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Published in final edited form as: ACS Macro Lett. 2024 Jul 18;13(8):959–965. doi: 10.1021/acsmacrolett.4c00240

Integration of Melt Electrowritten Polymeric Scaffolds and Bioprinting for Epithelial Healing via Localized Periostin Delivery

Nileshkumar Dubey 1,▽, Maedeh Rahimnejad 2,▽, W Benton Swanson 3, Jinping Xu 4, Mylène de Ruijter 5, Jos Malda 6, Cristiane H Squarize 7, Rogerio M Castilho 8, Marco C Bottino 9
PMCID: PMC13020899  NIHMSID: NIHMS2154584  PMID: 39024469

Abstract

Management of skin injuries imposes a substantial financial burden on patients and hospitals, leading to diminished quality of life. Periostin (rhOSF), an extracellular matrix component, regulates cell function, including a proliferative healing phase, representing a key protein to promote wound healing. Despite its proven efficacy in vitro, there is a lack of scaffolds that facilitate the in situ delivery of rhOSF. In addition, there is a need for a scaffold to not only support cell growth, but also to resist the mechanical forces involved in wound healing. In this work, we synthesized rhOSF-loaded mesoporous nanoparticles (MSNs) and incorporated them into a cell-laden gelatin methacryloyl (GelMA) ink that was bioprinted into melt electrowritten poly(ε-caprolactone) (PCL) microfibrous (MF-PCL) meshes to develop mechanically competent constructs. Diffraction light scattering (DLS) analysis showed a narrow nanoparticle size distribution with an average size of 82.7 ± 13.2 nm. The rhOSF-loaded hydrogels showed a steady and controlled release of rhOSF over 16 days at a daily dose of ~40 ng/mL. Compared with blank MSNs, the incorporation of rhOSF markedly augmented cell proliferation, underscoring its contribution to cellular performance. Our findings suggest a promising approach to address challenges such as prolonged healing, offering a potential solution for developing robust, biocompatible, and cell-laden grafts for burn wound healing applications.

Graphical Abstract

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Skin injuries, such as burns, compromise the skin’s primary role of protecting the body against biological, physical, and chemical insults by impairing the functional epidermal barrier.1 Among these injuries, burns afflict more than 1 million Americans annually and rank as the seventh leading cause of injury and death in the United States.2 After wounding, the deposition of a provisional matrix and the coordinated response of the epidermis, including its stem cells, are required for rapid resurfacing and re-establishment of the epidermal barrier.3 The challenge of managing burn wounds includes the limited amount of available donors, along with continuous discomfort and consequent morbidity.3 Regenerative medicine and tissue engineering approaches potentially tackle these challenges by facilitating rapid restoration of the epidermal barrier without the need for donor tissue.

From a molecular signaling perspective, we aim to reactivate the local PI3K signaling pathway through the localized delivery of recombinant human Osteoblast-Specific Factor-2 (rhOSF/Periostin). The PI3K signaling is a potent molecular circuitry involved in skin homeostasis and tissue regeneration.4 Squarize and Castilho have shown that activation of the PI3K pathway directly impacts the mTOR signaling, an integral component of skin closure during wound healing.5 We also previously characterized the ability of rhOSF to activate the mTOR pathway.6 Periostin (rhOSF), an extracellular matrix (ECM) component, has emerged as a key player in skin wound healing and scar formation.7 It is expressed in various normal tissues, often associated with fibroblasts, and is involved in remodeling the ECM environment.8 rhOSF is up-regulated after skin injury and is considered one of the essential mediators of skin wound healing. It peaks in expression around 7 days after injury.9 Previous studies have shown that rhOSF promotes the proliferation and differentiation of various cells in the wound, leading to appropriate re-epithelialization and myofibroblast differentiation in the granulation tissue, thereby facilitating wound closure.10 Developing complex 3D living scaffolds for the sustained delivery of rhOSF to activate mTOR may represent a significant advancement in burn wound management.

