Injectable Short Nanofiber Fragments Enable Conformal Fibrous Scaffolds for Tissue Engineering on Complex Surfaces

ABSTRACT

Polymeric short nanofibers are widely utilized in drug delivery due to their biocompatibility and sustained release properties; however, their application as scaffold‐forming biomaterials for tissue engineering remains limited. Here, short nanofiber fragments (SNFs) derived from electrospun poly(D‐lactide)/gelatin (PG) nanofiber mats are developed and evaluated for conformal fibrous network formation on complex substrates. SNFs are generated via probe sonication and deposited onto impermeable (carbon tape‐mounted aluminum foil) and porous (Ti–6Al–4 V alloy) substrates through drop casting. Scanning electron microscopy reveals that SNFs uniformly coat both substrate types, forming extended, interconnected fibrous networks with effective infiltration into porous structures, unlike direct electrospinning. Surface wettability is significantly enhanced following fragmentation of the nanofiber mat into SNFs, as evidenced by a reduction in water contact angle of 11.5°. In vitro studies using normal human dermal fibroblasts (nHDF) and preosteoblasts (MC3T3‐E1) demonstrate that PG3 SNF‐coated substrates exhibit excellent cytocompatibility and support time‐dependent cell proliferation, comparable to PG3 nanofiber mats. No statistically significant differences in proliferation are observed for either nHDF or MC3T3‐E1 at any of the investigated time points. These findings demonstrate that SNFs enable conformal scaffold formation on complex surfaces, offering a promising strategy for advanced tissue engineering applications.

Short nanofiber fragments (SNFs) derived from electrospun PDLA/gelatin (PG) mats dynamically reassemble into conformal fibrous networks on complex substrates. This approach overcomes the geometric limitations of conventional electrospinning, enabling uniform surface coverage, enhanced wettability, and cytocompatible scaffold formation, thereby offering a versatile and translational strategy for advanced tissue engineering applications.

1. Introduction

Electrospun polymeric nanofibers have been extensively explored for biomedical applications owing to their structural similarity to the native extracellular matrix (ECM), high surface area‐to‐volume ratio, and adjustable physicochemical properties [1, 2, 3, 4]. The widespread adoption of electrospinning stems from its simplicity, versatility, and cost‐effectiveness in producing polymeric nanofibers, making it highly attractive for large‐scale manufacturing, as evidenced by several commercial nanofiber‐based products [5, 6, 7]. Based on the number of fluid phases involved, electrospinning techniques are broadly classified into single‐fluid, dual‐fluid, and multi‐fluid modalities [8, 9]. Compared with other nanofiber fabrication techniques, including template‐assisted methods, phase separation, nanofiber printing, and self‐assembly, electrospinning remains a highly versatile and scalable approach for fabricating nanofibers with controlled architectures and diverse functionalities [10, 11].

Polymeric short nanofibers (SNs) have recently attracted considerable interest in drug delivery systems due to their superior biocompatibility relative to conventional inorganic nanocarriers, such as metal‐based nanoparticles [12, 13]. Their high aspect ratio and rod‐like morphology, analogous to carbon nanotubes, facilitate efficient drug loading and promote favorable cellular interactions, thereby enhancing therapeutic performance. Short nanofibers (SNs) are typically derived from electrospun nanofiber mats through various fragmentation strategies, including electrospraying, mechanical homogenization, ultrasonic treatment, and cryo‐sectioning [14, 15]. For instance, Yabuki et al. reported the fabrication of four distinct nanofiber morphologies—short, short‐beaded, short aggregated‐beaded, and continuous‐beaded—by incorporating 1.2 µm silica microspheres into cellulose acetate solutions of varying concentrations during electrospinning [16]. Li et al. fabricated graphene/Fe₃O₄/PLGA SNs via an in situ deposition approach after homogenization, yielding fibers with an average length of 11.90 ± 2.03 µm and a diameter of 256.5 ± 13.7 nm [17]. In some cases, these approaches are combined to achieve precise control over fiber dimensions and morphology. For example, Wei et al. reported the preparation of SNs by freezing electrospun poly(lactic‐co‐glycolic acid) (PLGA) fiber mats in water, followed by sectioning into 50 µm slices and subsequent ultrasonic dispersion [18]. Drug incorporation to SNs can be achieved either by blending therapeutic agents with polymer solutions prior to electrospinning or through post‐fragmentation strategies [19].

