A Thermosensitive Hydrogel Capable of Releasing bFGF for

May 11, 2012 - We hypothesized that enhancing MSC survival under low nutrient and low oxygen conditions would restore the differentiation. To enhance ...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/Biomac

A Thermosensitive Hydrogel Capable of Releasing bFGF for Enhanced Differentiation of Mesenchymal Stem Cell into Cardiomyocyte-like Cells under Ischemic Conditions Zhenqing Li, Xiaolei Guo, and Jianjun Guan* Department of Materials Science & Engineering, The Ohio State University, Columbus, Ohio 43210, United States ABSTRACT: A thermosensitive hydrogel capable of differentiating mesenchymal stem cells (MSCs) into cardiomyocytelike cells was synthesized. The hydrogel was based on Nisopropylacrylamide (NIPAAm), N-acryloxysuccinimide, acrylic acid, and hydroxyethyl methacrylate-poly(trimethylene carbonate). The hydrogel was highly flexible at body temperature with breaking strain >1000% and Young’s modulus 45 kPa. When MSCs were encapsulated in the hydrogel and cultured under normal culture conditions (10% FBS and 21% O2), the cells differentiated into cardiomyocytelike cells. However, the differentiation was retarded, and even diminished, under low nutrient and low oxygen conditions, which are typical of the infarcted heart. We hypothesized that enhancing MSC survival under low nutrient and low oxygen conditions would restore the differentiation. To enhance cell survival, a pro-survival growth factor (bFGF) was loaded in the hydrogel. bFGF was able to sustainedly release from the hydrogel for 21 days. Under the low nutrient and low oxygen conditions (1% O2 and 1% FBS), bFGF enhanced MSC survival and differentiation in the hydrogel. After 14 days of culture, survival of 70.5% of MSCs remained in the bFGF-loaded hydrogel, while only 4.9% of MSCs remained in the hydrogel without bFGF. The differentiation toward cardiomyocyte-like cells was completely inhibited at 1% FBS and 1% oxygen. Loading bFGF in the hydrogel restored the differentiation, as confirmed by the expression of cardiac markers at both the gene (MEF2C and CACNA1c) and protein (cTnI and connexin 43) levels. bFGF loading also up-regulated the paracrine effect of MSCs. VEGF expression was significantly increased in the bFGF-loaded hydrogel. These results demonstrate that the developed bFGF-loaded hydrogel may potentially be used to deliver MSCs into hearts for regeneration of heart tissue. factor and stem cells,17 genetic modification of stem cells with pro-survival genes,18−20 and pretreating stem cells with growth factor21 or hypoxic conditions22 have been utilized to increase the survival of injected stem cells. Among these, genetic modification has potential safety issues. Pretreatment of cells with growth factor or hypoxic conditions can improve shortterm cell survival, but has failed to show continuous effect in the long term.22 Co-delivery of growth factor and stem cells in a matrix may be a suitable approach to support stem cell survival before angiogenesis is established, as sustainably released growth factor continuously supports cell survival. Injectable hydrogels are good matrices for growth factor and stem cell delivery. Upon injection, the viscous hydrogel solution can better retain growth factor and cells in the heart than nonviscous saline.23 In addition, hydrogel properties can be tailored to control growth factor release kinetics, by modulating hydrogel composition and degradation,24 or by introducing protein affinitive molecules such as aptamers and peptides.25−27 Cardiac differentiation of delivered stem cells is necessary for regeneration of new heart tissue. Direct injection of MSCs into

1. INTRODUCTION Stem cell therapy has been applied to treat myocardial infarction (MI), one of the major cardiovascular diseases affecting millions of people in the Western world.1 The ultimate goal of stem cell therapy is to improve cardiac function that has deteriorated due to the loss of heart muscle caused by MI. Many types of stem cells have been used for cardiac stem cell therapy, such as hematopoietic stem cells,2 cardiosphere derived cells (CDCs),3 cardiac stem cells,4−6 and mesenchymal stem cells (MSCs).7 Among these, MSCs have been widely investigated in animal studies and clinical trials.8−10 Injection of MSCs into hearts has been demonstrated to improve heart function.11 However, current stem cell therapy has low efficacy, due to the extremely low survival rate and inferior cardiac differentiation.12,13 Various studies have shown that less than 2% of injected cells survived after two weeks,11,14,15 and few of the surviving cells differentiated into cardiac cells for regeneration.16 Such a low efficacy significantly limits the application of MSCs in clinics. The poor cell survival is the result of the harsh ischemic environment of the infarcted heart. Angiogenesis is an ultimate solution to address this issue. However, before angiogenesis can be established, a temporary method is needed to support cell survival. Approaches such as co-delivery of pro-survival growth © 2012 American Chemical Society

Received: April 13, 2012 Revised: May 10, 2012 Published: May 11, 2012 1956

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

Figure 1. Scheme for synthesis of the thermosensitive and injectable hydrogel.

the heart often yields an extremely low cardiac differentiation. One of the possible causes is that scar tissue inhibits the differentiation, because its stiffness is much higher than that of healthy heart tissue.28 Various studies have demonstrated that the differentiation of MSCs is sensitive to matrix stiffness.29,30 To address the cardiac differentiation issue, strategies including predifferentiation of MSCs into cardiomyocytes in vitro before injection,31,32 and co-delivery of differentiation stimulator and MSCs,33 have been used. In vitro MSC differentiation can be achieved by co-culturing with mature cardiomyocytes,31 or demethylization with 5-azacytidine (5-aza).32 The major drawbacks of this strategy lie in the low differentiation rate (30−40%) and in the fact that it is time-consuming.34 Codelivery of differentiation stimulator and MSCs for in vivo differentiation is more convenient, but the cardiac differentiation efficacy is highly dependent on the stimulator dosage, release kinetics, and bioactivity.33 On the basis of the above, new approaches to cardiac differentiation of MSCs are necessary. Recent progresses in tissue engineering and stem cell biology have shown that stem cells differentiate into specific lineages when cultured on/in hydrogels with moduli matching the corresponding tissues.28,29,35,36 This matrix stiffnessinduced stem cell differentiation has great clinical significance for heart regeneration, as the matrix and cells may be directly transplanted into the heart for regeneration, without the need for predifferentiation of stem cells into cardiac lineage in vitro. Kraehenbuehl et al. studied the cardiac differentiation of embryonic stem cells in cross-linked PEG gels with various moduli and found that the optimal modulus is less than 1 kPa.36 We have demonstrated that 31 kPa is optimal for CDCs to differentiate into cardiac cells in a hydrogel based on polycaprolactone, N-isopropylacrylamide, 2-hydroxyethyl methacrylate, and dimethyl-γ-butyrolactone acrylate.37 While these studies have demonstrated that stem cells can be directed to differentiate into cardiac cells, the differentiation was achieved under standard culture conditions. It is unknown whether the differentiation can be achieved under the low nutrient and low oxygen conditions that are typical of the infarcted heart.

