Acellular lung matrix scaffold coated by fibronectin enhances hepatic differentiation of adipose-derived mesenchymal stem cells

Abstract

End-stage liver disease is a multifactorial disorder and leading cause of death. The development of effective tissue models has been a focus of research for treating liver disease. This study investigated the use of acellular lung matrix (ALM) and fibronectin (FN) to culture adipose-derived mesenchymal stem cells (ADMSCs) and promote their differentiation into hepatic-like cells. In this experimental study, sodium deoxycholate and NaCl were used to decellularize the lungs, and scaffolds were characterized by histology, scanning electron microscopy, and DNA quantification. ADMSCs were differentiated into hepatocyte-like cells using a two-step protocol under three conditions: in two-dimensional culture, on ALM scaffolds, and on FN-coated ALM scaffolds. Hepatic differentiation was confirmed by RT-PCR for specific genes (alpha-fetoprotein [AFP], albumin [ALB], cytokeratin-18 [CK-18]), ALB and urea levels were measured using specific commercial kits. The main extracellular matrix components (collagen and glycosaminoglycans) were preserved, and residual DNA was minimal (P<0.001), indicating effective decellularization. ADMSCs cultured on FN-coated ALM exhibited significantly higher expression of hepatic markers (AFP, P<0.01; ALB, P<0.01; CK-18, P<0.01). Functional assay confirmed higher ALB and urea production (P<0.05), while MTT assay demonstrated that the scaffold was non-cytotoxic. Collectively, these results demonstrate that FN-coated ALM holds considerable potential for facilitating the hepatic differentiation of ADMSCs under the applied induction conditions.

Introduction

Cirrhosis is a liver condition and is characterized by the gradual destruction of liver tissue over time. Persistent damage leads to hepatic scarring (fibrosis), which, if unopposed, progresses to cirrhosis and the failure of liver function. End-stage liver fibrosis is characterized by cirrhosis, in which fibrotic bands, parenchymal nodules, and vascular distortion disrupt normal liver architecture. Portal hypertension and hepatocyte dysfunction are consequences that lead to significant systemic issues and early mortality [1, 2]. The primary etiological agents of cirrhosis worldwide are infection with hepatitis B and C viruses and the excessive consumption of alcohol. Egypt exhibits the highest global prevalence of hepatitis C virus, reaching approximately 20% in some populations [3].

Orthotopic liver transplantation (OLT) represents the sole therapeutic intervention once liver disease progresses to the irreversible cirrhotic stage. However, the growing shortage of donor organs limits liver transplantation. However, the utility of OLT is constrained by a critical shortage of donor organs. This disparity between supply and demand is widening amid rising rates of liver disease, condemning many patients to a lifetime of immunosuppression and poor outcomes. Consequently, these limitations underscore the urgent need to develop novel therapeutic strategies to complement or supplement OLT [2, 4].

Liver transplantation remains the only curative treatment for end-stage liver disease; however, its application is severely limited by a critical donor organ shortage. This challenge has generated substantial interest in developing alternative therapeutic strategies. A promising approach to address the donor shortage is liver tissue engineering based on decellularized liver matrix (DLM) scaffolds. In this strategy, hepatic cells are seeded into DLM scaffolds to regenerate functional liver-like tissue for supplementing liver function [5, 6]. The decellularized scaffold offers a more favorable environment for cell attachment, proliferation, maturation, and function than previously employed synthetic or natural three-dimensional (3D) materials [7-11] since it retains site-specific natural extracellular matrices (ECM) and fundamental organ structural frameworks, which are difficult to reproduce artificially.

Additionally, the vascular networks preserved in the decellularized scaffold also provide pathways for perfusion and implantation in vivo. Research has shown that decellularized matrix-based tissue regeneration is achievable for many hollow and parenchyma organs, including the bladder [12], skin [13], blood vessels [14], heart [15], lung [16], kidney [17], as well as the liver [18].

However, decellularized scaffolds often exhibit critical limitations, including incomplete repopulation of the parenchymal compartment, and a tendency toward thrombus formation. To address these widespread challenges, we have developed a simple and safe surface modification strategy to significantly improve the recellularization efficiency of decellularized organ scaffolds.

Previous research has established that surface modification can significantly enhance the biocompatibility of scaffolds without compromising their fundamental structural properties. A range of techniques exists for this purpose, including enzymatic and chemical methods [19, 20].

