MOLECULAR MASS DEPENDENT MECHANICAL PROPERTIES OF METALFREE CLICK HYDROGELS
A Thesis
Presented to
The Graduate Faculty at The University of Akron
In Partial Fulfillment
of the Requirements for the Degree
Master of Science
Huifeng Wang
May, 2015
MOLECULAR MASS DEPENDENT MECHANICAL PROPERTIES OF METALFREE CLICK HYDROGELS
Huifeng Wang
Thesis
Approved:
Accepted:
_____________________________
Advisor
Dr. Matthew Becker
_____________________________
Department Chair
Dr. Coleen Pugh
_____________________________
Committee Member
Dr. Abraham Joy
_____________________________
Dean of the College
Dr. Eric J. Amis
_____________________________
Interim Dean of the Graduate School
Dr. Rex Ramsier
_____________________________
Date
ii
ABSTRACT
A series of PEG-based hydrogels were fabricated via copper-free strain-promoted
alkyne-azidecycloaddition. Characterization of the hydrogels using NMR, IR, UV and
MALDI-TOF mass spectroscopy revealed the chemical structure of the small
molecules and polymers. Rheology was used to determine the mechanical properties
and gelation kinetics of the hydrogels. The gelation time of the hydrogels was
observed by taking the crossover point between the storage modulus (G’) vs. time (t)
curve the loss modulus (G”) vs. t curve. The smooth plateau of both curves confirms
the plateau modulus of the hydrogels. The relationship between G’ and t exhibited
molecular mass dependent behavior of PEG-based hydrogels. The differences in the
gelation point and plateau modulus were attributed to the difference in the molecular
mass of the PEG-based hydrogels as a result of trapped entanglements.
KEYWORDS hydrogels, mechanical properties, gelation kinetics, copper-free strainpromoted alkyne-azidecycloaddition
iii
ACKNOWLEDGEMENTS
I would like to take this opportunity to express my heartfelt gratitude to my advisor,
Dr. Matthew L. Becker, for his continuous guidance, support and encouragement through
the course of my research. He gave me a lot of inspiration, freedom and supports to
investigate the research I am interested in. His enthusiastic and optimistic personalities
also influence my philosophy towards science and life.
I would like to thank the reader of my senior thesis, Dr. Abraham Joy, who
graciously gave his time to evaluate my work and provided helpful suggestions.
I would like to thank all my former and current research group members for their
friendships and help both in research and in my life. Acknowledgements are especially
given to Mr. JukuanZheng and all the group members in Dr. Becker’s research group for
their assistance in proceeding with my research successfully and constructing my life in
Akron colorfully.
Last but not the least, I would like to express my deep appreciation and love to my
parents, my friends. Thanks for their support, understanding and sacrifice. They always
stand by me and encourage me to overcome all the difficulties to achieve my dream.
iv
TABLE OF CONTENTS
Page
LIST OF TABLES ........................................................................................................... viii
LIST OF FIGURES ........................................................................................................... ix
LIST OF SCHEMES........................................................................................................ .xii
CHAPTER
I. INTRODUCTION ........................................................................................................... 1
II. BACKGROUND ............................................................................................................ 5
2.1 “Click” Chemistry..................................................................................................... 5
2.1.1Azide–Alkyne Huisgen Cycloaddition ............................................................... 6
2.1.2Copper-Catalyzed Azide–Alkyne Cycloaddition ............................................... 6
2.1.3 Copper-free Strain-promoted Alkyne-AzideCycloaddition ............................... 8
2.2 Introduction of Hydrogels ......................................................................................... 9
2.3 PEG-based Hydrogels ............................................................................................. 11
2.4 Hydrogels for Cell Encapsulation ........................................................................... 14
2.5 Objectives, Motivations, Innovations and Hypothesis ........................................... 14
III. EXPERIMENTAL SECTION .................................................................................... 20
3.1 Materials and apparatus .......................................................................................... 20
v
3.2 Synthesis of DIBO .................................................................................................. 23
3.2.1 Synthesis of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene (1) ............. 23
3.2.2Synthesis of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6-dibenzocyclocta1,5,7-triene. (2) ......................................................................................................... 24
3.2.3Synthesis of 11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol.
(3) .............................................................................................................................. 24
3.2.4 Synthesis of 5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol. (4) 24
3.3 Synthesis of PNPc activated DIBO......................................................................... 25
3.4 Synthesis of PEG-DIBO. ........................................................................................ 25
3.5 1HNMR of DIBO, PNPc activated DIBO and PEG-DIBO. ................................... 26
3.6 MALDI-TOF MS of PEG-DIBO. ........................................................................... 26
3.7Preparation of PEG-DIBO Hydrogels. .................................................................... 27
3.8Rheology. ................................................................................................................. 29
3.9 Swelling ratio experiments. .................................................................................... 30
IV. RESULTS AND DISCUSSION ................................................................................. 31
4.1 1HNMR of DIBO. ................................................................................................... 31
4.1.1 1HNMR of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene. .................... 32
4.1.2 1HNMR of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6-dibenzocyclocta1,5,7-triene. ............................................................................................................... 33
4.1.3 1HNMR of 11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol.
................................................................................................................................... 34
4.1.4 1HNMR of 5, 6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol. ....... 35
4.2 1HNMR of PNPc activated DIBO........................................................................... 36
4.3 1HNMR of PEG-DIBO. .......................................................................................... 37
4.4 MALDI-TOF MS of PEG-DIBO incorporating various molecular mass. ............. 38
vi
4.5 Rheological studies of PEG-DIBO hydrogels. ....................................................... 45
4.6 Swelling ratio experiments of PEG-DIBO hydrogels. ............................................ 49
V. CONCLUSIONS .......................................................................................................... 51
VI. SUMMARY................................................................................................................ 52
REFERENCES ................................................................................................................. 53
APPENDIX ....................................................................................................................... 62
vii
LIST OF TABLES
Table
Page
3.1 Materials ..................................................................................................................... 21
3.2 Apparatus .................................................................................................................... 22
3.3 Mole quantities and actual masses of PEG-DIBO and tetra-arm azide functionalized
PEG to form different molecular mass PEG-basedhydrogel ............................................ 29
viii
LIST OF FIGURES
Figure
Page
1.1 (a) Synthesis of PEG-DIBO.(b) The chemical structure of bi-DIBO- functionalized
PEG (PEG-DIBO); (c) The chemical structure of tetra-arm azide functionalized PEG
cross-linker; (d) A schematic of hydrogel network formation via SPAAC; (e) Inverted
tube test contains hydrogel formation for the5kPEG-DIBO............................................... 5
2.1 Mechanism of the copper(I)-catalyzed alkyne-azidecycloaddition .............................. 9
2.2 Structure of PEG (polyethylene glycol) ...................................................................... 15
4.1 1HNMR spectrum of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene ................ 33
4.21HNMR spectrum of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2:5,6-dibenzocyclocta1,5,7-triene ........................................................................................................................ 34
4.3 1HNMR spectrum of 11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5ol ....................................................................................................................................... 35
4.41HNMR spectrum of
5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol……………….....….….. ..... 36
4.51HNMR spectrum of Carbonic acid,
5,6-dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-yl ester, 4-nitrophenyl ester... .. 37
4.6 1HNMR Spectrum of Dibenzylcyclooctyne Polyethylene glycol ............................... 38
4.7 MALDI-TOF mass spectrum of 2k PEG-DIBO.(a) full spectrum (b) expansion ...... 39
4.8 MALDI-TOF mass spectrum of 3.5k PEG-DIBO.(a) full spectrum (b) expansion ... 41
4.9 MALDI-TOF mass spectrum of 5k PEG-DIBO.(a) full spectrum (b) expansion ...... 42
4.10 MALDI-TOF mass spectrum of 6k PEG-DIBO.(a) full spectrum (b) expansion. ... 44
4.11 MALDI-TOF mass spectrum of 7.5k PEG-DIBO.(a) full spectrum (b) expansion .....
........................................................................................................................................... 45
x
4.12MALDI-TOF mass spectrum of a series of PEG-DIBOs ranging in molecular mass
from 2k to 7.5k.................................................................................................................. 46
4.13 (a)Gelation processfor PEG-DIBO hydrogels with 3.5k molecular mass.(b) Storage
modulusfor aqueous PEG-DIBO hydrogels with different molecular masses ................. 50
4.14 Plateau modulus and gelation kineticsof hydrogels prepared using various molecular
mass................................................................................................................................... 51
4.15 Swelling ratio of PEG-DIBO hydrogels ................................................................... 52
A1Gelation processfor PEG-DIBO hydrogels with 2k molecular mass ........................... 65
A2 Gelation processfor PEG-DIBO hydrogels with 5k molecular mass .......................... 66
A3 Gelation processfor PEG-DIBO hydrogels with 6k molecular mass .......................... 67
A4 Gelation processfor PEG-DIBO hydrogels with 7.5k molecular mass ...................... .68
x
LIST OF SCHEMES
Scheme
Page
3.1 Synthesis of DIBO ...................................................................................................... 23
xi
CHAPTER I
INTRODUCTION
Hydrogels are a unique category of polymeric networks that can absorb and retain a
significant amount of water.1 Due to its high water content and mechanical stability,
hydrogels have been widely used as a suitable polymeric material for biomedical and
pharmaceutical applications, including contact lenses, tissue engineering and cell
regeneration.2-5 The cross-linking methods for in situ formed hydrogels are divided into
physical cross-linking and chemical cross-linking.6-9Physical cross-link between
polymers can be obtained using various non-covalent interactions, such as van der Waals
forces, hydrogen bonding, ionic and hydrophobic interactions.4,11 However, the assembly
of polymer chains through physical cross-linking are weak and reversible interactions,
which the hydrogels are inclined to revert to their sol phase by changes of pH,
temperature and ionic concentration.12-14 Chemical cross-links will yield covalent bonds
among polymer chains and the resulting cross-linked network is more resistant to
mechanical
forces
than
physically
cross-linked
1
network.11,14
Various
types
of covalent cross-linking strategies have emerged to form hydrogels, including Michaeladdition, thiol-ene radical addition, copper (Ⅰ)-catalyzed alkyne-azidecycloadditon
(CuAAC) or metal-free strain-promoted alkyne-azidecycloaddition (SPAAC), etc.12,14-16
While the presence of metal catalysts, organic solvents and residual functional groups
limit its use in living systems.13
There have been many reports using alginates and physically cross-linked gels in neural
and stem cell engi-neering.6,17-19,21-25 However, neural cells are extremely sensitive to
photochemical and catalytic activation methods frequently used to induce cross-linking
and the resulting modulus ranges have been limited in their range and versatility.7,20
Poly(ethylene glycol) (PEG)-based hydrogels are one of the most widely studied
polymeric materials in biomedical applications, owing to their cytocompatibility, low
toxicity and ease of end group modification for cross-linking or network formation.14,26-30
In addition, PEG-based hydrogels display tunable mechanical properties in the
appropriate range for soft tissue regeneration.12,14-16,30-32 PEG-based hydrogels can be
formed by cross-linking or entanglement of polymer chains, through either physical or
chemical bonding.33,34 Cycloadditionsare considered to be ideal reactions due to
theirregiospecificity and chemoselectivity.35Huisgen developed the [3+2] cycloaddition
between an azide and an acyclic alkyne, but a good condition, such as metal catalyst and
high temperature, is required to overcome the activation barrier to form
triazole.36Sharpless and co-workers developed a Cu-catalyzed alkyne-azidecycloaddition,
but the presence of metal catalyst may lead to cytotoxicityissues.37 Boon’s group
accelerate the rate of reaction by increasing strain energy through a functionalized
2
derivative of dibenzocyclooctyne (DIBO), which is metal-free strain-promoted alkyneazidecycloaddition (SPAAC).38 In recent years, hydrogels produced by SPAAC crosslinking of multi-functional polymers and multifunctional small molecules, or cross-linked
multi-functional polymer-polymer system have grown in popularity.39-43The reaction
proceeds in aqueous solutions without side reactions, and the triazole is stable to
oxidation and acid hydrolysis.41 Previously, this reaction was widely used for surface
immobilization,26 oligonucleotide functionalization,27dendrimer modification29 and the
decoration of nanostructures.28 The gel formation via SPAAC is particularly attracting
due to its molecular mass dependent behavior.44 Currently, significant efforts have
focused on replicating the mechanical properties of tissues with synthetic hydrogels.45-47
These unique features inspire us to create hydrogels with tunable mechanical properties
and gelation kinetics by modulating the molecular mass.
