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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Biotechnol Bioeng. Author manuscript; available in PMC 2010 May 1.
Published in final edited form as:
PMCID: PMC2847036
NIHMSID: NIHMS179219

Microporous Cell-laden Hydrogels for Engineered Tissue Constructs

Abstract

In this paper, we describe an approach to generate microporous cell-laden hydrogels for fabricating biomimetic tissue engineered constructs. Micropores at different length scales were fabricated in cell-laden hydrogels by micromolding fluidic channels and leaching sucrose crystals. Microengineered channels were created within cell-laden hydrogel precursors that contained agarose solution mixed with sucrose crystals. The rapid cooling of the agarose solution was used to gel the solution and form micropores in place of the sucrose crystals. The sucrose leaching process generated micropores that were homogeneously distributed within the gels, while enabling the direct immobilization of cells within the gels. We also characterized the physical, mechanical, and biological properties (i.e. microporosity, diffusivity, and cell viability) of cell-laden agarose gels as a function of engineered porosity. The microporosity was controlled from 0% to 40% and the diffusivity of molecules in the porous agarose gels increased as compared to controls. Furthermore, the viability of human hepatocyte cells that were cultured in microporous agarose gels corresponded to the diffusion profile generated away from the microchannels. Based on their enhanced diffusive properties, microporous cell-laden hydrogels containing a microengineered fluidic channel could be a useful tool for generating tissue structures for regenerative medicine and drug discovery applications.

Keywords: Microporous, Agarose, Cell-laden hydrogel, Tissue engineering

1. Introduction

Hydrogels hold great potential as scaffolding materials for a number of biological applications such as regenerative medicine, drug discovery, and biosensors since they can provide physiological environments with characteristics such as high water content, high porosity, and mechanical support(Khademhosseini and Langer 2007; Peppas et al. 2006). Hydrogels have been used for tissue engineering fields for a number of tissue types, including bone(Burdick and Anseth 2002; Burdick et al. 2003; Burdick et al. 2002), cartilage(Bryant and Anseth 2003), liver(Liu Tsang et al. 2007), brain(Bakshi et al. 2004; Ford et al. 2006; Tian et al. 2005; Woerly 1993), and others(Changez et al. 2004; Elisseeff et al. 2000; Mann et al. 2001). Due to their excellent properties, hydrogels derived from natural sources (i.e. collagen, hyaluronic acid (HA), chitosan, alginate, and agarose) or synthetic methods (i.e. poly(ethylene glycol) (PEG)) have been extensively used for various tissue engineering applications(Khademhosseini et al. 2006; Lee and Mooney 2001; Wu et al. 2008).

Agarose, a temperature-sensitive and water soluble hydrogel, is a polysaccharide extracted from marine red algae(Aymard et al. 2001) and is used as a cell culture substrate(Uludag et al. 2000). The mechanical properties of agarose gels can be controlled by gelling temperatures and curing times(Aymard et al. 2001). Furthermore, the diffusion properties of macromolecules such as proteins, polymer beads, and DNAs within agarose gels have been characterized by using fluorescence based methods(Pluen et al. 1999) and movement of nanoparticles(Fatin-Rough et al. 2004; Labille et al. 2007). Biocompatible agarose gel has been used for the cell encapsulation and in vivo transplantation applications(Rahfoth et al. 1998; Uludag et al. 2000).

Porous structures in biomaterials are potentially useful for mimicking native tissues. Pores improve protein transport and diffusion in agarose gels. To make porous scaffolds, several methods have been previously developed. For example, colloidal suspension was used to create pores within hydroxyapatite (HA) scaffolds(Cordell et al. 2009). The mechanical bending and compression analysis of this scaffold has shown that strength of the bulk microporous HA with smaller micropore sizes was higher as compared to HA with larger micropore sizes. This was consistent with reported results for HA and other porous materials(Bignon et al. 2003; Sopyan et al. 2007). Poly(methyl methacrylate) (PMMA) beads were used to generate microporous structures within fibrin scaffolds(Linnes et al. 2007). To create micropores within scaffolds, PMMA beads were removed by using toxic chemical processes.

