Mechanical Properties and Cell Compatibility of Agarose Hydrogels

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Mechanical Properties and Cell Compatibility of Agarose Hydrogels Containing Proteoglycan Mimetic Graft Copolymers Hannah M. Pauly, Laura W. Place, Tammy L. Haut Donahue, and Matt J Kipper Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.7b00643 • Publication Date (Web): 15 Jun 2017 Downloaded from http://pubs.acs.org on June 17, 2017

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Mechanical Properties and Cell Compatibility of Agarose Hydrogels Containing Proteoglycan Mimetic Graft Copolymers Hannah M. Pauly,†,‡ Laura W. Place,†,‡ Tammy L. Haut Donahue,†,§ and Matt J. Kipper*,†,ǁ †

School of Biomedical Engineering, §Department of Mechanical Engineering, and ǁDepartment

of Chemical and Biological Engineering, Colorado State University, 1370 Campus Delivery, Fort Collins, Colorado, United States KEYWORDS: Hydrogel, agarose, proteoglycan, glycosaminoglycan, polysaccharide, stress relaxation, cyclic compression

ABSTRACT

Proteoglycans have vital biochemical and biomechanical functions. Their proteolytic degradation results in loss of these functions. We have previously reported non-protein proteoglycan-mimetic graft copolymers that stabilize and deliver growth factors and are not subject to proteases. Here we expand our investigation of these proteoglycan mimics by also investigating their effects on hydrogel mechanical properties. Four polysaccharide side chains – chondroitin sulfate, heparin,

ACS Paragon Plus Environment

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dextran, and dextran sulfate – are each grafted to a hyaluronan backbone. The polysaccharides and graft copolymers are added to agarose hydrogels. Cyclic compression and stress relaxation tests reveal how the addition of the polysaccharides and graft copolymers influences hydrogel modulus. Cells encapsulated in agarose hydrogels containing chondroitin sulfate and the chondroitin sulfate graft copolymer have decreased cell viability and metabolic activity compared to cells in unmodified agarose hydrogels. These multifunctional additives can be used to improve both the biochemistry and biomechanics of materials, warranting further optimization to overcome the potentially negative effects these may have on cell viability and activity.

INTRODUCTION Proteoglycans are polyanionic glycosylated proteins composed of a core protein with covalently attached glycosaminoglycan (GAG) side chains.1 The GAG side chains are linear polysaccharides bearing anionic carboxylate and sulfate groups.2 Proteoglycans are found throughout the extracellular matrix of many human musculoskeletal tissues. The molecular weight of proteoglycan core proteins varies greatly. Decorin, biglycan, and fibromodulin core proteins have molecular weights from 36-42 kDa while larger proteoglycans such as versican, perlecan, and aggrecan can have core protein molecular weights of up to 220-470 kDa.1 Furthermore, the degree of modification with GAG side chains also varies. For example, decorin and biglycan have just one or two GAG side chains, while some isoforms of versican have over 10, and aggrecan can have over 100 GAG side chains.2 The degree of negative charge, the type, and the number of GAG side chains can all affect the structure, biochemical activity, and biomechanical functions of proteoglycans.


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Articular cartilage is a load-bearing, joint lubricating tissue that covers the articulating ends of long bones. The principle functions of articular cartilage are to provide a smooth lubricated surface for bone articulation and to facilitate load transfer between bones.3,4 Cartilage consists of type II collagen, chondrocytes, interstitial fluid, and proteoglycans, with the proteoglycans accounting for 10% to 15% of the weight of cartilage.3 Perlecan is a cartilage proteoglycan with heparan sulfate side chains that has been implicated in organizing and stabilizing the cartilage extracellular matrix, promoting cell attachment, and sequestering growth factors.5 Versican is a chondroitin sulfate proteoglycan implicated in creating an environment conducive to cell migration, proliferation, and differentiation during joint development.6,7 The largest and most abundant proteoglycan found in cartilage is aggrecan, which consists of a protein core with attached negatively charged chondroitin sulfate and keratan sulfate side chains.8,9 Aggrecan can interact with hyaluronan via link proteins which allows multiple aggrecan proteoglycans to connect together to form a larger proteoglycan aggregate.8,9 The architecture of these aggrecan aggregates plays an important role in the mechanical properties of cartilage, particularly in compression. In addition to the solid matrix components listed above, articular cartilage is 70% interstitial fluid by volume, and that fluid consists of a variety of inorganic ions, including sodium, calcium, chloride, and potassium.3 The electrostatic forces between the negatively charged proteoglycan aggregates and the osmotic pressure imparted by ions in the interstitial fluid enable cartilage to resist compressive loading.3,10 The functional integrity of articular cartilage is dependent on the presence and structure of these proteoglycans. Many proteoglycans are subject to enzymatic degradation. In cartilage, the loss and destruction of proteoglycans is primarily due to proteolytic cleavage of the core protein by either matrix metalloproteinases or aggrecanases.11 The structure of the GAG side chains may also be altered,


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changing the function of the proteoglycans. Proteoglycan degradation has been shown to occur as a result of traumatic loading, and has been observed in the early stages of osteoarthritis.9,10. Breakdown of cartilage proteoglycans may prevent the cartilage tissue from properly performing its load bearing function.12 Because proteoglycans serve such a wide variety of biological functions, there is interest in synthesizing proteoglycan mimics which are not subject to natural proteolysis. Proteoglycan mimics have been investigated by other research groups for a variety of purposes, including acting as signaling or cell adhesion molecules, sequestering growth factors, or establishing chemokine gradients.13 The bioactivity of chondroitin sulfate proteoglycans, for example, has been mimicked using a novel end-functionalized ring-opening metathesis polymerization polymer.14 Additionally these chondroitin sulfate-mimics can be incorporated into collagen scaffolds to be used for tissue engineering applications.15 Sharma et al. created an aggrecan-mimetic molecule using magnetically aligned collagen with peptidoglycans to enhance the bulk modulus of collagen scaffolds.16 These molecules were also resistant to cytokineinduced degradation, which would be ideal for long term in vitro or in vivo use.16 Heparin-bound star-poly(ethylene glycol) (star-PEG) gels have been investigated for a variety of uses including releasing growth factors and anchoring proteins and peptides that enable cell growth.17,18 Relatively few studies have assessed the influence of proteoglycan mimics on hydrogel mechanics. We are interested in developing proteoglycan-mimetic, GAG-containing nanomaterials with tunable compositions and functions.19–21 We recently reported graft copolymers formed by covalently attaching GAG chains to a modified hyaluronan backbone.19 These proteoglycanmimetic graft copolymers were formed with either heparin or chondroitin sulfate GAG side