In this study, we fabricate PEG-based hydrogels with different molecular mass using
copper-free strain-promoted alkyne-azidecycloaddition (SPAAC). The synthesis strategy
is outlined in Figure 1.1 We have designed and synthesized two precursors: bidibenzocyclooctyne(DIBO) functionalized polyethylene glycol (PEG-DIBO) (Figure
1.1(a)) and tetra-arm azide cross-linker (Figure 1.1(b)) molecule. With identical
precursor chemistries, we found that the mechanical properties and gelation kinetics were
influenced greatly by molecular mass of PEG-based hydrogels.We discuss the details of
the synthesis and the mechanical properties of the networks with emphasis on the
relationship between the plateau modulus and molecular mass, which is the condition
expected in cell regeneration.
3
Figure 1.1 (a) Synthesis of PEG-DIBO was achieved by reaction between DIBO and acid
chloride and subsequent reaction with poly(ethylene glycol) diamine. (b) The chemical
structure of bi-DIBO functionalized PEG (PEG-DIBO); (c) The chemical structure of
tetra-arm azide functionalized PEG cross-linker; (d) A schematic of hydrogel network
formation via SPAAC; (e) Inverted tube test contains hydrogel formation for the 5k PEGDIBO.
4
CHAPTER II
BACKGROUND
2.1 “Click” Chemistry
“ Click Chemistry” is a term that was introduced by K.B.Sharpless in 2001 to describe
reactions that are high yielding, wide in scope, create only byproducts that can be
removed without chromatography, are stereospecific, simple to perform, and can be
conducted in easily removable or benign solvents. 45, 48 This concept was developed in
parallel with the interest within the pharmaceutical, materials, and other industries in
capabilities for generating large libraries of compounds for screening in discovery
research.37,49-51 The most used “click” reaction that can fulfill these conditions is by far
the Cu(I)-catalyzed azide/alkyne cycloaddition (CuAAC).28,52,53 Other “click” reactions
are the thiol-ene, oxime, Diels-Alder, Michael addition and pyridyl sulfide reactions.54-60
The copper catalyzed azide/alkyne cycloaddition (CuAAC) is thermodynamically and
kinetically favorable,which its energy is 50 and 26kcal/mol, respectively.52,53 In addition,
the regiospecific and chemoselective properties make the CuAAC more widely used in
the chemical synthesis.37,46 There is a 107 rate enhancement over the non-catalyzed
5
reaction, which proves the role of the metal catalysts is significant.37 Moreover, the
triazoles are stable to oxidation and acid hydrolysis.41
2.1.1Azide–Alkyne HuisgenCycloaddition
Azides react as 1,3-dipoles when converted with alkynes in a concerted [3+2]
cycloaddition, leadingto aromatic triazole products as proposed by Rolf Huisgen in the
1950s, when he introduced hisconcept of 1,3-dipolar cycloadditions.54 The particular
attractivity ofthe alkyne and azide functionalities as complementary coupling partners is
based on the ease of theirsynthesis, their kinetic stability, and their tolerance to a wide
variety of functional groups andreaction conditions. Although this reaction has been
served to form hydrogels by cross-linkage ofazide- and alkyne-functionalized macromonomers,55 the high temperatures or pressures that arerequired to promote this reaction
are not compatible with living systems. To address this problem,Kiser and Clark
developed a Huisgencycloaddition for hydrogel formation that works at
physiologicaltemperature by using electron-deficient alkynes;56 the reaction rates,
however, are relatively slow.
2.1.2Copper-Catalyzed Azide–Alkyne Cycloaddition
Independent of each other, the groups of Sharpless57 and Meldal58 discovered a dramatic
rateacceleration of the azide–alkyne coupling along with the selective formation of 1,4disubstituted1,2,3-triazoles under copper(I) catalysis. This is because the alkyne is
activated by the formation of acopper acetylide toward the reaction with azides; as a
result, the reaction even occurs at roomtemperature. The mechanism of this reaction,
which is termed copper-catalyzed azide–alkynecycloaddition (CuAAC), has remained
6
difficult to establish and is still subject to ongoingresearch.59-61The proposed mechanism
for the CuAAC is shownin Figure 2.1.
The CuAAC has become the most widely used click reaction in organic synthesis,62
polymerchemistry,63 materials chemistry,63 chemical biology,64 and in medicinal65 and
pharmaceuticalchemistries.66 Its potential for hydrogel preparation could be
demonstrated, for example, by Hawkeret al.,67 who synthesized well-defined PEG
hydrogels by cross-linking alkyne-terminated tetra-armPEG and diazide-terminated PEG
by CuAAC. Yang and co-workers synthesized PEG–peptide hydrogelsby cross-linking
diazide-terminated peptides and alkyne-terminated tetra-arm
PEGs.68Theobtainedhydrogels exhibit improved mechanical properties in comparison to
photo-chemicallycross-linked PEGgels.
Figure 2.1 The mechanism of the copper(I)-catalyzed alkyne-azide cycloaddition.
The CuAAC methods have been used in many applications, such as organic synthesis,
dendrimer assembly, and surface functionalization.69-71For example, the DNA
microarrays are really useful for large scale parallel analysis of gene expression. While
the chemistry used for immobilization is limited by cross-reactivity on surface, the
7
efficiency and bioorthogonality of CuAAC could overcome existing limitations of
immobilization.72-75 Besides, The CuAAC reaction has been widely used in organic
syntheses including several types of macromolecules.66In immobilization reactions,
amino resins are converted to azido resins by diazo transfer followed by a CuAAC
reaction. Alkyne-containing proteins have been immobilized by triazole formation.7679
The peptide functions have been modified with triazoles. Triazoles are excellent
mimetics of the peptide bonds which can provide efficient inhibitors of key mammalian,
bacterial and viral proteases.79,80Natural products and drugs have been modified with
triazoles in order to enhance the solubility or bioavailability, to attach a biolabel or a
fluorophore, or to diversify the drug structure. 81-85
2.1.3 Copper-free Strain-promoted Alkyne-AzideCycloaddition
In 2004, Bertozzi et al.initially employed this reaction toward in-vitro labeling of
biomolecules at physiologicalconditions.86,87Recently, the study of live cells and
organisms has been the primary application of cyclooctyne reagents; however, the simple
reaction conditions and purification procedures of copper-free click chemistry have
resulted in its expansion to a variety of disciplines including surface immobilization,gel
synthesis,oligonucleotide functionalization, dendrimer modification, and the decoration
of nanostructures.88-93 This reaction was selective and mild, but its relatively slow rate
remained a liability for many applications. So, enhancing the rate of the azidecyclooctynecycloaddition by applying classical physical organic chemistry principles to
further activate the strained alkynes for reaction with azides is really important.38
Through lots of experimental efforts, two rate-enhancement strategies have emerged: the
8
addition of electron-withdrawing groups and the augmentation of strain energy.94
Therefore, the copper-free strain-promoted alkyne-azidecycloaddition click reaction has
emerged to utilize the click chemistry in tissue engineering and pharmaceutical
applications.
