza virus in biodegradable microspheres. J Infect Dis 1993;167(1):84-90. 90. Plosker GL, Brogden RN. Leuprorelin. A review of its pharmacology and therapeutic use in prostatic cancer, endometriosis and other sex hormone-related disorders. Drugs 1994;48(6):930-67. 91. Zhu G, Schwendeman SP. Stabilization of proteins encapsulated in cylindrical poly(lactide-co-glycolide) implants: mechanism of stabilization by basic additives. Pharm Res 2000;17(3):351-7. 92. Herrlinger M. Polymerabbau un Wirksttoffreigabe von Poly-DL-Lakid- Formlingen: University of Heidelberg, Germany; 1994. 93. Jaenicke R. Stability and stabilization of globular proteins in solution. J Biotechnol 2000;79(3):193-203. 94. Determan AS, Wilson JH, Kipper MJ, Wannemuehler MJ, Narasimhan B. Protein stability in the presence of polymer degradation products: consequences for controlled release formulations. Biomaterials 2006;27(17):3312-20. 95. Singh M, O'Hagan DT. Recent advances in veterinary vaccine adjuvants. Int J Parasitol 2003;33(5-6):469-78. 38
CHAPTER 3
RESEARCH OBJECTIVES AND ORGANIZATION
3.1 Research Objectives
As demonstrated in the previous chapters, polyanhydrides microspheres have shown great promise for development as vaccine delivery vehicles. The overall goal of this work was to study the effect of polyanhydride chemistry on protein release kinetics, protein stability, and immune activation, leading to rational design of carriers for specific applications. This study focuses on polyanhydrides based on the anhydride monomers of sebacic acid (SA), 1,6-bis(p-carboxyphenoxy)hexane (CPH), and 1,8-bis(p- carboxyphenoxy)3,6-dioxaoctane (CPTEG). The two specific goals of this thesis research project were:
Specific goal 1: Determine how polymer chemistry and fabrication methods affect the release kinetics of proteins from polyanhydride microspheres and the stability of the released protein.
Specific goal 2: Investigate the surface marker expression of murine bone marrow- derived dendritic cells by polyanhydride microspheres and prove that different chemistries can lead to distinct activation pathways of these immune cells.
3.2 Thesis Organization
Specific goal 1, discussed in Chapter 4, will focus on the controlled released of ovalbumin, a model antigen, from polyanhydride microspheres. This study will compare 39
two non-aqueous microsphere fabrication methods. In addition, the chapter will present information on which polyanhydride chemistries preserve the primary structure and epitope availability of ovalbumin.
Specific goal 2, discussed in Chapter 5, will address the effect of polyanhydride chemistry on the activation of dendritic cells. The degree to which stimulation occurs is chemistry dependent, and varies for each surface marker investigated. 40
CHAPTER 4
EFFECT OF POLYMER CHEMISTRY AND FABRICATION METHOD ON
PROTEIN RELEASE AND STABILITY FROM
POLYANHYDRIDE MICROSPHERES
A paper to be submitted to Journal of Biomedical Materials Research, Part B
Senja K. Lopac1,2, Maria P. Torres1, Jennifer H. Wilson-Welder3,
Michael Wannemuehler3, Balaji Narasimhan1,4
4.1 Abstract
The release kinetics and protein stability of ovalbumin-loaded polyanhydrides
microspheres with varying chemistries were studied. Polymers based on the anhydride
monomers sebacic acid (SA), 1,6-bis(p-carboxyphenoxy)hexane (CPH), and 1,8-bis(p- carboxyphenoxy)-3,6-dioxaoctane (CPTEG) were chosen. Microspheres were fabricated using two non-aqueous methods: a solid/oil/oil emulsion technique and cryogenic atomization. Studies found no significant difference in release kinetics of ovalbumin.
Ovalbumin released from microspheres prepared by cryogenic atomization was studied for preservation of primary structure by SDS-PAGE and availability of immunogenic epitopes by western blot. The more hydrophilic polyanhydrides containing CPTEG
1 Department of Chemical and Biological Engineering, Iowa State University 2 Primary researcher and author 3 Department of Veterinary Microbiology and Preventive Medicine 4 Author for correspondence 41
showed more favorable protein stability, preserving both the immunological epitopes and the primary structure.
4.2 Introduction
Biodegradable polymers have been used as carriers for the controlled delivery of drugs and proteins for over two decades. These carriers have the advantages of providing sustained release over long periods of time, well-controlled release profiles, and biocompatibility. The most common biodegradable polymers used in drug delivery applications are polyesters such as poly(D,L-lactide-co-glycolide) (PLGA), polyanhydrides, and poly(ortho esters). A potential draw-back in using the bulk-erodible
PLGA for protein delivery is that the water penetration into the bulk is fast and that the degradation products are fairly acidic; for example, a pH of less than 3 for degradation products 1 and a pH of 2 inside a PLGA drug delivery device 2 have been reported.
Studies have shown that at these pH values, some proteins can undergo denaturation by
unfolding, and in some cases, irreversible aggregation 3. This is problematic for most
proteins because a loss in structure is detrimental to function. In comparison, the pH
values produced by polyanhydride degradation products are much higher, notably 4.2 for
sebacic acid (SA) and 5.5 for 1,6-bis(p-carboxyphenoxy)hexane (CPH) 4. The hydrophobic chemistry of polyanhydrides helps prevent water-induced covalent aggregation of proteins since water penetration into the bulk is negligible; however, non- covalent aggregation due to hydrophobic interactions may result 5,6. This has motivated
research to make these materials less hydrophobic 7,8. This was achieved by
incorporating oligomeric ethylene glycol units, such as triethylene glycol, into the 42
backbone of hydrophobic aromatic polyanhydrides, such as Poly(CPH), leading to amphiphilic polymers with mixed erosion mechanisms and both covalent and non- covalent aggregation of proteins can be reduced 7.
Polyanhydrides were not studied for drug delivery applications until 1983, when Langer
and co-workers reported their potential for controlled drug delivery, based on their
biodegradable properties, and non-toxic and non-mutagenic nature 9. Due to their ability
to surface erode, polyanhydrides exhibit a predictable zero-order release rate, making
them attractive candidates for drug delivery applications 10. Degradation of
polyanhydrides occurs by base-catalyzed hydrolysis of their anhydride linkages, in the
presence of water, to form dicarboxylic acids; their rate of degradation depends upon on
the monomers used 11-13. Poly(sebacic acid) (SA) and Poly(1,6-bis-(p- carboxyphenoxy)hexane (CPH) tablets, for example, degrade in 54 days and 1 year, respectively 14. In contrast, the ethylene glycol containing polyanhydride, Poly(1,8-bis(p-
carboxyphenoxy)-3,6-dioxaoctane) (Poly(CPTEG)) degrades by 80% in 28 days 7. Thus, by combining different anhydride monomers in various ratios, copolymer degradation rates can be tailored for specific applications 15.
Biodegradable polymers are also preferred for parenteral drug delivery systems as there is
no need to remove them following implantation. A size of less than 25 μm is normally preferred for such applications; this size also allows for delivery into the tissue by the use of a syringe and needle 16. Typical methods for microsphere fabrication include hot melt
microencapsulation 17, double emulsion 10,18-22, spray drying 23-24, and cryogenic 43
atomization 20,25-28. In particular, previous research has shown that double emulsion
methods in which water/organic interfaces are present are potentially detrimental for
protein stabilization 21,29-32. Thus, several groups have focused on developing non-
aqueous methods for preparing protein-loaded microspheres. Two commonly used non-
aqueous techniques for fabricating microspheres include solid-oil-oil (S/O/O) double
emulsion and cryogenic atomization (CA); these techniques prevent hydrolysis of the
polymer by eliminating water from the process 33.
The objective of this work is to define the effects of polymer chemistry and fabrication
methods on the release kinetics of proteins from polyanhydride microspheres and the stability of the subsequent released protein. Polymer chemistries based on the anhydride monomers SA, CPH, and CPTEG were chosen (Fig. 4.1). Ovalbumin (ova) from chicken egg white was selected as the model protein. Microspheres were fabricated by solid-oil- oil and cryogenic atomization techniques.
O O O O O O O n O n
O O O O O O O n Figure 4.8 – Chemical structures of polymers used, from top, left to right: poly (sebacic acid), poly (1,6-bis(p-carboxyphenoxy)hexane), and poly (1,8-bis(p-carboxyphenoxy)- 3,6-dioxaoctane), where n represents the number of repeating monomer units.
44
4.3 Materials and Methods
4.3.1 Materials
Albumin from chicken egg whites (ovalbumin), 1,6-dibromohexane, 4-hydroxybenzoic acid, 1-methyl-2-pyrrolidinone, sebacic acid (99%), monoclonal anti-chicken egg albumin (clone Ova-14), rabbit anti-chicken egg albumin, alkaline phosphatase conjugated goat anti-rabbit IgG, and tri-ethylene glycol were purchased from Sigma-
Aldrich (St. Louis, MO). 4-p-fluorobenzonitrile was purchased from Apollo Scientific
(Cheshire, UK). Acetic acid, acetic anhydride, acetone, acetonitrile, dimethyl formamide, ethyl ether, heptane, hexane, methylene chloride, petroleum ether, potassium carbonate, sodium hydroxide, sulfuric acid, and toluene were purchased from Fisher
Scientific (Fairlaw, NJ). Dialysis cassettes, bicinchoninic acid (BCA) assay reagents, and
GelCode blue were purchased from Pierce (Rockford, IL). Low protein molecular weight standards were purchased from BioRad (Hercules, CA). 12% tris-glycine PAGEr
Duramide Precast Gels were purchased from Lonza Bioscience (Basel, Switzerland).
