- Email: [email protected]
Gas foamed open porous biodegradable polymeric microspheres
ARTICLE IN PRESS
Biomaterials 27 (2006) 152–159 www.elsevier.com/locate/biomaterials
Gas foamed open porous biodegradable polymeric microspheres Taek Kyoung Kima, Jun Jin Yoona, Doo Sung Leeb, Tae Gwan Parka, a
Department of Biological Sciences, Korea Advanced Institute of Science and Technology, Daejeon 305-701, South Korea Department of Polymer Science and Engineering, Sungkyunkwan University, Suwon, Kyungki-do 440-746, South Korea
b
Received 10 March 2005; accepted 27 May 2005 Available online 14 July 2005
Abstract Highly open porous biodegradable polymeric microspheres were fabricated for use as injectable scaffold microcarriers for cell delivery. A modified water-in-oil-in-water (W1/O/W2) double emulsion solvent evaporation method was employed for producing the microspheres. The incorporation of an effervescent salt, ammonium bicarbonate, in the primary W1 droplets spontaneously produced carbon dioxide and ammonia gas bubbles during the solvent evaporation process, which not only stabilized the primary emulsion, but also created well inter-connected pores in the resultant microspheres. The porous microspheres fabricated under various gas foaming conditions were characterized. The surface pores became as large as 20 mm in diameter with increasing the concentration of ammonium bicarbonate, being sufficient enough for cell infiltration and seeding. These porous scaffold microspheres could be potentially utilized for cultivating cells in a suspension manner and for delivering the seeded cells to the tissue defect site in an injectable manner. r 2005 Elsevier Ltd. All rights reserved. Keywords: Biodegradable polymer; PLGA; Porous; Microspheres; Gas foaming; Polylactic acid; Scaffold; Biodegradation; Microcarrier
1. Introduction Porous, biodegradable polymer scaffolds have been extensively utilized as temporal templates for regeneration of various tissues [1]. A highly open porous structure with well inter-connected pores is required not only to achieve sufficient cell seeding density within the scaffold, but also to facilitate in- and out-transport of nutrients and oxygen for subsequent cell proliferation and differentiation. A family of poly(lactic-co-glycolic acid) (PLGA) copolymers were popularly used for fabrication of such biodegradable scaffolds due to their controllable biodegradability and proven biocompatibility [2]. A wide range of PLGA scaffolds having diverse morphological characters were fabricated by various methods. Among them, a porogen leaching method was most extensively used; a variety of Corresponding author. Tel.: +82 42 869 2621; fax: +82 42 869 2610. E-mail address: [email protected] (T.G. Park).
0142-9612/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2005.05.081
particulate porogens, such as salts [3,4], carbohydrates [5], and hydrocarbon waxes [6], were imbedded into a polymer/solvent mixture and they were leached out after solvent evaporation for formation of pores. Other methods including emulsion/freeze drying [7], expansion in supercritical fluid [8,9], and 3 D-guided ink-jet printing [10] were also reported. We previously reported that macroporous PLGA scaffolds could be produced by a gas foaming/salt leaching method [11]. Effervescent salt particulates such as ammonium bicarbonate were homogeneously mixed, as a gas-evolving salt porogen, into a semi-solidified PLGA mass precipitated in a non-solvent, and the mixture was immersed in an acidic aqueous solution. Vibrant evolution of ammonia and carbon dioxide gas bubbles from the PLGA/salt mixture resulted in producing a large open porous structure throughout the PLGA scaffold. These scaffolds afforded high cell seeding density and subsequently showed promising potentials for regenerating liver [11] and cartilage tissues [12]. However, the aforementioned PLGA scaffolds were fabricated
ARTICLE IN PRESS T.K. Kim et al. / Biomaterials 27 (2006) 152–159
primarily for implantation in the body that requires an open surgery process. Their size and shape were tailored to fit into the dimension of a tissue defect. Recently, soft, hydrophilic, and biodegradable polymer materials were paid much attention to deliver cells into the defect site with a syringe in an injectable manner. After injection, they were photo-crosslinked or in situ gelled at the site to form cell-containing hydrogels [13,14]. A wide range of natural and synthetic microcarriers have been used to cultivate anchorage-dependent mammalian cells for producing therapeutic proteins [15,16]. They not only provide a large surface area for cell attachment, but also make feasible for suspension culture. However, these microcarriers are not biocompatible, unsuitable for human applications. To this end, biocompatible, biodegradable, and open porous PLGA microspheres are expected to be ideal scaffold carriers for cell delivery, because they can be utilized not only for suspension microcarriers for cell cultivation, but also for injectable vehicles for the cultivated cells. The porous structure of microcarriers is highly desirable for enhancing initial cell seeding density, and for promoting cell growth by facilitating mass transport of nutrients and oxygen. In this study, we fabricated highly open porous PLGA microspheres for injectable cell delivery. They were produced by a modified water-in-oil-in-water (W1/ O/W2) double emulsion solvent evaporation method. An effervescent salt was used as a gas foaming agent. Ammonium bicarbonate was incorporated in the primary W1 droplets to generate carbon dioxide and ammonia gas bubbles during the solvent evaporation process. The resultant porous microspheres prepared under different gas foaming conditions were characterized and their potentials for cell cultivation and delivery were explored.