2.2 Introduction of Hydrogels
Hydrogels are cross-linked networks of hydrophilic polymers capable of retaining large
amounts of water yet remaining insoluble and maintaining their three-dimensional
structure.95,96Since their discovery and application in the biomedical field by Wichterle et
al. in the early 1950s, an immense number of hydrogels have been developed, and they
have been studied for a wide range of biomedical and pharmaceutical applications,
including contact lenses, tissue engineering, diagnostics, drug delivery, vascular
prostheses, and coating for stents and catheter.97-101 The hydrophilic polymers used to
create hydrogels need to be physically and/or chemically cross-linked to prevent their
dissolution.102 Hydrogels can be prepared from natural and synthetic polymers and they
can consist of homo-polymers, copolymers, and interpenetrating or double polymeric
networks.103Hydrogels can be made biodegradable by a proper selection of their building
blocks as well as the applied cross-linking strategy. 104
Hydrogels are generally regarded as non-cytotoxic materials because their high water
content and soft nature render them similar to natural extracellular matrices and minimize
tissue irritation and cell adherence.105 Furthermore, their porous structure, along with
their water content, are extremely suitable properties to accommodate high loads of
9
water-soluble compounds, like therapeutically active proteins and peptides.106 Unlike
other delivery systems (microparticles, emulsions, etc.), where preparation conditions are
sometimes detrimental to proteins (i.e., use of organic solvents and protein denaturating
processes, like homogenization, exposure to interfaces, and shear forces, etc.), hydrogel
preparation procedures are beneficial in preserving protein stability, as very mild
conditions aqueous environment, room temperature) are normally adopted.107Finally,
proteins have a limited mobility or are immobilized in the hydrogel network, which is
favorable for preservation of their mostly fragile 3D structure.108 All these unique
properties of hydrogels have raised increasing interest in their use as reservoir systems
for proteins that are slowly released from the hydrogel matrix in a controlled fashion to
maintain a therapeutic effective concentration of the protein drug in the surrounding
tissues or in the circulation over an extended period of time.109 Proteins can be physically
incorporated in the hydrogel matrix, and their release is governed by several mechanisms,
such as diffusion, swelling, erosion/degradation, or a combination of these
mechanisms.110 Hydrogels allow fine-tuning of the protein release by tailoring their
cross-link density via changes in polymer architecture, concentration, molecular weight,
or chemistry.111Other strategies to tailor drug release from hydrogels exist and they rely
on reversible protein−polymer interaction or encapsulation of the protein in a second
delivery system (e.g., micro- or nanoparticles) dispersed in the hydrogel network.112
10
2.3 PEG-based Hydrogels
PEGs are the most frequently used synthetic macro-monomers in the biological area,
becausePEGs and their resulting hydrogels have been believed to be non-immunogenic,
non-toxic, and inertto protein and cell interactions113 As a result, PEG, PEG-hydrogels,
and PEG-containing formulationshave been approved by the U.S. Food and Drug
Administration (FDA) and thereafter widely beenused for the encapsulation of proteins
and cells.114Poly(ethylene glycol) (PEG) is also a commercially available polyether
backbone polymer that is soluble ina variety of different solvents including water,
methanol, ethanol, toluene, and dichloromethane. Itcan be prepared in a well controllable
way by anionic polymerization of ethylene oxide entailing awide range of accessible
molecular weights from 300 g/mol to 10,000,000 g/mol along withnarrow molecular
weight distributions. By appropriate choice of the initiator, mono-functional,
homoandheterobi-functional linear PEG, as well as PEG dendrimers and multi-arm PEGs
can be prepared.97Post-modification of hydroxy-terminated PEG provides access to a
large variety of functionalities, suchas acrylates,115 methacrylates,116 vinyl sulfones,117
thiols,118 maleimides,119 alkynes,120 azides,120 amines,121carboxylates, and active esters.122
Thesefunctional groups can serve to form three-dimensionalcross-linked polymer
networks by using linear PEG-acrylates or -methacrylates and applying
freeradicalchemistry123,124 or by mixing functional multi-armPEGs with complementary
functional linearPEGs.115 The resulting polymer networks often display polydisperse
mesh sizes, along with structuralimperfections such as loops and dangling chains
entailing low mechanical strength of the gels.125-128
11
Hydrogels are a class of biomaterial scaffolds that have been widely used in complex
device fabrication, drug release, and tissue engineering.129,130PEG-based hydrogels, in
particular, have proven extremely versatile for tissue engineering applications.131,132 In
addition, PEG-based hydrogels display tunable mechanical properties in the range
appropriate to soft tissue regeneration. 45-47
Figure 2.2 The structure of PEG(polyethylene glycol).
PEG hydrogels have also been used to modify biomaterial surfaces to provide protein
resistance and to enhance surface biocompatibility, due to low levels of nonspecific
binding to a range of biological molecules such as proteins and polypeptides.133,134
PEG hydrogels can be formed by cross-linking of polymer chains, through either
physical or chemical means.33,34 PEG hydrogels prepared by physical cross-linking are
usually free of impurities arising from initiator fragments, and therefore are favorable for
biomedical applications.135,136 However, the polymer chains in physically cross-linked
PEG hydrogels are connected through weak, reversible interactions such as van der
Waals forces, hydrogen bonding, ionic, or hydrophobic interactions.4,11,137 Therefore, the
gels can often revert to their sol phase by application of relatively small mechanical
forces, or by changes in temperature and solvent conditions.12-14,138 Chemically cross12
linked networks prepared by a free radical process, for example, by UV irradiation of
PEG precursors and copolymers, also produce gels with low levels of
impurities.11,14,139However, there is generally a high occurrence of defects within the
hydrogel network such as loops and entanglements.140,141 The consequence of the
conventional radical cross-linking mechanism where cross-linking monomers can either
react to form cross-links (the ideal case), loops, or remain unreacted (both non-ideal).142144
For this reason, it is difficult to ascertain with great accuracy the relationship between
the concentration of cross-linking agent and properties of the resulting hydrogels, such as
mechanical properties, rates of solute transport, and network porosity.145,146
PEG hydrogels can also be prepared by reaction of functionalized polymeric precursors
with cross-linkers that possess complementary reactivity.147 Such complementary
systems include, radical-based thiol-ene, Michael addition, 1,3-dipolar cycloaddition of
azides with ring strained alkynes, 1,3-dipolar cycloaddition of azides with electron
deficient alkynes and copper-catalyzed Huisgen’s 1,3-dipolar cycloaddition of azides
with alkynes.54-60,148The use of complementary functional groups for cross-linking
eliminates the occurrence of loop-based defects, and the cross-link density can be
controlled by changing the molecular weight of the polymeric precursors.149 The “click”
chemistry reactions described above have been shown to be good candidates for the
cross-linking reactions to prepare hydrogels, because of their high efficiency and
selectivity. 150
The mechanical properties of the PEG-based hydrogel are
molecularweightdependent.44,151Therefore, we can modulate the mechanical properties of
13
the hydrogels by changing the molecular weight of the PEG, which can be used in
distinct fields such as drug delivery, tissue scaffold and neuron cultivation.45-47,152
Generally speaking, the storage modulus will decrease along the increase of the
molecular weight of the PEG. The neurons are sensitive to the soft hydrogels, which the
high-molecular-weight PEG-based hydrogels can be used in nerve applications.
2.4 Hydrogels for Cell Encapsulation
Several in-vivo studies in animals153,154 and even in humans155 have confirmed the
potential ofmicroencapsulated cells for therapeutic treatment of diabetes, but have
coincidently shown thenecessity for the development of new polymeric matrixes. For this
purpose, highly biocompatible, bio-inert,and tunable synthetic polymeric precursors such
as polyglycerol and PEG should be used along withreversible and responsive chemical or
physical cross-links.
2.5 Objectives, Motivations, Innovations and Hypothesis
The thesis outlines our efforts to fabricate the PEG-based hydrogels via copper-free
strain-promoted alkyne-azide cycloaddition “click” chemistry that are soft and
dynamically stable, which can be used in neuron cultivation. We aim to change the
molecular weight of the PEG and this tunable molecular weight allows for the increase of
the mechanical properties. These novel polymeric materials will allow development of
hydrogels with continuous stability to the neurons and improvement of mechanical
properties to control a balance of hydrogel formation and neuron cultivation.
14
Our hypothesis is that the change of the molecular weight of the PEG in the PEG-based
hydrogels will decrease the mechanical properties and gelation kinetics in tissue
engineering.
The current surgical options to address defects in long nerves resulting from trauma,
defect, resection, or disease have significant limitations. Versatile, resorbablepolymeric
materials for hydrogel formation with sufficient mechanical properties to minimize
supplementary external fixation could be used for numerous clinical applications where
current off the shelf solutions are not adequate. So these novel biodegradable and
biocompatible PEG-based hydrogels by changing the molecular weight of the PEG offer
a cost effective solution to the current recombinant technologies. The reinforcement of
mechanical properties and aid in the healing process giving them more potential
applications to be an effective treatment for neuron cultivation.
In this thesis, the primary innovation is the development of PEG-based hydrogels for the
neuron cultivation utilizing different molecular weight of PEG to increase the mechanical
properties, biodegradation rate and biocompatibility. The advantages include:
1). Polymeric materials with different molecular weight of PEG can increase the
stiffness and strength compared with the polymeric hydrogels currently widely used in
nerve connectivity.
2). The tunable molecular weight of PEG will also increase the flexibility and
biocompatibility which provide us a cost effective way to apply these polymeric materials
into biological application.
15
CHAPTER III
EXPERIMENTAL SECTION
3.1 Materials and apparatus
The materials and instruments used in the research are shown below.The materials were
purchased from Sigma-Aldrich (St. Louis, MO), Fisher (Pittsburgh, PA) and Jekem
(Beijing, China) and used as received.
20
Table 3.1Materials
Name
Formula
Purity
Source
Phenylacetaldehyde
C8H8O3
98%
Sigma-Aldrich
Trimethylsilyl iodide
C3H9SiI
98%
Sigma-Aldrich
n-butyl lithium
C4H9Li
99%
Sigma-Aldrich
bromine
Br2
99%
Sigma-Aldrich
Lithium
C6H14LiN
99.5%
Fisher
96%
Sigma-Aldrich
diisopropylamide
4-Nitrophenyl
ClCO2C6H4NO2
chloroformate
pyridine
C3O3Cl6
98%
Fisher
Magnesium Sulfate
MgSO4
99%
Sigma-Aldrich
Sodium Sulfate
Na2SO4
98%
Chemstore
21
Chloroform, Anhydrous
CHCl3
99%
Fisher
Tetrahydrofuran
C4H8O
99%
Fisher
C2nH4n+6On+1N2
96%
Jenkem
C2nH4n+6On+1N2
96%
Jenkem
C2nH4n+6On+1N2
96%
Jenkem
C2nH4n+6On+1N2
96%
Jenkem
Anhydrous
Polyethylene glycol
bisamine Mw2000
Polyethylene glycol
bisamine Mw3500
Polyethylene glycol
bisamine Mw5000
Polyethylene glycol
bisamine Mw7500
Table 3.2The apparatus used in the work.
Apparatus
1
Type
H-NMR
Varian Mercury 300 Spectra
Rheology
TA Instruments ARES-G2 rheometer
MALDI-TOF MS
BruckerUltraFlexⅢ tandem time-of-flight
(TOF/TOF) mass spectrometer
22
3.2 Synthesis of DIBO
The synthesis of DIBO was achieved by the following four steps, which the synthesis
strategy was shown in Scheme 3.1.
O
H
iodotrimethylsilane
chloroform
O
n-BuLi
THF
HO
Br2
chloroform
Br
Br
LDA
THF
HO
HO
Scheme 3.1 Synthesis of DIBO.
3.2.1 Synthesis of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene (1)
A 250 mL flask was flame dried and charged with argon. Phenylacetaldehyde (18.52 g,
0.154 mol) and 100 mL of chloroform (anhydrous) were then added via syringe. The
reaction flask was cooled in an ice bath. Trimethylsilyl iodide (25 mL, 37.5 g, 0.188mol)
was added to the solution and the reaction was allowed to stand at 5 °C for 7 days. The
reaction was monitored by TLC. After 7 days, sodium thiosulfate (1.0 M, 160 mL) and
chloroform (200 mL) were added, and the mixture was stirred until the iodine color was
discharged. The organic phase was separated, dried (sodium sulfate), and concentratedin
vacuum. Chromatography on silica gel eluting with chloroform yielded 6.1 g of the
crystalline ether compound (35%).