Despite these advantages, the application of SNs in tissue engineering remains limited. Their nano‐ to micro‐scale dimensions facilitate cellular internalization, which may lead to undesirable cytotoxicity at higher concentrations [20, 21, 22, 23]. In contrast, continuous electrospun nanofiber mats have been widely employed as scaffolds for tissue regeneration due to their ability to support cell adhesion, proliferation, and differentiation. However, these conventional non‐woven mats are inherently restricted to forming planar, two‐dimensional architectures and exhibit limited accessibility to conform to porous substrates. This limitation becomes particularly significant in the context of complex biomedical devices and implants, where uniform surface coverage and structural integration are critical. These challenges underscore the need for alternative strategies capable of generating conformal, stable, and biocompatible fibrous networks across diverse substrate geometries.

In this regard, we hypothesize that short nanofiber fragments (SNFs), which retain partial structural continuity unlike fully individualized SNs, can reassemble into interconnected fibrous networks upon simple deposition. Such SNFs are expected to bridge the gap between dispersed nanofiber segments and continuous electrospun mats, thereby enabling the formation of scaffold‐like architectures on both impermeable and porous surfaces. The ability of SNFs to reconstruct fibrous networks is likely governed by parameters such as polymer composition, fragmentation conditions, and deposition dynamics [14].

Poly(D‐lactide) (PDLA) is a well‐established biodegradable polymer distinguished by its mechanical stability and relatively slow degradation under physiological conditions, making it suitable for long‐term biomedical applications. It is a stereoregular, biodegradable aliphatic polyester synthesized via the ring‐opening polymerization of D‐lactide, the cyclic dimer of D‐lactic acid. A defining feature of PDLA is that all chiral centers along its polymer backbone possess the R‐configuration, rendering it an isotactic polymer and the enantiomeric counterpart of poly(L‐lactide) (PLLA) [24, 25]. Gelatin, derived from collagen, provides bioactive motifs that promote cell adhesion and proliferation, particularly in soft tissue engineering [26, 27]. By combining PDLA with gelatin, composite nanofibers can integrate structural integrity with biological functionality. In the present study, PDLA/gelatin electrospun nanofiber mats are fabricated and subsequently converted into short nanofiber fragments (SNFs) via probe sonication. The ability of SNFs to form extended fibrous networks upon drop casting is evaluated on two model substrates: an impermeable planar surface (carbon tape on aluminum foil) and a three‐dimensional porous biomaterial (Ti–6Al–4 V alloy). The cytocompatibility and proliferative responses of normal human dermal fibroblasts (nHDF) and preosteoblasts (MC3T3‐E1) are assessed on these SNF‐fabricated substrates. This study establishes a versatile SNF‐based platform for generating conformal fibrous scaffolds on complex surfaces, thereby advancing the translational potential of electrospun nanofibers in tissue engineering and regenerative medicine.

2. Results and Discussion

2.1. Evaluation of Scaffold's Physicochemical Properties

Electrospun nanofiber mats fabricated from varying PDLA/gelatin compositions exhibited thicknesses in the range of 0.06–0.08 mm, as measured using a digital caliper. SEM was employed to evaluate surface morphology and fiber diameter distribution (Figure S1). All samples displayed smooth, cylindrical fibers with partial alignment along a preferred direction. The pristine PDLA sample (P) exhibited fiber diameters ranging from 208 to 411 nm. In contrast, PDLA/gelatin composite fibers showed broader and more heterogeneous diameter distributions, with ranges of 174–993 nm (PG1), 139–1155 nm (PG2), and 86–1276 nm (PG3), indicating the influence of gelatin incorporation on fiber formation and jet instability during electrospinning.

ATR‐FTIR analysis confirmed the presence of characteristic functional groups and intermolecular interactions within the composite system (Figure 1a). PDLA exhibited distinct peaks at 1759 cm⁻¹ (C = O stretching) and 1184 and 1085 cm⁻¹ (C–O–C stretching). Gelatin showed characteristic amide I and amide II bands at 1640 and 1521 cm⁻¹, respectively. These observations are consistent with previous reports on electrospun PDLA/gelatin‐based systems [28, 29]. In the composite (PG3), these peaks were retained but shifted to 1666 and 1541 cm⁻¹, indicating hydrogen bonding interactions between the carbonyl groups of PDLA and amide groups of gelatin macromolecules. Surface wettability was assessed via water contact angle measurements (Figure 1b). PDLA exhibited a hydrophobic surface (134.1 ± 0.9°), whereas gelatin showed comparatively higher wettability (92.8 ± 1.6°). The composite PG3 displayed an intermediate contact angle of 118.2 ± 1.4°, reflecting the combined physicochemical characteristics of both polymers.