2. Materials and methods 2.1. Materials Poly(D,L-lactide-co-glycolide) (lactide:glycolide ratio 75:25, RG756, Mw 100,000) (PLGA) was purchased from Boehringer Ingelheim (Ingelheim, Germany). Polyvinyl alcohol (PVA, 87–89% hydrolyzed, Mw 31,000–50,000) was from Sigma (St. Louis, MO). Fetal bovine serum (FBS) was purchased from Gibco (Gaithersburg, MD), Dulbeccos’s modified Eagle’s medium (DMEM) with 4.5 g/l glucose from Sigma (St. Louis, MO). NIH 3T3 mouse embryo fibroblasts were obtained from the Korean Cell Line Bank (KCLB). All other chemicals were of analytical grade. 2.2. Preparation of W/O emulsion film Ammonium bicarbonate was dissolved in 0.6 ml deionized water at a 5 wt% concentration, which was added to 2 ml of methylene chloride containing 125 mg of PLGA. Water-in-oil
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emulsion was prepared by a Powergen 700 homogenizer (Fisher Scientific Co., USA) at 5000 rpm for 2 min. The emulsion was then poured into a glass Petri dish (F ¼ 50 mm) and immediately examined by using the optical microscope (Eclipse TE300, Nikkon Co., Japan). The casting emulsion film was dried under a laminar flow at room temperature for 3 days and under vacuum for 2 days. After dried, the film was frozen with liquid nitrogen and fractured. Scanning electron microscopy (Phillips XL30S, Netherlands) was employed to view the internal structure of the fractured film. 2.3. Preparation of microspheres Microspheres were prepared by a W/O/W double emulsion method. To 8 ml of methylene chloride containing 500 mg PLGA, 2.5 ml of deionized water containing different amounts of NH4HCO3, was added. The first w/o emulsion was prepared using a Powergen 700 homogenizer at 5000 rpm for 3 min. This primary emulsion was immediately poured into a beaker containing 300 ml of 0.1% (w/v) PVA solution and then was re-emulsified by using an overhead propeller (LR-400A, Fisher Scientific Co., USA) for 4 h at 200 rpm. After the solvent was evaporated, the microspheres were separated by centrifugation, washed three times with distilled water and lyophilized using a freeze dryer. 2.4. Characterization of microspheres During solvent removal process, gas evolution from hardening microspheres was visualized on an optical microscope with a digital camera. To observe the surface and internal structures by scanning electron microscopy, dried porous microspheres were frozen with liquid nitrogen and fractured with a surgical blade. Samples were mounted on a metal stub and coated with gold. With an image software (Image J, developed at the US National Institute of Health), the size distribution and the average diameter of microspheres were determined by image analysis (n ¼ 100). In order to measure the surface pore size of microspheres, five microspheres were analyzed using Image J software. The total number of the surface pores was 100. Because the pore size of microspheres had a binary distribution, small size surface pores (below 5 mm) were ignored. Dried microspheres were first wetted by immersion in 70% ethanol, washed three times with cold deionized water, and incubated in phosphate buffer solution for 12 h. Water contents were measured by weighing wet microspheres after wiping the excess water on the surface. The water uptake ratio was calculated from the following equation: Water uptake ratio ¼ ðW w W d Þ=W d , where Ww and Wd are the wet weight and dry weight of the microspheres, respectively. 2.5. Cell culture Dry PLGA microspheres were sterilized by soaking into 70% ethanol at 4 1C for 6 h, and then ethanol was exchanged with excess amount of deionized water and subsequently 33 mM phosphate buffered saline (PBS, 0.1 M NaCl, pH 7). After removing PBS, the microspheres were resuspended in
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FBS at 4 1C for 12 h and then microspheres were washed with PBS three times. For seeding, 2.1 106 cells were inoculated into 1.3 ml of culture media containing 26 mg of microspheres in siliconized 12 well cell culture plate. The sample was maintained at 37 1C under 5% CO2 condition with continuous agitation. After 24-h incubation, the microspheres were rinsed with culture media and were filtered with a nylon mesh to remove unattached cells. The cell-attached microspheres were resuspended into 80 ml culture media in a siliconized spinner flask. The microsphere suspension was stirred at 50 rpm. The culture medium consisted of DMEM supplemented with 10% (v/v) FBS, 100 units/ml penicillin, and 100 g/ml streptomycin. 2.6. Cell attachment analysis Attached cells on the microspheres were stained with LysoTracker Reds (Molecular Probes) for cytological observation. The cell-attached microspheres were rinsed with PBS twice, and then the microspheres were resuspended culture media containing 75 nM LysoTracker probe. The cellattached microspheres were incubated for 30 min with agitation. Subsequently, the cell-attached microspheres were centrifuged at 50g for 3 min, and then resuspended into fresh media. Reddish color of the cells was observed using a confocal microscope (Carl Zeiss LSM5100, Germany). Scanning electron microscopy was employed to examine the attached cells on the microspheres. Visualization was carried out by using a scanning electron microscopy.