Despite five decades of advancements in skin tissue constructs, several unresolved challenges related to physiological structure and function, mechanical strength, and the development of readily accessible and reproducible constructs for research and clinical applications persist.1 Recently, there has been growing interest in using 3D bioprinting technologies to engineer biologically functional and highly organized tissue constructs in custom-designed patterns using biomaterials (e.g., hydrogels) to deliver cells and biomolecules. While hydrogels have demonstrated a significant capability in fostering extensive cellular growth, their intrinsic fragility renders them unsuitable for applications involving mechanical forces, which are prevalent during wound healing.6 To overcome this disadvantage, we have proposed a reinforcing approach using biodegradable MF-PCL scaffolds integrated into hydrogels to address this fundamental problem.11,12 Moreover, the evolving landscape of 3D printing explores the convergence of different biofabrication technologies to produce tissue constructs to mimic native ECM features to restore damaged tissues effectively.13 Our scaffold incorporates MEW fibers to emulate fibrous proteins, a GelMA hydrogel to mimic water content, and mesoporous silica nanoparticles loaded with periostin to represent biomolecular components typically found in ECM structures. This convergence of techniques enhances the reproducibility and scalability across batches, thereby saving time when fabricating biological constructs. It facilitates the development of tissue constructs and biomedical devices tailored to patient-specific needs. Building on this evidence, we postulate that a converged biofabrication approach, combining MEW and droplet bioprinting, could result in engineered skin constructs integrated with rhOSF-loaded MSNs.11,12 We aim to engineer biodegradable MF-PCL scaffolds housing rhOSF-loaded MSNs and epidermal cells encapsulated within a GelMA hydrogel, a light-sensitive polymerizable hydrogel composed of altered natural extracellular matrix (ECM) constituents,14 to develop mechanically competent epidermal constructs for effective wound management.

In this study, we first investigated the size distribution of the nanoparticles and the release profile of rhOSF from drug-loaded mesoporous silica nanoparticles (MSNs). Proteins are crucial in various clinical applications, such as wound healing.15 However, the widespread use of proteins for therapeutic purposes could be improved by their inherent challenges, such as low stability and large size, which can compromise their therapeutic efficacy.15 Effectively delivering active proteins and drugs to specific target sites in a controlled manner is challenging. To overcome these hurdles, nanoparticle-based delivery systems are a promising solution. Among these, MSNs stand out due to their exceptional biocompatibility, high stability, rigid framework, well-defined pore structure, easily controllable morphology and size, and tunable surface chemistry.15

MSNs have emerged as a desirable option for protein delivery. In this regard, rhOSF and FITC-BSA-loaded MSN were synthesized utilizing a sol–gel emulsion method,16 with FITC-BSA serving as a model drug to assess the release profile of rhOSF. The loaded MSN were isolated through centrifugation, and their surface morphology was characterized using scanning electron microscopy (SEM; Figure 1A). The SEM images revealed well-defined spherical structures on a scale of 100 nm, indicating the successful fabrication of MSNs. Additionally, the size distribution of the nanoparticles was determined by diffraction light scattering (DLS) (Figure 1B). A narrow size distribution was demonstrated with an average of 82.7 ± 13.2 nm, emphasizing the uniformity of the synthesized MSNs. Internalization of FITC-BSA within the MSNs was confirmed by confocal microscopy. This technique allowed for the visualization of FITC-BSA localized within discrete spheres when drop-cast from a solution of MSNs (Figure 1C). The choice of FITC-BSA as a drug model enabled facile tracking and quantification of the drug release dynamics.

Figure 1.

Figure 1.

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(A) scanning electron microscopy image of loaded MSN surface morphology that was isolated by centrifugation (scale = 100 nm); (B) Size distribution of MSN particles determined by diffraction light scattering (DLS); (C) Morphological and chemical characterization of MSN: (i) SEM image of MSNs, (ii–iv) Scanning transmission electron microscopy (STEM) images under dark field (DF) mode and energy-dispersive X-ray spectroscopy (EDS) mapping images of the elements Si and O; (D) Loaded spheres studied by laser confocal microscopy to visualize FITC-BSA localized in discrete spheres when drop-cast from a solution of MSN; (E) FITC-BSA and rhOSF-loaded MSN incorporation into a GelMA hydrogel mesh with a reinforced matrix examined using confocal microscopy, where FITC signal is observed throughout the matrix in 3D Z-stack renderings; (F) Remaining FITC signal is observed most intensely near the printed thermoplastic mesh compared to the more hydrated GelMA after 16 days of release; (G, H) The release of rhOSF from both MSN and the 3D printed GelMA mesh incorporating MSN, measured by ELISA (R&D Systems, Human Periostin kit).

