Mucoadhesive chitosan-coated Fe3O4 magnetic nanoparticles for the treatment of intestinal dysmotility

Intestinal dysmotility represents a significant health burden, often leading to severe and life-threatening complications. Current therapies are limited, highlighting the need for smart treatment strategies. We propose a novel, minimally invasive approach involving the oral delivery of mucoadhesive magneto-responsive nanoparticles, which can be actuated by external magnets. As a proof of concept, Fe₃O₄ nanoparticles were coated with chitosan to optimise their interaction and adhesion to the intestinal epithelium and to enhance cytocompatibility. To protect the chitosan-coated particles from the acidic conditions of the stomach and enable their targeted release further along the gastrointestinal tract, they were encapsulated in pH responsive alginate beads. Using an advanced 3D multi-layered in vitro model of the intestinal wall, we confirmed the absence of nanoparticle translocation across the barrier. Moreover, epithelial para- and transcellular transport pathways were unaltered in the presence of the chitosan-coated particles, suggesting that nutrient passage is conserved. Finally, a hydrogel simulating the intestinal submucosal layer was loaded with the nanoparticles and then actuated with external magnets to verify their capacity to generate physiological strains. These findings demonstrate the feasibility of magnetically actuated mucoadhesive nanoparticles as a foundation for new therapeutic strategies to restore intestinal motility, paving the way for externally controlled, minimally invasive interventions in gastrointestinal disorders.

Introduction

Visceral smooth muscle, composed of two layers of longitudinally and circumferentially aligned fibres, plays a crucial role in intestinal functions. Its contractile capability, driven by a cytoskeletal protein network, is essential for propelling substances through the digestive tract via peristalsis and segmentation. Enteric nervous system disorders, enzymatic deficits, intestinal pacemaker cell alteration, or genetic mutations affecting these cytoskeletal proteins can lead to debilitating conditions, such as Irritable Bowel Syndrome (IBS), Hirschsprung disease, or Visceral Myopathies (VM) (Benarroch, 2007; Hashmi et al., 2021; Oh et al., 2019; Yadak et al., 2019). VM, which compromises smooth muscle function in the bowel resulting in diminished tone and impaired motility, is one of the most aggressive and debilitating intestinal pathologies, often emerging in childhood.

Currently, besides intestinal transplantation, there is no cure. The unavailability of treatment is due to the unknown nature of the disease’s progression and development (Hashmi et al., 2021). To manage symptoms and prevent life-threatening complications, current care relies on supportive and nutritional strategies. Ileostomies may be considered to alleviate obstructions (Dreyer et al., 2021), and intravenous total parenteral nutrition or enteral feeding, when some digestive function remains (Thapar et al., 2018), represent the primary nutritional support options for patients with severely impaired bowel motility.

In parallel, several invasive or device-based approaches have been explored to restore intestinal motility. The sacral nerve stimulator is a device implanted under the skin that emits mild electrical pulses to the sacral nerve (Noblett and Cadish, 2014), although its applicability is limited to the terminal part of the bowel. Alternatively, transcutaneous electrical stimulation devices deliver mild electrical pulses to the abdominal skin to stimulate the nerves that control the bowel (Ng et al., 2016). Mechanical stimulation strategies have also been proposed. Swallowable vibrating capsules, controlled magnetically or via Bluetooth, can vibrate in situ and, in their latest generation, integrate wireless modules for motor control and inertial sensing. However, they remain unable to deform the bowel wall effectively due to their limited contact area. Moreover, they may induce non-physiological deformation, producing direct radial contraction of the lumen while the capsule itself moves in the opposite direction (Math et al., 2023).

A comprehensive recent review by Garbati et al. (2024) summarises current strategies for gut-targeted nanocarriers, highlighting mucoadhesive coatings, pH-responsive delivery platforms and the use of biocompatible polymers for local retention and targeted release. Complementing this overview, several recent studies have explored natural, hybrid or magnetically functionalized nanomaterials for biomedical applications: biogenic silver nanoparticles for antimicrobial and anticancer uses (Doğan et al., 2025), activated-carbon-coated magnetic nanocomposites for drug delivery (Doğan et al., 2024), chitosan-based formulations incorporating natural extracts (Evcil et al., 2025), and activated-carbon-supported magnetic nanocarriers derived from plant biomass (Öziç et al., 2024). Although these studies address different targets, they collectively support the rationale of combining biopolymer coatings (e.g., chitosan), magnetic cores (Fe₃O₄) and pH-sensitive hydrogels (e.g., alginate) to obtain biocompatible, environmentally green nanosystems for localized gastrointestinal applications.