Among these, one of the most widely utilized approaches is the physical adsorption of coatings with desirable biological functionalities onto the scaffold material [21].

One of the most important components of the ECM is fibronectin (FN). FN is classified as a glycoprotein characterized by its high molecular weight. The main function of FN is to serve as a scaffold for cell adhesion and migration, thereby also regulating cell proliferation and differentiation [19]. Surface modification with FN promotes cell attachment by presenting specific binding sites for cell-surface integrins, thereby stabilizing adhesion [22]. This anchored connection is vital for cell viability, as it inhibits apoptosis induced by cell detachment [23]. Furthermore, integrin-mediated binding to FN activates intracellular signaling cascades that support cell proliferation and survival. For example, FN-coated surfaces have been shown to markedly enhance the attachment and expansion of diverse cell types, such as endothelial cells, hepatocytes, fibroblasts, and osteoblasts [23].

FN demonstrates superior biocompatibility and bioactivity compared to synthetic alternatives, minimizing immunogenic response while actively supporting the cellular processes essential for tissue integration and function [24]. It can be adapted into various formats—such as coatings, scaffolds, and hydrogels—to meet specific requirements in tissue engineering. This multifunctional capability establishes FN as a highly effective biomaterial for applications demanding rapid tissue regeneration and integration [25]. The aim of the present study was to evaluate the feasibility of a FN-acellular lung matrix (ALM) as a 3D extracellular matrix scaffold for supporting the in vitro adhesion, proliferation, and hepatogenic differentiation of adipose-derived mesenchymal stem cells (ADMSCs) under defined induction conditions. We hypothesized that FN coating would improve cell adhesion and viability on the ALM scaffold, thereby facilitating a more stable and supportive microenvironment for hepatogenic induction without implying enhancement of lineage-specific specification beyond the applied differentiation protocol.

Materials and Methods

Lung decellularization

The Naval Medical Research Institute mice were obtained from the animal center of Ahvaz Jundishapur University of Medical Sciences. The experiment was approved by the Animal Ethics Committee of Ahvaz Jundishapur University of Medical Sciences (code: IR.AJUMS.AEC.1404.035). The lung of the healthy mice under deep anesthesia was removed and excessive tissues around the lung were removed. The whole lung underwent a decellularization process according to an established protocol [26]. Initially, the tissue was rinsed in distilled water (DDW; INOCLON) for 30 minutes to eliminate residual blood cells. Decellularization was then achieved by incubating the tissue in a 1% sodium deoxycholate (302-95-4, Sigma-Aldrich) solution at 4°C for 72 hours. Following this, the tissues were treated with a 1 M NaCl solution (7647-14-5; Sigma-Aldrich) for one hour. Upon completion of these steps, the resulting acellular lung matrix was thoroughly rinsed with DDW over 48 hours. A brief immersion in 70% ethanol (64-17-5; Merck) for 10 seconds was performed for disinfection. Finally, to remove any residual ethanol, the samples were washed with phosphate-buffered saline (PBS; 10010023, Gibco) every 12 hours for three days. The decellularized lung samples were cut into small pieces and stored at 4°C until further use.

Histological staining

Five decellularized lung samples were fixed in 4% formaldehyde for 4 hours, processed, paraffin-embedded, and sectioned into 6 µm slices. Sections were stained with hematoxylin and eosin (H&E) and DAPI to verify cell removal, Masson’s trichrome for collagen, and Alcian Blue for GAGs. The stained sections were examined and imaged using a light microscope (BX51; Olympus) equipped with a DP72 digital camera.

Assessment of residual DNA

The saline extraction method was used to isolate DNA from both decellularized and native tissues [27]. To assess and quantify total DNA content, 30 mg of lyophilized native and decellularized lung tissues were treated with digesting buffer solution containing 1 M Tris, 5 M NaCl, 0.5 M EDTA, pH 8 in the presence of proteinase K and 10% sodium dodecyl sulfate (lysis buffer) overnight at in water bath. DNA was precipitated with 5 M NaCl, while the proteins were eliminated from the mixture. The purified DNA was redissolved in 100 μl of distilled water, and its concentration was assessed using a NanoDrop spectrophotometer (Multiskan spectrum; Thermo Scientific).