FITC-BSA and rhOSF-loaded MSNs were loaded into a GelMA hydrogel. The successful incorporation of FITC-BSA-loaded MSNs into the GelMA hydrogel was confirmed through confocal microscopy, and the observed FITC signal distribution within the 3D Z-stack renderings is illustrated in Figure 1D–F. This observation provides a 3D distribution of the nanoparticles within the GelMA hydrogel. Notably, a significant decrease in the abundance of FITC-BSA-loaded spheres was observed over a 16-day time frame. The remaining FITC signal was most intensely concentrated near the printed thermoplastic polycaprolactone (PCL) mesh, with a discernible contrast in signal intensity relative to that of the more hydrated GelMA regions (Figure 1D–F). This observed spatial variation in FITC signal intensity suggests that the release dynamics of FITC-BSA from the MSN-loaded hydrogel matrix are influenced by the substrate’s hydrophilic or hydrophobic properties.17 This influence could potentially be attributed to the hydrophilic properties of GelMA. The enhanced signal intensity near the printed PCL mesh may also be attributed to altered diffusion kinetics and interactions between the nanoparticles and the surrounding matrix, resulting in a more concentrated release in that region. This spatial variation in FITC signal intensity highlights the drug distribution over the scaffold and the potential for tailoring drug release profiles.

To assess the release kinetics over an extended period, we evaluated both MSNs alone and a 3D-printed GelMA matrix incorporating MSNs over 23 and 16 days, respectively, as shown in Figure 1G and 1H. This comparison between the release profiles from MSNs (Figure 1G) and the 3D-printed GelMA mesh incorporating MSNs (Figure 1H) facilitates an understanding of the impact of the GelMA matrix on the release kinetics of rhOSF. ELISA results provided insights into the temporal release patterns of rhOSF from both delivery systems. The release profile from MSNs showed an initial burst release followed by a more sustained and gradual release over time. This initial burst release is likely due to the rapid diffusion of rhOSF from the outer layers of the nanoparticles, a behavior consistent with the inherent properties of MSNs, which can offer controlled release due to their mesoporous structure.

Similar to our results, the literature indicates that MSNs experience a three-phase degradation following administration, including initial burst release, subsequent steady release, and increasing gradually.18 In contrast to MSNs, the release profile from the 3D-printed GelMA matrix-incorporating MSNs demonstrated a potentially modified release pattern compared to that of MSNs alone. This modulation of release kinetics is likely a result of the interplay between the GelMA matrix and the embedded MSN. The GelMA matrix could influence the diffusion dynamics and retention of rhOSF, introducing complexity and an additional layer of control to the release process.19

Collagen is the major natural ECM component in skin, including collagen types I, III, IV, VII, and XIV.20 After being secreted into extracellular space, collagen molecules assemble into higher-order fibrils, and then, the fibrils are assembled into collagen fibers with diameters ranging from 50 to 500 nm. The fiber structure of collagen is critical for cell attachment, proliferation, and viability in vitro and in vivo and is, therefore, a candidate of interest for tissue reconstruction. However, to address concerns over potential pathogen transmission and immune rejection associated with natural collagen (animal or cadaver sources), an additive manufacturing technique, i.e., MEW, has been implemented to create highly porous MF scaffolds of medical grade material and has been integrated with hydrogels and cells.12 Melt electrowritten scaffolds have been used to increase handleability (Figure 2D), enhance mechanical properties,21 and guide cell alignment.22 Figure 2A presents a schematic representation of the developed skin construct, highlighting the incorporation of the MSN-loaded GelMA hydrogel. This construct synergizes the benefits of MF-PCL, MSNs, and GelMA for enhanced wound healing and tissue regeneration. Previously, our research showcased the capacity of biodegradable and synthetic MF-PCL porous scaffolds to facilitate the attachment and proliferation of human bone marrow-derived stem cells (hMSCs).23 Additionally, we demonstrated the efficacy of MF-PCL scaffolds, characterized by precisely controlled porosity and pore size, in enhancing the mechanical resilience of GelMA hydrogels.11Notably, these scaffolds boast up to 98% porosity levels and exhibit fiber diameters akin to those found in native ECM collagen fibers.24

Figure 2.

