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. 2026 Sep 12;26(9):e70257. doi: 10.1002/mabi.70257

Stimulation Based Drug Release From Dual‐Layered Microspheres

Ahmad Zunnu Rain 1, Harini G Sundararaghavan 1,✉
PMCID: PMC13570566  PMID: 42730697

ABSTRACT

The ability to mimic biological release of growth factors during regeneration and repair has important applications in several tissue‐engineered systems. PLGA microspheres hydrolyze in water, resulting in high burst release. Gelatin microspheres maintain controlled release but are susceptible to enzymatic degradation, resulting in nonlinear release. The goal of this project is to develop a bioengineered scaffold that allows sustained, linear growth‐factor release that is fully controlled via electrical stimulation. We have fabricated a hyaluronic acid‐carbon nanotube (HA‐CNT) conductive nanofiber material that is seeded with our dual‐layered PLGA‐gelatin microspheres loaded with growth factors that can be controlled via electrical stimulation to maintain linear release. Electrical stimulation of the HA‐CNT fibers allows for electrophoretic transport of positively charged proteins through the gelatin layer to initiate growth‐factor release, and the PLGA allows continual linearized release. Growth factor release was tested in both fibroblasts and chick dorsal root ganglia model. Both cell types showed activity under electrically released growth factors.

Keywords: biomedical engineering, biophysics, controlled release, gelatin, materials science, nanofiber, plga, stimulation, tissue engineering

The goal of this study is to develop a dual‐layered microsphere system that can be controlled by electrical stimulation and allows for sustained, linear growth factor release. GelMA‐PLGA microspheres are co‐electropsun with aligned HA‐CNT nanofibers to provide topographical cues. A custom electrical stimulation chamber is developed to control growth factor release and provide electrical cues to the cells.

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1. Introduction

Peripheral nerve injuries affect millions of people in the United States annually [1] leading to loss of sensation and eventually loss of function. Current treatments include direct suture repair, autograft/allograft transplantation, and the use of nerve guided conduits [2, 3, 4]. Shorter defects (<5 mm) may be repaired using direct end‐to‐end suturing if they can be properly aligned and reconnected without tension [2]. Critical size defects (>5 mm) cannot be sutured together in this manner and usually require an implant to bridge this gap. The gold standard treatment for these larger defects is the use of an autologous nerve graft, however, this is only partially successful because of size mismatch and donor site morbidity [5, 6]. Several studies have shown that neuronal cells respond positively to chemical, electrical, mechanical, and topographical biomaterial [2, 7, 8, 9]. This study is focused on developing a peripheral nerve graft with topographical, electrical, and chemical cues.

The use of chemical cues has been primarily studied through the utilization of growth factor‐releasing microspheres. Several recent studies show the use of dual‐layered microspheres for sequential growth factor release [10, 11]. Chen et al. developed a GelMA hydrogel‐PLA microsphere composite to deliver dual growth factors for bone regeneration, where the gelatin hydrogel delivered NGF and the internal PLA microspheres exhibited delayed release of Yoda1 similar to our study. However, release was only initiated through hydrolysis [10]. We have previously shown gelatin [12] and Poly(lactic‐co‐glycolic acid) (PLGA) microspheres [8] can enhance neurite growth when combined with aligned hyaluronic acid (HA) nanofibers, but neither is able to give controlled, linear drug release. For nerve regeneration, growth factor release should be slow and continual [13, 14]. Additionally, as found in previous studies, sustained release of growth factors is difficult to achieve using gelatin and PLGA microspheres alone [8, 12, 15, 16, 17]. The goal of this study is to control drug release, particularly to reduce burst release and delay release following the body's initial inflammatory response.

Electrical stimulation therapy can play an important role in promoting neuromuscular functional recovery [18]. Studies have confirmed that utilization of electrical stimulation within safe limits enhances electrical activity of neuronal cells and promotes growth of nerve cells [19, 20, 21]. Neurotrophic factors after electrical stimulation are highly expressed and improve the nerve regeneration microenvironment, accelerating axon outgrowth across the nerve injury site [22, 23, 24, 25]. This study looks at combining this phenomenon with electrically triggered release of growth factors. We have previously developed an electrically conductive fibrous scaffold that can direct neurite growth through stimulation [24]. Several recent studies have looked at electrical stimulation in peripheral nerves, spinal cord, and in neural stem cells [26, 27, 28]. All showed positive results of low‐level stimulation. The goal of this study is to develop and incorporate a dual‐layered microsphere system that can be controlled by electrical stimulation and allows for sustained, linear growth factor release. This is combined with aligned nanofibers for topographical cues and an electrically conductive HA‐CNT scaffold that will also provide electrical cues to the cells.

2. Materials and Methods

2.1. PLGA Microspheres

PLGA microspheres were manufactured using a water/oil/water emulsion of 75:25 PLGA (adopted and adjusted from Whitehead et al.) [8]. Briefly, PLGA was thoroughly dissolved in dichloromethane (DCM) to create the initial PLGA solution. An aqueous solution of 1 mg/mL protein (BSA, NGF, bFGF) and 2% Polyvinyl Acetate (PVA) was suspended in the PLGA solution, creating a loaded aqueous PLGA phase, and sonicated. The solution was dispersed and stabilized in 0.5% (w/v) aqueous PVA followed by 30 s of pulse vortexing to separate the microspheres, resulting in the second oil‐in‐water emulsion, that is stirred for 1 h. The solution was subsequently rinsed with distilled (DI) water two times to remove excess DCM, and the PLGA microspheres were collected via centrifugation for further modification.