Building on these concepts, we propose a novel, minimally invasive strategy to restore intestinal motility. By using mucoadhesive magnetic nanoparticles, localized, temporary magneto-responsiveness are imparted to the bowel wall, enabling controlled deformation through the application of external magnetic fields. This paper focuses on the coating of magneto-responsive nanoparticles with a mucoadhesive material (chitosan) and their characterisation in terms of stability over time at different pH levels, mucoadhesion, and cell viability.

In the biomedical field, magnetic nanoparticles (NPs) are typically characterized and used for hyperthermic effects on cancer cells or for drug delivery to various tissues. Here we explore their application as wireless motion induction agents, assessing their suitability for this purpose. Using an advanced 3D multi-layered in vitro intestinal model (Ferreira et al., 2025; Macedo et al., 2020; Macedo et al., 2022), we investigated nanoparticle translocation across multiple cell layers and assessed the potential impact of NPs on cellular nutrient absorption. Additionally, we evaluated a protective, pH-responsive hydrogel designed to encapsulate the NPs, ensuring their structural integrity and controlled release under target pH conditions.

Figure 1 illustrates how the formulation will be used in the proposed strategy. Alginate protects the chitosan-coated nanoparticles (CS-NPs) in the stomach’s acid environment, ensuring their targeted release in the bowel. Once there, the CS-NPs are released (phase 1) and, due to their mucoadhesive properties, temporarily bound to the inner mucosal layer (phase 2). Using external magnets, the bowel can then be deformed as needed (phase 3). Finally, after the magnetic stimulus is removed, the CS-NPs are cleared from the body (phase 4). Clearance is expected to occur within approximately 24 h due to a combination of mucus turnover (Lai et al., 2008), intestinal peristalsis, and gradual degradation of the chitosan coating. These processes concur to facilitate their removal (Subramanian et al., 2022; Xu et al., 2019).

Figure 1

Figure 1. Schematic representation of the proposed minimally invasive strategy for restoring intestinal motility. NPs coated with chitosan (CS-NP) are encapsulated in alginate and can be ingested. After passing the stomach, the CS-NPs are released (phase 1) and bind to intestinal mucosa (phase 2). In phase 3 external magnetic fields induce wall motion. In the final phase (4) the CS-NPs are eliminated through the colon.

Materials and methods

Nanoparticle coating and characterisation

Iron oxide magnetic NPs of 200 nm diameter were selected to prevent their passage through the paracellular space and into the intestinal submucosa or vessels, thereby limiting their migration to other body regions. The Fe₃O₄-112 NPs were purchased from Atomicles (United States), through its European distributor Oocap France SAS (38 Rue Saint-Sébastien, 13,006 Marseille, France; LOT X03 240.129). Chitosan (CS, medium molecular weight, 190–300 kDa, Sigma-Aldrich Co.) was chosen as a biocompatible material to coat the NPs and confer mucoadhesion (Garbati et al., 2024). The protocol described by Khedri et al. (2018) was adapted to coat the NPs by adsorption. Briefly, a 0.2 mg/mL CS solution was prepared in 1 M acetic acid and stirred overnight. A 0.1 mg/mL NPs suspension was prepared in deionized water and sonicated for 5 min at 60% amplitude output intensity (VibraCell model VCX 130W equipped with a 6 mm probe, Sonics & Materials, Inc., Newtown, CT, United States) to disperse the NPs. The NPs were magnetically separated from the water and added to the CS solution, followed by overnight stirring. Chitosan-coated NPs (CS-NPs) were then collected using a magnet.

