Development of porous PEG hydrogels that enable efficient, uniform cell-seeding and permit early neural process extension

Available online at www.sciencedirect.com

Acta Biomaterialia 5 (2009) 1884–1897 www.elsevier.com/locate/actabiomat

Development of porous PEG hydrogels that enable efficient, uniform cell-seeding and permit early neural process extension R.M. Namba a, A.A. Cole b, K.B. Bjugstad c, M.J. Mahoney d,* a

Department of Chemistry & Biochemistry, University of Colorado, ECCH 111, Boulder, CO 80309, USA b Department of Physics, University of Colorado, 390 UCB, Boulder, CO 80309, USA c Department of Pediatrics, Mail Stop F8313, University of Colorado at Denver, Aurora, CO 80045, USA d Department of Chemical & Biological Engineering, University of Colorado, ECCH 111, Boulder, CO 80309, USA Received 21 August 2008; received in revised form 14 January 2009; accepted 16 January 2009 Available online 1 February 2009

Abstract Three-dimensional polymer scaffolds are useful culture systems for neural cell growth and can provide permissive substrates that support neural processes as they extend across lesions in the brain and spinal cord. Degradable poly(ethylene) glycol (PEG) gels have been identified as a particularly promising scaffold material for this purpose; however, process extension within PEG gels is limited to late stages of hydrogel degradation. Here we demonstrate that earlier process extension can be achieved from primary neural cells encapsulated within PEG gels by creating a network of interconnected pores thoughout the gel. Our method of incorporating these pores involves co-encapsulating a cell solution and a fibrin network within a PEG gel. The fibrin is subsequently enzymatically degraded under cytocompatible conditions, leaving behind a network of interconnected pores within the PEG gel. The primary neural cell population encapsulated in the gel is of mixed composition, containing differentiated neurons, and multipotent neuronal and glial precursor cells. We demonstrate that the initial presence of fibrin does not influence the cell-fate decisions of the encapsulated precursor cells. We also demonstrate that this fabrication approach enables simple, efficient and uniform seeding of viable cells throughout the entire porous scaffold. Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. Keywords: Porosity; Biomaterials; Nerve tissue engineering; Cell encapsulation; Cell morphology

1. Introduction Three-dimensional polymer scaffolds serve as useful culture systems for neural cell growth as they provide substrates that mimic in vivo growth conditions more closely than monolayer culture [1,2]. Polymer scaffolds are also useful for improving regeneration of the damaged central nervous system (CNS) as they can both increase survival of transplanted cells [3,4] and provide permissive substrates that support the integration of neural processes as they extend across lesions in both the spinal cord and the brain [5,6]. In the absence of a permissive substrate, neural pro-

*

Corresponding author. Tel.: +1 303 492 3573; fax: +1 303 492 4341. E-mail address: [email protected] (M.J. Mahoney).

cesses are not readily able to extend long distances in the central nervous system [7,8], but previous researchers have demonstrated that neural processes more readily extend when donor cells are implanted with guidance substrates such as laminin tracks [9] or peripheral nerve segments [10,11]. These and other published works have highlighted several requirements that a supportive scaffold material should fulfill to improve regeneration in the CNS. Ideally, the material should have a low stiffness similar to that of native brain tissue in order to simultaneously encourage neuronal process growth [12–14] while discouraging astrocytic growth that could lead to glial scar formation [14–16]. In addition, the material should be biocompatible by being composed of non-immunogenic materials. Lastly, the ideal

1742-7061/$ - see front matter Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.actbio.2009.01.036

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

material should have the potential to biodegrade over time as transplanted tissue integrates into the host environment. Recently, biodegradable poly(ethylene) glycol (PEG)based hydrogels have been proposed as a particularly promising material that can potentially fulfill these requirements [17]. Neural cells can be encapsulated within three-dimensional PEG hydrogels under physiological conditions by dissolving PEG macromer in medium that contains isolated cells, and then polymerizing this mixture in the presence of a photoinitiator and ultraviolet (UV) light [17]. The stiffness of PEG hydrogels is easily controlled by altering the weight percent of PEG macromer used to form the hydrogel, and can be matched to the stiffness of brain tissue (around 260–490 Pa [18]). PEG hydrogels also have the potential to decrease the immunogenicity of the implanted tissue because PEG hydrogels are bioinert and have a very tight 0 ˚ ) that may isolate mesh size upon polymerization (70 A donor cells from the host’s immune cells [19], thus minimizing the major cause of acute neural transplant rejection [20]). Finally, degradable PEG hydrogels also have the ability to biodegrade over time as new tissue regenerates and eventually will leave no trace in the patient [21]. PEG hydrogels also provide a permissive environment through which neural processes can extend, albeit only at very late stages of hydrogel degradation after the overall mesh size of the hydrogel increases [17]. We hypothesize that processes are not able to extend earlier because the initial mesh size of the hydrogel is too small to permit process growth into the bulk of the hydrogel. It is necessary to achieve earlier process extension within PEG hydrogels in order to ensure that neural cells are capable of extending processes throughout the bulk of the hydrogel to reach targets and effectively integrate with the surrounding host tissue before the protective hydrogel scaffold degrades. In an effort to facilitate process extension within PEG hydrogels, our approach is to eliminate the physical constraints set by the small mesh-size of the PEG hydrogel by incorporating interconnected pores sized large enough for processes to extend through (1 lm in diameter). Although not the focus of the current study, these pores would still be smaller than the diameter of an immune cell (10 lm) and could provide acute immune protection upon grafting. Scaffolds containing highly interconnected porous structures have been investigated for a wide variety of tissue engineering applications [22–27], and a number of approaches have been developed for their fabrication [28]. These techniques include methods such as electrospinning [29,30], solvent casting-particulate leaching [31], gas foaming [32], fiber bonding [33], freeze-drying [34,35], phase-separation [36] and melt molding [37]. Unfortunately, these fabrication methods are not ideal for our specific purpose as they are toxic to cells and require that cells be seeded into the scaffold post-fabrication. This results in difficulties achieving a homogeneous distribution of cells throughout the scaffold. In addition, it also requires that the pores created in these scaffolds must be large enough to allow cells to pass through, a factor which may afford

1885

donor cells less protection from an acute immune response immediately following transplantation. In a recent study, a porous structure containing interconnected pores sized between 200 and 500 lm was created within a PEG hydrogel in order to encourage the growth of microvascular networks that will be critical to the success of neural grafts [38]. Their fabrication protocol involved polymerizing PEG–polylysine around a salt-leached polylactic-co-glycolic acid scaffold that was then degraded in a sodium hydroxide solution [38]. This study therefore used a two-polymer system in which one of the polymer networks was degraded in order to create a porous network. Our work describes a porous scaffold fabrication protocol based on the same premise, only with the alternative specifications that the degradation process will be cytocompatible and will result in smaller pores (1 lm). The porous scaffold fabrication technique described here is designed to fulfill several additional criteria as well. The fabrication technique needs to ensure very high cell-seeding and cell-survival efficiencies – two important criteria, given the scarcity of available donor tissue for neural cell transplantation. In addition, the hydrogel should degrade at such a rate that it remains present for a minimum time period of approximately 2 weeks. This design factor is important in order to provide the grafted cells with protection from the blood-borne immune cells that are highly active shortly after implantation [20]. It is important that the porous scaffold fabrication technique ensures that the introduction of pores does not accelerate the rate of hydrogel degradation. With these considerations in mind, we have developed an alternative method of fabricating porous scaffolds that enables simple, efficient, reproducible and uniform seeding of viable neural cells throughout the entire scaffold. Our approach involves photopolymerizing PEG in the presence of neural cells and a fibrin network. The fibrin is then enzymatically degraded away under cytocompatible conditions, leaving behind a network of interconnected pores appropriate in size to permit neural process extension and to discourage cell migration into or out of the scaffold. The primary neural cells used for this study have been shown via gene expression and immunohistochemistry assays to be capable of differentiating to form cholinergic neurons (choline acetyltransferase positive), GABAergic neurons (glutamate decarboxylase positive) or adrenergic neurons (tyrosine hydroxylase positive, dopamine-b-hydroxylase positive) (data not shown). Therefore, the cell population used in this study could be relevant for neural cell transplantation applications such as Huntington’s disease, Alzheimer’s disease or spinal chord injury, where GABAergic neurons, cholinergic neurons and adrenergic neurons, respectively, need to be replaced. A fibrin network was chosen for this study because the diameter of the protein fibrils is on the order of 1 lm. We demonstrate that when primary neural cells are cultured within these porous scaffolds, process extension occurs significantly earlier relative to non-porous PEG scaffolds.