2.2. Dual‐Layered Microspheres

Fabrication of the dual‐layered microspheres was accomplished by coating the previously prepared PLGA microspheres with gelatin (type A or B) via three different methodologies: Adsorption, Absorption, and Conjugation.

2.2.1. Adsorption

Gelatin (type A or B) was dissolved in phosphate‐buffered saline (PBS) buffer to create a 1 mg/mL gelatin solution. The collected PLGA microspheres were dispersed into the gelation (type A or B) solutions for 24 h. Once dispersed into the gelatin bath, the PLGA microspheres were covered and left unagitated at room temperature to allow a complete exterior coating (Figure 1A). The gelatin‐coated PLGA microspheres were rinsed with distilled (DI) water two times and crosslinked with 0.1% (w/v) genipin. The resultant microspheres were freeze‐dried and stored at −20°C.

FIGURE 1.

FIGURE 1

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Schematic of different fabrication methods for making dual‐layered microspheres. (A) Adsorption. (B) Absorption. (C) Conjugation.

2.2.2. Absorption

The collected PLGA microspheres were freeze‐dried for 24 h to remove remaining liquid from the porous interior core. These freeze‐dried PLGA microspheres were dispersed into a 1 mg/mL gelatin (type A or B) solution for 24 h. The gelatin was pulled into the interior porous region of the freeze‐dried PLGA microspheres and coated with gelatin from the inside out (Figure 1B). The gelatin‐coated PLGA microspheres were rinsed with water two times crosslinked with 0.1% (w/v) genipin. The resultant microspheres were freeze‐dried and stored at −20°C.

2.2.3. Conjugation

A 1 mg/mL gelatin (type A or B)/PBS buffer solution was treated with 20 mL of 0.5 mg/mL 1‐ethyl‐3‐(dimethylaminopropyl) carbodiimide (EDC) for 30 min to initiate gelatin modification. Simultaneously, the collected PLGA microspheres were dispersed into 20 mL of 0.5 mg/mL EDC for 30 min to initiate surface modification of the PLGA. The microspheres were added to the gelatin (type A or B) solution and left unagitated for 24 h at room temperature. Treatment of EDC created an amide bond between the PLGA and gelatin (Figure 1C). The conjugated gelatin and PLGA microspheres were rinsed with water two times and crosslinked with 0.1% (w/v) genipin. The resultant microspheres were freeze‐dried and stored at −20°C.

2.2.4. Size Determination

The dual‐layered microspheres were imaged using scanning electron microscopy (SEM). The fabricated dual‐layered microspheres were placed onto a 6‐sample holder and gold sputter‐coated for 30 s. Once coated, the samples were loaded into the SEM, imaged, and measured (Image J v1.53e, n = 200).

2.2.5. Gelatin Concentration

Gelatin concentrations of dual‐layered microspheres fabricated with type A and B gelatin via adsorption, absorption, and conjugation were evaluated using a BCA assay (Thermo Scientific Pierce BCA Protein Assay Kit 23225) and plotted against standard absorbance of gelatin using UV spectrophotometry at 540 nm. PLGA microspheres were also evaluated as a control.

2.2.6. Release From Dual‐Layered Microspheres

Fabricated microspheres (0.05 g) were collected in a 12 well‐plate and fully submerged in 1 mL PBS for 24 h to ensure no burst release was occurring. After 24 h, the PBS was collected with a syringe‐filtered pipette (0.08 µm) to ensure no microspheres were removed. 1 mL of calcium (0.05 mg/mL CaCl2) containing PBS with 0.5 mg/mL collagenase was added to each well to trigger the release mechanism from the dual‐layered microspheres (adopted and adjusted from Mladenovska et al.) [29]. The plate was placed in the incubator at 37°C. At set time intervals, the plate was placed on a horizontal orbital shaker for 1 min to thoroughly stir the dispersion. A pipette was used to collect the supernatant. One milliliter of the calcium‐containing PBS with collagenase was added to the wells again until the next collection time. This process was repeated and collected for 7 days. The BSA concentration was observed by using a BCA assay and plotted against the standard absorbance of BSA using UV spectrophotometry at 562 nm [15].

2.3. Scaffold Fabrication

2.3.1. Electrospinning

HA (ECM Science, Detroit, MI) was dissolved in DI water and methacrylated to about 30% methacrylation by adding methacrylic anhydride (Sigma) while maintaining a basic pH (8‐10) using NaOH (as adopted by Steel et al.) [9]. A solution containing 2% methacrylated HA, 2% polyethylene oxide (PEO, 900 kDa), 0.05% Irgacure 2959 crosslinker, and 0.01% multi‐walled carbon nanotubes (CNTs) (all w/v) was thoroughly dissolved to create the electrospin solution. The 0.05 g of the manufactured dual‐layered microspheres were added to the fiber spin solution before electrospinning. Aligned HA‐CNT fibers were prepared via electrospinning the fiber spin solution on a rotating mandrel (1400 rpm) with a flow rate of 0.9 mL/hr and a voltage of 27 kV from a blunt 18‐gauge needle, 11 cm away from the mandrel (Figure 2). The HA‐CNT fiber spin solution was electrospun on methacrylated 12 mm square coverslips. All nanofibers spun on the coverslips were removed carefully and UV‐crosslinked for 30 min prior to storage at room temperature in a desiccant chamber.

