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SYNTHESIS OF NOVEL BIODEGRADABLE COPOLYMERS:
POST-POLYMERIZATION MODIFICATION OF POLY(L-LACTIDE-
CO-DIALLYLGLYCOLIDE)

presented by

Christopher Paul Radano

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SYNTHESIS OF NOVEL BIODEGRADABLE COPOLYMERS: POST-
POLYMERIZATION MODIFICATION OF POLY(L-LACTIDE-CO-
DIALLYLGLYCOLIDE)

By

Christopher Paul Radano

A DISSERTATION

Submitted to
Michigan State University
In partial fulfillment of the requirement
for the degree of

DOCTOR OF PHILOSOPHY
Department of Chemistry

2003

ABSTRACT
DESIGN OF NOVEL BIODEGRADABLE COPOLYMERS: POST-
POLYMERIZATION MODIFICATION OF POLY(L-LACTIDE-CO-
DIALLYLGLYCOLIDE)
By

Christopher Paul Radano

Poly(lactic acid) (PLA) represents a class of biodegradable polymers, which are
used extensively in biomaterials such as dissolvable sutures, surgical implants, matrices
for drug delivery, and scaffolds for tissue engineering. In order to improve the
biocompatibility of these materials, research groups have designed copolymers of PLA
with the goal of introducing unique functionality and enhancing cell adhesion properties.
Our group has also made progress in this area of research. However, the approach of our
research is different, in that we are employing a post-polymerization modification of a
common biodegradable polymer based on PLA. Using diallylglycolide as a comonomer,
we have synthesized a single biodegradable copolymer, which is able to undergo
functional group transformations through existing methods commonly used in organic
chemistry.

We have demonstrated that olefin cross metathesis and hydroboration/oxidation
transformations are both successful methods of functionalizing PLA. However, the most
successful method of functionalizing PLA was readily achieved via DCC coupling of
various bioactive substrates with our functional copolymer. After synthesizing a series of
copolymers containing bioactive substrates we prepared thin films of the polymers and

examined their ability to support tissue growth, specifically osteoblast growth and

differentiation. The synthesis and characterization of these copolymers as well as the
initial results of physiological experiments using our biodegradable copolymers are

reported.

ACKNOWLEDGMENTS

I would like to acknowledge my advisor Mitch Smith for his guidance during my
graduate career. I value tremendously the training he has imparted to me, teaching me to
carefully think and effectively execute scientific research. I would also like to thank him
for his time and patience, as I had the opportunity to work on a very unique project for
someone who entered as an inorganic chemist. Secondly, I would like to thank Greg
Baker for his guidance, and for his time answering many of my questions. I would like to
thank my committee members, Aaron Odom and Ned Jackson, for helpful discussions
and suggestions. I would also like to thank Professor Michael Mackay and his students
for their time and use of their equipment, as well as Professor Laura McCabe for her
discussions, and her students for carrying out the physiological studies on the polymer
surfaces. Also, I would like to thank former group members Dean, Carl, and Baixin for
their training, J ian-Yang Cho his friendship during the toughest five years of my life, and
colleagues Kin, Dan, and Jim for their friendship and profitable scientific discussions. I
would also like to thank Mei for her very helpful scientific insight and Abbas for his
diligent efforts in “taking over the family business.” I would also like to collectively
thank the MSU faculty and staff for catering to my needs for five years. Finally and most
importantly, I would like to thank my family and my extended family for their prayers,

support, and wisdom for me during my graduate career.

iv

“..F or by him all things were created: things in heaven and on earth, visible and invisible,
whether thrones or powers or rulers or authorities; all things were created by him and for

him. He is before all things, and in him all things hold together..”

TABLE OF CONTENTS

LIST OF ABBREVIATIONS .......................................................................................... viii
LIST OF FIGURES ........................................................................................................... xi
LIST OF TABLES ............................................................................................................ xv
CHAPTER ONE ................................................................................................................. 1
1 Materials Used in Biomedical Applications ............................................................... l
1.1 Engineering Surfaces for Cell Growth ................................................................ 3
1.2 Biodegradable Polymers as Scaffolding Materials ............................................. 6

1 .2. 1 Three-Dimensional Scaffolds ..................................................................... 6

1.2.2 Hybridization of Synthetic Polymers With Natural Polymers .................... 8
1.2.3 Substrate Encapsulation .............................................................................. 9

1.3 Biodegradable Polymers as Drug Delivery Matrices ........................................ 11
1.4 Modification of Poly(Lactic Acid) .................................................................... 14
1.4.1 Endgroup Modification of Biodegradable polymers ................................ 15
1.4.1.1 Polymer Macroinitiation ....................................................................... 15

1.4.1.2 Functional Group Initiation ................................................................... 17

1.4.1.3 Coupling Functional Endgroups ........................................................... 18

1.4.1.4 Backbone Modification of Poly(Lactic Acid) ....................................... 18
1.4.1.4.] Amino Acid-Based Copolymers ..................................................... 18

1.4.1.4.2 Non-Peptidic-Based Copolymers .................................................... 21
CHAPTER TWO .............................................................................................................. 23
2 Modification of Poly(Lactic Acid) ............................................................................ 23
2.1 Monomer Synthesis .......................................................................................... 24
2.2 Polymer Synthesis ............................................................................................. 26
2.3 ‘ Copolymer Synthesis and Characterization ...................................................... 27
2.4 Post-Polymerization Modification .................................................................... 33
2.4.1 Olefin Cross Metathesis ............................................................................ 33
2.4.1.1 General .................................................................................................. 33
2.4.1.2 Synthesis and Characterization ............................................................. 35

2.4.2 Hydroboration-Oxidation .......................................................................... 48
2.4.2.1 General .................................................................................................. 48
2.4.2.2 Synthesis and Characterization ............................................................. 50

2.4.3 DCC Coupling .......................................................................................... 53
2.4.3.1 General .................................................................................................. 53
2.4.3.2 Side-Chain Esterification of Fatty Acids .............................................. 54
2.4.3.3 Side-Chain Esterification of Phosphonic and Amino Acids ................. 64
2.4.3.3.1 Side Chain Esterification of Phosphonoacetic Acid ....................... 64

2.4.3.3.2 Side Chain Esterification of N,N'-diBoc-Lysine ............................ 67
CHAPTER THREE .......................................................................................................... 73
3 Cell Biology on Modified Poly(Lactic Acid) Surfaces ............................................ 73
3.1 ‘ Copolymer Thin Films ...................................................................................... 73
3.2 Osteoblast Cell Shape ....................................................................................... 75
3.3 Osteoblast Proliferation .................................................................................... 76

3 .4 Osteoblast Differentiation ................................................................................. 78

4 Conclusion ................................................................................................................ 80

vi

5 Experimental Section. ............................................................................................... 82

5.1 General Considerations. .................................................................................... 82
5.2 Synthesis ........................................................................................................... 86
6 References ............................................................................................................... 101

vii

ADMET
BBA

br

BrdU
CM

DAG
DCC
dd

DMAP
DMSO
DSC
ECM
EI
EtzO
EtOAc
EtOH
GC/MS
GPC

HPA
Hz

LA
LLA

LRMS ‘

MHz

LIST OF ABBREVIATIONS

Acyclic Diene Metathesis Polymerization
4-tert-Butylbenzylalcohol

broad

Bromodeoxyuridine

Cross Metathesis

ppm

Doublet

3,6-Diallyl-2,5-dioxan-1 ,4-dione (Diallylglycolide)
Dicyclohexylcarbodiimide

Doublet of Doublets

Heat of Fusion (crystallinity)
N,N-Dimethyl-4-aminopyridine
Dimethylsulfoxide

Differential Scanning Calorimetry
Extracellular matrix

Electron Impact

Diethyl Ether

Ethyl Acetate

Ethanol

Gas Chromatography/Mass Spectrometry
Gel Permeation Chromatography
Hydroxyapatite

2-Hydroxypent-4-enoic acid

hertz

Coupling Constant

Lactide

L-lactide

Low resolution mass spectrometry
multiplet

Megahertz

viii

PCL

PDAG

PDI

PDLLA
PDLLA-co-DAG

PEG

PGA

PHB

PLA

PLGA

PLLA

PLLA-co-DAG
PLLA-g-AB
PLLA-g-ArPr
PLLA-g-BocLysPr
PLLA-g-decenal
PLLA-g-decenyloxy-NB
PLLA-g—decenyloxy-TBS

PLLA-g-DPAPr
PLLA-g—HP
PLLA-g-LnPr
PLLA-g-LysTFAPr
PLLA-g-MyPr
PLLA-g—OlPr
PLLA-g-PAPr
PLLA-g-PB
PLLA-g-StPr

9

Number average molecular weight

Weight average molecular weight

Nuclear magnetic resonance

Poly(e-caprolactone)

Poly(diallylglycolide)

Polydispersiy index (MW/Mn)

Racemic poly(lactide)
Poly(D,L-lactide-co-diallylglycolide)
Poly(ethylene glycol)

Poly(glycolide)

Poly(hydroxybutyrate)

Poly(lactide)

Poly(lactide-co-glycolide)

Poly(L-lactide)
Poly(L-lactide-co-diallylglycolide)
Poly(L-lactide-g-allylbenzene)
Poly(L-lactide-g-arachidonic acid propyl ester)
Poly(L-lactide-g-N,N'-di-Boc-lysine propyl ester)
Poly(L-lactide-g—decenal)
Poly(L-lactide-g-decenyl-1 -oxy-(4-nitrobenzene))

Poly(L-lactide-g—decenyl- 1 -oxy-TBS)
Poly(L-lactide-g-diethylphosphonoacetic acid propyl
ester)

Poly(L-lactide-g—hydroxypropane)
Poly(L-lactide-g-linoleic acid propyl ester)
Poly(L-lactide-g-lysine bis(trifluoroacetate) propyl ester)
Poly(L-lactide-g-myristic acid propyl ester)
Poly(L-lactide-g-oleic acid propyl ester)
Poly(L-lactide-g-phosphonoacetic acid propyl ester)
Poly(L-lactide-g—Z-(2-propenyloxy)benzaldehyde)
Poly(L-lactide-g-stearic acid propyl ester)

quartet

ix

RCM
RGD
ROMP
ROP

Ring-Closing Metathesis
Arginine-Glycine-Aspartic Acid
Ring-Opening Metathesis Polymerization
Ring-Opening Polymerization
Room Temperature

singlet

Self-Assembled Monolayer
Tin(II)-2-ethylhexanoate

triplet

tert-Butyldimethylsilyl
Crystallization Temperature
Glass Transition Temperature
Tetrahydrofuran

Melting Temperature
Ultraviolet-Visible

LIST OF FIGURES

Figure 1. Natural biodegradable polymers ......................................................................... 2
Figure 2. Synthetic biodegradable polymers. .................................................................... 2
Figure 3. Correlation of the physiological function of cells with respect to cell shape ..... 6

Figure 4. Illustration of the common methods used to synthesize three-dimensional

scaffolds. ..................................................................................................................... 7
Figure 5. Different three-dimensional structures of PLLA-collagen hybrids. ................... 9
Figure 6. Tissue regeneration through the templation of bioactive substrates. ............... 10

Figure 7, Plasma concentration of a drug as a function of time. (—) Prolonged delivery

system. (—) Traditional delivery system with repetitive administration (*) ............ 11

Figure 8. Schematic illustration of microsphere preparation by the oil-in-water solvent

evaporation technique. .............................................................................................. 12
Figure 9. Synthetic route to poly(anhydrides) via polycondensation. ............................. 13
Figure 10. Block copolymer of poly(lactic acid) and poly(ethylene glycol). .................. 15

Figure 11. Block copolymers of poly(lactide) and poly(ethylene glycol) synthesized by
PEG macroinitiation. (a) -AB- block copolymer (b) -ABA- block copolymer. ...... 16

Figure 12. Endgroup modification of poly(lactic acid) using a functional coinitiator. 17

Figure 13. Copolymerization and modification of poly(lactic acid) using an amino acid
based functional comonomer. ................................................................................... 19

Figure 14. RGD amino acid sequence — Arginine-Glycine-Aspartic Acid. .................... 20
Figure 15. Synthetic route to 2-hydroxypent-4-enoic acid and 3,6-diallylglycolide. ...... 25

Figure 16. IH NMR (300 MHz, CDC13) spectrum of 3,6-diallylglycolide enriched in the
R,S isomer. Ha“ denotes the R,S isomer. Ha,M denotes the R,R and 8,8 isomers. 25

Figure 17. 1H NMR (300 MHz, CDC13) spectrum of poly(diallylglycolide) (PDAG). 27
Figure 18. 1H NMR (300 MHz, CDC13) spectrum of poly(L-lactide-co-allylglycolide). 30

Figure 19. Fox equation describing the glass transition temperature of a random
copolymer. w = weight fraction, and TEA and TgB are the glass transiton
temperatures for the pure homopolymers. ................................................................ 31

xi

Figure 20. Glass transition temperature of poly(rac-lactide-co-diallylglycolide) with
respect to the weight fraction, w, of diallylglycolide. (El) Experimental T35 of
PDLLA-co-AG copolymers. (0) Theoretical Tgs predicted by the Fox equation... 31

Figure 21. (El) Crystallinity (AH) of poly(L—lactide-co—allylglycolide) with respect to the
mole fraction, x, of diallylglycolide. (O) Melting point, Tm, of poly(L-lactide-co-

allylglycolide) with respect to the mole fraction, x, of diallylglycolide. .................. 32
Figure 22. (a) Grubbs’ first generation ruthenium carbene olefin metathesis catalyst. (b)
Grubbs’ second generation ruthenium carbene olefin metathesis catalyst. .............. 33
Figure 23. General scheme for ruthenium-catalyzed olefin cross metathesis. ................ 34

Figure 24. Schematic of polymer-bound ruthenium-catalyzed cross metathesis (CM),
followed by cleavage from the polymer support. ..................................................... 35

Figure 25. Substrates used for polymer-bound olefin cross metathesis to PLLA-co-AG.
................................................................................................................................... 35

Figure 26. Polymer-bound olefin cross metathesis between PLLA-co-AG and 9-decenyl-
1-oxy-(4-nitrobenzene). Conversion of allylglycolide sub-units (isolated yield). 36

Figure 27. 1H NMR (300 MHz, CDC13) spectrum of polymer-bound cross metathesis
between PLLA-co-AG and 9-decenyl-l-oxy—(4-nitrobenzene). (*) Unreacted
PLLA-co-AG. ........................................................................................................... 37

Figure 28. UV-Vis spectrum of polymer-bound olefin cross metathesis between PLLA-
co-AG and 9-decenyl-1-oxy-(4-nitrobenzene) .......................................................... 38

Figure 29. Degradation of linear copolymer PLLA-co-AG and cross-linked copolymer
PLLA-co-AG copolymer. ......................................................................................... 39

Figure 30. (a) Possible polymer ring-closing metathesis reactions. (b) Ring-closing
metathesis of the methyl ester of the ring-opened diallylglycolide performed under
polymer-bound cross metathesis conditions. ............................................................ 41

Figure 31. Percentage of olefin conversion of 9-decenyl-1-oxy-(4-nitrobenzene) with
respect to Grubbs 1St generation catalyst loading ...................................................... 42

Figure 32. (a) Polymer-bound cross metathesis between PLLA-co-AG and allylbenzene.
Conversion of allylglycolide sub units (isolated yield). (b) 1H NMR (500 MHz,
CDC13) Spectrum of polymer-bound cross metathesis between PLLA-co-AG and
allylbenzene. (*) Unreacted PLLA-co-AG. ............................................................. 43

Figure 33. (a) Polymer-bound cross metathesis between PLLA-co-AG and 2-(2-

propenyloxy)benzaldehyde. Conversion of allylglycolide sub-units (isolated yield).
(b) 1H NMR (500 MHz, CDCl3) spectrum of polymer-bound cross metathesis

xii

between PLLA-co—AG and 2-(2-propenyloxy)benzaldehyde. (*) Unreacted PLLA~
co-AG. ....................................................................................................................... 44

Figure 34. (a) Polymer-bound cross metathesis between PLLA-co-AG and 9-decenyl-1-
oxy-tert-butyldimethylsilyl ether. Conversion of allylglycolide sub units (isolated
yield). (b) 1H NMR (300 MHz, CDC13) spectrum of polymer-bound cross metathesis
between PLLA-co-AG and 9-decenyl-1-oxy—t-butyldimethylsilyl ether. (*)
Unreacted PLLA-co-AG. .......................................................................................... 46

Figure 35. (a) Polymer-bound cross metathesis between PLLA-co-AG and 9-decenal.
Conversion of allylglycolide sub units (isolated yield). (b) 1H NMR spectrum of
polymer-bound cross metathesis between PLLA-co-AG and 9-decenal. (*)

Unreacted PLLA-co-AG. .......................................................................................... 47
Figure 36. Synthetic route to poly(lactide-g-hydroxypropane) via hydroboration-
oxidation of poly(lactide-co-diallylglycolide). ......................................................... 50
Figure 37. ‘H NMR (300 MHz,CDCl3) spectra of (a) PLLA-g-HP; poly(lactide-g-
hydroxypropane) and (b) PLLA-co-AG; poly(l-lactide-co-diallylglycolide). .......... 52
Figure 38. General synthetic route to polymer-bound esters using DCC coupling. ........ 54

Figure 39. Fatty acid-modified polymers. (a) medium chain length
poly(hydroxyalkanoate). (b) biodegradable polyester based on glycerol, sebacic
acid, and a fatty acid. (c) polyanhydride containing fatty acid moieties. ................ 55

Figure 40. Synthetic route to saturated and unsaturated fatty acid-modified PLLA using
DCC coupling. .......................................................................................................... 56

Figure 41. ‘H NMR (300 MHz, CDC13) spectra of (a) PLLA-g-MyPr; Myristic acid-
modified PLLA-g—HP and (b) PLLA-g-StPr; Stearic acid-modified PLLA-g-HP. .. 58

Figure 42. 1H NMR (300 MHz, CDCl3) spectra of (a) PLLA-g-OlPr; Oleic acid-
modified PLLA-g-HP and (b) PLLA-g-LnPr; Linoleic acid-modified PLLA-g-HP.59

Figure 43. 1H NMR (300 MHz, CDC13) spectrum of PLLA-g-ArPr; Arachidonic acid-
modified PLLA-g-HP. .............................................................................................. 60

Figure 44. Differential scanning calorimetry of 6 mol % fatty acid-modified copolymers.
Second heating scan at 10°C/min under helium atmosphere. (a) PLLA (6% D-
lactide); (b) PLLA-g-HP; (c) PLLA-g—MyPr; (d) PLLA-g-StPr; (e) PLLA-g-OlPr; (f)
PLLA-g-LnPr; (g) PLLA-g-ArPr. ............................................................................. 61

Figure 45. Simplified synthetic route to the cross-linking of polyunsaturated fatty acid
modified PLLA copolymers. Proposed by Vogl and Blanksby.216 .......................... 62

Figure 46. Differential scanning calorimetry of 6 mol % fatty acid-modified copolymers.
Second heating scan at 10°C/min under helium atmosphere. (a) PLLA~g-LnPr;

xiii

linear polymer sample. (b) PLLA-g-LnPr; cross-linked sample. (c) PLLA-g-ArPr;
linear polymer sample. (d) PLLA-g-ArPr; cross-linked sample. Linear polymer
samples were prepared and stored under a nitrogen atmosphere prior to DSC
analysis. Cross-linked polymer samples were exposed to air during workup and
allowed to cross-link until the polymer became insoluble. ....................................... 63

Figure 47. Synthetic route to PLLA-g-DAPPr; Poly(lactide-g—diethylphosphonoacetic
acid, propyl ester) and PLLA-g—PAPr; Poly(lactide-g—phosphonoacetic acid, propyl
ester). ......................................................................................................................... 65

Figure 48. 1H NMR (300 MHz, CDC13) and “P NMR (120 MHz, CDC13) spectra of (a)
PLLA-g—DPAPr; Diethylphosphonoacetic acid-modified PLLA-g-HP and (b)
PLLA-g-PAPr; Phosphonoacetic acid-modified PLLA-g-HP. ................................. 66

Figure 49. Synthetic strategy for designing biodegradable polymers with amino acid
functionality. ............................................................................................................. 67

Figure 50. Synthetic route to PLLA-g-BocLysPr; Poly(lactide-g—N,N'-diBoc-lysine
propyl ester), and PLLA-g-LysTFAPr; Poly(lactide-g-lysine bis(trifluoroacetate)
propyl ester). ............................................................................................................. 69

Figure 51. 1H NMR (300 MHz, CDC13) spectra of (a) PLLA-g-BocLysPr; N,N'-diBoc-
lysine-esterified PLLA-g-HP and (b) PLLA-g-LysTFAPr; lysine

bis(trifluoroacetate)-modified PLLA-g-HP. ............................................................. 70
Figure 52. 1H NMR (300 MHz, dg-tht) spectrum of PLLA-g-LysTFAPr; lysine
bis(tri fluoroacetate)—modified PLLA-g—HP. ............................................................. 71
Figure 53. Microscope images of osteoblasts cultured for 48 hours on (a) silicon oxide
and (b) 5% phosphonate modified PLLA surfaces. .................................................. 76
Figure 54. Illustration of cell replication incorporating the BrdU fluorescent label. ...... 77

Figure 55. Digital photo images of fluorescent BrdU-labeled osteoblasts on (a) silicon

oxide and (b) 5% phosphonate modified PLLA surfaces. ........................................ 78
Figure 56. Gene expression levels of osteocalcin, runx2, and alkaline phosphatase as a
fimction of osteoblast culture time. ........................................................................... 79
Figure 57. Surface comparison of osteoblast differentiation on different surfaces. ........ 79

xiv

 

LIST OF TABLES

Table 1. Properties of biodegradable polymerslg'2| “ Time to complete mass loss. Time
also depends on part geometry. ................................................................................... 3

Table 2. Patterning sizes typically used to control cell adhesions in materials. ................ 5

Table 3. Copolymerization of rac-Lactide and 3,6-diallylglycolide ([M]/[I] = 300). "
Molar composition of DAG determined by integration of the methylene resonance of
the allyl repeat unit compared to the methine resonance of the lactide repeat unit. b
Molecular weight determined by GPC. " PDI = Polydispersity index (MW/Mn). d
Molecular weight determined by intergration of 4-tert-butylbenzyl endgroup
resonance compared to methine resonances of the lactide and allylglycolide sub-
units. ePoly(diallylglycolide) prepared by solution polymerization in toluene at
80°C for 72 hours using Sn(Oct)2 as the catalyst. .................................................... 28

Table 4; Copolymerization of L-Lactide and 3,6-diallylglycolide ([M]/[I] = 200). "
Molar composition of DAG determined by integration of the methylene resonances
of the allyl repeat unit compared to the methine of the lactide repeat unit.
Molecular weight determined by GPC. c Polydispersity index (MW/Mn).
Poly(diallylglycolide) prepared by solution polymerization in toluene at 80°C for 72
hours using Sn(Oct)2 as the catalyst .......................................................................... 29

I!