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chains, and the side-chain graft density could be altered to modulate the graft-copolymer size.19 We showed that these graft copolymers could bind, stabilize, and deliver the GAG-binding growth factor, fibroblast growth factor-2.19 The effects of the side chain composition on mechanical properties and cellular response have not yet been investigated. In the present work, we expand the composition of these graft copolymers to also include the non-GAG polysaccharides, dextran and dextran sulfate as side chains, and we investigate their influence on the mechanical properties of agarose hydrogels. Dextran sulfate is included because it has been used as a GAG analog to deliver growth factors.22 Dextran is included to compare the polyanionic GAGs to a non-charged polysaccharide. By grafting polysaccharide side chains (either chondroitin sulfate, dextran, dextran sulfate, or heparin) to a hyaluronan backbone, proteoglycan-mimetic graft copolymers can be synthesized with tunable side chain compositions. The goal of the present work is to discern how varying the side chain composition affects their mechanical properties and cellular response in the context of an agarose hydrogel.

MATERIALS AND METHODS Polysaccharide and Graft Copolymer Size and Zeta Potential Characterization. The synthesis of the proteoglycan-mimetic graft copolymers followed our previously published procedure.19,23 The details of the modification of hyaluronan (HA; MW = 470 kDa, Thermo Fisher Scientific, Waltham, MA) to contain pendent thiol groups (HA-SH), followed by conversion to hydrazide-modified hyaluronan, are provided in the Supporting Information (Scheme SI.1, and SI.2). These are analyzed by 1H NMR and an Ellman’s reagent assay, as reported in the Supporting Information.19,24,25 To create the graft copolymers (co-Hep, co-CS, coDex, and co-DS) the hydrazide-modified hyaluronan was combined with either heparin sodium


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(Hep; MW = 14.4 kDa, 12.5% sulfur by mass, Sigma-Aldrich, St Louis, MO), chondroitin sulfate sodium salt (CS; MW = 84 kDa, 6% sulfur, 6-sulfate/4-sulfate = 1.24, Sigma-Aldrich, St. Louis, MO), dextran (Dex; MW = 40 kDa, Sigma Aldrich, St. Louis, MO), or dextran sulfate (DS; MW = 36 – 50 kDa, 3.8 sulfate groups per disaccharide, MP Biomedicals, Solon, OH). The aldehyde form of the reducing end of the side chain polysaccharides was reacted with the hydrazide functional group by reductive animation (Scheme 1).26,27 The side chain polysaccharide (Hep, CS, Dex, or DS) was added to the hydrazide-modified HA at a ratio of one side chain per two-disaccharide repeat units in the HA backbone. (This represents a 1:1 stoichiometry of side chain to functional groups on the HA backbone, according to our analysis of the HA modification in the Supporting Information). On a mass basis, the ratio of side-chain to backbone ranges from about 1:19 for the heparin, to about 1:99 for the CS. The molecular weights, hydrodynamic diameters, and sulfate substituent composition of these four side chain polymers are listed in Table 1.


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Scheme 1. Formation of graft copolymer by reductive amination to couple the reducing end of a polysaccharide to a hydrazide-modified hyaluronan. Thiolation and hydrazide modification of the hyaluronan are detailed in the Supporting Information (Schemes SI.1, and SI.2). Table 1. Molecular weights, hydrodynamic diameters, and sulfate composition of proteoglycanmimetic graft polymer components. Molecular weight

Hydrodynamic

Average number of SO4−

(kDa)

diameter, Dh (nm)

per disaccharide

Heparin (Hep)

14.4

4.1

2.5

Dextran (Dex)

40

6.0

0

Dextran sulfate (DS)

36-50

5.5

3.9

Chondroitin sulfate

84

7.8

1-2

740

13.0

0

Polysaccharide

(CS) Hyaluronan (HA)

Dynamic light scattering (DLS) and electrophoretic light scattering (ELS) were conducted using a ZetaSizer Nano ZS (Malvern Instruments, Westborough, MA) to measure the hydrodynamic diameter of the constituent polysaccharides and the resulting graft copolymers, and the zeta potential of the graft copolymers. Samples were dissolved in PBS at 5 mg/mL and measurements were taken at 25 °C.


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Acellular Hydrogel Fabrication. To fabricate acellular hydrogels, a 2.25 weight % agarose solution was created by mixing agarose (Fisher Scientific, Hampton, NH) with phosphatebuffered saline (PBS) and heating the solution in an autoclave to 260 °C, to ensure complete agarose dissolution. The solution was allowed to cool to 60 °C. The solution was maintained at 60 °C while an additional 0.25 weight % of the polysaccharide (HA, CS, Hep, Dex, or DS) or graft copolymer (co-CS, co-Hep, co-Dex, or co-DS) was added to the 2.25 weight % agarose solution and stirred for 10 min. This produced a final 2.5 weight % solution. A plain 2.25% agarose gel was included as a control. Agarose hydrogels with 2.0 weight % and 2.5 weight % were also prepared as additional controls. Hydrogel solutions were cast into a stainless steel mold, allowed to cool for 15 min at room temperature, and then released to form a 10-mm thick hydrogel sheet. Cylindrical samples were created using a 10-mm diameter biopsy punch. These samples were stored submerged in phosphate-buffered saline (PBS) to prevent dehydration. We assume that this method of incorporating the polysaccharide and copolymer additives during the gelation results in a uniform and stable distribution of additive throughout the gel. This is confirmed by mechanical property measurements on multiple replicate samples of the same type, over the course of several hours and days. Mechanical Evaluation – Cyclic Compression. Acellular agarose hydrogels (n = 4 replicates per sample type) containing polysaccharides or graft copolymers were mechanically tested between steel platens using a servo-hydraulic mechanical testing system (MTS; Bionic Model 370.02 MTS Systems Corporation, Eden Prairie, MN) equipped with a 0.25 lb load cell (Futek LRF300, Irvine, CA). Hydrogels were kept hydrated during testing via immersion in a roomtemperature PBS bath. Hydrogels were fixed to the lower platen with a 50 µl drop of cyanoacrylate adhesive.