FIGURE 2.

FIGURE 2

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Schematic of the electrospinning process from syringe pump to rotating mandrel.

2.3.2. Nanofiber Characterization

The bioengineered nanofibers were characterized by SEM imaging. The electrospun samples were placed on a sample holder (6‐mount specimen holder) and gold sputter‐coated for 30 s. Images were captured at 1500 and 2000 magnifications. Scaffold nanofiber diameter and angle were measured using ImageJ (v1.53e) with at least 200 nanofibers per microsphere and fiber type.

2.4. Release Profile From HA‐CNT Scaffold via Enzymatic Trigger

This procedure largely follows the release profile collection described above. 12 mm coverslips with the electrospun scaffolds were placed in a 12 well‐plate and fully submerged in 1 mL PBS for 24 h to ensure there was no burst release. This process was repeated for 10 days.

2.5. Electrical Stimulation

The custom electrical stimulation plate was designed using 12‐well cell culture plates as previously described by Steel et al. [9]. (Figure 3A). The copper electrodes were never in contact with the culture media, but only with the nanofibers that were electrospun onto the coverslips. A function generator acted as the power source for electrical stimulation and was connected to a breadboard with 2 binding posts, 7.25“ × 7.5” (Jameco). Biphasic stimulus (voltage in) and the stimulus after passing through the plate (voltage out) were captured from Channel 1 and Channel 2 of the oscilloscope (Rigol) that was connected to a Raspberry Pi running a Python script to store data in real time via a USB port. The entire custom plate was connected in series to the function generator using alligator clips at the copper tape electrode ends of the plate (Figure 3). All connections were shielded via liquid electrical tape followed by standard electrical tape to prevent leaking, or a voltage drop throughout the plate. Electrical stimulation regiments of 50, 75, 100, and 125 mV/mm were charge‐balanced using 20 Hz biphasic square waves of ±25 ms pulse width and a ±50% duty cycle.

FIGURE 3.

FIGURE 3

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Schematic of custom electrical stimulation culture plate. (A) Illustration of entire stimulation plate. (B) Illustration of each individual well (I) 12‐well plate with 6 mm holes drilled through the bottom of well, (II) high acrylic adhesive, (III) copper tape overlaying (IV) electrospun nanofibers on coverslip.

2.5.1. Release Profile From HA‐CNT Scaffold Via Electrical Stimulation

Prior to electrical stimulation, all of the nanofibers in the 12‐well plate were hydrated using 1× PBS for 24 h to ensure there was no burst release. The plate was placed in the incubator at 37°C to emulate in vivo conditions. After 24 h, the PBS was collected and replaced with 1 mL of fresh PBS. Electrical stimulation was applied across the plate at different intervals to evaluate how the electrospun nanofibers seeded with the dual‐layered microspheres would react in terms of release. One time 30‐min stimulation, 2‐min stimulation every set interval of time, one‐time 5‐min stimulation, and 5‐min stimulation every 24 h were tested to evaluate release from the electrospun nanofibers. After each time interval, the supernatant was eluded and replaced with fresh PBS. This process was repeated for 14 days. BSA concentration was observed using a BCA assay and plotted against the standard absorbance of BSA using UV spectrophotometry at 562 nm.

2.5.2. NGF ELISA

NGF was loaded into the microspheres during fabrication instead of BSA and electrospun onto the HA‐CNT nanofibers. Stimulation plates were made as described above. Electrical stimulation was applied to the plate at different intervals, and fresh media was replaced after the supernatant was collected. ELISA was used to calculate the concentration of NGF release over 14 days. Standard curves were prepared with known concentrations of NGF dissolved in DMEM and plotted via linear regression to calculate the concentration of NGF in the supernatant.

2.6. L929 Fibroblast Cell Culture

Coverslips of the electrospun nanofibers with dual‐layered microspheres loaded with bFGF (1 mg/ml) were placed in the custom stimulation plate and fully submerged in L929 media (DMEM media with 10% FBS). Electrical stimulation was then applied across the nanofibers using a function generator connected to the copper electrodes of the custom stimulation plate. Electrical stimulation regiments of 50, 75, 100, or 125 mV/mm were applied across the wells. The plate was subsequently placed in an incubator and left overnight for 24 h. Fibroblasts (L929) were seeded (8000 cells/cm2) onto wells of a 12‐well cell culture plate in 500 µL of L929 media and incubated for 2 h to allow cellular adhesion to the individual wells. 1 mL of electrically stimulated media from the custom stimulation plate was collected 24 h following stimulation and placed into the fibroblast‐seeded 12‐well plate. Fresh L929 media was added to the stimulation plate, and electrical stimulation was administered. Both plates were returned to the incubator until the next stimulation time point 24 h later. This process was repeated for two additional 24‐h time periods and the fibroblasts were fixed after 72 h in culture. Unstimulated fibroblast cultures and HA‐CNT nanofiber scaffolds with empty microspheres and without dual‐layered microspheres using the same methodology were used as controls, as explained above. Cellular proliferation was observed via AlamarBlue to evaluate fibroblast growth after 72 h in culture. Phalloidin staining, followed by DAPI (1:1000, Fisher) was used to label the samples, and samples were imaged using microscopy (Nikon Eclipse Ti).