Figure 2.

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(A) Schematic image of developed skin graft using MSN-loaded GelMA C hydrogel; (B, C) Optical microscope images of MEW PCL mesh and mesh infused with printed GelMA C; (D) Punched construct captured with a tweezer, demonstrating excellent handleability; (E–G) Morphological characterization (SEM) of GelMA C loaded with mesoporous silica nanoparticle; red arrow (nanoparticle), yellow arrow (nanofibrillated cellulose); (H, I) SEM images showing the hydrogel phase uniformly infiltrated into the pores of the polymer mesh.

Of note, the utilization of MEW for fabricating MF-PCL mesh demonstrates a robust foundation for bolstering cell-laden hydrogels amidst mechanical strains encountered during wound healing processes stability.25 Figures 2B–C showcases optical microscope images of the MEW PCL mesh before and after GelMA infusion. The macroscopic evaluation indicates successful integration, with the GelMA visibly enhancing the structural integrity of the polymer mesh. To obtain the morphological insights, we used SEM (Figure 2E–G), presenting images of GelMA loaded with MSNs. Notably, red arrows denote the presence of nanoparticles within the GelMA matrix, while yellow arrows indicate the incorporation of nanofibrillated cellulose. This characterization underscores the effective loading and dispersion of MSNs within the GelMA hydrogel, forming a composite with promising structural attributes. Figure 2H,I further elaborates on the morphological features, illustrating the uniform infiltration of the hydrogel phase into the pores of the MF-PCL mesh. The close integration of GelMA into the MF-PCL mesh suggests an effective interaction between the hydrogel and the polymer, potentially enhancing the mechanical properties and overall construct stability.25

To investigate the effects of rhOSF and GelMA hydrogel on the viability and proliferation of bioprinted HaCaT cells, MTS and live–dead assays were performed. Figure 3A displays the results of the Calcein AM (live cells exhibit green fluorescence) and PI (dead cells are marked by red fluorescence) staining assay for live and dead cell analysis of bioprinted HaCaT cells (1 × 106 cells mL−1) after 1 and 3 days, treated with rhOSF and non-rhOSF culture media. The qualitative assessment revealed high cell viability with minimal cell death in both groups, implying that the shear stress that occurred during the bioprinting of the encapsulated cells did not cause cell death. The viability of HaCaT cells in bioprinted constructs on days 1 and 3 in the presence of rhOSF (50 ng/mL) and culture media are presented in Figure 3B. Significantly enhanced cell proliferation was observed in the presence of rhOSF compared to culture media alone at both time points. This observation suggests the potential role of rhOSF in promoting HaCaT cell survival and proliferation within the bioprinted constructs.

Figure 3.

Figure 3.

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(A) Calcein AM (green) and PI (red) staining assay for live and dead analysis of HaCaT cells printed at treated with rhOSF (50 ng/mL) and culture media after 1 and 3 days; (B) Graph showing HaCaT cell proliferation of bioprinted constructs at days 1 and 3 in rhOSF (50 ng/mL) and culture media (mean ± SD; N ≥ 2, n ≥ 6; ***p < 0.001, ****p < 0.0001; two-way ANOVA, post hoc Tukey test).