1886

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

2. Materials and methods 2.1. Primary neural cell isolation and culture Primary neural cells were isolated from embryonic rats in compliance with institutional ethical use animal protocols. Briefly, a Cesarean section was performed on timedpregnant Sprague–Dawley rats at 14–15 days gestation (Charles River). Embryos were retrieved and decapitated, and the brains were isolated. The hindbrain, midbrain and meninges were removed to yield the forebrain. The freshly dissected forebrain tissue was then mechanically and enzymatically dissociated. The dissociated tissue was then transferred to fresh culture medium and the tissue fragments were triturated with a glass Pasteur pipette until all tissue fragments were dissociated. The single cell suspension was then strained to remove any remaining connective tissue. In a recent study, direct counting of positively labeled cells in acutely dissociated cell cultures indicates that 53 ± 4% of this cell population is positive for b-tubulin-III, an immunocytochemical marker for post-mitotic neurons. The remainder of the cell population (57 ± 3%) is positive for nestin, an intermediate filament protein found in precursor cells (no cells stained positively for glial fibrillary acidic protein, a marker of glial cells at this time) [39]. All dissection procedures were carried out in cold dissection medium. All enzymatic dissociation procedures were performed at 37 °C. The dissociated cells were encapsulated into scaffolds as described in Section 2.2 and subsequently cultured in medium consisting of a 50:50 Dulbecco’s modified Eagle’s medium (DMEM):F12 (MediaTech) supplemented with 1 N2 (Invitrogen), 100 U ml1 penicillin–streptomycin (Hyclone), 1 mM Lglutamine (Invitrogen), and 20 ng ml1 basic fibroblast growth factor (bFGF; Sigma). Culture medium was replaced every 2–3 days. Aprotinin (Sigma) was added to the culture medium at a concentration of 25 lg ml1 for select experiments. 2.2. Preparation of porous scaffolds and non-porous control scaffolds Photoreactive, degradable-lactic acid containing PEG macromers were synthesized using 4600 g mol1 PEG (Sigma) as described previously [21]. Impurities were removed via rotary evaporation and subsequent filtration, and H1 NMR was employed to assess purity, average number of added lactide units (2.55) on each end of the PEG monomer, and the methacrylation efficiency (81%) of the macromer. 2.2.1. Preparation of non-porous control scaffolds Non-porous PEG scaffolds were made by combining PEG macromer (7.5 wt.%), photoinitiator (0.025 wt.%) (Irgacure 2959, Ciba) and sterile medium containing dissociated cells (1  107 cells ml1) to make 500 ll of pre-polymerization solution. The solution was then transferred to

an optically transparent mold, which in our experiments was a 0.5 ml syringe with the tip removed. The solution was then polymerized by exposure to UV light (365 nm, 4 mW cm2) for 10 min. The polymerized scaffold cylinder was cut into 17 cross-sections 3 mm thick and 4 mm in diameter. These hydrogel discs were then cultured in medium described in Section 2.1. 2.2.2. Preparation of porous scaffolds Porous scaffolds were fabricated using the simple threestep approach shown in Fig. 1a. First, two solutions were made: (i) a fibrinogen solution and (ii) a thrombin–PEG– photoinitiator + cells solution. These two solutions were quickly mixed upon injection through a single 18-gauge needle into an optically transparent mold, wherein the fibrinogen and thrombin reacted to form a fibrin network (step 1 in Fig. 1). The fibrin was allowed to coagulate within the mold for 10 min while rotating to prevent cell settling. The PEG gel was then photopolymerized (step 2 in Fig. 1) effectively encapsulating both the fibrin network and the cells. Finally, the polymerized scaffold cylinder was cut into cross-sections 3 mm thick and 4 mm in diameter, and incubated in a solution of 1.25 U ml1 collagenase (Sigma) at 37 °C for 10 min in order to digest the fibrin network into fragments low enough in molecular weight to be released from the gel (step 3 in Fig. 1). The gels were gently mixed during this incubation step to facilitate collagenase entry into the gel and uniform degradation of fibrin throughout the entire thickness of the hydrogel disc. Collagenase activity was then inhibited by incubation for 10 min in medium supplemented with 10 mg ml1 bovine serum albumin and 10 mg ml1 trypsin inhibitor (no collagenase is present in this inhibitor solution). Finally, gels were rinsed in phosphate-buffered saline (PBS; MediaTech) then cultured in medium as described in Section 2.1. The fibrinogen and thrombin materials used to make the fibrin network were generously provided by Baxter Biosciences. Unless otherwise stated, the final fibrinogen and thrombin concentrations were 5 mg ml1 and 1 U ml1 respectively. For select experiments, the fibrinogen and thrombin concentrations were increased to 25 mg ml1 and 125 U ml1 respectively. Final concentrations of PEG macromer and photoinitiator in the porous scaffolds were 7.5 and 0.025 wt.% respectively. 2.3. Quantification of the amount of fibrin remaining in the scaffold after enzyme treatment To monitor the release of fibrin fragments from the scaffolds following the 10 min treatment with collagenase, porous scaffolds were made as described above (Section 2.2) with the additional incorporation of fibrinogen Alexafluor-546 (Invitrogen) (50:1 unconjugated:conjugated fibrinogen) into the fibrinogen solution. At select time intervals after fabrication, n = 12 scaffolds from three different experiments were collected and individually homogenized in 200 ll of PBS using a disposable pestle (Kimble-Kontes).

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

1887

Fig. 1. Porous scaffold fabrication. (a) Porous scaffolds were fabricated using a simple three-step approach. First, a fibrinogen solution was mixed with a thrombin–PEG–photoinitiator–cell solution upon injection through a single needle into an optically transparent mold, wherein the fibrinogen and thrombin reacted to form a fibrin network (step 1). The filled mold was then placed under UV light in order to induce photopolymerization of the PEG gel (step 2). Finally, the polymerized scaffold was ejected from the mold, sliced and incubated in an enzyme solution (for 10 min) in order to digest the fibrin network (step 3). (b) After this 10 min incubation, the diffusion of fluorescent fibrin fragments from the scaffold was monitored using a spectrophotometer until they were no longer detectable.