2.7. In Vitro Neuron Cell Culture

The following in vitro neural cell culture was similar to the fibroblast cell culture protocol outlined above. The bioengineered nanofibers were electrospun with dual‐layered microspheres loaded with NGF (1 mg/ml) on coverslips, which were placed in the stimulation plate and fully submerged in NGF media composed of 1:1 DMEM/F‐12 supplemented with 50 ng/mL NGF. The stimulation plate was placed in the incubator and left overnight for 24 h. Lumbar dorsal root ganglia (DRG) were dissected from E11 chick embryos as previously described by Whitehead et al. [30]. The DRGs were dissociated and seeded (8000 cells/cm2) onto collagen‐coated (25 µg/mL Collagen type I dissolved in 0.01 M acetic acid) wells of a 12‐well cell culture plate in 500 µL of media and incubated for 2 h to allow adhesion to the wells. The 1 mL of electrically stimulated media from the stimulation plate was collected 24 h following stimulation and placed into the DRG‐seeded 12‐well cell culture plate. Fresh NGF media was added back to the stimulation plate, and electrical stimulation was administered (50, 75, 100, or 125 mV/mm) again. Both plates were then returned to the incubator until the next stimulation 24 h later. This process was repeated for two more 24‐h periods. The neurons were fixed after 72 h in culture. The electrical stimulation regimes studied were 50, 75, 100, and 125 mV/mm applied across the HA‐CNT scaffold seeded with dual‐layered microspheres. Unstimulated neuronal cultures and HA‐CNT nanofiber scaffolds without dual‐layered microspheres using the same configuration were used as controls. Neurons were labeled with primary antibody anti‐neurofilament 200 (1:400, Sigma), followed by secondary antibody Alexa Fluor 488 goat anti‐mouse IgG (1:1000, Invitrogen), and DAPI (1:1000, Fisher). The samples were then imaged and quantified using microscopy for neurite length and number of neurite‐bearing neurons, using Image J.

3. Results

3.1. Microsphere Fabrication

SEM was performed on the microspheres to evaluate surface properties and size. (Figure 4). Dual‐layered microspheres fabricated via adsorption (Figure 4A), absorption (Figure 4B), and conjugation (Figure 4C) were evaluated for surface properties and size to determine the best‐fit dual‐layered microsphere for further testing, electrospinning, and growth factor release. Dual‐layered microspheres fabricated via adsorption, absorption, and conjugation produced spheres that were 22.70 ± 1.13 µm, 36.23 ± 1.29 µm, 24.75 ± 1.26 µm, respectively (Figure 4G). Microspheres that were used in previous studies were also produced as controls to evaluate the differences in size and morphology. Gelatin type A (Figure 4D), gelatin type B (Figure 4E), and PLGA (Figure 4F) microspheres were fabricated with average sizes of 52.18 ± 1.84 µm, 54.49 ± 2.22 µm, 20.23 ± 0.91 µm, respectively.

FIGURE 4.

FIGURE 4

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Surface morphology and size differences between the various fabrication methods used to produce the microspheres. Dual‐layered microspheres fabricated via (A) adsorption, (B) absorption, and (C) conjugation (scale bar = 50 µm, 250× magnification). Control microspheres produced via single‐material fabrication; (D) gelatin type A, (E) gelatin type B, and (F) PLGA (scale bar = 10 µm, 2500× magnification). (G) Table and (H) graph depicting the average size with error of the different fabrication methods used to make the microspheres. * Indicates p < 0.01.

The differences in the fabricated microsphere diameter were quantified in Figure 4H. It is evident that the dual‐layered microspheres that were made via adsorption, absorption, and conjugation were significantly smaller than the two gelatin control microspheres because they were fabricated with an inner PLGA core that was coated with a gelatin layer. The dual‐layered microspheres fabricated via absorption were significantly larger than the other fabrication methods due to gelatin being pulled into the PLGA core during the production process. This resulted in much larger microspheres as compared to the other two fabrication methods.

In addition to surface morphology and comparing gelatin microspheres to the fabricated dual‐layered microspheres, gelatin concentration was also evaluated to ensure the microspheres produced via adsorption, absorption, and conjugation were in fact coated with gelatin (Figure 5A) The microspheres fabricated via absorption had the highest concentrations of gelatin at 461.94 ± 97.56 µg/mL and 404.02 ± 61.30 µg/mL for gelatin type A and B, respectively. Gelatin was absorbed by the lyophilized inner PLGA core and coated from the inside out, resulting in higher concentrations of gelatin and swollen, and nonspherical microspheres. There was no significant difference observed between the dual‐layered microspheres fabricated via adsorption and conjugation. The microspheres made by adsorption yielded gelatin concentrations of 89.89 ± 44.92 µg/mL and 57.40 ± 47.36 µg/mL for gelatin type A and B, respectively. In addition, the conjugated dual‐layered microspheres yielded gelatin concentrations of 141.47 ± 41.37 µg/mL and 110.74 ± 54.70 µg/mL for spheres made with gelatin type A and B, respectively. There was no significant difference between the microspheres made with gelatin type A or B. As a control, we measured protein content in our PLGA‐only microspheres, and there was no gelatin present. The dual‐layered microspheres fabricated via conjugation were used throughout the following steps in this study.