Scanning electron microscope

Both cell-seeded and unseeded scaffolds (FN-coated ALM and ALM) were fixed with 2.5% glutaraldehyde prepared in PBS (pH 7) and subsequently washed twice with PBS. Subsequently, the samples underwent post-fixation with 1% osmium tetroxide, were dehydrated through a series of graded ethanol solutions, and were then subjected to critical-point drying with liquid carbon dioxide. The dehydrated samples were coated with a thin layer of gold and subsequently examined for surface topography using a scanning electron microscope (Hitachi).

Cell cytotoxicity using the MTT assay

The biocompatibility of the ALM scaffold was assessed using the MTT assay after 10 days of cell culture, with cells cultured in scaffold-free wells serving as controls. Briefly, 1×10⁴ cells per well were placed into ALM scaffolds within 24-well plates. Following incubation, the medium was removed, and 5 g/L MTT solution was added to each well and incubated in the dark for 4 hours. The solution was then replaced with 1 ml DMSO to dissolve the formazan crystals, followed by a 10-minute incubation in the dark. Absorbance was measured at 570 nm using a microplate spectrophotometer (Epoch; BioTek).

Differentiation potential assay of ADMSCs

The ADMSCs were acquired from Iran’s National Center for Genetic and Biological Reserves. To confirm the stemness of the purchased cells, their ability to differentiate into the osteogenic lineage was assessed. Osteogenic induction was performed in culture medium supplemented with 1 nM dexamethasone (Sigma), 2 mM β-glycerophosphate (Sigma), and 50 mM ascorbate-2-phosphate (Sigma) for 3 weeks. Following fixation with 4% formalin, mineral deposition was visualized using 40 mM Alizarin Red staining (pH 4.1, Sigma) [28].

Fibronectin coating and recellularization of scaffolds

In this study, we defined three experimental groups: a two-dimensional (2D) culture group, an ALM scaffold group, and a FN-coated ALM scaffold group. Scaffolds were coated with FN (Sigma-Aldrich) by incubation in a 50 µg/ml solution prepared in sterile PBS for 24 hours at 37°C under static conditions, followed by three washes with PBS to remove unbound protein prior to cell seeding [29]. The ALM scaffolds were sterilized with 70% ethanol for 24 hours and placed in a 24-well culture plate. Following a 24-hours preincubation in complete medium, passage-3 ADMSCs (1×10⁶ cells) were resuspended in 100 μl of culture medium and seeded onto the scaffolds. Subsequently, 1 ml of culture medium was added to each well. The differentiation protocol was conducted as follows: The culture medium was changed 24 h later to Iscove’s modified Dulbecco’s medium containing 20 ng/ml epidermal growth factor, 20 ng/ml hepatocyte growth factor, 10 ng/ml basic fibroblast growth factor and 0.61 g/L nicotinamide (Sigma) and the ADMSCs were cultured in this for 7 days. Thereafter, the maturation step consisted of treatment with IMDM containing 20 ng/ml oncostatin M, 1 µmol/L dexamethasone (Sigma) and 1% insulin–transferrin–selenium premix for 14 days. For each step, the culture medium was changed every 3 days [30].

Gene expression analysis

Total cell RNA was extracted using the RNeasy Mini Kit (Qiagen) in accordance with the manufacturer’s guidelines. Subsequently, reverse transcription reactions were performed with 1 µg of total RNA utilizing the Prime Script^TM RT reagent kit (Takara). Real-time RT-PCR was conducted with SYBR premix ExTaq (Takara Bio) using an ABI one-step system alongside specific primers. The primers used in this research are as follows: GCAGACAATCCAGCACATCTC (forward alpha-fetoprotein [AFP]), GCCAAAGTGAAGAGGGAAGAC (reverse AFP); CAGACACCACTTTGCCATC (forward cytokeratin-18 [CK-18]), ACAGTCTGCTGAGGTTGGA (reverse CK-18); TGGCAAGTCTCAGCAGCA (forward albumin [ALB]), TCAAGTGTGCCAGTCTCCAA (reverse ALB); CATCACTGCCACCCAGAAGACTG (forward GAPDH), ATGCCAGTGAGCTTCCCGTTCAG (reverse GAPDH). The PCR amplification was conducted using a 45-cycle program consisting of the following steps: 95°C for 10 sec, 95ºC for 15 sec, and 60°C for 20 sec. Relative gene expression levels were calculated using the 2^-∆∆Ct method, with normalization to GAPDH. The specificity of qPCR amplicons was confirmed by melting curve analysis, with no primer-dimer formation observed.