3. Results and discussion The porous PLGA microspheres were produced by using a water-in-oil-in-water (W1/O/W2) double emulsion method by dissolving ammonium bicarbonate in the inner W1 droplets as a gas foaming agent. The oil phase was methylene chloride containing PLGA. We hypothesized that the dissolved ammonium bicarbonate in the W1 droplets generated gas bubbles upon contacting the primary W1/O emulsion with W2 phase, thereby stabilizing the emulsion and creating open porous morphology throughout the PLGA microspheres. There have been a number of studies about preparing PLGA microspheres by using the W1/O/W2 double emulsion method, with aiming at delivering hydrophilic and macromolecular protein and peptide drugs in a sustained manner [17–19]. These PLGA microspheres (normally less than 100 mm in diameter), however, did not have an open porous structure. The inner core had closed pores surrounded by PLGA polymer phase and the surface had very tiny pores. They are not suitable for a cell-delivering vehicle. To test the stabilization effect of dissolved effervescent salts in the primary W1 phase emulsion droplets on the porous morphology of a PLGA membrane, the W1/O emulsion solution was cast and dried onto a glass surface. As shown in Fig. 1, a highly porous PLGA film
was obtained. The ammonium bicarbonate included in the aqueous droplets spontaneously produced ammonia and carbon dioxide gas bubbles during solvent evaporation, resulting in the formation of open pores in the interior region. It can be seen in the optical microscopic picture that small gas bubble droplets are generated from the water droplets in the organic phase, which are likely to stabilize the primary emulsion droplets against coalescence. Since no surfactant was used in the experiment, the emulsion stabilization effect can be attributed to the spontaneous evolution of gas bubbles in the oil phase [20]. The coalescence of emulsion droplets is mainly caused by inherent thermodynamic instability of an interface between water and oil phases, which tends to reduce the interfacial area between the two immiscible phases. Two metastable emulsion droplets must be collided prior to coalescence. In this regard, small gas bubbles evolved from the surface of water droplets might play a critical role in sterically preventing them from the coalescence. In the SEM picture, it is clearly visible that many small pores are located in the vicinity of large pores, indicating that during the solvent evaporation, the small gas bubble droplets protected the primary aqueous droplets from aggregation by acting as coalescence barriers. When pure water or sodium chloride at the same concentration level was added in the W1 phase, the resultant membranes exhibited the formation of large isolated and scattered pores in the inner region due to the immediate coalescence of the aqueous droplets during the solvent removal. The results above revealed that aqueous emulsion droplets containing dissolved effervescent salts gradually generated gas filled bubbles, which stabilized inner pore structures, developing well inter-connection between the pores. Upon re-emulsifying the primary W1/O emulsion solution containing ammonium bicarbonate in the W1 phase into W2 phase containing poly(vinyl alcohol) (PVA), highly porous microspheres were successfully produced. In the optical microscopic observation, it was evident that gas bubbles were generated and were evolved through the surface during the in-water solvent evaporation. A foamy surface structure having many gas bubbles can be seen clearly. The resulting PLGA microspheres showed open pores with maintaining interconnectivities between the pores. The porous morphology in the cross-section was similar to that on the surface, indicating that a porous structure was produced homogeneously throughout the bulk phase of microspheres. Fig. 2 shows SEM pictures of porous PLGA microspheres prepared by incorporating different amounts of ammonium bicarbonate in the W1 aqueous phase. With increasing amount of ammonium bicarbonate, the more porous structure on the surface could be attained. There was no discernible difference in open porous morphology between the surface and the internal
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Fig. 1. A schematic diagram of porous membrane and microsphere prepared by a gas foaming method. Left panel: porous PLGA membrane prepared by W/O emulsion. Right panel: porous PLGA microsphere prepared by W/O/W double emulsion: (a) optical microphotograph of W/O primary emulsion droplets during solvent evaporation; (b) optical microphotograph of W/O/W double emulsion droplets during solvent evaporation; (c) cross-sectional SEM image of dried porous PLGA film; and (d) cross-sectional SEM image of dried porous PLGA microsphere.
region of PLGA microspheres (data not shown). At 10% ammonium bicarbonate concentration, large open pores on the surface can be seen. The SEM images of PLGA microspheres revealed the existence of a binary pore distribution composed of small and large pores, similar to the SEM images of the porous membrane as shown in Fig. 1. The small pores, presumably originated from evolved gas bubbles, were located in-between large pores that had been produced from the W1 aqueous phase droplets. It was likely that the small gas-filled droplets prevented the coalescence of the larger primary emulsion droplets as demonstrated in the porous PLGA film study. Fig. 3 shows morphological characteristics of PLGA microspheres prepared by adding increasing
amount of ammonium bicarbonate in the W1 aqueous phase of primary emulsion solution. For 0%, 1%, 5%, and 10% (w/v) ammonium bicarbonate, average diameters of PLGA microspheres were 343760, 403770, 439776, and 535762 mm, respectively (Fig. 3a), suggesting that larger porous PLGA microspheres were produced by increasing the amount of ammonium bicarbonate. This was obviously due to the effect of internal gas foaming that expanded the dimension of gas evolving primary aqueous droplets and generated additional gas bubbles in the solidifying polymer phase [11]. During solvent evaporation in the second emulsification step, the oil phase (methylene chloride dissolved with PLGA), being initially fluidic, became viscous and
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Fig. 2. SEM images of porous PLGA microspheres prepared by W/O/W double emulsion method containing different amounts of ammonium bicarbonate in the W1 phase. (a,b) 0%; (c,d) 1%; (e,f) 5%; (g,h) 10%. Left panel: gross morphology; right panel: surface.