23
3.2.2Synthesis of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6-dibenzocyclocta-1,5,7triene. (2)
2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene 1 (2.00 g, 5.84 mmol) in anhydrous
THF (60 mL) was placed into a three-necked round bottom flask and cooled in an ice
bath under argon. n-butyl lithium (4.92 mL, 2.5 M, 12.4 mmol) was added slowly via
syringe. The reaction mixture was stirred at room temperature under argon for 4 h. The
reaction was quenched by careful addition of water and extracted with 2 x 50 mL CHCl3.
The combined organic phases were washed with 30 mL of brine, dried over Na2SO4,
concentrated under vacuum and purified by column chromatography on silica gel CHC13
to yield 1.83 g of 3-hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6-dibenzocyclocta- 1,5,7triene (90%).
3.2.3Synthesis of 11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol. (3)
Bromine (0.51 mL, 10 mmol) was added dropwise to a stirred solution of 2 (2.22 g, 10
mmol) in CHCl3 (50 mL). After stirring the mixture for 0.5 h, TLC analysis indicated
completion of the reaction. The solvent was evaporated under reduced pressure and the
residue was purified by flash chromatography over silica gel (2:1/1:2, v/v,
hexanes/CH2Cl2) to yield 3 as lightyellow oil (60%).
3.2.4 Synthesis of 5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol. (4)
Lithium diisopropylamide in tetrahydrofuran (2.0 M; 8.0 mL, 16 mmol) was added
dropwise to a stirred solution of 3 (1.53 g, 4.0 mmol) in tetrahydrofuran (40 mL) under
an atmosphere of argon. The reaction mixture was stirred for 0.5 h, after which it was
quenched by the dropwise addition of water (0.5 mL). The solvents were removed under
24
reduced pressure, and the residue was purified by flash chromatography on silica gel
(hexanes/CH2Cl2 2:1/0:1, v/v) to yield 4 as a white amorphous solid (0.52 g, 60%).
3.3 Synthesis of PNPc activated DIBO.
4-Nitrophenyl chloroformate (0.182 g, 2 mmol) and pyridine (0.182 mL, 5 mmol) were
added to a solution of DIBO (0.10 g, 1 mmol) in CH2Cl2 (14 mL). After being stirred for
4 hours at room temperature, the mixture was washed with deionized water (2×10 mL)
and the organic layer was dried (MgSO4). The solvents were evaporated under reduced
pressure, and the residue was purified by silica gel column chromatography (hexane/ethyl
acetate, 10:1, v/v) to afford 4-nitrophenyl DIBO (0.22g, 80%).
3.4 Synthesis of PEG-DIBO.
PEG-DIBOs were prepared from the reaction of various PEG bisamine and nitrophenyl
activated Dibenzylcyclooctyne (DIBO), respectively. An example synthesis of a
3.5×103g/molPEGbisamine is noted below. PEG bisamine (100mg, 0.029mmol),
nitrophenyl activated DIBO (42mg, 0.112mmol), and triethylamine(19.50μL,
0.116mmol) were reacted in 34mL of dichloromethane over 100mL flask for 24h at room
temperature. The solvent was removed under vacuum and water was added to dissolve
the product. The product was dialyzed in water with a pore sizecut off of 1000g/mol
molecular mass for 2 days. The product was then lyophilized for 2 days at -52℃.
25
3.5 1HNMR of DIBO, PNPc activated DIBO and PEG-DIBO.
High-resolution, 300 MHz proton NMR spectra were acquired on a Varian Mercury 300
MHz spectrometer. Deuterated chloroform was used as a solvent, and the polymer
concentrations were varied between 2.5% and 3.0% by mass. All samples were run at
room temperature, 15Hz sample spinning, 90°angle, for 128 scans.
3.6 MALDI-TOF MS of PEG-DIBO.
The matrix-assisted desorption ionization(MALDI) matrix,
2-[(2E)-3-(4-tert-butylphenyl)-2-methylprop-2-enylidene] malonitrile (DCTB)
(20mg/mL), PEG-DIBO (20.0 mg/mL) and cationizing agent, sodium trifluoroacetate
(NaTFA) were prepared in aqueous solution and mixed in a 10:5:1 ratio (mass),
respectively. MALDI samples were deposited on the target using electrospray. The
MALDI time-of-flight mass spectroscopy (MALDI-TOF) MS was carried out on
BruckerUltraFlexⅢ
tandem
time-of-flight
(TOF/TOF)
mass
spectrometer
(BruckerDaltonics, Billerica, MA), equipped with a Nd:YAG laser emitting at a
wavelength of 355nm. MS experiments were performed by using reflectron mode and
linear mode for samples with different molecular mass without additional collision gas.
The data analysis was conducted with flexAnalysissoftware.
26
3.7 Preparation of PEG-DIBO Hydrogels.
Chemically cross-linked hydrogels were prepared via metal-free stain-promoted
alkyne-azidecycloaddition (SPAAC). PEG-DIBO (20% by mass fraction) was dissolved
in distilled deionized water (50.0μL). Based on a 1:1 molar ratio DIBO group:azide group
(Table 3.3), the tetra-arm azide functionalized PEG in same amount of deionized water
was mixed with the PEG-DIBO solution in 20.0 mL vial. After shaking for 5min, the
hydrogel was completely formed. The inverted tube test for a 5.0×103 g/mol hydrogel is
shown in Figure 1.1(e).Actual mass of PEG-DIBO is 20% by fraction. Based on 1:1
molar ratio DIBO group:azide group, the actual mass of tetra-arm azide functionalized
PEG was calculated.
27
Table 3.3 Mole quantities and actual masses of PEG-DIBO and tetra-arm azide
functionalized PEG to form different molecular mass PEG-based hydrogels.
Mole quantity
Mole quantity
Actual mass of Actual mass of
of PEG-DIBO
of tetra-arm
PEG-DIBO
tetra-arm azide
(μmol)
azide
(mg)
functionalized
functionalized
PEG (mg)
PEG (μmol)
2k
6.25
3.12
12.5
6.25
3.5k
3.57
1.78
12.5
3.57
5k
2.5
1.25
12.5
2.5
6k
2.08
1.04
12.5
2.08
7.5k
1.67
0.83
12.5
1.67
28
3.8Rheology.
In situ rheological studies were utilized for assessing the gelation kinetics and storage
modulus. Measurements were performed atambienttemperature with a frequency of 1
rad/s and a strain of 10%. The aqueous solution of 100μL mixture of PEG-DIBO and
tetra-arm azide functionalized PEG was loaded between the 8.0 mm parallel plate
geometry on a TA Instruments ARES-G2 rheometer. The measurement time for different
sample was based on the speed of gel formation. The storage and loss modulus were
monitored during the gelation process. Multiple experiments demonstrated excellent
reproducibity with relative standard uncertainty of 1% for each sample incorporating
various molecular mass (n= 3)
29
3.9 Swelling ratio experiments.
For swelling studies, PEG-DIBO hydrogels were made 10mm in diameter and 2mm in
height using a precursor mixing method. Solutions of PEG-DIBO were prepared by
dissolving 12.5mg in 50μL deionized water. Solutions of tetra-arm azide functionalized
PEG were prepared by dissolving the cross-linker precursor in 50μL deionized water
based on a 1:1 molar ratio DIBO group:azide group (Table 3.3). After 24 h, the samples
were weighed then placed in a freeze-dryer and lyophilized prior to being weighed again.
The swelling ratio, Q, was calculated with the following equation:
Q= (MS－ MD)/MD
where MS is the mass after swelling, MD is the mass after lyophilizing.
30
CHAPTER IV
RESULTS AND DISCUSSION
4.1 1HNMR of DIBO.
1
HNMR spectrums of DIBO are shown below, which exhibit the chemical structure of
each step.
31
7.27
4.1.1 1HNMR of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene.
hw38cyclorate7202014second
olvent
a
b
c,d
a
O
c,d
b
a
b
7.0
6.5
6.0
c
3.61
3.58
3.55
3.53
5.32
5.30
7.15
7.08
7.07
7.07
7.00
6.98
7.5
nt
2.81
2.76
7.12
7.10
c,
5.5
5.0
4.5
Chemical Shift (ppm)
4.0
3.5
3.0
2.5
2.0
9
Figure 4.11HNMR spectrum of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene.
1H NMR of 2,3:6,7-Dibenzo-9-oxabicyclo[3.3.l]nona-2,6-diene:
1H NMR (300 MHz, CDC13) δ= 7.09 (m, 8H), 5.30(d, 2H, J=5.9 Hz, CH), 3.55(dd, 2H,
J= 6.3, 16.2 Hz, CH2), 2.75(d, 2H, J=16.4 Hz, CH2).
32
4.1.2 1HNMR of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6-dibenzocyclocta- 1,5,7-
7.27
triene.
solvent
hw38cyclodiene2142014
b
a
c
d
HO
e
7.5
7.0
d
6.5
6.0
e
5.5
5.0
4.5
Chemical Shift (ppm)
4.0
3.5
3.0
1.87
1.85
3.50
3.48
3.45
3.43
3.38
3.34
3.33
3.30
c
5.31
5.29
5.28
5.26
7.28
7.48
7.45
a
7.18
7.10 7.12
7.12
6.91
6.87
6.81
6.85
b
2.5
2.0
Figure 4.21HNMR spectrum of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2 :5,6dibenzocyclocta- 1,5,7-triene.
1
H NMR of 3-Hydroxy-2’,3’,2’’,3’’-tetramethox1y,-2:5,6-dibenzocyclocta- 1,5,7-triene:
1
H NMR (300 MHz, CDCl3): δ=7.48 (m, 1 H), 7.10–7.30 (m, 7H), 6.86 (q, 2H, J=2.7,
12.0 Hz, CH), 5.31 (q, 1H, J=6.1, 10.0 Hz, CHOH), 3.45 (m, 2H, CH2).
33
7.19
4.1.3 1HNMR of 11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol.
hw38bromocyclorate10282014
solven
Br
Br
b
d
a
a
c
e
7.5
7.0
6.5
6.0
c
d
5.39
5.26
5.24
5.23
5.23
5.21
5.5
e
e
3.70
3.65
3.62
3.60
3.56
3.54
3.50
3.48
3.06
3.05
3.00
2.99
2.81
2.75
b
5.81
5.79
5.75
5.69
7.62
7.60
7.54
7.53
7.33
7.31
7.13
a
6.98
6.95
6.93
6.89
6.81
7.01
HO
4.0
4.5
5.0
Chemical Shift (ppm)
3.5
3.0
2.5
2.0
Figure 4.31HNMR spectrum of 11,12-Dibromo-5,6,11,12-tetrahydrodibenzo[a,e]cycloocten-5-ol.