FIGURE 5.

FIGURE 5

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Microsphere characterization and BSA release. (A) Gelatin concentration (in µg/mL) was measured via a BCA assay for the different fabrication methods. The dual‐layered microspheres fabricated via absorption had significantly higher concentrations of gelatin for spheres made with type A and B gelatin, respectively, than those made via adsorption and conjugation. Microspheres made via adsorption and conjugation had no significant difference in gelatin concentration. There was no significant difference in gelatin concentration between the microspheres made with gelatin type A or B. (B) Cumulative BSA release was observed from the different microspheres fabricated via conjugation. Collagenase was added as a trigger to initiate BSA release from the microspheres. PLGA microspheres had an initial burst release as they were not coated with gelatin. The dual‐layered microspheres had significantly more linearized release as compared to the controls and were not susceptible to burst release. * Indicates p < 0.05.

BSA release was observed in Figure 5B. Burst release was observed from the PLGA microspheres (control) before the collagenase was added. In contrast, the gelatin microspheres (control) and the dual‐layered microspheres did not display this burst release. BSA release was only initiated after the collagenase was used to trigger release. The dual‐layered microspheres had significantly more linearized release profiles as compared to the controls of the microspheres made with gelatin (type A and B) and PLGA, as shown in Figure 5B. Linearization for the dual‐layered microspheres started almost immediately after the addition of the collagenase smart trigger at 26 h. This linearized release was maintained until around 168 h for both types of dual‐layered microspheres. BSA release for all microspheres plateaued at around 75% cumulative release. There was no significant difference between the BSA release of dual‐layered microspheres made with gelatin type A or B.

Since there was no significant difference between gelatin type A or B in terms of concentration of gelatin and cumulative BSA release, fabrication of the dual‐layered microspheres was performed with an inner PLGA core coated with gelatin type A. For this study, gelatin type A was used because an overall positive charge can be favorable for neural outgrowth and protein interactions, however, either type of gelatin can be used in future studies.

3.2. Nanofiber Scaffold Fabrication

The dual‐layered microspheres made via conjugation were electrospun with HA to produce our bioengineered scaffold, as can be seen in Figure 6B. The microspheres were embedded into the HA scaffold without altering the alignment and direction of the nanofibers. Figure 6A shows a SEM image of our electrospun HA scaffold without the microspheres, while Figure 6B, shows the electrospun HA with our manufactured dual‐layered microspheres. The addition of the microspheres in the spin solution does not alter the size of the nanofibers formed and maintains the exact same properties of the original nanofibers.

FIGURE 6.

FIGURE 6

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SEM images and release profile for our bioengineered scaffold electrospun with dual‐layered microspheres. SEM images of HA (A) and HA electrospun with dual‐layered microspheres (B) (scale bar = 10 µm, 2000× magnification). (C) Cumulative BSA release using collagenase as a smart trigger for different types of microspheres electrospun onto HA nanofibers. Linearization is achieved by the scaffold that has the dual‐layered microspheres. * Indicates p < 0.05.

Cumulative BSA release was observed from the electrospun scaffold with collagenase as a smart trigger to initiate the release of BSA. HA was electrospun with microspheres made from gelatin type A and PLGA as controls. Figure 6C shows the three different release profiles from the three materials manufactured. The electrospun HA nanofibers with PLGA showed an initial burst release as soon as it was exposed to PBS. In contrast, the gelatin type A and dual‐layered microsphere scaffolds required collagenase to initiate release. HA nanofibers electrospun with dual‐layered microspheres had a more linearized release profile as compared to that of the controls. The burst release was mitigated, and linearization was achieved.

3.3. Electrical Stimulation Parameters

Electrical stimulation was used, instead of a collagenase trigger, to initiate release from the bioengineered scaffolds. The different electrical stimulation regimens were evaluated as can be shown in Figure 7. one‐time 30‐min [12, 24] and 5‐min electrical stimulation (Figure 7A,B), 2‐min electrical stimulation every time interval (Figure 7C), 5‐min electrical stimulation applied every 24 h (Figure 7D), and 5‐min electrical stimulation applied every 48 h (Figure 7E). Once the ideal electrical stimulation parameters were determined, NGF was loaded into the dual‐layered microspheres, electrospun onto the HA nanofibers, and release was evaluated using the determined ideal electrical stimulation regimen (Figure 7F).

FIGURE 7.

FIGURE 7

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Cumulative BSA and NGF release over a span of 14 days with different electrical stimulation regiments. The yellow dotted lines represent when electrical stimulation was applied across the scaffolds. Cumulative BSA release was observed when applying (A) a one‐time 30‐min electrical stimulation, (B) a one‐time 5‐min electrical stimulation, (C) multiple 2‐min electrical stimulations at varying intervals, (D) multiple 5‐min electrical stimulations every 24 h, and (E) 5‐min electrical stimulation every 48 h across our bioengineered scaffolds. (F) Cumulative NGF release was observed applying multiple 5‐min electrical simulations every 24 h. Voltages of 50, 75, 100, and 125 mV/mm were applied for all observed releases (A–F). Release was observed in ng/mL.