The amount of fluorescent fibrin present in each homogenized gel solution was quantified by measuring the fluorescence intensity of the solution using a spectrofluorometer (Fluostar Optima) with 544 nm excitation and 590 nm emission wavelengths. The fluorescence values for fibrinloaded PEG scaffolds that were not treated with enzyme were also quantified at select time points as positive controls. In addition, enzyme-treated PEG scaffolds containing no fibrin were quantified to obtain blank fluorescence values. The change in fluorescence intensity over time for each gel condition is expressed relative to the initial fluorescence intensity of the gel at t = 0 minus the blank (Fig. 1b).The detection limit for this assay was determined to be approximately 25 lg ml1 by measuring the resulting fluorescence when known concentrations of fluorescent fibrin were encapsulated into n = 5 gels (data not shown). 2.4. Estimation of the diameter and density of entrapped fibrils The structure of an encapsulated fluorescent fibrin network before enzymatic degradation was visualized via confocal microscopy (Zeiss) using a 40 water-immersion objective with a 280 nm lateral resolution limit and a 1.2 lm axial resolution limit. The average fibril diameter was estimated using ImageJ analysis software by manually measuring the diameter of n = 100 fibrils from three different images of entrapped fibrin networks made with 1 U ml 1 thrombin and either 5 or 25 mg ml–1 fibrinogen. The images were taken on a single x–y plane in order to ensure that the measured fibril diameters were well above the lateral resolution limit. Because the entrapped fibrin network resulted in a three-dimensional (3-D) network of interconnected fibrils, the density of the fibrils was estimated by determining the total cross-sectional area in each image

occupied by fibrils using ImageJ analysis software. Three different images were analyzed to obtain the average fibril density for fibrin networks made from 5 mg ml1 fibrinogen and either 1 or 125 U ml1 thrombin. Data are presented as mean ± SE (Fig. 2). 2.5. Estimation of the diameter and density of entrapped pores To ensure that enzymatically removing the fibrin network actually created a network of pores within the PEG scaffold, the structure of a porous scaffold was visualized using low vacuum scanning electron microscopy (LV-SEM). Porous scaffolds were made as described in Section 2.2 except the weight percent of PEG macromer was increased to 25 wt.% in order to better maintain the spatial dimensions of the porous scaffold during dehydration. The porous scaffolds were dialyzed in dH2O to remove residual salts and then dehydrated over night in a vacuum chamber (Fisher Scientific, model 280A) at room temperature and 12 in. Hg. The dehydrated scaffolds were mounted on aluminum stubs using adhesive carbon tape and visualized using a low vacuum scanning electron microscope (Evisa, JSM-6480LV). The average pore diameter was estimated using ImageJ analysis software by manually measuring the diameter of n = 100 pores from 3 different images of porous networks made with 1 U ml1 thrombin and either 5 or 25 mg ml1 fibrinogen. The density of the resulting pores was determined by manually counting the number of pores present in n = 9 superimposed boxes with an area of 100 lm2 box1. Three different images were analyzed to obtain the average pore density for fibrin networks made from 5 mg ml1 fibrinogen and either 1 or 125 U ml1 thrombin. Data are presented as mean ± SE (Fig. 2).

1888

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

Fig. 2. Porous scaffold characterization. Confocal microscopy was used to determine the structure of the fluorescent fibrin network and SEM was used to determine the structure of the resulting porous scaffold. Fibril and pore diameter and density were manually measured using ImageJ analysis software. (a) Increasing the concentration of fibrinogen in the initial fibrin network from 5 to 25 mg ml–1 increased both the diameter of the fibrin fibrils and the diameter of the pores in the resulting scaffold. Thrombin concentration was held constant at 1 U ml1. Asterisks indicate a statistically significant difference in measured pore diameters relative to the measured fibril diameters. (b) Increasing the concentration of thrombin in the initial fibrin network from 1 to 125 U ml1 increased both the density of the fibrin fibrils and the density of the pores in the resulting scaffold. Fibrinogen concentration was held constant at 5 mg ml1. (c–h) Representative images of (c) the no fibrin controls; (d) the no pore controls; (e, g) the initial red fibrin networks; and (f, h) the resulting black pore structures as the thrombin concentration is increased from 1 U ml–1 thrombin (e, f) to 125 U ml1 thrombin (g, h). Large and small arrows point to representative fibrils and pores respectively.

2.6. Measurement of cell viability within scaffolds

2.7. Measurement of cell proliferation within scaffolds

The impact of the enzyme treatment on cell viability was assessed using a standard MTT assay. First, cells were encapsulated within non-porous control scaffolds. Next, n = 5 scaffolds were incubated at 37 °C for 45 min in enzyme-containing medium with collagenase concentrations of 0, 5, 10, 15 and 25 U ml–1. The scaffolds were then separately incubated with 1 ml of MTT medium composed of 0.5 mg ml–1 MTT (3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide) (Sigma) and phenol-red free DMEM (Gibco). Samples were incubated at 37 °C for 4 h, during which time the MTT was reduced to purple formazan by reductases active only in the mitochondria of living cells. A 0.1 M HCl solubilization solution was then added to dissolve the purple formazan product into a colored solution. The absorbance of this colored solution was quantified using a spectrophotometer. Data represent the average measured absorbance value normalized with respect to the untreated control (Fig. 3a).

The influence of fibrin on the proliferative capacity of cells in hydrogels was assessed by measuring total DNA content on days 0 and 7 in cell-laden hydrogels filled with a fibrin network. The fibrin-loaded scaffolds were prepared with 1 U ml1 thrombin and either 0, 0.2, 1 or 5 mg ml1 fibrin. The goal of this experiment was to identify any possible influence that fibrin exposure might have on neural cell proliferation. Understanding what role fibrin exposure may have on neural cell proliferation will help to determine whether undetectable, but biologically active levels of fibrin are present in porous scaffolds. After establishing a proliferative effect of fibrin on neural cells, the amount of DNA present in porous scaffolds was measured immediately after forming the porous scaffold (day 0) and then again on day 7. This value was directly compared to the amount of DNA present on days 0 and 7 in control hydrogels formed in the absence of a fibrin network. For each experiment, the percent increase in total DNA content from day 0 to day 7 was

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

1889

Fig. 3. Influence of porous scaffold fabrication on cell behavior. (a) Cell viability was assessed after a 45 min incubation in medium containing increasing concentrations of collagenase using the MTT cellular toxicity assay as a measure of the number of living cells present in the scaffolds. The asterisk indicates a statistically significant difference relative to the untreated control. (b) The proliferative effect of fibrin is demonstrated in PEG scaffolds when neural cells are cultures in scaffolds that contain fibrin concentrations greater than 0.2 mg ml1. (c) No changes in proliferation were observed when cells were cultured in the porous scaffolds when compared to non-porous controls. Here, proliferation is defined as the percent increase in [DNA] from day 0 to day 7. (d) Cell composition was assessed by qRT-PCR analysis of nestin, b-tubulin or GFAP mRNA expression and is expressed as gene expression in porous scaffolds normalized with respect to gene expression in non-porous control scaffolds.

taken as an indicator of cell proliferation. At each time point, n=10 scaffolds from two different experiments were transferred to Eppendorf tubes containing 300 ll of cell lysis buffer. Each scaffold was homogenized using a disposable pestle until no visible gel fragments remained. The homogenized solution was then sonicated (Misonix) for 15 s at 3 W on ice to ensure complete cell lysis. This homogenate solution contained a DNA concentration that is directly proportional to the number of cells within the scaffold. DNA concentration was measured according to the manufacturer’s instructions using the PicoGreen reagent (Invitrogen). Data represent the average percent increase in DNA concentration from day 0 to day 7 (Fig. 3b and c). 2.8. Assessment of cell composition within scaffolds The relative abundance of cell-specific genes present in gel cultures was assessed on both day 0 and upon hydrogel degradation by quantifying the expression of cell-type-specific mRNA transcripts in both porous scaffolds (n = 6) and non-porous control scaffolds (n = 6) obtained from two different experiments. Neural precursor cells (NPCs) were identified by nestin expression, neurons with b-tubulin (III) expression and glial cells with glial fibrillary acidic

protein (GFAP) expression [39]. Primers were obtained from invitrogen and the sequences are as follows (50 –30 ): b-tubulin (fwd-gtttgtgatgggtgtgaacc, rev-tcttctgagtggcagt gatg); GFAP (fwd-agggacaatctcacacagg, rev-gactcaaccttc ctctcca); nestin (fwd-gtggcctctgggatgatg, rev-ttgaccttcctcc ccctc). Standard phenol:chloform RNA extraction was performed using TriReagent (Sigma). Isolated RNA was treated with DNAse (Ambion) and accurately quantified using RiboGreen Quantification Reagent (Invitrogen). Equal amounts of RNA (100 ng) were placed in each reverse transcription reaction, thus alleviating the need for the inclusion of house-keeping genes [40]. cDNA synthesis and SYBRgreen polymerase chain reaction (PCR) was carried out according to manufacturer’s recommendation (Biorad). Data represent the average fold change in gene expression relative to day 0 and are normalized with respect to the non-porous control scaffolds. Data are presented as the mean ± SE. 2.9. Characterization of encapsulation efficiency To evaluate the encapsulation efficiency within porous scaffolds, cells were entrapped within 0.5 ml of either porous or non-porous control scaffolds at a concentration of