Sucrose is a promising crystal as a pore or particle forming agent(Huang et al. 2003; Kwok et al. 2000). It has been used to create particles and pores within polylactideglycolic acid (PLGA) sponges during a gas foaming process(Huang et al. 2003). The elastic modulus of gas-foamed PLGA sponges was decreased with increasing sucrose concentrations. In addition to sucrose crystal leaching method, salt crystals have been previously used to create interconnected pores within polymeric scaffolds(Murphy et al. 2002). Porous scaffolds of poly(lactide-co-glycolide) were fabricated by solvent casting/particulate leaching or gas foaming leaching methods using a salt. Fusion of salt crystals in the solvent casting process enhanced pore interconnectivity within polymeric scaffolds. The pore size was controlled by using NaCl microparticles. The mechanical properties (i.e. compressive modulus) of scaffolds were strongly dependent on salt fusion and processes, such as solvent casting and gas foaming. However, although these previous methods enable the control of mechanical properties of scaffolds, they have potential limitations, such as the inability of cell encapsulation due to toxic chemical processes, e.g. gas foaming and solvent casting method.

With agarose gels, several chemical methods have been also used to create pores(Shi et al. 2005; Zhou et al. 2006). For example, pores have been made by water-in-oil emulsification using solid granules of calcium carbonate(Shi et al. 2005) and metal oxides have also been used for macropore within agarose gels(Zhou et al. 2006). However, these methods suggested to create pores within agarose gels could not be useful for the cell-laden hydrogel applications due to their non-biocompatible processes. Cell-laden hydrogel microfluidic devices can mimic the 3D microenvironment of the in vivo tissue constructs(Cabodi et al. 2005; Choi et al. 2007; Gillette et al. 2008; Golden and Tien 2007; Hwang et al. 2008; Ling et al. 2007). The integration of microfabricated devices and biocompatible hydrogels offers the potential for recreating the spatial complexity and diffusion properties of macromolecules. We have previously developed a cell-laden agarose microfluidic system and analyzed the diffusion profiles of molecules from the microchannels(Ling et al. 2007). Given this feature, we hypothesize that the ability to create micropores within the gels around the microchannels may provide potential improvements in biomolecular diffusion and oxygen transport.

In this paper, we describe a method to fabricate a cell-laden agarose gel system containing engineered constructs with a microvascular structure and micropores that are created by dissolving sucrose crystals without the use of any organic solvents. For this purpose, we developed the porous cell-laden agarose fluidic device and characterized the physical and mechanical properties of agarose gels with various micropores. We also analyzed the viability of hepatic cells encapsulated within agarose gels. Therefore, this porous cell-laden agarose gel system integrated with a microvascularized channel could be a potentially useful tool to study complex cell-microenvironment interactions and mimic microarchitectures of native tissues.

2. Materials and Methods

2.1. Fabrication of the microporous cell-laden agarose gels

We fabricated microporous cell-laden agarose gels containing a microengineered channel as shown in Figure 1. Briefly, sucrose crystals (200 μm in diameter) at varying concentrations of 0, 100, 200% (w/v) were mixed with 1 ml of the cell suspension (107cells/mL) and an additional 1 ml of 6 wt% agarose solution (Sigma-Aldrich, CA) at 40°C. The initial temperature for sample preparation was 25 °C, 37 °C, and 40 °C for sucrose crystals, cell suspension, and agarose solution respectively. Cells were only exposed to 40 °C agarose solution for short time and were cooled down to 4 °C after mixing with cell suspension and sucrose crystals as shown in Figure 1 (A). After mixing, the mixture was poured into a cylindrical poly(dimethylsiloxane) (PDMS) mold (2 cm diameter, 1 cm thick). To generate the hydrogel microchannel, a microneedle (0.38, 0.6 mm inner and outer diameter) was inserted in the middle of the PDMS side walls as a microcapillary (Figure 1B). The entire molds were then placed either at 25 °C for natural gelation or 4 °C for rapid gelation (Figure 1C). After the agarose gelation (~ 20 min), the microneedle was removed from the PDMS mold to create the microchannel (Figure 1C, D) and the cell-laden agarose gel was immersed within cell culture medium at 37 °C to dissolve the sucrose. The sucrose-leached medium in the bath was changed to fresh medium every 10 minutes. After 2 hours, the sucrose crystals remained within the agarose gels were completely removed (Figure 1E). For the continuous medium perfusion in the agarose microchannel, polyethylene tubing (1/16 inch inner diameter) was connected to metal tubes in PDMS molds. The culture medium was delivered into the microchannel by using a syringe pump (2 μl min−1). Hepatic cells encapsulated within microporous agarose gels were cultured for 5 days in vitro (Figure 1F).