3.3.1. The 30‐Min One‐Time Electrical Stimulation

One‐time 30‐min electrical stimulation (50, 75, 100, and 125 mV/mm) across the HA‐CNT nanofiber scaffolds seeded with dual‐layered microspheres at different levels of stimulation was observed via cumulative BSA release (Figure 7A). There was significant burst release until around the 30‐min time point, followed by linear release that can be seen after the electrical stimulation was applied for all voltages applied, followed by a plateau. For 50 and 75 mV, the release from the scaffold was incomplete as compared to the 100 and 125 mV releases. 50 mV and 75 mV displayed cumulative BSA releases of 139.60 ng/mL and 181.52 ng/mL, respectively, instead of close to 300 ng/mL that was observed at 100 mV and 125 mV of 311.75 ng/mL and 329.77 ng/mL, respectively (Figure 7A).

3.3.2. The 5‐Min One‐Time Electrical Stimulation

The 5‐min electrical stimulation (50, 75, 100, and 125 mV/mm) was applied once, 24 h after the scaffolds were fully submerged in 1× PBS. The 100 and 125 mV/mm data readings still had a burst release, but it was significantly lower than that found in Figure 6A. The cumulative BSA release from 50, 75, 100, and 125 mV/mm was 125.49 ng/mL, 148.05 ng/mL, 216.10 ng/mL, and 228.29 ng/mL, respectively (Figure 7B).

3.3.3. Multiple 2‐Min Electrical Stimulations at Varying Time Intervals

Two‐minute electrical stimulation was applied across the scaffold every time electrical stimulation was administered at varying time intervals (Figure 7C). The yellow‐dotted lines at every time point in Figure 7C represent 2‐min of electrical stimulation that was administered across the scaffold at various times to observe cumulative BSA release. Due to electrical stimulation at every time point, the scaffolds at all voltage levels displayed complete release around 300 ng/mL, as compared to the release in Figure 7A,B. The cumulative BSA release at 50, 75, 100, and 125 mV/mm was 322.14 ng/mL, 322.91 ng/mL, 308.53 ng/mL, and 313.51 ng/mL, respectively. The 2‐min electrical stimulation proved to be too high, and the microspheres were visually starting to erode from the scaffold at the higher voltages of 100 and 125 mV/mm (Figure 7C).

3.3.4. The 5‐Min Stimulations at Every 24‐h Interval

Electrical stimulation across the scaffolds was tested at a 5‐min stimulation that was applied every 24 h. (Figure 7D). The yellow‐dotted lines at every 24‐h interval in Figure 7D represent 5‐min electrical stimulation. The burst release that was observed in Figure 7A at 100 mV and 125 mV was significantly mitigated, and the release was much more linearized. In addition, complete cumulative BSA releases of 277.24 ng/mL and 326.18 ng/mL were achieved at 100 mV and 125 mV, respectively, as compared to Figure 7A,B as can be seen by the initiation of plateauing in Figure 7D. BSA release at 50 mV and 75 mV was still relatively incomplete at 138.89 ng/mL and 244.68 ng/mL, respectively. Varying the voltage of electrical stimulation that was applied proved that release from the scaffold was controllable.

3.3.5. The 5‐Min Stimulations at Every 48‐h Interval

To evaluate if release from the microspheres was controlled by the voltage applied, electrical stimulation was applied every 48 h for 5 min. This additional 24‐h interval was introduced to evaluate the ability of the microspheres to stop growth factor release on their own after electrical stimulation was applied across the scaffold. The yellow‐dotted lines at every 48‐h interval in Figure 7E represent 5‐min electrical stimulation. The releases observed for all voltages applied showed plateauing before electrical stimulation was reapplied. The releases observed at 100 mV and 125 mV were relatively linearized and plateaued at 285.53 ng/mL and 327.59 ng/mL, respectively. BSA release at 50 mV and 75 mV was still relatively incomplete at 141.52 ng/mL and 272.47 ng/mL. This meant that if different concentrations of release were required for any application, our scaffold has the capacity to adjust based on the voltage applied.

3.3.6. NGF Release

NGF was loaded into the dual‐layered microspheres and electrospun onto the HA‐CNT nanofibers. The 5‐min electrical stimulation was applied across the scaffold every 24 h, and cumulative NGF release was observed via ELISA (Figure 7F). The yellow dotted lines at every 24‐h interval depict the 5‐min electrical stimulation. The cumulative NGF releases at every voltage level maintained a nearly identical cumulative BSA release rate as shown in Figure 7D (NGF release had lower cumulative release amounts since less NGF was loaded into the dual‐layered microspheres). Nearly complete cumulative NGF release was observed for 100 and 125 mV/mm at 48.82 ng/mL and 49.95 ng/mL (Figure 7F), respectively. In addition, relatively incomplete cumulative NGF release was observed for 50 and 75 mV/mm at 18.82 ng/mL and 30.87 ng/mL (Figure 7F), respectively. These findings followed the same trend that was observed in the cumulative BSA release in Figure 7D.