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A preload of 0.01 N was applied followed by 10% strain for 100 cycles under dynamic uniaxial unconfined compression. Cyclic testing was performed at 0.05 Hz, 0.5 Hz, 1 Hz, 2 Hz, and 5 Hz to assess strain rate dependence of the mechanical properties. The height of each sample was measured before and after cyclic compression testing. Cyclic compression testing did not result in significant height change for any of the sample types (Supporting Information, Figure SI.3). Data were Fourier filtered to remove high-frequency noise. There were no significant differences between the data from the 2nd loading cycle and the 99th loading cycle, so only the 2nd loading cycle was analyzed. The filtered stress versus strain data for the second loading cycle was fit to an exponential of the form26 − = −1 −

(1)

Here, −σ33 is the compressive stress and (1−λ) is the compressive strain. This equation has been previously used for fitting compression experiments of bovine articular cartilage.26 The modulus at 0% strain (Emin) and the modulus at 10% strain (Emax) were calculated from the derivative of equation 1 as follows:

=

=

− = 1 −

− = . 1 − .

(2)

(3)

Mechanical Evaluation – Stress Relaxation. For stress relaxation testing samples were prepared similarly to the preparation technique for dynamic compression. During testing, a preload of 0.01 N was applied to ensure contact between the loading platen and the hydrogel.


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Hydrogels were then subjected to a ramp compression to 10% strain at a strain rate of 10% per s, and were then allowed to relax for 3600 seconds. The stress and strain data were smoothed using a tricube weighting function, described in the Supporting Information (Figure SI.1). This weighting was performed so that data spaced linearly in time were converted to data spaced logarithmically in time. The logarithmic data spacing facilitates fitting data to the Prony series (described below), with time constants that may vary over several orders of magnitude. The viscoelastic stress relaxation behavior of the hydrogels under uniaxial compression was modeled according to a Maxwell-Wiechert model (Figure 1). The Maxwell-Wiechert model contains a Hookean spring in series with n Maxwell elements. Each Maxwell element behaves according to: − = + " #

(4)

Where –σi(t) is the engineering compressive stress of the ith element, and ε=(1−λ) is the engineering compressive strain. The Hookean spring and each of the n Maxwell elements of the model experiences the same strain, ε(t). The total stress, −σ(t), is the sum of the stresses on each element, –σi(t). In stress relaxation the strain is held constant, and the stress-strain relationship is represented by a Prony series for the relaxation parameter E(t): =

−

= + $

% &'

(5)

Where t is the relaxation time (measured from the time at which the stress reaches a maximum during initial compression). The time constants in equation (5) are: ( =

"

(6)


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Figure 1. Maxwell-Wiechert model for viscoelasticity, composed of a Hookean spring in parallel with n Maxwell elements. Smoothed data from stress relaxation experiments were fit to the Prony series with three Maxwell elements (n = 3 in equation 5), and used to calculate a normalized relaxation parameter, E/Emax. The normalized relaxation parameter was fit with a normalized version of equation 5 using the Levenberg-Marquardt non-linear least squares algorithm implemented in Igor Pro 5.0.5.7 and 7.0.0 (Wavemetrics, Inc.), with E0, E1, E2, and all τi as adjustable parameters. The parameter E3 was constrained to E3 = Emax−E0−E1−E2. Cell-laden Hydrogel Fabrication. Adult human adipose-derived mesenchymal stem cells (ADSC) were used for creating cell-laden hydrogels. ADSCs (Zen-Bio ACS-F, Research Triangle Park, NC) at passage 2 were seeded at a density of 5000 cells cm−2 in 75 cm2 T flasks and expanded to passage 4. The cell growth media consisted of Dulbecco’s Modified Eagle Medium (DMEM) with 10% fetal bovine serum, 1% penicillin/streptomycin, and amphotericin B (2 µg mL−1). Cell-containing agarose hydrogels were fabricated to contain ADSCs and either CS or co-CS. CS and co-CS were chosen as additives in this initial study, because we have previously demonstrated that the co-CS proteoglycan mimic can bind, stabilize, and deliver the heparin binding growth factor FGF-2, and because CS is a major GAG found in aggrecan and versican in cartilage.23 To create the hydrogel solution, 0.5 weight % of CS or co-CS was added to a 4.5 weight % agarose solution and stirred for 10 min at 60 °C, similar to the procedure described


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above. The concentration of the polymers was doubled so that after combination with the cells, the final concentration would be the same as those used above for the cell-free hydrogels. Cells were trypsinized to release them from the 75 cm2 T flasks, counted with an automatic cell counter (Scepter 2.0, MilliporeSigma, Billerica, MA), and suspended in cell growth medium at a concentration of 20×106 cells mL−1. Equal volumes of cell suspension and 5% polymer solution were gently mixed at 40 °C and cast into stainless steel molds to create cylindrical constructs 5 mm in diameter and 3 mm thick. Hydrogels were allowed to cool for ~15 min at room temperature and were then transferred to a 48-well plate and bathed in 500 µL of cell growth medium. This procedure produced hydrogels consisting of 2.25% agarose, 0.25% polymer, and cells seeded at a density of 10×106 cells/mL. Each construct contained ~6000 cells. Cells encapsulated in plain 2.25% agarose hydrogels were included as controls. Cell-containing hydrogels were maintained at 37 °C and 5% CO2, and cell growth medium was changed every two days. Cell Viability and Metabolic Activity. A Live/Dead Cytotoxicity kit (Molecular Probes, Eugene, OR) was used to assess the number of live and dead cells present after 1 and 7 days of cell culture. One millimeter thick slices were taken from the cross-sections of cell-containing hydrogels (n = 4 replicates per condition) using stainless steel razor blades. After rinsing the slices in PBS they were stained with 500 µl of PBS solution containing calcein AM (0.25 µl mL−1) and ethidium homodimer (EthD-1, 2 µl mL−1) at 37 °C and 5% CO2. Prior to imaging, each sample was rinsed in PBS for 10 min to remove excess dye. Samples were imaged using a fluorescence microscope (Olympus IX70, Center Valley, PA). Viable cells were characterized by a green fluorescent calcein AM stain. Damaged cells were characterized by a red fluorescent EthD-1 stain. Counts of live and dead cell nuclei were determined from 20×