1890

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

1  107 cells ml1, as described in Section 2.2. After 24 h in culture within 4 ml of medium, the scaffolds were removed and the medium was collected to determine the number of loose, unencapsulated cells. Media collection involved trituration to remove cells adhered on the bottom of the culture wells and then centrifugation at 1000 rpm for 10 min to pellet the cells. Each cell pellet was resuspended in 50 ll of medium. Next, 90 ll of trypan blue (MediaTech) was added to the cell solution, and the number of unencapsulated cells was determined using a hemocytometer. The number of unencapsulated cells was then compared to the number of cells that was intended for encapsulation in the pre-polymerization solution, and the percent encapsulation was calculated as follows: %Encapsulation " ¼ 100 1 

ð# Loose cells on Day 1=mlÞð0:14 mlÞ ð1  107 cells=mlÞð0:5 mlÞ

!#

ð1Þ The average encapsulation efficiency was determined after quantifying unencapsulated cells in n = 8 wells from two different experiments. 2.10. Characterization of cell distribution and morphology Cells that had been cultured within non-porous control scaffolds and porous scaffolds were imaged via confocal microscopy to determine the distribution and morphology of live cells within the scaffolds. Live cells were labeled with calcein-AM (Invitrogen), which was determined to fully label both the cell bodies and extending processes of the neural cells. This was determined by plating neural cells in monolayer and labeling them with calcein-AM, then capturing both bright-field images as well as fluorescent images. These two image sets overlay completely (data not shown). In select experiments, aprotinin was added to the culture medium. To verify that aprotinin remains active when added to hydrogel cultures, we added aprotinin to PEG hydrogels containing a fibrin network. Process extension in fibrin-containing PEG hydrogels relies on cell-secreted protease degradation of the fibrin fibrils at early time points, and this early process extension was completely inhibited in the presence of aprotinin (data not shown). 2.10.1. Cell distribution images Cell distribution images were taken after 7 days of culture using a 10 water-immersion objective. Each analyzed image is a projection 100 lm thick compiled from 10 optical slices acquired along the z-axis. Images were acquired from three different gel tridrants (A, B, C) and at three different gel depths (surface, 1.5 mm and 3 mm) (see Fig. 3b). ImageJ analysis software was used to determine number of cells present per image on n = 9 different gels per tridrant and n = 9 different gels per gel depth. The concentration of cells in each image was determined by utilizing the Ima-

geJ software to calculate the total area of each image occupied by both single cells and cell clusters. The total area occupied by cells in each image was then divided by the average measured area of a single cell to determine the approximate number of cells present in each image. The volume of gel that was used to create each image was calculated based on the x, y and z dimensions of the image. 2.10.2. Cell morphology images Over the culture period, cells divide to form small aggregates of tissue with neural processes emerging from the aggregate core. Confocal microscopy was used to image each aggregate and associated processes. Images were taken at select time points using a 40 water-immersion objective. Each image is a projection spanning the entire thickness of the aggregate and was compiled from optical slices acquired every 1 lm along the z-axis. To determine the number of neural processes per aggregate, 15–50 different images from four different experiments were acquired for each time point for both porous and non-porous scaffolds. The mean number of processes per aggregate was determined by manually counting the number of processes in each image that extended at least 5 lm from the perimeter of the cell aggregate. At each time point, the data was verified to be normally distributed. The minimum sample size was calculated to be n=15 to acquire data that would ensure 90% confidence that the sample mean was within 5 processes per aggregate of the actual population mean. Images were also acquired on the ‘‘day of gel degradation”, which is the day before the hydrogel completely dissolves due to the hydrolysis of the degradable PEG network. Non-porous control scaffolds consistently degraded in week 3 of culture, while the porous scaffolds degraded in week 4 of culture. The images acquired on ‘‘day of degradation” were used to determine the average diameter of a single process extending from the cell cluster, and the average number of branch points per single process. The number of branch points per process was manually counted for every image, and the process diameter was manually measured at the thickest point on every process using Zeiss LSM image analysis software. 2.11. Statistical analysis For all data, statistical significance was determined using two-tailed Student’s t-test, with p < 0.05 considered significant. Statistically significant differences are denoted with an asterisk. Data are presented as mean ± standard deviation unless otherwise indicated. 3. Results 3.1. Estimation of size and density of pores in porous scaffolds Our porous scaffold fabrication approach involves encapsulating a fibrin network within a PEG hydrogel,

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

then enzymatically digesting the fibrin to reveal a porous PEG scaffold. A fibrin network made from 5 mg ml1 fibrinogen and 1 U ml1 thrombin was chosen for all experiments presented here that involve neural cells for the reasons explained below. A representative image of this encapsulated fibrin network and the resulting porous structure is shown in Fig. 2e and f, respectively. The initial fibrin fibrils were measured to be 0.7 ± 0.1 lm in diameter on average and the resulting pores were measured to be 1.1 ± 0.2 lm in diameter on average (Fig. 2a). The pores were measured in samples that had been dehydrated to allow for SEM visualization. Due to the high water content of PEG hydrogels, the scaffolds shrunk during the dehydration process by 68% in the x, y and z dimensions. When this shrinkage is taken into account, the average diameter of the pores in the hydrated scaffolds is estimated to be 1.6 ± 0.3 lm. It is not surprising that the measured pore diameters are slightly larger than the measured fibril diameters because the fibrils can inhibit the local polymerization of PEG [41]. These fibrils and pores were not observed in the control scaffolds (Fig. 2c and d). The fibrin network occupied 32 ± 2% of the total cross-sectional area of images taken of the entrapped fibrin network and resulted in a density of approximately 3 pores per 100 lm2 area in the SEM images of dehydrated scaffolds (Fig. 2b). When scaffold shrinkage is taken into account, it is expected that the average pore density in hydrated samples would be approximately 2 pores per 100 lm2. Given that the average diameter of a cell is approximately 10 lm, only one cell would be expected to occupy any given 100 lm2 area early in hydrogel culture. This means that each cell would be supplied with at least 2 pores when the fibrin network is made from 5 mg ml1 fibrinogen and 1 U ml1 thrombin. These results confirm that the porous scaffolds used in the experiments presented here contain an ample number of pores approximately 1.6 lm in diameter distributed throughout the scaffold to permit early process extension to occur. The porous structure that results after degrading the fibrin from the PEG gel can be easily modified by altering the structure of the initial fibrin network, which can be controlled by varying the concentrations of fibrinogen and thrombin used to form the network. For example, the diameter of the fibrin fibrils and resulting pores can be controlled by altering the concentration of fibrinogen used to make the initial fibrin network. When the concentration of fibrinogen used to make the fibrin network was increased to 25 mg ml1, the diameter of the fibrin fibrils increased to 1.4 ± 0.2 lm and the diameter of the resulting pores increased to 1.9 ± 0.2 lm (Fig. 2a) in dehydrated SEM images. This corresponds to a 2.8 ± 0.4 lm pore size in hydrated scaffolds. In addition, the density of the fibrin fibrils and resulting pores can be controlled by altering the concentration of thrombin used to make the initial fibrin network. Increasing the concentration of thrombin to 125 U ml1 resulted in a fibrin network that occupied 52 ± 3% of the total cross-sectional area of acquired