Table 5. Saturated and unsaturated fatty acid-modified PLLA copolymers containing 6
mol % fatty acid. a Molecular weight determined by GPC. PDI = Polydispersity
index. ‘ Differential scanning calorimetry of 6 mol % fatty acid-modified
copolymers. Second heating scan at 10°C/min under helium atmosphere.
Functional polymer not exposed to air. 9 Exposed to air until gellation renders the
functional polymer insoluble. f Hydroxylated copolymer starting material used to
couple fatty acids. 3 Independently prepared PLLA containing 6 mol % D units. h
Sample annealed at crystallization temperature. ....................................................... 57

Table 6; Protected and deprotected phosphonic and amino acid-modified PLLA
copolymers containing 6 mol % functional groups. a Molecular weight determined
by GPC. b PDI = Polydispersity index. ‘ Differential scanning calorimetry of
modified copolymers containing 6 mol % functionality. Tg was obtained after the
second heating scan at 10°C/min under helium atmosphere. Molecular weight was
not observed by GPC. d Independently prepared 6 mol % hydroxypropane-modified
copolymer. ................................................................................................................ 72

Table 7.~ Thin films of copolymers spin-coated on l-inch diameter silicon wafers from a
1% (w/w) solution in toluene unless otherwise noted. Spin rate: 2500 rpm, spin
time: 40 sec. “ Mole percent loading determined by 1H NMR spectroscopy.
Determined by ellipsometry. " Samples were prepared by heating above the polymer
melting point and quenching to room temperature to obtain amorphous samples.
Advancing (6m) and receding (0,“) contact angles were measured at a rate of 3

XV

uL/s up to a total volume of 30 uL using Milli-Q grade water. d Copolymer were
dissolved as a 1% (w/w) solution in tetrahydrofuran ................................................ 74

Images in this dissertation are presented in color.

xvi

CHAPTER ONE

1 Materials Used in Biomedical Applications

Material applications in the area of organ replacement represent one of the most
significant developments to medicine. In the area of tissue engineering, the primary goal
is to design a compatible synthetic environment for cells to proliferate, divide, and
differentiate, where the regenerated tissue remains physiologically indistinguishable from
tissue formed under natural conditions. In the area of drug delivery, biologically
compatible materials are used to support the controlled release of a variety of substrates,
thereby inducing the desired physiological effect. The requirements for these
applications are met in various biodegradable polymers.

Biodegradable polymers can be categorized into two classes: Natural and
Synthetic. Naturally occurring biodegradable polymers include, but are not limited to,
polypeptides, dextran, collagen, and chitosan (Figure 1). Many of these natural
biopolyrners constitute what is known as the extracellular matrix (ECM), which provides
the environment from which cells can grow into tissue. Often, this process is predicated
on the cells’ response to the proteins, polysaccharides, and glycosoaminoglycans which
make up the ECM.1 The most common synthetic biodegradable polymers are
poly(glycolide) (PGA), poly(D,L-lactide) (DL-PLA or PDLLA), poly(L-lactide) (L-PLA
or PLLA), poly(lactide-co-glycolide) (PLGA), poly(hydroxybutyrate) (PHB), and poly(e-

caprolactone) (PCL) (Figure 2).

Peptides Collagen
{70 0 OH OH
O
we) (ems (gm
HO n H0 NH
Dextran Chitin Chitosan

Figure 1. Natural biodegradable polymers.

bcoi‘ri row, row

Poly(a-hydroxy acid )5 Poly(hydroxyalkanoate)s Poly(a-caprolactone)
R = H, CH3

Figure 2. Synthetic biodegradable polymers.

Synthetic biodegradable polyesters such as PLA have emerged as an important
class of materials. Ultimately derived from readily available agricultural sources such as
corn and starch, PLA offers an environmentally benign alternative to polymers derived
from petrochemical sources.”4 AS one of a few biodegradable polyesters, PLA breaks
down into lactic acid, a naturally occurring by—product in human metabolism. As a
result, the biomedical field has benefited from these materials. Materials made from
PGA, PLA, PLGA, PHB, and PCL can be used for resorbable sutures,5 medical
implantsif'8 matrices for drug delivery,9'12 and in tissue engineering applications,“"3"'4
where PLA polymers and copolymers have been used as biodegradable scaffolds to
support tissue growth. Progress in both of these fields must be made through the design

of materials that emphasize favorable surface interactions between the material and the

targeted biological entity. Patterned surfaces, natural and synthetic biodegradable

polymers, and the modifications thereof constitute a significant portion of the
advancements made in biomaterial science implemented to optimize surface-cell
interactions in the areas of tissue engineering and drug delivery.

The uses of natural biodegradable polymers are limited primarily by their poor
enzymatic degradation profiles as well as their poor mechanical properties.15 Conversely,
because of their favorable degradation rates and flexibility in their mechanical properties,
development of synthetic biodegradable polymers is the preferred route used in tissue

engineering (Table 1).“5’17

 

Melting Glass Degrad. Density Tensile

 

Polymer point trans. time (g/cm’) strength Elongation Madgflus
type (°C) temp. (°C) (months)” (MP3) ( °) ( ‘0
PLGA Amorph. 45-55 Adjustable 1.30 41.4-55.2 3-10 1.4-2.8

DL-PLA Amorph. 55-60 12-16 1.25 27.6-41.4 3-10 1.4-2.8
L-PLA 173-178 60-65 >24 1.24 55.2-82.7 5-10 2.8-4.2
PGA 225-230 35-40 6-12 1.53 >689 15-20 >69
PCL 58-63 -65 >24 1.11 20.7-34.5 300-500 0.21-0.34

 

Table 1. ‘ Properties of biodegradable polymers.'8'2' " Time to complete mass loss. Time also depends on
part geometry.

1.1 Engineering Surfaces for Cell Growth

Surface chemistry plays an integral role in tissue engineering and drug delivery,
since the surface interaction of cells with any material, natural or unnatural, dictates the
biological response in either tissue engineering or drug delivery processes. The
development of materials addressing this important feature focuses on overcoming the
key challenge of uncontrolled adsorption of cells onto surfaces of materials rendering
them useless, characterized by the nonspecific reactivity with the desired biological
entity.1 The high reactivity and specificity with which biological processes operate, such
as enzymatic pathways and grth cycles, is quite remarkable. Man-made biomaterials

that would be able to mimic the reactivity and specificity seen in Nature would be highly

desirable. Since surface chemistry is of paramount importance, engineering materials
with specific surface properties has been an active area of research.

Surface modification using self-assembled monolayers (SAMS) has proved to be a
promising approach toward the design of new biomaterials. SAMS based primarily on
alkanethiolates attached to gold have been the starting materials for the modification of

many surfaces. Whitesides and Mrksich both have developed alkanethiolate SAMS that

22.23 24.25

present different groups such as oligomeric ethylene glycol units, carbohydrates,

26’” and hydrophobic ligands.28 All of these surfaces show some influence in

peptides,
regulating protein adsorption. For example, the hydrophilicity of carbohydrates and
ethylene glycol units attached to the termini of the hydrophobic SAMS help reduce cell
adhesion to the surface. However, the uniqueness between these two hydrophilic
terminal groups was further observed by Luk and Mrksich as they showed that patterned
hydroxyl-terminated SAMS are able to direct adhesion of 3T3 fibroblasts on the surface
of the material. Patterns using ethylene glycol terminated SAMS began to fade away
afier nine days, whereas mannitol-terminated SAMS retained the pattern integrity for over
twenty-five days.24 This work highlights the potential benefits of SAMS in controlling
cell adhesion and the use of well-defined patterns to regulate material-cell interactions.
Surface patterning is used primarily to invoke a geometrical constraint for cells on
the surface of the material in a way to influence the biology of the system. The types of

materials that have been patterned range from alkanethiolate SAMS,29 proteins,30‘3'

saccharides,32’33

and even minerals such as apatite. A resist patterned CaO-SiOz modified
glass substrate was exposed to an ionic solution containing simulated body fluid. The

ions in the Simulated body fluid induced apatite formation, and after removal of the resist

by dissolution in an organic solvent, the apatite appeared as well-defined patterns on the
glass surface. Moreover, an array of shapes and even alphanumeric characters with lines
as narrow as 2 pm were patterned.34 The size and scale of the pattern depends on the
desired purpose of the pattern. Surfaces can be patterned over a wide range of

dimensions (Table 2).

 

 

Substrate Pattern Size Research Topic
Alkanethiolate SAM 2 x 4 nm2 Study of organization and self-assembly
of ligands35
Alkanethiolate SAM 2-10 pm Measuring cell adhesion strength on
patterns coated with cell integrins.36
Polystyrenes 2-100 gm Cell growth on patterned polystyrene
using specific patterns.37
PLA/PEG 12- 70 m Patterning a biodegradable polymer
containing a biotingylated surface
group.3 .39
Poly(acrylic acid)/PEG- 5 0-1 00 m Regulating cell growth on poly(acrylic
coated Alkanethiolate acid)/PEG-modified SAMS.40
SAM

 

Table 2. Patterning sizes typically used to control cell adhesions in materials.

The result of patterning substrates, in essence, affects the shape of the cells by
controlling adhesion. This interaction has dynamic effects on the conformation of the
cell and consequently influences the cell’s physiological function.4143 Recognizing this,
Whitesides and Ingber have studied the effects of cell geometry on its physiological
function. Cells were seeded onto micro-patterned surfaces of different sizes (5-50 pm),
and the micron-sized surfaces were coated with adhesive proteins. Through control of

cell adhesion and consequently the size of the cell, they studied the effect of cell

geometry with respect to apoptosis (cell death), differentiation, and growth (Figure 3). In
their experiments, endothelial cells died (apoptosis) when their surface area was
SSOOumZ, and cell growth occurred when the surface area was 21500 umz. When the cell
surface area was approximately 1000 umz, both growth and apoptosis cycles were

stopped and differentiation started.

Apoptosis Differentiation Growth

Function

 

 

 

Cell Shape

Figure 3. Correlation of the physiological function of cells with respect to cell shape.

1.2 Biodegradable Polymers as Scaffolding Materials

1.2.1 Three-Dimensional Scaffolds

Intricate designs and structures of synthetic biodegradable polymers such as PGA,
PLA, and PLGA as scaffolding materials have been studied extensively.ls"7’44‘45 The use
of highly porous scaffolding materials as templates for cell growth is desirable because of
the high surface area for cellular attachment as well as the mechanical strength of the

material. The design of porous three—dimensional scaffolds can be achieved by different

methods (Figure 4).

   

Polymer Matrix Polymer Matrix +
Porogen/Solvent/CO2
. 0.0.. ..o.o..
’ ' o o. . ‘ ' o o. .
o . o o o . o o
o o . . o o . .
o . ' o
Porogen/Solvent/CO2 Porogen/Solvent/CO2

Figure 4. Illustration of the common methods used to synthesize three-dimensional scaffolds.

Porogen leaching is a common method used to create pores within the PLA
matrix. In this technique, a suspension of a water-soluble “porogen” such as a salt (salt
leaching) is mixed with the polymer. Once the solvent is removed, the water-soluble
particulates can be washed away leaving a porous, three-dimensional scaffold. Using this
technique, pore size can be easily controlled by the choice of the particulate used.

Mooney and coworkers have used gases such as carbon dioxide as a foaming
agent in the design of PLLA and PLGA porous materials.”47 Carbon dioxide gas diffuses
into the material creating a polymer/gas solution. As the pressure of carbon dioxide is
decreased, macropores form within the matrix. The pressure, gas type, and diffusivity all
determine pore size of the material. Park and Yoon also created foams by combining
porogen leaching and gas forming techniques. PGA/PLA copolymers containing
ammonium bicarbonate salts as a porogenous gas forming agent were immersed into an
aqueous citric acid solution to generate carbon dioxide in situ, generating scaffolds
containing pore sizes varying from 200-500 um.48‘49

A phase separation technique can also be used, where instead of mixing a solid

porogen additive, solvent molecules of a polymer solution can act as the additive.

Removal of the solvent by freeze-drying can result in a variety of polymer foams.
Depending on whether there are liquid-liquid or solid-liquid phase separations between
polymer and solvent during cooling, the overall infrastructure of the material can change.
Ma and coworkers have used this technique to synthesize highly porous nanofibrous
structures of PLLA and PLGA for tissue scaffoldingso’51

Although high surface area and highly porous materials can be prepared, the
limitation is that no Specific reactive sites or functionality is presented. Porous scaffolds
possessing framework functional sites would be beneficial. Nonetheless, porogen
leaching still remains a popular method for the design of three-dimensional porous
scaffolds.

Increased surface area can also be found in fibrous meshes of synthetic polymers.
These meshes can be woven in three-dimensional patterns or designs, perhaps most
commonly as sutures. Meshes of PLA, PGA, and PLGA have all been used to as
materials to aid the regeneration of many forms of human tissue.”56 This method
emphasizes the improved mechanical strength of PGA while utilizing the more
biocompatible surface properties of the PLLA or PLGA coating. Langer and Mooney
designed a bonding technique by spraying an atomized solution of PLLA or PLGA onto
PGA woven fibers.57 In Spite of the increase strength of the material, the strength is still
dependent on the efficiency of the connection between the PLLA and PGA layers.

Moreover, the lack of functionality still remains a limitation to such materials.

1.2.2 Hybridization of Synthetic Polymers With Natural Polymers

Although use of some biodegradable scaffolds has shown much promise in tissue

growth applications, with control over their degradation properties and cell-scaffold

interactions, they are deficient Since they lack functionality, and possess an overall
hydrophobic character. Hybrid materials based on PLA, PGA and a natural biologically
functional polymer, such as collagen, provide a material that possesses increased
biocompatibility and mechanical strength over just the PLA or collagen alone (Figure 5).
For example, PLLA-coated collagen fibers showed a 200% increase in tensile strength
and modulus over the PLLA alone.58 The inverse are also made, such as PLLA sponges
coated with collagen.59 In fact, collagen coated-PLLA Sponges interact more with mouse

fibroblast L929 cells than did the PLLA sponges alone.15

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

0
CC I ‘ ff 2 TD
<=( («:3 :r t: r: u:
Fibers Fibrous mesh Sponge
(I) PLLA/PGA
CO Collagen

Figure 5. Different three-dimensional structures of PLLA-collagen hybrids.

1.2.3 Substrate Encapsulation

A similar form of hybridization can be achieved using more specialized substrates
which possess a more specified type of reactivity. The biocompatibility of a polymer can
be improved by the blending or encapsulation of proteins,”62 growth factors,“66 or
minerals,50‘5"67‘68 directly with PLA or PGA (Figure 6). These methods all Show

improved activity in tissue regeneration when compared to PLA and PGA polymers and

copolymers alone.

DUDE] 13:11:11:
DUDE] [31:11:13
ENDED ’ 8CD —’ 5
DUDE] DE] "

 

 

 

 

Polymer Scaffold Polymer Scaffold Enhanced Cell Polymer degradation
Modified with a Growth onto Tissue regeneration
Bioactive substrate Modified Polymer

Figure 6. Tissue regeneration through the templation of bioactive substrates.

Growth factors such as bone morphogenic proteins (BMPs), for example, are used
precisely for this purpose. The growth factors used in combination with a defined
scaffold create a hybrid material which carries out a specific physiological role. For
example, Hollinger and Winn had reported that using a porous collagen-coated PLA
scaffold embedded with BMP-2 growth factors, produced more bone than the PLA-
collagen scaffold without the BMP-2.69 Peter and Mikos synthesized microspheres of
PLGA/PEG (95/5) containing transforming growth factor-Bl (TGF-Bl), which 70
subsequently improved both proliferation and differentiation. Mooney et a1. and Patrick
independently prepared functional porous PLGA scaffolds using a gas foaming process
containing embedded vascular endothelial growth factor (VEGF). The materials
ultimately led to increased proliferation of human umbilical vein endothelial cells."72
Ma has capitalized on the high porosity of a three-dimensional scaffold and doped
hydroxyapatite in order to enhance osteoblast growth.50 While not part of the framework

of the biomaterial itself, the growth factors and minerals offer a way to influence cell

proliferation and differentiation.

10

1.3 Biodegradable Polymers as Drug Delivery Matrices

Biodegradable polymers in drug delivery are largely based on PLA, PLGA or
PCL copolymers. Many approaches have been taken to obtain the optimal drug delivery
device. The use of biodegradable polymers is effective because the by—products are
safely eliminated in vivo. In drug delivery, a polymeric system can deliver a

pharmaceutical agent, via a controlled or systematic release (Figure 7).73

 

 

 

 

 

A
OD
g Toxic
o ... ...........
t: / \ / \ \
3% l \ l ‘\ l/ \'
b / \ \ / \
CI
8 / I \ / \ Therapeutic
8. / \ / \ / \
E: / \ ’ \ / \
g ...... / ........................ \ .............. l ............ I f ..... ‘
£3 \ /
o. \ /
\ / °
_, \ / Ineffective
Time

Figure 7. Plasma concentration of a drug as a function of time. (—) Prolonged delivery system. (-—)
Traditional delivery system with repetitive administration (*).

The therapeutic agent can be delivered as the polymeric system undergoes slow
degradation, via diffusion of the agent through the matrix.”75 Polymeric drug delivery
systems can be derived from natural polymers such as polysaccharides,76 chitosan,77
collagen76 or synthetic polymers which include PGA, PLA, PLGA, or PCL.”82

Typically, microspheres of these polymers are prepared using an oil-in water evaporation

technique (Figure 8).73

11

()0

U U

 

 

 

 

 

 

Addition Sonrcatron
, ,

__—————p O O O O
. , 0 ~ 'O
O O O O
O O
O O
Polymer, Drug, and
Organic Solvent Water and Stabilizer
Precipitation
0 O
o o. o
O. O . O
O O O
O O
Microspheres

Figure 8., Schematic illustration of microsphere preparation by the oil-in-water solvent evaporation

technique.

Copolymers which combine natural and synthetic polymers are also very common
as matrices. Li and coworkers have prepared comb-like copolymers containing a dextran
backbone with PGA and PLA oligomeric side chains.83 These copolymers are good drug
delivery matrices for the continuous release of fluorescein isothiocyanate labeled dextran
and bovine serum albumin (BSA), with accelerated degradation not seen in the pure PGA
or PLA. Since polyesters like PGA and PLA are very hydrophobic, hydrophilic domains
are generally incorporated in some way. Copolymers and blends of PLA and
polyethylene glycol (PEG) are also commonly used to deliver proteins and drugs.”87 For
example, Yeh has prepared microspheres from blends of PLGA, PEG and insulin. Stable
microspheres (3-8 um in diameter) provided a steady release of insulin over the course of

28 days.88 PLA-chitosan blendsgg'gl and PLA-peptide blends92 are some of the other

12

polymeric systems which have been prepared as microspheres for the delivery of proteins
and drugs.

Incorporating hydrophobicity in polymeric matrices can also improve drug delivery
processes by promoting surface erosion as opposed to internal degradation. Hydrophobic
regions maintain the integrity of the matrix by protecting the interior from the aqueous
media while the hydrophilic regions interact with the aqueous media to promote surface
erosion. In order to address this issue, Langer prepared polyanhydrides using a melt

polycondensation technique under reduced pressure (Figure 9).”95

Ho’lOLRjiou + fioi ‘—f_’ \(‘OfkaLI\ R: @OMOQ-

-CH3C02H
(p-carboxyphenoxypropane)

bi

(sebacic acid)

 

Figure 9. Synthetic route to poly(anhydrides) via polycondensation.

Polyanhydrides have become attractive drug delivery matrices because of the
variability in their properties. For example, aliphatic polyanhydrides can degrade within
days to weeks, while aromatic polyanhydrides may take several months to years.96 This
interesting feature can be exploited to design materials to fit the desired degradation
profile. ,Polyanhydrides derived from p-carboxyphenoxypropane and sebacic acid have
generated the most pharmaceutical interestf’m‘w’98 However, has been introduced by
Domb and coworkers introduced other functionality by incorporating fatty acids and
other hydrophobic groups through end group modification and copolymerizations with

polyanhydrides.99'103

13

These fatty acid based polyanhydrides have been used as carriers for anesthetics

98,104 while the FDA approved poly(p-

and chemo-therapeutic agents,
carboxyphenoxypropane-co-sebacate) has been used as an implant to deliver drugs for
brain cancer treatments.105 A PLA-coated hybrid polyanhydride sheet loaded with
heparin resulted in the controlled local delivery of heparin in laboratory rats over the
course of 20 days, as opposed to 4 days for the uncoated heparin loaded polyanhydride
sheet.106

Natural and synthetic biodegradable polymers have numerous advantages as both
tissue engineering scaffolds and drug delivery matrices. Often, copolymers or
hybridizations of these two classes of polymers are used in complementary fashion.