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magnification images (Particle count tool, ImageJ v1.48, Bethesda, MD) and were used to calculate the percentage of live cells. Cell metabolic activity after 1 and 7 days of culture was determined by a CellTiter-Blue Cell Viability Assay (Promega G808A, Madison, WI), which uses the indicator dye resazurin to measure the metabolic capacity of cells as an indicator of cell viability. Viable cells retain the ability to reduce resazurin into resorufin, which is fluorescent. Non-viable cells with no metabolic capacity, do not reduce the indicator dye, and thus do not generate a fluorescent signal. Thus a higher level of fluorescence is associated with greater cell metabolic activity. According to the manufacturer’s instructions, 20 µL of CellTiter-Blue dye was added to cell-containing hydrogels (n = 4 replicates per condition) for every 100 µL of cell culture media (100 µL dye per sample) and samples were incubated for 4 hr at 37 °C and 5% CO2. Acellular hydrogels were included as controls. Sample fluorescence was read in a microplate reader (Molecular Devices SpectraMax M3, Sunnyvale, CA) at 560 nm excitation and 590 nm emission. Statistics. All mechanical property measures are reported as mean ± standard deviation from n ≥ 4 replicates for each condition. Student’s t-tests were performed to compare polysaccharide and copolymer groups to controls. Cell viability and metabolic activity data are presented as mean ± standard error of the mean. Data analysis was performed using Minitab (Minitab, Inc., State College, PA). Differences corresponding to a p-value of less than 0.05 were considered to be statistically significant.

RESULTS AND DISCUSSION Polysaccharide and Graft Copolymer Size and Zeta Potential Characterization. The effective hydrodynamic diameter of each polysaccharide and each copolymer was determined


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using DLS. Because the size distributions were not all monomodal for the graft copolymers, we report both the number and intensity size distributions in Figure 2. The dextran sulfate copolymer has a significant fraction (31%) in the number distribution potentially representing uncomplexed or poorly complexed polymer, with a hydrodynamic diameter of 14.5 nm. This is close to the hydrodynamic diameter of the original HA and HA-SH, and represents a significant number fraction of the total polymer chains (31%), however it represents a much smaller fraction of the total mass of the sample. All of the graft copolymers have average hydrodynamic diameters between 300 nm and 620 nm (Figure 2), and negative zeta potentials (Table. 2).

Figure 2. Size distributions of the polysaccharides and graft copolymers from dynamic light scattering, showing both the intensity distributions and number distributions, with average hydrodynamic diameters from the number distributions. Table 2. Zeta potential of graft copolymers.


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Copolymer

Zeta potential

Co-Hep

-21 ± 2

Co-Dex

-21 ± 1

Co-DS

-14 ± 1

Co-CS

-30 ± 3

The four polysaccharides used as side chains in the graft copolymers have different molecular weights and charge densities, as indicated in Table 1 above, in the Materials and Methods section. The hyaluronan used for the proteoglycan-mimetic graft copolymer backbone has a large molecular weight. Chondroitin sulfate is a GAG side chain present in many proteoglycans in the cartilage.8,9 Chondroitin sulfate is moderately sized and has a moderate charge density. Heparin is structurally similar to heparan sulfate, and because heparin is commercially available, it is often used in experimental work in place of heparan sulfate. Heparin has a relatively low molecular weight but a higher charge density than chondroitin sulfate. Dextran is a complex branched polyglucan and dextran sulfate is the highly sulfated version of dextran. Dextran and dextran sulfate are similar in size; however, dextran is uncharged while dextran sulfate has a high negative charge density. Although the dextran sulfate has a high negative charge density, it resulted in copolymers with relatively less negative zeta potential (Table 2) and potentially reduced degree of substitution, evidenced by the fraction with a smaller hydrodynamic diameter (Figure 2).

Mechanical Evaluation – Cyclic Compression. For all samples, the stress versus strain curves for cyclic loading experiments were well-described by equation 1, enabling determination of Emax and Emin, according to equations 2 and 3. Examples of the cyclic loading data at all five


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loading rates for a 2.25% agarose hydrogel are shown in Figure 3. Stress versus strain plots for the 2.25% agarose hydrogel, for the 2nd and 99th compression cycle at 5 loading rates are shown in Figure 4. Stress increases with increasing loading rate (Figures 3 and 4). Stress versus strain plots for the 2% and 2.5% agarose hydrogels are compared to the corresponding plots for the 2.25% hydrogel in the Supporting Information (Figure SI.2). The height of the hydrogels did not change significantly after cyclic compression, demonstrating that the testing regime did not permanently alter the dimensions (Supporting Information, Figure SI.3). Values of Emax and Emin increase with increasing loading frequency, up to 2 Hz. At 5 Hz, the stress values are similar to the values at 2 Hz. In all sample types investigated, there is no substantial increase in the stress values at 5 Hz compared to 2 Hz. Some samples even exhibited lower stress at the 5 Hz loading rate. The values of Emax, Emin, and σ33 for all sample compositions at all loading rates are reported in the Supporting Information (Table SI.1).


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Figure 3. Compressive stress versus time for 2.25% agarose hydrogel with the median measured modulus at 5 loading frequencies. The first ten of 100 loading cycles is shown at each frequency for the sample with the mean modulus value.