elastic with gradual removal of solvent, and ultimately hardened [21]. The generation of gas within W1 phase droplets surrounded by the viscous and elastic PLGA phase enabled oil phase droplets to enlarge their original volume. Meanwhile, small gas bubbles evolved and escaped from the aqueous droplets into the oil phase
also increased the volume of PLGA skeletal backbone. Thus, incorporated ammonium bicarbonate in the primary emulsion droplets as a dispersed aqueous phase certainly contributed to the enlarged size of PLGA microspheres. Fig. 3b shows water uptake ratios of PLGA microspheres as a function of ammonium
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(b)
(a)
(c)
Fig. 3. (a) Size distribution of porous PLGA microspheres prepared by W/O/W double emulsion method containing different amounts of ammonium bicarbonate in the W1 phase; (b) water uptake ratio; and (c) surface pore diameter.
bicarbonate concentration in the aqueous phase. The water uptake ratio, a value calculated by dividing the wet weight of microspheres by the dry weight of microspheres, increased with the concentration of ammonium bicarbonate. This indicates that PLGA microspheres prepared using greater amount of ammonium bicarbonate had larger internal pore volume per dry weight of microspheres, which was equivalent to the porosity of microspheres. Fig. 3c exhibits apparent pore size on the surface. Surface pore diameter values also increased with increasing the concentration of ammonium bicarbonate. For porous PLGA microspheres using 10% ammonium bicarbonate, open pores up to 20 mm in diameter were produced on the surface. This result is also consistent with the previous finding that the generation of gas foaming bubbles in the solidifying PLGA microspheres during solvent removal resulted in larger microspheres having greater porosity. Open porous PLGA microspheres were used as suspension microcarriers for cultivating NIH3T3 cells in a spinner flask [22]. As shown in Fig. 4, NIH3T3 cells were homogeneously adhered onto the surface in the early culture period (1 day). The confocal and SEM images (Fig. 4a and b) show that the adhered cells are well spread on the surface. After prolonged suspension cultivation of 7 days, however, it can be observed that spherical cells are densely present within the pores of the microspheres, not on the surface. This suggests that initially attached cells on the surface gradually migrated
towards surface and inner pores, wherein they proliferated [23]. The size of surface pores (ca. 20 mm) was slightly larger than that of a single cell, resulting in the facile penetration of cells towards the inner pores [24]. It was likely that the cells were present primarily on the surface pore region due to narrow inter-connecting channels between the inner pores. Since the cells were cultivated in a suspension manner, the cells weakly adhered onto the surface were detached into the medium due to exerted shear stress on them, whereas the cells residing within the pores survived. Thus, open porous PLGA microspheres provide favorable spatial environment for the proliferation of cells that require protection from shear stress. To further improve cell adhesion and penetration onto and into the microspheres, cell adhesive peptide ligands such as arginine–glycine–aspartic acid (RGD) peptides could be immobilized onto the surface of microspheres. We have previously reported RGD immobilized PLGA scaffolds to enhance cell adhesion and function, which were fabricated by the same gas foaming method used in this study [25]. This study primarily aims at the new fabrication method of open porous microspheres that can be utilized as injectable and biodegradable scaffold microcarriers. More detailed quantitative studies on cell seeding and cellular behaviors within the microspheres, such as cell seeding density, adhesion, viability, and proliferation, would be necessary. The long-term cell cultivation results will be reported in the near future.
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foaming agent. The incorporation of ammonium bicarbonate in the primary emulsion droplets of W/O/W double emulsion formulation stabilized the primary emulsion droplets and subsequently generated a highly open porous structure after solvent evaporation. The resultant open porous biodegradable microspheres can be potentially used as temporal scaffold microcarriers for proliferation of cells. They can be also used as injectable cell-containing microcarriers for tissue repair.
Acknowledgement This study was supported by the grant (KRF-2004005-D00070) from the Korea Research Foundation, Korea and the Polymer Technology Institute, Sungkyunkwan University, Korea. References
Fig. 4. Cells attached on porous microspheres: (a) confocal image of dye labeled cells attached on the surface after 1 day culture; (b) SEM image of cells attached on the surface after 1 day culture; and (c) SEM image of cells proliferated within the pores after 7-day culture.