Samples were diluted with deionized water to a concentration of 0.01 mg/mL, and their diameter and zeta potential were measured using a ZetaSizer Nano ZS (Malvern Instruments Ltd., Worcestershire, United Kingdom) at room temperature. All measurements were performed in triplicate immediately after preparation and after 7 days. Stability at different pH levels, mimicking the gastrointestinal tract, was analyzed using a titrator from the ZetaSizer Nano ZS. This instrument measures the pH of the solution and adds small quantities of 1 M NaOH to increase the pH to user-set values. pH levels were increased in 0.2-unit increments from 2.89 to 6.5. Size and zeta-potential were measured at each pH value.

Powdered samples, magnetically collected after drying the CS-NP and NP solutions at room temperature for 24 h, were immobilised on copper grids and dried again at room temperature for 2 h. In this analysis, the solutions were not sonicated to avoid the disintegration of the coating itself. For CS-NPs, negative staining was performed with 2% uranyl acetate, and the samples were imaged using transmission electron microscopy (TEM – Jeol JEM 1400, 80 kV) to highlight the organic CS-shell (Ong et al., 2017). For uncoated NPs, high atomic number and density provided strong contrast; therefore, imaging was performed in scanning TEM mode with annular dark-field detector (STEM ADF), which enhances contour sharpness, reduces phase contrast artifacts, and gives a clean, easily interpretable image of the inorganic cores (Yu et al., 2005). The TEM analysis was qualitative and conducted to confirm the presence of the chitosan coating around the Fe₃O₄ cores, while quantitative measurements of particle size and distribution were obtained as previously described using the ZetaSizer.

FTIR spectra analysis (Fourier Transform Infra-Red Spectrometer – Frontier, Perkin Elmer) was performed on coated and uncoated NPs to further confirm the presence of chitosan functional groups on the surface of the synthetic core.

Evaluation of mucoadhesion

To evaluate the mucoadhesive properties of CS-NPs compared to (untreated) NPs, we adapted protocols from Vieira et al. (2018) to assess the time-dependent surface absorption of mucins through an increase in particle size and zeta-potential. A 0.2 mg/mL suspension of CS-NPs or NPs in PBS (phosphate-buffered saline) at pH 7 was mixed with an equal volume of 40 mg/mL porcine stomach mucin (Type II, Sigma-Aldrich, M2378) in PBS at pH 7. The mixture was incubated at 37 °C while orbital shaking at 100 RPM (Panasonic, MIR-S100-PE, Kadoma, Japan). At 1, 2, and 4 h, 200 μL samples were collected, centrifuged at 10,000 RPM for 5 min, and the supernatant was collected. The nanoparticle pellet was resuspended in deionized water and analyzed for size and zeta-potential. In addition, the supernatant was analyzed using a UV-VIS detector (Varian, United States) at 255 nm to determine the amount of bound mucin using a turbidimetric method. A 20 mg/mL mucin solution in deionized water was used as a reference to establish the maximum mucin concentration (considered as 100%). The amount of mucin bound to the NPs was estimated using Equation 1, where Abs_s is the absorbance of the supernatant and Abs₀ is the absorbance of the initial (40 mg/mL) mucin solution.

$Mucoadhesion=\frac{1-{Abs}_s}{{Abs}_0}·100(1)$

pH-responsive hydrogel

Sodium alginate with low glucuronic acid content (FG = 0.39, Sigma-Aldrich) was chosen as a sacrificial material to preserve the integrity of the CS-NPs until they reach the bowel. A 0.1 mg/mL suspension of CS-NPs was added to a 20 mg/mL sodium alginate solution in deionized water, adjusted to pH 3 with HCl, and stirred overnight. 10 μL droplets of this solution were added to a 0.1 M CaCl₂ solution to crosslink the alginate. The distance between the pipette tip and the water surface was kept constant to ensure uniform droplet size. The crosslinking process was monitored at 15, 30, and 60 min. The alginate-encapsulated CS-NPs (Alg-CS-NPs) were then transferred to a simulated stomach environment (0.1 mg/mL pepsin, 1.5 mg/mL mucins, 8.78 mg/mL NaCl, adjusted to pH 2 with 6N HCl, Sigma-Aldrich) (Ferrua and Singh, 2015). To determine any changes in size, over typical gastric transit times (Hellmig et al., 2006), they were monitored at 0, 1, 2, and 3 h using a Leica DMI6000 FFW microscope.