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Figure 4. Compressive stress versus strain for a 2.25% agarose sample with the median measured modulus at 5 different loading frequencies. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. Black line is the fit of the 2nd loading cycle to equation 1. The polysaccharide (Hep, Dex, DS, CS, and HA) additives modulate the mechanical properties of the hydrogels, in a molecular weight and charge density dependent manner. Adding hyaluronan to the agarose hydrogel increased the stress by more than 100% at the 1 Hz loading rate. Similarly adding chondroitin sulfate and dextran sulfate both increased the stress by more than 10% at the 1 Hz loading rate (Figure 5). Similar increases in stress were seen at the other loading rates for each of these three polysaccharide additives. Data for other loading rates is provided in the Supporting Information.

Figure 5. Compressive stress versus strain for an agarose sample with the median measured modulus at 1 Hz compared to hydrogels with added hyaluronan (+HA), chondroitin sulfate


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(+CS), and dextran sulfate (+DS). All three of these negatively charged additives increase the modulus at all loading rates. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. The black line is a fit to equation 1. The hyaluronan used in this work is a large weak polyanion (740 kDa, with one carboxylic acid group per disaccharide). Its inclusion in the hydrogel increases the Emin and Emax, likely due to the hyaluronan forming entanglements with the agarose, effectively engaging the hydrogel network. The CS and DS polysaccharides are smaller than hyaluronan, and similar in size (84 kDa for CS and 36-50 kDa for DS). Both are strong polyanions containing multiple sulfate groups per disaccharide. They may increase the Emin and Emax of agarose gels by both entangling with the agarose network, by electrostatic repulsion, or by increasing the osmotic pressure of the gel. However, based on the results for dextran (see below), which has no apparent effect on the gel modulus, the electrostatic and osmotic pressure effects likely dominate for these additives. Other research groups have found that the inclusion of chondroitin sulfate in a material allowed for modulus enhancement.29–31 An aggrecan mimic consisting of chondroitin sulfate functionalized with hyaluronan-specific binding peptides was found to increase the compressive strength of cartilage extracellular matrix-based constructs as measured by rheological frequency sweep.29 The inclusion of chondroitin sulfate in a genipin crosslinked collagen hydrogel increased the compressive modulus of the hydrogel.31 The authors hypothesized that this compressive modulus enhancement was due to attraction of water molecules to the negatively charged chondroitin sulfate.31 Alternatively, adding lower molecular weight polymers heparin (14.4 kDa) and uncharged dextran (40 kDa) does not increase the Emin and Emax of the hydrogel. Even though the heparin


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has a high charge density, with about 2.5 sulfate groups per disaccharide, it is apparently unable to engage with the agarose network. In fact, adding heparin decreases the Emin and Emax at all loading rates (Figure 6, top). Previous studies have shown that the inclusion of heparin in alginate and polyvinyl alcohol (PVA) hydrogels had no effect on the compressive modulus, however the decreased modulus observed in the present study may be due to the relative large amount of heparin that was added.32–34 Dextran is an un-charged polysaccharide, and also has no discernable effect on the mechanical properties at all loading rates (Figure 6, bottom). While the dextran used here has a similar molecular weight to the DS, DS increases the gel modulus, while dextran does not. This is perhaps due to charge repulsion among DS chains and the ability of DS to increase osmotic pressure, as described above. Dextran has previously been shown to enhance the modulus of PVA hydrogels, which may be due to the differing architecture of PVA compared to agarose.35 Although dextran and heparin do not increase the gel modulus, when either heparin or dextran is included in the gel in the form of the corresponding graft copolymer, the compressive modulus increases (Figure 6). This is strong evidence that the large size of the graft copolymers influences their ability to modulate the mechanical properties of the agarose hydrogel.


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Figure 6. Compressive stress versus strain for an agarose sample with the median measured modulus at 1 Hz compared to hydrogels with added heparin (+ Hep) and heparin graft copolymer (+ co-Hep) (top), and compared to hydrogels with added dextran (+ Dex) and dextran graft copolymer (+ co-Dex) (bottom). Adding either of the two polysaccharides to the agarose reduces the modulus at all loading rates. However, adding the corresponding graft copolymer increases the modulus. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. The black line is a fit to equation 1. The heparin used here is a much smaller molecular weight polyanion (14.4 kDa), and is relatively stiff, likely making it unable to form entanglements. So the added heparin mass does not effectively engage the hydrogel network to elastically absorb compressive force. The corresponding graft copolymer is very large and may behave as an elastic nanoparticle. Compression of these large polyanionic nanoparticles within the gel results in electrostatic


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repulsive restoring forces which causes an increase in the compressive modulus. CS has a similar negative charge density as heparin but a much higher molecular weight and increases the modulus, possibly suggesting the large size enables it to form entanglements, as descried above. The Emin and Emax of the agarose hydrogels with added polysaccharides increases with increasing molecular weight of the additive (Figure 7, left). While the low molecular weight heparin significantly reduces both Emin and Emax, the higher molecular weight dextran sulfate, chondroitin sulfate and hyaluronan additives significantly increase both Emin and Emax, (Figure 7, left). Furthermore, for polysaccharides of comparable size (dextran and dextran sulfate) we find that the negatively charged dextran sulfate increases the modulus, while the neutral dextran does not. We conclude from this that the charge on the polyanions can increase the modulus of the gel if the polymer is of sufficiently large size. All of the copolymers also significantly increase Emin, and three of the four significantly increase Emax, when added to the agarose gel (Figure 7, right). The co-DS copolymer additive does not significantly increase Emax. This is also the copolymer with a significant number fraction of smaller complexes and a less negative zeta potential (Figure 2 and Table 2). Although the copolymers are much larger than the hyaluronan, adding the copolymers to the hydrogel does not result in the same enhancement of the mechanical properties, as adding hyaluronan alone. The grafted chains on the copolymers cause them to adopt very extended chain conformations in solution (Figure 2), potentially reducing their ability to form entanglements. Furthermore, all of the copolymers have negative zeta potential (perhaps in part due to the hyaluronan backbone). Therefore, we attribute this mechanical property enhancement to the copolymer particles acting as repulsive elastic nanoparticles within the hydrogel network, and not to a relative increase in entanglements. Furthermore, all of the additives were included in


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the agarose gels in an equal mass ratio, and the copolymers have a much larger molar mass than any of the corresponding polysaccharides. Therefore including the same mass of the copolymers means that far fewer individual polymer chains were added in the case of the graft copolymers compared to the corresponding polysaccharide-containing gels.