Dow Corning oil, ethanol, and liquid nitrogen were obtained from in-house bulk chemical supplies.
4.3.2 Monomer/polymer synthesis
To produce the CPH monomer, the method described by Conix 34 for synthesizing 1,3-
bis(p-carboxyphenoxy)propane was altered, using 1,6-dibromohexane instead of 1,3-
dibromopropane. Prepolymers for both CPH and SA were synthesized using a method
outlined by Shen 35; copolymers of these compositions and Poly(SA) were synthesized by
melt condensation using a procedure outlined by Kipper and coworkers 36. The CPTEG 45
monomer and polymers of CPTEG and CPH were produced using a technique described by Torres et al 7. Polymers, pre-polymers, and diacids were characterized by 1H NMR,
using a Varian VXR-300 NMR (Palo Alto, CA), to ensure purity; a Waters GPC
(Milford, MA) was also used to measure the polymer molecular weight.
4.3.3 Protein
Ova from chicken egg whites was lyophilized prior to use. Lyophilization occurred by
pumping ova (50 mg) in 50 mM ammonia bicarbonate solution (10 mL) over 400 mL of
liquid nitrogen. The liquid nitrogen was allowed to boil off, and the remaining protein
was placed in a dryer oven overnight; denaturation of freeze-thawed ova at a neutral pH
is highly unlikely 37.
4.3.4 Contact angle measurements
To characterize the relative hydrophobicity of the polymers, contact angle measurements
were carried out. Polymers were dissolved in a 2.5 w/v% solution of tetrahydrofuran (for
Poly(CPTEG) and CPTEG-containing copolymers), or methylene chloride (for
Poly(CPH), Poly(SA), and their copolymers). After filtering solutions with 0.2 µm filters,
the solutions were pipetted onto separate round glass cover slides. After the solvent dried,
more solution was added until a suitable polymer thickness was obtained. To measure the
contact angle, a water droplet was carefully placed on the surface of the polymer film
immediately prior to imaging with a CCD camera. Image J software (NIH, Bethesda,
MD) was used to measure the contact angle. The experiment was performed in triplicate.
46
4.3.5 Microsphere fabrication methods
Two non-aqueous methods were used to fabricate polyanhydride microspheres: solid-oil- oil emulsion and cryogenic atomization. Previous research has demonstrated that these methods are effective at encapsulating and stabilizing proteins 4,20.
4.3.5.1 Solid-oil-oil emulsion
This method was modified from a previously published procedure 21. 100 mg of polymer
and 6 mg of ova were dissolved in methylene chloride. A Tissue-TearorTM homogenizer
(Biospec Products Inc., Bartlesville, OK) was used agitate the solution for one minute.
For the second emulsion, Dow Corning oil and methylene chloride were added while the homogenizer was turned down to 10000 rpm, to allow for thorough mixing during
addition. Again, the homogenizer was used for one minute. The various parameters used
for each emulsion step for the different polymer chemistries are shown in Table 4.1. The solution was added drop wise to a beaker of 200 mL of heptane immersed in an ice bath and stirred for two hours using a Caframo overhead stirrer (Wiarton, Ontario, Canada) set at 300 rpm. Finally, the microspheres were filtered and placed in a vacuum oven overnight to eliminate any residual heptane.
47
Table 4.1 – Parameters for solid/oil/oil double emulsion Methylene Rate for inner Rate for outer emulsion chloride for emulsion homogenization inner emulsion homogenization 20,000 rpm 3 mL oil/4 mL MeCl 20:80 CPTEG:CPH 2 mL 2 3 min 20,000 rpm, 3 min Poly(CPTEG) and 20,000 rpm 4 mL oil/6 mL MeCl 2 mL 2 50:50 CPTEG:CPH 3 min 20,000 rpm, 3 min 30,000 rpm 3 mL oil/4 mL MeCl Poly(SA) 3 mL 2 1 min 20000 rpm, 1 min 30000 rpm 3 mL oil/4 mL MeCl 20:80 CPH:SA 2 mL 2 1 min 30000 rpm, 1 min 20000 rpm 3 mL oil/4 mL MeCl 50:50 CPH:SA 2 mL 2 1 min 30000 rpm, 1 min
4.3.5.2 Cryogenic atomization
Cryogenic atomization, which employs an ultrasonic generator to produce a fine mist,
was also modified from previously published work 21. 100 mg of each polymer was
dissolved in methylene chloride as shown in Table 4.2, with 6 mg of ova. Using a glass
syringe, 20 gauge capillary tube, and. programmable syringe pump (KD Scientific,
Holliston, MA), the polymer solution was pumped over 200 mL of 200 proof ethanol
(frozen by liquid nitrogen), leaving a small layer of liquid nitrogen overlaying the
ethanol. The atomizing mist was provided by an ultrasonic atomizing nozzle (SonoTek
Corporation, Milton, NY). The beakers were placed in a -80 °C freezer for three days to allow the liquid nitrogen to boil off, the ethanol to thaw, and the methylene chloride to slowly be extracted. Afterwards, the microspheres were filtered and placed in a vacuum oven to dry overnight.
48
Table 4.2 – Parameters for cryogenic atomization Methylene Flow rate Wattage chloride 20:80 CPTEG:CPH, 50:50 CPTEG:CPH, 7 mL 3 mL/min 1.5 W and Poly(CPTEG)
50:50 CPH:SA 3 mL 1.5 mL/min 2.5 W
Poly(SA) and 3 mL 3 mL/min 1.5 W 20:80 CPH:SA
4.3.6 Microsphere characterization
A JEOL 840A scanning electron microscope (SEM) was used to determine relative size and shape of microspheres. Microspheres were smeared onto carbon stubs, coated with
200Å of gold, and imaged. Size distribution was performed using Image J software (NIH,
Bethesda, MD).
Figure 4.2 – 50:50 CPH:SA microspheres fabricated by S/O/O (left) and CA methods. Scale bars represent 50 μm. 49
Figure 4.3 – 20:80 CPTEG:CPH microspheres fabricated by S/O/O (left) and CA methods. Scale bars represent 20 μm.
4.3.7 Ova release
Microspheres (10 mg) of each chemistry and fabrication method were suspended in 1mL of phosphate buffer solution (pH 7.4) with 0.01% sodium azide and placed in an incubator at 37 °C and 100 rpm. Release samples were collected two hours after the initial time point, daily for one week, and finally every other day for 30 days. An aliquot of 750 µL was sampled each time and subsequently replaced with 750 µL of fresh phosphate buffer solution; samples were also centrifuged before sampling to ensure no microspheres were removed from the system. In order to assay for the amount of protein release, a bicinchoninic acid (BCA) assay was run on each sample, in duplicate, as described by the manufacturer (Pierce).
After one month of release, the samples were added to 10 kDa molecular weight cut-off dialysis cassettes to determine the amount of protein remaining inside the microspheres.
The left over microspheres were suspended in 3 mL of 17 mM NaOH and sonicated to break up any microsphere aggregates. The exposure to a high pH allows for fast 50
degradation of the polymer, since polyanhydride degradation is base-catalyzed 23. Each release sample was added to dialysis cassettes and incubated for one week at 40 °C and
100 rpm. A BCA assay was run on each sample in triplicate. The total protein loaded into the microspheres was calculated by adding the protein that was released in one month to the protein extracted from the remaining microspheres. The release data is presented as a cumulative fraction of protein released, which is normalized by the total protein loaded into the microspheres.
4.3.8 Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE)
Ova-loaded microspheres (5%) were synthesized using cryogenic atomization. 15 mg of
these microspheres were added to 1 mL of .phosphate buffer solution (pH 7.4), with
0.01% sodium azide, and placed in an incubator at 37 °C and stirred at 100 rpm for 2
hours; 750 μL of solution was removed and replaced with 2.75 mL of 0.01 mM NaOH
solution. This was performed to rid the solution of absorbed protein and to ensure that all
analysis was performed on released protein. The microspheres were added to 10 kDa
molecular weight cut-off dialysis cassettes, placed in 1 L of 0.01 mM NaOH solution,
and incubated for 2 weeks at 40 °C and 100 rpm. After two weeks, the solution was
removed from the dialysis cassette and centrifuged for 10 minutes at 10,000 rpm to
isolate the polymer remaining in the solution. A BCA assay, performed in triplicate, was
used to determine the concentration of ova released from the microspheres.
Using the concentration from the BCA assay, 2 μg of protein from each sample was
placed on a rotovap until completely dry. Samples were prepared under reducing 51
conditions by adding 20 μL of 2-mercaptoethanol sample buffer to each sample and placing samples on a heating block at 90 °C for 10 minutes to break up the disulfide linkages. The gels were made in triplicate, to allow for two to be used for western blot analysis (described below). Gels were run at a constant voltage of 120 V until the dye front reached the bottom. The gel set aside for SDS-PAGE analysis was rinsed with DI water and placed in gel fixative (50% methanol, 36% DI water, 14% acetic acid) overnight. The next day, the gel was stained with Gelcode blue for a few hours and destained with water overnight; the staining process was repeated to obtain a darker stained gel. The gel was placed between cellophane sheets and dried in a jet drier for 2 hours.