CS-NP release

After 60 min of crosslinking and 3 h in the simulated stomach environment, the Alg-CS-NPs were transferred to a simulated small intestine environment (pH 7). At 1, 2, and 3 h (the normal range for small bowel transit time is 2–6 h (Lee et al., 2014)), 100 μL of the solution was collected and analyzed using an Iron Kit (Spectroquant®, HC300920). Absorbance was measured at 564 nm using Thermo Scientific™ Varioskan™ LUX multimode microplate reader. A calibration curve was constructed to correlate absorbance with CS-NP concentration in solution.

3D multi-layered in vitro intestinal model

An advanced 3D multi-layered in vitro intestinal model (Figure 2) was developed using Transwells® following a previously reported protocol (Ferreira et al., 2025; Macedo et al., 2020; Macedo et al., 2022). The model is composed of an endothelial layer seeded on the external basolateral side of the membrane insert. A hydrogel encapsulating fibroblast-like cells is applied to the internal apical side to mimic the submucosal layer. Finally, a coculture of intestinal cells forms the epithelial layer.

Figure 2

Figure 2. Advanced 3D multi-layered in vitro intestinal model (Ferreira et al., 2025; Macedo et al., 2020; Macedo et al., 2022). The model reproduces the structure of the intestinal barrier. Transcellular and paracellular transport pathways are indicated by red and blue arrows, respectively. Image not to scale.

Briefly, standard 12-well Transwell® inserts (cellQART®, 1 µm pore size) were used to recreate the barrier model (Pereira et al., 2016). Each insert was inverted onto 6-well plates and was coated with 100 μL of 0.2% porcine gelatin (Merck) for 1 h at room temperature to promote cellular adhesion. After this period, the excess gelatin was removed, and 100 μL of human pulmonary microvascular endothelial (HPMEC-ST1.6R) cell suspension was added, with a resulting density of 5 × 10⁴ cells/cm². Cells were allowed to adhere for 2 h at room temperature, and 1 mL of PBS was added to each well to maintain humidity and prevent drying. Then, the insert was returned to its normal configuration in 12-well plates and 1 mL of DMEM was added to the basolateral compartment. A hydrogel composed of 6 mg/mL collagen I (Rat Tail, Corning) and 1 mg/mL alginate (Merck), encapsulating 10⁵ cells/mL of human intestinal fibroblasts (HIF) was applied to the internal apical side to mimic the submucosal layer (Chalkidi et al., 2022; Ferreira et al., 2025). After a 30-min gelation process, a mixture of Caco-2 and HT29-MTX cells in a 9:1 ratio was seeded on top of the hydrogel layer at a seeding density of 10⁵ cells/cm² in 0.5 mL of DMEM. Then, 1.5 mL of DMEM was added to the basolateral side, and the plates were incubated with a humidified atmosphere at 37 °C and 5% CO₂.

The model was maintained for 21 days to promote the differentiation of Caco-2 cells into enterocyte-like cells, resulting in the formation of dense microvilli on their surface (Lozoya-Agullo et al., 2017), while HT29-MTX cells produced a functional mucus layer, completing the structure of the intestinal barrier. Details on cell maintenance and sources are provided in the Supplementary Material (SI).

Cell viability

CS-NPs and NPs were tested for cell viability at concentrations of 3.125, 6.25, 12.5, 25, 50, and 100 μg/mL on the different cell types. A CS-only control was not included since the cytocompatibility of chitosan is well documented in the literature (Howling et al., 2001; Noi et al., 2018). Cell viability was assessed using a resazurin sodium salt (Sigma-Aldrich) assay in standard 96-well plates (Garizo et al., 2021). Resazurin fluorescence was measured at 530–590 nm using a microplate reader (Synergy MX, BioTek). The procedure was compliant with ISO 10993-5:2009 standards for biocompatibility testing (ISO 1099ISO 10993-5, 2024).

Translocation and nutrient absorption

After 21 days, a solution of 0.1 mg/mL of CS-NPs and NPs, prepared separately in Hanks’ Balanced Salt Solution (HBSS, Corning), was applied to the apical compartment of the 3D model and left for 24 h. On day 22, at 1, 2, and 4 h, 200 μL samples were collected from the basolateral compartment and analyzed using an Iron Kit (Spectroquant®, HC300920), according to the manufacturer’s specifications. Absorbance was determined using a Thermo Scientific™ Varioskan™ LUX multimode microplate reader and data were normalized to the initial concentration in the apical compartment.