1H NMR of 11, 12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol:
1H NMR (500 MHz, CDCl3): δ=7.70–7.68 (2 H, aromatics), 7.39–6.88 (6 H, aromatics),
5.88 (d, 1H, J=5.4 Hz, CHBr), 5.47 (dd, 1H, J=3.6, 15.9 Hz, CHOH), 5.30 (d, 1H, J=5.4
Hz, CHBr), 3.60 (dd, 1H, J=3.7, 16.1 Hz, CH2), 2.87 (dd, 1H, J=3.7, 16.1 Hz, CH2).
34
7.27
4.1.41HNMR of 5, 6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol.
hw38cyclotriene1142014
solven
a
a
c
b
HO
d
7.5
7.0
6.5
6.0
b
c
d
5.5
5.0
4.5
Chemical Shift (ppm)
4.0
3.5
3.0
2.16
2.14
3.15
3.14
3.10
3.09
2.97
2.96
2.92
2.91
4.65
7.77
7.75
7.46
7.44
7.35
7.37 7.34
a
2.5
2.0
Figure 4.41HNMR spectrum of 5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5ol.
1
H NMR of 5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol:
1
H NMR (500 MHz, CDCl3): δ=7.77 (1 H, aromatics), 7.44–7.23 (7 H, aromatics), 4.66
(dd, J=2.1, 14.7Hz, 1H, CHOH), 3.10 (dd, J=2.1, 14.8 Hz, 1H, CH2), 2.94 (dd, J=2.1,
14.8 Hz, 1H, CH2).
35
hw38nitrocyclo1162014
7.27
4.2 1HNMR of PNPc activated DIBO.
solven
a
b
c
O
O
O
d
e
8.0
7.5
d
7.0
6.5
6.0
c
b
3.38
3.33
3.09
3.08
3.03
7.65
7.62
8.11
8.31
8.28
e
a
5.59
7.44
7.40
7.35
NO2
4.5
5.0
5.5
Chemical Shift (ppm)
4.0
3.5
3.0
2.5
2.0
Figure 4.51HNMR spectrum of Carbonic acid, 5,6-dihydro-11,12-didehydrodibenzo[a,e]cycloocten-5-yl ester, 4-nitrophenyl ester.
1
HNMR of Carbonic acid, 5,6-dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-yl
ester, 4-nitrophenyl ester:
1
HNMR (500 MHz, CDCl3): δ=8.28–8.14 (2 H, aromatics), 7.64–7.63 (2 H, aromatics),
7.45–7.23 (8 H, aromatics), 5.61 (dd, J=3.9, 15.3 Hz, 1H, CHOH), 3.37 (dd, 1H, J=3.9,
15.3 Hz, CH2), 2.05 (dd, 1H, J=3.9, 15.3 Hz, CH2).
36
b
a
a
3.67
3.66
3.65
3.64
PEG-DIBO.txt
7.27
4.3 1HNMR of PEG-DIBO.
e
d
O
c
n
HN
O
3.67
solve
b
O
HN
a
3.80
3.79
3.74 3.73
7.30
7.29
7.29
7.28
a
5.56
5.51
e
7.5
7.0
6.5
6.0
4.5
5.0
5.5
Chemical Shift (ppm)
4.0
c
ds
3.52
3.51
3.51
3.50
3.41
3.20
3.16
3.70
3.70
a
3.64
3.64
3.62
3.69
3.68
O
O
3.5
3.0
2.5
Figure 4.61HNMR Spectrum of Dibenzylcyclooctyne Polyethylene glycol.
1
H NMR of Dibenzylcyclooctyne Polyethylene glycol:
1
H NMR (500 MHz, CDCl3): δ=7.51–7.23 (16H, aromatics), 5.51 (2H, dd,
J=3.9, 15.3 Hz, CHOH), 3.70-3.60 (~550H, s, OCH2CH2O), 3.40 (4H, d, J=6.4 Hz,
CH2NH), 3.20 (dd, 2H, J=3.9, 15.3 Hz, CH2), 2.91 (dd, 2H, J=3.9, 15.3 Hz, CH2).
37
4.4 MALDI-TOF MS of PEG-DIBO incorporating various molecular mass.
Figure 4.7MALDI-TOF mass spectrum of 2k PEG-DIBO.(a) full spectrum shows three
distributions with mass range from 2800 to 4600. (b) expansion showing the one main
series of peaks corresponding to K+ cationized molecules, where the other two minor
series correspond to the H+ and Na+ cationized molecules, respectively.
The molecular mass, molecular mass distribution and conversion of the functional
groups can be analyzed by matrix-assisted desorption ionization time-of-flight mass
spectroscopy MALDI-TOF MS. A MALDI-TOF MS of 2k PEG-DIBO (Figure 4.7(a))
clearly illustrates both high degree of purity and narrow molecular mass distribution. The
38
minor series correspond to variations in the molecular ion cationization. The main series
corresponds to K+ cationized PEG-DIBO. A minor second series of peaks corresponds to
H+cationized PEG-DIBO, and the third minor series may be attributed to Na+ cationized
species. H+ is due to the matrix acid, and K+ is in the matrix salt. The expanded mass
spectrum (Figure 4.7(b)) shows 44Da which represents the molecular mass of the repeat
unit for PEG.
Based on the MALDI-TOF MS data, the end-group analysis was performed to exhibit
a DIBO group is on each end.156The expansive spectrum contains a major distribution, A,
whose m/z values correspond to the K+ adducts of PEG-DIBO with DIBO end groups.
For example, the 38-mer of this distribution is expected to produce a signal at m/z
(monoisotopic) 38×44 (38×C2H4O)+2×276 (2×C18H14O2N) + 39.008 (39K+) = 2262, as
indeed observed. For the minor distribution, B, whose m/z values correspond to the H+
cationized PEG-DIBO. For example, the 38-mer of this series is expected to produce a
signal at m/z (monoisotopic) 38×44.02 (38×C2H4O)+2×276.86 (2×C18H14O2N) + 1.008
(1H+) = 2226, as indeed observed.
39
Figure 4.8MALDI-TOF MS of 3.5k PEG-DIBO. (a) full spectrum shows three
distributions with mass range from 2800 to 4600. (b) expansion showing the one main
series of peaks corresponding to K+ cationized molecules, where the other two minor
series correspond to the H+ and Na+ cationized molecules, respectively. The expansive
spectrum contains a major distribution, A, whose m/z values correspond to the K+
adducts of PEG-DIBO with DIBO end groups. For example, the 63-mer of this
distribution is expected to produce a signal at m/z (monoisotopic) 63×44 (63
×C2H4O)+2×276 (2×C18H14O2N) + 39.008 (39K+) = 3363, as indeed observed. For the
minor distribution, B, whose m/z values correspond to the H+cationized PEG-DIBO. For
example, the 64-mer of this series is expected to produce a signal at m/z (monoisotopic)
40
64×44 (64 ×C2H4O)+2×276.86 (2×C18H14O2N) + 1.008 (1H+) = 3369, as indeed
observed.
Figure 4.9MALDI-TOF MS of 5k PEG-DIBO. (a) full spectrum shows three distributions
with mass range from 3750 to 5750. (b) expansion showing the one main series of peaks
corresponding to K+ cationized molecules, where the other minor series correspond to the
H+ cationized molecules. The expansive spectrum contains a major distribution, A,
whose m/z values correspond to the K+ adducts of PEG-DIBO with DIBO end groups.
For example, the 91-mer of this distribution is expected to produce a signal at m/z
(monoisotopic) 91×44 (91 ×C2H4O)+2×276 (2×C18H14O2N) + 39.008 (39K+) = 4595, as
41
indeed observed. For the minor distribution, B, whose m/z values correspond to the
H+cationized PEG-DIBO. For example, the 92-mer of this series is expected to produce a
signal at m/z (monoisotopic) 92×44.02 (92 ×C2H4O)+2×276 (2×C18H14O2N) + 1.008
(1H+) = 4601, as indeed observed.
Figure 4.10MALDI-TOF MS of 6k PEG-DIBO. (a) full spectrum shows three
distributions with mass range from 4000 to 7000. (b) expansion showing the one main
series of peaks corresponding to K+ cationized molecules. The expansive spectrum
contains a major distribution, whose m/z values correspond to the K+ adducts of PEG42
DIBO with DIBO end groups. For example, the 114-mer of this distribution is expected
to produce a signal at m/z (monoisotopic) 114×44 (114 ×C2H4O)+2×276 (2×C18H14O2N)
+ 39.008 (39K+) = 5607, as indeed observed.
Figure 4.11MALDI-TOF MS of 7.5k PEG-DIBO. (a) full spectrum shows three
distributions with mass range from 6500 to 9000. (b) expansion showing the one main
series of peaks corresponding to K+ cationized molecules. The expansive spectrum
contains a major distribution, whose m/z values correspond to the K+ adducts of PEG43
DIBO with DIBO end groups. For example, the 150-mer of this distribution is expected
to produce a signal at m/z (monoisotopic) 150×44 (150 ×C2H4O)+2×276 (2×C18H14O2N)
+ 39.008 (39K+) = 7191, as indeed observed.
Figure 4.12MALDI-TOF mass spectrum of a series of PEG-DIBOs ranging in molecular
mass from 2k to 7.5k.
The MALDI-TOF MS of PEG-DIBOs synthesized from various molecular mass PEGs
are shown in Figure 4.12The relative signal intensities decrease as molecular mass
increases. The molecular mass of each sample can be clearly distinguished with all
polymers displaying the expected single molecular mass distribution.
44
4.5 Rheological studies of PEG-DIBO hydrogels.
The rheological studies have been used to characterize the mechanical properties and
gelation kinetics of the PEG-based hydrogels. Hydrogels are prepared by loading PEGDIBO in aqueous solution with the same volume of tetra-arm azide functionalized PEG
between two parallel plates by the strain to cross-link the PEG-DIBO. Since the solutions
are loaded between parallel plates, a certain amount of normal forces always applied in
the hydrogel to promote cross-linking.
Three rheological measurements were examined for the hydrogels as follows: a) a
frequency-sweep measurement using small strain allows us to determine a frequency to
ensure that the results we observe are gel formation and not influenced by the rheometer.
b) The subsequent strain-sweep measurement under a certain frequency obtained from
the frequency-sweep experiment suggests that subsequent time sweeps are carried out in
the linear viscoelastic region, and does not influence the gel formation and c) The timesweep measurement in which the storage modulus (G’) and loss modulus (G”) are
monitored as a function of time, using the value of strain and frequency obtained from the
first two measurements, will be used to quantitatively evaluate the gelation kinetics.