4.3.9 Western blot
For the polyclonal western blot analysis, the gels were immediately removed after gel electrophoresis, placed between filter paper and a PVDF membrane, and placed back in the electrophoresis chamber for 3 hours at a constant current of 70 mA. The membranes were blocked with a casein solution of TBST (tris buffer solution with 0.05% Tween, pH
7.6) and milk powder overnight. The following day, the membranes were rinsed in DI water, placed in a 50 mL centrifuge tube, and 12 μL of primary antibody (anti-Ova developed in rabbit) in TBST (1:1000) was added. The membranes were spun for four hours, washed thrice with TBST to remove any unbound antibody, and placed back on the spinner with 12 μL of secondary antibody (anti-rabbit IgG alkaline phosphate developed in goat) in TBST (1:1000). After two hours, the membranes were removed and rinsed thrice with TBST. A colorimetric detection method with napthol phosphate 52
and fast red solution was used to reveal bands. The membranes were air-dried between paper towels.
4.4 Results
4.4.1 Contact angle
In order to assess the hydrophobicities of the various polymers, the contact angles of each of the polymers were measured, as shown in Figure 4.4. As expected, Poly(CPTEG), the most hydrophilic and bulk-erodible polymer with a fast degradation profile (within weeks7), has a contact angle of 29°, the smallest among all the polymer chemistries
tested. In contrast, Poly(CPH), which is the most hydrophobic and surface-erodible
polymer, and takes years to degrade38 has the highest contact angle of 60°. As Figure 4.5
demonstrates, an increase in CPH content within the CPTEG:CPH copolymers results in
an increase in contact angle, which is consistent with an increase in hydrophobicity. In
the surface erodible CPH:SA system, since both CPH and SA homopolymers are
hydrophobic, their copolymers have relatively similar hydrophobicities, as evidenced by
contact angles that are statistically indistinguishable from each other.
Figure 4.4 – Optical microscope images of Poly(CPH) (left) and Poly(CPTEG) (right) used for contact angle measurements 53
80
70
60
50
40
30 Contact Angle
20
10
0 Poly SA 50:50 20:80 Poly CPH Poly CPTEG CPTEG:CPH CPTEG:CPH 50:50 CPH:SA 20:80 CPH:SA Figure 4.5 – Contact angle of all polyanhydride compositions. Error bars indicate standard deviations.
4.4.2 Release of ovalbumin from polyanhydride microspheres
SEM images of 50:50 CPH:SA and 20:80 CPTEG:CPH fabricated by S/O/O and CA methods are shown in Figures 4.2 and 4.3. Size distributions of the CPTEG particles ranged from 4 to 60 μm for S/O/O, with the majority being between 10 and 15 μm in diameter, and 2 to 16 μm in diameter for CA microspheres 20. For CPH:SA, the majority
of the microspheres fell in the 6 to 10 μm diameter range for both fabrication methods.
Figures 4.6 and 4.7 demonstrate the release profiles of ova from Poly(SA), 20:80
CPH:SA and 50:50 CPH:SA copolymer microspheres fabricated by solid-oil-oil and 54
cryogenic atomization fabrication methods. As shown, all these chemistries exhibit near zero-order release kinetics after the initial burst of protein, which is consistent with previous work 21. Each polyanhydride chemistry exhibited a different release rate, but
upon comparing the two methods, this was found to be unrelated to the fabrication
method. However, protein was released from Poly(SA) microspheres for a longer time
(30 days) for solid-oil-oil fabrication than cryogenic atomization (6 days); however, both
systems released 90% of the encapsulated protein at the end of the thirty day time period.
In the CPH:SA system, as the polymer hydrophobicity increased, the release rate of the
protein decreased.
In addition, the choice of method also influences the size of the burst. Microspheres
produced by the S/O/O technique experienced a smaller initial burst of protein. This
could be attributed to the interplay between two phenomena: the rate at which the
polymer precipitated during microsphere formation, and the rate at which the methylene
chloride was extracted into the non-solvent to form the microspheres. Also,
hydrophobicity seemed to have an influence, as a greater difference in initial bursts was
noted for the more hydrophilic Poly(SA) than the copolymers.
55
1 1
0.9 0.9
0.8 0.8
0.7 0.7
0.6 0.6
0.5 0.5
0.4 0.4 Poly SA SOO Poly SA CA 20:80 CPH:SA SOO 20:80 CPH:SA CA 0.3 0.3 Mass fractionof ova released
Mass fraction of ova released 50:50 CPH:SA SOO 50:50 CPH:SA CA
0.2 0.2
0.1 0.1
0 0 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time, days Time, days Figures 4.6 and 4.7 – Release of ova from Poly(SA) and CPH:SA copolymer microspheres using solid/oil/oil double emulsion (left) and cryogenic atomization (right) fabrication techniques. Error bars indicate standard deviations.
Figures 4.8 and 4.9 demonstrates the ova release profiles from Poly(CPTEG), 20:80
CPTEG:CPH and 50:50 CPTEG:CPH copolymer microspheres fabricated by S/O/O and
CA methods. Since Poly(CPTEG) is bulk-eroding, the protein release kinetics are not
directly proportional to the degradation kinetics, but rather depend upon a combination of
degradation, water swelling, and diffusion 7. While one would expect the Poly(CPTEG)
to have the fastest release profile, the 50:50 CPTEG:CPH copolymer actually releases
protein at the same rate (for S/O/O) or slightly faster (for CA) than the Poly(CPTEG) homopolymer. Previous work has shown that even though the mass loss (i.e., erosion) was consistent with the hydrophobicities of Poly(CPTEG) and 50:50 CPTEG:CPH copolymer, the water swelling and polymer degradation rates of both chemistries were very similar7. Our data is consistent with these observations (Figures 4.8 and 4.9). On
the other hand, the 20:80 CPTEG:CPH microspheres released ova at a slower rate, and 56
both fabrication methods correlated in their sustained release profiles by releasing ~50% of protein in 1 month.
Again, the only variation of release kinetics as a result of the fabrication methods is in the initial burst; Poly(CPTEG) microspheres demonstrated the largest difference in burst (8% in S/O/O vs. 42% in CA). This is consistent with the observations reported for the
CPH:SA system, considering Poly(CPTEG) is the most hydrophilic polyanhydride tested.
1 1
0.9 0.9
0.8 0.8
0.7 0.7
0.6 0.6
0.5 0.5
0.4 0.4
20:80 CPTEG:CPH SOO 20:80 CPTEG:CPH CA 0.3 0.3
Mass fractionof ova released 50:50 CPTEG:CPH CA 50:50 CPTEG:CPH SOO Mass fractionof ova released CPTEG CA 0.2 CPTEG SOO 0.2
0.1 0.1
0 0 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time, days Time, days Figures 4.8 and 4.9 – Release of ova from Poly(CPTEG) and CPTEG:CPH copolymer microspheres using solid/oil/oil emulsion (left) and cryogenic atomization (right) fabrication techniques. Error bars indicate standard deviations.
4.4.3 SDS-PAGE
Since the release kinetics studies did not show a significant difference between the solid- oil-oil and cryogenic fabricated microspheres, cryogenic atomized microspheres were used for the protein stability studies due to the ease of scale-up and the simplicity of fabrication.
57
Ova has a tendency to form moisture-induced covalent aggregates 39, which is shown by the presence of characteristic bands between 54 and 97 kDa (lane 2), in addition to the normal Ova band at 48kDa.
The ova released from the Poly(SA), 20:80 CPH:SA, and 50:50 CPH:SA microspheres
(lanes 3, 4, and 5, respectively) show the same higher molecular weight band as the
unencapsulated ova (lane 2), but failed to display the bands at 45 kDa, which is the
molecular weight of non-aggregated ova. This indicates that the CPH:SA system fails to
prevent non-covalent aggregation of the protein, presumably due to hydrophobic interactions; however, considering no bands are displayed at a lower molecular weight, it means these chemistries did not promote hydrolysis or degradation of the protein. This is consistent with the surface erodible nature of these polymers.
The amphiphilic polymers (Poly(CPTEG), 20:80 CPTEG:CPH, and 50:50 CPTEG:CPH)
all showed normal ova bands at 45 kDa as well as the aggregated state, which are both
observed in the unencapsulated ova. The bands for the unaggregated ova became darker with an increase in CPTEG content. Once again, no low molecular weight bands were present, meaning these polyanhydride chemistries did not degrade or cause hydrolysis of the protein. 58
Figure 4.10 – SDS-Page of ova released from microspheres over the course of 2 weeks. Lane 1 – Marker; lane 2 – ova at pH 10; lane 3 – Poly(SA); lane 4 – 20:80 CPH:SA; lane 5 – 50:50 CPH:SA; lane 6 – Poly(CPTEG); lane 7 –20:80 CPTEG:CPH; lane 8 – 50:50 CPTEG:CPH.
4.4.4 Western Blot
Figure 4.11 shows the Western blot analysis conducted on the released protein from each
of the fabricated polyanhydride preparations. Again, ova shows strong bands at both an aggregated (54 to 97 kDa) and unaggregated (45kDa) states. Strong bands are noted for both states for the protein released from each of the CPTEG-containing polymers (lanes
6-8), indicating that the protein epitopes are readily conserved, and that the protein structure is not perturbed. 50:50 CPH:SA preserved the epitopes at the unaggregated ova state, but produced only faint bands for the aggregated protein. Poly(SA), due to the 59
acidic nature of its degradation product, sebacic acid, degrades the protein below detection of the polyclonal western blot; 20:80 CPH:SA also showed a similar effect.
Figure 4.11 – Polyclonal western blot of ova released from microspheres over the course of 2 weeks. Lane 1 – Marker; lane 2 – ova at pH 10; lane 3 – Poly(SA); lane 4 – 20:80 CPH:SA; lane 5 – 50:50 CPH:SA; lane 6 – Poly(CPTEG); lane 7 –20: 80 CPTEG:CPH; lane 8 – 50:50 CPTEG:CPH.
