Idiopathic pulmonary fibrosis (IPF) is a progressive and fatal interstitial lung disease (ILD) characterized by excessive extracellular matrix (ECM) deposition, loss of alveolar structure, and respiratory failure. The disease primarily affects older adults and has a poor prognosis, with a median survival of only 3–5 years after diagnosis ^[1]. Current anti-fibrotic therapies can slow progression but neither halt nor reverse fibrosis ^[2]. The lack of curative options highlights the urgent need for human-derived disease models that recapitulate IPF pathogenesis ^[3].

Alveolar type II (AT-II) epithelial cells play a central role in alveolar homeostasis and fibrotic remodeling. These cuboidal surfactant-producing cells constitute ~ 5% of the alveolar surface, secrete surfactant, regulate immune responses, and act as progenitors for alveolar type I (AT-I) cells ^[4]. Under normal conditions, AT-II cells self-renew and transdifferentiate into AT-I cells after injury ^[4]. In IPF, however, AT-II cells undergo apoptosis, senescence, or aberrant activation, contributing to chronic epithelial damage and fibrosis ^[5]. In addition to these processes, AT-II cells in fibrotic lungs can give rise to aberrant basaloid cells, a transitional epithelial state observed in distal fibrotic regions of IPF ^[6–8]. Recent studies, including single-cell transcriptomic analyses, suggest that matrix stiffening and altered mechanical cues drive this basaloid differentiation program ^[6]. Such stiffness-induced epithelial transitions have been demonstrated in vitro and in vivo, where direct contact with rigid substrates promotes loss of AT-II identity and activation of basal markers ^{[6, 8–9]}. They may further sustain the fibrotic niche by secreting profibrotic cytokines, undergoing epithelial–mesenchymal transition (EMT), and serving as progenitors of myofibroblasts ^[10]. AT-II cells can be identified by lineage-defining markers such as surfactant proteins (SFTPC, SFTPB), ABCA3 ^[11], LAMP3 ^[12], NKX2-1/TTF-1, and the surface marker HT-II-280 used for human isolation ^[13–14]. Morphologically, the presence of lamellar bodies is a characteristic hallmark of AT-II cells ^[15–16]. AT-I cells, in contrast, cover ~ 95% of the alveolar surface and express RAGE/ AGER, HT-I-56, and HOPX^17–18]. In addition to AT-II–derived fibroblast-like states, injured AT-I cells in IPF adopt dysfunctional/senescent programs that secrete pro-fibrotic mediators (e.g., TGF-β, CTGF, WNT ligands), thereby activating fibroblasts and amplifying ECM deposition; AT-I–restricted regulators such as AGER (RAGE) and caveolin-1 further modulate this crosstalk, and their dysregulation has been linked to heightened fibrosis^[19–22].

Recent studies showed that IPF-derived AT-II cells undergo metabolic reprogramming from oxidative phosphorylation to aerobic glycolysis, a “Warburg-like” phenotype. This is linked to mitochondrial dysfunction, elevated glycolytic enzyme expression (LDHA, PDK1), lactate accumulation, and suppression of pyruvate dehydrogenase via the PDK1–HIF-1α axis ^[23–25]. Such energetic shifts are thought to drive fibrotic remodeling and represent potential therapeutic targets.

Organoids—self-organizing 3D epithelial culture systems—have become valuable models for disease research. Pulmonary organoids derived from fetal, adult, or induced pluripotent stem cell (iPSC) sources can recapitulate alveolar architecture and function ^[26–28]. They allow long-term expansion, passaging, and cryopreservation while maintaining lineage identity, enabling applications in disease modeling, drug testing, and personalized medicine ^[29–30]. Several lung organoid models reproduced features of fibrosis, epithelial plasticity, and anti-fibrotic responses ^[31–32]. Compared to 2D cultures, organoids better preserve epithelial polarity, surfactant secretion, and progenitor functions. However, robust primary human AT-II organoid models from end-stage fibrotic lungs are still lacking. Most existing models rely on murine or iPSC-derived epithelium and do not reflect the native metabolic or regenerative state of diseased AT-II cells.

Moreover, current systems often lack reproducibility across laboratories and limited scalability, restricting their broader translational use. A reproducible, feeder-free human AT-II model that allows cryopreservation and biobanking would therefore be highly valuable for precision medicine and cross-center validation.