During the process of gel formation, G’ and G” both increase. At the beginning, G” is
larger than G’, since the sample is viscous liquid. Subsequently, a crossover of G’ and G”
can be observed, which exhibits the gelation point. After the gelation point, G’ becomes
greater than G”, which demonstrates the formation of a cross-linking network, suggesting
that the sample is becoming an elastic solid.G’ and G” continuously increase until they
both reach a plateau, indicating that the gel formation process is completed. The gelation
45
process is shown in Figure 4.13(a)The time at which the G’ reaches the plateau provides
a coarse estimate of the time it takes for complete network formation
Figure 4.13(b) shows storage modulus of PEG-DIBO hydrogels prepared with various
molecular mass (2.0×103 g/mol, 3.5×103 g/mol, 5.0×103 g/mol, 6.0×103 g/mol, and
7.5×103 g/mol). Under the certain PEG-DIBO, the mass fraction storage modulus was
expected to decrease as the molecular mass increased, presumably due to the lower crosslink density of high molecular mass. However, the variation between molecular mass and
storage modulus is not pronounced for hydrogels at lower molecular mass. Figure
4.13shows that hydrogels prepared from the 6.0×103 g/mol and 7.5×103 g/mol PEGDIBO do have a lower shear modulus than those prepared from the 2. 0 ×103 g/mol,
3.5×103 g/mol and 5.0×103 g/mol PEG-DIBO. There are no significant differences in the
shear modulus of hydrogels prepared from the 2. 0 ×103 g/mol, 3.5×103 g/mol and
5.0 × 103 g/mol PEG-DIBO. For quantification, Figure 4.14show the differences in
plateau modulus and gelation time for PEG-DIBO hydrogels, with various molecular
masses ranging from 2.0×103 g/mol to 7.5×103 g/mol. As the molecular mass of the
PEG-DIBO increases, it takes more time to form cross-links and reach the plateau
modulus. Thus, higher molecular mass PEG-DIBO resulted in hydrogels with lower
cross-link density, yielding a lower plateau modulus.
Higher molecular mass of a polymer corresponds to longer chains. As the polymer
chain length increases, distances between the cross-links also increase, which means the
cross-link density decreases. Lower cross-link density results in reduced mechanical
46
properties and hence a lower plateau modulus. In dilute solution, entropic effects
dominate, therefore the reaction time between DIBO end groups and azide groups
increases leading to longer gelation time.
(a)
47
(b)
Figure 4.13(a)Time sweep conducted under 10% strain and 1rad/s using 8mm parallel
plate. Gelation process was monitored by the time sweep for 15min for PEG-DIBO
hydrogels with 3.5k molecular mass.Gelation points were demonstrated by the crossover
of the storage modulus and loss modulus. (b)Storage modulus measured as a function of
time for aqueous PEG-DIBO hydrogels with different molecular masses (20% by mass
fraction). Higher molecular mass required longer time to reach plateau modulus.
48
Figure 4.14 Plateau modulus of hydrogels prepared using various molecular mass. Higher
molecular mass lead to higher plateau modulus.Gelation kinetics of hydrogels
incorporating various molecular mass. Higher molecular mass required longer gelation
time.
4.6 Swelling ratio experiments of PEG-DIBO hydrogels.
The swelling ratio experiments were also utilized to characterize the water uptake of
the hydrogels incorporating various molecular mass. In the current study the swelling
ratio of PEG-based hydrogel was adjusted by changing the initial PEG molecular mass
for PEG-DIBO synthesis. Theoretically, as the PEG molecular weight increases, the
spacing between the cross-linkers becomes larger and therefore the resulting hydrogel
will have a larger mesh size, exhibiting as a higher swelling ratio.157 Figure 4.15shows
49
the differences in swelling ratio for the PEG-DIBO hydrogels. The results of this study,
as expected, demonstrated a significant difference in swelling ratio between PEG-DIBO
hydrogels with different molecular mass. Especially, PEG-DIBO 6k and PEG-DIBO 7.5k
hydrogels were shown to have a statistically higher swelling ratio than that of PEG-DIBO
2k and PEG-DIBO 3.5k hydrogels.
Figure 4.15Swelling ratio of PEG-DIBO hydrogels. Higher molecular mass result in
higher swelling ratio.
50
CHAPTER V
CONCLUSIONS
PEG-based hydrogels with highly tunable mechanical properties were synthesized via
cross-linking reaction between dibenzocyclooctynol (DIBO) end-functionalized
poly(ethylene glycol) (PEG) and tetra-arm azide functionalized PEG using metal-free
strain-promoted alkyne-azidecycloaddition. The characterization of 1H NMR and
MALDI-TOF MS provided detailed information of chemical structure and molecular
mass for the PEG-DIBO molecular mass series. The PEG-DIBO covalently cross-linked
hydrogel was formed in aqueous solution, at 20% fraction mass. The storage modulus for
the molecular mass series decreased from 1.6kPa to 0.3kPa with increasing molecular
mass, which effectively increase the cross-link density. Gelation time increased from 20
seconds to 1 minute with increasing molecular mass as a result of the entropic effect.
Hydrogels prepared from the PEG-DIBO could be used as scaffolds in neuron cell
regeneration, because the mechanical properties can mimic the substrate elasticity of the
neuron cells.
In vivo studies, further characterizations of the network structure, as well as cell response
studies are currently being examined using this strategy for hydrogel formation.
51
CHAPTER VI
SUMMARY
This thesis outlines our efforts to develop PEG-based hydrogels that are soft,
biodegradable and biocompatible for cultivate neuron cells. We changed the molecular
weight of PEG, in the monomer, synthesizing three derivatives of Dibenzylcyclooctyne
Polyethylene glycol monomers and their PEG-based hydrogels using copper-free strainpromoted alkyne-azidecycloaddition. The chemical structure of the monomers has been
characterized by 1H-NMR. In addition, the mechanical properties and gelation kinetics
have been measured by oscillatory shear rheology experiments. The rheology result
showed that this tunable molecular weight allows for the variation of mechanical
properties. This further demonstrates the potential of PEG-based hydrogels as a suitable
polymeric material in neuron cultivation.
Further studies on the change of more molecular weight of PEG-based hydrogels and
microenvironment are also designed and will be done in the next following months.
52
REFERENCES
(1) Mullarney, M. P., Seery, T. A. P., & Weiss, R. A. Polymer. 2006, 47, 3845–3855.
(2) Li, Y., Huang, G., Zhang, X., Li, B., Chen, Y., Lu, T., Xu, F., Adv. Funct.
Mater.2013, 23, 660–672.
(3) Ladet, S., David, L., Domard, A., Nature. 2008, 452, 76–79.
(4) Rodell, C. B., Kaminski, A. L., Burdick, J. A., Biomacromolecules. 2013, 14, 4125–
4134.
(5) Grover, G. N.; Lam, J.; Nguyen, T. H.; Segura, T.; May-nard, H. D.
Biomacromolecules. 2012, 13, 3013-3017.
(6) Little, L.; Healy, K. E.; Schaffer, D., Chem. Rev. 2008, 108, 1787-1796.
(7) Balakrishnan, B.; Banerjee, R., Chem. Rev. 2011, 111, 4453-4474.
(8) Lee, K. Y.; Mooney, D. J., Chem. Rev. 2001, 101, 1869-1879.
(9) Kharkar, P. M.; Kiick, K. L.; Kloxin, A. M., Chem. Soc. Rev. 2013, 42, 7335-7372.
(10) Rehmann, M. S.; Kloxin, A. M., Soft Matter. 2013, 9, 6737-6746.
(11) Zheng, J. K.; Callahan, L. A. S.; Hao, J. K.; Guo, K.; Wesdemiotis, C.; Weiss, R. A.;
Becker, M. L. ACS Macro Letters. 2012, 1, 1071.
(12) Truong, V., Blakey, I., Whittaker, A. K,Biomacromo-lecules. 2012, 13, 4012–4021.
(13) Jewett, J. C.; Bertozzi, C. R. Chem. Soc. Rev. 2010, 39, 1272.
(14) Lin, F., Yu, J., Tang, W., Zheng, J., Defante, A., Guo, K., Wesdemiotis C., Becker,
M. L. Biomacromolecules. 2013, 14, 3749–3758.
53
(15) Alge, D. L., Azagarsamy, M. A., Donohue, D. F., An-seth, K. S.,
Biomacromolecules. 2013, 14, 949–95.
(16) Vermonden, T.; Censi, R.; Hennink, W. E. Chem. Rev. 2012, 112, 2853.
(17) Lutolf, M. P., P. M. Gilbert, et al., Nature. 2009, 462, 433-441.
(18) Place, E. S., N. D. Evans, et al. Nat Mater. 2009, 8, 457-470.
(19) Prang, P., R. Muller, et al., Biomaterials. 2006, 27, 3560-3569.
(20) Engler, A. J., Sen, S., Sweeney, H. L., Discher, D. E. Cell. 2006, 126, 677–689.
(21) Atzet, S.; Curtin, S.; Trinh, P.; Bryant, S.; Ratner, B., Biomacromolecules. 2008, 9,
3370-3377.
(22) Choh, S. Y.; Cross, D.; Wang, C., Biomacromolecules. 2011, 12, 1126-1136.
(23) Ekenseair, A. K.; Boere, K. W. M.; Tzouanas, S. N.; Vo, T. N.; Kasper, F. K.;
Mikos, A. G., Biomacromolecules. 2012, 13, 1908-1915.
(24) Jiang, Y. J.; Chen, J.; Deng, C.; Suuronen, E. J.; Zhong, Z. Y., Biomaterials. 2014,
35, 4969-4985.
(25) Lin, Y. H.; Liang, H. F.; Chung, C. K.; Chen, M. C.; Sung, H. W., Biomaterials.
2005, 26, 2105-2113.
(26) Ma, Y., Zheng, J., Amond, E. F., Stafford, C. M., & Becker, M. L.,
Biomacromolecules. 2013, 14, 665–671.
(27) Ornelas C., Ruiz Aranzaes J., Cloutet E., Alves S., Astruc D., Angew. Chem. Int. Ed.
2007, 46, 872–877.
(28) Meldal M., Tornøe C.W. Chem. Rev. 2008, 108, 2952.
(29) Tang, W.; Policastro, G. M.; Hua, G.; Guo, K.; Zhou, J.; Wesdemiotis, C.; Doll, G.