4.5 Discussion
As expected, the higher the hydrophobicity of the polymer, the slower the degradation
rate, as the rate at which water penetrates into the bulk slows in correlation 40. In regards
to the release kinetics of ova, both fabrication methods were consistent with each other.
Cryogenic atomization is a preferential method of preparing microspheres, due to its ease of scale up. Burst profiles are correlated with the polymer hydrophobicity, as the most
hydrophobic microspheres, Poly(SA) and CPH:SA copolymers display the largest burst
regardless of the fabrication method used. This may be attributable to the 60
thermodynamic incompatibility of the protein with hydrophobic copolymers. Therefore, though the actual amount of protein released at the start of the degradation/erosion cycle may vary, the trend of hydrophobicity correlates with the observed burst effect.
When a drug or a protein is incorporated into a microsphere, the drug/protein molecules may be non-uniformly distributed due to thermodynamic incompatibility with the polymer carrier. Therefore, when drug-loaded microspheres immersed into a solution, the drug that is closer to the surface immediately escapes into the bulk solution, resulting in a large, instantaneous release of the drug. Microspheres fabricated with the solid/oil/oil emulsion method exhibited different burst characteristics than CA, with higher initial bursts resulting from the cryogenic atomized microspheres. This could be attributed to the differences in polymer precipitation and solvent extraction rate kinetics for each method. For CA, for example, the polymer solution is sprayed into frozen ethanol with an overlaying layer of liquid nitrogen, which slows the precipitation of polymer in the non-solvent. In addition, the beaker is placed in a -80 °C freezer for three days, over which the methylene chloride is slowly extracted into the ethanol to form and harden the microspheres. Due to the slow rate kinetics of the polymer precipitation and solvent extraction, the protein is also extracted to the surface instead of being evenly dispersed, resulting in a greater burst effect. Microspheres made by the S/O/O method may have more uniformly distributed protein, as the process is conducted in an ice bath, and the extraction occurs over 2 h. Polymer chemistry affected both of these rate kinetics; the more hydrophobic the polyanhydride, the less likely these kinetics had an effect on the initial burst. 61
As previous studies have shown, hydrophobic polymers affect the stability of the protein10; and the data reported here is consistent with the literature. Overall, these
studies indicate that polymer chemistry affects protein stability. The acidity of the SA and 20:80 CPH:SA degradation products affected both the primary structure and recognization of epitopes; only 50:50 CPH:SA fared better at epitope conservation at the unaggregated ova state. Protein structure and epitope availability of ova was better maintained in microspheres fabricated using CPTEG regardless of composition of
method. This is likely due to the amphiphilic nature of the polymer, which has been
shown to be conducive to protein stability 20,41.
These studies are of particular importance when designing protein delivery carriers. As
discussed earlier, the polymer chemistry plays an important role that can be beneficial or
detrimental for proteins. A balance between hydrophobic and hydrophilic environment
(i.e. amphiphilic) is necessary to ensure protein stability, as discussed elsewhere 41.
Drugs such as insulin have important stability implications; it has been proven to undergo structural changes upon release from encapsulated PLGA microspheres, due to the acidic nature of the polymer 42. Insulin can also undergo both covalent and noncovalent
aggregation when introduced to moisture-rich environments 43. Uterocalin, an acute phase protein being investigated for therapeutic use in wound healing applications, has also been theorized to become biologically inactivated upon structural modification 44,45.
Since proteins are well structured and ordered, their integrity must not be upset in order for it to function as intended; thus, it is imperative that the delivery device must not cause any disruptions to the structure. This is especially crucial in the areas of vaccination, 62
where multi-epitope vaccines have been proven more effective than their single-epitope counterparts for diseases such as cancer 46 and AIDS 47; By constructing a multi-epitope antigen, antibodies learn to recognize all epitopes, thus becoming more effective. In addition, mutation or evasion of cells is decreased dramatically; mutant epitopes can even be designed and introduced into the protein structure to avoid such problems 48.
However, if the polymer delivery vehicle is not capable of preserving the availability of epitopes, the multi-epitope antigen is not able to deliver at its full capacity.
4.6 Conclusion
These studies established the effects of polyanhydride chemistry and microsphere fabrication methods on release kinetics and protein stability. Cryogenically atomized polyanhydride microspheres containing CPTEG demonstrated the best preservation of epitopes and primary structure, thus confirming previous work done on amphiphilic environments being the best suited for ensuring protein stability 41.
4.7 Acknowledgements
The authors would like to thank undergraduate students Kristina Staley for help with polymer synthesis and release studies and Andrew Glowacki for help with size distribution analysis. The authors would also like to acknowledge funding from ONR-
MURI, the Hunke Fellowship, and NIH.
63
4.8 References
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32. Castellanos IJ, Cuadrado WO, Griebenow K. Prevention of structural perturbations and aggregation upon encapsulation of bovine serum albumin into poly(lactide-co-glycolide) microspheres using the solid-in-oil-in water technique. J Pharm Pharmacol 2001;53(8):1099-107. 33. Mathiowitz E, Langer R. Polyanhydride microspheres as drug delivery systems. In: Donbrow M, editor. Microcapsules and Nanoparticles in Medicine and Pharmacy. Boca Raton, FL: CRC Press; 1992. p 100-122. 34. Conix A. Poly[1,3-bis(p-carboxyphenoxy)-propane anhydride]. Macromolecular Synthesis 1966;2:95-98. 35. Shen E, Pizsczek R, Dziadul B, Narasimhan B. Microphase separation in bioerodible copolymers for drug delivery. Biomaterials 2001;22(3):201-10. 36. Kipper MJ, Shen E, Determan A, Narasimhan B. Design of an injectable system based on bioerodible polyanhydride microspheres for sustained drug delivery. Biomaterials 2002;23(22):4405-12. 37. Koseki T, Kitabatake N, Doi E. Freezing denaturation of ovalbumin at acid pH. J Biochem 1990;107:389-394. 38. Leong KW, Brott BC, Langer R. Bioerodible polyanhydrides as drug-carrier matrices. I: Characterization, degradation, and release characteristics. J Biomed Mater Res 1985;19:941-955. 39. Stotz C, Winslow SH, ML, D'Souze A, Ji J, Topp E. Degradation pathways for lyophilized peptides and proteins. In: Costantino HR, Pikal M, editors. Lyophilization of Biopharmaceuticals. Biotechnology: Pharmaceutical Aspects. Springer; 2004. 40. Gopferich A. Mechanisms of polymer degradation and erosion. Biomaterials 1996;17:103-114. 41. Kissel T, Li Y, Volland C, Gorich S, Koneberg R. Parenteral protein delivery systems using biodegradable polyesters of ABA block structure, containing hydrophobic poly(lactide-co-glyocolide) A blocks and hydrophilic poly(ethylene oxide) B blocks. J Control Release 1996;39:315-326. 42. Uchida T, Yagi A, Oda Y, Nakada Y, Goto S. Instability of bovine insulin in poly(lactide-co-glycolide) (PLGA) microspheres. Chem Pharm Bull (Tokyo) 1996;44(1):235-6. 43. Costantino HR, Langer R, Klibanov AM. Moisture-induced aggregation of lyophilized insulin. Pharm Res 1994;11(1):21-9. 44. Flower D, North A, Attwood T. Mouse oncogene protein 24p3 is a member of the lipocalin family. Biochem Biophys Res Commun 1991;180:69-74. 45. Playford R, Belo A, Poulsom R, Fitzgerald A, Harris K, Pawluczyk I, Ryon J, Derbi T, Nilsen-Hamilton M, Marchbank T. effects of mouse and human lipocalin homologues 24p3/lcn2 and neutrophil gelatinase-associated lipocalin on gastrointenstinal mucosal Integrity and repair. Gastroenterology 2006;131:809- 817. 46. Harandi A. Immunoplacental therapy, a potential multi-epitope cancer vaccine. Med Hypotheses 2006;66(6):1182-7. 66
47. Lu Y, Xiao Y, Ding J, Dierich M, Chen Y. Multiepitope vaccines intensively increased levels of antibodies recognizing three neutralizing epitopes on human immunodeficiency virus-1 envelope protein. Scan. J. Immunol. 2000;51:497-501. 48. Xiao Y, Lu Y, Chen Y. Epitope-vaccine as a new strategy against HIV-1 mutation. Immunol. Lett. 2001;77:3-6. 67
CHAPTER 5
EFFECT OF POLYMER CHEMISTRY ON IMMUNE ACTIVATION OF
DENDRITIC CELLS
5.1 Introduction
Over the last 200 years, the use of vaccines has proven to be one of the most successful medical interventions in the reduction of disease caused by infectious agents. However, many challenges still remain with regard to fully realizing the health benefits of active immunization programs. Some of these obstacles include the implementation of improved adjuvants, development of single dose vaccines, methods to overcome the poor immunogenicity of recombinant and subunit immunogens, and the ability to rapidly and rationally develop vaccines against emerging pathogens. In this regard, the mechanisms underpinning the effective modulation of cellular and molecular events associated with adjuvant enhancement of immune responses is still unclear. What is well known is that the first step in the immune response process is the activation of antigen presenting cells
(APCs), the most prolific of them being dendritic cells (DCs).
Dendritic cells are responsible for the induction of both T and B cell mediated immune responses. They are found in large numbers in the lymph, spleen, skin, and bloodstream.