Figure 7. Emin and Emax calculated according to equations 2 and 3 at the 1 Hz loading rate for the polysaccharides (left) and graft copolymer (right) additives in agarose hydrogels, compared to the agarose hydrogel control. * indicates statistically significant difference compared to the agarose hydrogel control (p < 0.05). Mechanical Evaluation – Stress Relaxation. The stress versus strain data from the stress relaxation experiments were fit to equation 5 (Figure 8). For all experiments, the time constants, (τ1 ~ 100 s) > (τ2 ~ 10 s) > (τ3 ~ 0.1 s) are generally separated by about one order of magnitude,


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confirming that three time constants are necessary to fit the full range of data for each sample. This also suggests that these time constants represent distinct relaxation phenomena. Examples of the agarose, agarose + Hep, and agarose + coHep stress relaxation data and fits are shown in Figure 8.

Figure 8. Smoothed stress relaxation data for an agarose hydrogel, a hydrogel containing heparin (+ Hep), and a hydrogel containing the heparin graft-copolymer (+ co-Hep), shown with the results from the corresponding fits to the normalized version of equation 5. The heparin-containing gels have significantly lower values of the relaxation parameter E0 and the sum of all relaxation parameters (E0, E1, E2, and E3), over the entire stress relaxation experiment (Figure SI.4). Similar to the cyclic loading experiments, when the heparin graft copolymer is added the relaxation parameters are significantly increased compared to the agarose control. In addition to significantly increasing the moduli, Ei, the time constants are also modulated by the additive. The addition of heparin to the hydrogels causes the longest time constant, τ1, to decrease, while the addition of the heparin copolymer causes the middle time constant, τ2, to increase (Figure 9). The higher E0 value in agarose hydrogels suggests that initially water is strongly bound within the agarose hydrogel and is not immediately expelled


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upon application of the load. When heparin is added to the hydrogel the small molecular weight of the polymer may disrupt the agarose network and allow water to flow out more quickly, resulting a lower E0. Additionally the disruption of the agarose network by heparin could then affect the overall relaxation profile, as evidenced by the differing values of the largest time constant as well as the relaxation parameters E1, E2, and E3. When the heparin graft copolymer is added to the agarose hydrogel, the significant increase in all relaxation parameters and the τ2 time constant could suggest that the large size of the polymer combined with the charge density reinforces the agarose network, encouraging fluid to remain trapped within the network. Similar effects can be seen where the addition of CS or HA polysaccharides to the agarose hydrogels increases the sum of all relaxation parameters (Figure SI.4). This again suggests that their large molecular weight may reinforce the hydrogel network. When co-CS is added to the agarose hydrogel it has no apparent effect on the relaxation parameters. This may be because fewer copolymer molecules were added to the hydrogel compared to the polysaccharide molecules alone, since the amount of additive is based on the mass. The copolymer therefore had less of an influence on the relaxation parameters. Additionally, adding any polysaccharide or graft copolymer to the agarose hydrogel significantly alters the initial relaxation response E0. This could suggest that a combination of the molecular weight and charge density of the additive may alter how fluid is able to flow in and out of the hydrogel. A plot of the mean modulus values, E0, E1, E2, and E3, obtained from fits to the normalized version of equation 5 for all compositions is in the Supporting Information (Figure SI.4).


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Figure 9. Time constants obtained from fits to equation 5 for stress relaxation in hydrogel samples. (mean ± s.d., for n = 4). * indicates statistically significant differences compared to the agarose hydrogel. Cell compatibility. CS-containing proteoglycans, such as versican and aggrecan, are important contributors to the health and development of cartilage.6–9,11 In our previous work, we showed that the CS-containing proteoglycan mimics can bind, stabilize, and deliver GAGbinding growth factors.19,20 GAGs can influence growth factor activity by binding to both the growth factor and its cognate receptor on the cell surface. Here, we also showed that they can modulate the mechanical properties of agarose hydrogels. Therefore, we selected the CScontaining copolymers for initial cell compatibility evaluation. The CellTiter Blue metabolic activity assay shows that ADSCs encapsulated in plain agarose hydrogels exhibit no change in metabolic activity from day 1 to day 7 (Figure 10, left). A steady level of metabolic activity suggests that the number of cells may remain fairly consistent over the 7-day period. For ADSCs encapsulated in agarose hydrogels containing either CS or co-CS additives the metabolic activity of the cells significantly decreased after 7 days in culture. This


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decrease in metabolic activity level could indicate lower numbers of live cells or cells with decreased metabolic activity. Calculating the percentage of live cells based on fluorescently stained images (Figure 10) yielded similar results: ADSCs encapsulated in control agarose hydrogels have a consistent number of live cells that did not change over the course of 1 week in culture (Figure 10, right). However when CS or co-CS was added to the hydrogels the percentage of live cells significantly decreased by day 7. However after 7 days of culture some cells did remain alive and metabolically active, suggesting that the presence of the CS and co-CS did not cause complete cytotoxicity. The results suggest that it may be necessary to make modifications to the gels or cell culture media in the future to increase cell viability. Other researchers investigating proteoglycan-mimetic materials have initially struggled with cell compatibility. Hydrogel materials with covalently cross-linked heparin and star-PEG hydrogel in which network characteristics can be gradually varied showed a similar initial low level of cell viability.36 However further modification of the heparin with the gels allowed attachment of RGD peptides which increased cell viability. Binding of growth factors such as FGF-2 and vascular endothelial growth factor (VEGF) further increased the percentage of live cells.17 One possible explanation for the decreased cell viability in the present study is that because chondroitin sulfate is highly sulfated the CS and co-CS present in the agarose hydrogels could be sequestering the calcium and magnesium ions present in the cell culture media that are necessary for cell proliferation. Additionally, the highly charged CS and co-CS could be interfering with cell adhesion and matrix deposition, thus decreasing cell viability. Additionally, in the present study, we also did not take advantage of the ability of CS and co-CS to potentially deliver survival cues and mitogenic growth factors to the cells.