Figure 1
Schematic of the fabrication process for cell-laden hydrogels containing micropores and a microchannel. (a) Sample preparation: Sucrose crystals (50–200 wt%), cells (107 cells ml−1) and agarose (6 wt%) solution were mixed at 40 °C. ...

2.2. Hepatic cell culture and cell viability

The hepatocelluar carcinoma cell line (HepG2) was purchased from American Tissue Type Collection (ATTC). All tissue culture components were purchased from Gibco-Invitrogen, CA, unless otherwise indicated. Culture medium for HepG2 cells consisted of Dulbecco's Modified Eagle Medium (DMEM) with 10% (v/v) fetal bovine serum (FBS), and 1% penicillin-streptomycin. Cells cultured in a tissue culture flask were fed by changing the medium every other day and were passaged when 90% confluency was reached. To analyze the viability of cells cultured within microporous agarose gels, a live/dead assay was used (Molecular Probes Inc., OR).

After culturing for 5 days, tubings for medium perfusion were disconnected and cell-laden agarose gels were cut (1cm×1cm×1cm) by a knife for the cell viability test. These cell-laden agarose gels were subsequently incubated in 2 μM calcein-AM and 4 μM ethidium homodimer for 10 min (37°C, 5% CO2). Live (green) and dead (red) cells around the microchannel of cell-laden agarose gels were analyzed by a fluorescence microscope. For the medium perfusion experiment, we analyzed the cross-sectional images of the microchannel in agarose gels. For the control, we characterized cell viability at 500 μm deep from agarose gel surface. Cell viability test was performed three times for each condition by using a single gel per condition.

2.3. Image analysis

Phase contrast and fluorescent images of cells encapsulated within agarose gels were obtained from an inverted microscope (Nikon, TE 2000). We observed the surfaces of agarose gels within cylindrical PDMS molds (2 cm diameter and 1 cm thick) (Figure 2A~C and Figure 3A(a)~(i)). For Figure 3A(g)~(i), the specimens were cross-sectioned by a razor blade and slightly dried before observation. To observe and identify micropores embedded in agarose gels, we used confocal microscope (Zeiss, LSM 510) and scanning electron microscope (Jeol, JSM-6500F). For fluorescent imaging (Figure 3A(j)~(l)) with the confocal microscope, an agarose solution was mixed with the fluorescein isothiocyanate (FITC)-dextran (0.5 mM, 2000 kDa, Sigma-Aldrich, CA). These phase contrast and fluorescent images for quantifying microporosity were analyzed by using the NIH Image J software with functions for contrast separation, area fractioning, and intensity profiling.

Figure 2
Micrographs of sucrose crystals embedded in agarose gels. (A) Phase contrast images of sucrose mixtures (0–300 wt%) after natural cooling (from 40 °C to 25 °C). (B) Phase contrast images of sucrose mixtures (0–200 wt%) ...
Figure 3
Microporosity and mechanical stiffness of hydrogels. (A) Images of microporosity: Phase contrast images of sucrose crystals (0–200 wt%, a–c), sucrose crystals dissolved within agarose gels (d–f), and cross-section images of agarose ...

2.4. Characterization of hydrogel mechanical properties

We characterized the mechanical stiffness of the gel constructs, which did not contain the cells, by using an Instron 5542 mechanical compression tester at a rate of 20%/min until failure occurred. The compressive modulus of agarose gels containing different sucrose concentrations (0–200 wt%) was obtained from the linear regime in the 10–15% strain.