Immature DCs constantly sample the surroundings for pathogens such as viruses and bacteria. Once they come into contact with such pathogens, they engulf the pathogen through phagocytosis, migrate to the lymph nodes, and undergo maturation. Upon maturation, DCs present the protein fragments or antigens, which are bound to MHC class II molecules, on their surface; this presentation leads to a surge of class II molecule 68
production. Consequently, CD4+ T cells recognize this MHC antigen complex, and activate B cells to differentiate into antibody secretingplasma cells or other T cells, such
as cytotoxic CD8+ T cells 1-5.
Naïve T cells are not activated by MHC antigen complexes alone; they also need co-
stimulatory signals expressed by APCs for initiation. The requirement of additional co-
stimulatory signals serves to verify the type of T cell being activated, thus minimizes the
chance for generating active T cells against self antigens. Co-stimulatory molecules are
upregulated on APCs following stimulation with a pathogen or pathogen mimicking
molecules, and are not specific towards a certain antigen. Co-stimulatory molecules
commonly presented for T cell activation include the B7 molecules, CD80 and CD86,
and CD40. The B7 molecules bind to the T cell receptor CD28, and CD40 binds to the
C40 ligand on T cells. The binding of CD40 to the CD40 ligand is not only important for
activation of the T cell, but also provides a necessary signal to initiate T cell proliferation
3-5.
Another important molecule recognized by DCs is the intercellular adhesion molecule
(ICAM). ICAM-3, in particular, is found on the surface of naïve T-cells, and has a strong
attraction for the DC specific mannose C-type lectin of DC-SIGN (dendritic cell specific
ICAM-3 grabbing non-integrin), designated in the mouse as CD209 3,6. Found in high
levels on the surface of mature DCs, DC-SIGN is also responsible for recruiting and
trafficking resting T cells through binding of ICAM-2 and promoting T cell proliferation
6,7. 69
Drug and protein delivery vehicles have been investigated in the past for providing a controlled release, parenteral delivery, and being biocompatible; biodegradable polymers, such as poly(D,L-lactide-co-glycolide) (PLGA), are able to provide all three. Due to
PLGA’s bulk-eroding properties and acidic degradation products, both of which can cause protein aggregation, PLGA can be considered detrimental to the protein’s structure and stability 8-10. Polyanhydrides, another class of biodegradable polymers, have been proven superior in these areas, and can be considered viable adjuvants for vaccine delivery. Polyanhydrides degrade by hydrolysis of their anhydride linkages, yielding dicarboxylic acids; this rate of hydrolysis can be altered based on monomer chemistry 11-
13. Considering that the byproducts of polyanhydride degradation are less acidic than
PLGA, there is less denaturing of proteins, thereby preserving the protein’s integrity 14.
Polyanhydride microspheres also have the ability to provide a sustained release of antigen, based on their surface erodible properties as demonstrated by the predictable near zero-order release kinetics of proteins 15. By changing the chemistry of the backbone, polyanhydrides can exhibit a wide range of release kinetics; copolymerization can result in tailored release kinetics for specific applications 16. Due to their surface
eroding nature, which allows no bulk water penetration into the polymeric device, proteins remain stable and not subjected to moisture induced-aggregation or hydrolysis.
However, the possibility of non-covalent aggregation may still occur, due to the
hydrophobic chemistry of polyanhydrides 17,18. Therefore, work has been done to
incorporate hydrophilic oligomeric ethylene glycol units, in particular triethylene glycol
to polymers such as poly(carboxyphenoxy hexane) to make the polymer amphiphilic.
These changes to the chemical structure create a more favorable environment for the 70
proteins, leading to prevention of non-covalent and covalent aggregation19. An in vivo
study with tetanus toxoid loaded polyanhydride microspheres by Kipper et al demonstrated the immunomodulatory properties of polyanhydride delivery vehicles; by changing the copolymer chemistry, a balanced immune response (i.e., both shift in serum antibody profile) was observed 20. The ability to modulate the immune response makes
these polyanhydride delivery vehicles versatile, especially when compared to traditional
21 alum-based vaccines, which are only able to induce/enhance a TH2 (humoral) response .
The purpose of this study was to investigate the surface marker expression of murine bone marrow- derived DCs by polyanhydride microspheres based on the anhydride monomers sebacic acid (SA), 1,6-bis(p-carboxyphenoxy)hexane (CPH), and 1,8-bis(p- carboxyphenoxy)3,6-dioxaoctane (CPTEG), and study the effect of polymer chemistry on
DC activation. E.coli lipopolysaccharide (LPS), a microbial cell wall component known to activate DCs, was used as positive control 22. The surface markers investigated
included MHC II, DC SIGN (CD209), CD86, and CD40, all associated with DC
maturation 3-5; statistical analysis was performed by assessing the changes in the
percentage of CD11c+ DCs expressing a given cell surface marker as opposed to changes
in the mean fluorescence intensity.
5.2 Materials and Methods
5.2.1 Materials
Sebacic acid (99%) and β2-mercaptoethanol were purchased from Sigma-Aldrich (St.
Louis, MO). E.coli lipopolysaccharide (LPS) and rat immunoglobulin were purchased 71
from Sigma (St. Louis, MO). Granulocyte macrophage colony stimulating factor (GM-
CSF) was purchased from PeproTech (Rocky Hill, NJ). Unlabeled mouse IgG was purchased from Pharmingen (Becton Dickinson, Franklin Lakes, NJ). Mouse serum and unlabeled CD36/16 FcγR was purchased from (Southern Biotech, Birmingham, AL).
Methylene chloride was purchased from Fisher Scientific (Fairlaw, NJ). RPMI 1640,
7.5% sodium bicarbonate, penicillin, streptomycin, and L-glutamine were purchased from Mediatech (Herndon, VA). Heat inactivated fetal calf serum was purchased from
Valley Biomedical (Winchester, VA). Unlabeled hamster IgG, Alexa Fluor® 700 anti- mouse CD11c (clone N418), FITC conjugated anti mouse/rat MHC Class II (I-Ek) (clone
14-4-4S), PE/Cy7 anti-mouse CD86 (clone GL-1), allophycocyanin (APC) anti-mouse
CD40 (clone 1C10), PE conjugated anti-mouse CIRE (DC-SIGN 209) (clone 5H10), and corresponding isotypes Alexa Fluor® 700 conjugated Armenian Hamster IgG (clone eBio299Arm), FITC IgG2a K (clone eBM2a), PE/Cy7 conjugated rat IgG2b isotype
(clone KLH/G2b-1-2), APC Rat IgG2a κ (clone eBR2a), and PE-conjugated rat IgG2a
(clone eBR2a) were all purchased from eBioscience (San Diego, CA).
5.2.2 Polyanhydride synthesis
To synthesize 1,6-bis(p-carboxyphenoxy)hexane (CPH) monomer, a technique to produce 1,3-bis(p-carboxyphenoxy)propane was altered. A method described by Shen and co-workers 23 was used to produce prepolymers of CPH and sebacic acid (SA); a method described by Kipper et al. 24 was used to produce copolymers of CPH and SA and
the SA homopolymers by melt condensation. All monomers, pre-polymers, and 72
polymers were characterized by 1H NMR spectroscopy using a Varian VXR-300 NMR to
ensure purity.
5.2.3 Microsphere fabrication and characterization
Microspheres were fabricated by the cryogenic atomization technique, similar to a
method previously published by Determan 25 with minor modifications. Briefly, 100 mg
of polymer was dissolved into an amount of methylene chloride given in Table 1 for each
polymer. The polymer solution was pumped over 200 mL of frozen ethanol (200 proof)
with a programmable syringe pump (KD Scientific, Hollison, MA), glass syringe, and 20
gauge capillary tube and atomized by a ultrasonic atomizing nozzle (SonoTek
Corporation, Milton, NY). Flow rates and wattage settings for each of the polymers are
shown in Table 1. The liquid nitrogen was allowed to boil off over three days in an -80
°C freezer. The solutions was then filtered and dried overnight in a vacuum oven. The
resulting microspheres were imaged by a JEOL 840A scanning electron microscope
(SEM, Tokyo, Japan), to verify spherical shape and size distribution.
Table 5.3 – Parameters for microspheres for cryogenic atomization Methylene Flow rate Wattage chloride Poly CPTEG 50:50 CPTEG:CPH 7 mL 3 mL/min 1.5 W 20:80 CPTEG:CPH 10:90 CPTEG:CPH 4 mL 1.5 mL/min 2.5 W 50:50 CPH:SA 3 mL 1.5 mL/min 2.5 W Poly(SA) 3 mL 3 mL/min 1.5 W 20:80 CPH:SA
73
5.2.4 Isolation and culture of dendritic cells
A method by Lutz et al. 26 was used to obtain bone marrow DCs. Briefly, C3H/HeOuJ
mice (ISU Laboratory Animal Resource breeding colony) were euthanized with CO2 gas and wet with 70% ethanol. The femur and tibia were used to collect the DCs; bone cavities were washed thrice with a total of 3 mL of wash media. Cells were centrifuged, racked, and plated at 4x106 cells per Petri dish with 10 mL of dendritic cell media
consisting of RPMI 1640 supplemented with 1% L-glutamine, 1% penicillin-
streptomycin solution, 1% HEPES, 10% heat inactivated fetal calf serum, 200 µL 1:100
dilution β2-mercaptoethanol, and 10 ng/mL granulocyte macrophage colony stimulating
factor (GM-CSF). Plates were incubated at 37 °C with 5% CO2. On the third day and
sixth days, 10 mL of warm media was added to each Petri dish to supplement DCs. On
the seventh day, the DCs were harvested from the Petri dish and placed in 24-well plates for stimulation.