The objectives of this work were to develop a hydrogel-based matrix capable of stimulating MSCs differentiating into cardiomyocyte-like cells, to investigate whether MSCs can differentiate into these cells under low nutrient and low oxygen conditions, and to explore whether enhancing cell survival under these conditions would restore the differentiation. We have synthesized a poly(N-isopropylacrylamide)-based thermosensitive hydrogel capable of differentiating MSCs into cardiomyocyte-like cells under standard culture conditions. However, the differentiation was retarded under low nutrient and low oxygen conditions. A pro-survival growth factor (bFGF) was loaded in the hydrogel aiming to restore the differentiation by enhancing MSC survival under these conditions. bFGF can be sustainedly released from the hydrogel over a 21-day period. The differentiation was reestablished with the presence of bFGF in the hydrogel, as confirmed by the expression of cardiac markers at both the gene and protein levels.

2. MATERIALS AND METHODS 2.1. Materials. N-Isopropylacrylamide (NIPAAm, TCI) was purified with hexane three times before use. Acrylic acid (AAc, VWR) and hydroxyethyl methacrylate (HEMA, VWR) were purified by passing through an inhibitor-remover column to remove the inhibitor. Acyloxy chloride (Sigma), N-hydroxysuccinicimide (NHS, Sigma), trimethylene carbonate (TMC, Boehringer Ingelheim), Type I collagen (Kensy Nash), human basic fibroblast growth factor (bFGF, Peprotech), and heparin (Fisher Scientific) were used as received. 2.2. Hydrogel Polymer Synthesis. The hydrogel polymer was based on NIPAAm, N-acryloxysuccinimide (NAS), AAc, and hydroxyethyl methacrylate-poly(trimethylene carbonate) (HEMAPTMC) (Figure 1). NAS was synthesized by the reaction of acyloxy chloride with NHS using methylene chloride as a solvent.38 HEMAPTMC was synthesized by ring-opening polymerization of TMC initiated by HEMA.17,39 The molar feed ratio of the TMC/HEMA was 2. The polymerization was conducted at 110 °C using stannous octoate as a catalyst. The obtained HEMA-PTMC had an average 2.1 TMC units calculated from the 1H NMR spectrum. The hydrogel polymer was synthesized by free radical polymerization.17,40 In brief, a 250 mL round flask was charged with tetrahydrofuran (THF), NIPAAm, AAc, NAS, and HEMA-PTMC. The molar ratio of 1957

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

Table 1. Primers Used for Quantification of Early Cardiac Markers name

forward (5′−3′)

reverse (5′−3′)

Tm (fwd/rev °C)

product size (bp)

CACNA1c VEGFA MEF2C β-actin

CAGAAACTACAGGAGAAGAGG CAGAATCATCACGAAGTGGT GATCATCTTCAACAGCACC AAGATCAAGATCATTGCTCCTC

AAGAAGAGGATCAGGTTGGT CATTCACATTTGTTGTGCTG GTTCAATGCCTCCACGA GGACTCATCGTACTCCTG

59.5/60.5 61.1/59.5 59.5/60.5 61.2/59.5

207 286 132 110

were used for encapsulation in hydrogels. Our previous reports demonstrated that these MSCs remained multipotent.17 2.7. Encapsulation of MSCs into Hydrogels. MSCs were trypsinized and resuspended in the culture medium at a density of 32 million/mL. Before encapsulation, hydrogel solution with or without bFGF loading was sterilized under UV light in a laminar flow hood for 30 min. A total of 0.25 mL of cell suspension was added to 1 mL of hydrogel solution. After thoroughly mixing, the mixture was incubated at 37 °C for 30 min to form solid hydrogel. The supernatant was replaced with a low serum medium (DPBS supplemented with 1% FBS and 1% penicillin/streptomycin). The hydrogels were then cultured in hypoxia incubators with different oxygen levels (20%, 5%, and 1%) for 14 days. The CO2 level was maintained at 5%. The culture medium was changed at days 1, 3, and 7. 2.8. MSCs Survival in Hydrogels. The survival of MSCs in hydrogel with and without bFGF was assessed by double-strand DNA (dsDNA, for live cell) content.48 In brief, hydrogels were taken out at days 1, 7, and 14. The hydrogels were then digested by papain solution at 60 °C for 24 h. The dsDNA concentration in the digested solution was measured by PicoGreen assay (Invitrogen), and normalized to the wet weight of the hydrogel. 2.9. The Differentiation of MSCs to Cardiomyocyte-like Cells in Hydrogels. MSC differentiation into cardiomyocyte-like cells was characterized at the gene level by real-time RT-PCR and protein level by immunohistochemistry. For real-time RT-PCR analysis, after 1, 7, and 14 days of culture, hydrogels were homogenized in TRIzol to isolate total RNA. The quantity and quality of total RNA were assessed by Nanodrop (Thermo). Approximately 1 μg of total RNA was used for cDNA synthesis using a High Capacity cDNA Reverse Transcription kit (ABI). Primers of forward and reverse pairs of MEF2C, CACNA1c, VEGFA, and β-actin were designed using PerlPrimer software by spanning the intron/exon boundary. The sequences, melting temperatures, and expected product sizes are listed in Table 1. Real-time RT-PCR was performed in triplicate for each sample with Maxima SYBR Green/fluorescein master mix on an Applied Biosystem 7900 system. β-Actin was used as the housekeeping gene. Fold differences were calculated using standard ΔΔCt method. For immunohistochemistry, the hydrogel constructs were fixed with 4% formaldehyde solution after 14 days of culture. The constructs were then cryo-sectioned at 10 μm thickness. Sections were permeabilized with 0.1% Triton X-100 and blocked by 10% goat serum. Sections were then incubated, first with primary antibodies anti-cardiac troponin I (cTnI, Abcam) and anti-mouse connexin 43 overnight at 37 °C, followed by secondary antibody Dylight488 goatanti mouse IgG for 1 h at 37 °C. Hoechst 33328 was used to counterstain nuclei. All images were taken with a confocal microscope (Olympus FV1000). The ratio of cells expressing cardiac markers was calculated by dividing the number of cells expressing markers by the total number of cells. 2.10. Statistical Analysis. All data were reported as mean ± standard deviation. One-way ANOVA with posthoc Tukey−Kramer test was used in analysis of MSC survival in hydrogels with or without bFGF and gene expression levels.