FIGURE 1.

Physicochemical characterization of electrospun nanofiber mats. (a) ATR‐FTIR spectra of PDLA, gelatin, and PDLA/gelatin (PG3) composites, showing characteristic functional groups and peak shifts indicative of intermolecular interactions. Dashed lines highlight the shifts in characteristic absorption bands. (b) Static water contact angle measurements of the corresponding samples, demonstrating differences in surface wettability.

During the preparation of short nanofiber fragments (SNFs) via probe sonication, only the PG3 composition yielded well‐dispersed SNFs, whereas P, PG1, and PG2 predominantly formed aggregated clumps with limited fragmentation under the same conditions. Therefore, PG3 was selected as the optimized formulation for subsequent studies. Upon standing the PG3 dispersion for 30 min, phase separation was observed, with larger, denser fragments sedimenting and smaller fragments remaining suspended. Representative images of nanofiber mats, fragmented SNFs, and drop‐casted substrates are shown in Figure 2.

FIGURE 2.

Digital images illustrating SNF preparation and deposition. (a) Electrospun nanofiber mat, cut fiber segments, fragmented nanofibers following probe sonication, and phase‐separated SNF fractions (sedimented and suspended) after standing. (b) Drop‐casting of PG3 SNFs onto an impermeable substrate (carbon tape‐mounted aluminum foil). (c) Deposition of PG3 SNFs onto a porous Ti–6Al–4 V substrate.

SEM analysis further revealed distinct fragmentation behavior between P and PG3 samples (Figure 3). The PDLA sample retained its original fibrous alignment with minimal fragmentation, whereas PG3 exhibited effective disintegration into individual or loosely entangled nanofiber fragments. Notably, SNFs collected from the suspended fraction demonstrated more complete fragmentation compared to those from the sedimented fraction. Upon drop‐casting, the suspended PG3 SNFs formed continuous, interconnected fibrous networks resembling electrospun mats. A significant change in surface morphology was observed following sonication. The initially smooth PG3 nanofibers transformed into rough, irregular structures (Figure S2), leading to enhanced surface wettability. The water contact angle decreased from 118.2 ± 1.4° to 106.7 ± 1.3°, representing a reduction of approximately 11.5° (Figure S3). This increase in surface roughness is likely to enhance protein adsorption and subsequent cell–material interactions [30].

FIGURE 3.

SEM images of fragmented nanofibers from (a) pristine PDLA (P) and (b) PDLA/gelatin (PG3) samples following probe sonication. SNFs derived from PG3 are shown for both the sedimented and suspended fractions, highlighting differences in fragmentation behavior and fiber morphology.

Ti–6Al–4 V alloy is a widely utilized orthopedic implant material owing to its excellent mechanical strength, corrosion resistance, and biocompatibility. Porous Ti–6Al–4 V architectures have emerged as promising alternatives for bone implants, as they mitigate stress‐shielding effects compared to dense counterparts [31, 32]. When deposited onto porous Ti–6Al–4 V substrates, the suspended PG3 SNFs formed extended fibrous networks that not only covered the outer surfaces but also penetrated into the internal pores (Figure 2c). The SNFs exhibited strong adhesion to the metallic substrate, as evidenced by SEM analysis (Figure 4). In contrast, directly electrospun nanofibers showed poor adhesion and were easily detached upon mechanical handling (Figure S4), highlighting the advantage of SNF‐based deposition for conformal coating of complex geometries.

FIGURE 4.

SEM images of (a) non‐coated (control) and (b) PG3 SNF‐coated three‐dimensional (3D) porous Ti–6Al–4 V substrates. The SNF coating forms an interconnected fibrous network across the surface and within the pore architecture. Regions within the inner pore at different depths are indicated by a star (*) and a solid dot (•).

2.2. Cell Proliferation on SNF‐Fabricated Structures

The biological performance of PG3 SNF‐coated substrates was evaluated using normal human dermal fibroblasts (nHDF) (Figure 5a) and MC3T3‐E1 preosteoblasts (Figure 5b) via a CCK‐8 assay, and the results were compared with cell growth on PG3 nanofiber mats.

FIGURE 5.