Here, we establish a defined feeder-free 3D culture system for primary human HT-II-280 + AT-II cells isolated from both healthy and fibrotic (IPF and secondary fibrosis) lung tissue adapted from previous work by Konishi et al.^[33] and Katsura et al.^[27]. We aimed to (i) maintain AT-II identity across passages using canonical markers (HT-II-280, proSP-C, lamellar bodies), (ii) assess cryopreservation and potential for organoid biobanking, and (iii) evaluate metabolic differences between healthy and fibrotic AT-II organoids using extracellular flux analysis. We hypothesized that fibrotic AT-II cells exhibit enhanced glycolytic metabolism compared to controls, reflecting disease-specific metabolic reprogramming. By providing a robust and transferable protocol, our study delivers a reproducible human lung organoid resource that supports mechanistic and translational research in ILD.

Primary human alveolar type II (AT-II) and type I (AT-I) cells were isolated from freshly explanted fibrotic or tumor-distant “healthy” control lung tissue by magnetic-activated cell sorting (MACS) using the AT-II marker HT-II-280 and the AT-I marker HT-I-56, respectively ^{[13, 17, 27, 33]}. All steps were performed on ice or at 4°C unless otherwise specified and all buffers and consumables were pre-cooled.

Peripheral lung tissue was dissected free of pleura, bronchi, and large vessels, mechanically minced, and enzymatically digested (Miltenyi Biotec, gentleMACS and No. 130-110-201 Multi Tissue Dissociation Kit 1) [Miltenyi Biotec, Cat. No. 130-110-201]). After sequential filtration (100/70 µm), centrifugation (450 g, 7 min, 4°C), and erythrocyte lysis with ACK lysis buffer (Thermo Fisher Scientific/Gibco, Cat. No. A10492-01) followed by sequential filtration (40 µm), cells were washed and resuspended in MACS buffer (1% BSA, 2 mM EDTA in PBS).

Following Fc receptor blockade, (10 µL/sample; Miltenyi Biotec) for 15 min at 4°C, cells were incubated with 5 µL mouse IgM anti-HT-II-280 antibody (Terrace Biotech, Cat. No. TB-27AHT2-280; 1:50 in MACS buffer) for 1 h at 4°C with gentle rotation (60 rpm). For AT-I isolation, the primary antibody was replaced by mouse IgG anti-HT-I-56 (Terrace Biotech, Cat. No. TB-29AHT1-56) at the same dilution.

Cells were washed with 1.5 mL MACS buffer, centrifuged, and resuspended in 2 mL MACS buffer containing anti-mouse IgM or anti-mouse IgG MicroBeads (Miltenyi Biotec; 20 µL beads per 10⁷ cells, diluted 1:10 in MACS buffer). Bead incubation was carried out for 30 min at 4°C with gentle rotation. After washing, suspensions were applied to pre-equilibrated LS columns in an OctoMACS separator. Negative fractions were collected as flow-through, while positive fractions were eluted after column removal from the magnet. Both fractions were pelleted, resuspended in MACS buffer, and used for counting or downstream applications. MACS was selected over FACS to reduce shear stress and preserve viability for organoid culture.

Both positive and negative fractions were pelleted (450 g, 7 min, 4°C) and resuspended in 200 µL MACS buffer for counting or immediate downstream applications.

MACS-isolated HT-II-280⁺ AT-II cells were pelleted (450 g, 7 min, 4°C) and resuspended in serum-free, feeder-free (SFFF) medium based on Advanced DMEM/F-12 (Gibco, Cat 12634028) supplemented with a variety of proliferation enhancers e.g. GSK3β-Inhibitor CHIR99021 and AT-I cell differentiation inhibitors such as ROCK-Inhibitor Y-27632, TGF-β-Receptor I (ALK5)-Inhibitor SB431542 and p38 MAPK-Inhibitor BIRB796 (see Supplementary Table S1). SFFF medium was adapted from Konishi et al. ^[33] using the same components but with modified concentrations to optimize maintenance of AT-II cell phenotype and increased proliferation. Cells in SFFF medium were mixed with Corning® Matrigel® Growth Factor Reduced Basement Membrane Matrix (Cat. No. 356231) at a 5:8 ratio (cell suspension:matrigel) and seeded as 100–130 µL domes in 24-well plates (60,000 cells/dome) in initial passage (P0). After polymerization (20–30 min, 37°C), 500 µL SFFF medium was added per well. For the first 4–5 days, IL-1β was added to enhance alveolosphere formation. Culture medium also contained EGF and FGF-10 to support Wnt and ERK/AKT pathway activation, essential for AT-II expansion. Organoids were grown for up to 16 days and passaged up to P3, with medium exchanged every 2–3 days.