Simple adaptations of biodegradable polyesters such as PLA and PGA with different

forms of functionality have been explored.

1.4 Modification of Poly(Lactic Acid)

Due to their biocompatibility, PLA, PGA, and PLGA, are the most common
materials from which biodegradable tissue scaffolds are madam“lo However, a
significant drawback to using the materials of PLA and PGA in tissue engineering is the
lack of control in cell adhesion, where the hydrophobic polymer (e.g. PLA) will adhere
too strongly to cells and cell adhesion proteins, thereby decreasing its bioactivity.m’”2
This exposes a core problem for researchers who use biodegradable polymers such as
PLA in the areas of tissue engineering and drug delivery. While the biocompatibility and
biodegradability of these polymers may be adequate enough to actually grow tissue or

release drugs, the specificity with which cells bind is minimal. The current solution to

this problem is to design firnctionalized copolymers to counteract the lack of specific

14

functionality in the PLA or PGA polymers or copolymers. Since the only site available
for chemical modification on poly(a-hydroxy acids) are the endgroups, the synthesis of
block and graft copolymers, random copolymers using functional monomers, and

coupling between a polymer and bioactive substrate must be pursued.
1.4.1 Endgroup Modification of Biodegradable polymers

1.4.1.1 Polymer Macroinitiation

Both diblock1 ‘3 " '5 and triblockgb'l ”"118 copolymers of PLA/PGA and
poly(ethylene glycol) (PEG) have been prepared in order to create biomaterials with both

hydrophobic and hydrophilic domains (Figure 10).

o
W): Nb
oooooooooooom

Hydrophobic Hydrophilic
PLA PEG

Figure 10; Block copolymer of poly(lactic acid) and poly(ethylene glycol).

The PEG segments in the copolymers serve as a means to better control the protein and
cell adhesion while maintaining the biocompatibility of the material.112 The synthesis of
these block copolymers is achieved by polymerizing lactide using the hydroxylated chain

end of the PEG unit as a macroinitiator (Figure 11).

15

(a) 10 O
0 0I o
H3CO/(’\/OIF/\OH & H3CO/(TVOIEAO’6HOWAO’):
0

PEG

Moooooooooooo

O O O O
raver. ———-~ toflofiotvottfofisem

PEG
OOOOOOOOOOW 0000000000

Figure 11. Block copolymers of poly(lactide) and poly(ethylene glycol) synthesized by PEG
macroinitiation. ((1) -AB- block copolymer (b) -ABA- block copolymer.

Recently, Mikos has shown the use of a block copolymer, PLA-b-PEG, as a tissue
scaffold for the differentiation of bone marrow cells. The copolymer was synthesized
through the ring-opening polymerization of D,L-lactide with tin(II)octanoate as the
catalyst and poly(ethylene glycol)-monomethyl ether as the macroinitiator. The PEG
domain Serves to regulate cell adhesion of adsorbed serum proteins (e.g. fibronectin),
thus resulting in differences in cell shape and increased osteoblast differentiation as
compared to PLA, PLGA and tissue culture polystyrene dishes.'”‘115 This result
emphasizes that controlled adhesion of serum proteins to the biomaterial affects the shape
and conformation of the cell, thereby influencing differentiation. The hydrophilicity of
the PEG domains can also be used as a way to synthesize hydrogels.l '2 These hydrogels
are copolymers of PLA and PEG which are then photochemically cross-linked through

119.120

reactive endgroups. These gels are stronger materials than linear block copolymers,

providing a porous material that does not dissolve, but swells in aqueous systems.

16

Terminal hydroxyl groups of poly(hydroxybutyrate) (PHB),121

122 124

poly(dimethylsiloxane), and poly(e-caprolactone),'23 poly(vinylalcohols), and

poly(saccharides)‘25

can also be used as macroinitiators to prepare block copolymers and
graft copolymers of lactide. These copolymers offer applications in both tissue scaffolds

and as drug delivery matrices.

1.4.1.2 Functional Group Initiation

The hydroxyl groups of more specified substrates can be used as coinitiators with
tin(II)-octanoate to prepare polymers with specific endgroups (Figure 12).

\ Substrate /

\\
M‘

\IAO Sn(Oct)2 ”on(00
O Bloacli>
UI/‘\ Substrate

O

 

Figure 12. Endgroup modification of poly(lactic acid) using a functional coinitiator.

Stupp and coworkers incorporated a cholesterol endgroup in PLA simply by using

'26 The cholesterol

cholesterol itself as the initiator in the polymerization of lactide.
endgroup takes advantage of the cholesterol-cell membrane interaction in fibroblasts.
The cholesterol-filnctionalized PLA affected the overall shape fibroblasts and improved
their adhesion compared to the PLA control.‘27 To further demonstrate the utility of
endgroup modification, Gardella and coworkers functioalized the endgroups of PLA with
fluorinated alkyl chains through both post-polymerization coupling reactions between
flouroalkyl chains and the PLA as well as the use of a fluorinated alcohol as a

128

coinitiator. The fluorinated segments can be used to regulate the surface adhesion

properties of the material. Groups with multiple functionality containing groups such as

17

primary amines, diols, and carboxylic acids are incompatible with the catalyst, thus

limiting their use in functional endgroup initiation.

1.4.1.3 Coupling Functional Endgroups

It is also possible to use standard coupling chemistry to link functional endgroups
to biodegradable polymers. Utilizing a DCC coupling procedure, galactosyl derivatives
have been successfully coupled to carboxylic acid endgroups of PLA to prepare

29 Langer has also used DCC

microspheres suitable for drug delivery applications.l
coupling to link polylysine directly with PLA.130 Coupling chemistry offers a mild way
to introduce diverse functionality, however the only modification site on PLA is the

endgroup, which means that the incorporated functionality is limited to coupling at this

site.
1.4.1.4 Backbone Modification of Poly(Lactic Acid)

1.4.1.4.] Amino Acid-Based Copolymers

In order to expand the applications of these biomaterials, two different methods
have been developed to modify the structure of PLA using amino acid chemistry. The
first is the copolymerization of lactide with a morpholinedione-based comonomer
containing amino acid moietiesm'133 For example, Langer and coworkers synthesized a
functional comonomer bearing a protected lysine.132 Copolymerization of the protected
comonomer with L-lactide followed by the deprotection of the copolymer resulted in low
loadings (1%-5%) of lysine-functionalized copolymer. The lysine is incorporated into
the backbone of the polymer, and sites for adhesion are distributed along the polymer

chain, where further chemical modification can be achieved (Figure 13). Chemical

18

attachment of amino acids is interesting in that it can serve as a model for more specific
binding of proteins.

The second approach to modifying biodegradable copolymers with amino acids is
through direct coupling of the polymer with amino acid groups.'3°’I34 One specific type
of functionality that is important in the area of cell biology and cell adhesion is found in

RGD (arginine-glycine-aspartic acid) containing peptide sequences.
Protecting Group

0 ) 1. Copolymerization
‘NH 2. Deprotection
O-
O

Lactic Acid Aminll :‘\Cltl

 

 

Protected Morpholinedione
Comonomer

CH3 H 0
Functional Tissue Scaffold e CNN
0 R

Figure 13. Copolymerization and modification of poly(lactic acid) using an arrrino acid based functional

 

 

comonomer.

RGD is a specific sequence of amino acids which shows activity in binding to
integrins within a cell’s membrane, thereby affecting the conformation of the cell, and

”5"37 While using a peptidic functional comonomer

ultimately its function (Figure 14).
that incorporates the whole RGD sequence is not practical, modifiying the backbone of

the polymer using coupling chemistry is the preferred direction that research groups have

taken. In fact, Langer and coworkers have also synthesized a biodegradable copolymer

containing the RGD sequence,138 which involved the transformation of their previously
synthesized PLA-co-lysine copolymer.132 In this study, the side chain amino groups (-
NHz) from lysine incorporation react with carbonyldiimidazole, which is followed by
coupling of the 5-unit RGD-containing peptide. Other research groups have also
performed such RGD modifications on acrylate-based polymers and ethylene glycol

based polymers using an RGD coupling procedurem'143

R G D
A A A

r —\l \f N

H 9 i? 9
—N-CH—C——NH-(IZH—C—NH-CH—C-O—

(EH2 H (EH2
9H2 9:0
9H2 OH
IIIH

(Er-NH

NH;

Figure 14. RGD arrrino acid sequence — Arginine-Glycine-Aspartic Acid.

The physiological effect of this modification was seen in the significantly increased
surface area of bovine aortic endothelial cells (BAECs) on the RGD modified PLA-co-
lysine compared to the surface area of BAECs on pure PLA, pure PLA-co-lysine, and
control RGD-peptide sequences.‘38 The coupling procedures of amino acids to existing
biodegradable copolymers are an alternative way to incorporate amino acid firnctionality
compared to the design of a protected comonomer.

This work also provides a model for the modification of biodegradable polymers
with other large groups such as pharmaceutical agents, growth factors, and other proteins.
From a biodegradation standpoint, the lactic acid and amino acid products upon
degradation can be processed in viva. Applications of these copolymers in tissue

engineering enhance biocompatibility through cell-peptide interactions as well as the cell-

20

polymer interactions. The latter interface can be altered by the direct coupling of specific

substrates.

1.4.1.4.2 Non-Peptidic-Based Copolymers

Although the functionalization of biodegradable polymers has been established as
a viable strategy in the design of amino acid/peptide-modified biomaterials, this approach
is not limited to amino acids. The synthesis of other functional comonomers and
copolymers has also been shown. Gonsalves has synthesized a biodegradable copolymer
containing pendant phosphonate groups.'44 The phosphonate groups are incorporated to
help induce the nucleation and growth of hydroxyapatite, Ca10,(POa)6(OH)2, an essential
mineral in the regeneration of tissue responsible for bone formation. Gross and
coworkers have demonstrated the incorporation of carbohydrate functionality through the
copolymerization of a protected cyclic carbonate with L-lactide.I45 Deprotection results
in the random copolymer, PLA-co-pentafuranose, with pendant sugars along the
backbone. The hydroxyl groups of the pendent sugar are now accessible for further
reactivity. Carbohydrates have found their niche as biologically compatible fimctional
groups in part due to their biodegradability, where the degradation rates are much faster
in carbohydrate-modified copolymers than in PLA and PGA polymers.‘29

Specific drawbacks to the design of specialized comonomers are (1) addressing
the differences in physiological function requires the constant synthesis of new
monomers which reflect the function, and (2) functional monomers must be compatible
to the conditions of the polymerization. The majority of biodegradable copolymers fall
under one of these two categories. The design of a chemically inert monomer possessing

Sites which could be modified after the polymerization would enable us to incorporate

21

diverse functionality in the framework of the polymer, and prevent a complicated
monomer and copolymer synthesis. Once the functional copolymer is prepared, routine
organic methodology can provide an avenue for synthesizing a broad range of materials.
By employing a combinatorial approach to polymer synthesis, our aim is to develop a
biodegradable polymer system based on a single copolymer by which we can introduce
hydrophilicity, hydrophobicity, pendant reactive sites, and macroinitiators for different
polymer architectures, all through simple and efficient reactions common to synthetic
organic chemists. Post-polymerization modification offers us the possibility of using a
functional monomer whose synthesis and copolymerization with lactide can be achieved

through the same simple protocol as lactide itself.

22

CHAPTER TWO
2 Modification of Poly(Lactic Acid)

Biodegradable polymers have found their niche as important materials with wide
applicability in the biomedical field. Medical devices made from synthetic biodegradable
polymers such as PGA, PLA, PLGA, PHB, and PCL have become standard in the
medical community.5'7 In tissue engineering applications, scaffolds made from these
synthetic polymers, have been improved by designing unique three-dimensional
structures. Despite this improvement, the scaffold itself remains a hydrophobic material,
possessing neither control over cell adhesion nor any biospecific functionality.

”"133 of biodegradable polymers with biologically relevant materials,

Copolymerizations
have helped improve control over cell adhesion, but these materials do not possess any
functionality other than a single reactive endgroup. Hybridizations‘r’o'66 may offer more
Specificity, however, a new material must be made for each new application, and
opportunities for multiple functionalization within a given matrix are not available. Our
approach towards improving current methods of functionality is to design a single
copolymer which possesses sites for modification to introduce functionality by more than
one way. Starting with a single biodegradable copolymer, the functional side chain can
be modified in a number of different ways through a multitude of organic
transformations. This would allow the modification of the biodegradable polymer and
subsequent function to be differentiated through established organic methodology.

Moreover, at biologically relevant loadings of functionality, the physical properties of the

polymer can be maintained. Post-polyrnerization modification is our method to access a

23

library of copolymers by simple covalent attachment of the desired firnctional group to

the polymer chain. This would allow us to work from a single copolymer.

2.1 Monomer Synthesis

The functional comonomer, 3,6-diallylglycolide (DAG), was synthesized by the
acid catalyzed condensation of the allyl substituted a-hydroxy acid, 2-hydroxypent-4-
enoic acid. This acid catalyzed condensation is the industrial process by which L-lactide,
the monomer precursor of PLA is synthesized, and is the general approach to a variety of
alkyl substituted glycolides.'46 Using a different approach, Emrick and coworkers have
synthesized a-allyl(5-valerolactone) via allylation of 8-valerolactone.147 However, in our
case, the a-hydroxy acid was readily prepared from the bismuth-zinc mediated allylation
of glyoxylic acid monohydrate using allyl bromide as the allyl source. The acid was
obtained“ in excellent yield and could be used directly in the next step or recrystallized
from ether and hexanes give a pure crystalline material. The acid-catalyzed condensation
of the allyl-substituted or-hydroxy acid was carried out in toluene for 3 days with the
water from condensation being removed using a Dean-Stark trap. The crude reaction
mixture contained both oligomers and the monomer. An isomer mixture of 3,6-
diallylglycolide, was isolated as a colorless oil by distillation of the reaction mixture

under reduced pressure using zinc oxide, ZnO, as a transesterification catalyst (Figure

15).

24

Allyl Bromide,

O
HBJZOHWO 8031" I option, :pJ a? (3?

THF, 16h 0H 2. A, ZnO

 

93% yield Meso (45%)
Racemic NFj(55%)

40°/o yield

Figure 15. Synthetic route to 2-hydroxypent-4-enoic acid and 3,6-diallylglycolide.

The monomer contains a near statistical mixture of RR, RS, and SS stereoisomers
which can be distinguished by NMR spectroscopy (Figure 16). This is seen mostly
clearly by the observation of the methine region of the proton NMR spectrum. These

protons both are observed as doublets of doublets.

 

 

 

 

 

 

 

 

 

 

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Hb,Hc
Hd Haunt:
1T1IITIIYTIIIITII’TITII1’111’11ll'TlllaTllll'rlllIrIl11171IIIIT‘IT[VIII]ITIITI’IWIIII’IIIITIITI‘ITITIII'TII'T’II—TITTTTWIT
6.0 5.0 4.0 3.0 ppm

Figure 16. lH NMR (300 MHz, CDCl3) spectrum of 3,6-diallylglycolide enriched in the R,S isomer. H,*

denotes the R,S isomer. H,M denotes the R,R and 8,8 isomers.

25

2.2 Polymer Synthesis

The statistical mixture of the monomers was polymerized in solution to give the
homopolymer, poly(diallylglycolide), PDAG. The polymerization of 3,6-diallyglycolide
in toluene at 80°C for 3 days using tin(II)-2-ethylhexanoate as the initiator gave PDAG as
a tacky solid with a number average molecular weight (Mn) of 24,100 and a glass
transition temperature (T3) of -10°C. Given the allyl side chain of the homopolymer, the
low Tg of PDAG is consistent with results obtained by Yin and Baker for alkyl-
substituted polyglycolides.146 The melt polymerization of 3,6-diallylglycolide can also be
performed, however, at high temperatures side reactions lead to poor control over the
molecular weight and molecular weight distribution. The resultant polymers, whether
prepared by solution or the melt polymerization are discolored and their physical texture
is tacky making it very difficult to handle. The 1H NMR spectrum shows three sets of
broad peaks with complicated multiplicities (Figure 17), and a quantitative 13C NMR
spectrum shows multiple resonances for the carbonyl, indicative of a mixture of
stereochemistry throughout the polymer. The physical consistency of PDAG and the
difficulty with which it is handled, render the use of the low Tg polymer impractical.
Greater control over the glass transition, melting point, and other properties can be
achieved by copolymerizing DAG with a monomer such as rac-lactide or L-lactide.
Moreover, the benefit of a copolymer incorporating DAG as the minor component in a
copolymer is to maintain similar properties to the homopolymer of the major component,

with the added benefit of functional sites for modification.

26

 

 

 

 

 

d

(i
'lWi’IliiiFI‘lll’lTITli'lrl‘ll1"I'Tl'll“I'l’Tl"er—_I"Tl["l’lrll'”
6.0 5.0 4.0 3.0 ppm 2.0

Figure 17. 1H NMR (300 MHz, CDCl;) spectrum of poly(diallylglycolide) (PDAG).

2.3 Copolymer Synthesis and Characterization

Melt polymerizations of mixtures of L-lactide with 3,6-diallylglycolide in the
desired ratio using tin(II)-2-ethylhexanoate and 4-tert-butylbenzyl alcohol (BBA) as
coinitiators affords the copolymer poly(L-lactide-co-diallylglycolide), PLLA-co-AG.
The reaction is carried out at 145°C for approximately 15-20 minutes. Since the ROP of
lactide and glycolides is considered a living polymerization, the molecular weights of the

4 49
18" Copolymers

polymers can be controlled by the monomer-initiator ratio (M/I).
containing up to 15% of the allyl comonomer were synthesized using rac-lactide (Table

3) and 32% using L-lactide (Table 4) as the lactide sources. In both cases, the amounts of

allyl in the monomer feed ratio compares well to the allyl content in the copolymer.

27

IBBA
Sn(Oct)2 >

 

O
PLA 0 O PLAoOBBA
O

 

 

145°C
0 O I
BBA = 4-tert-butylbenzyl alcohol
Ho’ C {
Monomer Copolymer 1 Weight
Feed Ratio Composition“ A3153)? 5 Fraction of T g / °C
(LA:DA G) [LA'DA G) A llylgycolide
100 : O 100 : 0 32,000 (1.67) 0.00 50.1
96 : 4 96 : 4 30,400 (1.27) 0.06 42.3
92:8 91 :9 21,600 (1.27) 0.12 40.9
89: 11 86: 14 23,800 (1.35) 0.18 35.1
85: 15 81 :19 16,200(l.36) 0.24 31.9
021009 0:100 24,100 (1.22) 1.00 ~10.0

 

Table 3. Copolymerization of rec-Lactide and 3,6-diallylglycolide ([M]/[I] = 300). " Molar composition
of DAG determined by integration of the methylene resonance of the allyl repeat unit compared to the
methine resonance of the lactide repeat unit. b Molecular weight determined by GPC. ‘ PDI =
Polydispersity index (MW/Mn). ‘1 Molecular weight determined by intergration of 4-tert-butylbenzyl
endgroup resonance compared to methine resonances of the lactide and allylglycolide sub-units.
Toly(diallylglycolide) prepared by solution polymerization in toluene at 80°C for 72 hours using Sn(Oct)2

as the catalyst.

28

 

 

Copolymer 1
Monomer Feed . . 0 Mn /gmof o -,
Ratio (LLA-DAG) 3:303:23 (PDI)"" T'" / C A” ”g
100:0 100:0 17,000(1.16) 175 44.1
95 : 5 96 : 4 17,900 (1.24) 158 28.2
90: 10 92 :8 18,800 (1.29) 141 15.5
85: 15 89:11 14,100 (1.32) 131 4.4
50:50 68 :32 12,800 (1.18) - -
0:100 0:100 24,100 (1.22) - -

 

Table 4. Copolymerization of L-Lactide and 3,6-diallylglycolide ([M]/[I] = 200). “ Molar composition of
DAG determined by integration of the methylene resonances of the allyl repeat unit compared to the
methine of the lactide repeat unit. h Molecular weight determined by GPC. ‘ Polydispersity index
(MW/Mn). d Poly(diallylglycolide) prepared by solution polymerization in toluene at 80°C for 72 hours

using Sn(Oct)2 as the catalyst.

The copolymers were characterized by 1H NMR spectroscopy and clearly show
the presence of the allyl subunits in the copolymer (Figure 18). At 5.7-5.8 ppm, the
resonance for the internal proton of the olefin is observed as a broad multiplet, while the
resonances for the terminal olefin and the methine protons are buried underneath the
large methine resonance of the PLA backbone. Also, the methylene resonances of the
pendant allyl group are observed between 2.6-2.7 ppm. At low enough monomer to
initiator ratios, the molecular weights of the polymers can be measured by integrating the
tert-butyl resonance of the BBA endgroup as the reference. However, at higher monomer
to initiators ratios, integration of the tert-butyl resonance was unreliable. Since molecular
weights of our copolymers determined by the integration of the BBA resonance gave
inconsistent results, the molecular weights of our polymers were determined using gel

permeation chromatography (GPC).

29

 

 

 

 

 

j 0
We
CH3
0 l
H u
1*?" l
a...)
d,e O l
at.
c ill a,b ’U\Cfi3 ’Bu
- A - D. .4
llVYFI I I i l T'I I I I I I 17 Y‘l I I I l l I l I l l I I‘I l V I l l I l I I77 VII I II V I771 '7 I_I I fl!1 r
6.0 5.0 4.0 3.0 2.0 1.0 ppm

Figure 18. 1H NMR (300 MHz, CDC l3) spectrum of poly(L-lactide-co-allylglycolide).