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Figure 10. CellTiter Blue metabolic assay results and percent of live cells based on live/dead fluorescent stained images. * indicates significant difference from Day 1 value. CONCLUSIONS Proteoglycans, particularly those containing chondroitin sulfate have important biochemical and biomechanical functions. In this work we expanded the composition of our previously reported proteoglycan mimics, and we investigated their ability to modulate the mechanical properties of hydrogels. We also investigated the effect of chondroitin sulfate and chondroitin sulfate graft copolymer additives on cells encapsulated within the agarose hydrogels. Additives affected mechanical properties in a molecular weight and charge density dependent manner. Specifically the smaller molecular weight polysaccharides decreased the modulus of the hydrogels while the larger molecular weight polysaccharides and graft copolymers increased the modulus.

Individual

polysaccharide

additives

may increase modulus

by increasing


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entanglements, while graft copolymers may increase the modulus by acting as elastic nanoparticles that absorb compressive loads. Charge density of the polysaccharides is also an important contributor to the mechanical properties. Negatively charged polysaccharides were able to increase the hydrogel modulus, possibly by means of electrostatic repulsion and osmotic pressure, but only when the molecular weight of the polysaccharide is sufficiently high. Cells encapsulated in agarose hydrogels containing chondroitin sulfate and the chondroitin sulfate graft copolymer showed lowered metabolic activity and viability. While these proteoglycan mimics can both deliver growth factors and modulate mechanical properties of hydrogels, further optimization, including combining these with growth factors, will be conducted to develop their use in cartilage tissue engineering. These graft copolymers can be used to study how the structure and composition of proteoglycans and proteoglycan mimics can be used to tune both the biomechanical and biochemical properties of materials.

SUPPORTING INFORMATION Synthesis of HA-SH, hydrazide-modified HA, and copolymers; Description of tricube weighting function; Compressive stress versus strain for cyclic loading of 2 %, 2.25 %, and 2.5 % agarose hydrogels; Gel sample heights before and after cyclic loading; Emin, Emax, and σ33 obtained from cyclic loading of all sample types at all loading rates; stress relaxation results for all sample types. AUTHOR INFORMATION Corresponding Author *Tel: 970-491-0870; E-mail: [email protected]


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Present Addresses ‡Present address: Tufts University, Medford, Massachusetts, United States

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ‡These authors contributed equally.

Funding Sources Funding for this work was provided by the National Science Foundation (DMR 0847641), and by the Walter Scott Jr. College of Engineering at Colorado State University. REFERENCES (1) Schaefer, L.; Schaefer, R. M. Cell Tissue Res. 2010, 339, 237–246. (2) Gasimli, L.; Linhardt, R. J.; Dordick, J. S. Biotechnol. Appl. Biochem. 2012, 59, 65–76. (3) Sophia Fox, a. J.; Bedi, a.; Rodeo, S. a. Sports Health Multidiscip. Approach 2009, 1 (6), 461–468. (4) Cohen, B.; Lai, W. M.; Mow, V. C. J. Biomech. Eng. 1998, 120, 491–496. (5) Costell, M.; Gustafsson, E.; Asz??di, A.; M??rgelin, M.; Bloch, W.; Hunziker, E.; Addicks, K.; Timpl, R.; F??ssler, R. J. Cell Biol. 1999, 147 (5), 1109–1122. (6) Choocheep, K.; Hatano, S.; Takagi, H.; Watanabe, H.; Kimata, K.; Kongtawelert, P.; Watanabe, H. J. Biol. Chem. 2010, 285 (27), 21114–21125. (7) Shepard, J. B. Int. J. Biol. Sci. 2007, 380–384. (8) Handley, C. J.; Lowther, D. A.; McQuillan, D. J. Cell Biology International Reports. 1985, pp 753–782. (9) Hardingham, T.; Bayliss, M. Semin. Arthritis Rheum. 1990, 20 (3), 12–33. (10) Pearle, A. D.; Warren, R. F.; Rodeo, S. A. Clin. Sports Med. 2005, 24 (1), 1–12. (11) Lark, M. W.; Bayne, E. K.; Flanagan, J.; Harper, C. F.; Hoerrner, L. A.; Hutchinson, N. I.; Singer, I. I.; Donatelli, S. A.; Weidner, J. R.; Williams, H. R.; Mumford, R. A.; Lohmander, L. S. J. Clin. Invest. 1997, 100 (1), 93–106. (12) Franz, T.; Hasler, E. M.; Hagg, R.; Weiler, C.; Jakob, R. P.; Mainil-Varlet, P. Osteoarthritis Cartilage 2001, 9 (6), 582–592. (13) Weyers, A.; Linhardt, R. J. FEBS J. 2013, 280, 2511–2522. (14) Lee, S.-G.; Brown, J. M.; Rogers, C. J.; Matson, J. B.; Krishnamurthy, C.; Rawat, M.; Hsieh-Wilson, L. C. Chem. Sci. 2010, 1, 322–325. (15) Merrett, K.; Liu, W.; Mitra, D.; Camm, K. D.; McLaughlin, C. R.; Liu, Y.; Watsky, M. a.; Li, F.; Griffith, M.; Fogg, D. E. Biomaterials 2009, 30 (29), 5403–5408. (16) Sharma, S.; Panitch, A.; Neu, C. P. Acta Biomater. 2013, 9 (1), 4618–4625. (17) Nie, T.; Baldwin, A.; Yamaguchi, N.; Kiick, K. L. J. Controlled Release 2007, 122, 287– 296.