Functional assay

Cell culture medium supernatants were collected on Day 14 after induced differentiation. Albumin secretion was quantified using an albumin assay kit (H-7650; HITACHI) according to manufacturer’s instructions. To evaluate the hepatocyte-mediated biotransformation of ammonia to urea, the concentration of urea in the medium was measured using a urea assay kit (Abcam).

Glycogen staining

The functional assay of the hepatocyte-like cells was conducted using periodic acid–Schiff (PAS) staining to identify glycogen storage. Both 3D and 2D cultures were fixed with 4% paraformaldehyde. Following this, samples were rinsed with distilled water and then incubated in 0.5% periodic acid for 5 minutes. Subsequently, the cultures were washed again and treated with Schiff’s reagent for 15 minutes. The cells were then counterstained with hematoxylin.

Statistical analysis

Data analysis was performed using SPSS software (version 22.0; IBM Co.). One-way analysis of variance was applied, followed by post hoc pairwise comparisons using Tukey’s or least significant difference tests. For non-parametric data, the Kruskal–Wallis test was employed. A P-value <0.05 was considered statistically significant. Each assay was conducted a minimum of three times.

Results

Characteristics of acellular lung matrix scaffold

After approximately one week of decellularization, the lung tissues became completely transparent (Fig. 1). H&E staining confirmed the effective removal of both nuclear and cytoplasmic contents from the ALM following decellularization (Fig. 1). In Masson’s Trichrome-stained sections, the native lung shows dense cellular components and abundant collagen fibers distributed around alveolar walls, bronchioles, and vascular structures, indicating the complex cellular and stromal organization of intact tissue. After decellularization, nuclei and cytoplasmic components are completely removed, while the collagenous framework remains intact, confirming the preservation of the 3D ECM network essential for structural integrity (Fig. 2). Alcian Blue staining reveals a prominent presence of glycosaminoglycans (GAGs) in the native tissue, visible as intense blue coloration within the interstitial matrix. In contrast, a marked reduction in blue staining is observed in the acellular scaffold, signifying partial loss of GAGs during detergent-based decellularization while maintaining the overall ECM architecture (Fig. 2). PAS staining further illustrates the presence of carbohydrate-rich components, such as basement membrane glycoproteins, in native tissue. Following decellularization, the faint PAS signal indicates efficient removal of cellular remnants and cytoplasmic glycoconjugates while preserving the fine alveolar microstructure (Fig. 2).

Fig. 1.

Optical and microscopic (H&E-stained) images of native and decellularized lungs. Representative optical (macroscopic) images of native mice lung and decellularized lung scaffold. H&E staining of native lung tissue shows dense cellularity, intact alveolar septa, and the presence of nucleated cells. In contrast, H&E staining of decellularized lung demonstrates acellular alveolar walls with preserved extracellular matrix architecture and only rare residual nuclei. Scale bar=200 μm.

Fig. 2.

Histological characterization of native and decellularized lung tissues using Masson’s Trichrome, Alcian Blue, and periodic acid–Schiff (PAS) staining. All images were acquired at 200× magnification; scale bar=200 μm. Masson’s Trichrome staining: Native lung tissue shows collagen fibers surrounding airways. Decellularized lung retains a well-preserved blue collagen network, indicating intact extracellular matrix (ECM) after decellularization. No significant disruption of the collagen framework was observed. Alcian Blue staining: Native lung exhibits weak to moderate staining of glycosaminoglycans (GAGs) in the basement membrane and peri-bronchial regions. Decellularized lung shows retained Alcian Blue positivity, although with slightly reduced intensity, suggesting partial preservation of GAGs within the ECM. PAS staining: Native lung demonstrates strong PAS positivity in the basement membranes, reflecting the presence of neutral glycoproteins. Decellularized lung shows markedly reduced but still detectable PAS staining, indicating partial loss of carbohydrate-rich components while preserving the underlying ECM architecture.

Residual DNA assessment

Representative DAPI staining images of native and ALM show the complete removal of nuclear material after decellularization. The native tissue exhibits abundant blue fluorescence, indicating intact nuclei, whereas the ALM shows an absence of nuclear staining, confirming effective cellular removal. Quantitative analysis of DNA content revealed a significant reduction in residual DNA in the ALM (192.86±18.3 ng/mg) compared to the native lung tissue (4,520.3±123.6 ng/mg) (P<0.001), demonstrating the efficiency of the decellularization process (Fig. 3).