To better understand the cellular performance of encapsulated HaCaT cells within rhOSF-loaded hydrogel and bioprinted on top of MEW MF-PCL meshes, four different conditions were studied, including (1) cell-laden constructs treated with non-rhOSF media and (2) rhOSF media-treated constructs (3) laden with blank nanoparticles and (4) constructs incorporating rhOSF-loaded MSNs. Figure 4A presents the live–dead results of encapsulated HaCaT cells after 1 and 5 days in cell-laden GelMA and GelMA/MSN constructs. The red arrows in the images highlight dead cells, providing visual insights into the cell viability within the constructs over 5 days. Incorporating MSN alone into the GelMA matrix improved the cell behavior and survival. Cell proliferation was presented in Figure 4B, assessed through the MTS assay, and exhibited a significant difference in proliferation over time and a notable increase in the constructs containing rhOSF-loaded MSNs. The incorporation of MSN positively influences cell proliferation, suggesting a potential role in supporting cellular activities by providing topological cues within the GelMA matrix. The findings agree with previous studies.26 Moreover, the enhanced cell viability and proliferation observed in constructs incorporating rhOSF-loaded MSNs imply that rhOSF in the scaffold provides a conducive microenvironment, emphasizing promising implications for advanced tissue engineering applications.

Figure 4.

Figure 4.

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(A) Calcein AM (green) and PI (red) staining assay for live and dead analysis of HaCaTs after 1 and 5 days in the cell-laden GelMA C and GelMA C/MSN (containing 50 ng/mL of rhOSF; red arrows: dead cells); (B) MTS assay showing the proliferation of HaCaT cells of the printed constructs at days 1, 3, and 5 (mean ± SD; N ≥ 2, n ≥ 6; ***p < 0.001, ****p < 0.0001; two-way ANOVA, post hoc Tukey test).

In conclusion, this research leverages advanced biofabrication technologies to tackle existing challenges in wound dressing, such as poor mechanical strength, nonviability, and the controlled release of drugs and biologically active agents. Incorporating rhOSF-loaded MSNs into GelMA bioink, bioprinted on MF-PCL meshes, enables sustainable and continuous release of Periostin, further mimicking the natural wound healing environment, which is promising for skin tissue engineering. While further research and preclinical/clinical studies are warranted to validate the efficacy and safety of this novel approach, this work is foundational to advances in the development of biomimetic skin constructs.

Regarding the limitations, it is important to note that PCL has a relatively long biodegradation timeline of ca. 2 years for complete degradation, which exceeds the typical duration of wound healing processes. This extended degradation period poses potential challenges in clinical applications, where rapid biodegradation is preferable. Further exploration of this aspect is crucial for optimizing the scaffold’s application in wound healing, thus, alternative polymers (e.g., amino acid-based poly(ester urea).27–29

MATERIALS AND METHODS

Synthesis and Morphological Characterization of MSNs.

Mesoporous nanoparticles (MSNs) were prepared in two batches, including the drug and blank groups. In a 20 mL vial, 0.3 mL of pH 4 buffer (phosphate-citrate), 8.5 mL of cyclohexane, 2.0 mL of Triton X-100, and 1.0 mL of hexanol were added and stirred at 800 rpm until clear (~30 min). Then, 200 μL of TEOS was added to the vial and stirred for 1 h at 800 rpm. 50 μL of 100 μg/mL drug and ddH2O in the case of “blank” and 8 μL of 25% NH4OH were added to the system and stirred for 24 h at 800 rpm. At 24 h, acetone was added to the system to break the emulsion. The solution was centrifuged at 13000 rpm for 30 min to collect the nanoparticles. After every centrifugation, nanoparticles were rinsed with 50/50 EtOH: ddH2O five times. Finally, the nanoparticles were lyophilized for collection. All materials were purchased from Sigma-Aldrich, St. Louis, MO, U.S.A.

Electron microscopy was used to study the morphology of the synthesized MSNs. To determine the chemical composition, HAADF images, and EDS elemental mapping analyses were performed using a scanning transmission electron microscope (STEM; JEOL 3100 R05, Akishima, Tokyo, Japan). The rhOSF release profiles from MSNs and GelMA hydrogels were monitored and quantified using an Enzyme-Linked Immunosorbent Assay (ELISA) with a Human Periostin kit from R&D Systems. Both the MSNs and GelMA hydrogels, each containing 25 μg of rhOSF-loaded MSNs, were soaked in 1 mL of DI water and incubated at 37 °C. For the GelMA hydrogels, the rhOSF-loaded MSNs were first encapsulated in 1 mL of hydrogel and printed into a PCL mesh before being soaked and incubated. At predetermined intervals, the solutions were centrifuged at 4500 rpm for 5 min to separate the released rhOSF, which was then quantitatively determined as a function of the release time (t) using an ELISA kit.