NIPAAm/AAc/NAS/HEMA-PTMC was controlled at 85/6/5/4. Initiator benzoyl peroxide (BPO) was then added. The reaction was conducted at 70 °C overnight with stirring. The polymer solution was precipitated in hexane. The polymer was purified twice with THF/ ethyl ether. 2.3. Hydrogel Characterization. The composition of the polymer was determined from 1H NMR spectrum. The spectrum was obtained on a 300 MHz spectrometer using CDCl3 as the solvent. The molecular weight of the polymer was measured by gel permeation chromatography (GPC). THF was used as the solvent. The thermal transition temperature of the hydrogel solution (20%) was tested using differential scanning calorimetry (DSC) over a temperature range of 0−60 °C with a heating rate of 10 °C/min. The injectability of the hydrogel solution was tested by injecting the solution through a 26gauge needle.41,42 The hydrogel solution gelation time was determined using an Olympus IX71 microscope at 37 °C.40 The time needed for a drop (20 μL) of hydrogel solution to become full opaque (i.e., transmittance equal to zero) is referred to as the gelation time.37,40 The tensile properties of the hydrogel were tested at 37 °C using an Instron tensile tester. A cross-head speed of 50 mm/min was used. 2.4. Loading bFGF into Hydrogels. Hydrogel solution (20 wt %) was prepared by dissolving the synthesized hydrogel polymer in Dubucceo’s modified phosphate buffer saline (DPBS, pH 7.4). To improve hydrogel biocompatibility, Type 1 collagen was added to the hydrogel solution at a final concentration of 6% of polymer weight. After mixing completely, the solution was set at 4 °C overnight. To load bFGF into the hydrogel, heparin was added to the hydrogel solution first at a concentration of 1 mg/mL. The function of the heparin was to bind to bFGF and preserve its bioactivity.43 The hydrogel solution was then mixed with bFGF solution to reach a final bFGF concentration of 10 or 50 μg/mL. The bFGF-loaded hydrogel solution was then incubated at 37 °C for 30 min to achieve gelation. The supernatant was collected and the amount of bFGF was measured using a bFGF ELISA kit (Peprotech).44 2.5. bFGF Release Kinetics and Bioactivity of Released bFGF. To measure the release kinetics of the bFGF, the hydrogel sample (∼40 mg) was placed in a 2 mL microcentrifuge tube, and 200 μL of release medium DPBS was added. The sample was incubated at 37 °C. At predetermined time points, the release medium was collected. bFGF concentration in the release medium was measured using a bFGF ELISA kit. Bioactivity of the released bFGF was assessed using rat smooth muscle cell (SMC) proliferation assay.45 The cells were cultured in a T-175 flask supplemented with a culture medium containing Dulbecco’s Modified Eagle Medium (DMEM) and 10% fetal bovine serum (FBS). After digestion, SMCs were seeded to a 96-well plate at a density of 2 × 105 cells/mL. After 24 h, the culture medium was removed and replaced with the collected release medium supplemented with 0.5% FBS. After 48 h of incubation, cell viability was measured by MTT assay.46 Release media collected from the hydrogel without bFGF, and 1 ng/mL bFGF, were used as controls. Relative cell viability was determined by normalizing MTT absorbance of the release media to that of the 1 ng/mL bFGF. 2.6. Human Mesenchymal Stem Cells Culture. Human bone marrow mesenchymal stem cells (MSCs) were provided by Dr. Prockop at the Texas A&M University Health Science Center. The cells were positive to CD105, CD73, and CD90, while negative to CD45, CD34, and CD14.47 Cells were cultured in T-175 flasks using alpha modified minimum essential medium (αMEM) supplemented with 20% FBS, 2% L-glutamine, and 1% penicillin/streptomycin. Cells were expanded at 90% confluence. MSCs between passages 8 and 10

3. RESULTS 3.1. Hydrogel Properties. The synthesized hydrogel polymer had a NIPAAm/NAS/AAc/HEMAPTMC ratio of 85/5.5/5.2/4.3. The polymer had a number average molecular weight of 13.7 kDa and a polydispersity index of 1.3. The hydrogel solution (20 wt %) was injectable. It can be easily 1958

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

containing bFGF had significantly higher viability than those cultured in 1 ng/mL bFGF and release medium from hydrogel without bFGF loading. This result demonstrates that the released bFGF had a greater stimulatory effect than 1 ng/mL bFGF. 3.4. Effect of Nutrient and Oxygen Level on MSC Survival. The MSC encapsulated hydrogels were cultured under different FBS content and oxygen levels to investigate how these conditions affect MSC survival. We first investigated how nutrient level affects MSC survival. Under 10% FBS and 21% oxygen conditions, MSCs showed a remarkable proliferation during a 14-day culture period. Adding bFGF to the hydrogel slightly increased MSC proliferation (Figure 4). When

injected through a 26-gauge needle. The hydrogel solution possessed a thermal transition temperature of 23.8 ± 0.6 °C, and can rapidly gel (within 7 s) at 37 °C. The formed solid gel was highly flexible with a Young’s modulus of 45 ± 3 kPa and a breaking strain greater than 1000% (beyond the testing limit). When incubated in the DPBS (pH 7.4), the hydrogel had a weight loss of 8 ± 2% over a 14-day period. The completely degraded hydrogel (obtained by hydrolysis in NaOH solution) was soluble at 37 °C, since its thermal transition temperature was elevated to 49.4 °C, higher than 37 °C. 3.2. Release Kinetics of bFGF. The encapsulation efficiency of bFGF was 95 ± 1% and 94 ± 2% for concentrations of 10 and 50 μg/mL, respectively. The bFGF was able to sustainably release from the hydrogel during a 21day release period. An initial burst release occurred in the first 8 h, followed by a slower and sustained release until day 21 (Figure 2). Interestingly, linear release kinetics was observed for

Figure 4. MSC survival in hydrogels cultured under different conditions. Culture conditions: 10% FBS and 21% oxygen, 1% FBS and 21% oxygen, 1% FBS and 5% oxygen, and 1% FBS and 1% oxygen. dsDNA content was used to quantify cell number in the hydrogels. The dsDNA content at day 1 was used for normalization.