Fig. 3.

Assessment of residual nuclei and DNA content in native and decellularized lung tissues. Fluorescence microscopy of DAPI-stained samples: representative fluorescence images of native lung (A) and decellularized lung (B) stained with DAPI, a nuclear dye. Native lung tissue shows abundant bright blue nuclei, confirming high cellularity. In contrast, decellularized lung tissue exhibits only a few scattered residual nuclei, demonstrating efficient removal of cellular material. Magnification 200×. (C) Quantitative DNA content analysis: bar graph showing DNA content (ng/mg of dry tissue weight) in native versus decellularized lung tissues. Decellularized lung tissue shows a dramatic reduction in DNA content compared to native tissue. ***P<0.001.

Scanning electron microscopy

The scanning electron microscopy analysis confirms that the decellularization protocol successfully generated a porous ECM scaffold while preserving its intrinsic ultrastructure. The subsequent FN coating and recellularization with ADMSCs resulted in a biocompatible construct where the cells actively engage with the matrix. This combination of native lung ECM topographical guidance and FN-derived pro-adhesive signals creates a synergistic microenvironment, thereby supporting the hypothesis that this scaffold is a highly conducive platform for the hepatic differentiation of ADMSCs (Fig. 4).

Fig. 4.

Scanning electron microscopy micrographs of acellular lung matrix (ALM) before and after fibronectin (FN) coating and following recellularization with adipose-derived mesenchymal stem cells (ADMSCs). (A) ALM-uncoated: the decellularized scaffold retains a porous, fibrous extracellular matrix (ECM) architecture with open pore structures. No cellular debris or surface deposits are observed. (B) FN-coated ALM (FN-coated): following coating with FN, the scaffold surface appears denser and shows increased surface roughness with fine fibrillar deposits, indicating successful FN adsorption onto the ECM fibers. (C) ALM recellularized with ADMSCs (uncoated): ADMSCs seeded onto uncoated ALM show cell attachment and spreading. Cells appear round or ovoid with few cytoplasmic extensions, suggesting suboptimal adhesion and differentiation. (D) FN-coated ALM recellularized with ADMSCs: ADMSCs seeded onto FN-coated ALM exhibit extensive cell attachment, spreading, and elongation. Scale bar=20 μm.

Osteogenic potential assay

The plasticity of ADMSCs was also assessed by induction of osteogenic differentiation. Three weeks after osteogenic medium, we observed extracellular calcium phosphate precipitates, as shown by Alizarin-Red staining (Fig. 5).

Fig. 5.

Osteogenic differentiation of adipose-derived mesenchymal stem cells (ADMSCs). Representative images showing undifferentiated and differentiated ADMSCs under osteogenic induction conditions. (A) Undifferentiated ADMSCs: ADMSCs cultured in standard growth medium without osteogenic supplements. (B) Differentiated ADMSCs: without staining; ADMSCs subjected to osteogenic differentiation medium. (C) Differentiated ADMSCs: Alizarin Red staining; following osteogenic induction. Scale bar=40 μm.

MTT assay

The viability of cells seeded on ALM and FN-coated ALM was not significantly changed compared with the 2D culture at various time points. Over a culture period of up to 10 days, we periodically assessed cell viability. The results demonstrated that cells seeded on both ALM and FN-coated ALM consistently exhibited viability levels that were not statistically different from those maintained in 2D culture, indicating that the 3D ALM and FN-coated ALM environment does not compromise cell health (Fig. 6).

Fig. 6.

Assessment of scaffold cytotoxicity: viability of adipose-derived mesenchymal stem cells (ADMSCs) cultured on uncoated acellular lung matrix (ALM) and fibronectin (FN)-coated ALM (FN-coated ALM) over 10 days. ADMSCs were seeded on tissue culture plastic (two-dimensional [2D] culture; control), uncoated ALM, or FN-coated ALM. Cell viability was assessed at days 2, 4, 6, 8, and 10 using a standard cytotoxicity assay, and values are expressed as a percentage relative to the 2D culture control at each time point (set to 100%).