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Yamaguchi, N.; Kiick, K. L. Biomacromolecules 2005, 6, 1921–1930. Place, L. W.; Kelly, S. M.; Kipper, M. J. Biomacromolecules 2014, 15 (10), 3772–3780. Place, L. W.; Sekyi, M.; Kipper, M. J. Biomacromolecules 2014, 15 (2), 680–689. Volpato, F. Z.; Almodovar, J.; Erickson, K.; Popat, K. C.; Migliaresi, C.; Kipper, M. J. Acta Biomater. 2012, 8 (4), 1551–1559. Takada, T.; Katagiri, T.; Ifuku, M.; Morimura, N.; Kobayashi, M.; Hasegawa, K.; Ogamo, A.; Kamijo, R. J. Biol. Chem. 2003, 278 (44), 43229–43235. Kipper, M.; Place, L. In Macro-Glycoligands; Sun, X.-L., Ed.; Methods in Molecular Biology; Springer New York, 2016; pp 69–86. Pomin, V. H. Anal. Chem. 2013, 86 (1), 65–94. Damodaran, V. B.; Joslin, J.; Reynolds, M. M. J. Mater. Chem. 2012, 22, 5990. Sisu, I.; Udrescu, V.; Flangea, C.; Tudor, S.; Dinca, N.; Rusnac, L.; Zamfir, A.; Sisu, E. Open Chem. 2009, 7 (1). Dalpathado, D. S.; Jiang, H.; Kater, M. A.; Desaire, H. Anal. Bioanal. Chem. 2005, 381 (6), 1130–1137. Park, S.; Hung, C. T.; Ateshian, G. a. Osteoarthritis Cartilage 2004, 12, 65–73. Bernhard, J. C.; Panitch, A. Acta Biomater. 2012, 8, 1543–1550. Strehin, I.; Nahas, Z.; Arora, K.; Nguyen, T.; Elisseeff, J. Biomaterials 2010, 31 (10), 2788–2797. Zhang, L.; Li, K.; Xiao, W.; Zheng, L.; Xiao, Y.; Fan, H.; Zhang, X. Carbohydr. Polym. 2011, 84 (1), 118–125. Jeon, O.; Powell, C.; Solorio, L. D.; Krebs, M. D.; Alsberg, E. J. Controlled Release 2011, 154 (3), 258–266. He, C.; Cheng, C.; Ji, H.-F.; Shi, Z.-Q.; Ma, L.; Zhou, M.; Zhao, C.-S. Polym Chem 2015, 6, 7893–7901. Kim, M.; Lee, J. Y.; Jones, C. N.; Revzin, A.; Tae, G. Biomaterials 2010, 31 (13), 3596– 3603. Cascone, M. G.; Maltinti, S.; Barbani, N.; Laus, M. J. Mater. Sci. Mater. Med. 1999, 10, 431–435. Freudenberg, U.; Hermann, A.; Welzel, P. B.; Stirl, K.; Schwarz, S. C.; Grimmer, M.; Zieris, A.; Panyanuwat, W.; Zschoche, S.; Meinhold, D.; Storch, A.; Werner, C. Biomaterials 2009, 30 (28), 5049–5060.


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Figure 1. Maxwell-Wiechert model for viscoelasticity, composed of a Hookean spring in parallel with n Maxwell elements. 39x25mm (600 x 600 DPI)


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Figure 2. Size distributions of the polysaccharides and graft copolymers from dynamic light scattering, showing both the intensity distributions and number distributions, with average hydrodynamic diameters from the number distributions. 106x79mm (300 x 300 DPI)


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Figure 3. Compressive stress versus time for 2.25% agarose hydrogel with the median measured modulus at 5 loading frequencies. The first ten of 100 loading cycles is shown at each frequency for the sample with the mean modulus value. 140x159mm (300 x 300 DPI)


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Figure 4. Compressive stress versus strain for a 2.25% agarose sample with the median measured modulus at 5 different loading frequencies. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. Black line is the fit of the 2nd loading cycle to equation 1. 48x13mm (300 x 300 DPI)


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Figure 5. Compressive stress versus strain for an agarose sample with the median measured modulus at 1 Hz compared to hydrogels with added hyaluronan (+HA), chondroitin sulfate (+CS), and dextran sulfate (+DS). All three of these polyanions increase the modulus at all loading rates. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. The black line is a fit to equation 1. 48x13mm (300 x 300 DPI)


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Figure 6. Compressive stress versus strain for an agarose sample with the median measured modulus at 1 Hz compared to hydrogels with added heparin (+ Hep) and heparin graft copolymer (+ co-Hep) (top), and compared to hydrogels with added dextran (+ Dex) and dextran graft copolymer (+ co-Dex) (bottom). Adding either of the two polysaccharides to the agarose reduces the modulus at all loading rates. However, adding the corresponding graft copolymer increases the modulus. The 2nd (red squares) and 99th (blue circles) loading cycles are shown with filled symbols for the loading and open symbols for the unloading. The black line is a fit to equation 1. 96x67mm (300 x 300 DPI)


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Figure 7. Emin and Emax calculated according to equations 2 and 3 at the 1 Hz loading rate for the polysaccharides (left) and graft copolymer (right) additives in agarose hydrogels, compared to the agarose hydrogel control. * indicates statistically significant difference compared to the agarose hydrogel control (p < 0.05). 110x83mm (300 x 300 DPI)


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Figure 8. Smoothed stress relaxation data for an agarose hydrogel, a hydrogel containing heparin (+ Hep), and a hydrogel containing the heparin graft-copolymer (+ co-Hep), shown with the results from the corresponding fits to the normalized version of equation 5. 67x54mm (300 x 300 DPI)


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Figure 9. Time constants obtained from fits to equation 5 for stress relaxation in hydrogel samples. (mean ± s.d., for n = 4). * indicates statistically significant differences compared to the agarose hydrogel. 73x59mm (300 x 300 DPI)


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Figure 10. CellTiter Blue metabolic assay results and percent of live cells based on live/dead fluorescent stained images. * indicates significant difference from Day 1 value. 100x87mm (300 x 300 DPI)


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Scheme 1. Formation of graft copolymer by reductive amination to couple the reducing end of a polysaccharide to a hydrazide-modified hyaluronan. Thiolation and hydrazide modification of the hyaluronan are detailed in the Supporting Information (Schemes SI.1, and SI.2). 85x42mm (300 x 300 DPI)


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Mechanical Properties and Cell Compatibility of Agarose Hydrogels

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