2.5. Modeling of the diffusion profiles in hydrogels

Diffusion in the extracelluar space of cell-laden hydrogels is analogous to diffusion in a porous medium. To measure the diffusion properties of agarose gels, the integrative optical imaging (IOI) technique(Nicholson 2001) could be useful for analyzing macromolecules. In case of which a few nanoliters of dextran labeled with fluorescent dye diffuses away from the agarose gel, the concentration of the fluorescent dye is decreased as a function of time. If the concentration profile is extracted from the agarose, the diffusion can be easily characterized as a diffusion coefficient. For diffusion equation, the Fick's law and the conservation of material with the space average leads to the diffusion coefficient. If a representative elementary volume of hydrogels in the narrow space is assumed to be V and the extracellular space is defined as V0, the diffusion model(Nicholson 2001) can be expressed as

tC0=D^2C0+sα
(1)

Where the operator is 2=2xi2, C0 is the concentration in the extracellular space, s is the source density,α is the porosity defined in the porous medium as V0V, the operator left angle bracket right angle bracket is space average, and D^=DK^ is the effective diffusion coefficient of the hydrogel which is a second-order tensor. The tensor K^ is a reciprocal proportion to the tortuosity of the hydrogels. If the hydrogels are uniform in the averaging space of interest, the tortuosity (K^) is simplified as a scalar being in the inverse ratio to the square of tortuosity (1λ2). In addition, Nicholson and Phillips showed that the diffusion equation in the extracellular space could be described in a free medium as follows(Nicholson and Phillips 1981):

tC=D^2C+sα
(2)

The equation is simplified by dropping the term (s) in cases where there is no source density in the extracellular space.

tC=D^2C
(3)

Note that the diffusion coefficient (D^) is a vector in a space.

Although non-uniform transport partially brings out convective term due to partial inhomogeneous pores, the spatially averaged intensity allows for the diffusivity in a specimen to be considered as a uniform transport. In addition, the evaluation of diffusion properties in a static condition is important because diffusion coefficient should be satisfied in a condition which excludes convection effect due to an infusion rate. In other words, diffusion takes place due to the Brownian motion which is caused by the concentration difference. In our experiment and simulation, the infusion effect is not considered and the intensity is only measured in the x-y plane of the hydrogel specimen after starting the diffusion of FITC-dextran into an agarose microchannel due to the concentration difference.

2.6 Assumption of the modeling

This model neglects the source density which contributes to the transient diffusion profiles except the initial concentration of the fluorescent dye. The hydrogel for experiment has a uniform porous size, so that the diffusion coefficient (D^) is considered as constant in a space. In addition, there is no evaporation of the fluorescent dye into the environment during experiment.

3. Results and Discussion

3.1. Morphology and mechanical property of microporous gels

The microporosity within hydrogels plays a significant role in controlling the delivery of nutrient and oxygen transport to the cells. The microporosity was created by leaching sucrose crystals within agarose gels. Sucrose concentrations enable the control of the percentages of microporosity and mechanical stiffness of hydrogels. Agarose solution was gelled as temperature was decreased. During natural cooling from 40 °C to 25 °C for gelation of agarose, hydrogels derived from the formulation with 200 wt% sucrose contained homogeneous crystals, while sucrose crystals were aggregated in 300 wt% sucrose (Figure 2A (c–d)).

The gelation was performed by decreasing temperature from 40 °C to 25 °C (~ 2 h). However, 2 hours for the gelation process in hydrogels derived from the formulation with 200 wt% sucrose might result in physiologically osmotic shock in an initial stage. The alternative method for addressing this challenge is to decrease the gelation temperature, as solubility of crystals was dominated by the temperature. The gelation time significantly decreased when temperatures were decreased. Here, we used 4°C which is suitable for rapid gelation while maintaining cell viability. Therefore, we performed rapid cooling to 4 °C for fast gelation. For the rapid cooling (40 °C →4 °C), the densities of the sucrose crystals in Figure 2B was similar to the half densities of the sucrose crystals during natural cooling (40 °C → 25 °C) as shown in Figure 2A. Sucrose crystals within hydrogels derived from the formulation with 100 wt% sucrose were relatively homogeneously distributed, while they were aggregated in hydrogels derived from the formulation with 200 wt% sucrose. The gelation time was also reduced to 20 min during the rapid cooling to prevent the potential osmotic shock caused from the natural cooling process. Figure 2C shows phase contrast images of hydrogels derived from the formulation with 100 wt% sucrose which remains within agarose gels. It was revealed that most sucrose crystals within hydrogels derived from the formulation with 100 wt% sucrose in the agarose gels were completely dissolved after 90 min.