Cellular response on PG3 nanofiber mat and PG3 SNF‐coated impermeable substrate. CCK‐8 assay showing time‐dependent proliferation of (a) normal human dermal fibroblasts (nHDF) and (b) MC3T3‐E1 preosteoblasts cultured on both samples. Corresponding SEM images illustrating cell attachment and spreading are shown for (c) nHDF and (d) MC3T3‐E1.

A time‐dependent increase in cell growth was demonstrated on both SNF‐coated impermeable substrate and nanofiber mats. However, the difference in cellular growth between them was different depending on the cell types. The growth of fibroblasts and preosteoblasts on both samples did not show any remarkable difference on the investigation period. The corresponding SEM images (Figure 5c,d) showing the cellular growth reflecting the CCK assay findings. The images confirmed that both cell types adhered to the SNF networks within 24 h and progressively spread with well‐defined filopodia over time, indicating favorable cell–material interactions. Further, the cellular morphology was not affected on both samples.

As PG3 nanofiber mats cannot be directly fabricated on porous Ti–6Al–4 V substrates, and given that this metallic substrate is widely used for bone tissue regeneration [33], cell proliferation studies were conducted using MC3T3‐E1 cells exclusively on PG3 SNF‐fabricated substrates. The CCK‐8 assay results demonstrated that PG3 SNFs exhibited a similar time‐dependent proliferation trend on the porous implant, consistent with observations on impermeable substrates (Figure 6a).

FIGURE 6.

Cellular response on PG3 SNF‐coated porous substrates. (a) CCK‐8 assay showing time‐dependent proliferation of MC3T3‐E1 preosteoblasts cultured on SNF‐coated porous structures. (b) SEM images and (c) CLSM images illustrating cell attachment and growth across different regions of the SNF‐fabricated scaffold; cells are indicated by arrows. In CLSM images, F‐actin and nuclei were stained with Alexa Fluor 594 phalloidin and DAPI, respectively.

SEM and confocal microscopy images further confirmed uniform cell attachment and spreading across complex topographical features, including both crests and pores, facilitating effective cell colonization throughout the inner and outer regions of the scaffold (Figure 6b,c).

Collectively, these results demonstrate that SNF‐fabricated networks provide an amicable microenvironment for cell attachment and proliferation. The reconstructed fibrous architecture mimics the non‐woven structure of electrospun mats while overcoming their limitations, such as poor penetration into porous substrates and weak surface adhesion. Importantly, the absence of freely dispersed nanoscale particles minimizes the likelihood of cellular internalization‐associated cytotoxicity.

Previous studies have primarily utilized short nanofiber fragments as reinforcing components within composite biomaterial systems rather than as standalone scaffold‐forming platforms. For instance, the research group led by Ko et al. incorporated poly(lactic‐co‐glycolic acid) (PLGA) nanofiber fragments into gelatin methacrylate hydrogels for 3D bioprinting applications, demonstrating enhanced cell proliferation compared to control hydrogels [34]. Similarly, Chen and co‐workers developed gelatin/PLGA‐derived nanofiber fragments to reinforce cartilage‐derived decellularized matrix‐based bioinks, resulting in improved scaffold performance for cartilage tissue regeneration [35]. In another study, Boda et al. fabricated nanofiber fragments from electrospun poly(D,L‐lactide‐co‐glycolide)/collagen/gelatin composites and further functionalized them through mineralization and conjugation with calcium‐binding osteoinductive peptides [36]. These nanofiber fragments were employed as fillers in alveolar bone defects, demonstrating their potential for bone tissue regeneration. Collectively, these studies highlight the utility of nanofiber fragments as functional additives to enhance the performance of biomaterial systems; however, their direct application as self‐supporting, scaffold‐forming networks on complex substrates remains largely unexplored.

The present study addresses this gap by demonstrating that SNFs can be engineered to form conformal, interconnected fibrous networks while retaining key structural and biological attributes of electrospun mats. Although comparative cellular studies with non‐coated controls were not extensively performed, future work will focus on incorporating bioactive agents (e.g., antimicrobial, osteoconductive, or electroconductive components) and evaluating in vivo performance in relevant tissue regeneration models.