Figure 2. Release kinetics of bFGF loaded in the hydrogels. bFGF loading was 10 and 50 μg/mL, respectively. The error bars are small.

both concentrations after day 3. The release kinetics was dose dependent. The concentration of released bFGF was much higher in the hydrogel loaded with 50 μg/mL bFGF than in the hydrogel loaded with 10 μg/mL bFGF. 3.3. Bioactivity of the Released bFGF. The bioactivity of released bFGF was evaluated in terms of its stimulatory effect on SMC growth. A total of 1 ng/mL bFGF was used as a control, as this concentration significantly stimulated SMC growth (Figure 3). Figure 3 shows that all of the released bFGF remained bioactive. SMCs cultured in the release medium

the FBS content was decreased to 1% and the oxygen level was kept at 21%, significant cell death was observed for hydrogel without bFGF, with 68.8% and 8.5% survival after 7 and 14 days of culture, respectively. However, all of the MSCs survived in the hydrogel with bFGF after 7 days of culture. Interestingly, MSCs even proliferated after 7 days (Figure 4). These results demonstrate that, while low nutrient conditions caused significant cell death, bFGF addition overcame the effect. We further investigated how oxygen level affects MSC survival under the low nutrient condition (1% FBS). The decrease in oxygen level from 21% to 5% did not significantly affect MSC survival in the hydrogels without bFGF loading, although significant cell death was observed. Similarly, decreasing the oxygen level from 21% to 5% did not affect MSC survival in the hydrogels loaded with bFGF. At 5% oxygen level, there was no cell death in the first seven days, and cell number increased slightly after 14 days. When the oxygen level was further decreased to 1%, an oxygen level comparable to that in the infarcted heart, massive cell death was observed in the hydrogel without bFGF. Only 4.9% of cells survived after 14 days. bFGF loading significantly enhanced cell survival, with 70.5% of cells surviving after 14 days. These results demonstrate that bFGF largely improved MSC survival under low nutrient and low oxygen conditions, typical of the infarcted heart.

Figure 3. Bioactivity of released bFGF. Release media from hydrogel without bFGF, and 1 ng/mL bFGF were used as controls. Relative cell viability was determined by normalizing MTT absorbance of the release media to that of the 1 ng/mL bFGF. 1959

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

3.5. MSC Differentiation to Cardiomyocyte-like Cells in Hydrogel. The differentiation of MSCs to cardiomyocytelike cells in hydrogel was evaluated at the mRNA level by realtime RT-PCR, and at the protein level by immunohistochemistry. To characterize the differentiation at the mRNA level, the cardiac markers MEF2C and CACNA1c were used. At 10% FBS and 21% O2 culture conditions, MEF2C and CACNA1c expressions in the hydrogel were up-regulated more than 330 and 308 times, respectively, compared to those on the tissue culture plate. This demonstrates that MSCs differentiated into cardiomyocyte-like cells in the hydrogel (Figure 5). Adding

low nutrient condition (1% FBS), a decrease in oxygen level from 21% to 5% and 1% completely inhibited the expression of MEF2C, and remarkably decreased the expression of CACNA1c. However, adding bFGF into hydrogels significantly improved the expression. These results demonstrate that adding bFGF into hydrogel reverted MSC differentiation to cardiomyocyte-like cells in the hydrogel under low oxygen and low serum conditions. The differentiation of MSCs to cardiomyocyte-like cells in hydrogel was further assessed by the expression of cardiac protein cTnI (Figure 6). Under 10% FBS and 21% oxygen conditions, 93.2% of MSCs were cTnI positive. Adding bFGF into hydrogel increased the ratio to nearly 100%. Under 1% FBS and 21% oxygen conditions, the ratio of cells positive to cTnI was 92.9%, but the incorporation of bFGF into the hydrogel increased the ratio to 93.9%. Under 1% FBS condition, the decrease in O2 levels to 5% and 1% remarkably decreased cTnI expression. Only 63.2% of cells were positive to cTnI at the 5% oxygen level, and none of the cells were positive to cTnI at the 1% oxygen level. However, adding bFGF into the hydrogels allowed for 94.8% and 30.5% of the cells to be positive to cTnI at 5% and 1% oxygen levels, respectively. Under 10% FBS and 21% oxygen conditions, the differentiated MSCs also developed cell−cell communication as confirmed by the expression of cell−cell gap junction protein connexin 43 (Figure 7). At 21% oxygen level, a decrease in FBS to 1% did not substantially decrease the expression. At 1% FBS condition, a decrease in the oxygen level to 5% dramatically decreased the expression. When the oxygen level was further decreased to 1%, the expression was completely silenced. However, adding bFGF restored the expression at both oxygen levels. 3.6. Effect of bFGF on VEGF Expression of MSCs. Under low nutrient and oxygen conditions, MSCs show paracrine effect aimed at augmenting cell survival by upregulating the expression of angiogenic growth factors like VEGF.49 We investigated how oxygen and nutrient levels affected the VEGF expression, and whether bFGF loading in the hydrogels could significantly enhance the VEGF expression. Under 10% FBS and 21% O2, bFGF addition significantly increased VEGF expression. When the FBS content was decreased to 1% while keeping O2 at 21%, the VEGF expression did not significantly change in the hydrogels with or without bFGF loading, although the expression was significantly higher in the bFGF loaded hydrogel (Figure 8). When the FBS content was kept at 1%, the decrease in oxygen level from 21% to 5% did not change the VEGF expression. However, when the oxygen level was further decreased to 1%, a remarkably decrease in VEGF expression was observed. At this condition, bFGF loading did not stimulate VEGF expression. These results demonstrate that bFGF can stimulate paracrine effect of MSC at a low nutrient level (1% FBS) and relatively high oxygen level (greater than 5%), but not under both low nutrient (1% FBS) and low oxygen (1%) conditions that are typical of the infarcted heart.

Figure 5. Differentiation of MSCs in hydrogels at mRNA level. (A) MEF2C expression; (B) CACNA1c expression. Cells were cultured under different conditions: 10% FBS and 21% oxygen, 1% FBS and 21% oxygen, 1% FBS and 5% oxygen, and 1% FBS and 1% oxygen. The MEF2C and CACNA1c expressions in hydrogels were normalized by those on the tissue culture plate (plate).

bFGF to the hydrogel further up-regulated the expression of MEF2C and CACNA1c (p < 0.05). When the FBS content was decreased from 10% to 1% at 21% oxygen level, the expression of MEF2C and CACNA1c in the hydrogel was remarkably decreased, yet was still more than 150 and 22 times that on the tissue culture plate, respectively. This indicates that low nutrient level decreased but still allowed MSC differentiation into cardiomyocyte-like cells. Under this culture condition, incorporation of bFGF in the hydrogel significantly increased the expression of MEF2C and CACNA1c (p < 0.05). Under

