Fabrication of Biodegradable Polymeric Microspheres with Controllable Porous Structure for Cell Delivery

Porous microspheres fabricated by biodegradable polymers show great potential
as carriers for cell cultivation in tissue engineering. We have attempted to prepare
porous microspheres using various biodegradable polymers, and the polymer used is
biodegradable Poly(D,L-lactide-co-glycolide) (PLGA), chitosan, hyaluronic acid, gelatin.
The influence of fabrication parameters, such as the concentration of crosslinker and
polymer, concentration and type of porogen, as well as the post-hydrolysis treatment
and thawing time of the porous structure of the microspheres are discussed. an active
spread of Hep3B-GFP cells on the microspheres is observed, which indicates that the
biodegradable polymeric microsphere with controllable porous structure are of great
potential as cell delivery carriers for tissue engineering.


Introduction
Many groups have demonstrated the potential of cell delivery in therapeutic applications by directly injecting a dispersion of cells into the diseased or injured site [1][2][3][4]. Despite its proven ability to improve organ and tissue function after damage, cell transplantation is limited by poor graft retention following the delivery of the cells.

A significant proportion of the transplanted cells leak out through
the hole that is made by the puncturing needle or enter systemic circulation [5,6]. How to precisely control the amount, distribution, and viability of the injected cells has become a major challenge in maximizing the therapeutic effects and elucidating the healing mechanism [7,8].
To prevent loss of cells, various injectable hydrogels, formed from native or synthetic polymers, have been co-implanted with the cells [9][10][11][12]. Upon injection, the hydrogels undergo an in-situ sol-gel transition and thus embed the transplanted cells intramuscularly.
As a major drawback, the cells encapsulated within the interior of a bulk hydrogel may lack a sufficient supply of oxygen or nutrient, resulting in massive cell death and a non-uniform cell distribution [13]. In addition, their low mechanical strength and durability are still problematic for practical applications [14]. In comparison, porous constructs made of synthetic and/or natural polymers with a three-dimensionally(3D) porous structure are attractive carriers for cell delivery because they allow for quantification of the number of cells injected, as well as improvement of cell viability and retention [15]. In particular, injectable scaffolds show promise for this application, as cells can be mixed homogeneously with the scaffold formulation prior to injection. The ability to deliver injectable scaffolds in a minimally invasive manner to a cavity of any size or shape renders them especially attractive for clinical use in tissue repair [16]. One type of injectable scaffold for tissue engineering applications involves the use of discreet polymer microspheres. Microspheres can be fabricated using a variety of different biodegradable polymers such as chitosan, gelatin and PLGA, and their use for delivery of cells and growth factors for repair of tissues such as bone, skin and brain has been reported [17][18][19][20][21]. To establish a polymer and porosity that would produce structurally stable porous scaffolds, we tried to manufacture microspheres from various polymers. Finally, an in vitro Hep3B-evaluate the influence of the various polymer microspheres on cell growth and to assess the microspheres biomedical potentials as microcarriers.

Preparation of Porous PLGA Microspheres by Gas-Foaming Method
Gas-foaming method was used with the ammonium bicarbonate as porogen. To 8 mL of methylene chloride containing 6.25% (w/v) PLGA, 2.5 mL of deionized water containing different amounts of ammonium bicarbonate (5, 10, 20 and 30%, w/v) was added. The first W-O emulsion was prepared using a homogenizer (Dispenser T 10 basic, IKA Works, Inc.) at 5,000 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 a magnetic stirrer (HS 10, IKA Works, Inc.) for 4 hours 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 [22]. The lyophilized microspheres were immersed in 80 mL of various concentrations of NaOH solution for 2 hours for control of pore size. Afterward, the hydrolyzed microspheres were washed with distilled water to remove the remaining NaOH solution [23].

Preparation of Porous PLGA Microspheres by Microfluidic System
On the other hand, porous PLGA microspheres were prepared using a microfluidic system that consisted of a Tygon® tube (0.8 and needles (25G). The device with two-way flow channels was fabricated by inserting the needle and the capillary tube into the Tygon® tube, followed by sealing with epoxy adhesive [24]. The respectively. The W-O-W droplets, formed at the tip of the needle, flowed along the capillary tube into ice-cold water (collection phase) and were gently stirred overnight to remove the organic solvent by evaporation. To remove the residual gelatin, the acquired microspheres were gently stirred for 3 hours in a warm water bath.
The resultant microspheres were washed with deionized water three times and collected for further application [25].

