Cell isolation procedures followed recently described protocols (Menzel et al., 2017; Kreimendahl et al., 2019; Kniebs et al., 2020; Kniebs et al., 2021; Schuhenn et al., 2022). For information on ethical approval and relevant guidelines, please refer to the declarations section.

Briefly, HRE were obtained from lung transplant donors post mortem at the University Medical Center Essen, Germany. Selection criteria for donors are listed in the Eurotransplant guidelines. Cells were isolated and cultivated following a well-established, verified and commonly used method (Mertens et al., 2017). They were expanded in keratinocyte-SF medium (KSFM, Gibco) supplemented with human EGF (Gibco, 2.5 ng/ml), bovine pituitary extract (BPE, Gibco, 25 μg/ml), isoproterenol (Sigma-Aldrich, 1 μM) and a mixture of Penicillin, Streptomycin, Ciprofloxacin and Amphotericin B (PanBiotech, 2.5 μg/ml). Afterwards, cells were frozen in liquid nitrogen using a mixture of 90% KSFM, 10% dimethyl sulfoxide (DMSO, Sigma) and BPE (0.3 mg/ml) until further use. To verify cell identities of isolated HRE, they are screened regularly by immunohistochemical staining for basal cell marker P40/chromogranin A. Furthermore, HRE were verified by their ability to differentiate in monoculture under ALI conditions into ciliated cells and mucus producing goblet cells (see Supplementary Figure S1). HUVEC were isolated from human umbilical cords provided by the Clinic for Gynecology and Obstetrics (RWTH Aachen University Hospital) and expanded in endothelial cell growth medium 2 (EGM2, Promocell) up to passage 3.

BM-MSC were isolated from femoral heads supplied by the Clinic for Orthopedic, Trauma and Reconstructive Surgery (RWTH Aachen University Hospital) and cultured in Mesenpan medium (PAN-Biotech) supplemented with 2% FCS (Gibco) and 1% antibiotic/antimycotic solution (ABM, Gibco) up to passage 4.

ASC were isolated from abdominal fat tissue obtained from the Clinic for Plastic, Hand and Burns Surgery (RWTH Aachen University Hospital) and expanded in supplemented Mesenpan medium up to passage 3.

HNF were isolated from nasal conchae received from the Clinic for Otorhinolaryngology (RWTH Aachen University Hospital) and cultured in Dulbecco`s Modified Eagle Medium (DMEM, Gibco) supplemented with 10% FCS and 1% ABM up to passage 3.

Cell identities of HUVEC, BM-MSC, ASC and HNF were verified in previous studies using the same isolation protocols (Weinandy et al., 2014; Kreimendahl et al., 2019; Kniebs et al., 2020; Thiebes et al., 2021). Additionally, isolation procedures for MSC are regularly verified by flow cytometry according to the minimal criteria for the classification of MSC given by Dominici et al. (2006): ASC and BM-MSC gave positive signals for CD90, CD73 and CD105 with ≥97.2%. The markers CD45, CD34, CD11b, CD79α and HLA-DR surface molecules were found to be negative (<1.3%).