Gene expression

The mRNA expression levels of characteristic hepatic markers, including AFP, CK-18, and ALB, were assessed to evaluate the hepatic differentiation of ADMSCs. Gene expression analysis demonstrated that culture on ALM and FN-coated ALM not only maintained cell viability but also substantially improved differentiation efficiency. Specifically, cells differentiated on the 3D scaffold, including ALM and FN-coated ALM environment, showed a substantial upregulation in mRNA levels compared to those in 2D culture, with a 3.5-fold and 4.2-fold (ALM and FN-coated ALM, respectively) increase for AFP, a 4-fold and 4.5-fold increase for CK-18, and a 3-fold and 3.8-fold increase for ALB (Fig. 7).

Fig. 7.

Relative mRNA expression of hepatic markers (alpha-fetoprotein [AFP] and albumin [ALB]) and cytokeratin-18 (CK-18) in adipose-derived mesenchymal stem cells (ADMSCs) differentiated on two-dimensional (2D) culture, uncoated acellular lung matrix (ALM), and fibronectin (FN)-coated ALM. ADMSCs were differentiated toward hepatocyte-like cells under three conditions: conventional 2D tissue culture plastic (2D culture), ALM, and FN-coated ALM (FN-ALM). Fold changes in gene expression were calculated relative to undifferentiated ADMSCs (set to 1.0) after normalization to a housekeeping gene (GAPDH). Data are presented as mean±SD. **P<0.01.

Functional assay

The functional capacity of the differentiated ADMSCs was evaluated by measuring the secretion of key hepatic markers, albumin and urea, using specific commercial kits. The results confirm the higher performance of cells cultured on the ALM and FN-coated ALM scaffolds compared to those in 2D culture (Fig. 8).

Fig. 8.

Serum protein excretion of urea and albumin in two-dimensional (2D) and three-dimensional (3D) differentiated cell (acellular lung matrix [ALM] and fibronectin (FN)-coated ALM). Bar graph illustrating the significant difference between 2D and 3D differentiated cell. Data are presented as mean± SD. *P<0.05.

Periodic acid–Schiff staining

To evaluate hepatic differentiation, glycogen production and storage were assessed using PAS staining, which is commonly used to identify this functional feature. The assay was performed in parallel on cells differentiated under 2D monolayer conditions and within 3D culture (ALM and FN-coated ALM). The results of the PAS staining revealed a distinct and significant difference in glycogen content between the two culture systems. Hepatocyte-like cells differentiated in the 3D culture configuration demonstrated markedly intensified PAS-positive staining compared to their 2D counterparts (Fig. 9).

Fig. 9.

Representative micrographs of periodic acid–Schiff (PAS) staining demonstrating intracellular glycogen storage in differentiated cells after 14 days of hepatic induction. Cells cultured under three different conditions were fixed and subjected to PAS staining, a histochemical method used to detect glycogen and other neutral glycoproteins through oxidation of carbohydrate residues and subsequent Schiff reagent reaction, resulting in a characteristic magenta/purple coloration. (A) Cells differentiated in two-dimensional culture; (B) cells differentiated on uncoated acellular lung matrix (ALM) scaffold; (C) cells differentiated on fibronectin (FN)-ALM scaffold. PAS-positive staining indicates intracellular glycogen accumulation, reflecting a metabolic feature associated with hepatic lineage differentiation. Scale bar=100 μm.

Discussion

In this study, we developed a 3D differentiation platform using ALM and FN-coated ALM (FN-ALM) as supportive extracellular matrix scaffolds for the differentiation of ADMSCs into hepatocyte-like cells under hepatogenic induction conditions. Rather than serving as a tissue-specific inducer of hepatic lineage commitment, ALM was employed as a permissive 3D microenvironment capable of supporting cell attachment, survival, and structural organization during the differentiation process. Our findings demonstrated that the ALM-based scaffolds enhanced hepatic differentiation outcomes compared with 2D culture conditions, as reflected by increased expression of hepatic-associated markers, elevated secretion of liver-related proteins, and improved hepatocyte-like functional properties.

The success of any acellular scaffold depends on the effective removal of cellular material while maintaining the intricate structure and biochemical composition of the native ECM [31, 32]. The native lung ECM possesses key biochemical and biophysical characteristics that confer biocompatibility, elasticity, and porosity, thereby facilitating cell attachment and influencing cell fate decisions [33, 34].