Preparation of Porous Chitosan Microspheres
Porous chitosan microspheres were prepared using freeze-

Preparation of Porous Hyaluronic Acid Microspheres
Porous hyaluronic acid microspheres were prepared according to the procedure that was described for the fabrication of chitosan.
A series of hyaluronic acid solutions (1, 2 and 4%, w/v) in distilled water was used. To investigate the control of pore size, the lyophilized hyaluronic acid microspheres were stirred in ethanol of various concentrations (0.5, 10, 20 and 30% crosslink yield) of cross-linking agent for 24 hours. After the cross-linking agent was removed, the microspheres were separated by centrifugation and re-lyophilized using a freeze dryer.

Preparation of Porous Gelatin Microspheres
Porous gelatin microspheres were prepared according to the procedure that was described for the fabrication of chitosan.
A gelatin solution (20%, w/v) in distilled water was used.

Characterization of Microspheres
Scanning electron microscopy (SNE-4000M, SEC, KR) was used to observe the surface morphology and pore size of the microspheres. The morphology of microspheres was observed after gold coating using a sputter-coater (MCM-100, SEC, KR). The distribution of pore and microspheres size calculated using ImageJ.

Cell Culture onto Porous Microspheres
The various porous microspheres were sterilized by 70% ethanol solution, washed with phosphate-buffered saline, and preincubated overnight in the culture medium at 37℃, respectively. To        Figure 6 shows the morphologies of porous chitosan microspheres obtained; that the pores were well interconnected;

Preparation of Porous Chitosan Microspheres
and that their distribution was fairly homogeneous. As the chitosan concentration increased from 1 to 4 wt%, the diameter of the porous microspheres was increased and the pore size and porosity were decreased. As shown in Figure 7, the pore size was dramatically increased by the thaw-refreeze method. It was evident that a longer thawing time results in the formation of larger interconnected pores inside the microspheres. When the chitosan microspheres were frozen with liquid nitrogen, the water crystallized into ice rapidly, which resulted in a small pore size. However, when the frozen chitosan microspheres were slightly thawed under specific conditions, the melted ice crystals and chitosan molecules restored the mobility, combined with each other, and finally reformed their structure [26].

Preparation of Porous Hyaluronic Acid Microspheres
As shown in Figure 8 and HA (hydroxyl group). It is known that the porous network can be adjusted by the amount of cross-linking agent [28].

Preparation of Porous Gelatin Microspheres
The SEM images of the obtained porous gelatin microspheres are shown in Figure 10. It can be found that the microspheres have an interconnected porous structure. The pore size range was 20-80 μm, which is suitable for cell attachment. The free water in the gelatin microspheres was frozen, which caused the polymer chains to gather and condense. The mean pore diameter could be controlled by varying the freezing temperature which is related to the cooling rate [29]. since the ice crystal growth rate and the pore diameters are functions of the temperature gradient. The pore size of gelatin microspheres decreased with freezing temperature for -80℃. The free water in the gelatin microspheres was frozen more densely at low temperatures, which caused the polymer chains to gather and condense.

Cell Culture onto Porous Microspheres
To evaluate the efficiency of cell seeding onto porous microspheres formed from the various polymer microspheres, the fluorescence images of Heb3B-GFP attached to the porous microspheres were observed after 7 days of seeding. As time elapsed, the inoculated cells continued to migrate, proliferate, and eventually spread throughout the microspheres. After 7 days, it was found that most cells attached uniformly to the surface and cross section of the microspheres except hyaluronic acid ( Figure 11). In the case of hyaluronic acid, low crosslink-yield induced swelling under cell culture conditions and high crosslink-yield reduced pore size. As shown in Figure 12

Conclusion
In this study, porous microspheres were used to ensure good retention of the engrafted cells as a platform for cell delivery.
The pore size could be controlled, depending on the polymer and method of preparation. This study demonstrates the feasibility of using porous microspheres as a cell culture substrate to expand cells and as a cell transplantation vehicle. These findings may contribute to the development of injectable tissue engineering solutions for therapy and repair of tissue.