4. Conclusions Highly open porous PLGA microspheres were produced by using ammonium bicarbonate as a gas
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Biomaterials 27 (2006) 152–159 www.elsevier.com/locate/biomaterials
Gas foamed open porous biodegradable polymeric microspheres Taek Kyoung Kima, Jun Jin Yoona, Doo Sung Leeb, Tae Gwan Parka, a
Department of Biological Sciences, Korea Advanced Institute of Science and Technology, Daejeon 305-701, South Korea Department of Polymer Science and Engineering, Sungkyunkwan University, Suwon, Kyungki-do 440-746, South Korea
b
Received 10 March 2005; accepted 27 May 2005 Available online 14 July 2005
Abstract Highly open porous biodegradable polymeric microspheres were fabricated for use as injectable scaffold microcarriers for cell delivery. A modified water-in-oil-in-water (W1/O/W2) double emulsion solvent evaporation method was employed for producing the microspheres. The incorporation of an effervescent salt, ammonium bicarbonate, in the primary W1 droplets spontaneously produced carbon dioxide and ammonia gas bubbles during the solvent evaporation process, which not only stabilized the primary emulsion, but also created well inter-connected pores in the resultant microspheres. The porous microspheres fabricated under various gas foaming conditions were characterized. The surface pores became as large as 20 mm in diameter with increasing the concentration of ammonium bicarbonate, being sufficient enough for cell infiltration and seeding. These porous scaffold microspheres could be potentially utilized for cultivating cells in a suspension manner and for delivering the seeded cells to the tissue defect site in an injectable manner. r 2005 Elsevier Ltd. All rights reserved. Keywords: Biodegradable polymer; PLGA; Porous; Microspheres; Gas foaming; Polylactic acid; Scaffold; Biodegradation; Microcarrier
1. Introduction Porous, biodegradable polymer scaffolds have been extensively utilized as temporal templates for regeneration of various tissues [1]. A highly open porous structure with well inter-connected pores is required not only to achieve sufficient cell seeding density within the scaffold, but also to facilitate in- and out-transport of nutrients and oxygen for subsequent cell proliferation and differentiation. A family of poly(lactic-co-glycolic acid) (PLGA) copolymers were popularly used for fabrication of such biodegradable scaffolds due to their controllable biodegradability and proven biocompatibility [2]. A wide range of PLGA scaffolds having diverse morphological characters were fabricated by various methods. Among them, a porogen leaching method was most extensively used; a variety of Corresponding author. Tel.: +82 42 869 2621; fax: +82 42 869 2610. E-mail address: [email protected] (T.G. Park).
0142-9612/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2005.05.081
particulate porogens, such as salts [3,4], carbohydrates [5], and hydrocarbon waxes [6], were imbedded into a polymer/solvent mixture and they were leached out after solvent evaporation for formation of pores. Other methods including emulsion/freeze drying [7], expansion in supercritical fluid [8,9], and 3 D-guided ink-jet printing [10] were also reported. We previously reported that macroporous PLGA scaffolds could be produced by a gas foaming/salt leaching method [11]. Effervescent salt particulates such as ammonium bicarbonate were homogeneously mixed, as a gas-evolving salt porogen, into a semi-solidified PLGA mass precipitated in a non-solvent, and the mixture was immersed in an acidic aqueous solution. Vibrant evolution of ammonia and carbon dioxide gas bubbles from the PLGA/salt mixture resulted in producing a large open porous structure throughout the PLGA scaffold. These scaffolds afforded high cell seeding density and subsequently showed promising potentials for regenerating liver [11] and cartilage tissues [12]. However, the aforementioned PLGA scaffolds were fabricated
ARTICLE IN PRESS T.K. Kim et al. / Biomaterials 27 (2006) 152–159
primarily for implantation in the body that requires an open surgery process. Their size and shape were tailored to fit into the dimension of a tissue defect. Recently, soft, hydrophilic, and biodegradable polymer materials were paid much attention to deliver cells into the defect site with a syringe in an injectable manner. After injection, they were photo-crosslinked or in situ gelled at the site to form cell-containing hydrogels [13,14]. A wide range of natural and synthetic microcarriers have been used to cultivate anchorage-dependent mammalian cells for producing therapeutic proteins [15,16]. They not only provide a large surface area for cell attachment, but also make feasible for suspension culture. However, these microcarriers are not biocompatible, unsuitable for human applications. To this end, biocompatible, biodegradable, and open porous PLGA microspheres are expected to be ideal scaffold carriers for cell delivery, because they can be utilized not only for suspension microcarriers for cell cultivation, but also for injectable vehicles for the cultivated cells. The porous structure of microcarriers is highly desirable for enhancing initial cell seeding density, and for promoting cell growth by facilitating mass transport of nutrients and oxygen. In this study, we fabricated highly open porous PLGA microspheres for injectable cell delivery. They were produced by a modified water-in-oil-in-water (W1/ O/W2) double emulsion solvent evaporation method. An effervescent salt was used as a gas foaming agent. Ammonium bicarbonate was incorporated in the primary W1 droplets to generate carbon dioxide and ammonia gas bubbles during the solvent evaporation process. The resultant porous microspheres prepared under different gas foaming conditions were characterized and their potentials for cell cultivation and delivery were explored.