4. DISCUSSION Stem cell therapy for MI has shown promising results in animal studies and clinical trials. However, it remains impractical for widespread clinical application, due to extremely low cell survival and cardiac differentiation in infarcted hearts. Various reports have demonstrated that less than 2% of transplanted cells survived after two weeks, and few of them differentiated 1960

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

Figure 6. Immunohistological staining of cTnI for MSCs in hydrogels with or without bFGF. Cells were cultured under different conditions: 10% FBS and 21% oxygen, 1% FBS and 21% oxygen, 1% FBS and 5% oxygen, and 1% FBS and 1% oxygen. Scale bar 25 μm.

into cardiac cells for regeneration of new heart tissue.11,14,15,50 The low nutrient and oxygen environment in the infarcted heart is responsible for low cell survival, and stiffness of the scar tissue is the cause of low cardiac differentiation.51,52 To address these issues, it is necessary to deliver an external microenvironment into the heart that not only supports cell survival under low nutrient and low oxygen conditions, but also stimulates stem cell differentiation into cardiac cells. In this report, we have shown that the developed PNIPAAm-based hydrogel is capable of differentiating MSCs into cardiomyocyte-like cells at high efficiency when the hydrogel matched the stiffness of the heart muscle (45 kPa) and was cultured under 10% FBS and 21% O2 conditions. However, we found that cell differentiation was decreased, even diminished, when cultured under lower FBS and low oxygen conditions (Figures 5−7), indicating that cells will not differentiate into cardiomyocyte-like cells under the low nutrient and low oxygen conditions of infarcted hearts. We hypothesized that improving cell survival under low nutrient and low oxygen conditions will restore the differentiation. In this report, we incorporated a pro-survival growth factor, bFGF, into the hydrogel, and found that it enhanced cell survival and differentiation. In addition, the use of bFGF may also stimulate angiogenesis in vivo, which in turn will stimulate cell survival and differentiation. 4.1. bFGF Release and Bioactivity. The hydrogel used in this work demonstrated high bFGF encapsulation efficiency (>94%). The encapsulated bFGF showed a sustained release profile during a 21-day testing period (Figure 2). The released bFGF preserved its bioactivity throughout the release period (Figure 3). The amount of released bFGF was dependent on

the initial loading dosage. A higher release amount was obtained at a higher loading dose. A two-stage release kinetics was observed for all of the bFGF loaded hydrogels: a burst release before 8 h, followed by a slower and sustained release for up to 21 days. The initial burst release may possibly result from diffusion of bFGF at the surface of the hydrogel. The ensuing slow and linear release may be due to strong interactions between heparin and bFGF attenuating bFGF release. Heparin consists of negative charges (carboxylic and sulfate) that form interactions with amine groups in the bFGF.43 In this report, the amount of heparin was 100 times of bFGF. Such a large amount of heparin may form strong interactions with bFGF, leading to slow bFGF release. 4.2. MSC Survival under Low Nutrient and Oxygen Conditions. After transplant into the infarcted heart, cells experience low nutrient and low oxygen conditions, leading to massive cell death.52,53 We investigated how nutrient and oxygen levels affect cell survival both individually and in concert. MSC-encapsulated hydrogels were cultured under normal (10% FBS) and low (1% FBS) nutrient conditions to study the effect of nutrients on cell survival. In addition, cells were cultured under three oxygen levels (21%, 5%, and 1%) to study the effect of oxygen level on cell survival under low nutrient conditions (1% FBS). Figure 4 shows that MSCs proliferated in the hydrogel under normal nutrient and oxygen conditions (10% FBS and 21% O2). However, the low nutrient condition (1% FBS) significantly decreased MSC survival even when the oxygen level was normal. This indicates that nutrients play an important role in MSC survival. Under low nutrient conditions (1% FBS), a decrease in oxygen level from 21% to 5% did not significantly affect cell survival. This may be due to 1961

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

Figure 7. Immunohistological staining of connexin 43 for MSCs in hydrogels with or without bFGF. Cells were cultured under different conditions: 10% FBS and 21% oxygen, 1% FBS and 21% oxygen, 1% FBS and 5% oxygen, and 1% FBS and 1% oxygen. Scale bar 25 μm.

FBS). Cells in bFGF-loaded hydrogels showed significantly higher survival rates at all oxygen levels. Surprisingly, MSCs even proliferated at 21% and 5% oxygen levels, while significant cell death was observed in control gels under the same conditions (Figure 4). At a 1% oxygen level, bFGF-loaded hydrogel retained 70.5% cell survival, compared to 4.9% in nonbFGF loaded hydrogel. These results demonstrate that using bFGF may remarkably improve MSC survival in the infarcted heart. These results are also consistent with previous reports that FGFs, including bFGF and FGF-9, can inhibit cell apoptosis under serum depletion conditions in vitro and in vivo.58−60 The improved cell survival is likely due to the activation of PI3k/Akt59 and/or hypoxia-induced factor 1 (HIF-1) pathways by bFGF.61 4.3. Effect of bFGF on MSC Differentiation to Cardiomyocyte-like Cells under Low Nutrient and Oxygen Conditions. Under normal culture conditions (10% FBS and 21% O2), MSCs in the hydrogel differentiated into cardiomyocyte-like cells, confirmed by the expression of cardiac markers at both the gene and protein levels (Figures 5−7). The differentiation is probably initiated by stiffness and chemical environment of the hydrogel. Various studies have shown that stem cells can differentiate into corresponding lineages in the matrices whose stiffnesses match those of the tissues in which the cells reside.28−30,62−66 The hydrogel used in this report had a stiffness of 45 kPa, within the range of the native heart tissue (1−140 kPa).67 Chemical groups in the hydrogel may also be a cause of the differentiation. Benoit et al. demonstrated that chemical groups like acid and amine in poly(ethylene glycol) hydrogels can stimulate MSC differentiation without using

Figure 8. VEGF expression of MSCs in the hydrogels under different culture conditions. Cells were cultured under conditions of 10% FBS and 21% oxygen, 1% FBS and 21% oxygen, 1% FBS and 5% oxygen, and 1% FBS and 1% oxygen, respectively. The expression was normalized by that under 10% FBS with 21% oxygen culture condition.

the native bone marrow environment, where MSCs reside, having an oxygen level of 5%.54 The cells are thus tolerant of this oxygen level.55,56 A further decrease in oxygen to 1%, a level similar to that in the infarcted heart,57 caused significant cell death. Only 4.9% of MSCs had survived after 14 days. This result confirmed previous in vivo studies, where low nutrient and oxygen conditions caused extensive cell death.52,53 Introducing bFGF into the hydrogel did not significantly affect MSC proliferation under normal culture conditions (20% FBS and 21% O2, Figure 4). However, bFGF significantly improved MSC survival under low nutrient conditions (1% 1962