We identified micropores, which were substituted for the sucrose microcrystals, by using three microscopes: inverted microscope (Figure 3A (a)~(i)), confocal microscope (Figure 3A (j)~(l)), and scanning electron microscope (Figure 3A (m)~(o)). As expected, the relatively homogeneous distribution of pores was observed in hydrogels derived from the formulation with 100 wt% sucrose. In hydrogels derived from the formulation with 200 wt% sucrose, pores were interconnected due to aggregation of the sucrose crystals. Figure 3A (k–l) shows that the diameters of the average pores are approximately 200 μm which is similar to the original diameter of sucrose crystals. Furthermore, the microporosity was characterized with the sucrose concentrations (Figure 3B). Above 50 wt% of sucrose, the porosity percentage was linearly increased with sucrose concentrations. This result indicates that we can control pore sizes (i.e. single pores with 200 μm diameter and interconnected pores) and porosities by using various sucrose concentrations.

To characterize the effects of sucrose concentrations on the mechanical stiffness of the agarose gels, we performed the compressive testing by using Instron mechanical tester (Figure 3C). Hydrogels derived from the formulation with 100 wt% sucrose showed the half compressive modulus (63.6 ± 33.0 kPa) as compared to the compressive modulus (129.8 ± 7.0 kPa) of non-porous agarose gels. The compressive modulus (14.7 ± 3.0 kPa) of hydrogels derived from the formulation with 200 wt% sucrose was lower than 15% of non-porous agarose gels. These microporosity and mechanical stiffness results demonstrated that percentages of the microporosity were directly proportional to sucrose concentrations, while compressive moduli were inversely proportional to sucrose concentrations. Although hydrogels derived from the formulation with 200 wt% sucrose contain 40% microporosity, the careful mechanical handling is required because they have the lowest compressive modulus. However, hydrogels derived from the formulation with 100 wt% sucrose show good mechanical robustness and microporosity (15%). Therefore, sucrose concentrations enabled the control of the microporosity and mechanical stiffness. The control of these properties could prove advantageous for tailoring the hydrogels to match specific tissue types.

The microporous hydrogels derived from 100 wt% sucrose-leaching show uniform pore sizes that were similar to the original pore size of the initial sucrose crystals. Nonetheless, we observed the relatively large deviation of the compressive modulus (Figure 3C). This deviation is probably due to small local connectivity among the micropores derived from the sucrose crystals.

3.2. Diffusion profiles from the microchannel within microporous agarose gels

Micropores enable the control of diffusion profiles of soluble molecules from the microchannel within agarose gels. We analyzed the diffusion profiles within agarose gels by using a fluorescent dye (FITC-Dextran, 0.25 mM, 20 kDa). In general, FITC-dextran has a similar molecular weight to soluble growth factors associated with metabolism in the body. The channel surface of hydrogels derived from the formulation with 100 wt% sucrose (Figure 4A (d)) was relatively rough as compared to that of 0 wt% sucrose mixtures (Figure 4A (b)) due to micropores around the microchannel.

Figure 4
Diffusion profiles in agarose gels containing the microchannel and micropores. (A) Phase contrast images of a microchannel within agarose gels. (B) Phase contrast and fluorescent images of diffusion profiles in the agarose microchannels containing different ...

To characterize the diffusion patterns as a function of time at each sucrose concentration, we performed diffusion experiments in a static condition after infusion of FITC-dextran into an agarose microchannel (Figure 4B). The evaluation of diffusion properties in a static condition is important, because it can exclude the surface roughness effect that may cause non-pure diffusion, including a convection term derived from shear stress or friction. As expected, in hydrogels derived from the formulation with 100 wt% and 200 wt% sucrose (Figure 4B (d)–(i)), diffusion patterns were not uniform due to micropores around the microchannel. The diffusion coefficient can be defined as the diffusion occurs in a Brownian motion by pure diffusion. Thus, the convective effect by non-uniform pores brings about an undesirable diffusion coefficient. In our experiment, non-uniform transport partially makes convective term due to partial inhomogeneous pores during the diffusion process. To minimize the convective effect, Nicholson et al. (Nicholson and Tao 1993; Thorne and Nicholson 2006) introduced a diffusion model in partial inhomogeneous model by space average. In this paper, the spatially averaged intensity was obtained after t=10 min and was applied to the diffusion Eqs.(1)(3). Note that the equations in this paper are modified from general pure diffusion equation for spatially averaged pure-diffusion.