Experiments were conducted with HRE in passage 1, with HUVEC, ASC and HNF in passage 4 and with BM-MSC in passage 5. BM-MSC, ASC and HNF were tested using three different donors per cell type while for HUVEC and HRE, one donor was tested per cell type leading to three repetitions per culture and supporting cell type. As high cell numbers are needed for three-dimensional cell culture and the amount of primary cells that can be isolated from fresh tissue is limited, only the supporting cell types were tested with regard to donor variation in order to evaluate their influence on the differentiation. The HUVEC donor and the HRE donor were kept constant to avoid a donor-related impact on the results not originating from the supporting cell types. Especially with regard to HRE, donor variability can be high and availability of cells derived from healthy donors is limited. In tri-culture experiments, HUVEC were mixed with either BM-MSC, ASC or HNF in a fibrin gel before HRE were seeded on top of the gel. In co-culture experiments, either HUVEC were combined with one supporting cell type (BM-MSC, ASC or HNF) in a fibrin gel or HRE were seeded on top of a gel containing one of the supporting cell types. Additionally, a co-culture of HUVEC and HRE as well as monocultures of HUVEC and HRE were prepared as controls. Fibrin gels were molded according to previously published protocols (Kniebs et al., 2020; Kreimendahl et al., 2021). Briefly, HUVEC and/or one of the supporting cell types were seeded with a final concentration of 3 ⋅ 10 6  cells/mL each in a fibrin gel containing 5 mg/ml fibrinogen (VWR), 3 U/mL thrombin (Sigma-Aldrich) and 3.75 mM CaCl₂. Gels were either molded in 24-well plates (VWR) for co-cultures with HUVEC and monocultures (350 µL) or in Transwell^® Inserts (PET membrane, 0.4 µm pore size, Corning^®) combined with ThinCert™Plates (Greiner Bio One) when used for ALI (206 µl). Polymerization of the gels was achieved by incubation at room temperature for 20 min followed by incubation at 37°C and 5% CO₂ in a humidified atmosphere for another 20 min. In case of co-cultures with HRE and tri-cultures, HRE were then seeded on top of the gels with a final density of 80,000 cells/cm². For the HRE monoculture, cells were seeded onto Transwell inserts coated with collagen IV from human placenta (12 μg/cm², Sigma-Aldrich). Detailed media composition was described previously (Kreimendahl et al., 2019; Luengen et al., 2020). In short, cultures with HRE except the monoculture were proliferated under submerged conditions in a 1:1-mixture of EGM2 and airway epithelial cell growth medium (AECGM, Promocell) for 1 week before switching to ALI-conditions in a 1:1-mixture (EMM) of EGM2 and MucilAir medium (Epithelix) for 4 weeks. HRE monoculture was performed using AECGM in the proliferation phase and a modified version of AECGM in the differentiation phase (mAir). Co-cultures with HUVEC as well as monocultures of BM-MSC, ASC and HNF were cultured in EMM medium for 2 weeks. In all cultures, medium was exchanged every 2–3 days. All media were supplemented with 0.05%–0.1% Gentamicin (Rotexmedica) and 0.16% tranexamic acid (Carinopharm) to avoid gel degradation.

Cell differentiation was analyzed by scanning electron microscopy (SEM), two-photon scanning electron microscopy (TPLSM) and histologic and immunohistochemical stainings. To evaluate ciliation of HRE after 4 weeks of ALI, SEM analysis was conducted by the facility for electron microscopy of the medical faculty of RWTH Aachen University. After fixation in 3% glutaraldehyde (Agar scientific), samples were rinsed with 0.1 M sodium phosphate buffer (Merck) followed by dehydration in an ascending ethanol series. Subsequent to critical point drying in liquid CO₂, samples were coated with a 10 nm gold/palladium layer (Sputter Coater EM SCD500, Leica). SEM was carried out in a high vacuum environment at 10 kV acceleration voltage with an environmental scanning electron microscope (ESEM XL30 FEG, FEI).

Vascularization of the samples was analyzed using TPLSM after previously established protocols (Kreimendahl et al., 2019; Kniebs et al., 2020). After fixation in ice-cold methanol, samples were first stained with mouse anti-CD31 (PECAM-1, 1:100; Sigma-Aldrich) and Alexa Fluor^® 594 goat anti-mouse IgG (1:400, Invitrogen). A selection of tri-culture samples was further stained sequentially using mouse anti-vimentin (1:100; Sigma-Aldrich), Alexa Fluor^® 488 goat anti-mouse IgG (1:400, Invitrogen) and DAPI (Molecular Probes). The TPLSM was performed using a two-photon laser scanning microscope (Olympus FluoView 1000 MPE) with a ×25 water-objective NA 1.05 (Olympus Optical) and a MaiTai Deep-See Titan-Sapphire-laser. FluoView FV 4.2 Software was used for the visualization while images were processed and evaluated with Imaris 9.6.0 software (Bitplane Inc. South Winsor). Vascularization was quantified by evaluating the average volume, surface area, length and number of branching points of vascular-like structures. Three images per gel were taken for the statistical result evaluation.