1891

images, and resulted in a scaffold with approximately 5 pores per 100 lm2 area in dehydrated SEM images (Fig. 2b, g and h). This corresponds to approximately 3.5 pores per 100 lm2 area in hydrated scaffolds. 3.2. Influence of the porous scaffold fabrication protocol on cell behavior We have previously demonstrated that high cell viability is possible when neural cells are encapsulated and cultured within non-porous PEG scaffolds [17], so it was necessary to ensure that the enzyme treatment used in the porous scaffold fabrication protocol did not impair the survival of encapsulated cells. To directly assess how vulnerable the cells are to enzyme treatment, cells were encapsulated within non-porous scaffolds and incubated in the presence of increasing concentrations of enzyme (Fig. 3a). The resulting viability of the enzyme-treated cells was then determined using a standard MTT cell viability assay. The results demonstrate that incubating cells within an enzyme solution at a concentration of 25 U ml1 negatively affects cell viability, as the MTT absorbance value is only 55 ± 9% of the non-treated control (Fig. 3a). However, when the enzyme concentration is decreased to 5 U ml1, cell viability is not significantly (p > 0.05) different than the untreated control, with MTT absorbance values at 92 ± 17% of the untreated control (Fig. 3a). The porous scaffold fabrication protocol described above was therefore specifically designed to use enzyme concentrations less then 5 U ml1 (1.25 U ml1 was chosen) to ensure that cell viability was not significantly impaired, while also ensuring that the enzyme concentration was high enough to efficiently remove the fibrin network. We also performed a monolayer culture experiment to verify that exposure of the neural cell population to collagenase does not impair the cell’s ability to extend processes, an effect that would confound interpretation of the experimental results presented in this paper. When cells are incubated in a solution of collagenase (1.25 U ml1) for 10 min and then plated on a monolayer surface, process extension occurs over the course of 7 days in a manner identical to that which occurs in the absence of collagenase (data not shown). Though Fig. 1b demonstrates that an enzyme concentration of 1.25 U ml1 efficiently removes all detectable fibrin from the scaffold, it was nevertheless essential to validate that any remaining undetectable fibrin did not influence the long-term behavior of the encapsulated neural cells. In addition, it is possible that changes in cell behavior might be observed, given that cells were exposed to fibrin during the first 24 h as fibrin fragments were released from the scaffold. To make certain that the porous scaffold fabrication protocol did not influence encapsulated cells in this way, cell proliferation and differentiation in both porous scaffolds and non-porous control scaffolds were directly compared on day 7 of culture. To evaluate cell proliferation within the scaffolds, DNA content was assessed after culturing neural cells for 7 days

1892

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

Fig. 4. Cell seeding within a porous scaffold. (a) Cell encapsulation efficiency was determined by counting unencapsulated cells after 24 h of culture. (b–e) Live cells were evenly distributed throughout the gel with dimensions shown in (b). (c) On day 7, confocal microscopy was employed to image the distribution of live cells labeled with calcein-AM in 0.1 mm optical sections at three different depths along the z-axis of the gel. The number of live cells at each gel depth were counted using ImageJ analysis software. (d) Additionally, confocal microscopy was employed to image the distribution of live cells within the x/y-axis by counting live cells present within 0.1 mm optical sections obtained from three different tridrants (A, B, C) of a single plane with approximate locations shown in (b). (e) A representative image of the cell distribution on day 1 of culture in a 0.1 mm optical gel slice.

in porous scaffolds, non-porous negative control scaffolds or fibrin-loaded positive control scaffolds. A proliferative effect of fibrin was observed within PEG scaffolds when fibrin concentrations of greater than 0.2 mg ml1 were encapsulated in PEG scaffolds (Fig. 3b). Thus, it was important to verify that any fibrin that remained in the porous scaffolds would not influence the proliferation of the encapsulated cells. When cells were encapsulated in non-porous control scaffolds, a 45 ± 8% increase in total DNA content was observed over time (Fig. 3b). This increase is consistent with previous observations, and has been associated with the proliferation of neural precursor cells [17]. When cells were encapsulated in porous scaffolds, a similar increase (42 ± 7%) in total DNA content was observed (Fig. 3b). Therefore, total DNA content in porous scaffolds was not significantly (p > 0.05) different from that in non-porous control scaffolds, a finding that suggests that cell exposure to fibrin and/or fibrin fragments does not significantly impact cell proliferation after 1 week of culture. It was also important to ensure that the cell-fate decisions of the neural precursor cells were not affected by exposing the cells to fibrin during the porous scaffold fabrication protocol. We have previously shown that when the cell population used in this study is encapsulated within non-porous control

scaffolds, a fraction of the cells differentiate over time into glial cells and neurons, while the remainder are maintained as undifferentiated neural precursor cells [17]. In order to evaluate the relative cell composition in both the porous and control scaffolds, qRT-PCR was used to assess the expression of cell-type specific genes. If significant changes in the expression of nestin, b-tubulin or GFAP were observed, this would indicate that the fibrin exposure influenced the fraction of neural precursor cells, neurons or glial cells respectively that were present in the porous scaffolds relative to the control scaffolds. The results show that exposing the cells encapsulated within the porous scaffolds to fibrin during the fabrication protocol did not significantly alter the expression of nestin (p > 0.05), b-tubulin (p > 0.05) or GFAP (p > 0.05) (Fig. 3c) relative to the non-porous control scaffolds after 3–4 weeks of culture. 3.3. Cell seeding in porous scaffolds The simple approach to fabricating porous scaffolds described here may be able to address some limitations of other porous scaffold fabrication protocols pertaining to low seeding efficiencies and uneven cell distributions. Results in Fig. 4a demonstrate that the porous scaffold fabrication approach described here enabled high encapsula-

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

1893

Fig. 5. Neural process extension within porous scaffolds. Cells were encapsulated and cultured in either porous scaffolds or non-porous control scaffolds and then labeled using calcein-AM, which fully labels the cell bodies and extending processes of neural cells. The cells were then visualized using confocal microscopy to assess the morphology of live cells over time. (a) The average number of processes per aggregate was manually determined at select time intervals in both non-porous control scaffolds and porous scaffolds. Asterisks indicate statistically significant differences relative to time-matched nonporous control values. (b) Upon scaffold degradation, the process diameter and number of branch points present on a single process were measured in both non-porous control scaffolds and porous scaffolds. The results are plotted as a percent increase relative to the non-porous control scaffold. (c) Cell morphology was also monitored in porous scaffolds cultured in the presence (gray squares) and absence (black diamonds) of aprotinin. The number of processes per aggregate was counted at select time intervals in each condition. (d–k) Representative images of the cell aggregates growing in both porous and non-porous scaffolds respectively after 1 day (d, e), 3 days (f, g), 13 days (h, i) and when the scaffold was largely degraded as a result of hydrolysis (j, k).

tion efficiency (99%), as assessed at 24 h. This fabrication approach also easily enabled an even distribution of cells in all three dimensions of the porous scaffolds (Fig. 4b–d) as assessed quantitatively on day 7 of culture. Fig. 4e demonstrates a representative image of the cell distribution observed on day 1.