To evaluate nutrient absorption in the presence of CS-NPs, we assessed the paracellular and transcellular passage of substances across the layers (Figure 2). To this end FITC-dextran 40 (FD40 - Sigma-Aldrich), a commonly used molecule for investigating paracellular transport, and Rhodamine 123 (Rho123 – Gibco, MA, United States), a P-glycoprotein substrate, used to assess transcellular transport were employed as markers. Different solutions of FD40 and Rho123 were prepared and applied to the apical compartment of the 3D model. These were 0.5 mg/mL FD40 in HBSS, 0.5 mg/mL FD40 combined with 0.1 mg/mL CS-NPs in HBSS, 10 μM Rho123 in HBSS, and 10 μM Rho123 combined with 0.1 mg/mL CS-NPs in HBSS. The solutions were left in contact with the apical compartment for 24 h. On day 22, at 1, 2, and 4 h, 200 μL samples were collected from the basolateral compartment and analyzed using a Synergy MX multimode microplate reader (BioTek) in fluorescence mode, at wavelengths of 490–520 nm for FD40 and 488–529 nm for Rho123.

Actuation, proof of concept

As a preliminary demonstration of the feasibility of magnetic actuation of the Alg-CS-NPs, we fabricated a 2 mm thick hydrogel membrane that mimics the 1.6 mm thick alginate-collagen hydrogel used in the fibroblast layer of the 3D model. We encapsulated CS-NPs (0.1 mg/mL) and added 0.2 mg/mL of exogenous mucins. After cutting the hydrogel into a bar shape (2 × 2 × 10 mm³), it was placed in a Petri dish and magnetically actuated using an NdFeB magnet (BMN-42, 25 mm diameter, 186 N holding force).

To quantify the bar strain, we recorded the magnetic stimulation using a camera (see Supplementary Video S1, Supplementary Material) as the magnet was brought towards and then retracted from the dish. The video was processed in MATLAB. Each frame was binarized by applying an optimal threshold, and the images were cropped to isolate the region containing the hydrogel bar. The bar area was estimated by calculating the number of pixels covering the bar surface. Finally, the bar strain was calculated as the difference in area before and after magnetic stimulation, normalized to the initial area. This process was repeated for ten approach-retract cycles, and the resulting strain value was obtained as the means of the strains for each cycle.

Statistical analysis

Experiments were performed in triplicate. Statistical analyses were performed on GraphPad Prism through 2-way ANOVA tests and simple t-tests, with a 95% confidence interval for the multiple comparisons (Tukey’s method).

Results

Table 1 shows the dimensions of CS-NPs and NPs in deionized water at days 0 and 7. The NP average size approximately doubles after coating with CS (204.4–390.6 nm), indicating a shell thickness of around 100 nm. The presence of CS is confirmed by the higher zeta-potential value, exceeding 50 mV, compared to the 1–5 mV range of uncoated NPs. CS-NPs were stable over time and at room temperature. CS-NPs show a polydispersity index (PDI) of approximately 0.22, while uncoated NPs present a higher value of 0.43.

Table 1

Table 1. Size, zeta-potential, and polydispersity index of CS-coated and uncoated NPs immediately after preparation and after 7 days. The size distributions of coated and uncoated NPs, obtained from ZetaSizer analysis, are shown in Supplementary Figure S1 of the Supplementary Material.

The size of both coated and uncoated NPs remained stable as the pH increased (394.73 ± 24.67 nm and 220.31 ± 10.38 nm, respectively), as shown in Figure 3A. Figure 3B illustrates the zeta-potential of CS-NPs and NPs with pH. While the zeta-potential of uncoated NPs remains stable, it decreases with increasing pH in CS-NPs due to the deprotonation of CS (Oviedo et al., 2021).

Figure 3

Figure 3. Effect of pH on the size (A) and zeta potential (B) of CS-NPs and uncoated NPs, measured by the ZetaSizer. TEM BF images of CS-NPs (C) negatively stained with uranyl acetate reveal the grey chitosan shell surrounding the iron oxide core. In uncoated NPs, where the organic coating is absent, STEM ADF images (D) show bright, high-contrast cores on a dark background.