3.4. Electrical Stimulation via Cellular Models

3.4.1. Electrical Stimulation of L2929 Fibroblasts

The dual‐layered microspheres were loaded with bFGF and electrospun onto HA‐CNT nanofibers to create the scaffold that will be utilized for the electrical stimulation of the L929 fibroblasts. A control well with no electrical stimulation was used to evaluate fibroblast proliferation without any external stimuli (Figure 8A). Cellular proliferation data were obtained after 72 h in culture as observed through alamarBlue (Figure 8F). Fibroblasts that were subject to 100 mV/mm of stimulated media displayed the highest amount of growth overall, with about 206.28 × 103 cells. The next highest in terms of cellular proliferation were the fibroblasts that were subject to 75 mV/mm of stimulated media with about 176.19 × 103 cells, followed by the fibroblasts that were subject to 125 and 50 mV/mm, which had 169.09 × 103 cells and 139.28 × 103 cells, respectively. Surprisingly, 50 mV/mm condition did not have significantly higher proliferation than the two controls without any electrical stimulation.

FIGURE 8.

FIGURE 8

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Depiction of L929 fibroblasts after 72 in culture. Primary phalloidin stain, followed by DAPI was performed on all of the samples. L929 fibroblast growth of (A) control on plastic, (B) 50 mV/mm, (C) 75 mV/mm, (D) 100 mV/mm, and (E) 125 mV/mm was performed via microscopy (scale = 200 µm, 20× magnification). (F) Cellular proliferation of the L929 fibroblasts via alamarBlue. * Indicates p < 0.05.

3.4.2. Electrical Stimulation of DRGs

Neurite outgrowth was observed after 72 h via microscopy. A control well with HA‐CNT nanofibers without the dual‐layered microspheres was used in the stimulation well plate, and 100 mV/mm was used to evaluate neurite outgrowth (Figure 9A). The differing voltage levels of 50 (Figure 9B), 75 (Figure 9C), 100 (Figure 9D), and 125 (Figure 9E) mV/mm were evaluated for neurite outgrowth. The electrically stimulated DRGs at 100 and 125 mV/mm had significantly higher levels of growth than the other stimulation conditions of around 280.66 µm and 325.91 µm, respectively. Additionally, 50 and 75 mV/mm also proved to have significantly larger neurite outgrowths of 184.68 µm and 206.21 µm, respectively, than the control well on plastic, which only had 91.43 µm of neurite outgrowth (Figure 9F).

FIGURE 9.

FIGURE 9

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E11 DRG neurite outgrowth after 72 h in culture. Primary antibody anti‐neurofilament 200, followed by secondary antibody AlexaFlour 488 goat anti‐mouse IgG, and DAPI was performed on all samples. Neurite outgrowth of (A) control on plastic, (B) 50 mV/mm, (C) 75 mV/mm, (D) 100 mV/mm, and (E) 125 mV/mm was performed via microscopy (scale = 100 µm, 20x magnification). (F) Comparison of neurite outgrowth at different levels of electrical stimulation. * Indicates p < 0.05.

3.5. Statistical Methods

All data are reported as mean ± standard deviation unless otherwise specified. For experiments with 2 or more groups, statistical analysis was performed using RStudio for 2‐way and 3‐way ANOVA, followed by Tukey posthoc. Significance was accepted at * p < 0.05 unless otherwise stated.

4. Discussion

In this study, we explored three different fabrication methodologies (adsorption, absorption, and conjugation) for making our dual‐layered microspheres with gelatin type A and B. We completed microsphere characterization and conducted drug release studies to determine the ideal fabrication method. Based on SEM imaging, size and surface morphology were determined for the dual‐layered microspheres for all three fabrication methods for gelatin type A and B (Figure 4). We concluded that the dual‐layered microspheres that were fabricated via conjugation with gelatin type A yielded the best‐suited microspheres for electrospinning and our scaffold manufacturing methodology. The chemically conjugated amide bonds can maintain higher stability under acidic and basic conditions and can withstand the high temperatures that occur during the electrospinning process [31, 32]. Drug release studies showed that (Figure 5). the dual‐layered microspheres were significantly better than the single‐layered microspheres in terms of linearizing release and preventing an initial burst release from occurring. Drug release profiles did not change drastically when electrospun on HA nanofibers (Figure 6), and we tested a wide range of electrical parameters in Figure 7.

We concluded that 100 mV/mm is the ideal voltage at which release is the most linearized while minimizing burst release (Figure 7). Over the 2 week time period measured, we do have some ability to control the release of growth factors and, at lower stimulation levels, we see that microspheres still have growth factors loaded that can be released at a future time point. Nerve regeneration occurs over several months, and therefore, we could use this system to delay growth factor release.

Several recent papers have described the phenomenon of electrophoretic transport of positively charged proteins through conductive GelMA [33, 34, 35]. Electrophoretic delivery involves applying voltage to push charged proteins through GelMa meshes to increase the diffusion rate. Cabot et al. showed the ability to push charged molecules through a 5% GelMA mesh by measuring fluorescein flow through microscopy. By varying the electric field and the concentration of GelMA in the mesh, they showed control of molecular flow [33]. Cheah et al. used a similar approach to develop a conductive scaffold with GelMA and poly(3,4‐ethylenedioxythiophene) where they were able to control protein delivery by varying electrical stimuli [34]. Chen et al. showed that the composite hydrogel of gelatin cryogel and PEDOT:PSS (conductive material) could be used to deliver drugs through electrically triggered release [35]. In this study, we move this work one step further by incorporating protein‐loaded GelMA onto electrically conductive nanofibers giving us better control of topography and the amount of protein (microspheres) incorporated.