3.4. Neural process extension in porous scaffolds By providing micron-scale pores that are similarly sized to processes that extend from neural cells, it was expected that cells encapsulated within porous scaffolds would be able to achieve earlier process extension relative to cells

1894

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

encapsulated within non-porous control scaffolds. This hypothesis was tested by encapsulating neural cells in both porous and non-porous scaffolds and monitoring the resulting process extension using confocal microscopy which enabled the direct visualization of cell morphology within the scaffold (Fig. 5d–k). The results demonstrate that cells encapsulated in porous scaffolds were able to extend processes earlier (by day 3 in culture) than cells encapsulated in non-porous control scaffolds (week 2) (Fig. 5a, f and g). In addition, at every subsequent time point thereafter, more processes were present in the porous scaffolds than in the non-porous scaffolds (Fig. 5a, h–k). Interestingly, the porous scaffolds consistently degraded at a slower rate (1 week slower) than the non-porous scaffolds and therefore allowed more time for process extension and maturation to occur before complete scaffold degradation. The slower degradation rate of the porous scaffolds can be attributed to our fabrication protocol, as incorporating the fibrin network into the pre-polymerization solution effectively concentrates the PEG present in the porous scaffolds. The end result is that when the scaffolds degrade due to hydrolysis of the degradable PEG network, neural cells cultured within porous scaffolds were able to extend roughly two times more processes per aggregate of cells (Fig. 5a, j and k) than when cells were cultured in non-porous control scaffolds. In addition, by the time the hydrogel degrades, the single processes (no bundled processes were observed) that extend from the cell aggregates in porous scaffolds demonstrate a 40% increase in process diameter and an 80% increase in the number of branch points relative to the processes that extend from cells cultured within the non-porous control scaffolds (Fig. 5b, h and i). 3.5. Necessity of provided pores for achieving earlier process extension Having demonstrated that earlier process extension is achieved when neural cells are cultured within porous scaffolds, it was important to verify that the cells were extending processes into pores that had been cleared of fibrin during the fabrication of the porous scaffold. Alternatively, it is feasible that low, undetectable levels of fibrin fragments remain after enzyme treatment, even though the fluorescence assay (Fig. 1b) and biological activity assays (Fig. 3c and d) suggest that all fibrin is cleared from the gel. If this is the case, the observed early process extension may be the result of cells clearing their own path for process extension by secreting proteases that degrade the fibrin remnants. To determine whether or not cell-derived protease activity is required in order to observe early process extension, porous scaffolds were cultured both with and without the presence of a protease inhibitor, aprotinin. The resulting process extension was examined via confocal microscopy over time. The results show that even in the presence of the protease inhibitor, neural cells encapsulated within porous scaffolds were able to extend processes by

day 3 (Fig. 5c). This finding indicates that cell-derived protease activity is not necessary to achieve earlier process extension within the porous scaffolds, and provides evidence that the pores used to achieve earlier process extension are actually created during the scaffold fabrication process. In contrast, when the fibrin network was not enzymatically removed prior to culturing in the presence of aprotinin, the fibrin effectively blocked any early process extension (data not shown). The processes presumably were unable to penetrate through the fibrin when their cell-secreted proteases were inactivated with aprotinin. 4. Discussion The research presented here describes an innovative method of fabricating porous scaffolds. The traditional fabrication approach involves first fabricating a porous scaffold, then seeding cells within the pre-made scaffold. This conventional approach has resulted in reported difficulties achieving efficient and uniform seeding of viable cells throughout the scaffold [28,42–45]. The alternative method described here instead uses cytocompatible fabrication conditions in which cells can simply be encapsulated during the scaffold fabrication process. The idea of incorporating cells during the fabrication of porous scaffolds was previously introduced by researchers utilizing 3-D-cell printing and microintegration techniques to fabricate porous scaffolds [28]. Unfortunately, both of these technologies are still in the early developmental stage and require specialized instrumentation [23]. In contrast, the fabrication approach described here represents an important advance for researchers interested in utilizing porous scaffolds for tissue engineering purposes because our approach utilizes simple, inexpensive and easily accessible technology; and provides efficient (99%) and uniform seeding of viable cells throughout the entire scaffold. Our method involves photopolymerizing PEG in the presence of neural cells and a fibrin network, then enzymatically degrading away the fibrin under cytocompatible conditions to leave behind a network of interconnected pores. This fabrication method is modeled on previous work in which porosity was imparted upon a two-polymer system by preferentially degrading one polymer via hydrolytic means [46]. Other researchers have used concepts similar to this approach in order to selectively degrade entrapped polymers enzymatically [47,48]. The experiments reported here additionally incorporate cells into the two-polymer system (PEG and fibrin) prior to enzymatic degradation of the entrapped fibrin. We provide evidence that the fibrin can be enzymatically degraded from the PEG hydrogel without damaging cell viability by showing that the metabolic activity of cells encapsulated within porous hydrogels is not significantly different to the metabolic activity of cells encapsulated within non-porous control scaffolds that sustained no enzyme treatment. In previous work, we have shown that metabolic activity can be used as a quantitative indicator of cell viability, and that cells encapsulated within

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

non-porous control scaffolds maintain metabolic activity levels comparable to a population of cells that is 97% viable for at least 16 days as assessed by trypan blue exclusion [17]. We also demonstrate that exposing the entrapped cells to fibrin during the first 24 h as the fibrin fragments are released from the gel does not significantly influence cell behavior. Specifically, we have demonstrated that cell proliferation is not affected by exposure to fibrin. In addition, the assessment of cell-type specific gene expression further indicates that fibrin exposure has no effect on the cell fate decisions of encapsulated precursor cells as there were no significant differences found between the composition of cells encapsulated in non-porous control scaffolds and those encapsulated in the porous scaffolds 3 weeks after encapsulation. The details of this protocol can easily be modified by other researchers who are interested in using synthetic porous scaffolds for a wide range of tissue engineering applications. For example, though our study focuses on pore sizes on the order of 1 lm, we have demonstrated that other pore sizes and pore densities are obtainable by altering the thickness and density of the fibrils used as the pore template. Likewise, though this study describes the use of an interconnected fibrin network as the pore template, oriented channel structures could be achieved by aligning fibrin or collagen fibrils prior to encapsulation [49,50]. Other researchers may be interested in obtaining porous structures with interconnected spherical pores and could create such structures using an entrapped template of spherical microparticles composed of polymerized protein that could subsequently be enzymatically degraded [51–52]. A potentially bioactive protein could be avoided altogether by degrading entrapped synthetic polymers that contain enzymatically degradable peptide sequences [53]. The primary objective of this work was to achieve earlier process extension in PEG hydrogels. We hypothesized that neural processes in the non-porous control scaffolds were sterically inhibited from extending, given the very small 0 ˚ ) of PEG hydrogels upon polymeraverage mesh size (70 A ization [17]. Neural cells encapsulated within control scaffolds are unable to extend processes until the second week of culture, when the mesh size has increased. Interestingly, it has been determined that the mesh0 size after about ˚ , which should 2 weeks of culture is approximately 150 A still be small enough to sterically prohibit neural process extension [17]. This paradox is also apparent when considering the quantifiable and visually apparent proliferation of cells encapsulated within PEG hydrogels despite their being imbedded within a hydrogel that has a mesh size small enough to prohibit cell proliferation. One explanation may be that considerable heterogeneity in the local crosslinking density exists within a given area of a PEG hydrogel [54]. It is possible that these heterogeneities in crosslinking density may be especially profound when gelation occurs in the presence of cells. Cells would be able to contribute to these heterogeneities by either passively or actively terminating local radicals via the production of