L.; Becker, M. L., J. Am. Chem. Soc. 2014, 136, 16357.
(30) Long, R., Mayumi, K., Creton, C., Narita, T., Hui, C-Y.,Macromolecules. 2014, 47,
7243−7250.
(31) Lin, Z. F.; Cao, S. Q.; Chen, X. Y.; Wu, W.; Li, J. S., Biomacromolecules. 2013, 14,
2206-2214.
(32) Kraehenbuehl, T. P.; Zammaretti, P.; Van der Vlies, A. J.; Schoenmakers, R. G.;
Lutolf, M. P.; Jaconi, M. E.; Hubbell, J. A., Biomaterials.2008, 29, 2757-2766.
54
(33) Moffat, K. L.; Marra, K. G., J Biomed Mater Res B. 2004, 71, 181-187.
(34) Missirlis, D.; Spatz, J. P., Biomacromolecules. 2014, 15, 195-205.
(35) Drury, J. L.; Mooney, D. J.; Biomaterials. 2003, 24, 4337-4351.
(36) Langer, R; Tirrell, D. A.; Nature. 2004, 428, 487-492.
(37) Kolb, H.C.; Finn, M.G.; Sharpless, B.K.Angew. Chem. Int. Ed.2001, 40, 2004-2021.
(38) Ning, X., Guo, J., Wolfert, M., Boons, G. J,Angew. Chem., Int. Ed. 2008, 47, 22532255.
(39) Oshima, K., Fujimoto, T., Minami, E., Mitsukami, Y., Macromolecules.2014, 47,
7573−7580.
(40) DeForest, C. A.; Polizzotti, B. D.; Anseth, K. S. Nat. Mater. 2009, 8, 659–664.
(41) Fairbanks, B. D.; Sims, E. A.; Anseth, K. S.; Bowman, C. N., Macromolecules.
2010, 43, 4113–4119.
(42) Huang, Z. J.; Liu, X. W.; Chen, S. S.; Lu, Q. H.; Sun, G., PolymChem-Uk. 2015, 6,
143.
(43) Lin, C. C.; Metters, A. T., Biomacromolecules. 2008, 9, 789-95.
(44) Gibson, S. L., Bencherif, S., Cooper, J. A., Wetzel, S. J., Antonucci, J. M., Vogel, B.
M., Horkay, F., Washburn, N. R.,Biomacromolecules. 2004, 5, 1280-1287.
(45) Wu, P., Feldman, A. K., Nugent, A. K., Hawker, C. J., Scheel, A., Voit, B., Pyun, J.,
Jean M. J., Sharpless, K. B., Fokin, V. V., Angew. Chem. Int. Ed. 2004, 43, 3928 –3932.
(46) Frisman, I.; Shachaf, Y.; Seliktar, D.; Bianco-Peled, H., Langmuir.2011, 27, 69776986.
(47) Benoit, D. S. W.; Schwartz, M. P.; Durney, A. R.; Anseth, K. S., Nat. Mater. 2008,
7, 816-823.
(48) Mason, M. N.; Metters, A. T.; Bowman, C. N.; Anseth, K. S.; Macromolecules2001,
31, 4630.
(49) Moses, J. E.; Moorhouse, A. D., Chem.Soc. Rev. 2007, 36, 1249-1262.
(50) Zhao, D.; Tan, S. W.; Yuan, D. Q.; Lu, W. G.; Rezenom, Y. H.; Jiang, H. L.; Wang,
L. Q.; Zhou, H. C., Adv Mater. 2011, 23, 90.
55
(51) Bernardin, A.; Cazet, A.; Guyon, L.; Delannoy, P.; Vinet, F.; Bonnaffe, D.; Texier,
I., Bioconjugate Chem. 2010, 21, 583-588.
(52) Tron, G. C.; Pirali, T.; Billington, R. A.; Canonico, P. L.; Sorba, G.; Genazzani, A.
A., Med Res Rev.2008, 28, 278-308.
(53) Ackermann, L.; Potukuchi, H. K.; Landsberg, D.; Vicente, R., Org Lett.2008, 10,
3081-3084.
(54) Huisgen, R. Angew. Chem. Int. Ed. Engl. 1963, 2, 565–598.
(55) Sisson, A. L.; Papp, I.; Landfester, K.; Haag, R. Macromolecules 2008, 42, 556–559.
(56) Clark, M.; Kiser, P. Polym. Int. 2009, 58, 1190–1195.
(57) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. Angew. Chem. Int.
Ed. 2002, 41, 2596–2599.
(58) Tornoe, C. W.; Christensen, C.; Meldal, M. J. Org. Chem. 2002, 67, 3057–3064.
(59) Himo, F.; Lovell, T.; Hilgraf, R.; Rostovtsev, V. V.; Noodleman, L.; Sharpless, K.
B.; Fokin, V. V. J. Am. Chem. Soc. 2004, 127, 210–216.
(60) Rodionov, V. O.; Fokin, V. V.; Finn, M. G. Angew. Chem. Int. Ed. 2005, 44, 2210–
2215.
(61) Rodionov, V. O.; Presolski, S. I.; Diaz Diaz, D.; Fokin, V. V.; Finn, M. G. J. Am.
Chem. Soc. 2007, 129, 12705–12712.
(62) Bock, V. D.; Hiemstra, H.; van Maarseveen, J. H. Eur. J. Org. Chem. 2006, 2006,
51–68.
(63) Fournier, D.; Hoogenboom, R.; Schubert, U. S. Chem. Soc. Rev. 2007, 36, 1369–
1380.
(64) Lutz, J.-F. Angew. Chem. Int. Ed. 2007, 46, 1018–1025.
(65) vanBerkel, S. S.; Dirks, A. J.; Debets, M. F.; van Delft, F. L.; Cornelissen, J. J. L.
M.; Nolte, R. J. M.; Rutjes, F. P. J. T. ChemBioChem2007, 8, 1504–1508.
(66) Tron, G. C.; Pirali, T.; Billington, R. A.; Canonico, P. L.; Sorba, G.; Genazzani, A.
A. Med. Res. Rev. 2008, 28, 278–308.
(67) Hein, C.; Liu, X.-M.; Wang, D. Pharm. Res. 2008, 25, 2216–2230.
56
(68) Malkoch, M.; Vestberg, R.; Gupta, N.; Mespouille, L.; Dubois, P.; Mason, A. F.;
Hedrick, J. L.; Liao, Q.; Frank, C. W.; Kingsbury, K.; Hawker, C. J. Chem. Commun.
2006, 2774–2776.
(69) Liu, S. Q.; Rachel Ee, P. L.; Ke, C. Y.; Hedrick, J. L.; Yang, Y. Y. Biomaterials
2009, 30, 1453–1461.
(70) Jonkheijm, P.; Weinrich, D.; Kohn, M.; Engelkamp, H.; Christianen, P. C.;
Kuhlmann, J.; Maan, J. C.; Nusse, D.; Schroeder, H.; Wacker, R.; Breinbauer, R.;
Niemeyer, C. M.; Waldmann, H., AngewChemInt Ed Engl.2008, 47, 4421-4424.
(71) Killops, K. L.; J Am Chem Soc. 2008, 130, 5062-5064.
(72) Kade, M. J.; Burke, D. J.; Hawker, C. J., J PolymSci Pol Chem. 2010, 48, 743-750.
(73) Zheng, Z. L.; Perkins, B. L.; Ni, B. K., J. Am. Chem. Soc.2010, 132, 50-51.
(74) Park, S.; Yousaf, M. N., Langmuir. 2008, 24, 6201-6207.
(75) Kagan, H. B.; Riant, O., Chem. Rev. 1992,92, 1007-1019.
(76) Brieger, G.; Bennett, J. N., Chem. Rev. 1980, 80, 63-97.
(77) Hein, J. E.; Fokin, V. V., Chem. Soc. Rev.2010, 39, 1302-15.
(78) Finn, M. G.; Fokin, V. V., Chem. Soc. Rev.2010, 39, 1231-2.
(79) Xiao, J.; Tolbert, T. J., Org. Lett.2009, 11, 4144-4147.
(80) Shao, C.; Wang, X.; Xu, J.; Zhao, J.; Zhang, Q.; Hu, Y., J. Org. Chem. 2010,75,
7002-5.
(81) Tan, H.; Rubin, J. P.; Marra, K. G.Macromol. Rapid Commun.2011, 32, 905−911.
(82) Nimmo, C. M.; Shoichet, M. S.Bioconjugate Chem. 2011, 22,2199−2209.
(83) Hashidzume, A.; Tomatsu, I.; Harada, A. Polymer2006, 47,6011.Vol.
(84) Iha, R. K.; Wooley, K. L.; Nystrom, A. M.; Burke, D. J.; Kade, M.J.; Hawker, C. J.
Chem. Rev. 2009, 109, 5620−5686.
(85)] Azagarsamy.M.A.;Anseth.K. S.;Angew. Chem. Int. Ed.2013, 52, 13803 –13807.
(86) Sletten, E. M.; Bertozzi, C. R. Angew. Chem. Int. Ed. 2009, 48, 6974–6998.
(87) Agard, N. J.; Prescher, J. A.; Bertozzi, C. R. J. Am. Chem. Soc. 2004, 126, 15046–
15047.
57
(88) Ornelas, C.; Broichhagen, J.; Weck, M., J. Am. Chem. Soc. 2010, 132, 3923-31.
(89) Yao, J. Z.; Uttamapinant, C.; Poloukhtine, A.; Baskin, J. M.; Codelli, J. A.; Sletten,
E. M.; Bertozzi, C. R.; Popik, V. V.; Ting, A. Y., J. Am. Chem. Soc.2012, 134 (8), 37208.
(90)Zheng, J.; Liu, K.; Reneker, D. H.; Becker, M. L., J. Am. Chem. Soc. 2012, 134,
17274-7.
(91) Manova, R.; van Beek, T. A.; Zuilhof, H., Angew. Chem. Int. Ed. Engl.2011, 50,
5428-30.
(92) Manova, R. K.; Pujari, S. P.; Weijers, C. A.; Zuilhof, H.; van Beek, T. A., Langmuir.
2012, 28, 8651-63.
(93) Marks, I. S.; Kang, J. S.; Jones, B. T.; Landmark, K. J.; Cleland, A. J.; Taton, T. A.,
Bioconjug Chem. 2011, 22, 1259-63.
(94) Pratt, M. R.; Bertozzi, C. R., J. Am. Chem. Soc.2003, 125 (20), 6149-59.
(95) Bencherif, S. A.; Washburn, N. R.; Matyjaszewski, K.Biomacromolecules2009, 10,
2499−2507.