The copolymers based on rac-lactide were amorphous polymers with varying
glass transition temperatures, while the copolymers based on L-lactide showed some
crystallinity up to at least 11% comonomer incorporation. Both sets of copolymers based
on rec-lactide and L-lactide were characterized by differential scanning calorimetry
(DSC), and can be seen in Table 3 and Table 4, respectively. In the case of poly(rac-
lactide-co-diallylglycolide), (PDLLA-co-AG), an increase in the allyl content in the
copolymer results in a decrease in the observed Tg. This decrease behaves linearly

according to the Fox equation'46‘150"5' (Figure 19):

3O

1 WA WB

T Ts” T33

8

 

 

 

Figure 19. Fox equation describing the glass transition temperature of a random copolymer. w = weight

fraction, and TgA and TgB are the glass transiton temperatures for the pure homopolymers.
The glass transition of the pure PDLLA was measured at 501°C and the pure

PDAG was measured at -10°C. Analysis of the copolymer Tgs with respect to the allyl

weight fraction reveals a nice fit according to Fox’s model (Figure 20).

 

55

9

g45~ .

a

E [:1 a
8.404

E

P

 

 

 

30 T r , T
0.00 0.05 0.10 0.15 0.20 0.25
W Diallylglycolide

Figure 20. Glass transition temperature of poly(rac-lactide-co-diallylglycolide) with respect to the weight
fraction, w, of diallylglycolide. (Cl) Experimental Tgs of PDLLA-co-AG copolymers. (0) Theoretical Tgs

predicted by the Fox equation.
In the case of poly(L-lactide-co-allylglycolide) (PLLA-co-AG), DSC reveals that
the melting temperature, Tm, and the crystallinity, AH, decrease as the allyl content in the

copolymer increases. When the mole fraction of DAG exceeds 0.11, the crystallinity

disappears and the polymer becomes amorphous. In a similar fashion to the PDLLA-co-

31

AG copolymers, Tm and AH for the PLLA-co-AG copolymers are related linearly (Figure
21). This result is not uncommon to copolymers where the addition of a comonomer
decreases the crystallinity compared to the homopolymer.152 Both sets of data are

consistent with a random copolymerization as opposed to a block copolymer.

 

 

 

 

50$ 180
40 4
+ 160
«TA 30 F a A
a: e
E
:g 20 1 L E“
8 ~ 140
10 i .
l D
0 .r i 1 120
0.00 0.04 0.08 0.12

X. Diallylglycolide

Figure 21. (D) Crystallinity (AH) of poly(L-lactide-co-allylglycolide) with respect to the mole fraction, x,
of diallylglycolide. (O) Melting point, Tm, of poly(L-lactide-co-allylglycolide) with respect to the mole
fraction, x, of diallylglycolide.

Spectroscopic and thermal characterizations suggest the presence of a random

copolymer. Pendant olefins located randomly along the backbone are now ready to

undergo functional group transformation.

32

2.4 Post-Polymerization Modification
2.4.1 Olefin Cross Metathesis

2.4.1.1 General

Carbon-carbon bond formation via ruthenium-catalyzed olefin metathesis has
become standard procedure for the synthetic organic chemist in the design of simple and
complex organic molecules, such as key intermediates in the synthesis of natural

153,l54

products and for polymer synthesis.15 5'15 8 The use of versatile, air-stable ruthenium

159"“ research group has made olefin

carbene catalysts developed by the Grubbs
metathesis a common synthetic technique. Since the discovery of Grubbs’ first
generation carbene catalyst, Clz(PCy3)2Ru(=CHPh) (Figure 22a), a new, more active
second generation catalyst, C12(PCy3)(IMes)Ru(=CHPh), was developed by Grubbs, in
which one of the tricyclohexylphosphine ligands is replaced by a less labile N-

heterocyclic carbene ligand. (Figure 22b).

f—\
Mes-N N-Mes
PCy3 T
Ph Ph
CIH'RUZ/ CllhRuz/
Cl’ I Cl’ l
PCY3 PCY3
Cy = cyclohexyl Mes = 2,4,6-trimethylbenzene
(a) (b)

Figure 22. (a) Grubbs’ first generation ruthenium carbene olefin metathesis catalyst. (b) Grubbs’ second

generation ruthenium carbene olefin metathesis catalyst.

Compared to ring-closing (RCM) metathesis, ring-opening metathesis polymerization
(ROMP), and acyclic diene metathesis (ADMET), olefin cross metathesis (CM) has not

been applied nearly as often. The reaction offers a unique method of carbon-carbon bond

33

formation, yet the lack of selectivity with which CM operates makes widespread
applications impractical (Figure 23). Therefore, until general conditions for highly
selective CM have been discovered, it will remain a reluctantly approached technique.

Ru=CH(R) (cat)
>
- H20=CH2

2% R2

 

2°
/
/
I:

desired undesired
Figure 23. General scheme for ruthenium-catalyzed olefin cross metathesis.

Achieving selectivity for olefin cross metathesis must be found in choosing in the
appropriate substrate. Selectivities may be improved by adjusting the cross metathesis
conditions to decrease a substrate’s reactivity via its inherent electronic or steric
propertieslf’z’163 Interesting applications of cross metathesis used in biologically relevant
systems include the CM of allyl-modified amino acids164 and natural productsms’166 In
spite of the limitations, there are examples of CM that provide useful avenues to highly
desirable compounds. An exquisite example of cross metathesis is found in the
“dimerization” of the immunosuppresant FKSO6, which is used as a chemical inducer of

7 Like its small molecule analogues, CM on polymer substrates is not

dimerization.16
utilized nearly as much as RCM or ROMP. However, the benefits of performing CM on
polymer substrates are useful when it comes to purification of the products, since the
products can be much more easily separated from the small molecule by-products.
Blechert and coworkers have demonstrated the use of allyldimethylsilylphenyl-
functionalized polystyrene as a template for the synthesis of organic substrates by

ruthenium-catalyzed cross metathesis (Figure 24).168

Afier the cross metathesis product is
synthesized, the silyl group is subjected to electrophilic cleavage of the olefinic substrate,

resulting in the formation of a new C-C bond. Breed has also demonstrated CM on

34

polymers by reacting a hydrophilic triethyleneglycol allyl ether side chain bound to a

POE/styrene resin with simple olefins.169

Cleavage

O/\ M OA/R —————> AR

Figure 24. Schematic of polymer-bound ruthenium-catalyzed cross metathesis (CM), followed by

cleavage from the polymer support.

2.4.1.2 Synthesis and Characterization

In preparing the allyl-functionalized copolymer, PLLA-co-AG, we designed a
biodegradable copolymer able to participate in olefin cross metathesis. In order to test its
metathesis reactivity and demonstrate a proof of concept, we chose a group of olefin
substrates to perform cross metathesis with PLLA-co—AG: 9-decenyl-l-oxy-(4-
nitrobenzene), allylbenzene, 2-(2-propenyloxy)benzaldehyde, 9-decene-1-oxy—(tert-
butyldimethylsilane), and 9-decen-l-al (Figure 25).

/\/O H
M0 / / 6 Moms m

0

9-Decen-1-oxy- Allylbenzene 2-(2-propenyloxy) 9-Decen-1-oxy- 9—Decene-1-al
(4-nitrobenzene) benzaldehyde (tert-butyldimethylsilane)

Figure 25. Substrates used for polymer-bound olefin cross metathesis to PLLA-co-AG.

Quantification of the metathesis conversion of the PLLA-co-AG copolymer was
carried by 1H NMR and UV-Vis spectroscopy. For all substrates, 1H NMR analysis was
performed by comparing the integration of the resonances between newly formed
functionalized copolymer and the unreacted copolymer. In order to demonstrate the
validity of quantitative analysis using UV-Visible spectroscopy, we chose an olefin

substrate possessing considerable reactivity and a UV-Vis-active chromophore. The

35

olefin cross metathesis between PLLA—co-AG and 9-decenyl-1-oxy-(4-nitrobenzene) was
carried out at room temperature in dichloromethane for 24 hours using Grubbs’ 1St

generation ruthenium carbene catalyst (Figure 26).

0“”
Mo

C12(PPh3)2RU=CHPh ONOZ
(30 eq.) (0.10 eqi , EWO
CH2 C12, RT, 24b 7

 

T

26% (79%)

Figure 26. Polymer-bound olefin cross metathesis between PLLA-co-AG and 9-decenyl-l-oxy-(4-

nitrobenzene). Conversion of allylglycolide sub-units (isolated yield).

Purification of poly(L-lactide-g-9-decenyl-l-oxy—(4-nitrobenzene) (PLLA-g-
Decenyloxy-NB) required dissolving the polymer in dichloromethane and passing it
through a plug of silica gel. The unreacted olefin and cross metathesis by-products
passed through the column while the polymer remained bound to the silica gel. The
polymer was then extracted from the silica gel using dichloromethane and ethyl acetate
and was-precipitated from hexanes. Once the polymer was filtered it was dissolved in
dichloromethane a second time and reprecipitated from methanol. A second precipitation
the addition of hexanesequential precipitation from hexanes followed by methanol. The
isolated yield of the polymer was 79% and the Mn increased from 10,500 to 13,300 gmol'
' (MW/Mn = 1.40). The molecular weights provide no evidence for degradation, but the
polymer was discolored, indicating that trace amounts of ruthenium still may be present.

After work up of the polymer, 1H NMR clearly shows the incorporation of the
decenyloxy nitrobenzene olefin substrate onto the copolymer (Figure 27). Integration of
the olefinic regions around 5.2-6.0 ppm and the substrate methylene resonance at 4.0

ppm, with respect to the functionalized and unfunctionalized copolymer methylene

36

resonances between 2.5-2.7 ppm, revealed the conversion of the pendant allyl groups of

the PLLA-co-AG copolymer.

 

 

- $1-. Jk

 

 

 

W0
ii A .1 4 All. ML JLU
I—TTfiIW I7TI I I T ITITI—YI I I I TrfITIfI I I I I I I I IfI rI rfI I l I I I I I fiI I ITI I ITITI—I I I I IjT rFT—T
8.0 7.0 6.0 5.0 4.0 3.0 2.0 ppm

Figure 27. lH NMR (300 MHz, CDCl;) spectrum of polymer-bound cross metathesis between PLLA-co-

AG and 9-decenyl-l-oxy-(4-nitrobenzene). (*) Unreacted PLLA-co-AG.

Using GPC, we were also able to measure the olefin metathesis conversion
quantitatively using ultraviolet-visible spectroscopy. Since GPC is a chromatographic
separation technique, we measured the absorbance of the polymer using the UV-Visible
detector and correlated it to the concentration of the sample using 9-decenyl-1-oxy-(4-
nitrobenzene) as the calibration standard. From these data we can determine the percent
conversion of the olefins that underwent metathesis. The prerequisite for reliable UV-Vis
quantification is that the side chain chromophore has a strong absorption at a wavelength

where the polymer backbone is transparent. In our case, Am“ is 232 nm for PLLA-co-

37

AG, while the 2mm is 307 nm for the 9-decenyl-l-oxy—(4-nitrobenzene) (Figure 28).
Thus, we were able to quantify the amount of conversion using UV-Visible spectroscopy.

NMR and UV-Vis analytical methods give consistent results for the conversion of 26.3

and 29.4%, repectively.
0.08
s
0'06 ’ (reference) / N02

/

Q)
§
.0
< N02
(reference)
0.02
0.00 z - fl .-

 

240.0 280.0 320.0 360.0 400.0 440.0 480.0
Wavelength (nm)

Figure 28. UV-Vis spectrum of polymer-bound olefin cross metathesis between PLLA-co-AG and 9-
decenyl-l~oxy-(4-nitrobenzene).

The order of the addition of reactants turns out to be quite important. Routinely,
the polymer is dissolved in dichloromethane with approximately five equivalents of the
olefin substrate. Grubbs’ second generation ruthenium catalyst is then added to the
stirred solution and a syringe pump slowly delivers additional substrate at a rate of 0.1
mL/min. If the ruthenium catalyst is added in the absence of the five equivalents of
olefin substrate, the solution becomes viscous and ultimately forms a gel after ten

minutes, presumably as a result of cross-linking the pendant allyl groups. Interestingly,

38

this rapid gellation was not observed with Grubbs’ first generation catalyst. Degradation
of the polymer to its alkyl esters was carried out by refluxing the polymer in ethanol and
excess of Sn(Oct)2 for 48 hours (Figure 29). Both alkyl ester products were observed by
gas chromatography in the case of the linear copolymer. In the case of the cross-linked
gel, the dihydroxy-diethyl ester degradation product that arises from allyl metathesis was
observed and the absence of the allylhydroxy ethyl ester indicated the full consumption
of the terminal olefin sub-units.

Linear Copolymer:

CH3 0 CH3
0 9, Sn(0ct)2 O o
0 o EtOH 08 051
I 48 h

 

OH OH

C ross-linked Copolymer:

 

  

CH3 0 CH3
0 I
(HAD W0 n
O O SMOCtlz O OH O
EtOH ' H3C\(U\OEI + EtOMOEt
o o 48" OH 0 OH
(VLHO ‘5”)
CH3 n o 2 CH3 n

Figure 29. Degradation of linear copolymer PLLA-co-AG and cross-linked copolymer PLLA-co-AG
copolymer.

The mechanism of cross-linking is similar to the mechanism of ADMET, where
diene monomers undergo metathesis to combine as in a step growth mechanism.158
Without the presence of a monomeric olefin to participate in cross metathesis reactions,
intermolecular metathesis of the side chains on the polymer is a likely occurrence. Poché
observed a similar cross-linking phenomenon when he reacted polypeptides containing

olefin side chains with Grubbs’ first generation ruthenium catalyst.170 Therefore, for

39

cross metathesis reactions, five equivalents of olefin substrate were added prior to the
catalyst in order to favor cross metathesis instead of cross-linking.

Because of the proximity of the terminal olefins in the diallylglycolide sub-unit,
we did not rule out the possibility of ring-closing metathesis along the polymer chain
(Figure 30). In our model of ring-closing metathesis between adjacent allyl sub-units, the
eight-membered ring-closed product was not observed by GC/MS. While five-, six-, and
seven-member rings are thermodynamically stable, an eight-member ring is less stable,
and consequently, not as easily formed by ring-closing metathesis.171 Macrocycle
formation (> 8-membered rings) is also possible, but alone, would not account for the gel
formation. Therefore, it is likely that intermolecular metathesis between polymer chains

is primarily responsible for the gel formation.

40

[Ru]=CHPh ¥
CHZCIZ. 24h

 

     

       

[Ru]=CHPh

CHZCIZ, 24h k /

 

   

 

   

 

(a)
l
0 O 0
R0 0 OCH3 [Ru]=CHPh > R0 0 OCH3
O CHzclz, 24h _
|
R = H, tetrahydropyran Not Observed
(b)

Figure 30. (a) Possible polymer ring—closing metathesis reactions. (b) Ring-closing metathesis of the
methyl ester of the ring-opened diallylglycolide performed under polymer-bound cross metathesis

conditions.

The catalyst loading is an important variable in metal-catalyzed olefin metathesis.
High catalyst loadings require more extensive purification steps to remove the metal.
This is even more important when the metathesis reaction involves a biodegradable
material. The effect of catalyst loading on the conversion of the polymer-bound olefins
to products was investigated (Figure 31). Conversion increases with catalyst loading, but

conversions exceeding 50% require impractically high catalyst loadings.

41

 

    

 

 

 

 

 

 

50.0— I 1H NMRI
DUV-Vis

4o.oe
E
g 30.04
e
O
>
C
O
0
g 20.04
h—
2
o

10.0—/

0.0 . i i

3.0 7.6 18.0
[Ru] (mol%)

Figure 31. Percentage of olefin conversion of 9-decenyl-l-oxy-(4-nitrobenzene) with respect to Grubbs
1’t generation catalyst loading.

Although Grubbs’ catalysts are known to be functional group tolerant, we wanted
to evaluate polymer reactivity with a substrate possessing no functionality. Allylbenzene
is a simple aromatic molecule, where the ring is essentially nonfunctional. Furthermore,
allylbenZene metathesis products can be characterized spectroscopically. The metathesis
was carried out by the slow addition of the olefin in dichloromethane solution of the
polymer using 10% ruthenium catalyst relative to the amount of allyl sub-units in the
copolymer. Afier 24 hours, the polymer underwent successful cross metathesis as

determined by 1H NMR (Figure 32).

42

*VQ C12(PPh3)2Ru=CHPh

(30 eq.) (0.10 eq.) > EAWQ

CH2 (:12, RT, 24h

 

T

75% (90%)
(a)

(like) @AA/Q:

CHCl3

of ML. 1..

IIIIIIIIIIIIIIIIIIII"IIII IIIlIIIIIII IIIIIIjIIIIIIWIITITIIIIITTT

4.0 3.0ppm

 

 

( b)

Figure 32. (a) Polymer-bound cross metathesis between PLLA-co-AG and allylbenzene. Conversion of
allylglycolide sub units (isolated yield). (b) 'H NMR (500 MHz, CDCl3) spectrum of polymer-bound cross

metathesis" between PLLA-co-AG and allylbenzene. (*) Unreacted PLLA-co-AG.

Polymer purification was carried out by a simple washing of the polymer with
hexanes, followed by precipitation from methanol. This procedure washes away the self-
metathesis by-product of allylbenzene, 1,4-diphenyl-2-butene. After multiple
precipitations, residual ruthenium was likely responsible for the discoloration of the
material. After determining the reactivity of the allylbenzene, we looked for a substrate
which could be covalently attached and further modified by exploiting some form of
secondary reactivity. We chose the allyl ether of salicylaldehyde, 2-(2-

propenyl"oxy)benzaldehyde. Once the aldehyde was linked to the polymer, we could

43

envision the use of Schiff-base chemistry to link a broad range of molecules to the
polymer.

While the metathesis of allylbenzene proceeded well with good percent
conversion and in excellent yield, the 2-(2-propenyloxy)benzaldehyde conversions and
yields were significantly lower. Spectroscopically, we are able to observe the 2-(2-
propenyloxy) benzaldehydic 1H NMR resonances, however, there is a substantial amount

of unreacted polymer and substrate (Figure 33).

HO

A0
é C12(PPh3)2RU=CHPh

H 0
0.10 .
EM (3O eq.) ( eq) > W0
CH2 C12, RT, 24h

 

 

 

 

 

 

 

 

(a) 15% (63%)
0
fl
. (he)
a C H C CH3 as“
W06 1 i
b d e g
f
CHCl3 d
i M 1'"; L-
’IjIII—IYIIW WIIIIIITI—IIIIITWIIIIIIIIIT IIIWII'IYTTTIIITTIIITIIITTIIIITTI‘TI‘TII'IT"
ll .0 10.0 7.0 6.0 5.0 4.0 3.0 ppm
(b)
Figure 33. (a) Polymer-bound cross metathesis between PLLA-co-AG and 2-(2-

propenyloxy)benzaldehyde. Conversion of allylglycolide sub-units (isolated yield). (b) 'H NMR (500
MHz, CDC13) spectrum of polymer-bound cross metathesis between PLLA-co-AG and 2—(2-

propenyloxy)benzaldehyde. (*) Unreacted PLLA-co-AG.

44

In order to assess the reactivity of the 2-(2-propenyloxy)benzaldehyde, the self-
metathesis reaction was carried out under similar conditions to the polymer-bound cross
metathesis reactions. The result showed only 15% consumption of the olefin after 14
hours. Thus, the slow self-metathesis rate for 2-(2-propenyloxy)benzaldehyde explains
the low yield by cross metathesis products on the polymer. We speculate that the lower
reactivity may be chelation of the carbonyl oxygen of the aldehyde to the active
ruthenium catalyst during metathesis. This is consistent with literature assertions that
stable ruthenium-carbonyl chelates during metathesis can adversely affect activity.172

In order to make a substrate more accessible to the polymer, and thus more
reactive, we have incorporated a ten-carbon alkyl chain as a spacer between the olefin
and the desired functionality of the substrate. Substrates possessing the 9-decenyl spacer
such as" the 9-decenyloxy—tert-butyldimethylsilane and 9-decenal have improved
conversions compared to the benzaldehyde case and similar conversions compared
allylbenzene. However, a more detailed purification procedure must be used. The 9-
decenyloxy-t-butyldimethylsilane modified copolymer (PLLA-g-decenyloxy-TBS) and
9-decenal modified copolymer (PLLA-g-decenal) required precipitation from
dichloromethane into hexanes followed by a second precipitation from dichloromethane
into methanol in order to remove the by-products. The dimerized alkyl chain in both
cases is insoluble in methanol, and co-precipitated when only methanol was used. Once
the purification was complete, the 1H NMR spectrum of PLLA-g—decenyloxy-TBS
showed the presence of the TBS groups as well as the other characteristic resonances for
the alkyl chain spacer (Figure 34). The percent conversion of the allyl groups was

determined by the integration of the TBS groups with respect to the methylene

45

resonances of the copolymer at 2.5-2.7 ppm, which represent both metathesized and

unmetathesized allyl groups along the copolymer.

\ ,~‘
,3-
M0 "6 C12(PPh3)2Ru=CHPh

(0.10 eq.) \
EM (3O eq.) = Mofiiv
CH2 C12, RT, 24h 7 ’

 

 

 

 

 

 

 

 

84% (26%)
(a)
O
o H
H o
- o
W i W. T
CH3 ’3 m
a c e g l k l ,1
‘3? m j
’l / 0’ 76
J b d f h j 1
e-i
c,b dj fl
' k l
1 \ a,* l \ l
Till[Ii—lllIlli'lIllllIlllTlll'illlliv—lllllT—Illlirlll_llllT_TITYY"T1_1 I_T'T“I
6.0 5.0 4.0 3.0 2.0 1.0 0.0 ppm

(b)

Figure 34. (a) Polymer-bound cross metathesis between PLLA-co—AG and 9-decenyl-l-oxy-tert-
butyldimethylsilyl ether. Conversion of allylglycolide sub units (isolated yield). (b) IH NMR (300 MHz,
CDC13) spectrum of polymer-bound cross metathesis between PLLA-co-AG and 9-decenyl-l-oxy-t—

butyldimethylsilyl ether. (*) Unreacted PLLA-co-AG.