For histologic and immunohistochemical evaluation, samples were embedded in 2% agarose, fixed in Carnoy’s solution and dehydrated in an ascending ethanol series followed by incubation in xylene and embedding in paraffin. Native human nasal concha tissue was fixed in 4% paraformaldehyde solution before dehydration and paraffin embedding as described above. Afterwards, 5 µm-thick deparaffinized sections were stained by periodic acid Schiff’s reaction (PAS reaction) to analyze epithelial cell differentiation using a bright field light microscope. Briefly, deparaffinized samples were first hydrolyzed in 1% w/v periodic acid solution before staining with Schiff’s reagent (both Merck). After a washing step in 35°C warm tap water, cell nuclei were counterstained with Mayer’s hematoxylin (Sigma-Aldrich). Immunohistochemical analysis was also performed on Carnoy-fixed sections after a previously described protocol (Luengen et al., 2020). After deparaffinization, samples were blocked with 5% normal goat serum (NGS, Dako). Following incubation in primary antibody solutions overnight at 4°C and three washing steps, sections were incubated protected from light in secondary antibody solutions for 1 h at 37°C. Primary antibodies included rabbit anti-pan-cytokeratin (1:200, Acris), mouse anti-mucin-5AC (1:800, Acris), rabbit anti-claudin-1 (1:800, Biorbyt), mouse anti-acetylated tubulin (1:800, Sigma-Aldrich) and mouse anti-CD31 (PECAM-1, 1:100; Sigma-Aldrich). Fluorescently labelled secondary antibodies Alexa Fluor 488 goat anti-rabbit IgG and Alexa Fluor 594 goat anti-mouse IgG (both Invitrogen) were used for detection. All antibodies were diluted in PBS containing 1% w/v bovine serum albumin (BSA, Sigma-Aldrich) and 0.1% w/v sodium azide (Sigma-Aldrich). Counterstaining of cell nuclei was carried out using DAPI. For observation, an inverted fluorescence microscope (Axio Observer. Z1, Carl Zeiss) was used.

To evaluate differences in signaling patterns between cultures, a LEGENDPlex™ bead-based immunoassay (BioLegend) for eight human cytokines including vascular endothelial growth factor (VEGF), epidermal growth factor (EGF), interleukin-6 (IL6), interleukin-8 (IL8), angiopoietin 1, angiopoietin 2, fibroblast growth factor basic (FGF-b), placenta growth factor (PIGF), interleukin-33 (IL33), thymic stromal lymphopoietin (TSLP) and transforming growth factor-beta1 (TGF-β1) was performed on media supernatant at the end of the ALI culture following the manufacturer’s instructions. Results were evaluated according to the manufacturers’ instructions.

To compare the ciliation amount between different cultures, ten independent and in terms of mucociliary differentiation well educated observers classified SEM images blinded for the condition as previously described (Luengen et al., 2020). The scoring system provided in Table 1 was used for classification of the proportion of the ciliated area in relation to the total surface area in one general overview image of the sample. Afterwards, mean values per image and for each condition were calculated.

Statistical analysis was performed using GraphPad Prism software 9.2.0. For vascularization and ciliation quantification, statistics were performed by one-way analysis of variance (ANOVA) with Tukey’s post-hoc test. Standard deviation of ciliation scores was determined by Gaussian error propagation. Cytokine production results were analyzed by two-way ANOVA with Tukey’s post-hoc test. A p-value of p < 0.05 was considered as statistically significant.

Cell morphology, presence of goblet cells and mucus production in co- and tri-cultures were analyzed using PAS reaction (Figure 2). All cultures were established with three different supporting cell donors, variability between donors was found to be low. HRE-co-cultures and tri-cultures with BM-MSC revealed a characteristic and pseudostratified epithelial layer. Mucus-secreting goblet cells and basal cell morphology resembled the native respiratory epithelium (Figures 2A,B, M, N). Both HRE-co-cultures and tri-cultures with ASC showed a multilayered morphology. Mucus production confirmed in the HRE-co-cultures with ASC was mostly absent in the corresponding tri-cultures (Figures 2C,D, O, P). In addition, co-cultures with HRE and ASC lacked the clear morphology of a basal cell layer in terms of conventional histology which was observed in cultures with BM-MSC or in ASC-tri-cultures. Formation of mucus was reduced in tri-cultures with HNF in comparison to the corresponding co-cultures with HRE. However, both had a thin multilayered and rather non-pseudostratified epithelium (Figures 2E,F, Q, R). Co-cultures with HUVEC and different supporting cell types exhibited a comparable morphology (Figure 2G–L). PAS reaction images of all donors per condition as well as of native human nasal concha tissue as positive control, a co-culture of HRE and HUVEC as well as of a monoculture of HRE on inserts are shown in the supplementary (see Supplementary Figures S1–S4).