5.2.5 Stimulation of dendritic cells with polyanhydrides
Polyanhydride microspheres (Poly(SA), 20:80 CPH:SA, 50:50 CPH:SA, 10:90
CPTEG:CPH, 20:80 CPTEG:CPH, 50:50 CPTEG:CPH, and Poly(CPTEG)) were added
to separate wells of immature DCs at concentrations of 0.25 mg/mL, which allowed for a
cell:microsphere ratio of 6:1. For a positive control, 200 ng/mL of LPS was added to
stimulate immature DCs; cells were also left without stimulants to serve as a negative
control. Well plates were placed in the incubator at 37 °C with 5% CO2 for 48 hours.
74
5.2.6 Staining of dendritic cells
After 48 hours of stimulation, the well plates were checked for viability visually using an inverted microscope. DCs were transferred into tubes and placed in the centrifuge (2500 rpm 5 minutes) to rid the cells of the culture media. To ensure specific staining, 250 µL of flow block (100 mL FACS, 1% Rat Immunoglobulin G (IgG), 100 µL unlabeled hamster IgG, 100 µL unlabeled mouse IgG, 2% mouse serum, and 250 µL unlabeled
CD36/16 FcγR) was added to each tube, and tubes were incubated at 4 °C for 1 hour.
The antibodies specific for MHCII (.25 μg), CD86 (.24 μg), CD11c (.12 μg), CD40 (.48
μg), and CD209 (.6 μg) were added, and the cells were incubated on ice for 1 hour; respective antibody and isotype controls were also prepared and added to separate aliquots of cells as negative controls. Following incubation with the antibodies, cells were washed twice by centrifugation with cold FACS/FBS buffer (500 mL phosphate buffer saline, 500 mg sodium azide, 1% fetal calf serum). Cells were resuspended in 200
µL of FACS/FBS, and cells were run immediately on the Becton-Dickinson FACSCanto flow cytometer (San Jose, CA). Propidium iodide (PI), was added just before analysis to differentiate live from dead cells. Data was analyzed using FlowJo flow cytometry analysis software (Tree Star Inc., Ashland, OR).
5.3 Results
5.3.1 Size distribution
Figure 5.1 demonstrates the typical size of polyanhydride microspheres used in the study; ideally, microspheres should be less than 10 μm, which can lead to efficient phagocytosis 75
by dendritic cells, and thus enhance presentation to immune cells 27. Figure 5.2 shows the
percentage of microspheres that are under 10 μm for each polyanhydride formulation.
Figure 5.9 – Poly(SA) (top left), 20:80 CPH:SA (top right), 20:80 CPTEG:CPH (bottom left) and 50:50 CPTE:CPH (bottom right) microspheres. Scale bar for top microspheres represents 2 μm; for bottom microspheres, 5 μm.
90 80 70 60 50 40 30
% of microspheres % of 20 10 0 20:80 50:50 Poly SA CPH:SA CPH:SA 10:90 20:80 50:50 Poly CPTEG CPTEG:CPH CPTEG:CPH CPTEG:CPH Figure 5.10 – Percentage of microspheres smaller than 10 μm for each polyanhydride chemistry incubated with DCs
76
5.3.2 Assessment of surface markers by flow cytometry
After six days in culture, bone marrow cultures proliferated from microcolonies and developed into semi-adherent single cells. Many cells developed hair like-projections, known as dendrites, which were visible with light microscopy (Figure 5.3, left).
Microspheres appeared to be readily phagocytosed by DCs within 24 hours after addition
(Figure 5.3, right). Preliminary results from companion studies have shown that polyanhydride microspheres are contained within these cells in a multitude of intracellular compartments (data not shown; manuscript forthcoming).
Figure 5.3 – Unstimulated bone marrow dendritic cells (left) and after incubation with CPTEG:CPH microspheres (right). The dark areas in the picture on the right are polymer microspheres.
Flow cytometric analysis of the BMDCs revealed that 86% of cultured cells expressed
CD11c on their cell surface (i.e., were DCs), as shown in Figure 5.4; all subsequent analysis were performed by gating on PI-negative, CD11c+ population. 77
Figure 5.4 – Histogram of percent positive CD11 cells (dark line). CD11c isotype control (light line) is shown for comparison.
Significant shifts occurred for MHCII surface marker expression upon maturation of DCs with polyanhydrides. Cells incubated in the presence of the more hydrophobic polyanhydrides (10:90 CPTEG:CPH, 20:80 CPTEG:CPH, 50:50:CPH:SA) exhibited two subpopulations, a broader and taller peak at a dimmer fluorescence, and a smaller peak at a higher fluorescent intensity. For the relatively more hydrophilic polyanhydrides, only a narrow peak at the lower fluorescence appeared.
78
Figure 5.5 – Representative plot of MHCII surface marker expression of CD11c+ cells following stimulation with polyanhydrides microspheres.
In the CPH:SA system, an increase in the CPH content was found to increase CD86 surface marker expression; in the CPTEG:CPH system, an increase in CPTEG was found to cause the same effect. Therefore, while SA did not enhance surface marker expression,
CPTEG proved to be exceptional at inducing this co-stimulatory molecule, achieving expression near that of the LPS positive control. Poly(CPTEG) and 50:50 CPTEG:CPH, the two highest stimulating polyanhydrides, showed a 2.6 and 3.0 fold increase over the non-stimulated negative control, respectively. The only chemistry not proficient at inducing stimulation of DCs was Poly(SA).
79
Figure 5.6 – Representative plot of percentage of CD11c-positive cells also expressing CD86 following stimulation by polyanhydride microspheres.
Overall, a decrease in hydrophobicity decreases surface marker expression of CD40.
Poly(CPTEG) and 50:50 CPTEG:CPH were adept in activating mature DC, showing a
11.4 and 12.3 fold increase, respectively, indicating that DCs are activated by amphiphilic chemistries. Even the lower expressing polymers were able to express more than 2.7 times that of non-stimulated cells; this indicates that polyanhydrides are proficient at activating co-stimulatory molecules. 80
Figure 5.7 – Representative plot of percentage of CD11c-positive cells also expressing CD40 following stimulation by polyanhydride microspheres.
Figure 5.8 demonstrates the results of staining DCs with DC-SIGN CD209. Once again, the amphiphilic chemistry was preferential in expression of DC-SIGN CD209. The more
amphiphilic polyanhydrides Poly(CPTEG) and 50:50 CPTEG:CPH surpassed the LPS
positive control, both tallying more than five times that of the non-stimulated DCs,
compared to 1.4 times for LPS. 81
Figure 5.8 – Representative plot of percentage of CD11c-positive cells also expressing CD209 following stimulation by polyanhydride microspheres.
5.4 Discussion
Immature dendritic cells are phagocytic in nature, and once mature, express high levels of
MHC class II molecules, the adhesion molecule DC-SIGN, and the co-stimulatory
molecules CD86 and CD40 3-5. As demonstrated, all polyanhydride microspheres,
regardless of chemistry, were shown to induce maturation of murine dendritic cells; the
degree to which each surface marker tested was activated was a function of polymer
chemistry. In comparison, PLGA stimulates all the markers as well, but not to the same
magnitude; for the MHCII, CD40, and CD209 markers, it induced the least amount of
stimulation of all polymers tested, and enhanced only slightly more CD86 marker expression than 20:80 CPH:SA (Data not shown).
Stimulation of DCs was enhanced with the amphiphilic chemistries, as shown by
increased surface marker expression of co-stimulatory molecule CD40, the adhesion 82
molecule DC-SIGN, and antigen presenting MHCII molecules. CD86 was the only marker to show a different response, with a trend related to hydrophobicity. Since it has been shown that increase expression of CD40 also up-regulates CD86, there must be more than one pathway activated by these polymers since we observed enhanced stimulation of CD86 by some polymers without the increase in CD40 5,28,29. Another
observation is the high levels of surface expression of CD40 for all the DCs incubated
with polyanhydrides ; even if high levels of MHCII and co-stimulatory molecules are
present, the CD40-CD40L relation is still required for both CD4+ and CD8+ T cell
immunity 30. For MHC class II molecules, high levels were especially noted with the
more hydrophilic molecules, nearing 90% of dendritic cells testing positive with
Poly(CPTEG).
Considering it is well known that the more hydrophobic a polymer, the more
inflammation can occur upon implantation 31, hydrophobicity corresponds to
immunostimulatory properties. Inflammation also can correspond with high levels of the
surface marker CD40 32. All polyanhydrides-incubated DCs were able to express CD40,
which is appropriate, since polyanhydrides are hydrophobic molecules. What is
interesting to note is that the molecules with more hydrophilic entities (CPTEG-
containing polymers) were actually better at expressing CD40 than the more relatively
hydrophobic polyanhydrides.
Polyanhydrides are relatively hydrophobic polymers, and as proven in the results, are
able to stimulate dendritic cells to different degrees. The well-known “danger signal” 83
model by Matzinger may provide some explanation as to why polyanhydrides are able to stimulate dendritic cells. A decade later, an extension of this model was made to include
Janeway’s self/non-self model 33, in which the common thread of hydrophobic molecules
activating ties both models together 34. Therefore, once a hydrophobic molecule is
exposed to innate cells, they become activated, seeing this exposure as a danger signal 35.
LPS, a well known microbial immunostimulatory molecule and a positive control for these experiments, is one such example of this, due to its hydrophobic portions.