To evaluate our scaffold's ability to maintain cellular growth, we used L929 fibroblasts to observe cellular proliferation (Figure 8). We used stimulated FGF media that was released using the multiple 5‐min electrical stimulation regiment every 24‐h to evaluate cellular proliferation via alamarBlue at varying voltages (50, 75, 100, and 125 mV/mm). We concluded that 100 mV/mm resulted in the highest cellular proliferation after 72 h in culture. Next, we used stimulated NGF media that was released using multiple 5‐min electrical stimulations every 24‐h to observe neurite outgrowth in E11 chick DRGs at varying voltages (50, 75, 100, and 125 mV/mm) as can be seen in Figure 9. We observed that 100 and 125 mV/mm produce the longest neurite outgrowth of DRGs after 72 h in culture. The ability to control growth factor release and delay release could help guide regenerating axons along the neural pathway to promote proper reinnervation [36, 37].

Several previous studies have investigated nerve guided conduits constructed via electrospun fibers [8, 12, 24, 30, 38]. Work done by Zhu et al. suggested that utilizing NGF concentration gradients direction on their nanofibers provided chemotactic cues for accelerating cellular migration of sciatic nerves [39]. However, there is no control over delivery. In addition, work done by Xue et al. cited peripheral nerve growth on aligned nanofibers coated with particles consisting of a mixture of collage and fibronectin in a bidirectional gradient promoted extension of neurites from a DRG body in the direction of increasing particle density [40]. Here, they are utilizing particles that facilitate the directional growth of the DRGs, similar to our study, however, this is a surface coating and does not involve sustained protein delivery.

Studies have been conducted to demonstrate the in vivo feasibility of extended 60‐min electrical stimulation to significantly accelerate axonal outgrowth [41, 42, 43]. Additionally, others have seen the positive effects of short‐term electrical stimulation that implicates enhanced growth initiation. These studies suggest that short‐period, lower‐frequency electrical stimulation could potentially be a new therapeutic approach to accelerate neural regeneration after injury and ultimately improve functional recovery [41, 44]. To our knowledge, our results provide the first in‐depth look at an in vitro demonstration of dual‐layered microspheres facilitating drug release via electrical stimulation to aid in neural regeneration. We propose that the HA‐CNT nanofiber scaffold seeded with our bioengineered dual‐layered microspheres can maintain controlled release of growth factors to aid in neural regeneration for at least 14 days in culture (Figure 7).

The HA‐CNT nanofibers developed in this study only degrade under enzymatic degradation, as is common for methacrylated HA‐based biomaterials [45]. One of the limitations to consider is the use of CNTs in our biomaterial and cellular toxicology concerns [46]. However, the toxicity concerns are minimized by our fabrication process [9, 24] and ensuring to eliminate the catalyst impurities that exist in the CNTs. In our study, we have not observed any cytotoxic effects after 48 h in culture for both the L929 fibroblast and DRGs cultured with stimulated media released from our scaffold. We are also limited by the length of the study. Though we only looked at electrical stimulation over 14 days, we could consider reducing the stimulation and extending the study over several months, which would be more in line with nerve regeneration in vivo. We are interested in further understanding the electrophysiology involved in protein release and neurite behavior and will investigate this in future studies.

5. Conclusions

In conclusion, we have demonstrated that multiple 5‐min electrical stimulations every 24 h delivered through our HA‐CNT nanofibers seeded with dual‐layered microspheres can significantly enhance neural outgrowth over HA nanofiber and plastic controls. We have demonstrated that applying at least 100 mV/mm of electrical stimulation through our HA‐CNT nanofibers can linearize release and help improve neuron growth relative to unstimulated and lower levels of electrical stimulation. Future work will quantify the mechanical properties of the bioengineered nanofiber scaffold that we have developed. We plan to further understand neuronal activity under electrical stimulation and NGF exposure through electrophysiological studies (Supporting Information).

Author Contributions

Harini G. Sundararaghavan: conceptualization, funding acquisition, Writing – review and editing, visualization, project administration, supervision, resources. Ahmad Zunnu Rain: investigation, writing – original draft, data curation, formal analysis, writing – review and editing.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: mabi70257‐sup‐0001‐SuppMat.docx.

MABI-26-e70257-s001.docx (174.5KB, docx)

Acknowledgements

This work was supported by the WSU Graduate School, the NF Michigan Foundation, and partially funded by a Department of Defense grant (W81XWH221056).

Data Availability Statement

The data that support the findings of this study are openly available in DRYAD.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting File: mabi70257‐sup‐0001‐SuppMat.docx.

MABI-26-e70257-s001.docx (174.5KB, docx)

Data Availability Statement

The data that support the findings of this study are openly available in DRYAD.

Articles from Macromolecular Bioscience are provided here courtesy of Wiley

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