Uranyl acetate, used as a negative staining agent for TEM BF imaging, binds to the chitosan shell but not to the iron oxide core (Ong et al., 2017). TEM BF images show the presence of a CS shell (in grey) surrounding the black iron oxide core (Figure 3C). For uncoated NPs, STEM ADF provides high-contrast, sharp images of dense iron oxide cores, appearing white against a dark background (Figure 3D). Other TEM BF images of coated and uncoated NPs are shown in Supplementary Figures S2, 3 in Supplementary Material.

FTIR spectra are shown in Supplementary Figure S4 in the Supplementary Material. The FTIR spectrum of CS-NPs shows additional characteristic bands at 1,385 and 1,632 cm^-1 that are absent in uncoated NPs. These peaks are attributed to C–H bending vibrations and the amide I band (C=O stretching) of chitosan, respectively (Rahimzadeh et al., 2016).

The mucoadhesion results show that the size of CS-NPs increases by about 100% during the first hour and then stabilizes over time. In contrast, the size of NPs continuously increases over time, resulting in a 50% increase in size after 4 h (Figure 4A). The zeta-potential of CS-NPs decreases significantly (** = p < 0.01, **** = p < 0.0001) after 1 h and then stabilizes, while that of NPs gradually decreases over time (Figure 4B). The turbidimetric method further supports these findings, showing a significant (**** = p < 0.0001) difference between coated and uncoated NPs and a consistent trend with the other two measurements (Figure 4C).

Figure 4

Figure 4. Analysis of CS-NPs and NPs after 1–4 h of incubation in 40 mg/mL porcine stomach mucin. (A) Percentage change in particle size with respect to initial size, (B) zeta potential (** = p < 0.01, **** = p < 0.0001), and (C) mucoadhesion from turbidity measurements expressed according to Equation 1 (**** = p < 0.0001).

Figure 5A shows that Alg-CS-NPs are essentially spherical. Figure 5B illustrates how the size of the pH-responsive hydrogel changes over time due to swelling. Alg-CS-NPs crosslinked for 15 min triple in size after 1 h of exposure to the simulated stomach environment and dissolve completely after 1.5 h. When alginate is crosslinked for 30 min, a similar trend is observed but with a 30-min delay, and the gel dissolves completely after 2 h. Increasing the alginate crosslinking time to 60 min results in increased stability. Alg-CS-NPs crosslinked for 60 min double in size after 3 h of exposure without complete dissolution.

Figure 5

Figure 5. Brightfield microscopy images of Alg-CS-NPs (A). Effect of crosslinking time on the size changes of the Alg-CS-NPs in simulated gastric fluid (B). Quantitative analysis of CS-NPs released into simulated small intestinal fluid, measured as nanoparticle concentration in solution over time, approaching complete release after 2 h (C).

Figure 5C shows the release profile of CS-NPs from 60-min crosslinked Alg-CS-NPs. Approximately 50% of the initial CS-NP concentration is released after 1 h, with complete release occurring after 2 h. No detectable CS-NP release was observed during exposure to the gastric-like environment.

The results of the cell viability assays are presented in Figure 6. Uncoated NPs exhibit a significant decrease in cell viability, particularly for HT29-MTX and HPMEC cells (*** = p < 0.0001). In contrast, CS-NPs maintain a viability above 70% (the threshold according to ISO 10993-5:2009) for all tested cell types and concentrations.

Figure 6

Figure 6. Resazurin assay results for the different cell types used in the advanced 3D multi-layered in vitro intestinal model depicted in Figure 2. (A) Caco-2; (B) HT29-MTX; (C) HIF; (D) HPMEC.The dashed line indicates the 70% viability limit, defined as the biocompatibility threshold in ISO 289 10993-5:2009 (**** = p < 0.0001).

Figures 7A,B indicate that neither NPs nor CS-NPs cross the intestinal barrier model in detectable amounts. The basal nanoparticle concentration is essentially zero, considering the iron kit’s 5% sensitivity limit. Conversely, the apical concentration is approximately 100%.