Our decellularization protocol provides a translucent ALM with minimal residual DNA content, confirming efficient cell removal. Removing residual DNA and cellular debris results in improved cell positioning within the scaffold, reduces the immune reaction, and enhances the viability of the cultured cells [35]. The primary components of the ECM, including collagen and GAGs, were also detected in the ALM. Our findings demonstrated that the ALM microenvironment can preserve cell viability throughout the entire differentiation process, suggesting that ALM scaffold is capable of supporting both cell differentiation and long-term culture.

While tissue-engineered constructs represent a pioneering approach for replacing damaged tissues, their clinical success is often limited by persistent hurdles, including the difficulty of establishing functional vascular networks and the subsequent risk of thrombosis [36]. Therefore, improving scaffold biocompatibility to enhance recellularization efficacy is critical for overcoming these challenges. FN-coating provides notable benefits, such as stronger cellular adhesion, increased cell growth, and lower rates of programmed cell death. Together, these effects support more consistent recellularization and help decrease the potential for clot formation [29]. The application of FN-coating in this study was intended to enhance the scaffold’s functionality by improving cell attachment, proliferation, and survival [37].

In a previous study, Xiang et al. [38] examined the efficacy of acellular spleen matrix in culturing bone marrow mesenchymal stem cells (BMSCs) and enhancing their differentiation into hepatic-like cells. The findings indicated that acellular spleen matrix might have considerable potential in fabricating hepatic-like tissue, especially since it can facilitate hepatic differentiation of BMSCs, which exhibited higher levels and more stable functions [38].

We evaluated liver-specific gene expression to compare hepatic differentiation between 2D and 3D cultures (ALM and FN-coated ALM). At the molecular level, the significant upregulation of hepatocyte-specific genes, including ALB, CK-18, and AFP, in the 3D cultures underscores the scaffold’s ability to promote a more robust and specific hepatic differentiation pathway.

Furthermore, gene expression in cells differentiated on the FN-coated ALM was higher than in those cultured on uncoated ALM scaffolds, indicating that the FN coating may provide a bioactive advantage in supporting hepatic differentiation.

Our functional assays indicate enhanced hepatic differentiation in the 3D system. The differentiated cells on the ALM and FN-coated ALM showed higher levels of albumin and urea secretion compared to 2D culture, which are commonly used indicators of hepatocyte-like function. Furthermore, PAS staining indicated glycogen storage, reflecting a metabolic feature associated with hepatic differentiation.

The choice of acellular lung matrix, as opposed to a liver matrix (DLM), is a notable aspect of our study. While acellular liver matrix is the ideal scaffold for liver engineering, its widespread application is severely hampered by the global shortage of donor livers. In contrast, lung tissue can be more readily obtained from various sources, such as patients undergoing resection for trauma or disease. Our research demonstrates that ALM, despite its origin, possesses a biocompatible ECM that can effectively support the differentiation and function of hepatocyte-like cells. This “cross-organ” approach could potentially circumvent the critical limitation of donor organ scarcity for scaffold generation in liver tissue engineering. However, this study has several limitations. The scaffold used is derived from mouse lung, and all experiments were performed in vitro, which limits the direct translational relevance of the findings. No in vivo implantation or evaluation of immune response, vascular integration, or long-term functional stability was conducted. Future studies should therefore include in vivo assessment to determine biological performance under physiological conditions, as well as consider the use of human-derived acellular matrices to improve clinical relevance.

Conclusion

We have established a proof-of-concept for the use of FN-ALM as a supportive 3D scaffold for generating hepatocyte-like cells from ADMSCs under defined hepatogenic induction conditions. In the present study, the preserved native extracellular matrix architecture of the lung was utilized primarily to provide a permissive microenvironment that facilitates cell attachment, survival, and spatial organization.

Limitation

Several limitations should be acknowledged. The mouse- derived lung scaffold raises concerns regarding xenoantigens and species-specific ECM signaling, and all experiments were performed in vitro without in vivo validation of biocompatibility or vascular integration. The study lacks a direct comparison with the gold standard DLM and does not include proteomic characterization of liver-specific ECM components (e.g., laminin isoforms, collagen IV). Consequently, the observed supportive effects on hepatic differentiation cannot be attributed to hepatic-specific signals, and the underlying mechanisms remain speculative. Future studies should address these limitations before any clinical translation can be considered.