To generate single-cell suspensions for passaging of AT-II cells, organoids were segregated using TrypLE™ Select Enzyme (Gibco, Cat. No. 12563011). Per matrigel dome, 500 µL TrypLE™ Select Enzyme were utilized to firstly mechanically disrupt domes and incubate them at 37°C for 15–30 min with regard to organoid sizes and numbers. Suspensions were again mechanically disrupted, washed 3x in PBS, and centrifuged at 500 g for 5 min before further processing.

Technical note: For all histological investigations and especially for flow cytometry labeling, we strongly recommend the utilization of Cell Recovery Solution (Corning, Cat. No.: 354253) before further processing. 300 µL Cell Recovery Solution were added to the domes to dissolve organoids from matrigel for more surface efficient stainings (e.g. such as for HT-II-280 labeling) and minimization on unlabeled cells in FACS to to residual matrigel. Organoids free of matrigel can further undergo immediate trypsination.

For cryopreservation, organoids were mechanically and enzymatically dissociated as described above. Up to 500,000 single cells were resuspended in 1 ml cryomedium (70% FBS, 20% SFFF medium, 10% DMSO, added dropwise). Cryomedium was incubated with AT-II for 20min at RT to allow for proper diffusion of DMSO. Cryovials were placed in a Mr. Frosty™ container at − 80°C and transferred after 24 h to − 150°C for long-term storage. To assess safety and viability, samples were stored ≥ 28 days at − 150°C. For recovery, cells were thawed in a 37°C water bath for 2 min, diluted in 14 ml Advanced DMEM/F-12 at room temperature, incubated for 10 min, centrifuged (7 min, 450 g), and directly reseeded in Matrigel at a 5:8 ratio.

To confirm the identity and purity of alveolar organoids over multiple passages and to evaluate cryostorage potential, a multitude of techniques was applied including conventional histology, microcomputed tomography, transmission electron microscopy, image analysis, immunofluorescence & flow cytometry as well as viability assays. For the sake of space, these techniques are described in detail in the supplement.

To assess the metabolic properties of fibrotic vs control organoids, glycolytic capacity and ATP production of fibrotic and healthy tissue-derived AT-II cells measured by extracellular flux assays using the Agilent Seahorse XF HS Mini Analyzer S7852A platform (Agilent, Santa Clara, CA). Organoids from P0-P1 were separated to single-cell suspension using trypsination as previously described in method section. Next, 12,000 cells from P1-P2 were plated 10–12 d prior to experiments in each well of XFp PDL Miniplates (Agilent, Cat. No.: 03722 − 100) in matrigel domes with 50 µl total volume as suggested by Ludikhuize et al. ^[34] and provided with SFFF media. By day of experiment, each well therefore roughly contains 85,000-120,000 cells as organoids. Samples were prepared in duplets. Cartridges were prepared corresponding to the manufacturer’s instructions for Seahorse XFp Glycolysis Stress Test Kit (Agilent, Cat. No.: 103017-100) and Seahorse XFp Real-Time ATP Rate Assay Kit (Agilent, Cat. No.: 103591-100). On the day of experiment, assay media were freshly prepared. SFFF was aspirated from matrigel domes exhibiting normal sized alveolospheres after cultivation miniplates and exchanged with 100 µl assay media 45 min prior to measurement, resulting in a total volume of 150 µl per well. Of note, after serial titration experiments for Seahorse XFp Glycolysis Stress Test Kit, we increased the concentrations of glucose to 15 mM, oligomycin to 5 µM and 2-DG to 200 mM per well, each resuspended in assay medium. Each injection port contained 20 µl volume. We measured 3 cycles each basal curve, glycolysis, and glycolytic capacity followed by 5 cycles of glycolytic reserve asessement. Similarly, for Seahorse XFp Real-Time ATP Rate Assay Kit, oligomycin (5µM) and Retenone + Antimycin A (2µM) per well were utilized in higher concentration than recommended from Agilent for measurement of 2D cell culture and with a volume of 20 µl per injection port. These concentrations are roughly in line with concentration utilized to measure extracellular flux of intestinal organoids by Ludikhuize et al. ^[34]. ATP production were assessed by 3 cycles each basal ECAR and OCR, inhibition of mitochondrial ATP synthesis, and, finally, mitochondrial-associated acidification. Following measurements, organoids were collected from matrigel using Cell Recovery Solution (Corning, Cat. No.: 354253) and harvested for DNA isolation using QIAamp DNA micro kit (Qiagen, Cat. No.:56304) and samples were measured by Nanodrop to enable a normalization of experimental values relative to approximated number of cells. Additionally, BrdU assays (Cell Signaling Technology, Cat. No.: 6813S) were performed to confirm unaltered cell proliferation and viability post-extracellular flux assays (data not shown).