2. Materials and methods 2.1. Materials Poly(D,L-lactide-co-glycolide) (lactide:glycolide ratio 75:25, RG756, Mw 100,000) (PLGA) was purchased from Boehringer Ingelheim (Ingelheim, Germany). Polyvinyl alcohol (PVA, 87–89% hydrolyzed, Mw 31,000–50,000) was from Sigma (St. Louis, MO). Fetal bovine serum (FBS) was purchased from Gibco (Gaithersburg, MD), Dulbeccos’s modified Eagle’s medium (DMEM) with 4.5 g/l glucose from Sigma (St. Louis, MO). NIH 3T3 mouse embryo fibroblasts were obtained from the Korean Cell Line Bank (KCLB). All other chemicals were of analytical grade. 2.2. Preparation of W/O emulsion film Ammonium bicarbonate was dissolved in 0.6 ml deionized water at a 5 wt% concentration, which was added to 2 ml of methylene chloride containing 125 mg of PLGA. Water-in-oil
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emulsion was prepared by a Powergen 700 homogenizer (Fisher Scientific Co., USA) at 5000 rpm for 2 min. The emulsion was then poured into a glass Petri dish (F ¼ 50 mm) and immediately examined by using the optical microscope (Eclipse TE300, Nikkon Co., Japan). The casting emulsion film was dried under a laminar flow at room temperature for 3 days and under vacuum for 2 days. After dried, the film was frozen with liquid nitrogen and fractured. Scanning electron microscopy (Phillips XL30S, Netherlands) was employed to view the internal structure of the fractured film. 2.3. Preparation of microspheres Microspheres were prepared by a W/O/W double emulsion method. To 8 ml of methylene chloride containing 500 mg PLGA, 2.5 ml of deionized water containing different amounts of NH4HCO3, was added. The first w/o emulsion was prepared using a Powergen 700 homogenizer at 5000 rpm for 3 min. This primary emulsion was immediately poured into a beaker containing 300 ml of 0.1% (w/v) PVA solution and then was re-emulsified by using an overhead propeller (LR-400A, Fisher Scientific Co., USA) for 4 h at 200 rpm. After the solvent was evaporated, the microspheres were separated by centrifugation, washed three times with distilled water and lyophilized using a freeze dryer. 2.4. Characterization of microspheres During solvent removal process, gas evolution from hardening microspheres was visualized on an optical microscope with a digital camera. To observe the surface and internal structures by scanning electron microscopy, dried porous microspheres were frozen with liquid nitrogen and fractured with a surgical blade. Samples were mounted on a metal stub and coated with gold. With an image software (Image J, developed at the US National Institute of Health), the size distribution and the average diameter of microspheres were determined by image analysis (n ¼ 100). In order to measure the surface pore size of microspheres, five microspheres were analyzed using Image J software. The total number of the surface pores was 100. Because the pore size of microspheres had a binary distribution, small size surface pores (below 5 mm) were ignored. Dried microspheres were first wetted by immersion in 70% ethanol, washed three times with cold deionized water, and incubated in phosphate buffer solution for 12 h. Water contents were measured by weighing wet microspheres after wiping the excess water on the surface. The water uptake ratio was calculated from the following equation: Water uptake ratio ¼ ðW w W d Þ=W d , where Ww and Wd are the wet weight and dry weight of the microspheres, respectively. 2.5. Cell culture Dry PLGA microspheres were sterilized by soaking into 70% ethanol at 4 1C for 6 h, and then ethanol was exchanged with excess amount of deionized water and subsequently 33 mM phosphate buffered saline (PBS, 0.1 M NaCl, pH 7). After removing PBS, the microspheres were resuspended in
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FBS at 4 1C for 12 h and then microspheres were washed with PBS three times. For seeding, 2.1 106 cells were inoculated into 1.3 ml of culture media containing 26 mg of microspheres in siliconized 12 well cell culture plate. The sample was maintained at 37 1C under 5% CO2 condition with continuous agitation. After 24-h incubation, the microspheres were rinsed with culture media and were filtered with a nylon mesh to remove unattached cells. The cell-attached microspheres were resuspended into 80 ml culture media in a siliconized spinner flask. The microsphere suspension was stirred at 50 rpm. The culture medium consisted of DMEM supplemented with 10% (v/v) FBS, 100 units/ml penicillin, and 100 g/ml streptomycin. 2.6. Cell attachment analysis Attached cells on the microspheres were stained with LysoTracker Reds (Molecular Probes) for cytological observation. The cell-attached microspheres were rinsed with PBS twice, and then the microspheres were resuspended culture media containing 75 nM LysoTracker probe. The cellattached microspheres were incubated for 30 min with agitation. Subsequently, the cell-attached microspheres were centrifuged at 50g for 3 min, and then resuspended into fresh media. Reddish color of the cells was observed using a confocal microscope (Carl Zeiss LSM5100, Germany). Scanning electron microscopy was employed to examine the attached cells on the microspheres. Visualization was carried out by using a scanning electron microscopy.