3. Conclusions

In this study, PDLA/gelatin nanofiber mats were fabricated and converted into short nanofiber fragments (SNFs), enabling the identification of an optimized formulation (PG3) capable of forming conformal nanofibrous networks on both impermeable and porous substrates. Unlike conventional electrospun fibers, PG3 SNFs effectively infiltrated porous architectures and reassembled into interconnected fibrous networks, providing uniform coverage across complex surfaces. Enhanced wettability, attributed to increased surface roughness, further supported favorable cell interactions. This work introduces SNFs as reconstructive building blocks that bridge the gap between dispersed nanofibers and continuous mats, validating the hypothesis that they can form scaffold‐like networks on complex geometries. In contrast to previous studies where nanofiber fragments serve mainly as fillers or reinforcement phases within hydrogels or composite matrices, SNFs here function as standalone scaffold‐forming materials. In vitro studies confirmed cytocompatibility, with cell proliferation comparable to electrospun mats. This platform shows potential for coating porous orthopedic implants, injectable wound dressings, and minimally invasive tissue repair. Future work will focus on the incorporation of bioactive agents (e.g., osteoinductive, antimicrobial, or angiogenic molecules) into SNFs and their validation in in vivo models to assess tissue integration, biodegradation kinetics, and long‐term regenerative efficacy. Such advancements are expected to further establish SNFs as a versatile and translational platform for next‐generation tissue engineering and regenerative medicine.

4. Experimental Section

4.1. Chemicals and Materials

Poly(D‐lactide) (PDLA; Mw = 85 000–160 000 Da, BMG Incorp., Kyoto Japan), gelatin from porcine skin (Type A, gel strength ≈300 Bloom), 1,1,1,3,3,3‐hexafluoro‐2‐propanol (HFIP, ≥99%), and trifluoroacetic acid (TFA, 99%) were purchased from Sigma–Aldrich and used as received without further purification. Deionized (DI) water was used throughout the study. Ti–6Al–4 V Grade 23 powder with a particle size range of 15–45 µm was obtained from Tekna. Porous Ti–6Al–4 V structures were fabricated via powder bed fusion (PBF) using selective laser melting (SLM).

4.2. Preparation of PDLA/Gelatin Electrospun Nanofibrous Mats

Electrospun nanofibrous mats were prepared from PDLA/gelatin polymeric blends (denoted as P, PG1, PG2, and PG3) dissolved in a solvent mixture of 7 mL HFIP and 1 mL TFA under magnetic stirring at 40°C for 30 min to obtain homogeneous solutions (compositions are provided in Table S1). Electrospinning was carried out at an applied voltage of 15 kV under ambient conditions (temperature: 25°C; relative humidity: ≈30%). The polymer solution was delivered using a syringe pump at a constant flow rate of 0.6 mL h⁻¹ through a stainless‐steel needle (21 gauge). The tip‐to‐collector distance was maintained at 10 cm. A rotating cylindrical collector covered with aluminum foil was employed, operating at a rotational speed of 1000 rpm to ensure uniform fiber deposition. The collected nanofiber mats were dried in a hot air oven at 60°C for 24 h to remove residual solvent.

4.3. Preparation of PDLA/gelatin Short Nanofiber Fragments

Electrospun nanofiber mats were cut into small segments using scissors and transferred into a 10 mL glass vial containing deionized water. Fragmentation was performed using a probe sonicator (Vibra‐Cell ultrasonic disperser) operated at a frequency of 20 kHz and an amplitude of 28% for 90 s, resulting in the formation of short nanofiber fragments (SNFs). The resulting dispersion was freeze‐dried to obtain lyophilized SNF powders. Subsequently, 0.1 g of lyophilized SNFs was re‐dispersed in 10 mL of distilled water. Upon standing for 10–15 min, larger and denser SNFs sedimented, while smaller fragments remained suspended in the supernatant. The supernatant (containing smaller SNFs) and the sedimented fraction were separately collected and deposited onto solid substrates by drop casting (200 µL per deposition, repeated three times) to evaluate differences in fiber assembly following solvent evaporation. The preparation of SNFs from electrospun nanofiber mats and their deposition onto impermeable and porous substrates are schematically illustrated in Scheme 1.

SCHEME 1.

Schematic illustration of the fabrication process of poly(D‐lactide)/gelatin (PG) electrospun nanofiber mats, followed by the generation of short nanofiber fragments (SNFs) and their subsequent deposition onto impermeable and 3D porous substrates via a drop‐casting approach. The schematic was created using BioRender.com and Blender software.