1895

antioxidants [54]. It is also possible that cells secrete esterases that are able to cleave the lactide units present in degradable PEG gels, thus creating room for proliferation and/or process extension. Further studies are needed to determine how the cells proliferate and extend processes within unmodified PEG hydrogels despite the small overall average mesh size of these gels. Nonetheless, this work demonstrates that, when primary neural cells are cultured within porous scaffolds, neural processes begin to extend earlier than when cultured in control scaffolds. Processes may be able to extend earlier in the porous scaffolds because their extension is no longer physically inhibited by the small mesh size of the hydrogel and they now have space to extend. This possibility is validated by previous research using alginate gels showing that process growth can be inhibited if the average gel mesh size is too narrow [55]. Another potential explanation for the earlier process extension that is observed in the porous scaffolds is that the porous network is providing a distribution of 2-D surfaces within the 3-D PEG gel. It has been previously proposed that such an approach may be a useful way to achieve more extensive neural process extension within 3D matrices, given the observed preference of processes to extend along entrapped 2-D surfaces rather than through 3-D matrices [56]. Incorporating an interconnected porous network within the 3-D PEG matrix would effectively increase the 2-D surface area available for cell adhesion as processes are extending. It is also feasible that the neural processes extend earlier when cultured in porous scaffolds because undetectable fibrin fragments that remain in the hydrogel act as adhesive contacts that assist the extension of neural processes [57]. Our data cannot exclude the possibility that increased 2-D surface area or adhesive contacts contribute to the earlier process extension observed in our porous scaffolds, and these may be useful design parameters for other researchers interesting in improving process extension within 3-D matrices. Achieving earlier process extension in PEG hydrogels is a critical step towards improving the suitability of degradable PEG hydrogels as a material capable of regenerating lesioned areas of the CNS. Our results show that, relative to non-porous PEG hydrogels, porous PEG hydrogels permit neural cells to extend processes earlier and allow more time for the processes to mature before the hydrogel degrades, as evidenced by the observed increase in two indicators of in vitro neurite maturation [58,59]: the neural process diameter and the number of branch points present on single processes upon gel degradation. Together, the results described here will contribute to the development of improved PEG hydrogel scaffolds for treating multiple diseases and injuries in the CNS. 5. Conclusions In conclusion, this work describes an alternative porous scaffold fabrication method involving co-encapsulating a

1896

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897

solution of neural cells and a fibrin network within a PEG gel. The fibrin is subsequently enzymatically degraded under cytocompatible conditions, leaving behind a network of interconnected pores within the PEG gel. We demonstrate that the initial presence of fibrin does not influence the proliferation or differentiation of the encapsulated precursor cells. We also demonstrate that incorporating pores into PEG hydrogels is an effective approach to achieve earlier and more extensive process extension from encapsulated neural cells. This work is significant in that it describes the implementation of a broadly applicable concept for the fabrication of porous scaffolds that avoids the seeding efficiency and uniformity difficulties that occur when seeding cells into pre-made porous scaffolds. Acknowledgements We thank the National Institutes of Health for funding this research (R01 NS052597-02). We are also greatly appreciative to Bill Tawil from Baxter’s Bioscience division for supplying fibrin kits. Also, we would like to thank Dr. Linda Watkins for sharing the qRT-PCR primer sequences with us. References [1] Bard JB, Hav ED. The behavior of fibroblasts from the developing avian cornea. Morphology and movement in situ and in vitro. J Cell Biol 1975;67:400–18. [2] Discher DE, Janmey P, Wang YL. Tissue cells feel and respond to the stiffness of their substrate. Science 2005;310:1139–43. [3] Teng YD, Lavik EB, Qu X, Park KI, Ourednik J, Zurakowski D, et al. Functional recovery following traumatic spinal cord injury mediated by a unique polymer scaffold seeded with neural stem cells. Proc Natl Acad Sci 2002;99:3024–9. [4] Lavik EB, Klassen H, Warfvinge K, Langer R, Young MJ. Fabrication of degradable polymer scaffolds to direct the integration and differentiation of retinal progenitors. Biomaterials 2005;26:3187–96. [5] Geller HM, Fawcett JW. Building a bridge: engineering spinal cord repair. Exp Neurol 2002;174:125–36. [6] Duconseille E, Cressant A, Kelche C, Woerly S, Will B, Poucet B, et al. Homotopic septal grafts combined with a hydrogel bridge to promote functional recovery in rats with fimbria–fornix lesions: a unit recording study. Restor Neurol Neurosci 1999;15:305–17. [7] Bjorkland A, Stenevi U, Schmidt RH, Dunnett SB, Gage FH. Intracerebral grafting of neuronal cell suspensions. II. Survival and growth of nigral cell suspensions implanted in different brain sites. Acta Physiol Scand 1983;522:9–18. [8] Fawcett JW, Asher RA. The glial scar and central nervous system repair. Brain Res Bull 1999;49:377–91. [9] Yoshii S, Yamamuro T, Ito S, Hayashi M. In vivo guidance of regenerating nerve by laminin-coated filaments. Exp Neurol 1987;96:469–73. [10] Gage FH, Stenevi U, Carlstedt T, Foster G, Bjorkland A, Aguayo AJ. Anatomical and functional consequences of grafting mesencephalic neurons into a peripheral nerve bridge connected to the denervated striatum. Exp Brain Res 1985;60:584–9. [11] Cheng H, Cao Y, Olson L. Spinal cord repair in adult paraplegic rats: partial restoration of hind limb function. Science 1996;273:510–3. [12] Balgude AP, Yu X, Szymanski A, Bellamkonda RV. Agarose gel stiffness determines rate of DRG neurite extension in 3-D cultures. Biomaterials 2001;22:1077–84.