We also characterized spatio-temporal diffusion patterns at each sucrose mixture as a function of distance away from the channel surface (Figure 4C). The simulation results were in agreement with experimental results of diffusion patterns. The diffusion coefficient of the microporous cell-laden agarose gels was calculated by using finite element method (FEM, Comsol) and was subsequently compared to the diffusion experiments in agarose gels over time. The simulation for Eq. (2) can be conducted in the finite 3D rectangular domain (2 × 1 mm2). The channel was located at the center of the specimen and its diameter and length were about 500 μm and 20 mm, respectively. The normalized initial concentrations were applied inside the channel as 1 mM and the boundaries of the specimen were considered to be zero concentrations. The temporal pattern of the diffusion was calculated inside the gels and channel boundary. The diffusion coefficients were also calculated by simulating hydrogel environments with three different sucrose concentrations (0, 100, and 200 wt%). Since the diffusion is originated from the pure diffusion at the boundary of specimen, it is natural that the concentration decreases as a function of channel length and time in Eq. (3). The simulations were conducted by changing diffusion coefficient to fit the experimental results of Figure 4C (b), (d) and (f). In addition, the diffusion profile was extracted from the channel surface to the boundary of the specimen. These results revealed that diffusion velocities increased as porosity was increased. Our experimental results are in good agreement with the previous studies (Nicholson and Tao 1993; Thorne and Nicholson 2006), where the diffusion coefficients of FITC-dextran in agarose gels were reported between 4.2 and 13.5×10−11 m2 s−1. In our experiment, we aimed to confirm the similar pattern for the diffusivity of FITC-dextran in the cell-laden structure under our experimental conditions, such as temperature, hydrogels condition and perfusion method.

Furthermore, diffusion coefficients of FITC-dextran in the agarose gels were smaller than those in the water (8×10−11 m2 s−1 in 20 kDa dextran)(Cornelissen et al. 2008) (Figure 4D). We found that the diffusion coefficient of FITC-dextran in hydrogels derived from the formulation with 200 wt% sucrose was approximately 1.5 times higher than that in 0 wt% sucrose. Therefore, the diffusion coefficient was increased with increasing the sucrose concentrations. It seems plausible that biomolecules of similar size can increase the diffusivity in the highly porous hydrogels derived from the formulation with 200 wt% sucrose.

3.3. Cell viability of microporous cell-laden agarose gels with a microchannel

Micropores within cell-laden agarose gels enable the control of cell viability, because medium and nutrients can be diffused through micropores. To study the viability of cells encapsulated within agarose gels containing different micropore sizes, we compared static culture condition and medium perfusion condition. Figure 5A and B presents fluorescent images of cells and quantitative analysis of cell viability near the surfaces (500 μm deep from the surface) under static culture conditions (no medium perfusion). We found that cell viability in hydrogels derived from the formulation with 100 wt% sucrose was higher than that in hydrogels derived from the formulation with 0 and 200 wt% sucrose. Figure 5C shows quantitative analysis of cell viability as a function of distance away from the agarose surface in the static condition. It was revealed that cell viability was decreased with increasing the distance away from the agarose surface. However, at the 2,200 μm distance from the agarose surface, cells in hydrogels derived from the formulation with 100 wt% sucrose remained viable (68%) as compared to those in hydrogels derived from the formulation with 0 and 200 wt% sucrose (44%). This result was similar to cell viability near the surface in the static condition (Figure 5B), because agarose gels derived from the formulation with 100 wt% sucrose had 15% micropores and high mechanical stiffness (60 kPa) (Figure 3B, C). Although hydrogels derived from the formulation with 200 wt% sucrose showed the interconnected pores, their mechanical stiffness was approximately 5 times lower than the stiffness of agarose gels with 100 wt% sucrose. Thus, hydrogels derived from the formulation with 200 wt% sucrose might contain weak microstructures (10 kPa). In addition, cell viability in non-porous agarose gels was low, because medium and oxygen could not be easily diffused through smaller pore sizes. We demonstrated that cell viability in hydrogels derived from the formulation with 100 wt% sucrose was gradually decreased (~30%) when increasing the distance away from the agarose surface, while cell viability in hydrogels derived from the formulation with 0 and 200 wt% sucrose was promptly reduced (~45%).

Figure 5
The viability of hepatic cells cultured within agarose gels containing different microporosities without medium perfusion. (A) Fluorescent images of the cell viability at initial time (a), after culturing for 5 days in 0 wt% (b), 100 wt% (c), and 200 ...