4.4. Instrumental Characterization

The thickness of the electrospun nanofiber mats was measured using a digital caliper (Mitutoyo 500‐197‐30 Absolute Scale). The morphology of the nanofibers was examined by scanning electron microscopy (SEM; SUPRA 25, Zeiss). Prior to imaging, samples were mounted on carbon tape and sputter‐coated with platinum to enhance conductivity. Fiber diameter distribution and three‐dimensional (3D) surface reconstructions were analyzed using ImageJ software. Functional group analysis was performed using attenuated total reflectance Fourier transform infrared spectroscopy (ATR‐FTIR; Spectrum GX, PerkinElmer, USA) over a wavenumber range of 4000–400 cm⁻¹. Surface wettability was evaluated by static water contact angle measurements using a contact angle goniometer (EasyDrop FM40Mk2, Krüss, Germany) equipped with a CCD camera (precision ±0.2°). Measurements were conducted using the sessile drop method, and at least three independent locations per sample were analyzed to obtain average values. Confocal laser scanning microscopy (CLSM) images of cells cultured on the scaffolds were acquired using an LSM 800 system (Zeiss, Germany).

4.5. Cell Maintenance and Culture on Scaffolds

Normal human dermal fibroblasts (nHDF; ATCC, USA) were cultured in Dulbecco's modified Eagle's medium (DMEM; Welgene, Korea) supplemented with 10% (v/v) fetal bovine serum (FBS; Welgene, Korea) and 1% (v/v) penicillin–streptomycin. MC3T3‐E1 preosteoblasts (CRL‐2593, ATCC, USA), derived from C57BL/6 mouse calvaria, were cultured in α‐minimum essential medium (α‐MEM; Gibco, USA) supplemented with 10% (v/v) FBS and 1% (v/v) antibiotic–antimycotic solution (10 000 U mL⁻¹ penicillin, 10 mg mL⁻¹ streptomycin, and 25 µg mL⁻¹ amphotericin B). Both cell lines were maintained in a humidified incubator at 37°C with 5% CO₂, and the culture medium was refreshed every 48 h.

SNF‐coated impermeable and porous substrates prepared via drop‐casting were placed in 48‐well tissue culture plates and sterilized under UV irradiation for 48 h. The scaffolds were equilibrated with 500 µL of Dulbecco's phosphate‐buffered saline (DPBS) for 1 h, after which the solution was removed. Cells were seeded onto each scaffold at a density of 3 × 10⁴ cells per well in 500 µL of culture medium. Culture durations were 1, 3, and 5 days for nHDF cells and 1, 4, and 7 days for MC3T3‐E1 cells.

4.6. Cell Proliferation Assay and Imaging

Cell viability was assessed using a Cell Counting Kit‐8 (CCK‐8; Dojindo, Japan) at predetermined time points. The culture medium was replaced with a mixture of 450 µL DPBS and 50 µL CCK‐8 reagent, followed by incubation at 37°C for 1.5 h. Subsequently, 100 µL of the supernatant was transferred to a 96‐well plate, and absorbance was measured at 450 nm using a microplate reader (SpectraMax 340, Molecular Devices, USA).

Cell morphology, orientation, and distribution on the scaffolds were evaluated by SEM and CLSM. For SEM analysis, cells were fixed with buffered formalin at 25°C for 15 min, washed twice with phosphate‐buffered saline (PBS), and dehydrated through a graded ethanol series (50%, 70%, 80%, 90%, and 100%). Samples were then air‐dried overnight prior to imaging. For CLSM analysis, cells were permeabilized with 0.5% (v/v) Triton X‐100 in PBS for 10 min and blocked with 2.5% (w/v) bovine serum albumin (BSA) in PBS for 10 min. Cells were then incubated with a staining solution containing Alexa Fluor 594 phalloidin and 4′,6‐diamidino‐2‐phenylindole (DAPI) (Thermo Fisher Scientific, USA) for 20 min to visualize F‐actin (λ = 516 nm) and nuclei (λ = 405 nm), respectively. Samples were washed twice with PBS after each step.

4.7. Statistical Analysis

All quantitative data are presented as mean ± standard deviation (SD) (n = 3). Statistical significance is evaluated using one‐way analysis of variance (ANOVA) followed by Tukey's post hoc test, with comparisons made against the control group. Differences are considered statistically significant at p < 0.05.

Funding

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (No. RS‐2026‐25468613) and by the Technology Innovation Program (RS‐2025‐04572968, Development and Commercialization of AI‐Based Rapid Neovascularization‐Inducing Bioink Materials to Improve the Engraftment Rate of Disease‐specific Organoids) funded by the Ministry of Trade, Industry & Energy (MOTIE, Korea).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: marc70337‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.