Therefore, it is likely the dendritic cells are reacting to the hydrophobicity of the polyanhydrides primarily, and can react slightly differently based on the individual chemistry of each polymer tested. Since Poly(CPTEG), the most amphiphilic tested, is still considered hydrophobic compared to sugars and lipids, it has a balance between the two environments that activates dendritic cells to a higher level than its more hydrophobic counterparts.
The differing levels of maturation by each polyanhydride can serve a great purpose.
Polyanhydrides such as CPTEG with high levels of each surface marker, , are able to induce maturation of all dendritic cells. This makes Poly(CPTEG) a prime candidate for diseases that need a high levels of APC activation for overcoming immune tolerance, such as in cancer therapies. However, it is unclear if the lower stimulating polyanhydrides are capable of inducing a similar magnitude of T cell stimulation. A group of cell displaying CD86 marker with Poly(SA) incubation, for example, may not be the same group expressing CD209. Or, there may be a group of cells present that are still immature. Partially activated cells are adequate enough to provide a tolerance 84
against a certain disease (such as rheumatoid arthritis). In order to achieve a desired level of immune activation, using a cocktail approach of various polyanhydrides may be desired.
5.5 Conclusion
As demonstrated, all the polyanhydride chemistries studied were shown to induce maturation of murine dendritic cells; the degree to which each surface marker tested was activated was a function of polymer chemistry. In addition to supporting Matzinger’s danger model, the maturation of DCs also showed a preference for the more amphiphilic polyanhydrides. Further investigation is needed to elucidate the effect of differing activation on T cell responses. Taken together, polyanhydrides show considerable promise for a wide variety of protein delivery applications by tailoring not just the release kinetics, but also the level of stimulation of key antigen presenting cells.
85
5.6 References
1. Pierre P, Turley SJ, Gatti E, Hull M, Meltzer J, Mirza A, Inaba K, Steinman RM, Mellman I. Developmental regulation of MHC class II transport in mouse dendritic cells. Nature 1997;388(6644):787-92. 2. Winzler C, Rovere P, Rescigno M, Granucci F, Penna G, Adorini L, Zimmermann VS, Davoust J, Ricciardi-Castagnoli P. Maturation stages of mouse dendritic cells in growth factor-dependent long-term cultures. J Exp Med 1997;185(2):317-28. 3. Janeway CA, Travers P, Walport M, Shlomchik MJ. Immunology: the immune system in health and disease. New York: Garland Science; 2005. 4. Banchereau J, Briere F, Caux C, Davoust J, Lebecque S, Liu YJ, Pulendran B, Palucka K. Immunobiology of dendritic cells. Annu Rev Immunol 2000;18:767- 811. 5. Banchereau J, Steinman RM. Dendritic cells and the control of immunity. Nature 1998;392:245-252. 6. Geijtenbeek TB, Torensma R, van Vliet SJ, van Duijnhoven GC, Adema GJ, van Kooyk Y, Figdor CG. Identification of DC-SIGN, a novel dendritic cell-specific ICAM-3 receptor that supports primary immune responses. Cell 2000;100(5):575- 85. 7. Geijtenbeek TB, Krooshoop DJ, Bleijs DA, van Vliet SJ, van Duijnhoven GC, Grabovsky V, Alon R, Figdor CG, van Kooyk Y. DC-SIGN-ICAM-2 interaction mediates dendritic cell trafficking. Nat Immunol 2000;1(4):353-7. 8. Zhu G, Schwendeman SP. Stabilization of proteins encapsulated in cylindrical poly(lactide-co-glycolide) implants: mechanism of stabilization by basic additives. Pharm Res 2000;17(3):351-7. 9. Jaenicke R. Stability and stabilization of globular proteins in solution. J Biotechnol 2000;79(3):193-203. 10. Costantino HR, Langer R, Klibanov AM. Moisture-induced aggregation of lyophilized insulin. Pharm Res 1994;11(1):21-9. 11. Kumar N, Langer RS, Domb AJ. Polyanhydrides: an overview. Adv Drug Deliv Rev 2002;54(7):889-910. 12. Leong KW, D'Amore PD, Marletta M, Langer R. Bioerodible polyanhydrides as drug-carrier matrices. II. Biocompatibility and chemical reactivity. J Biomed Mater Res 1986;20(1):51-64. 13. Laurencin C, Peirrie-Jacques H, Langer R. Toxicology and biocompatibility considerations in the evaluation of polymeric materials for biomedical applications. Clin Lab Med 1990;10:549-570. 14. Determan AS, Wilson JH, Kipper MJ, Wannemuehler MJ, Narasimhan B. Protein stability in the presence of polymer degradation products: consequences for controlled release formulations. Biomaterials 2006;27(17):3312-20. 15. Determan AS, Trewyn BG, Lin VS, Nilsen-Hamilton M, Narasimhan B. Encapsulation, stabilization, and release of BSA-FITC from polyanhydride microspheres. J Control Release 2004;100(1):97-109. 86
16. Eldridge JH, Staas JK, Meulbroek JA, McGhee JR, Tice TR, Gilley RM. Biodegradable microspheres as a vaccine delivery system. Mol Immunol 1991;28(3):287-94. 17. Ron E, Turek T, Mathiowitz E, Chasin M, Hageman M, Langer R. Controlled release of polypeptides from polyanhydrides. Proc Natl Acad Sci USA 1993;90:4176-4180. 18. Tabata Y, Gutta S, Langer R. Controlled delivery systems for proteins using polyanhydride microspheres. Pharm Res 1993;10(4):487-96. 19. Torres MP, Vogel BM, Narasimhan B, Mallapragada SK. Synthesis and characterization of novel polyanhydrides with tailored erosion mechanisms. J Biomed Mater Res A 2006;76(1):102-10. 20. Kipper MJ, Wilson JH, Wannemuehler MJ, Narasimhan B. Single dose vaccine based on biodegradable polyanhydride microspheres can modulate immune response mechanism. J Biomed Mater Res A 2006;76(4):798-810. 21. Singh M, O'Hagan DT. Recent advances in veterinary vaccine adjuvants. Int J Parasitol 2003;33(5-6):469-78. 22. Rescigno M, Granucci F, Citterio S, Foti M, Ricciardi-Castagnoli P. Coordinated events during bacteria-induced DC maturation. Immunol Today 1999;20(5):200- 3. 23. Shen E, Pizsczek R, Dziadul B, Narasimhan B. Microphase separation in bioerodible copolymers for drug delivery. Biomaterials 2001;22(3):201-10. 24. Kipper MJ, Shen E, Determan A, Narasimhan B. Design of an injectable system based on bioerodible polyanhydride microspheres for sustained drug delivery. Biomaterials 2002;23(22):4405-12. 25. Determan AS, Graham JR, Pfeiffer KA, Narasimhan B. The role of microsphere fabrication methods on the stability and release kinetics of ovalbumin encapsulated in polyanhydride microspheres. J Microencapsul 2006;23(8):832-43. 26. Lutz HU, Gianora O, Nater M, Schweizer E, Stammler P. Naturally occurring anti-band 3 antibodies bind to protein rather than to carbohydrate on band 3. J Biol Chem 1993;268(31):23562-6. 27. Eldridge JH, Staas JK, Meulbroek JA, Tice TR, Gilley RM. Biodegradable and biocompatible poly(DL-lactide-co-glycolide) microspheres as an adjuvant for staphylococcal enterotoxin B toxoid which enhances the level of toxin- neutralizing antibodies. Infect Immun 1991;59(9):2978-86. 28. Caux C, Massacrier C, Vanbervliet B, Dubois B, Van Kooten C, Durand I, Banchereau J. Activation of human dendritic cells through CD40 cross-linking. J Exp Med 1994;180(4):1263-72. 29. Flores-Romo L, Bjorck P, Duvert V, van Kooten C, Saeland S, Banchereau J. CD40 ligation on human cord blood CD34+ hematopoietic progenitors induces their proliferation and differentiation into functional dendritic cells. J Exp Med 1997;185(2):341-9. 30. Fujii S-i, Liu K, Smith C, Bonito AJ, Steinman RM. The linkage of innate to adaptive immunity via maturing dendritic cells in vivo requires CD40 ligation in addition to antigen presentation and CD80/86 costimulation. J Exp Med 2004;199(12):1607-1618. 87
31. Hunter R, Strickland F, Kezdy F. The adjuvant activity of nonionic block polymer surfactants. I. The role of hydrophile-lipophile balance. J Immunol 1981;127(3):1244-50. 32. O'Sullivan B, Thomas R. CD40 and Dendritic Cell Function. Critical Reviews in Immunology 2003;23(1&2):83-107. 33. Janeway CA, Jr. Approaching the asymptote? Evolution and revolution in immunology. Cold Spring Harb Symp Quant Biol 1989;54 Pt 1:1-13. 34. Seong SY, Matzinger P. Hydrophobicity: an ancient damage-associated molecular pattern that initiates innate immune responses. Nat Rev Immunol 2004;4(6):469- 78. 35. Matzinger P. Tolerance, danger, and the extended family. Annu Rev Immunol 1994;12:991-1045. 88
CHAPTER 6
CONCLUSIONS AND FUTURE WORK
6.1 Conclusions
The overall objective of this research is to study the effect of polyanhydride chemistry on protein release and stabilization and on immune cell activation. The polyanhydrides studied in this work are based on Poly[1,6-bis(p-carboxyphenoxy)hexane] (CPH),
Poly(sebacic anhydride) (SA), and Poly[1,8-bis(p-carboxyphenoxy)3,6-dioxaoctane]
(CPTEG). Ova was studied as the model protein in these studies. The two specific goals of this thesis are to:
Specific goal 1: Determine how polymer chemistry and fabrication methods affect the release kinetics of proteins from polyanhydride microspheres and the stability of the released protein.