Similar to PLLA-g-decenyloxy-TBS, the 1H NMR spectrum of PLLA-g-decenal
showed the resonances of the alkyl chain spacer as well as the characteristic aldehyde

proton resonance at 9.74 ppm (Figure 35). Conversion of allyl groups was determined by

46

comparison of the methylene adjacent to the carbonyl at 2.39 ppm with respect to the

metathesized and unmetathesized methylene resonances of the copolymer at 2.5-2.9 ppm.

0

MH C12(PPh3)zRu=CHPh

7

(0.10 eq.)
@M (30 6‘“ > EWH

CH2 C12, RT, 24h

 

58% (48%)

(a)

 

l fl f l fl l l '"T l' T l ‘ l ’T l TF’ TF1 l l " l7 YdT l ' l l l 'T TTT
10.2 9.2 6.0 5.0 4.0 3.0 2.0 1.0 ppm
0?)

Figure 35. (a) Polymer-bound cross metathesis between PLLA-co-AG and 9-decenal. Conversion of
allylglycolide sub units (isolated yield). (b) lH NMR spectrum of polymer-bound cross metathesis between

PLLA-co-AG and 9-decenal. (*) Unreacted PLLA-co-AG.

Ruthenium-catalyzed olefin cross metathesis of the biodegradable copolymer
PLLA-co-AG with specific olefinic substrates has been successfully demonstrated. 1H
NMR and UV-Vis spectroscopic techniques have been shown to quantitatively determine
the conversion of the pendent olefins on the polymer. From a synthetic standpoint, we
have observed that the workup protocol varied from substrate to substrate. Furthermore,

we also see substrate dependence in the reactivity of different olefins. This is consistent

47

with what is known in the literature regarding the electronic and steric effects of olefin
metathesis.162"63 However, as a method for modifying biodegradable materials, even low
conversions using a bioactive substrate could be extremely valuable in tissue engineering
applications. Moreover, orthogonal functionalization strategies would further enhance

the utility of olefin cross metathesis for modifying DAG copolymers.
2.4.2 Hydroboration-Oxidation

2.4.2.1 General

In hopes of expanding our current method of post-polymerization modification,
we wanted to pursue an alternative means of functionalization which will allow us to
screen a broader range of biologically relevant substrates. While olefin metathesis can be
useful if the appropriate reactive substrate is used, not every substrate is metathesis
active. In certain cases, an appropriate bioactive ligand might first require chemical
modification before the polymer-bound olefin cross metathesis is applied (e.g.
synthesizing the alkenyl ether or ester of the ligand). Therefore, modification of the
terminal ~olefin through functional group transformation would allow us to explore other
types of coupling chemistry. Alcohols are a particularly useful functional group for
tethering substrates through reactions such as esterification or etherification, as well as
carbonate and carbamate linkages. The best known transformation for converting olefins
to alcohols is hydroboration followed by oxidation.

The first example of hydroboration was reported by H.C. Brown who discovered
the groundbreaking reaction for converting terminal olefins to alcoholsm'I76 In his
pioneering work, he used diborane (BzH6) to synthesize organoboranes from terminal

olefins followed by treatment of the organoborane with sodium hydroxide and aqueous

48

hydrogen peroxide to afford the primary alcohol. Hydroboration-oxidation has been one
of the most utilized methods to synthesize primary alcohols from olefins.

Similar to small molecule cases, polymer-bound hydroborations have also been
performed, mostly on polyolefins to generate block and graft copolymersm'180
However, the hydroboration reaction in the context of biodegradable polymers such as
poly(a-hydroxy acids) has not been exploited. The primary benefit of performing
hydroboration-oxidation on biodegradable polymers would be to incorporate hydroxyl
grafi sites for polymer modification. The chemistry of primary alcohols is plentiful, and
incorporating a hydroxyl site along the polymer backbone would position us to modify
our copolymer in a number of different ways using traditional synthetic organic
chemistry. However, preparing a hydroxyl monomer is impractical, as it will react
adversely under our polymerization conditions, namely with the Sn(Oct)2 catalyst. Since
a hydroxyl group such as an alcohol is required to initiate polymerization with Sn(Oct)2
as the catalyst, polymerizing a monomer containing a hydroxyl would prevent the
formation of high molecular weight polymer. Yang and coworkers have circumvented
this by synthesizing a glycolide monomer bearing a benzyl-protected vinyl alcoholm
After polymerization and deprotection, both homopolymer and copolymer with lactide

2 Gross has also

yielded the polyglycolide containing pendant hydroxyl groups.18
incorporated hydroxyl graft points by copolymerizing a carbohydrate-modified cyclic
carbonate monomer with lactide.I45 Subsequent deprotection of the benzyl groups yields

PLLA-co-pentofuranose. Jerome and coworkers synthesized a silyl-protected y—hydroxy-

e-caprolactone which they copolymerized with e-caprolactone in order to use the free

183

hydroxyl groups as macroinitiators for lactide grafts. Each of these approaches

49

achieves“ the goal of incorporating hydroxyl functionalization, however at the expense of
synthesizing a specialized monomer.

Utilizing a post-polymerization modification of our allyl-functionalized
copolymer PLLA-co-DAG, we report an efficient way to incorporate hydroxyl
functionality along our copolymer by avoiding the synthesis a complex monomer, and the
subsequent protection and deprotection schemes before and after polymerization. Simple
hydroboration of PLLA-co-DAG and further oxidation to the terminal alcohol has

enabled us to incorporate hydroxyl groups as grafting sites for additional modification.

2.4.2.2 Synthesis and Characterization

The hydroxylated copolymer of PLLA-co-DAG was synthesized by the
hydroboration of the copolymer followed by oxidation using aqueous sodium acetate and
hydrogen peroxide (Figure 36).

1. 9-BBN,THF, RT, 2h
HEM 2. NaOAc, H202, THF, RT, 13h _ UiMOH

 

PLLA-co-AG PLLA-g-HP

Figure 36. Synthetic route to poly(lactide-g-hydroxypropane) via hydroboration-oxidation of poly(lactide-

co-diallylglycolide).

In the first step of the hydroboration, the polymer was stirred at room temperature
for 1.5 to 2 hours using 9-borabicyclononane (9-BBN) as the borane source. The
progressof this reaction was monitored by the appearance of the trialkylboron resonance
centered at 88 ppm in the HB NMR spectrum. Once the hydroboration was determined to
be complete, the polymer was treated with a two-fold excess of aqueous base and

hydrogen peroxide. In our first attempt to prepare the hydroxylated copolymer, we used

50

sodium hydroxide. After 12 hours of oxidation, the polymer had degraded and could not
be precipitated. This prompted us to select a milder base such as sodium acetate, since
the polyester PLLA-co-AG is sensitive to harshly basic conditions. The reaction was
then stirred at room temperature for 13 hours or until the oxidation was complete.
Precipitation of the polymer from methanol two times gave the hydroxylated copolymer
poly(lactide-graft-hydroxypropane) (PLLA-g-HP). Under oxidation conditions using
sodium acetate and hydrogen peroxide, GPC analysis confirmed that our copolymer did
not degrade, where the molecular weight of the copolymer increased from Mn = 12,800
gmol'l for PLLA-co-AG to Mn = 13,800 gmol'l for PLLA-g-HP, an increase expected as
a result of copolymer fractionation through multiple precipitations. The 1H NMR
spectrum of the copolymer shows the disappearance of the allyl resonances and the
appearance of the primary alcohol at 3.6 ppm. Also, the alkyl methylene resonances of

the hydroxypropane grafi are seen between 1.5-2.0 ppm (Figure 37).

51

 

 

 

 

 

 

CH3 cg;
a,b e,f
MOH
(a) E c,d l
1 l
:3 e,f Ii
l
I c,d a,b ,111 L
I l
a,b d,e
(b) M
c
b O
3,
/U\Ct13 lg 'Bu
L wL___
l" I! T T IV’ITI I’ IT] I r11!” I I’l I ["1 I" l 1 II T [Tl IT I T TTI I IT’I I I’I T’IT’I T‘I’W T l"l I]
6.0 5.0 4.0 3.0 2.0 1.0 ppm

Figure 37. 1H NMR (300 MHz,CDCl3) spectra of (a) PLLA-g-HP; poly(lactide-g-hydroxy'propane) and
(b) PLLA-ico-AG; poly(l-lactide-co-diallylglycolide).

The advantages of a hydroxylated polymer are two-fold. First, the hydroxylated
copolymer can be used directly as a functional biomaterial, to improve biocompatibility
through hydrogen bonding, interactions with other bioactive groups, or through substrate
binding.‘84"86 Second, and perhaps more significant is the ability to support further
chemistry, such as macroinitiation or other coupling reactions. Primary alcohols have a
wealth of chemistry and this copolymer provides an ideal starting point to directly exploit
simple chemistry to create a variety of new materials.

Since initiation of lactide using Sn(Oct)2 requires an alcohol coinitiator, it is

possible to consider new polymer architectures using the pendant hydroxyl groups as

52

macroinitiators in the design of graft and dendrimeric-type copolymers!”189 A unique
example of this chemistry was can'ied out by Zheng and coworkers in designing
hydrophobic-hydrophilic microspheres of polydivinylbenzene and poly(hydroxyethyl
methacrylate) (poly(HEMA)). Hydroboration-oxidation of the pendant olefins and
subsequent esterification generated ATRP initiators for the grafting of the hydrophilic
acrylate.190

Using the hydroxyl sites merely as attachment points for further chemistry has
also been accomplished. Gross and coworkers have designed cyclic carbohydrate-
modified carbonate monomers, which have been copolymerized with lactide. After the
copolymerization of the carbonate monomer with lactide, the deprotection of the pendant
sugars are carried out to reveal hydroxyl groups now accessible for further

. - - 4 ,9 - 4
functionallzauon.l 5 l l 19

2.4.3 DCC Coupling

2.4.3.] General

Esters can be synthesized by a number of different methods, many of which
require acidic conditions. However, using dicyclohexylcarbodiimide (DCC) coupling,
we have used an established route to the efficient synthesis of esters, which are
covalently attached to our copolymer (Figure 38). DCC coupling is a well-known

l95-l 97

synthetic route applied in dendn'mer chemistry, peptide synthesis,'98’199 block

0 and coupling substrates to polymerszoo’zm An extremely

copolymer synthesis,‘3
attractive feature of DCC coupling is the ease of purification. Under the reaction

conditions of DCC coupling, the insoluble dicyclohexyl urea by-product is simply

removed by filtration, facilitating purification.

53

o o
0
Wm + JL DCC,DMAP(O.2 eq.), Woks: + CyHN/lkNHCy
H0 R CHZCIZ, RT, 20h

0
AR = Commercially Available Carboxylic acids

 

HO
Figure 38. General synthetic route to polymer-bound esters using DCC coupling.

The generality of this approach is also appealing. DCC coupling of our
00polymer with a variety of carboxylic acids affords a simple way to covalently attach
substrates containing a carboxylic acid functional group. An added benefit is the creation
of an ester linkage. Since our copolymer is a polyester, the ester linkage adjoining our
functional side chain to the polymer is an insignificant perturbation to the polymer itself.
Additionally, the esterified functional group may enhance the biocompatibility of our

copolymer as a material in tissue regeneration.

2.4.3.2 Side-Chain Esterification of Fatty Acids

There are a few classes of biodegradable materials which have utilized fatty acids

or long chain alkyl substituents. Such polymers have been used with the incorporated

103,202,203

alkyl chain acting as an inherent plasticizer, a bioactive substrate amenable for

100,101,204

drug delivery applications, or as a cross-linking agent?”207 These classes of

materials include medium chain length poly(hydroxyalkanoates) (mcl-PHAs),208

polyesters based on glycerol and diacid functional groups which incorporate fatty acids,

and polyanhydrides functionalized with substituted fatty acids (Figure 39).”)0'm3'205"206

54

(a) (b) M (c)
\L Q11} 1° 0 O 8 n
O n O=I

R

 

 

 

R = Medium length alkyl chain R = Fatty acid-based alkyl chain R = Fatty acid-based alkyl chain
Witholt et al. Kobayashi et a]. Domb et al.

Figure 39. Fatty acid-modified polymers. ((1) medium chain length poly(hydroxyalkanoate). (b)
biodegradable polyester based on glycerol, sebacic acid, and a fatty acid. (c) polyanhydride containing

fatty acid moieties.

Using commercially available fatty acids we have demonstrated DCC coupling as
a viable method for functionalizing PLLA-g-HP (Figure 40). The pendant hydroxyl
groups reacted with the fatty acid in the presence of DCC and a catalytic amount of N,N-
dimethyaminopyridine (DMAP) to form the ester linkages onto our copolymer. We
chose a group of long chain fatty acids, some of which have influence in the biology of

. 9
bone tissue growth.20 ’2”)

55

O O

0
Wm + JL occ.oMAP(o.2eq.) _r_ E/VOAR + CyHNJkNHCy
HO R CH2c12. RT. 20 h

 

O
PLLA-g-MyPr = EMOW
poly(l-lactide-g-myristic acid propyl ester)
0
poly(l-lactide-g-stearic acid propyl ester)
0
PLLA.g-Olpr = Mow—WW

poly(l-lactide-g-oleic acid propyl ester)

0
PLLA-g-LnPr = EMOMWW

poly(l-lactide-g-linoleic acid propyl ester)

0

PLLA-g-ArPr =

”3”

poly(I-lactide-g-arachidonic acid propyl ester)

Figure 40. Synthetic route to saturated and unsaturated fatty acid-modified PLLA using DCC coupling.

The reactions were carried out in dichloromethane for 20 hours to ensure
complete coupling. After the desired reaction time, the insoluble dicylcohexyl urea was
removed by filtration, and the polymer was concentrated, and reprecipitated twice from
methanol. The reaction conditions provide a simple and efficient method for ester
formation without harsh acidic or basic conditions. In spite of reports of polymer
degradation using DMAP?” we observe good yields, full conversions, and no molecular

weight loss for the polymer (Table 5).

56

 

 

‘ a c a c . T...‘/"C .
(Carbfzfgnfizfjrated) M"/g’"0[l(PDD '1) T8 / C T“ / C (AH/Jg" % Y’e’d
Myristic (14:0) 16,300 (1.37) 27 86 135 (24.3) 85
Stearic (18:0) 15,800 (1.37) 25 79 131 (29.5) 87
Oleic (18: 1) 17,700 (1.36) 23 90 135 (25.5) 89
Linoleic (18:2)d 18,300 (1.48) 21 87 132 (23.7) 81
Arachidonic (204)“ 20,000 (1.63) 18 86 132 (26.1) 80
Linoleic (18:2)e 18,300 (1.48) 32 - - 81
Arachidonic (20:4)e 19,900 (1.84) 52 - - 80
PLLA-g-HP’ 13,500 (1.58) 54 121 142 (12.9) 75
PLLA (6% D)? 17,700 (1.28) 53 -" 151 (25.6) 73

 

Table 5. Saturated and unsaturated fatty acid-modified PLLA copolymers containing 6 mol % fatty acid. “

Molecular weight determined by GPC. b PDI = Polydispersity index. ‘ Differential scanning calorimetry of

6 mol % fatty acid-modified copolymers. Second heating scan at 10°C/min under helium atmosphere. d

Functional polymer not exposed to air. e Exposed to air until gellation renders the functional polymer
insoluble. f Hydroxylated copolymer starting material used to couple fatty acids. ‘ Independently prepared

PLLA containing 6 mol % D units. h Sample annealed at crystallization temperature.

1H NMR spectra show the quantitative incorporation of the saturated fatty acids
myristic (C 14) and stearic (C18) acid (Figure 41). Spectral analysis of the C13 unsaturated
acids also reveals complete conversion of the fatty acids to their polymer-bound esters
(Figure 42). Additionally, the polyunsaturated arachidonic acid-modified (C20) polymer
was synthesized and spectroscopically characterized (Figure 43). Proton NMR
assignments for all the fatty acid modified polymers were made in order to identify the
protons corresponding to the side chain. Some notable features in the NMR are that
linoleic acid and arachidonic acid—modified polymers both have bisallylic methylene
protons appearing at 2.7-2.8 ppm, which were absent in the saturated fatty acid and oleic
acid polymers. Also, the increase in intensity of the olefinic proton region at 5.3-5.5 ppm

is a characteristic feature of the covalent attachment of the unsaturated fatty acids.

57

Cfla

(a) EMO . I

 

 

 

(D

 

 

 

 

LL
(b) HMOW f-s
t

 

II’I r17l'lIlYlI1lYlIYIlIITlIII—r1III'TIT’TIIIfiT’TIYITYiiITI'T'T—I

5.0 4.0 3.0 2.0 1.0 ppm

Figure 41. lH NMR (300 MHz, CDCl3) spectra of (a) PLLA-g-MyPr; Myristic acid-modified PLLA-g-HP

and (b) PLLA-g-StPr; Stearic acid-modified PLLA-g-HP.

58

(a) BMOW

 

 

 

 

 

 

 

 

 

a c o e g 1
b d f h
. kJ, ,
11,0 11
J C
5.0 4.0 3.0 2.0 1.0 ppm

Figure 42. lH NMR (300 MHz, CDCl;) spectra of (a) PLLA-g—OlPr; Oleic acid-modified PLLA-g-HP and

(b) PLLA-g-LnPr; Linoleic acid-modified PLLA-g-HP.

59

 

 

 

 

I Til "I I”I l”I TNT ' ’1 '1”1 1'71 Til f1 F 177' Ti'l 61 F‘I T ‘T 17T_'1’;" I'CTT'TWT": I I 1 1 '

5.0 4.0 3.0 2.0 1.0 ppm

Figure 43. lH NMR (300 MHz, CDC13) spectrum of PLLA-g-ArPr; Arachidonic acid-modified PLLA-g-

HP.

Thermal analysis of the copolymers using DSC showed the T8 of the materials
decreased from the original PLLA-g-HP (54°C) upon esterification of the fatty acids
(Figure 44). The glass transition, crystallization, and melting temperatures of the fatty
acid-modified materials are all very similar. The crystallinity seen here is due to the
crystallinity of the backbone, since both PLLA-g-HP and PLLA-co-AG containing 6%
allyl units are still crystalline. The two observed melting points are due to or- and [3-

structures of PLLA, and have been discussed by other groupsm'2M

60

 

 

 

 

Endo —-——>

 

 

+—— Exo
v\
Q,

 

-15 - 25 65 105 145 185

Temperature (°C)

Figure 44. Differential scanning calorimetry of 6 mol % fatty acid-modified copolymers. Second heating
scan at 10°C/min under helium atmosphere. (a) PLLA (6% D-lactide); (b) PLLA-g—HP; (c) PLLA-g-MyPr;
(d) PLLA-g—StPr; (e) PLLA-g-OlPr; (0 PLLA-g-LnPr; (g) PLLA-g—ArPr.

A noteworthy feature of the DSC results is the behavior of the linoleic acid and
the arachidonic acid-modified copolymers. When left standing for a prolonged period of
time in solution in the presence of oxygen, the polymers became viscous and gelled.
After longer periods of time, the polymer became completely insoluble in any solvent,
providing evidence of oxidative cross-linking. The mechanism of cross-linking occurs
via oxidatively-induced isomerization of the bisallylic methylene hydrogens to generate a
peroxy-radical.215 The peroxy-radical then acts as the propagating species to facilitate

cross-linking between double bonds on adjacent chains (Figure 45). This cross-linking

216

behavior, known as curing, is common in the cross-linking of certain oils, and can also

61

be thermally induced, as in the ease of Kobayashi’s glycerol-based biodegradable

206 These biodegradable polyesters were

polyester containing fatty acid side chains.
thermally cured to give cross-linked, transparent polymeric films showing increased

hardness. In his work, only samples containing bisallylic hydrogens underwent cross-

 

linking.
O
EMOM/WW
H ’H
l—> O
EM0WM
H
. . 'H
‘ rearrangement and ox1dat10n - 02 I

0 'O‘o

‘ L" crosslinked polymer

Figure 45. Simplified synthetic route to the cross-linking of polyunsaturated fatty acid modified PLLA

216

copolymers. Proposed by Vogl and Blanksby.

For the saturated fatty acid and the oleic derivatives we prepared, there are no
labile bisallylic hydrogens, and we observe no cross-linking. This is in agreement with
the findings of Kobayashi.206 Reproducible DSC scans of each copolymer suggests that
they are insensitive to oxidative and thennally-induced cross-linking. Consequently, this
cross-linking behavior provides a basis for the observed increase in the Tg of the linoleic
acid and arachidonic acid-modified copolymers to 32°C and 52°C, respectively. The
arachidonic acid containing three bisallylic methylenes should be more susceptible to the

cross-linking, which is reflected in the higher Tg. Conversely, when the linoleic acid- and

arachidonic acid-modified copolymers are kept from oxygen, there appears to be no
cross-linking as indicated by the solubility, and the measured Tgs of the copolymers are

much lower at 21°C and 18°C, respectively (Figure 46).

(CIM

E‘

 

 

 

\
/

<——— Exo
’\
{i

 

Temperature (°C)

Figure 46. Differential scanning calorimetry of 6 mol % fatty acid-modified copolymers. Second heating
scan at 10°C/min under helium atmosphere. (a) PLLA-g-LnPr; linear polymer sample. (b) PLLA-g-LnPr;
cross-linked sample. (c) PLLA-g-ArPr; linear polymer sample. (d) PLLA-g-ArPr; cross-linked sample.
Linear polymer samples were prepared and stored under a nitrogen atmosphere prior to DSC analysis.
Cross-linked polymer samples were exposed to air during workup and allowed to cross-link until the

polymer became insoluble.