[13] Flanagan LA, Ju YE, Marg B, Osterfeild M, Janmey PA. Neurite branching on deformable substrates. Neuroreport 2002;13:2411–5. [14] Ju YE, Jamney PA, McCormick ME, Sawyer ES, Flanagan LA. Enhanced neurite growth from mammalian neurons in three-dimensional salmon fibrin gels. Biomaterials 2007;28:2097–108. [15] Georges PC, Miller WJ, Meaney DF, Sawyer ES, Jamney PA. Matrices with compliance comparable to that of brain tissue select for neuronal over glial growth in mixed cortical cultures. Biophys J 2006;90:3012–8. [16] Woerly S, Doan VD, Sosa N, de Vellis J, Espinosa-Jeffrey A. Prevention of gliotic scar formation by NeuroGel allows partial endogenous repair of transected cat spinal cord. J Neurosci Res 2004;75:262–72. [17] Mahoney MJ, Anseth KS. Three-dimensional growth and function of neural tissue in degradable polyethylene glycol hydrogels. Biomaterials 2006;27:2265–74. [18] Miller K, Chinzei K, Orssengo G, Bednarz P. Mechanical properties of brain tissue in vivo: experiment and computer simulation. J Biomech 2000;33:1369–76. [19] Wilson JT, Chaikof EL. Challenges and emerging technologies in the immunoisolation of cells and tissues. Adv Drug Deliv Rev 2008;60:124–45. [20] Modo M, Rezaie P, Heuschling P, Patel S, Male DK, Hodges H. Transplantation of neural stem cells in a rat model of stroke: assessment of short-term graft survival and acute host immunological response. Brain Res 2002;958:70–82. [21] Sawhney AS, Chandrashekhar PP, Hubbell JA. Bioerodible hydrogels based on photopolymerized poly(ethylene glycol)-co-poly(alphahydroxy acid) diacrylate monomers. Macromolecules 1993;26:581–94. [22] Hollister SJ. Porous scaffold design for tissue engineering. Nat Mater 2005;4:518–24. [23] Soletti L, Nieponice A, Guan J, Stankus JJ, Wagner WR, Vorp DA. A seeding device for tissue engineered tubular structures. Biomaterials 2006;27:4863–70. [24] Karp JM, Dalton PD, Shoichet MS. Theme article–scaffolds for tissue engineering. Mater Res Soc 2003;28:94841. [25] Murphy WL, Dennis RG, Kileny JL, Mooney DJ. Salt fusion: an approach to improve pore interconnectivity within tissue engineering scaffolds. Tissue Eng 2002;8:43–52. [26] Xiong Z, Yan Y, Zhang R, Sun L. Fabrication of porous poly(lactic acid) scaffolds for bone tissue engineering via precise extrusion. Scripta Mater 2001;45:773–9. [27] Murugan R, Sampath TS, Ramakrishna S. Scaffolds for bone tissue restoration from biological apatite. Trends Biomater Artif Organs 2006;20:35–9. [28] Sachlos E, Czernuszka JT. Making tissue engineering scaffolds work. Review: the application of solid freeform fabrication technology to the production of tissue engineering scaffolds. Eur Cells Mater 2003;5:29–40. [29] Alvarez-Barreto JF, Linehan SM, Shambaugh RL, Sikavitsas VI. Flow perfusion improves seeding of tissue engineering scaffolds with different architectures. Ann Biomed Eng 2007;35:429–42. [30] Nisbet DR, Pattanawong S, Ritchie NE, Shen W, Finkelstein DI, Horne MK, et al. Interaction of embryonic cortical neurons on nanofibrous scaffolds for neural tissue engineering. J Neural Eng 2007;4:35–41. [31] Mikos AG, Thorsen AJ, Czerwonka LA, Bao Y, Langer R. Preparation and characterization of poly(L-lactic acid) foams. Polymer 1994;35:1068–77. [32] Mooney DJ, Baldwin DF, Suh NP, Vacanti JP, Langer R. Novel approach to fabricate porous sponges of poly(D,L-lactic co-glycolic acid) without the use of organic solvents. Biomaterials 1996;17:1417–22. [33] Mikos AG, Bao Y, Cima LG, Ingber DE, Vacanti JP, Langer R. Preparation of poly (glycolic acid) bonded fibres structures for cell attachment and transplantation. J Biomed Mater Res 1993;27:183–9. [34] Stokols S, Tuszynski MH. The fabrication and characterization of linearly oriented nerve guidance scaffolds for spinal cord injury. Biomaterials 2004;25:5839–46.

R.M. Namba et al. / Acta Biomaterialia 5 (2009) 1884–1897 [35] Whang K, Thomas CK, Nuber G, Healy KE. A novel method to fabricate bioabsorbable scaffolds. Polymer 1995;36:837. [36] Lo H, Ponticiello MS, Leong KW. Fabrication of controlled release biodegradable foams by phase separation. Tissue Eng 1995;1: 15–28. [37] Thompson RC, Yaszemski MJ, Powers JM, Mikos AG. Fabrication of biodegradable polymer scaffolds to engineering trabecular bone. J Biomater Sci-Polym 1995;7:23–38. [38] Ford MC, Bertram JP, Hynes SR, Michaud M, Li Q, Young M, et al. A macroporous hydrogel for the coculture of neural progenitor and endothelial cells to form functional vascular networks in vivo. PNAS 2006;103:2512–7. [39] Hartfuss E. Characterization of subtypes of precursor cells in the development of the central nervous system. PhD Thesis, Max Planck Institute for Neurobiology; 2003. [40] Hashimoto JG, Beadles-Bohling AS, Wiren KM. Comparison of RiboGreen and 18S rRNA quantitation for normalizing real-time RT-PCR expression analysis. Biotechniques 2004;1(54–56):58–60. [41] Bryant SJ, Anseth KS, Lee DA, Bader DL. Crosslinking density influences the morphology of chondrocytes photoencapsulated in PEG hydrogels during the application of compressive strain. J Orthop Res 2003;22:1143–9. [42] Burg KJ, Holder WDJ, Culberson CR, Beiler RJ, Greene KG, Loebsack AB, et al. Comparative study of seeding methods for threedimensional polymeric scaffolds. J Biomed Mater Res 2000;51:642–9. [43] Holy E, Shoichet MS, Davies JE, Chantal J. Engineering threedimensional bone tissue in vitro using biodegradable scaffolds: investigating initial cell-seeding density and culture period. Biomed Mater Res 2000;51:376–82. [44] Ito A, Ino K, Hayashida M, Kobayashi T, Matsunuma H, Kagami H. Novel methodology for fabrication of tissue-engineered tubular constructs using magnetite nanoparticles and magnetic force. Tissue Eng 2005;11:1553–61. [45] Li Y, Ma T, Kniss DA, Lasky LC, Yang ST. Effects of filtration seeding on cell density, spatial distribution, and proliferation in nonwoven fibrous matrices. Biotechnol Prog 2001;17:935–44. [46] Shastri VP, Hildgen P, Langer R. In situ pore formation in a polymer matrix by differential polymer degradation. Biomaterials 2003;24:3133–7.

1897

[47] Ren D, Yi H, Zhang H, Xie W, Wang W, Ma X. A preliminary study on fabrication of nanoscale fibrous chitosan membranes in situ by biospecific degradation. J Membr Sci 2006;280:99–107. [48] Coombes AGA, Verderio E, Shaw B, Li X, Griffin M, Downes S. Biocomposites of non-crosslinked natural and synthetic polymers. Biomaterials 2002:2113–8. [49] Matsumoto T, Sasaki J, Alsberg A, Egusa H, Yatani H, Sohmura T. Three-dimensional cell and tissue patterning in a strained fibrin gel system. PLoS ONE 2007;11:e1211. [50] Guo C, Kaufman LJ. Flow and magnetic field induced collagen alignment. Biomaterials 2007;28:1105–14. [51] Berthold A, Cremer K, Kreuter J. Collagen microparticles: carriers for glucocorticosteroids. Eur J Pharm Biopharm 1998;45:23–9. [52] Payne RG, Yaszemski MJ, Yasko AW, Mikos AG. Development of an injectable, in situ crosslinkable, degradable polymeric carrier for osteogenic cell populations. Part 1. Encapsulation of marrow stromalosteoblasts in surface crosslinked gelatin microparticles. Biomaterials 2002;22:4359–71. [53] Lee S, Moon JJ, Miller JS, West JL. Poly(ethylene glycol) hydrogels conjugated with a collagenase-sensitive fluorogenic substrate to visualize collagenase activity during three-dimensional cell migration. Biomaterials 2007;28:3163–70. [54] Bryant SJ, Anseth KS, Lee DA, Bader DL. Crosslinking density influences the morphology of chondrocytes photoencapsulated in PEG hydrogels during the application of compressive strain. J Orthop Res 2004;22:1143–9. [55] Dillon GP, Yu X, Sridharan A, Ranieri JP, Bellamkonda RV. The influence of physical structure and charge on neurite extension in a 3D hydrogel scaffold. J Biomater Sci Polym Ed 1998;9:1049–69. [56] Bellamkonda RV. Peripheral nerve regeneration: an opinion on channels, scaffolds and anisotropy. Biomaterials 2006;27:3515–8. [57] Ju YE, Jamney PA, McCormick ME, Sawyer ES, Flanagan LA. Enhanced neurite growth from mammalian neurons in three-dimensional salmon fibrin gels. Biomaterials 2007;28:2097–108. [58] Roisen FJ. The effects of dimethyl sulfoxide on neurite development in vitro. Ann NY Acad Sci 1975;243:279–96. [59] Roisen FJ, Murphy RA, Braden WG. Neurite development in vitro. I. The effects of adenosine 30 50 -cyclic monophosphate (cyclic AMP). J Neurobiol 1972;3:347–68.