In this study, we established a robust, cell-line-free, patient-derived 3D lung organoid model generated from primary human alveolar type II (AT-II) epithelial cells isolated from fibrotic and non-fibrotic lung tissues. Using HT-II-280-based magnetic bead sorting, adapted from Konishi et al.^[33], we achieved high-purity AT-II cell populations that were expanded in serum- and feeder-free conditions to form alveolospheres. These organoids recapitulated key AT-II features—proSP-C production, HT-II-280 localization, lamellar body formation—and preserved cell identity across passages. Compared to existing in vitro systems relying on iPSCs, immortalized lines, or murine cells ^{[26, 28, 40]}, our approach provides lineage fidelity and patient-specific disease signatures.

A major strength of our model is the high cellular purity and phenotypic stability. More than 80% of cells within alveolospheres remained HT-II-280⁺ or proSP-C⁺ across 3D-culture passaging, consistent with human AT-II organoid data ^[30]. Although HT-II-280–based magnetic sorting provided a highly enriched AT-II population prior to culturing, purity was not absolute. Immunofluorescence of cytospins revealed ~ 89% HT-II-280⁺ cells, while flow cytometry directly following sorting also detected strong enrichment of HT-II-280⁺ cells, however indicating that a small fraction of non–AT-II cells remained after isolation. These observations are consistent with the moderate colony formation rate (CFR) observed in the initial passage (P0), whereas both CFR and AT-II marker-specific labeling in flow cytometry notably increased in P1, suggesting that clonal expansion under serum- and feeder-free (SFFF) conditions—supplemented with proliferative boosters and AT-I inhibitors—further selects for bona fide AT-II progenitors during early culture. Ultrastructural analysis of organoids revealed abundant lamellar bodies, confirming mature AT-II identity ^{[16, 17]}. Thus, the combination of magnetic enrichment and medium-driven selection in adapted matrigel-culturing likely achieves a higher functional purity than sorting alone. With our chosen ratio of matrigel to SFFF for initital dome pouring, we in addition overcome mechanical influences on epithelial fate: Matrix stiffness and mechanical cues are key regulators of alveolar epithelial differentiation in fibrotic lung tissue. It has been shown that increased rigidity of the extracellular matrix can induce AT-II cell dedifferentiation and emergence of aberrant basaloid states, as also observed in vivo in fibrotic regions of IPF ^{[6, 8]}. Our in vitro findings mirror this behavior, as direct contact of organoids with the rigid plastic surface triggered loss of AT-II morphology and basaloid-like transformation. Increasing the Matrigel content to a 5:8 ratio prevented this effect by maintaining a compliant, fully embedded 3D environment. This adjustment thus represents a simple but essential refinement that preserves epithelial identity and enhances reproducibility in long-term culture of primary human AT-II organoids. Furthermore, differentiation into AT-I occurred solely modestly in later passages, with emergence of HT-I-56+/proSP-C + intermediates, indicating retained progenitor potential as described in other alveolar regeneration models ^[41–42]. Importantly, these results were reproducible across a large number of patient samples, supporting the transferability of this platform as a reliable technical resource.

We also observed morphological heterogeneity of alveolospheres, which could be categorized as lumen-containing (type A) or dense lobular (type B), similar to murine AT-II-derived spheroids ^[18] and alveolar epithelial progenitor-derived structures ^[14]. Despite using highly enriched HT-II-280⁺ ATII cells, we occasionally observed airway-like organoids in addition to alveolospheres. This may reflect inclusion of epithelial progenitors with dual potential ^[18], the influence of tissue origin (distal versus bronchiolar regions), and culture conditions that permit airway cell outgrowth, e.g. the addition of FGF10, consistent with previous reports on human airway organoids ^[29].and HT-II-280⁺-derived cultures ^[27]. Rather than a limitation, such morphological heterogeneity reflects the target tissue and patient-specific biology and increases the translational relevance of the model for personalized medicine approaches.