Increased Mn and PDI values for the linoleic acid and arachidonic acid-modified
copolymers may also be reflective of cross-linking between chains. This behavior is an

extremely interesting finding, and reveals a potential route to the synthesis of

63

polyunsaturated fatty acid-modified PLLA copolymers possessing higher Tgs, where only

small amounts of the coupled unsaturated side chain are required for cross-linking.
2.4.3.3 Side-Chain Esterification of Phosphonic and Amino Acids

2.4.3.3.1 Side Chain Esterification of Phosphonoacetic Acid

The synthesis of biodegradable phosphonated copolymers217 and biodegradable
polymer compositesm'221 have been targeted specifically for their ability to support
nucleation and growth of hydroxyapatite (HAp) and other minerals in the biology of
osteoblast tissue engineering. Hydroxyapatite, Ca10(PO4)6(OH)2, is the mineral deposited
by the osteoblast matrix in the formation of bone. Materials that nucleate HAp can help
in the proliferation of osteoblasts, which further differentiate into a mineral-depositing,
bone-producing matrix. Li and coworkers have designed biodegradable PLLA fibers
coated with HAp in order to help nucleate minerals secreted by the differentiated

'8 Gonsalves has designed a synthetic biodegradable copolymer

osteoblastic matrix.2
based on poly(e-caprolactone) which contains pendant vinyl phosphonate groups.217
Acrylate-based polymers containing pendant phosphonate groups are also used as
materials for dental compositesm’223 Regardless of the nature of the material, the
phosphonate groups are chosen for their ability to bind minerals such as calcium.
Consequently, the incorporation of phosphonate groups in the copolymer also increases
the hydrophilicity of the material. Incorporating hydrophilic moieties within the polymer
framework is a strategy employed to help control the cell adhesion properties of a given

material. PLLA is a hydrophobic polymer which cannot control cell adhesion. Thus,

incorporating pockets of hydrophilicity in the form of phosphonic acid groups to help

64

 

regulate cell adhesion as well as help in the templation of minerals which occur during
cell differentiation. This material can be used while maintaining the benefits of the
biodegradable polymer.

The synthesis of a phosphonic acid-modified copolymer was achieved by the
DCC coupling of diethylphosphonoacetic acid to PLLA-g—HP in the presence of a
catalytic amount of DMAP. After stirring for 20 hours the reaction was filtered and

precipitated twice from methanol (Figure 47).

o o
kamoenz, DCC, DMAP(O.2 eq.) 0 0
H0 11
OH > OJK/Hoa):
CHzClz. RT. 20 h
o o o o
H TMSBr lOe . .
Okploaiz ( q g OJK,'P(0H)2
CH2C12, RT, 14 h

Figure 47. Synthetic route to PLLA-g-DAPPr; Poly(lactide-g—diethylphosphonoacetic acid, propyl ester)

 

 

and PLLA-g-PAPr; Poly(lactide-g-phosphonoacetic acid, propyl ester).

This polymer was characterized by both 1H NMR and 3'P NMR spectroscopy
(Figure 48). A distinctive 1H NMR resonance for the methylene protons, which are
coupled to the adjacent phosphorous, gives rise to a doublet with a JP-” coupling constant
of 22 Hz. The ethyl resonances of the phosphodiester are also clearly seen at 1.2-1.4 and
4.1-4.2 ppm. A single resonance at 20.2 ppm in the 3'P NMR spectrum is also consistent
with the esterification product of diethylphosphonoacetic acid. The spectroscopic data
are in agreement with other phosphonated materials containing phosphonates and

phosphonodiesters.

65

 

 

 

 

3'P NMR ' a,b e f of k
(600)231ka
c,d
e,f,ij 1
I J. I
21.0 . I I
I“ gah a,b c,d II I (I
*1 11: ,. 1 . j 11 EU" J25». "1 .1ng 11
11 I I
a,b e,f 1 1
(b) 31? NMR 1 I OJE:B'\" “OH I
1 1 . .
3H Y ’3' l bed .1
26.0 20.0 . . e,f .’ I“
___,_,_,,../ WWW K“.
fi—fm—T—TI—r-w—Ffi wfifimfifi—Ifl’flWfl—p fl—wfivm_~__I_T__I_I T—F‘
5-0 4-0 3.0 2.0 1.0 ppm

Figure 48. ‘H NMR (300 MHz, CDCI3) and “P NMR (120 MHz, CDCI3) spectra of (a) PLLA-g-DPAPr;

Diethylphosphonoacetic acid-modified PLLA-g-HP and (b) PLLA-g-PAPr; Phosphonoacetic acid-modified

PLLA-g-HP.

The deprotection of the phosphodiester copolymer was achieved by simply
stirring the copolymer with a ten-fold excess of bromotrimethylsilane. After stirring for 8
hours, the polymer was quenched with methanol and precipitated from dichloromethane
into aqueous methanol. The labile silyl groups on the phosphodiester intermediate were
cleaved upon aqueous workup to yield pendant phosphonic acid groups covalently
attached to the PLLA backbone. The lH NMR shows the disappearance of the ethyl
group resonances. Notably, the phosphorous coupling to the methylene protons in the
side chain is not as prominent as in the protected phosphoester; however, the coupling is

observed when the polymer is characterized in another solvent such as d6-DMSO.

66

2.4.3.3.2 Side Chain Esterification of N,N'-diBoc-Lysine

Amino acids incorporated into biodegradable polymers are important bioactive

substrates in tissue engineering applications. They can serve as attachment sites for

138,224 65,225,226

adhesion peptides, growth factors, and provide a simple model for coupling
peptides to biodegradable materials. The known strategy for incorporating amino acids
into biodegradable polymers is to synthesize substituted morpholine-2,5-diones as the
amino acid-bearing functional comonomer.227 Copolymerization of the morpholine—2,5-
dione with a monomer such as lactide gives a biodegradable copolymer with amino acid
groups built into the polymer. Once polymerized, the material can be used directly in
biological applications or further modified by additional procedures (Figure 49). Many

o o o 0 ° .22
research groups have taken thls approach to incorporate amino acrds such as lysrne,I32 5

serine,'3 "228 aspartic
Protecting Group

1.Copolymerization CHSN o
2. Deprotection» '
H30 :N{R- PG)

Lactic Acid Amino Acid

Protected Morpholinedione _mm
Comonomer

CH3 H 0
Functional Tissue Scaffold 4 CNN
0 R

Figure 49. Synthetic strategy for designing biodegradable polymers with amino acid functionality.

229'” l and glutamic acidm'ng‘232 in polymers.

 

 

 

67

In an alternative approach, we have employed our DCC coupling chemistry to
couple lysine to our biodegradable copolymer. In order for our DCC coupling conditions
to work, we used N,N'-diBoc-lysine and carried out the DCC coupling procedure as
previously described (Figure 50). The DCC coupling proceeded smoothly, and the
polymer was precipitated twice from methanol. The 1H NMR spectrum shows the
expected peaks from the protected lysine copolymer (Figure 51). In a similar procedure,
Langer and Lavik used DCC coupling to modify the endgroup of PLGA, with

30 In our

carboxybenzoyl-protected poly(lysine) resulting in a block copolymer.1
reactions, the BOC protecting groups were removed by reacting the copolymer with an
excess of trifluoroacetic acid. The solution was directly poured into water containing an
excess of sodium bicarbonate to neutralize the excess acid. This precipitation was done
carefully since the evolution of carbon dioxide accompanies the acid-base reaction. The
polymer was obtained in nearly quantitative yield and contained the trifluoroacetate
ammonium salt of the deprotected lysine. '3 C NMR of the polymer showed two quartets
in the carbon spectrum at 162 ppm (Joy: = 33 Hz) and 118 ppm (J01: = 292 Hz) for the
carbonyl and trifluoromethyl carbon respectively. The coupling constants are diagnostic
for fluorine coupling with the carbonyl carbon and the trifluoromethyl carbon of the

acetate. The mixture was purified by centrifugation to give PLLA-g-LysTFAPr in nearly

quantitative yield (Figure 51).

68

 

O
N,N'-diBocLys-OH, DCC, DMAP (0.2 eq)
OH > OWNHBOC
CHZCIZ, RT, 20 h NHBoc
O Trifluoroacetic Acid (20% WV) 0
E/moMA/NHBOC > hoW/NHi’
NHBOC CH2Cl2, RT, l5 mm NH3‘

Figure 50. Synthetic route to PLLA-g-BocLysPr; Poly(lactide-g-N,N'-diBoc-lysine propyl ester), and

 

PLLA-g-LysTFAPr; Poly(lactide-g-lysine bis(trifluoroacetate) propyl ester).

Upon deprotection, the BOC resonance at 1.4 ppm disappeared and the other
proton resonances were shifted. The protected lysine-modified copolymer is soluble in
typical organic solvents such as dichloromethane, chloroform, and tetrahydrofuran. The
solubility of the deprotected form of the copolymer, however, can be dependent on the
molar composition of lysine units in the copolymer. We have observed that the lower
loading of lysine residues (<5%) increased the solubility in halogenated solvents.
However, if the lysine loading is high enough (>5%), the copolymer becomes more
insoluble in d-chloroform, and NMR characterization must be done in d4-methanol, d6-
dimethyl sulfoxide, or dg-thf. This solubility change was also observed by Langer with
his copolymers of PLLA-co-lysine, and thus this finding might be expected in our
case.l3?"233’234 The lH NMR of PLLA-g-LysTFAPr in dg-thf shows the alkyl resonances
more clearly (Figure 52). Spectral assignments of the proton resonances were based on

two-dimensional COSY NMR experiments.

69

 

I a c 0 d e g I: I
4; i - .
(a) '3 $N\OJ‘\I/\/\/NTOBU I .
b OYNH f h 0 I I
,i OBut ' e’f’g
c T: i.
h a b ,FZM /

_ a c 0 d e g . _ .
(b) $/\/\OJ\K\/\/NH3 02ch3 I
b +NH3 f h ‘i
. f,g

 

 

I' . 'ozccr3
all .1
'I‘ d h a’e b '
C , , l
I / «\f‘w " 'lw
_.._____..........~’ “\ , . -_,~___’_ ~J.._.—/' «‘h _ ,-~- __,___/\____________,_._.—/\.____.._-~'/ “"‘J.
‘ ' '1 If T "T I a I "" i Y r ‘ ‘ ‘ ‘ 7' ’ 'fi "—'f T fi—' —' — TT‘M'T " ‘7 l 7"". 7“? T fi‘ ""T 7" ' '#7 'T—l
4.0 3.0 2.0 1.0 ppm

5.0

Figure 51. 1H NMR (300 MHz, CDCl3) spectra of (a) PLLA-g-BocLysPr; N,N'-diBoc-lysine-esterified

PLLA-g-HP and (b) PLLA-g-LysTFAPr; lysine bis(trifluoroacetate)-modified PLLA-g—HP.

70

NW

C 0 d e _ CH3

_ W0 WWW WNW, 020CF3 l:
WW? +NH3 f h I ‘

   

 

OzCCFg l I
. l '
d -thf l
i -. I
i1 I, 3 i
l: I! l
‘x d7'thf l' II I
ii“ I; I f . fig
; .1 l 31 I '1
' II 5 i. i l
l . . i
i I C 4‘ h a b we]; 1
’ . d fl f“ V ”J l
‘I \ l “i H W"
“M; KL ._..___../ w"? W \anw
”‘T a F." T‘—'_ "_‘ V "‘ “TT T‘ Til fi—F'T—Tl’ " 'T—l ‘I—w—V é_ ‘I——'__ _.’—T_ j—TT— “— “—_r T "- "T—FT‘ —'_ i
5.0 4.0 3.0 2.0 1.0 ppm

Figure 52. 'H NMR (300 MHz, dg'thf) spectrum of PLLA-g-LysTFAPr; lysine bis(trifluoroacetate)-

modified PLLA-g-HP.

Table 6 shows the results of molecular weight and thermal analyses of the
phosphonic acid and lysine-modified copolymers. In the other copolymers, no apparent
decomposition or degradation of the polymer was noted under our DCC coupling or
deprotection steps. The DSC profiles for these copolymers change once the phosphonate
and the lysine modifications are made. The crystallinity is lost, and a small decrease in
the Tg is observed for the protected and deprotected copolymers compared to the PLLA-

g-HP copolymer.

71

 

Copolymer M,/ gmof’(PDI)“'b T; / ”C % Yield

 

PLLA-g-DPAPr 13,000 (1.45) 40 88
PLLA-g-PAPr 12,000 (1.37) 57 71
PLLA-g—BocLysPr 17,800 (1.42) 54 70
PLLA-g-LysTFAPrd -- 47 95
PLLA-g-HP" 13,500 (1.58) 54 75

 

Table 6. Protected and deprotected phosphonic and amino acid-modified PLLA copolymers containing 6
mol % functional groups. a Molecular weight determined by GPC. b PDI = Polydispersity index. "
Differential scanning calorimetry of modified copolymers containing 6 mol % functionality. T8 was
obtained after the second heating scan at 10°C/min under helium atmosphere. Molecular weight was not

observed by GPC. d Independently prepared 6 mol % hydroxypropane-modified copolymer.

Successful preparation of PLLA copolymers containing pendant hydrophobic
(fatty acids) and pendant hydrophilic (phosphonic and amino acid) groups was achieved
by DCC coupling and deprotection, where necessary, of commercially available and
biologically relevant starting materials. Spectroscopic characterization of the copolymers
confirm the transformation and covalent attachment of these diverse functional groups.
DSC analyses show the influence of the tethered substrates on the Tg of the material
compared to its copolymer precursors.

Our goal in creating a simple method of functionalization was achieved by
designing a simple comonomer and using simple organic chemistry such as
hydroboration and DCC coupling to covalently attach functionally diverse bioactive
substrates. The usefulness of this methodology is underscored by the simplicity with
which widely biologically-diverse materials can be made. The evaluation of these
materials in tissue engineering applications is the next step to determine where our

materials best fit into this developing area of research.

72

CHAPTER THREE

3 Cell Biology on Modified Poly(Lactic Acid) Surfaces

3.1 Copolymer Thin Films

In order to assess the physiology of our copolymers we prepared thin films to use
as surfaces for tissue growth studies. Thin films were prepared by spin-coating the
polymer on freshly cleaned polished silicon wafers. Osteoblasts were seeded on the
polymer-coated wafers and their proliferation, differentiation, and shape during the
culturing process were assessed.

The functional copolymers from which the films were prepared were chosen for
their potential interaction with osteoblasts. Osteoblasts are cells that are responsible for
the production of bone. The osteoblasts first proliferate (divide) and then reach a critical
point where they cease to proliferate and begin to differentiate and secrete the
extracellular matrix (ECM). The matrix provides an environment where cells can deposit
minerals, which ultimately comprise the material in bone formation. In choosing
substrates for our copolymer modification, we were hoping to influence proliferation and
differentiation by creating a unique surface over which the cells can spread. The
geometric shape of cells has been shown to greatly influence their proliferation and
differentiation.‘”'43 We have prepared these copolymers with unique functionality with
the hypothesis that they will interact with the cells to alter and influence their shape.

To determine whether or not our polymer surfaces possessed any functionality on
the surface, we characterized the surfaces of thin films using ellipsometry and contact
angle measurements (Table 7). The thicknesses of the films were uniform on a given

substrate, but varied slightly between copolymers. All of the films prepared were spin-

73

coated from a 1% (w/w) solution of the polymer in toluene. The film thicknesses were

measured using ellipsometry and had thicknesses generally around 50 nm.

 

 

Copolymer Loading 0 Thickness/ nm b Bad, ‘ 6,“ ‘
PDLLA -- 56 a: 0.5 70 i: 2.1 51 i: 1.7
PLLA-g-HP 6% 70 i 1.5 84 :l: 2.8 50 .4: 2.6
LLA-g-MyPr 6% 47 :8 0.8 82 :8 0.9 52 :i: 2.4
PLLA-g-StPr 6% 48 :i: 1.1 85 d: 1.1 50 :l: 2.7
PLLA-g-OlPr 6% 47 :l: 4.7 78 1: 0.7 53 i 1.4
PLLA-g-LnPr 6% 49 i 1.1 78 i 1.7 52 :1: 1.7
PLLA-g-ArPr 6% 42 :t 3.2 82 :l: 1.7 45 :t 3.2
PLLA-g-DPAPr 6% 45 i 0.5 75 i 1.5 37 :1 1.2
PLLA-g-PAPr" 6% 96 :l: 4.0 73 :t 1.1 35 :l: 1.5
PLLA-g-BocLysPr 6% 52 i 0.9 79 i 1.0 39 i 1.9
PLLA-g-LysTFAPrd 6% 68 :1: 3.3 69 :t 2.4 27 a: 0.6

 

Table 7. Thin films of copolymers spin-coated on l-inch diameter silicon wafers fi'om a 1% (w/w)
solution in toluene unless otherwise noted. Spin rate: 2500 rpm, spin time: 40 sec. " Mole percent loading
determined by IH NMR spectroscopy. b Determined by ellipsometry. ‘ Samples were prepared by heating
above the polymer melting point and quenching to room temperature to obtain amorphous samples.
Advancing (604,.) and receding (6,“) contact angles were measured at a rate of 3 uL/s up to a total volume
of 30 11L using Milli-Q grade water. d Copolymer were dissolved as a 1% (w/w) solution in
tetrahydrofuran.

The wettability of the polymer surface was monitored by recording the advancing
and receding contact angles of a droplet of Milli-Q grade water. The polymer samples
were all heated 160°C under an atmosphere of nitrogen then immediately placed on a
clean surface at room temperature in order to ensure a common sample history. Five
separate contact angle measurements were taken on each polymer-coated silicon wafer,

with the error set as one standard deviation from the mean. Compared to PLLA-g-HP,

the change in the advancing and receding contact angles for the esterified fatty acids is

74

not significant, suggesting that the hydrophobic grafis may not be present near the surface
of the film. The contact angle values compare well with side-chain fatty esters of
polysaccharides”5 and endgroup fatty esters of polylactides.236 Conversely, the more
polar phosphonate and lysine derivatives result in a notable decrease in the advancing and
receding contact angles, suggesting that the polar groups may reside close to the surface
of the film. For our lysine derivatives specifically, the advancing and receding contact
angles compare well with the PLLA-co-lysine polymers synthesized by Langer.138 The
contact angles reflect the wettability of each of the copolymer surfaces and may influence

cell shape.

3.2 Osteoblast Cell Shape

We have assessed the biocompatibility of our copolymers by seeding osteoblasts
on three different surfaces: PLLA, silicon oxide, and phosphonate-modified PLLA
(PLLA-g-PAPr). In view of the wide applicability of biodegradable polyesters used as
biomaterials, our modified PLLA copolymers are more suitable candidates to be used for
in vivo applications. Therefore, by first assaying the in vitro cell growth behavior on the
surfaces of modified PLLA copolymers, we hope to obtain information on osteoblast cell
growth in the presence of these materials.

MC3T3-El osteoblasts were seeded at a concentration of 4400 cells/cmz, cultured
for 2 days in a-MEM (minimal essential medium) containing 2 mM B-glycerolphosphate
and 25 ug/ml ascorbic acid, after which they were stained and visualized in order to
observe their appearance on the silicon oxide wafers, and the phosphonate-modified
PLLA. The differences in morphology of these sets of cells show how the osteoblasts

spread on each of the surfaces. Cells cultured on the cleaned silicon wafers are more

75

spread out, while the PLLA-g-PAPr coated wafers are intermediate of the other surfaces,
possessing characteristics of both the tissue culture plates and the silicon wafers.
Clearly, the silicon wafers and the polymer-coated wafers alter the shape of the
osteoblasts, providing positive evidence that the wafers and the polymer are influencing
the conformation of the cells and therefore, may be able to influence osteoblast

proliferation and differentiation.42

   

o 9
(a) Silicon oxide (b) EMOJK/HOHh
5% phosphonate

Figure 53. Microscope images of osteoblasts cultured for 48 hours on (a) silicon oxide and (b) 5%

phosphonate modified PLLA surfaces.
3.3 Osteoblast Proliferation

The quantification of cell proliferation can be carried out by measuring the DNA
of a dividing cell. This information is obtained by incorporating a fluorescent-labeled
DNA base pair, allowing the cells to divide, and watching the incorporation of the

fluorescent label. As the cells continue to replicate, the fluorescent label multiplies and is

measured (Figure 54). The fluorescent label used is Bromodeoxyuridine (BrdU).

76

Mama

0 = Adenosine
0 = Thymidine
0 = Bromodeoxyuridine

Figure 54. Illustration of cell replication incorporating the BrdU fluorescent label.

The cells were cultured for 48 hours and then incubated for 2 hours with BrdU.
The BrdU incorporated into the cell nuclei at the S phase of dividing cells. The
osteoblasts were then stained with propidium iodide, which was used to stain all cell
nuclei. Osteoblast proliferation on unmodified PLLA was poor due to the because of the
hydrophobicity of the polyester. However, the results on the other surfaces show that
proliferation on the surfaces of silicon oxide and phosphonate-modified PLLA is
essentially the same (Figure 55). The cell density viewed in the microscope images is
much higher indicating rapid cell division before the BrdU assay at 48 hours. Therefore,
while the BrdU assay shows no significant difference in proliferation between silicon
oxide and the phosphonate-modified PLLA surface, the phosphonate-modified PLLA

surface may have induced a proliferation burst within the first 48 hours.

77

Nuclei undergoing
Total Nuclei DNA replication

Silicon
a
() OXide - -
“Mo/936011):
5% phosphonate

0= Adenosine
O= Bromodeoxyuridine

Figure 55. Digital photo images of fluorescent BrdU-labeled osteoblasts on (a) silicon oxide and (b) 5%

phosphonate modified PLLA surfaces.

3.4 Osteoblast Differentiation

The process through which bone tissue forms begins with the cells’ ability to
differentiate. As cell proliferation precedes differentiation, longer culture times are
required to assess differentiation. As differentiation occurs, the osteoblasts form the
matrix for mineralization, and ultimately bone formation. After 14 days, RNA analysis
quantifies levels of expressed genes alkaline phosphatase, osteocalcin, and transcription
factor runx2 relative the internal control gene cyclophilin. The genes serve as markers to

quantitate osteoblast differentiation (Figure 56).