Figure 7

Figure 7. (A,B) Quantification of nanoparticle passage across the 3D intestinal barrier model through iron quantification assay. (C,D) Effects of CS-NPs on paracellular (FD40) and transcellular (RHO123) nutrient transport (values normalized to the initial concentration in the apical compartment). Different letters indicate statistically significant differences (p < 0.05).

Moreover, the addition of CS-NPs to the apical compartment did not exert significant effect on the passage of FD40 or RHO123 compared with controls at any time point (2-way ANOVA). In both treated and untreated control conditions paracellular transport (FD40) remained stable over time, whereas transcellular transport (RHO123) progressively increased. The observed permeability values were within the expected ranges (10%–20% for FD40% and 20%–40% for RHO123, respectively, Figures 7C,D) (Patient et al., 2019).

Finally, upon exposure to a neodymium magnet, the bar-shaped hydrogel sample visibly bent and progressively conformed to the contour of the Petri dish in response to the applied magnetic field (Supplementary Video S1, Supplementary Material) with a strain of approximately 5.45% ± 0.31%. The deformation was reversible for up to 10 cycles.

Discussion

Despite significant clinical advancements, effective treatments for intestinal motility disorders still represent a significant unmet need. Current therapeutic options often involve invasive procedures or have limited efficacy. For instance, sacral nerve stimulation, while beneficial for some patients, may not be suitable for those with small bowel dysmotility. Similarly, pharmacological interventions, such as prokinetic agents, often have limited efficacy and can be associated with adverse effects. Our proposed minimally invasive magnetic strategy offers a novel approach to address these limitations. By leveraging the properties of mucoadhesive magnetic NPs, we aim to provide a targeted and effective solution for restoring intestinal motility. This proof-of-concept study is focused on the development of mucoadhesive magnetic NPs that can be orally delivered and then adhere to the intestinal mucosa, providing a means for actuation of the intestinal wall via external magnets.

To prevent systemic absorption, we selected NPs larger than the paracellular spaces, which typically range from 10 to 15 nm (Madara and Pappenheimer, 1987) in physiological conditions but can increase up to 100 nm in pathological conditions such as Inflammatory Bowel Diseases (Garbati et al., 2024) or infections (Lee, 2015). Therefore, we chose 200 nm NPs, which increased to approximately 400 nm after coating with CS. Our translocation studies confirmed that CS-NPs remained confined to the apical chamber, indicating that they did not penetrate the intestinal barrier.

The CS-NP concentration was chosen as a compromise between obtaining a stable nano-range colloidal suspension above the cells and maximising the concentration to increase the magnetic response of NPs. In addition, in our experimental design, the CS-NPs are intended to reside in the intestinal mucosa for a limited period of up to 24 h (phase 4 of our proposed strategy), after which they are cleared from the body. Under these conditions, the CS coating markedly improved the cytocompatibility of the NPs, as demonstrated by the increased cell viability in all cell types within the 3D model. This finding is consistent with the well-established biocompatible and non-toxic properties of CS.

Our results show that the CS-NPs are stable at various pH levels and retain their mucoadhesive properties. The mucoadhesive properties of CS-NPs are attributed to the formation of multiple types of bonds with mucins, including ionic bonds (due to the different external charges of the molecules, positive for CS and negative for mucins), hydrogen bonds (due to hydrogen atoms bound to highly electronegative atoms like oxygen or nitrogen), and Van der Waals forces (weak but significant when present in large numbers) (Garbati et al., 2024). Despite deprotonation at higher pH, CS-NPs can still bind to mucins, confirming their suitability for oral delivery.

To deliver CS-NPs to the bowel, we employed an alginate-based pH-responsive hydrogel. Alginate, an anionic polymer with a pKa of approximately 3.5 (Lee and Mooney, 2012), forms strong ionic bonds with the positively charged CS coating of the NPs at low pH. The effects of this interaction and the shrunken form of alginate under pKa prevent the release of CS-NPs in the acidic gastric environment. As the pH increases in the small intestine, the alginate hydrogel undergoes deprotonation (Chuang et al., 2017), weakening the ionic bonds and facilitating the release of CS-NPs (Figure 5). Additionally, the deprotonation of CS at higher pH values (∼6.5 (Wang et al., 2006), depending on the deacetylated degree) further contributes to the release process.