3. Results and discussion The porous PLGA microspheres were produced by using a water-in-oil-in-water (W1/O/W2) double emulsion method by dissolving ammonium bicarbonate in the inner W1 droplets as a gas foaming agent. The oil phase was methylene chloride containing PLGA. We hypothesized that the dissolved ammonium bicarbonate in the W1 droplets generated gas bubbles upon contacting the primary W1/O emulsion with W2 phase, thereby stabilizing the emulsion and creating open porous morphology throughout the PLGA microspheres. There have been a number of studies about preparing PLGA microspheres by using the W1/O/W2 double emulsion method, with aiming at delivering hydrophilic and macromolecular protein and peptide drugs in a sustained manner [17–19]. These PLGA microspheres (normally less than 100 mm in diameter), however, did not have an open porous structure. The inner core had closed pores surrounded by PLGA polymer phase and the surface had very tiny pores. They are not suitable for a cell-delivering vehicle. To test the stabilization effect of dissolved effervescent salts in the primary W1 phase emulsion droplets on the porous morphology of a PLGA membrane, the W1/O emulsion solution was cast and dried onto a glass surface. As shown in Fig. 1, a highly porous PLGA film
was obtained. The ammonium bicarbonate included in the aqueous droplets spontaneously produced ammonia and carbon dioxide gas bubbles during solvent evaporation, resulting in the formation of open pores in the interior region. It can be seen in the optical microscopic picture that small gas bubble droplets are generated from the water droplets in the organic phase, which are likely to stabilize the primary emulsion droplets against coalescence. Since no surfactant was used in the experiment, the emulsion stabilization effect can be attributed to the spontaneous evolution of gas bubbles in the oil phase [20]. The coalescence of emulsion droplets is mainly caused by inherent thermodynamic instability of an interface between water and oil phases, which tends to reduce the interfacial area between the two immiscible phases. Two metastable emulsion droplets must be collided prior to coalescence. In this regard, small gas bubbles evolved from the surface of water droplets might play a critical role in sterically preventing them from the coalescence. In the SEM picture, it is clearly visible that many small pores are located in the vicinity of large pores, indicating that during the solvent evaporation, the small gas bubble droplets protected the primary aqueous droplets from aggregation by acting as coalescence barriers. When pure water or sodium chloride at the same concentration level was added in the W1 phase, the resultant membranes exhibited the formation of large isolated and scattered pores in the inner region due to the immediate coalescence of the aqueous droplets during the solvent removal. The results above revealed that aqueous emulsion droplets containing dissolved effervescent salts gradually generated gas filled bubbles, which stabilized inner pore structures, developing well inter-connection between the pores. Upon re-emulsifying the primary W1/O emulsion solution containing ammonium bicarbonate in the W1 phase into W2 phase containing poly(vinyl alcohol) (PVA), highly porous microspheres were successfully produced. In the optical microscopic observation, it was evident that gas bubbles were generated and were evolved through the surface during the in-water solvent evaporation. A foamy surface structure having many gas bubbles can be seen clearly. The resulting PLGA microspheres showed open pores with maintaining interconnectivities between the pores. The porous morphology in the cross-section was similar to that on the surface, indicating that a porous structure was produced homogeneously throughout the bulk phase of microspheres. Fig. 2 shows SEM pictures of porous PLGA microspheres prepared by incorporating different amounts of ammonium bicarbonate in the W1 aqueous phase. With increasing amount of ammonium bicarbonate, the more porous structure on the surface could be attained. There was no discernible difference in open porous morphology between the surface and the internal
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Fig. 1. A schematic diagram of porous membrane and microsphere prepared by a gas foaming method. Left panel: porous PLGA membrane prepared by W/O emulsion. Right panel: porous PLGA microsphere prepared by W/O/W double emulsion: (a) optical microphotograph of W/O primary emulsion droplets during solvent evaporation; (b) optical microphotograph of W/O/W double emulsion droplets during solvent evaporation; (c) cross-sectional SEM image of dried porous PLGA film; and (d) cross-sectional SEM image of dried porous PLGA microsphere.
region of PLGA microspheres (data not shown). At 10% ammonium bicarbonate concentration, large open pores on the surface can be seen. The SEM images of PLGA microspheres revealed the existence of a binary pore distribution composed of small and large pores, similar to the SEM images of the porous membrane as shown in Fig. 1. The small pores, presumably originated from evolved gas bubbles, were located in-between large pores that had been produced from the W1 aqueous phase droplets. It was likely that the small gas-filled droplets prevented the coalescence of the larger primary emulsion droplets as demonstrated in the porous PLGA film study. Fig. 3 shows morphological characteristics of PLGA microspheres prepared by adding increasing
amount of ammonium bicarbonate in the W1 aqueous phase of primary emulsion solution. For 0%, 1%, 5%, and 10% (w/v) ammonium bicarbonate, average diameters of PLGA microspheres were 343760, 403770, 439776, and 535762 mm, respectively (Fig. 3a), suggesting that larger porous PLGA microspheres were produced by increasing the amount of ammonium bicarbonate. This was obviously due to the effect of internal gas foaming that expanded the dimension of gas evolving primary aqueous droplets and generated additional gas bubbles in the solidifying polymer phase [11]. During solvent evaporation in the second emulsification step, the oil phase (methylene chloride dissolved with PLGA), being initially fluidic, became viscous and
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Fig. 2. SEM images of porous PLGA microspheres prepared by W/O/W double emulsion method containing different amounts of ammonium bicarbonate in the W1 phase. (a,b) 0%; (c,d) 1%; (e,f) 5%; (g,h) 10%. Left panel: gross morphology; right panel: surface.