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

differentiation medium.68 Addition of bFGF into the hydrogel enhanced the MSC differentiation into cardiomyocyte-like cells. This observation is consistent with previous studies where MSC differentiation was enhanced by bFGF under normal culture conditions.69 MSC differentiation to cardiomyocyte-like cells is dependent on nutrient level. Decreasing FBS from 10% to 1% significantly decreased MEF2C expression. Interestingly, it did not significantly decrease the ratio of cells positive to cTnI. These results demonstrate that MSCs remains at a high differentiation efficiency to cardiomyocyte-like cells under low nutrient conditions if the oxygen level is normal (21%). Under this condition, bFGF loading did not significantly increase the number of cells positive to cTnI, although it did significantly increase the MEF2C expression at the gene level. At low nutrient levels (1% FBS), MSC differentiation to cardiomyocyte-like cells is highly dependent on the oxygen level. When the oxygen was reduced from 21% to 5% and 1%, the MEF2C expression was completely silenced. The ratio of cells positive to cTnI was reduced from 93.9% to 63.2% and 0%, respectively. Similarly, connexin 43 expression was decreased. These results may explain the outcome in many in vivo studies, where an extremely low ratio of MSCs differentiated into cardiomyocytes. The differentiation is retarded by the low nutrient and low oxygen conditions in the heart, in addition to the stiff scar tissue. The nutrient- and oxygen-dependent differentiation of MSCs was also reported by other studies when MSCs were differentiated into osteoblasts and chondrocytes.10,20,70,71 Under low nutrient and low oxygen conditions, addition of bFGF remarkably enhanced MSC differentiation to cardiomyocyte-like cells. The MEF2C and CACNA1C expressions and the ratio of cTnI positive cells were significantly up-regulated. The extent of up-regulation was higher at the 5% oxygen level than at the 1% oxygen level. The stimulatory effect of bFGF on MSC differentiation may result both from bFGF directly and from enhanced cell survival when using bFGF. The results in this study demonstrate that simply delivering MSCs and the hydrogel capable of differentiating MSCs into cardiomyocyte-like cells may not lead to a significant differentiation. Improving cell survival under low nutrient and oxygen conditions may warrant the differentiation. 4.4. The Effect of bFGF to MSC Paracrine Effect. The paracrine effect is considered a key benefit of using MSCs for MI therapy.70,72 VEGF is one of the important cytokines involved in the paracrine effect. Under normal culture conditions, addition of bFGF substantially up-regulated VEGF expression. Reducing FBS from 10% to 1%, and oxygen levels from 21% to 5%, did not affect VEGF expression. Under these conditions, the addition of bFGF significantly increased VEGF. However, the VEGF expression was largely suppressed when the oxygen level was decreased to 1% (Figure 8). The addition of bFGF did not significantly improve the expression. These results demonstrate that the paracrine effect of VEGF is low in the infarcted heart environment. The improvement of the paracrine effect with the addition of bFGF at oxygen levels of 5% and higher is possibly due to the activation of the Akt pathway in MSCs.73−75 It was reported that MSCs, after being genetically modified with Akt, have an enhanced paracrine effect.10

5. CONCLUSIONS We reported a thermosensitive and injectable hydrogel with the ability to deliver bFGF for enhanced MSC differentiation to cardiomyocyte-like cells under the low nutrient and oxygen conditions, typical of the infarcted hearts. The hydrogel was capable of sustained release of bioactive bFGF for 21 days. The low nutrient and low oxygen conditions caused extensive cell death and retarded differentiation. bFGF incorporation significantly improved MSC survival and differentiation. These results suggest that bFGF-loaded hydrogel has the potential to be delivered into infarcted hearts together with MSCs for cardiac regeneration.

■

AUTHOR INFORMATION

Corresponding Author

*Department of Materials Science and Engineering, The Ohio State University, 2041 College Road, Columbus, OH 43210. Phone: 614-292-9743. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work was supported by the National Science Foundation (DMR1006734) and the Institute for Materials Research at The Ohio State University.

■

REFERENCES

(1) Chen, J.; Rizzo, J. A. Crit. Care Clin. 2012, 28, 77−88, vi. (2) Martinez, E. C.; Kofidis, T. J. Mol. Cell. Cardiol. 2011, 50, 312− 319. (3) Davis, D. R.; Zhang, Y.; Smith, R. R.; Cheng, K.; Terrovitis, J.; Malliaras, K.; Li, T.-S.; White, A.; Makkar, R.; Marbán, E. PLoS One 2009, 4, e7195. (4) He, J.-Q.; Vu, D. M.; Hunt, G.; Chugh, A.; Bhatnagar, A.; Bolli, R. PLoS One 2011, 6, e27719. (5) Leri, A.; Kajstura, J.; Anversa, P. Circ. Res. 2011, 109, 941−961. (6) Huang, C.; Gu, H.; Yu, Q.; Manukyan, M. C.; Poynter, J. A.; Wang, M. PLoS One 2011, 6, e29246. (7) Liu, X.-H.; Bai, C.-G.; Xu, Z.-Y.; Huang, S.-D.; Yuan, Y.; Gong, D.-J.; Zhang, J.-R. Microvasc. Res. 2008, 76, 23−30. (8) Williams, A. R.; Hare, J. M. Circ. Res. 2011, 109, 923−940. (9) Enoki, C.; Otani, H.; Sato, D.; Okada, T.; Hattori, R.; Imamura, H. Int. J. Cardiol. 2010, 138, 9−18. (10) Gnecchi, M.; He, H.; Noiseux, N.; Liang, O. D.; Zhang, L.; Morello, F.; Mu, H.; Melo, L. G.; Pratt, R. E.; Ingwall, J. S.; Dzau, V. J. FASEB J. 2006, 20, 661−669. (11) Wang, F.; Guan, J. Adv. Drug Delivery Rev. 2010, 62, 784−797. (12) Grinnemo, K.-H.; Månsson-Broberg, A.; Leblanc, K.; Corbascio, M.; Wärdell, E.; Siddiqui, A. J.; Hao, X.; Sylvén, C.; Dellgren, G. Ann. Med. 2006, 38, 144−153. (13) Dai, W.; Hale, S. L.; Martin, B. J.; Kuang, J.-Q.; Dow, J. S.; Wold, L. E.; Kloner, R. A. Circulation 2005, 112, 214−223. (14) Reffelmann, T.; Kloner, R. A. Cardiovasc. Res. 2003, 58, 358− 368. (15) Hofmann, M.; Wollert, K. C.; Meyer, G. P.; Menke, A.; Arseniev, L.; Hertenstein, B.; Ganser, A.; Knapp, W. H.; Drexler, H. Circulation 2005, 111, 2198−2202. (16) Noiseux, N.; Gnecchi, M.; Lopez-Ilasaca, M.; Zhang, L.; Solomon, S. D.; Deb, A.; Dzau, V. J.; Pratt, R. E. Mol. Ther. 2006, 14, 840−850. (17) Wang, F.; Li, Z.; Khan, M.; Tamama, K.; Kuppusamy, P.; Wagner, W. R.; Sen, C. K.; Guan, J. Acta Biomater. 2010, 6, 1978− 1991. (18) Zhang, D.; Fan, G.-C.; Zhou, X.; Zhao, T.; Pasha, Z.; Xu, M.; Zhu, Y.; Ashraf, M.; Wang, Y. J. Mol. Cell. Cardiol. 2008, 44, 281−292.