A 3D intestinal barrier model featuring a biomimetic multi-layered structure with fibroblasts embedded in a lamina propria-like hydrogel was used to investigate the translocation of NPs and their effect on nutrient transport. The presence of CS-NPs did not significantly impact nutrient absorption, as demonstrated by the unchanged paracellular (FD40) and transcellular (Rho123) transport rates (Figures 7C,D) across the barrier, making it suitable for testing nanoparticle-mediated mechanical stimulation aimed at restoring intestinal motility.

Future studies are required to address several limitations of the present work. First, the clearance of CS-NPs from the bowel must be investigated to determine their residence time in situ. Moreover, additional analyses of nutrient absorption involving other molecules and transport pathways, such as fatty acids, are needed to fully assess potential interactions with intestinal physiology. Finally, the proposed technology should be validated at the macro-scale, for instance through experiments in ex vivo intestinal tissues or intestinal phantoms, to evaluate the feasibility of the overall strategy.

Although it is limited to in vitro conditions, this proof of concept study represents a significant step towards the development of novel therapies for intestinal motility disorders and paves the way for further investigations using more physiological scenarios. The clearance and residence time of CS-NPs in the gastrointestinal tract must be validated in vivo, accounting for physiological mucus turnover and motility. While the 3D intestinal model supports early assessments of nanoparticle impact on barrier function, a broader range of nutrients -including lipids and peptides - and long-term exposure should be evaluated. Finally, actuation was demonstrated in a simplified hydrogel model; validation in ex vivo or bioinspired dynamic intestinal phantoms is necessary to assess mechanical performance under realistic tissue constraints.

Further research is also needed to assess the in vivo efficacy and safety of the NPs, optimize the formulation for enhanced delivery and reduced toxicity, and investigate the potential for targeted delivery to specific regions of the intestine. Finally, the development of a suitable external magnetic stimulation system is necessary for the successful application of this technology.

Conclusion

This study demonstrates the feasibility of using chitosan-coated magnetic nanoparticles (CS-NPs) for minimally invasive, targeted stimulation of intestinal motility. The CS-NPs showed strong mucoadhesive properties, pH-responsive protection via alginate encapsulation, and cytocompatibility across relevant gut cell types. Importantly, they did not disrupt intestinal barrier integrity or interfere with key transport pathways in a multi-layered intestinal model. The ability to generate physiologically relevant strain through external magnetic actuation further supports their potential as active biomechanical agents. These findings lay the groundwork for a new class of smart, responsive materials to support intestinal function and address motility disorders.

Data availability statement

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding author.

Ethics statement

Ethical approval was not required for the studies on humans in accordance with the local legislation and institutional requirements because only commercially available established cell lines were used.

Author contributions

PS: Data curation, Formal Analysis, Investigation, Methodology, Software, Validation, Writing – original draft, Writing – review and editing. LC: Investigation, Methodology, Supervision, Validation, Writing – review and editing. FF: Formal Analysis, Writing – original draft. CM: Methodology, Writing – review and editing. BS: Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Writing – review and editing. AA: Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Writing – original draft, Writing – review and editing.

Funding

The author(s) declared that financial support was received for this work and/or its publication. PS was funded by a STSM grant from COST Action IMPROVE (CA21139), supported by COST (European Cooperation in Science and Technology). The work was partially supported by the Italian Ministry of University and Research (MUR) in the framework of the FIS2 project INTERCELLAR (CUP I53C2400311000).

Conflict of interest

The authors declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The author AA declared that they were an editorial board member of Frontiers at the time of submission. This had no impact on the peer review process and the final decision.

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Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fnano.2025.1716360/full#supplementary-material

Footnotes

^{Abbreviations:}Alg-CS-NPs, Alginate-encapsulated CS-NPs; CS, Chitosan; CS-NPs, Chitosan-coated magnetic nanoparticles; HBSS, Hanks’ Balanced Salt Solution; HIF, Human intestinal fibroblasts; HPMEC, Human pulmonary microvascular endothelial cells; IBS, Irritable bowel syndrome; NPs, Magnetic nanoparticles; PBS, Phosphate-buffered saline; VM, Visceral myopathies.

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