elastic with gradual removal of solvent, and ultimately hardened [21]. The generation of gas within W1 phase droplets surrounded by the viscous and elastic PLGA phase enabled oil phase droplets to enlarge their original volume. Meanwhile, small gas bubbles evolved and escaped from the aqueous droplets into the oil phase
also increased the volume of PLGA skeletal backbone. Thus, incorporated ammonium bicarbonate in the primary emulsion droplets as a dispersed aqueous phase certainly contributed to the enlarged size of PLGA microspheres. Fig. 3b shows water uptake ratios of PLGA microspheres as a function of ammonium
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(b)
(a)
(c)
Fig. 3. (a) Size distribution of porous PLGA microspheres prepared by W/O/W double emulsion method containing different amounts of ammonium bicarbonate in the W1 phase; (b) water uptake ratio; and (c) surface pore diameter.
bicarbonate concentration in the aqueous phase. The water uptake ratio, a value calculated by dividing the wet weight of microspheres by the dry weight of microspheres, increased with the concentration of ammonium bicarbonate. This indicates that PLGA microspheres prepared using greater amount of ammonium bicarbonate had larger internal pore volume per dry weight of microspheres, which was equivalent to the porosity of microspheres. Fig. 3c exhibits apparent pore size on the surface. Surface pore diameter values also increased with increasing the concentration of ammonium bicarbonate. For porous PLGA microspheres using 10% ammonium bicarbonate, open pores up to 20 mm in diameter were produced on the surface. This result is also consistent with the previous finding that the generation of gas foaming bubbles in the solidifying PLGA microspheres during solvent removal resulted in larger microspheres having greater porosity. Open porous PLGA microspheres were used as suspension microcarriers for cultivating NIH3T3 cells in a spinner flask [22]. As shown in Fig. 4, NIH3T3 cells were homogeneously adhered onto the surface in the early culture period (1 day). The confocal and SEM images (Fig. 4a and b) show that the adhered cells are well spread on the surface. After prolonged suspension cultivation of 7 days, however, it can be observed that spherical cells are densely present within the pores of the microspheres, not on the surface. This suggests that initially attached cells on the surface gradually migrated
towards surface and inner pores, wherein they proliferated [23]. The size of surface pores (ca. 20 mm) was slightly larger than that of a single cell, resulting in the facile penetration of cells towards the inner pores [24]. It was likely that the cells were present primarily on the surface pore region due to narrow inter-connecting channels between the inner pores. Since the cells were cultivated in a suspension manner, the cells weakly adhered onto the surface were detached into the medium due to exerted shear stress on them, whereas the cells residing within the pores survived. Thus, open porous PLGA microspheres provide favorable spatial environment for the proliferation of cells that require protection from shear stress. To further improve cell adhesion and penetration onto and into the microspheres, cell adhesive peptide ligands such as arginine–glycine–aspartic acid (RGD) peptides could be immobilized onto the surface of microspheres. We have previously reported RGD immobilized PLGA scaffolds to enhance cell adhesion and function, which were fabricated by the same gas foaming method used in this study [25]. This study primarily aims at the new fabrication method of open porous microspheres that can be utilized as injectable and biodegradable scaffold microcarriers. More detailed quantitative studies on cell seeding and cellular behaviors within the microspheres, such as cell seeding density, adhesion, viability, and proliferation, would be necessary. The long-term cell cultivation results will be reported in the near future.
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foaming agent. The incorporation of ammonium bicarbonate in the primary emulsion droplets of W/O/W double emulsion formulation stabilized the primary emulsion droplets and subsequently generated a highly open porous structure after solvent evaporation. The resultant open porous biodegradable microspheres can be potentially used as temporal scaffold microcarriers for proliferation of cells. They can be also used as injectable cell-containing microcarriers for tissue repair.
Acknowledgement This study was supported by the grant (KRF-2004005-D00070) from the Korea Research Foundation, Korea and the Polymer Technology Institute, Sungkyunkwan University, Korea. References
Fig. 4. Cells attached on porous microspheres: (a) confocal image of dye labeled cells attached on the surface after 1 day culture; (b) SEM image of cells attached on the surface after 1 day culture; and (c) SEM image of cells proliferated within the pores after 7-day culture.
4. Conclusions Highly open porous PLGA microspheres were produced by using ammonium bicarbonate as a gas
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