78

Osteocalcin

"'runx2

Alkaline
Phosphatase

% Expression

 

Days in culture

Figure 56. Gene expression levels of osteocalcin, runx2, and alkaline phosphatase as a function of
osteoblast culture time.

After 14 days, the RNA of the cells was sampled and monitored for runx2 levels.
Comparing the four surfaces, the phosphonate-modified copolymers showed a marked
increase in differentiation over the unmodified PLLA, while it also exceeds

differentiation on the silicon oxide (Figure 57).

     

0.6
I: t:
.9 .9
E: 8
g :9“ 0.4
i=3 §
'0 .M
H 0.2
9 E
i?
75' 0

PLLA Silicon PLLA-g-PAPr PLLA-g-PAPr
Oxide

fl/voUm. Mk».

3% phosphonate 5% phosphonate

Figure 57. Surface comparison of osteoblast differentiation on different surfaces.

79

The increase in the observed differentiation from 3 mol % to 5 mol %
phosphonate suggests that under identical conditions for all of the surfaces, the
phosphonate groups are largely responsible for the increase. This result is consistent with
our hypothesis that the pendant phosphonate groups on the copolymer may help support
mineralization of the ions deposited (e. g. hydroxyapatite) during differentiation.

Our data has presently focused on physiology of osteoblasts on polymer-coated
silicon wafers which are essentially smooth surfaces. Since surface roughness is known

to be influential in regulating osteoblast proliferation and differentiation on polymer

237-239 220,240-242

surfaces and metal surfaces, similar experiments to the ones described on
rough surfaces may reveal differences in osteoblast proliferation and differentiation.
4 Conclusion

The post-polymerization modification of PLLA-co-AG using olefin cross
metathesis, and hydroboration-oxidation followed by the synthesis of ester using DCC
coupling has proven to be an effective way to screen substrates for biological activity
when tethered to biodegradable polymers such as PLLA. The synthetic ease with which
the modification takes place coupled with the commercial availability and diversity of the
functional side chains, make this modification widely applicable to the fields of tissue
engineering and drug delivery. Furthermore, the initial experiments conducted on the
physiological response to these functional copolymer surfaces appear to have potential in
the design of in vivo biodegradable materials.

Regardless of the modification, the common objective throughout the described

work is to design materials which will help regulate cell adhesion and influence the cell

conformation, and consequently, the function of the biological system. We have

80

designed a protocol which will allow us to fulfill this objective. Post-polymerization
modification has expanded the physiological utility of PLA in the area of tissue

engineering.

81

 

5 Experimental Section.

5.1 General Considerations.

1H NMR spectra were recorded on Varian Gemini—300 and Inova—300 (299.9
MHz) and Varian Unity-500 (499.9 MHz) spectrometers and referenced to residual
proton solvent signals. '3 C NMR spectra were recorded on Varian Gemini—300 and
Inova—300 (75.4 MHz) and Varian Unity-500 (125.7 MHz) spectrometers and referenced
to residual solvent signals. “P NMR spectra were recorded on a Varian Inova~300
(121.5 MHz) and referenced using H3PO4 as the external standard. 11B NMR spectra
were recorded on a Varian VXR—300 (160.6 MHz) and referenced using BF 3.0Et2 as the
external standard. 19F NMR spectra were recorded on a Varian Inova—300 (289.4 MHz)
and referenced using CFCl3 as the external standard. Differential Scanning Calorimetry
(DSC) analyses of the polymers were performed under a helium atmosphere at a heating
rate of 10°C/min on a Perkin-Elmer DSC 7, with the temperature calibrated using an
indium standard. The reported DSC curves are second heating scans, taken after an
initial heating to erase thermal history, and a slow cooling to at 10°C/min to -50°C.
Molecular weights of the polymers were determined by gel permeable chromatography
(GPC) using a Beckman 100A instrument, equipped with a Waters 410 differential
refractometer detector and a Waters 960 photodiode array detector. Measurements were
obtained in THF with a PLgel 5 u Mix column (Polymer Laboratories) at a flow rate of
1.0 mL/min at 35°C. Results were calibrated with monodisperse polystyrene standards.
Polished silicon wafers purchased from WaferWorldTM were cleaned by immersion in a
70:30 volume ratio solution of concentrated H2804 (98% by mass) and H202 (30% by

mass) and rinsed in deionized water. Polymers were dissolved in toluene (1% w/w),

82

filtered two times through 0.2 pm PTFE filters and spin-coated onto polished silicon
wafers (1" diameter) at a spin rate of 2500 rpm for 40 s. Ellipsometric measurements
were obtained with a rotating analyzer ellipsometer (model M-44, J .A. Woollam) using
WVASE32 software. The angle of incidence was 75° for all measurements. For the
calculation of the film thickness, a film refractive index of 1.50 was used.

Contact Angle Measurements. After spin-coating the polymers onto silicon
wafers, the wafers were held under vacuum at 40°C for 12 hours to ensure all the solvent
was removed from the wafers. The polymer-coated wafers were kept at room
temperature for 1 hour under a constant purge of nitrogen. Samples were heated under a
nitrogen atmosphere at a rate of 10°C/min to 160°C (15°C above the melting point of the
all polymer samples) and held for 1 minute. The samples were immediately removed
from the oven and placed on a clean surface at room temperature. Advancing and
receding contact angles were measured by dispensing a droplet of Milli-Q grade water
(18 M9 cm) at a rate of 3 uL/s to a total volume of 30 uL on the surface of polymer-
coated silicon wafers. Contact angles were measured by capturing the image as a movie
with a Sanyo® B/W CCD camera (model VC8 3512). The angles were measured using
the FTA200 software.

Cell Culture. MC3T3-E1 osteoblasts (OBS) were plated on silicon discs
(WaferWorldTM) at a concentration of 4,400 cells/cmz. These were either cultured for 2
days for the proliferation studies or 14 days for the differentiation studies, where at this
stage OBS exhibit maximal levels of alkaline phosphatase and mineralization). Cells
were fed every two days with a-MEM containing 2mM B-glycerolphosphate (Sigma, St.

Louis, MO) and 25 ug/ml ascorbic acid (Sigma).

83

 

Cell Growth and Shape. After plating for 48 hours, osteoblasts were incubated
with 10 uM BrdU (5-Bromo-2’Deoxyuridine) (Sigma, St. Louis, M0) for two hours.
Subsequently, cells were fixed with 70% ethanol. The cell culture plates were then spun
at 1500 rpm for 5 minutes to ensure all cells remained at the bottom of the plate, followed
by quick rinse with 1Xphosphate buffered solution (IXPBS). Cells were permealized
with 4 N HCl in IXPBS. Subsequently washed gently with 1 mL of IXPBS until the PBS
in the wells reached a pH=7. Osteoblasts were further incubated in 20% anti-BrdU (BD
Biosciences, San Jose, CA) antibody solution, in IXPBS, 0.5% BSA, 0.1% Tween-20,
for 45 minutes at 37°C under humid conditions. Brdu antibody was removed and cells
further introduced to propidium iodide stain at 200 pg/ml PBS (BD Biosciences, San
Jose, CA) immediately after which nuclei of the cells were visualized by fluorescence
microscopy and recorded with a digital camera. BrdU stains the nuclei at S phase of cell
cycle, i.e. when cells are dividing, and therefore, used to determine cell growth.
Propidium iodide stains all nuclei.

Actin and Hoechst Stain. Osteoblasts were fixed with 3.7% formaldehyde for
10 minutes at room temperature, 2 days after seeding, and subsequently permealized for
5minutes with 0.1% Triton X-100 in IXPBS. Cells were blocked with 1% bovine serum
albumin for 30 minutes and subsequently incubated for 20 minutes with rhodamine-
pahlloidin (Molecular Probes Inc. Euegene, OR) and further incubated with at 10 uM
stain in deionized H20, for 30 minutes. Actin localization and Hoechst stained
(Molecular Probes Inc. Eugene, OR) nuclei were visualized by fluorescence microscopy

and recorded with a digital camera.

84

 

RNA Analysis. RNA was extracted, on day 14, using TRI Reagent RNA
isolation reagent (Molecular Research Center, Inc., Cincinnati, OH). Integrity of the RNA
was verified by formaldehyde-agarose gel electrophoresis. Two-step quantitative RT-
PCR was performed to verify gene expression. First strand cDNA was synthesized by
reverse transcription of 2 ug RNA, using the Superscript II Kit and oligo d(T12-18)
primers (Invitrogen Life Technologies, Carlsbad, CA). 1 ul of cDNA was amplified by
Real Time PCR, by using the SYBR Green Core Reagents (PE Biosystems, Warrington,
UK) and Taq DNA polymerase (Invitrogen Life Technologies, Carlsbad, CA) with a final
volume of 25 ul. Genes associated with osteoblast differentiation, alkaline phosphatase,
osteocalcin, and runx2, were analyzed using cyclophilin as the internal control. Real time
PCR was canied out and analyzed for 40 cycles, using iCycler software (Biorad, Hecules,
CA). Gel electrophoresis and melting curves were used to verify the integrity of a single
PCR product (amplicon).

Copolymer Degradation. In a typical degradation, 0.20 g of polymer were
suspended in 15 mL of ethanol and Sn(Oct)2 (0.20 g, 0.49 mmol). The mixture was
refluxed for 48 hours or until complete degradation as determined by lH NMR. The
solution was then directly analyzed by GC/MS for the ethyl esters of L-lactic acid, 2-
hydroxypent-4-enoic acid, and 2,7-dihydroxyl-oct-4-enoic-1,8-diacid (cross-link
degradation product).

Solvents. Toluene, tetrahydrofuran, diethyl ether, and pentane were pre-dried
over sodium and distilled from sodium/benzophenone ketyl. Dichloromethane was
distilled from calcium hydride (CaHz). All other reagents were used as received unless

otherwise noted.

85

 

 

5.2 Synthesis

0

NOH
OH

2-Hydroxypent-4-enoic acid (2-HPA).243 Glyoxylic acid monohydrate (4.80 g,
0.053 mol) and fieshly distilled allyl bromide (9.44 g, 0.078 mol) were added to a stirring
suspension of zinc powder (7.12 g, 0.109 mol), and BiCl3 (24.6 g, 0.078 mol) in 60 mL
tetrahydrofuran. The reaction was initially kept at 0°C and allowed to warm to room
temperature while stirring under N2 for 16 h. After 16 h, the reaction is quenched with 40
mL of l M HCl, and the product is extracted with diethyl ether (3x80 mL). The organic
phases were decanted from the solid, combined, and washed with saturated NaCl(aq) until
the ether solution became colorless (5x20 mL). The ether layer was then dried over
MgSO4, and the solvent removed to give a colorless oil which crystallized under vacuum.
The a-hydroxy acid was then recrystallized from EtZO/Hexanes. Yield: 93%. 1H NMR
(300 MHz, CDCI3, 6): 5.65-5.85 (m, 1H, -CHCH2CH=CH2), 5.0-5.2 (m, 2H, -
CHCHzCH=CH2), 4.27 (dd, 1H, H02CCH(OH)CH2CH=CH2), 2.35-2.65 (m, 2H, -

CHCHzCH=CH2). 13C NMR (75.0 MHz, CDCl3, 8): 177.3, 132.2, 118.8, 69.7, 38.2.

3,6-Diallylglycolide (DAG). 2-HPA (3.0 g, 0.025 mmol), and a catalytic amount
(30 mg) of p-toluenesulfonic acid (stOH) were dissolved in 400 mL of toluene. The
mixture was refluxed for 3 days and the water removed by Dean-Stark trap. After 3 days
the mixture was cooled, washed 3 times with 10 mL saturated aqueous NaHCO3 and the

resultant oil distilled over ZnO under high vacuum at 80°C/0.01 mmHg. Gas

86

 

chromatography data indicates a statistical mixture of S/S, R/S, and R/R isomers of the
diallylglycolide. Yield: 40%. 1H NMR (500MHz, CDCl3, 6): 5.8 (m, 1H, -
CHCHZCH=CH2), 5.1-5.3 (m, 2H, -CHCH2CH=CH2), 5.00 (dd, 1H, -CHCH2CH=CH2, J
= 5.1 Hz), 4.91 (dd, 2H, *-CHCH2CH=CH2, J = 4.6 Hz), 2.60-2.90 (m, 4H, -
CHCHZCH=CH2). l3C NMR(125 MHz, c0013, 8): *166.1,164.8,*130.6,129.8, 121.2,
*120.0, 75.9, *75.1, 36.4, *34.1. LRMS (El, m/z): 197 (MW), 98, 81, 54, 41. Anal.
Calcd for C10H1204: C, 61.21; H, 6.18. Found: C, 61.29; H, 6.17. *Indicates RR/SS
isomer. (Note: Using a combination of L-lactic acid and 2-HPA in the desired ratio under
identical reaction conditions and workup results in a statistical mixture of L-lactide, 3-

allyl-6-methylglycolide, and 3,6-diallylglycolide and appropriate stereoisomers.)
l
o
0):”);
o
l
Polymer Synthesis. Poly(diallylglycolide) (PDAG). In a glovebox,
diallylglycolide (0.163 g, 0.832 mmol) was combined in a Schlenk flask with tin(II)-2-
ethylhexanoate (0.005 mmol) as the catalyst. The reaction tube was placed in an oil bath
at 80°C and stirred for 72 h. The reaction mixture was pumped dry, dissolved in the
minimal amount of CHzClz and precipitated from cold methanol two times. Yield: 90%.
1H NMR (500 MHz, CDCl3, 8): 5.8 (m, 1H, -CHCH2CH=CH2), 5.0-5.2 (m, 3H, -

CHCHZCH=CH2), 2.5-2.8 (m, 2H, -CH2CH=CH2). l3C NMR (125 MHz, CDCl;, 8):

l68.l,13l.6, 119.1, 72.0, 35.2.

87

11M

Poly(L-lactide-co-diallylglycolide) (PLLA-co-AG). In a typical synthesis of
PLLA-co-AG, freshly sublimed L-lactide (2.88 g, 20 mmol) and distilled diallylglycolide
(DAG) were combined in a glass tube with tin(Il)-2-ethylhexanoate (0.067 mmol) and 4-
tert-butylbenzylalcohol (BBA) (0.067 mmol) as the catalyst and initiator, respectively.
The reaction tube was flame-sealed under vacuum and placed in an oil bath at 145°C.
After 15 minutes, the sample was removed from the oil bath, cooled, and monitored for
conversion. Crude 1H NMR revealed nearly complete conversion (> 90%). Polymers
were dissolved in a minimal amount of CHzClz, and precipitated from cold methanol two
times. Yield: 90%. 1H NMR (500 MHz, CDCI3, 5): 5.8 (m, 1H, -CHCH2CH=CH2),
5.05-5.25 (q, *-CH(CH3), -CHCH2CH==CH2), 2.5-2.8 (m, 2H, -CH2CH=CH2), 1.4-1.7 (d,

*-CH(CH3)). 13C NMR (125 MHz, CDCl3, 8): *169.6,168.4,131.4, 119.1, 71.9, *69.0,

H 0
W06

2-(2-Propenyloxy)benzaldehyde.244 In a 500 mL round bottom flask,

35.2, *16.6. *PLLA.

salicylaldehdye (14.6 g, 0.152 mol) and allyl bromide (23.0 g, 0.191 mol) were dissolved
in 200 mL of acetone. Anhydrous potassium fluoride (35.4, 0.61 mol) was then added
and the mixture refluxed for 72 h. After 72 h., the reaction mixture was filtered, then the
solvent removed by rotary evaporation. The resultant oil was distilled twice at 75°C/0.1
mmHg to give pale yellow oil. Yield: 77%. 1H NMR (300 MHz, CDC13, 5): 10.5 (s,
1H, -C(O)H), 7.77 (dd, 1H, -HA,), 7.47 (m, 1H, JIM), 6.9-7.0 (m, 2H, -HA,), 6.02 (m,

1H, -CH=CH2), 5.24-5.44 (m, 2H, -CH=CH2), 4.59 (q, 2H, -OCH2CH=CH2). 13C NMR

88

 

(75 MHz, CDCI3, 8): 189.5, 160.7, 135.7, 132.2, 128.1, 124.8, 120.6, 117.8, 112.7, 68.9.
LRMS (EI, m/z): 163 (MH+), 161, 121.
Wows

9-Decenyl-l-oxy-tert-butyldimethylsilane.245 In an 250 mL Erlenmeyer flask, 9-
decen-l-ol (2.0 g, 12.8 mmol) was dissolved in 25 mL CHzClz. To this solution was
added Chloro-tert—butyldimethylsilane (2.12 g, 14.1 mmol) in 15 mL CHzClz. The
reaction was stirred for 8 hours at room temperature under a nitrogen atmosphere. After
8 hours, the solvent is removed under rotary evaporation. The product was extracted with
EtzO, filtered and dried in vacuo. The oil was purified by flash column chromatography
through silica gel using hexanes/EtOAc (96/4) as the eluent. The product was obtained
as an oil. Yield: 95%. 1H NMR (300 MHz, CDCI3, 5): 5.79 (m, 1H, -CH=CH2), 4.90-
5.00 (m, 2H, -CH=CH2), 3.58 (t, 2H, -OCH2C9H17), 2.01 (q, 2H, -CH2CH=CH2), 1.49
(m, 2H, -OCH2CH2C3H15), 1.2-1.4 (m, 10H, -(CH2)2(CH2)5CH2CH=CH2), 0.874 (s, 6H,
SiC(CH3)3), 0.027 (s, 6H, Si(CH3)2). 13C NMR (75 MHz, CDCl;, 6): 139.2, 114.1, 63.3,
33.8, 32.9, 29.5, 29.4, 29.1, 28.9, 26.0, 25.8, 18.4, -5.6. LRMS (El, m/z): 271 (MHI),

255, 214.

O

W”
9-Decen-1-al.246 In a round bottom flask, oxalyl chloride (13.35 mL, 0.03 84 mol)
was dissolved in CHzClz, and cooled to -78°C. DMSO (5.00 mL, 0.0704 mol) is slowly
added neat via syringe and then stirred for 15 min. 9-decen-1-ol (5.0 g, 0.032 mol) in 10
mL of CHzClz is added slowly, after which the reaction becomes cloudy. After stirring
for 15 minutes, NEt3 (22.6 g, 0.224 mol) is added via syringe and stirred for 10 minutes.

The cold bath is removed and stirred to room temperature. The solvent is then removed

89

by rotary evaporation to give a yellow oil, which is first purified by flash chromatography
through silica gel using hexanes as the eluent. The hexanes are removed by rotary
evaporation. The oil is further purified by column chromatography using hexanes/EtOAc
(80/20). Yield: 90%. 1H NMR (300 MHz, CDC13, 6): 9.57 (t, 1H, -C(O)H), 5.61 (m,
1H, -CH=CH2), 4.68-4.86 (m, 2H, -CH=CH2), 2.23 (m, 2H, -CH2C(O)H), 1.85 (m, 2H, -
CHzCH=CH2), 1.43 (m, 2H, -CH2CH2C(O)H), 1.05-1.25 (m, 8H, -(CH)4CH2CH=CH2).
13C NMR (75 MHz, CDC13, 6): 202.8, 138.9, 114.1, 43.78, 33.64, 29.09, 28.98, 28.77,

28.69, 21.92. LRMS (131, m/z): 155 (Mir), 135, 121,95, 81.

/ O

9-decenyl-l-oxy-(4-p-nitrobenzene). In a 500 mL round bottom flask, p-

 

nitrophenol (2.0 g, 14.4 mmol), 9-decene-1-tosylate247 (4.9 g, 15.8 mmol), K2CO3 (2.09
g, 15.12 mmol), and a catalytic amount of K1 were refluxed for 2 days in 300 mL of dry
acetone. After 2 days, the reaction is filtered and washed with CHzClz. The filtrate was
concentrated, washed with saturated NaHCO3, and extracted with CHzClz. The organic
phases were then stirred with anhydrous NaZSO4, filtered, and the solvent removed to
give an orange oil. The oil was purified by flash column chromatography through silica
gel using Hexanes/EtOAc (97/3) as the eluent. The resultant yellow oil crystallized upon
standing. Yield: 86%. lH NMR (300 MHz, CDC13, 6): 8.13 (d, 2H, -HA,), 6.89 (d, 2H,
-HA,), 5.76 (m, 1H, -CH=CH2), 4.85-5.00 (m, 2H, -CH=CH2), 4.00 (t, 2H, -OCH2C9H17),
2.00 (q, 2H, -CH2CH=CH2), 1.78 (m, 2H, -OCH2CH2C3H15), 1.2-1.5 (m, 10H, -
(CH2)2(CH2)5CH2CH=CH2). 13C NMR (75 MHz, CDCl:,, 6): 164.0, 141.0, 138.8, 125.6,

114.1, 113.9, 68.6, 33.5, 29.1, 29.0, 28.8, 28.7, 28.6, 25.6. LRMS (El, m/z): 278 (MH+),

90

232, 123. Anal. Calcd for CI6H23O3N: C, 69.27; H, 8.37; N, 5.05 Found: C, 69.23; H,

8.29; N, 5.13.

PLLA-g-AB (AB = Allylbenzene). In a small round-bottom flask, PLLA-co-AG
was dissolved in CH2C12 (0.01 M) with approximately 5 equivalents of the olefin
substrate. Grubbs’ first or second generation ruthenium catalyst was added to the
solution. An excess of olefin was introduced by a slow syringe pump addition (0.10
mLh"). Reaction was carried out for 24 h, quenched with ethyl vinyl ether and
precipitated multiple times from hexanes and methanol. Yield: 90%. 1H NMR (300
MHz, CDC13, 6): 7.5-7.1 (br, 5H, -CH2C6H5), 5653 (br, 2H, -CH=CH-), 5.3-5.0 (m, *-
CH(CH3), -CH(CH2CH=CHC7H7)), 3.33 (m, 2H, -CH2C6H5), 2.8-2.5 (m, 2H, -

CHzCH=CHC7H7), 1.8-1.4 (m, *-CH(CH3)). * PLLA.