Specific goal 2: Investigate the surface marker expression of murine bone marrow- derived dendritic cells by polyanhydride microspheres and study the effect of polymer chemistry on activation pathways of these immune cells.
Specific goal 1 was addressed in Chapter 4. Two non-aqueous methods of ova-loaded polyanhydride microspheres were studied: solid/oil/oil emulsion and cryogenic atomization. In addition, the stability of the released protein from cryogenic atomized microspheres was evaluated by SDS-PAGE and Western blot analysis to determine the state of the protein’s primary structure and epitope availability. As expected, the more 89
hydrophobic polyanhydrides were shown to provide a slower release rate of Ova. No significant difference was observed between the two methods. The more amphiphilic polyanhydrides, i.e., those containing the CPTEG monomer, preserved both the primary structure and epitope availability. Polymers containing SA were not able to preserve either; this is most likely attributed to the acidity of the degradation product.
Specific goal 2 was investigated in Chapter 5. The expression of the co-stimulatory molecules CD86 and CD40, the adhesion molecule DC SIGN, and MHCII molecule, all known to be expressed on mature dendritic cells, was analyzed after DCs were incubated with polyanhydride microspheres. All polyanhydride microspheres, regardless of chemistry, were shown to induce maturation of murine dendritic cells; the degree to which each surface marker tested was activated was a function of polymer chemistry.
For CD40, DC-SIGN, and MHCII molecules, the stimulation of DCs increased with increasing hydrophilicity. CD86 was the only marker to show a different response, with a trend related to chemistry rather than hydrophobicity. Matzinger’s “danger signal” theory was used to describe why polyanhydrides are able to stimulate DCs 1. Normally,
the hydrophobic portion of molecules is not exposed in the body; instead they aggregate
or form micelles to hide the segment. Therefore, when a hydrophobic segment or
molecule is uncovered, it is considered a sign of danger, and the immune system becomes
activated. Since polyanhydrides are relatively hydrophobic, the DCs may see them as an
alarm signal, and thus mature. An interesting observation is that Poly(CPTEG), the most
amphiphilic of the polyanhydrides tested, showed the most proficiency at activating DCs; 90
therefore, a balance between the hydrophobic-hydrophilic environment must be preferred to activate DCs.
The common thread through these two studies is that the chemistry of the polymer is of utmost importance when designing a protein vehicle, affecting aspects such as protein stability, release kinetics, and dendritic cell activation. The amphiphilic environment proved to be superior to the hydrophobic chemistries in achieving these attributes.
However, a cocktail of microspheres with different polyanhydride microspheres may be the most desired as it would provide control over the release rate and the burst, while also providing the preferred immune response.
6.2 Future Work
6.2.1 Interaction of polyanhydrides and the immune system
As discussed in Chapter 5, different polyanhydride chemistries are capable of stimulating surface markers on dendritic cells (DCs) to a different degree. Toll-like receptors, located on the surface of cells such as DCs, are pattern-recognition receptors; each receptor recognizes different molecular patters. TLR-4, for example, recognizes the bacterial-derived lipopolysaccharide; TLR-2 recognizes not only LPS, but other microbial components such as peptidoglycans. Recognition of these patterns aids in maturation of DCs and allows them to display co-stimulatory molecules essential to an appropriate immune response. While polyanhydrides were found to stimulate the same surface markers as LPS, and in some cases surpass the performance of LPS, it is hypothesized that they operate through the same TLRs. 91
In addition, while surface marker expression was shown by polyanhydrides in Chapter 5, it is unclear as to whether it was one group of cells emitting all the surface markers or various cells emitting markers separately. For polyanhydrides like CPTEG, which stimulated all the markers in ~90% of the cells, nearly all cells attained maturity. For cells stimulated by Poly(SA), for example, it is unclear whether all the cells tested were mature. Since ultimately, dendritic cell maturation will lead to T cell activation, it is important to actually test this by mixing T cells with the mature DCs.
Ideally, microspheres should be less than 10μm, which can lead to efficient phagocytosis by dendritic cells, and thus effective presentation to immune cells 2. As mentioned in
Chapter 5, at least 60% of all microspheres incubated with dendritic cells were of 10 μm in diameter or less. However, it is unclear as to whether smaller microspheres, of the nanometer size, would stimulate DCs to a higher level. Therefore, nanospheres of the same polyanhydride chemistries tested should be incubated with DCs to determine if size has an effect on maturation of DCs. Likewise, the degradation products, which are dicarboxylic acids, could also be responsible for stimulating DCs. The easiest way to test this concept would be to set up the experiment in transwells, in which two wells separated by a membrane. DCs would be placed in one well and polyanhydride microspheres in the second; by selecting a membrane with the appropriate pore size , only the degradation products of the polyanhydrides can be allowed through the membrane.
92
And finally, as mentioned throughout this thesis, is the investigation of cocktails of polyanhydrides. By allowing for a mixture of polyanhydride microspheres, one can tailor the release kinetics, burst of protein, and immune response while ensuring protein stability. Therefore, all of these past experiments should be repeated with mixtures of polyanhydrides with varying chemistries.
6.2.2 Polyanhydride coated drug-eluting stents
These properties of polyanhydrides also make them promising candidates for stents.
Stents are expandable metallic mesh structures or tubes that are utilized to prevent occlusion, or collapse of the body’s arteries or pathways and allow blood and other flow to remain unrestricted 3. One dilemma with using bare metal stents is the potential for
restenosis, or the reoccurrence of stenosis (narrowing of structure), due to tissue
hyperplasia and the regrowth of tumors. Another problem is the formation of neointima,
or thick smooth muscle tissue. Therefore, drugs are often incorporated to prevent
stenosis, inhibit inflammation and neointima formation, and often to block cell
proliferation and prevent thrombosis. A variety of drugs are used in drug-eluting stents,
depending on the application (palliative versus therapeutic, normally). Ideally, a drug
used should inhibit inflammation and prevent reoccurrence of stenosis, as well as block
cell proliferation and prevent thrombosis. Sometimes, more than one drug is
incorporated to receive the best combination of positive activity. Also, coating the drugs
on the stent and applying locally requires a smaller does than would an oral or
intravenously application. By combining with a degradable polymer, a controlled long
term drug release can be achieved, as when the polymer degrades over time, the drug is 93
slowly released. The drug release can be altered simply by changing the chemistry of the polymer. This also means that more drugs can be loaded onto the stent, thereby reducing the number of times the stent has to be replaced.
At this time, there are no known polyanhydride coated stents on the market. However, there are numerous patents and trials using polyanhydrides, specifically using these polymers as the actual stent. These stents are termed “bioabsorbable” rather than simply biodegradable, implying that as the polymer degrades, cells will uptake the monomeric units by phagocytosis 4. In addition to the normal characteristics of biodegradable
polymers, bioabsorbable stents must also have excellent tensile strength to keep restenosis from occurring.
Preliminary cytotoxicity tests were performed to determine the cellular proliferation of
polyanhydrides with human cells, using an MTT assay with the NL-20 human bronchial
epithelial cell line (CRL2503, American Type Culture Collection (ATCC), Manassas,
VA); the cells were incubated overnight with 20:80 CPH:SA and 50:50 CPH:SA
copolymers. Polymer concentrations varying from 10 mg/mL to 0.08 mg/mL were
incubated with 6 x 105 cells/mL; the assay was run with and without cells. Results are
shown in Figure 6.1 For the copolymer 50:50 CPH:SA, an optical density equivalence of
50% live was noted at a concentration of 2.5 mg/mL. The same optical density was
observed at a much lower concentration (0.16mg/mL) for 20:80 CPH:SA; the reason for
the lower concentration could be attributed to the higher in 20:80 a CPH:SA. This study 94
provides promising initial results regarding the suitability of using polyanhydrides as reabsorbable stents.
16 100% Live 14 50:50 CPH:SA 12 20:80 CPH:SA 50:50 CPH:SA w/o cells 10 20:80 CPH:SA w/o cells
8 50 % Live OD ratio 6
4
2
0 10 5 2.5 1.25 0.63 0.31 0.16 0.08 Concentration of polymer, mg/mL Figure 6.1 – MTT assay results
Future work could include tracking the degradation of polymer over the desired time
span, incorporation of suitable drugs into polymer matrix, additional in-vitro studies with
epithelial cells to test biocompatibility, and finally in-vivo studies with the stent to
confirm therapeutic value.
95
6.3 References
1. Seong SY, Matzinger P. Hydrophobicity: an ancient damage-associated molecular pattern that initiates innate immune responses. Nat Rev Immunol 2004;4(6):469- 78. 2. Eldridge JH, Staas JK, Meulbroek JA, Tice TR, Gilley RM. Biodegradable and biocompatible poly(DL-lactide-co-glycolide) microspheres as an adjuvant for staphylococcal enterotoxin B toxoid which enhances the level of toxin- neutralizing antibodies. Infect Immun 1991;59(9):2978-86. 3. Borovetz HS, Burke JF, Ming Swi Chang T. Applications of Materials in Medicine, Biology, and Artificial Organs. In: Ratner BD, Hoffman AS, Schoen FJ, Lemons JE, editors. Biomaterials Science: An Introduction to Materials in Medicine. San Diego: Elsevier Academic Press; 2004. 4. Middleton E, Tipton A. Synthetic biodegradable polymers as medical devices. Medical Plastics and Biomaterials 1998;2:30-39. 5. Abbot. Abbott announce positive one-year results from the world's first clinical trial of a fully bioabsorbable drug eluting coronary stent. 2007.