1963

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964

Biomacromolecules

Article

(19) He, Z.; Li, H.; Zuo, S.; Pasha, Z.; Wang, Y.; Yang, Y.; Jiang, W.; Ashraf, M.; Xu, M. Stem Cells Dev. 2011, 20, 1771−1778. (20) Zuo, S.; Jones, W. K.; Li, H.; He, Z.; Pasha, Z.; Yang, Y.; Wang, Y.; Fan, G.-C.; Ashraf, M.; Xu, M. Stem Cells Dev. 2012, 21, 598−608. (21) Rosenblatt-Velin, N.; Lepore, M. G.; Cartoni, C.; Beermann, F.; Pedrazzini, T. J. Clin. Invest. 2005, 115, 1724−1733. (22) van Oorschot, A. A. M.; Smits, A. M.; Pardali, E.; Doevendans, P. A.; Goumans, M.-J. J. Cell. Mol. Med. 2011, 15, 2723−2734. (23) Martens, T. P.; Godier, A. F. G.; Parks, J. J.; Wan, L. Q.; Koeckert, M. S.; Eng, G. M.; Hudson, B. I.; Sherman, W.; VunjakNovakovic, G. Cell Transplant. 2009, 18, 297−304. (24) Hoare, T. R.; Kohane, D. S. Polymer 2008, 49, 1993−2007. (25) Soontornworajit, B.; Zhou, J.; Zhang, Z.; Wang, Y. Biomacromolecules 2010, 11, 2724−2730. (26) Savla, R.; Taratula, O.; Garbuzenko, O.; Minko, T. J. Controlled Release 2011, 153, 16−22. (27) Asai, D.; Xu, D.; Liu, W.; Garcia Quiroz, F.; Callahan, D. J.; Zalutsky, M. R.; Craig, S. L.; Chilkoti, A. Biomaterials 2012, DOI: 10.1016/j.biomaterials.2012.03.083. (28) Engler, A. J.; Carag-Krieger, C.; Johnson, C. P.; Raab, M.; Tang, H.-Y.; Speicher, D. W.; Sanger, J. W.; Sanger, J. M.; Discher, D. E. J. Cell. Sci. 2008, 121, 3794−3802. (29) Engler, A. J.; Sen, S.; Sweeney, H. L.; Discher, D. E. Cell 2006, 126, 677−689. (30) Young, J. L.; Engler, A. J. Biomaterials 2011, 32, 1002−1009. (31) Li, X.; Yu, X.; Lin, Q.; Deng, C.; Shan, Z.; Yang, M.; Lin, S. J. Mol. Cell. Cardiol. 2007, 42, 295−303. (32) Yang, M.-C.; Wang, S.-S.; Chou, N.-K.; Chi, N.-H.; Huang, Y.Y.; Chang, Y.-L.; Shieh, M.-J.; Chung, T.-W. Biomaterials 2009, 30, 3757−3765. (33) Bartunek, J.; Croissant, J. D.; Wijns, W.; Gofflot, S.; de Lavareille, A.; Vanderheyden, M.; Kaluzhny, Y.; Mazouz, N.; Willemsen, P.; Penicka, M.; Mathieu, M.; Homsy, C.; De Bruyne, B.; McEntee, K.; Lee, I. W.; Heyndrickx, G. R. Am. J. Physiol.: Heart Circ. Physiol. 2007, 292, H1095−1104. (34) Balana, B.; Nicoletti, C.; Zahanich, I.; Graf, E. M.; Christ, T.; Boxberger, S.; Ravens, U. Cell Res. 2006, 16, 949−960. (35) Engler, A. J.; Griffin, M. A.; Sen, S.; Bönnemann, C. G.; Sweeney, H. L.; Discher, D. E. J. Cell Biol. 2004, 166, 877−887. (36) 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. (37) Li, Z.; Guo, X.; Matsushita, S.; Guan, J. Biomaterials 2011, 32, 3220−3232. (38) Pollak, A.; Blumenfeld, H.; Wax, M.; Baughn, R. L.; Whitesides, G. M. J. Am. Chem. Soc. 1980, 102, 6324−6336. (39) Guan, J.; Hong, Y.; Ma, Z.; Wagner, W. R. Biomacromolecules 2008, 9, 1283−1292. (40) Li, Z.; Wang, F.; Roy, S.; Sen, C. K.; Guan, J. Biomacromolecules 2009, 10, 3306−3316. (41) Pakulska, M. M.; Ballios, B. G.; Shoichet, M. S. Biomed. Mater. 2012, 7, 024101. (42) Homicz, M. R.; Watson, D. Facial Plast. Surg. 2004, 20, 21−29. (43) Faham, S.; Hileman, R. E.; Fromm, J. R.; Linhardt, R. J.; Rees, D. C. Science 1996, 271, 1116−1120. (44) Shi, Y.-H.; Bingle, L.; Gong, L.-H.; Wang, Y.-X.; Corke, K. P.; Fang, W.-G. Pathology 2007, 39, 396−400. (45) Pang, Y.; Wang, X.; Ucuzian, A. A.; Brey, E. M.; Burgess, W. H.; Jones, K. J.; Alexander, T. D.; Greisler, H. P. Biomaterials 2010, 31, 878−885. (46) Ormsby, R.; McNally, T.; O’Hare, P.; Burke, G.; Mitchell, C.; Dunne, N. Acta Biomater. 2012, 8, 1201−1212. (47) Dominici, M.; Le Blanc, K.; Mueller, I.; Slaper-Cortenbach, I.; Marini, F.; Krause, D.; Deans, R.; Keating, A.; Prockop, D.; Horwitz, E. Cytotherapy 2006, 8, 315−317. (48) Hoganson, D. M.; O’Doherty, E. M.; Owens, G. E.; Harilal, D. O.; Goldman, S. M.; Bowley, C. M.; Neville, C. M.; Kronengold, R. T.; Vacanti, J. P. Biomaterials 2010, 31, 6730−6737.