An important advance is the demonstration that organoids retained structural integrity and viability after cryopreservation. Cryostorage and successful revitalization were reproducible across both fibrotic and control tissues, demonstrating suitability for standardized biobanking and inter-center sharing. Following storage at − 150°C, we observed no loss of organoid-forming efficiency, viability, or metabolic activity, supporting the feasibility of biobanking. To our knowledge, this is among the first reports of successful cryorecovery of primary human AT-II organoids while preserving epithelial identity and functionality. This reproducibility across both control and fibrotic samples underscores the scalability of the approach and highlights its potential for inter-center biobanking, harmonized disease modeling, and standardized drug testing pipelines.

Most notably, we identified a pronounced consistent metabolic shift in fibrotic AT-II organoids—especially those from IPF—towards aerobic glycolysis, establishing the organoids as a reproducible human platform to capture disease-specific energetic reprogramming. While extracellular flux assays have been used in murine epithelial organoids ^[43], their application to primary human AT-II organoids from ILD patients is novel. By adapting extracellular flux analysis protocols from intestinal organoid studies ^[34], we showed that IPF-derived alveolospheres exhibited elevated basal ECAR, enhanced compensatory glycolysis, and increased glycolytic ATP production. This recapitulates the “Warburg-like” metabolic phenotype described in IPF epithelial cells, driven by upregulated LDHA and PDK1, lactate accumulation, and reduced mitochondrial pyruvate dehydrogenase activity ^{[24, 35–36]}. Because this phenotype was consistently detected across independent donors, our study provides a reproducible framework to interrogate metabolic reprogramming in patient-derived lung epithelium.

The uncoupling of glycolysis from oxidative phosphorylation suggests disease-associated mitochondrial dysfunction and adaptation to high energetic demands of proliferation and ECM remodeling ^{[26, 37]}. Sun et al.^[25] reported that PDK1-driven lactate accumulation suppresses mitochondrial pyruvate dehydrogenase, reinforcing glycolytic flux. Metabolic studies in IPF have also shown decreased late glycolytic intermediates and elevated lactate ^[37]. Such metabolic rewiring is implicated in epithelial-to-mesenchymal transition (EMT), apoptosis resistance, and profibrotic signaling. The PDK1–HIF-1α axis promotes glycolysis-dependent myofibroblast differentiation under hypoxic/inflammatory stress ^[38]. Recent organoid-based work by Choi et al. ^[39] further demonstrated decreased regenerative capacity of IPF-derived AT-II cells with disease progression, highlighting the clinical significance of metabolic dysfunction.

As a novel cellular model, some limitations must be noted. First, the model relies on end-stage explants, limiting insights into early pathogenic events ^[5]. Second, patient heterogeneity, despite normalization, introduces variability in sorting efficiency and expansion during cultivation. Third, while canonical markers validated AT-II identity, transcriptomic profiling is needed to capture sublineage states ^{[41, 44]}. Incorporating stromal or immune components via co-culture or organoid-on-chip approaches could improve physiological relevance and enable modeling of epithelial–mesenchymal and epithelial–immune interactions ^{[31, 45–46]}. Furthermore, possible transdifferentiation of AT-II cells into KRT5⁺ basal cells under profibrotic mesenchymal or TGF-β signals ^[32] underscores the need to explore epithelial plasticity in IPF organoids.

By allowing direct scalable and reproducible comparison of fibrotic versus control AT-II organoids under patient-matched conditions, our model offers a platform to interrogate epithelial dysfunction and therapeutic responses for mechanistic studies, multicenter validation, and translational drug screening.

Together, our findings establish a reproducible, feeder-free 3D organoid model of primary human AT-II cells that preserves lineage identity and captures fibrotic metabolic reprogramming. These organoids retained ultrastructural identity and remained viable after cryopreservation, enabling long-term storage and recovery. Importantly, fibrotic organoids consistently displayed a glycolytic shift, reflecting disease-specific metabolic reprogramming. By combining reproducibility, scalability, and compatibility with cryopreservation, this model provides a transferable resource for biobanking, multicenter validation, and translational drug discovery. Beyond serving as a disease model, our platform establishes a technical foundation for future precision medicine approaches in interstitial lung diseases.