(96) Jen, A. C.; Wake, M. C.; Mikos, A. G., BiotechnolBioeng. 1996, 50 (4), 357-64.
(97) Hoffman, A. S., Adv Drug Deliv Rev.2002, 54, 3-12.
(98) Nguyen, K. T.; West, J. L., Biomaterials. 2002, 23, 4307-14.
(99) Nicodemus, G. D.; Bryant, S. J., Tissue Eng Part B Rev. 2008, 14, 149-65.
(100)Van Vlierberghe, S.; Dubruel, P.; Schacht, E., Biomacromolecules.2011, 12, 13871408.
(101) Li, Y. L.; Rodrigues, J.; Tomas, H., Chem. Soc. Rev.2012, 41 (6), 2193-2221.
(102) Lin, C. C.; Metters, A. T., Adv Drug Deliver Rev.2006, 58, 1379-1408.
(103) Jeong, B.; Kim, S. W.; Bae, Y. H., Adv Drug Deliv Rev. 2002, 54, 37-51.
(104) Jen, A. C.; Wake, M. C.; Mikos, A. G., BiotechnolBioeng.1996, 50 (4), 357-64.
(105) Reinhoudt, D.A. et al. Chem. Bio.Chem.2007, 8, 1997-2002.
(106) Pochan, D. J.; Schneider, J. P.; Kretsinger, J.; Ozbas, B.;Rajagopal, K.; Haines, L.
J. Am. Chem. Soc. 2003, 125, 11802−11803.
58
(107)Peppas, N. A.; Hilt, J. Z.; Khademhosseini, A.; Langer, R. Adv.Mater.2006, 18,
1345.
(108) Seal, B. L.; Panitch, A.Biomacromolecules2003, 4, 1572−1582.
(109) Laughlin, S. T.; Bertozzi, C. R. ACS Chem. Biol. 2009, 4, 1068−1072.
(110) S. Punna, E. Kaltgrad, M.G. Finn, Bioconjugate Chem. 2005, 16,1536
(111) Malik, N. N. Drug Discovery Today 2008, 13, 909.
(112) Kim, I. L.; Mauck, R. L.; Burdick, J. A. Biomaterials2011, 32,8771−8782.
(113) Knop, K.; Hoogenboom, R.; Fischer, D.; Schubert, U. S. Angew. Chem. Int. Ed.
2010, 49, 6288–6308.
(114) Peppas, N. A.; Hilt, J. Z.; Khademhosseini, A.; Langer, R. Adv. Mater.2006, 18,
1345–1360.
(115) Metters, A.; Hubbell, J. Biomacromolecules2004, 6, 290–301.
(116) DiRamio, J. A.; Kisaalita, W. S.; Majetich, G. F.; Shimkus, J. M.
Biotechnol.Prog.2005, 21, 1281–1288.
(117) Lutolf, M. P.; Lauer-Fields, J. L.; Schmoekel, H. G.; Metters, A. T.; Weber, F. E.;
Fields, G. B.; Hubbell, J. A. Proc. Natl. Acad. Sci. USA 2003, 100, 5413–5418.
(118) Zustiak, S. P.; Leach, J. B. Biomacromolecules2010, 11, 1348–1357.
(119) Garcia, A. Ann. Biomed. Eng. 2014, 42, 312–322.
(120) Adzima, B. J.; Tao, Y.; Kloxin, C. J.; DeForest, C. A.; Anseth, K. S.; Bowman, C.
N. Nat. Chem. 2011, 3, 256–259.
(121) Elbert, D. L.; Hubbell, J. A. Biomacromolecules2001, 2, 430–441.
(122) Sakai, T.; Matsunaga, T.; Yamamoto, Y.; Ito, C.; Yoshida, R.; Suzuki, S.; Sasaki,
N.; Shibayama, M.; Chung, U.-i. Macromolecules 2008, 41, 5379–5384.
(123) Roberts, J. J.; Bryant, S. J. Biomaterials 2013, 34, 9969–9979.
(124) Lei, J.; Mayer, C.; Freger, V.; Ulbricht, M. Macromol.Mater. Eng. 2013, 298, 967–
980.
(125) Falender, J. R.; Yeh, G. S. Y.; Mark, J. E. J. Am. Chem. Soc. 1979, 101, 7353–
7356.
59
(126) Falender, J. R.; Yeh, G. S. Y.; Mark, J. E. Macromolecules 1979, 12, 1207–1209.
(127) Mark, J. E.; Tang, M. Y. J. Polym.Sci., Polym. Phys. Ed. 1984, 22, 1849–1855.
(128) Tang, M. Y.; Mark, J. E. Macromolecules 1984, 17, 2616–2619.
(129) Niece, K. L.; Czeisler, C.; Sahni, V.; Tysseling-Mattiace, V.;Pashuck, E. T.;
Kessler, J. A.; Stupp, S. I.Biomaterials2008, 29, 4501−4509.
(130) Khademhosseini, A.; Langer, R., Biomaterials.2007, 28 (34), 5087-92.
(131) ZhengShu, X.; Liu, Y.; Palumbo, F. S.; Luo, Y.; Prestwich, G. D.,
Biomaterials.2004,25, 1339-48.
(132) Zhu, J., Biomaterials.2010, 31, 4639-56.
(133) Lutolf, M. P.; Hubbell, J. A., Nat Biotechnol.2005, 23 (1), 47-55.
(134) Leach, J. B.; Schmidt, C. E., Biomaterials 2005,26 (2), 125-135.
(135) Salmaso, S.; Semenzato, A.; Bersani, S.; Matricardi, P.; Rossi, F.; Caliceti, P., Int J
Pharm 2007,345, 42-50.
(136) Bjugstad, K. B.; Redmond, D. E., Jr.; Lampe, K. J.; Kern, D. S.; Sladek, J. R., Jr.;
Mahoney, M. J., Cell Transplant. 2008,17, 409-15.
(137) Andreopoulos, F. M.; Roberts, M. J.; Bentley, M. D.; Harris, J. M.; Beckman, E. J.;
Russell, A. J., BiotechnolBioeng. 1999,65 (5), 579-588.
(138) Stahl, P. J.; Romano, N. H.; Wirtz, D.; Yu, S. M., Biomacromolecules.2010,11,
2336-2344.
(139) Pfister, P. M.; Wendlandt, M.; Neuenschwander, P.; Suter, U. W.,
Biomaterials.2007,28, 567-575.
(140) Zheng, Y. J.; Mieie, M.; Mello, S. V.; Mabrouki, M.; Andreopoulos, F. M.; Konka,
V.; Pham, S. M.; Leblanc, R. M., Macromolecules.2002,35 (13), 5228-5234.
(141) Burdick, J. A.; Anseth, K. S., Biomaterials.2002,23 (22), 4315-4323.
(142) Li, Y.; Yang, C.; Khan, M.; Liu, S. Q.; Hedrick, J. L.; Yang, Y. Y.; Ee, P. L. R.,
Biomaterials. 2012,33, 6533-6541.
(143) Rao, S. S.; Han, N.; Winter, J. O., J BiomaterSciPolym Ed. 2011,22, 611-25.
(144) Kloxin, A. M.; Kasko, A. M.; Salinas, C. N.; Anseth, K. S., Science 2009,324, 5963.
60
(145) Van Dijk, M.; van Nostrum, C. F.; Hennink, W. E.; Rijkers, D. T.; Liskamp, R. M.,
Biomacromolecules. 2010,11, 1608-1614.
(146) Russell, R. J.; Pishko, M. V.; Gefrides, C. C.; McShane, M. J.; Cote, G. L., Anal
Chem. 1999,71 (15), 3126-3132.
(147) Lutolf, M. R.; Weber, F. E.; Schmoekel, H. G.; Schense, J. C.; Kohler, T.; Muller,
R.; Hubbell, J. A., Nat Biotechnol. 2003,21, 513-518.
(148) Hahn, M. S.; Taite, L. J.; Moon, J. J.; Rowland, M. C.; Ruffino, K. A.; West, J. L.,
Biomaterials. 2006,27, 2519-2524.
(149) Nuttelman, C. R.; Benoit, D. S. W.; Tripodi, M. C.; Anseth, K. S.,
Biomaterials.2006,27, 1377-1386.
(150) Sanborn, T. J.; Messersmith, P. B.; Barron, A. E., Biomaterials.2002,23, 27032710.
(151) Peppas, N. A.; Hilt, J. Z.; Khademhosseini, A.; Langer, R., Adv Mater.2006,18,
1345-1360.
(152) Lu, S. X.; Anseth, K. S., Macromolecules.2000,33, 2509-2515.
(153) Qi, M.; Strand, B. L.; Morch, Y.; Lacik, I.; Wang, Y.; Salehi, P.; Barbaro, B.;
Gangemi, A.; Kuechle, J.; Romagnoli, T.; Hansen, M. A.; Rodriguez, L. A.; Benedetti,
E.; Hunkeler, D.; Skjak- Brak, G.; Oberholzer, J. Artif. Cells Blood
Substit.Biotechnol.2008, 36, 403–420.
(154) Elliott, R. B.; Escobar, L.; Calafiore, R.; Basta, G.; Garkavenko, O.; Vasconcellos,
A.; Bambra, C. Transplant. Proc. 2005, 37, 466–469.
(155) Qi, M. Advances in Medicine 2014, 2014, DOI: 10.1155/2014/429710.
(156) Arnould, M. A., Polce, M. J., Quirk, R. P., Wesdemio-tis, C., Int. J. Mass spectrom.
2004, 238, 245–255.
(157) Park, H., Temenoff, J. S., Tabata, Y., Caplan, A. I., Raphael, R. M., Jansen, J. A.,
Mikos, A.G., J. Biomed. Mater. Res. 2009, 88, 889–897.
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APPENDIX
Rheological studies of series PEG-DIBO hydrogels with different molecular mass.
Figure A1Time sweep conducted under 10% strain and 1rad/s using 8mm parallel plate.
Gelation process was monitored by the time sweep for 12min for PEG-DIBO hydrogels
with 2k molecular mass.
62
Figure A2Time sweep conducted under 10% strain and 1rad/s using 8mm parallel plate.
Gelation process was monitored by the time sweep for 20min for PEG-DIBO hydrogels
with 5k molecular mass.
63
Figure A3Time sweep conducted under 10% strain and 1rad/s using 8mm parallel plate.
Gelation process was monitored by the time sweep for 120min for PEG-DIBO hydrogels
with 6k molecular mass.
64
Figure A4Time sweep conducted under 10% strain and 1rad/s using 8mm parallel plate.
Gelation process was monitored by the time sweep for 150min for PEG-DIBO hydrogels
with 7.5k molecular mass.
65