To confirm mucociliary differentiation and formation of vascular-like structures, immunohistochemical stainings of endothelial cell marker CD31 and of characteristic epithelial cell markers pan-cytokeratin, mucin5AC, claudin-1 and α-tubulin were performed (Figure 3). Pan-cytokeratin acts as marker for the epithelial cytoskeleton, mucin5AC reveals mucus production, claudin-1 indicates barrier formation and α-tubulin marks cilia while CD31 confirms presence of endothelial cells (Kreimendahl et al., 2019). In all cultures with HRE containing BM-MSC (Figures 3A,B,G,H, M, N, S, T), staining for the four epithelial cell markers and CD31 was positive verifying the epithelial origin of the cells as well as the presence of cilia, mucus, tight junctions and vascular-like structures comparable to native human nasal tissue as shown in the supplementary (see Supplementary Figure S5). In BM-MSC-tri-cultures, claudin-1 and pan-cytokeratin expression was additionally found to be localized in the basal cell layer only as was also observed in the native control. In co-cultures with BM-MSC, these markers were detected throughout the whole epithelial cell layer. Some unexpected α-tubulin staining was observed in BM-MSC inside gels of BM-MSC tri-cultures and co-cultures with HRE (Figures 3G,H) (Zhou et al., 2017). While all HRE-cultures containing ASC were found to be positive for pan-cytokeratin and claudin-1, cilia and mucus were only detected in co-cultures with HRE (Figures 3C,D,I,J, P, U, V). ASC-cultures containing HUVEC were stained positively for CD31 (Figures 3O, P). Moreover, claudin-1 and pan-cytokeratin expression was detected in the whole epithelial cell layer lacking the localization of these cell markers in the basal cell layer found in the native control (see Supplementary Figure S5). Although the epithelial origin of the cells could be shown by the stainings, HRE in tri-cultures with ASC failed to develop a mucociliary phenotype (Figures 3D,J,V). Cultures containing HNF were again positive for all analyzed markers although epithelial cell layers were again found to be thinner than in cultures with MSC (Figures 3E,F,K, L, Q, R, W, X). Furthermore, the characteristic distribution pattern of claudin-1 and pan-cytokeratin was also absent in all HNF-cultures which instead show expression throughout the whole epithelium. Regarding CD31 staining in co-cultures of HUVEC with supporting cell types, expression was found to be reduced in cultures with ASC and HNF in comparison with BM-MSC (Figure 3M, O, Q). Controls with native tissue, HRE and HUVEC in monoculture as well as in co-culture with each other are depicted in the supplementary (see Supplementary Figure S5).

Cilia formation was additionally analyzed by SEM and could be confirmed for all cultures (Figure 4). In comparison with the HRE-co-cultures (Figures 4A,B), BM-MSC-containing tri-cultures (Figures 4G,H) were found to be slightly less ciliated. The same effect could be observed for cultures with ASC (Figures 4C,D,I,J), which additionally exhibited a lower level of ciliation in total than the cultures with BM-MSC. Cilia formation in cultures with HNF seemed to be comparable between co- and tri-cultures (Figures 4E,F,K, L).

Estimation of the differences in ciliation amounts between different culture conditions by independent observers based on SEM images revealed significant differences between almost all groups (Figure 5): only the tri-culture with BM-MSC (mean score 2.97 ± 0.10) was found to be non-significantly different from the HNF-co-culture (mean score 3.17 ± 0.11) and HNF-tri-culture (mean score 2.77 ± 0.19). The latter was additionally observed to be non-significantly different from the monoculture of HRE (mean score 2.5 ± 0.10) that was evaluated as control group. The highest score was estimated for the co-culture with BM-MSC (mean score 3.80 ± 0.07) while the lowest scores were observed for the co-culture (mean score 1.10 ± 0.08) and the tri-culture (mean score 0.30 ± 0.06) with ASC.