Microporosity within agarose gels can also control diffusion profiles that significantly affect cell viability in the medium perfusion condition. Figure 6A shows cell viability on the cross-sections of the agarose microchannel with 0–200 wt% sucrose mixtures. Cell viability in hydrogels derived from the formulation with 100 wt% sucrose (Figure 6B) was higher than that in hydrogels derived from the formulation with 0 wt% sucrose at all distances from the medium perfusion channel. Cells cultured near the microchannels showed similar cell viability (80–95%) in different sucrose mixtures. However, the viability in hydrogels derived from the formulation with 200 wt% sucrose at the 700–2200 μm distance from microchannels was 10–20% higher than that in non-porous agarose gels, because hydrogels derived from the formulation with 200 wt% sucrose contained interconnected pores that could easily deliver medium and oxygen to the cells.

Figure 6
The viability of hepatic cells exposed to continuous medium perfusion from a microchannel within agarose gels. Cells were cultured in the agarose gel microchannel for 5 days in vitro. (A) Phase contrast (a, c, e) and fluorescent images (b, d, f) of cells ...

For the static culture condition (Figure 5C), although hydrogels derived from the formulation with 200 wt% sucrose contained interconnected pores, cell viability was similar to non-porous agarose gels. In contrast, as we applied to medium perfusion in the agarose gel channel with 200 wt% sucrose, cell viability was higher than non-porous agarose gels (Figure 6B), because nutrients were easily delivered into the cells through interconnected pores. Thus, the homogeneous porosity derived from 100 wt% sucrose increased cell viability in the static culture condition, while the interconnected pores made by 200 wt% sucrose enabled the nutrient delivery into the cells in the medium perfusion condition, resulting in high cell viability. Furthermore, we found that patterns of the cell viability according to the distance away from the microchannel were corresponded to diffusion patterns generated from the microchannel (Figure 4). As compared to the shear stress in a microfabricated channel on a 2D surface, flow rate (2 μl/min) we used in this paper may not significantly affect cell viability, because the hydrogel acts as a resistance of the fluidic flow, reducing flow penetration into the gel as it has been previously reported(Mosadegh et al. 2007).

To confirm the cell viability as a function of the distance away from the microchannel and assess the effect of oxygen and waste transfer on cell viability independent of the medium components, we analyzed cell viability in PBS perfusion condition (Figure 6C). As expected, cell viability in a PBS perfusion condition was lower than the medium perfusion condition. Also, a similar trend was observed as cells closer to the channel better maintained their viability. Therefore, we demonstrated that pore sizes of agarose gels and variation of diffusion coefficient derived from the porosity played a significant role in controlling cell viability in a 3D cell-laden agarose gel device.

4. Conclusions

We developed a porous cell-laden hydrogel system with an engineered microporosity. Micropores were created by leaching sucrose crystals within cell-laden agarose gels and their distributions were controlled by varying sucrose concentrations (0–200 wt%). We controlled and optimized the solubility of sucrose crystals and gelation time to improve physiological condition via a rapid cooling process. The microporosity (0~40%) was directly proportional while mechanical stiffness was inversely proportional to sucrose concentration. The compressive modulus of hydrogels derived from the formulation with 200 wt% sucrose was lower than 15% of non-porous agarose gels. The diffusion of biomolecules in the porous gels was also analyzed as a function of the microporosity and the distance away from the microchannels. The diffusion coefficient in hydrogels derived from the formulation with 200 wt% sucrose containing interconnected pores was 1.5 times increased as compared to non-porous agarose gels. We demonstrated that microporous structures significantly affected the diffusion of biomolecules and the viability of cells cultured within microporous cell-laden agarose gels. Cell viability in the porous agarose gel microchannels (200 wt% sucrose) was 10–20% higher than in the non-porous agarose microchannels. Therefore, this approach may be potentially beneficial for engineering tissue constructs for regenerative medicine and drug discovery applications.

Acknowledgements

This paper was partly supported by the National Institutes of Health (DE019024, HL092836, and EB007249), US Army Core of Engineers, and the Charles Stark Draper Laboratory. Jae Hong Park was supported by the Korea Research Foundation Grant funded by the Korean Government (MOEHRD) (Grant Number: KRF-2007-357-D00101).

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