(49) Lindoso, R. S.; Araujo, D. S.; Adão-Novaes, J.; Mariante, R. M.; Verdoorn, K. S.; Fragel-Madeira, L.; Caruso-Neves, C.; Linden, R.; Vieyra, A.; Einicker-Lamas, M. Cell. Physiol. Biochem. 2011, 28, 267− 278. (50) Hou, D.; Youssef, E. A.-S.; Brinton, T. J.; Zhang, P.; Rogers, P.; Price, E. T.; Yeung, A. C.; Johnstone, B. H.; Yock, P. G.; March, K. L. Circulation 2005, 112, I150−156. (51) McGinn, A. N.; Nam, H. Y.; Ou, M.; Hu, N.; Straub, C. M.; Yockman, J. W.; Bull, D. A.; Kim, S. W. Biomaterials 2011, 32, 942− 949. (52) Zhu, W.; Chen, J.; Cong, X.; Hu, S.; Chen, X. Stem Cells 2006, 24, 416−425. (53) Potier, E.; Ferreira, E.; Meunier, A.; Sedel, L.; LogeartAvramoglou, D.; Petite, H. Tissue Eng. 2007, 13, 1325−1331. (54) Lennon, D. P.; Edmison, J. M.; Caplan, A. I. J. Cell. Physiol. 2001, 187, 345−355. (55) Salim, A.; Nacamuli, R. P.; Morgan, E. F.; Giaccia, A. J.; Longaker, M. T. J. Biol. Chem. 2004, 279, 40007−40016. (56) Rochefort, G. Y.; Delorme, B.; Lopez, A.; Hérault, O.; Bonnet, P.; Charbord, P.; Eder, V.; Domenech, J. Stem Cells 2006, 24, 2202− 2208. (57) Rumsey, W. L.; Pawlowski, M.; Lejavardi, N.; Wilson, D. F. Am. J. Physiol. 1994, 266, H1676−1680. (58) Matsunaga, S.; Okigaki, M.; Takeda, M.; Matsui, A.; Honsho, S.; Katsume, A.; Kishita, E.; Che, J.; Jishan, C.; Kurihara, T.; Adachi, Y.; Mansukhani, A.; Kobara, M.; Matoba, S.; Matoba, Y.; Tatsumi, T.; Matsubara, H. J. Mol. Cell. Cardiol. 2009, 46, 663−673. (59) Park, S. J.; Kim, S. H.; Choi, H. S.; Rhee, Y.; Lim, S.-K. Bone 2009, 45, 994−1003. (60) Bolitho, C.; Xu, W.; Zoellner, H. J. Vasc. Res. 2008, 45, 193− 204. (61) Egger, M.; Schgoer, W.; Beer, A. G. E.; Jeschke, J.; Leierer, J.; Theurl, M.; Frauscher, S.; Tepper, O. M.; Niederwanger, A.; Ritsch, A.; Kearney, M.; Wanschitz, J.; Gurtner, G. C.; Fischer-Colbrie, R.; Weiss, G.; Piza-Katzer, H.; Losordo, D. W.; Patsch, J. R.; Schratzberger, P.; Kirchmair, R. FASEB J. 2007, 21, 2906−2917. (62) Berry, M. F.; Engler, A. J.; Woo, Y. J.; Pirolli, T. J.; Bish, L. T.; Jayasankar, V.; Morine, K. J.; Gardner, T. J.; Discher, D. E.; Sweeney, H. L. Am. J. Physiol.: Heart Circ. Physiol. 2006, 290, H2196−2203. (63) Rowlands, A. S.; George, P. A.; Cooper-White, J. J. Am. J. Physiol.: Cell Physiol. 2008, 295, C1037−1044. (64) Kim, J.; Park, Y.; Tae, G.; Lee, K. B.; Hwang, S. J.; Kim, I. S.; Noh, I.; Sun, K. J. Mater. Sci. Mater. Med. 2008, 19, 3311−3318. (65) Pek, Y. S.; Wan, A. C. A.; Ying, J. Y. Biomaterials 2010, 31, 385− 391. (66) Wang, L.-S.; Boulaire, J.; Chan, P. P. Y.; Chung, J. E.; Kurisawa, M. Biomaterials 2010, 31, 8608−8616. (67) Chen, Q.-Z.; Harding, S. E.; Ali, N. N.; Lyon, A. R.; Boccaccini, A. R. Mater. Sci. Eng., R 2008, 59, 1−37. (68) Benoit, D. S. W.; Schwartz, M. P.; Durney, A. R.; Anseth, K. S. Nat. Mater. 2008, 7, 816−823. (69) Cucchiarini, M.; Ekici, M.; Schetting, S.; Kohn, D.; Madry, H. Tissue Eng., Part A 2011, 17, 1921−1933. (70) Weil, B. R.; Markel, T. A.; Herrmann, J. L.; Abarbanell, A. M.; Meldrum, D. R. Surgery 2009, 146, 190−197. (71) Psaltis, P. J.; Zannettino, A. C. W.; Worthley, S. G.; Gronthos, S. Stem Cells 2008, 26, 2201−2210. (72) Takahashi, M.; Li, T.-S.; Suzuki, R.; Kobayashi, T.; Ito, H.; Ikeda, Y.; Matsuzaki, M.; Hamano, K. Am. J. Physiol.: Heart Circ. Physiol. 2006, 291, H886−893. (73) Tsubokawa, T.; Yagi, K.; Nakanishi, C.; Zuka, M.; Nohara, A.; Ino, H.; Fujino, N.; Konno, T.; Kawashiri, M.; Ishibashi-Ueda, H.; Nagaya, N.; Yamagishi, M. Am. J. Physiol.: Heart Circ. Physiol. 2010, 298, H1320−1329. (74) Deng, J.; Han, Y.; Yan, C.; Tian, X.; Tao, J.; Kang, J.; Li, S. Apoptosis 2010, 15, 463−473. (75) Yin, Q.; Jin, P.; Liu, X.; Wei, H.; Lin, X.; Chi, C.; Liu, Y.; Sun, C.; Wei, Y. Mol. Biol. Rep. 2011, 38, 9−16.

1964

dx.doi.org/10.1021/bm300574j | Biomacromolecules 2012, 13, 1956−1964