H o
1W6

PLLA-g-PB (PB = 2-(2-propenyloxy)benzaldehyde). Polymer-bound olefin

cross metathesis reaction conditions were identical to PLLA-g-AB. After 24 h, the
reaction was quenched with ethyl vinyl ether and precipitated multiple times from
methanol. Yield: 63%. lH NMR (300 MHz, CDCI3, 6): 10.45 (s, 1H, -C(O)H), 7.80
(m, 1H, -HA,), 7.49 (m, 1H, -HA,), 7.1-6.9 (m, 2H, -HA,), 5.9-5.6 (m, 2H, -CH=CH), 5.3-
5.0 (m, *-CH(CH3), -CH(CH2CH=CHC7H7)): 4.7-4.6 (m, 2H, -CH20C6H4(C(O)H)), 2.8-

2.5 (m, 2H, -CH2CH=CHCH20C6H4(C(O)H)), 1.8-1.4 (m, *-CH(CH3)). * PLLA.

91

1W0-

PLLA-g-decenyloxy-TBS (Decenyloxy-TBS = 9-decen-l-oxy- tert-
butyldimethylsilane). Polymer-bound olefin cross metathesis reaction conditions were
identical to PLLA-g-AB. After 24 h, the reaction was quenched with ethyl vinyl ether
and precipitated from hexanes then precipitated from methanol. Yield: 26%. 1H NMR
(300 MHz, CDC13, 6): 5.5-5.3 (br, 2H, -CH=CH(CH2)gOTBS), 5.3-5.0 (m, *-CH(CH3), -
CH(CH2CH=CH(CH2)gOTBS), 3.57 (t, 2H, -CH20TBS), 2.8-2.5 (m, 2H, -
CHzCH=CH(CH2)3OTBS), 1.96 (m, 2H, -CH=CHCH2C7H14OTBS), 1.8-1.4 (m, *-
CH(CH3)), 1.26 (m, 12H, -(CH2)6CH20TBS), 0.868 (s, 9H, Si-C(CH3)3), 0.021 (s, 6H,

Si(CH3)2. * PLLA.

HEW“

PLLA-g-decenal. Polymer-bound olefin cross metathesis reaction conditions
were identical to PLLA-g-AB. After 24 h, the reaction was quenched with ethyl vinyl
ether and precipitated from hexanes then precipitated from methanol. Yield: 48%. 1H
NMR (300 MHz, CDCI3, 6): 9.74 (m, 1H, -C(O)H), 5.5-5.3 (br, 2H, -CH=CH-
C7H14C(O)H), 5.3-5.0 (m, *-CH(CH3), -CH(CH2CH=CH C7H14C(O)H)), 2.9-2.5 (m, 2H,
-CH2CH=CH-), 2.39 (t, 2H, -CH2C(O)H), 1.96 (m, 2H, -CH=CHCH2C6H12C(O)H), 1.8-

1.4 (m, *-CH(CH3)). 1.26 (m, 10H, -(CH2)5CH2C(O)H). * PLLA.

PLLA-g-decenyloxy- NB (Decenyloxy- NB = 9-decenyl-l-oxy-(4-

nitrobenzene). The polymer-bound olefin cross metathesis reaction conditions were

92

identical to PLLA-g-AB. After 24 h, the reaction was quenched with ethyl vinyl ether
and precipitated from hexanes then precipitated from methanol. Yield: 79%. 1H NMR
(300 MHz, CDC13, 6): 8.17 (d, 2H, -HA,), 6.92 (d, 2H, -HA,), 5.6-5.2 (br, 2H, -CH=CH),
5.3-5.0 (m, *-CH(CH3), -CH(CH2CH=CH(CH2)8OC6H4(N02)), 4.02 (t, 2H, -
CH2OC6H4(N02)), 2.8-2.5 (m, 2H, -CH2CH=CH(CH2)30C6H4(N02)), 1.96 (m, 2H, -
CH=CHCH2C7H14OC6H4(NO2)), 1.79 (m, 2H, -CH2C6H12OC61-14(NO2)), 1.7-1.4 (m, *-

CH(CH3)), 1.30 (m, 10H, -(CH2)5CH2OC6H4(NO2)). * PLLA.

18W?

Hydroboration of PLLA-co-AG. The hydroborated copolymer was monitored
using ”B NMR (160 MHz). In an NMR tube reaction, PLLA-co-AG (0.137 g, 2.59
mmolg", 0.355 mmol) was dissolved in CDCI3, after which 9-BBN (0.417 g, 0.342
mmol) was added. The reaction was gently warmed at 35°C for 2h. A broad resonance

centered at 88 6 in the 11B NMR spectrum indicated a trialkylboron moiety and therefore,

PLLA-g-HP (Hydroboration-Oxidation of PLLA-co-AG). In a round bottom

confirming the hydroboration step.

flask, PLLA-co-AG (4.05 g, 1.19 mmolg", 4.82 mmol) was dissolved in 100 mL of dry
THF. 9-BBN (9.73 mL, 0.50 M, 4.87 mmol) was slowly added to the polymer solution
via syringe and stirred at room temperature for 1.5 h. The reaction was then cooled to 0°C
and a 1 M aqueous solution of sodium acetate (9.64 mL, 9.64 mmol) and a 30% solution
of H202 (1.09 g, 9.64 mmol) were added. The mixture was and stirred for 13 h. The

reaction was quenched with 2 M HCl (9.64 mmol) and the polymer solution dried by

93

rotary evaporation. The polymer was then redissolved in CH2Cl2 and further extracted
from water and dried over Na2SO4. The polymer filtrate was concentrated and
precipitated from methanol two times. The polymer was filtered and dried under
vacuum. Avg. Yield (4 times): 60%. IH NMR (300 MHz, CDCl3, 6): 5.25-5.00 (q, *-
CH(CH3), -CHCH2CH2CH2OH), 3.66 (m, 2H, -(CH2)2CH2OH), 2.04 (m, 2H, -
CH2(CH2)2OH), 1.68 (m, 2H, -CH2CH2CH2OH), 1.6-1.4 (d, *-CH(CH3)). 13C NMR (75

MHZ, CDCl3, 6): *169.6, 72.5, *69.0, 61.8, 27.7, 27.4, *16.6. * PLLA.

O

’\/\J\/\/\/\/\/\/\
E o

PLLA-g-MyPr (MyPr = Myristic acid propyl ester). A small round bottom
flask containing PLLA-g-HP (0.352 g, 0.950 mmolg'l, 0.333 mmol), myristic acid (0.114
g, 0.50 mmol), and DMAP (8.0 mg, 0.066 mmol) was stirred in 10 mL of CH2C12 until
completely dissolved. 1,3Dicyclohexylcarbodiimide (DCC) (0.50 mL, 1.0 M, 0.50
mmol) was then added via syringe and the reaction mixture was stirred under N2 at room
temperature. (Note: After 15 minutes, the colorless, homogeneous solution became
cloudy, indicative of the esterification). After 12 h, the reaction mixture was filtered to
remove the insoluble dicyclohexyl urea byproduct, further concentrated, and filtered a
second time to fully remove the urea. The filtrate was then concentrated to dryness,
redissolved in the minimal amount of CH2CI2, precipitated and centrifuged from
methanol two times. The polymer was filtered and dried under vacuum. Avg. Yield (2
times): 89%. lH NMR (300 MHz, CDCI3, 6): 5.2-5.0 (q, *-CH(CH3), -
CH(CH2)3CO2C13H27), 4.06 (m, 2H, -CH2C02C13H27), 2.24 (td, 2H, -CO2CH2C12H25),
1.95 (m, 2H, -CH2(CH2)2CO2C13H27), 1.75 (m, 2H, -CH2CH2CH2CO2C13H27), 1.6-1.5

(m, 2H, -CO2CH2CH2C11H23), 1.56 (d, *-CH(CH3)), 1.23 (m, 20H, -(CH2).0CH3), 0.855

94

(t, 3H, -(CH2)|2CH3). 13C NMR (75 MHz, CDCI3, 6): 173.7, *169.6, 72.1, *69.0, 63.2,
34.2, 33.9, 31.9, 29.6, 29.5, 29.4, 29.3, 29.2, 27.7, 24.9, 24.8, 24.2, 22.7, *16.6, 14.1. *

PLLA.
gmoJk/WW

PLLA-g-StPr (StPr = Stearic acid propyl ester). The polymer-bound DCC
coupling procedure was identical to PLLA-g-MyPr. Avg. Yield (3 times): 88%. 1H
NMR (300 MHz, CDCl;, 6): 5.2-5.0 (q, *-CH(CH3), -CH(CH2)3CO2C17H35), 4.06 (m,
2H, -CH2CO2C17H35), 2.27 (td, 2H, -CO2CH2C16H33), 1.95 (m, 2H, -
CH2(CH2)2CO2C17H35), 1.75 (m, 2H, -CH2CH2CH2CO2C18H37), 1.6-1.5 (d, *-CH(CH3), -
CO2CH2CH2C15H31), 1.23 (m, 28H, -(CH2)14CH3), 0.856 (t, 3H, -(CH2)16CH3). 13C
NMR (75 MHz, CDCl;;, 6): 173.7, *169.6, 72.2, *69.0, 63.2, 34.2, 33.9, 31.9, 29.7, 29.5,

29.3, 29.2, 29.1, 27.6, 24.9, 24.7, 24.2, 22.7, *16.6, 14.1. * PLLA.
EMOWM/W

PLLA-g-OlPr (OlPr = Oleic acid propyl ester). The polymer-bound DCC
coupling procedure was identical to PLLA-g-MyPr. Avg. Yield (2 times): 89%. 1H
NMR (300 MHz, CDCl3, 6): 5.32 (m, 2H, -CH=CH), 5.2-5.0 (q, *-CH(CH3), -
CH(CH2)3CO2C17H33), 4.06 (m, 2H, -CH2CO2C17H33), 2.27 (td, 2H, -CO2CH2C16H31),
1.98 (m, 6H, -CH2CH2CH2CO2C17H33, -(CH2)(CH=CH)(CH2)), 1.75 (m, 2H, -
CH2CH2CH2CO2CI7H33), 1.6-1.5 (d, *-CH(CH3), -CO2CH2CH2C15H29), 1.27-1.24 (m,
20H, -(CH2)4(CH2)(CH=CH)(CH2)(CH2)6CH3), 0.855 (t, 3H, -(CH2)6CH3). 13C NMR

(75 MHz, CDCI3, 6): 173.7, *169.5, 129.9, 129.7, 72.2, *69.0, 63.2, 34.2, 33.8, 31.8,

95

29.7, 29.6, 29.5, 29.3, 29.2, 29.1, 27.6, 27.2, 27.1, 24.9, 24.7, 24.2, 22.6, *16.6, 14.1. *

PLLA.
EMOWM

PLLA-g-LnPr (LnPr = Linoleic acid propyl ester). The polymer-bound DCC
coupling procedure was similar to PLLA-g-MyPr. This polymer can be oxidized, and
therefore should be carried out and worked up in an inert atmosphere where exposure to
oxygen is minimal. Avg. Yield (3 times): 81%. IH NMR (300 MHz, CDCI3, 6): 5.35
(m, 4H, -CH=CH-(CH2)-CH=CH), 5.2-5.0 (q, *-CH(CH3), -CH(CH2)3CO2C17H31), 4.06
(m, 2H, -CH2CO2C.7H31), 2.75 (t, 2H, =CH-(CH2)-CH), 2.4-2.2 (m, 2H,
CO2CH2C16H29), 2.02 (m, 6H, -CH2CH2CH2CO2C17H31, -CH2(C5H6)CH2), 1.76 (m, 2H,
-CH2CH2CH2CO2C17H31), 1.6-1.5 (d, *-CH(CH3), -CO2CH2CH2C15H27), 1.29 (m, 14H, -
(CH2)4(CH2)(C5H5)(CH2)(CH2)3CH3), 0.866 (t, 3H, -(CH2)6CH3). ”C NMR (75 MHz,
CDCI3, 6): 173.7, *169.6, 130.2, 130.0, 128.0, 127.9, 72.2, *69.0, 63.2, 34.2, 32.5, 31.5,

29.6, 29.3, 29.5, 29.3, 29.2, 29.1, 27.6, 27.2, 25.6, 24.8, 24.2, 22.5, *16.6, 14.0. * PLLA.
8M0 _ —— — —

PLLA-g-ArPr (ArPr = Arachidonic acid propyl ester). The polymer-bound
DCC coupling procedure was similar to PLLA-g—LnPr. This polymer can be oxidized,
and therefore should be carried out and worked up in an inert atmosphere where exposure
to oxygen is minimal. Avg. Yield (3 times): 84%. 1H NMR (300 MHz, CDCI3, 6): 5.35
(m, 8H, -(CH=CH-CH2)3CH=CH), 5.2-5.0 (q, *-CH(CH3), -CH((CH2)3CO2C19H31), 4.06
(m, 2H, -CH2CO2C19H31), 2.79 (m, 6H, -(CH=CH-CH2)3), 2.4-2.2 (m, 2H, -

CO2CH2C13H29), 2.15-1.9 (m, 6H, -CH2CH2CH2CO2C19H31, -CH2(CH=CH-

96

 

CH2)3(CH=CH)CH2), 1.8-1.6 (m, 4H, -CO2CH2CH2C.7H27, -CH2CH2CH2CO2C19H3|),
1.6-1.5 (d, *-CH(CH3)), 1.28 (m, 6H, -(CH2)3CH3), 0.865 (t, 3H, -(CH2)4CH3). ”C
NMR (75 MHz, CDCI3, 6): 173.4, *169.6, 130.46, 128.85, 128.57, 128.22, 128.17,
128.12, 127.83, 127.52, 72.1, *69.0, 63.3, 33.58, 33.54, 33.25, 31.48, 29.29, 27.63, 27.19,

26.55, 26.44, 25.60, 24.76, 24.63, 24.20, 22.54, *16.6, 14.0. * PLLA.

° 1?
HWO/K/ P(OEt)2

PLLA-g-DPAPr (DPAPr = Diethylphosphonoacetic acid propyl ester). In a
small round bottom flask, PLLA-g-HP (0.500 g, 1.31 mmolg", 0.655 mmol),
diethyphosphonoacetic acid (0.14] g, 0.720 mmol) and DMAP (16 mg, 0.131 mmol)
were stirred in 10 mL of CH2C12 until completely dissolved. 1,3-
Dicyclohexylcarbodiimide (DCC) (0.720 mL, 1.0 M, 0.720 mmol) was then added via
syringe and the reaction mixture is stirred under N2 at room temperature. (Note: After
15 minutes, the colorless, homogeneous solution became cloudy, indicative of the
esterification). After 12 h, the reaction mixture was filtered to remove the insoluble
dicyclohexyl urea byproduct, further concentrated, and filtered a second time to firlly
remove the urea. The filtrate was then concentrated to dryness, redissolved in the
minimal amount of CH2CI2, precipitated and centrifuged from methanol two times. Avg.
Yield (3 times): 88%. 1H NMR (300 MHz, CDCI3, 6): 5.2-5.0 (m, *-CH(CH3), -
CH(CH2)3CO2CH2P(=O)(OEt)2), 4.14 (m, 6H, -CH2CO2CH2P(=O)(OCH2CH3)2), 2.94
(d, 2H, -CH2P(=O)(OEt)2, JP-” = 22 Hz), 1.99 (m, 2H, -CH2(CH2)2CO2CH2P(=O)(OEt)2),
1.78 (m, 2H, -CH2CH2CH2CO2CH2P(=—=O)(OEt)2), 1.6-1.4 (d, *-CH(CH3)), 1.4-1.2 (t, 6H,

-P(=O)(OCH2CH3). ”C NMR (75 MHz, CDC13, 8): *169.5, 169.2, 165.7 (Jp-c = 5.8

97

 

Hz), 72.0, 69.0, 64.5, 62.7 (Jp.c = 6.3 Hz), 34.2 (Jp.c = 134.6 Hz), 27.4, 24.0, *16.6, 16.3

(Jp-c = 5.9 Hz). 31P NMR (121 MHz, CDC13, 6): 20.2. * PLLA.

0 9
UWO/K/HOW

PLLA-g-PAPr (Phosphonoacetic acid propyl ester). In a small vial PLLA-g-
DPAPr (0.355 g, 0.665 mmolg", 0.236 mmol) was dissolved in 3 mL of CH2C12.
Bromotrimethylsilane (0.361 g, 2.36 mmol) was added via syringe and stirred at room
temperature for 8 hours. The solution was concentrated and precipitated from aqueous
methanol/water (90/10 v/v) two times. The polymer was then filtered and dried. Avg.
Yield (3 times): 71%. 1H NMR (300 MHz, CDCI3, 6): 5.2-5.0 (m, *-CH(CH3),
CH(CH2)3CO2CH2P03H2), 4.17 (m, 2H, (CH2)2CH2CO2CH2PO3H2), 3.00 (d, 2H,
CH2PO3H2), 1.98 (m, 2H, -CH2(CH2)2CO2CH2PO3H2), 1.78 (m, 2H, -
CH2CH2CH2CO2CH2P(=O)(OEt)2), 1.6-1.4 (d, *-CH(CH3)). l3C NMR (75 MHz, d6-
DMSO, 6): *169.2, 166.9 (Jp.c = 6.0 Hz), 72.0, *68.7, 63.6, 36.5 (Jp-c = 127.8 Hz), 27.1,

23.6, *16.5. 31P NMR (121 MHz, CDC13, 6): 23.3.

0
UWOWNHBOC
NHBoc

PLLA-g-BocLysPr (BocLysPr = N,N'-Di—Boc-lysine propyl ester). In a small
round bottom flask, PLLA-g—HP (0.470 g, 1.31 mmolg", 0.616 mmol), N,N'-di-Boc-
lysine (0.256 g, 0.740 mmol) and DMAP (15 mg, 0.123 mmol) were stirred in 10 mL of
CH2Cl2 until completely dissolved. 1,3Dicyclohexylcarbodiimide (DCC) (0.740 mL, 1.0
M, 0.740 mmol) was then added via syringe and the reaction mixture is stirred under N2
at room temperature. (Note: After 15 minutes, the colorless, homogeneous solution

became cloudy, indicative of the esterification). After 12 h, the reaction mixture was

98

 

filtered to remove the insoluble dicyclohexyl urea byproduct, further concentrated, and
filtered a second time to fully remove the urea. The filtrate was then concentrated to
dryness, redissolved in the minimal amount of CH2CI2, precipitated and centrifuged from
methanol two times. Avg. Yield (3 times): 70%. lH NMR (300 MHz, CDCI3, 6): 5.2-5.0
(m, *-CH(CH3), -CH(CH2)3CO2Lys(Boc)2), 4.66 (m, 1H, -CH2NHBoc), 4.23 (m, 1H, -
(CH2)3CO2CH(NHBoc)(CH2)4NHBoc), 4.12 (m, 2H, -(CH2)2CH2CO2Lys(Boc)2), 3.07
(d, 2H, -CH2NHBoc), 1.97 (m, 2H, -CH2(CH2)2CO2Lys(Boc)2), 1.77 (m, 2H, -
CH2CH2CH2CO2Lys(Boc)2), 1.7-1.5 (d, *-CH(CH3)), 1.41 (s, 18H, -C(CH3)3), 1.7-1.3 (s,
6H, -(CH2)3CH2NHBoc). 13C NMR (75 MHz, CDC13, 6): 172.7, *169.6, 156.1, 155.5,

79.8, 79.0, 72.0, *69.0, 64.2, 53.3, 40.1, 32.3, 29.6, 28.4, 28.3, 27.5, 24.1, 22.5, 22.1,

0 +
H
UEAAOW/N 3 -OzCCF3
+ NH3 '02CCF3

PLLA-g-LysTFAPr (LysTFAPr = lysine bis(trifluoroacetate) propyl ester).

* 16.6. * PLLA.

In a small round bottom flask, PLLA-g-BocLysPr (0.150 g, 1.31 mmolg'l, 0.616 mmol)
was dissolved in 4 mL of CH2CI2. Trifluoroacetic acid (1.0 mL, 12.98 mmol) was added
(20% v/v) and the solution stirred for 25 minutes, at which point the solution became
slightly turbid. The solution was then directly precipitated into an aqueous solution
containing a ten-fold excess of NaHCO3 (Note: Precipitation into the basic solution must
be performed slowly as CO2 is evolved). The polymer was centrifuged from water 3
times to give the pure polymer as the ammonium salt. Avg. Yield (2 times): 95%. 1H
NMR (300 MHz, dg-thf, 6): 5.3-5.0 (m, *-CH(CH3), -CH(CH2)3CO2Lys-NH3(TFA)),
4.20 (m, 2H, -CH2CO2Lys-NH3(TFA)), 3.81 (m, 1 H, -

(CH2)3CO2CH(NH3(TFA))(CH2)4NH3(TFA)), 2.98 (m, 2H, -

99

 

C02CH(NH3(TFA))(CH2)3CH2NH3(TFA)), 1.99 (m, 2H, -CH2(CH2)2CO2Lys-
NH3(TFA)), 1.83 (m, 2H, -CH2CH2CH2CO2Lys-NH3(TFA)), 1.9-1.3 (m, *-CH(CH3),
6H, -(CH2)3CH2NH3(TFA)). ”C NMR (75 MHz, dg-thf, 6): *170.1, 162.3 (.IC.F = 33.2
Hz), 118.0 (JC.F = 292 Hz), *69.7, 53.9, 39.8, 31.9, 28.5, 27.6, 22.7, 20.9, *16.9. ”F

NMR (289 MHz, dg-thf, 6): -76.8. * PLLA.

100

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