Fabrication of MEW PCL Mesh with GelMA.

The mesh with or without cells/nanoparticles was fabricated through the convergence of melt electrowriting MEW and droplet printing by using 3DDiscovery (RegenHU, Villaz St. Pierre, Switzerland). Briefly, PCL meshes of 0/90°crosshatch design with 500 μm of space were printed via MEW with parameters as previously described.11 Microvalve-based droplet printing generates arrays of GelMA bioink loaded with cells and MSNs.

Morphological and Biochemical Characterization.

The morphological assessment of the skin construct was performed by using scanning electron microscopy (SEM; MIRA3, FEG-SEM, and TESCAN). To assess the distribution profile of nanoparticles in the printed hydrogel and mesh, the fluorescein isothiocyanate FITC (Sigma-Aldrich, St. Louis, MO, U.S.A.)-BSA nanoparticles were fabricated and observed under a confocal microscope (Nikon Eclipse C2).

Cellular Assays.

The human immortalized keratinocyte cell line, HaCaT (ATCC, PCS-200-011, Manassas, VA, U.S.A.), was printed at a concentration of 1 × 106 cells/mL in GelMA C bioink (Purchased from Cellink containing nanocellulose) alone and on top of MEW MF-PCL meshes, cultured in DMEM medium (Thermo Fisher Scientific_Gibco) supplemented with 10% FBS and 1% P/S, for proliferation and viability using MTS assay and live and dead staining for up to 3 days and 5 days, respectively, without or with 50 ng/mL of rhOSF. Live cells were stained with Calcein-AM, and dead cells were stained with propidium iodide and imaged using CLM (Carl Zeiss). For proliferation at each time point, 50 μL of MTS reagent was added, and the constructs were incubated at 37 °C. The absorbance at 490 nm was recorded after 2 h using a microplate reader (Multiskan, Thermo Fisher Scientific, U.S.A.).

ACKNOWLEDGMENTS

The content is solely the authors’ responsibility and does not necessarily represent the official views of the National Institutes of Health.

Funding

National Institutes of Health (NIH), Grant Nos. R01GM143938 and R01DE031476.

ABBREVIATIONS

MF-PCL

poly(ε-caprolactone) microfibrous

PCL

poly(ε-caprolactone)

MSNs

mesoporous nanoparticles

SEM

scanning electron microscopy

ECM

extracellular matrix

DLS

diffraction light scattering

STEM

scanning transmission electron microscopy

DF

dark field

EDS

energy-dispersive X-ray spectroscopy

MEW

melt electrowriting

ELISA

enzyme-linked immunosorbent assay

Footnotes

Complete contact information is available at: https://pubs.acs.org/10.1021/acsmacrolett.4c00240

The authors declare no competing financial interest.

Contributor Information

Nileshkumar Dubey, Faculty of Dentistry, National University of Singapore, 119077, Singapore.

Maedeh Rahimnejad, Department of Cariology, Restorative Sciences, and Endodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States.

W. Benton Swanson, Department of Biologic and Materials Science, Division of Prosthodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States.

Jinping Xu, Department of Cariology, Restorative Sciences, and Endodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States.

Mylène de Ruijter, Regenerative Medicine Center Utrecht, 3584 Utrecht, The Netherlands; Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, 3584 Utrecht, The Netherlands; Department of Orthopedics, University Medical Center Utrecht, 3584 Utrecht, The Netherlands.

Jos Malda, Regenerative Medicine Center Utrecht, 3584 Utrecht, The Netherlands; Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, 3584 Utrecht, The Netherlands; Department of Orthopedics, University Medical Center Utrecht, 3584 Utrecht, The Netherlands.

Cristiane H. Squarize, Department of Periodontics and Oral Medicine, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States

Rogerio M. Castilho, Department of Periodontics and Oral Medicine, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States

Marco C. Bottino, Department of Cariology, Restorative Sciences, and Endodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan 48109, United States; Department of Biomedical Engineering, College of Engineering, University of Michigan, Ann Arbor, Michigan 48109, United States

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