The influence of different supporting cell types in co-cultures with HUVEC and in tri-cultures on formation of vascular-like structures was evaluated using TPLSM (Figure 6), followed by statistical analysis (Figure 7). In all cultures, elongated and branched vessel-like structures could be observed after staining with anti-CD31-antibody (Figure 6). However, tri-cultures with BM-MSC seemed to develop a larger number of thin structures that in addition appeared to be longer and more homogeneously outspread through the gel than the structures observed in other cultures (Figure 6D). While vascularization results were comparable in all co-cultures with HUVEC and in the tri-culture with HNF (Figures 6A–C,F), it seemed to be reduced in tri-cultures with ASC compared to the corresponding co-culture (Figure 6E). A selection of tri-culture samples was further stained with anti-vimentin-antibody to visualize the supporting cell type in the gel (Figures 6G–I). BM-MSC were found to be elongated and more concentrated on one side of the gel (Figure 6G) whereas ASC and HNF seemed to be smaller, not as spindle-shaped and to be more evenly distributed in the gel (Figures 6H,I). However, the observed cell distribution is caused by method-related effects of the TPLSM due to an uneven sample surface and thus not meaningful.

Statistical analysis of the TPLSM images showed significantly increased structure length and area for tri-cultures with BM-MSC in comparison to co-cultures with ASC or HNF (Figures 7A,C). In addition, the number of branching points was significantly higher in these tri-cultures in comparison to all other groups (Figure 7D). Differences observed in the volume of vessel-like structures were not found to be statistically significant (Figure 7B). With regard to functionality of formed vascular-like structures, we could prove lumen formation independent on the supporting cell type, the corresponding images are shown in the supplementary (see Supplementary Figure S6).

In order to investigate differences in cytokine production between the cell types, media supernatants were analyzed for levels of eight cytokines involved in vascularization processes and inflammation by LEGENDPlex bead-based immunoassay (Figure 8). Cytokine concentrations were normalized to cell-free media. Statistically significant differences could be observed for angiopoietin 2 (Figure 8D) and VEGF (Figure 8F). The HUVEC-co-culture with BM-MSC showed significantly increased levels of angiopoietin 2 (94,513.38 ± 4,400.76 pg/ml) in comparison to all other groups. HUVEC-co-cultures with ASC (41,359.83 ± 9,893.24 pg/ml) and HNF (37,400.70 ± 30,372.09 pg/ml) each also revealed to have significantly increased levels in angiopoietin 2 in comparison to tri-cultures and co-cultures with HRE (Figure 8D).

HUVEC-co-cultures with ASC or HNF in addition exhibited significantly higher levels of VEGF (28,160.08 ± 14,049.48 pg/ml resp. 74,956.75 ± 54,838.28 pg/ml) when compared to all other groups (Figure 8F). Differences in cytokine levels of IL6, IL8, angiopoietin 1, EGF, FGF-b and PIGF (Figures 8A–C,E, G-H) were found to be statistically non-significant. Nevertheless, trends can be observed: HUVEC-co-cultures with BM-MSC seemed to have increased levels in IL6, IL8 and angiopoietin 1 when compared to the other groups. The latter was also found to be high in the two other HUVEC-co-cultures whereas only the HUVEC-co-culture with ASC seemed to have increased IL8 levels. IL6 concentrations in HRE-co-cultures with ASC or HNF and in tri-cultures with HNF were close to the detection limit (Figures 8A–C). With exception of the HUVEC-co-culture with ASC, all cultures consumed EGF (Figure 8E) while FGF-b was consumed by all cultures without exception (Figure 8G). High levels of PIGF were found for all tri-cultures and co-cultures with HUVEC while concentrations in HRE-co-cultures with MSC were close to the detection limit (Figure 8H). However, these observations have to be described as numerical differences without statistical significance demonstrated in this study. Cytokine levels in media supernatants of monocultures with BM-MSC, ASC or HNF in fibrin gel are given in the supplementary (see Supplementary Figure S7). IL6 concentration was found to be significantly higher in BM-MSC-monocultures while angiopoietin 1 level was significantly increased in monocultures of HNF. In addition, concentrations of pro-inflammatory cytokines IL33, TSLP and TGF-β1 in media supernatants are given in the supplementary (see Supplementary Figure S8). While levels of IL33 and TSLP were found to be very low in all cultures, TGF-β1 could be detected to some degree in co-cultures with HUVEC as well as in monocultures of supporting cell types.

In summary, using BM-MSC as supporting cell type in tri-cultures led to well differentiated respiratory epithelium and a high degree of vascularization. HNF-supported tri-cultures also showed mucociliary differentiation and formation of vascular-like structures although to a lesser degree than with the BM-MSC. HRE in tri-cultures containing ASC were mostly undifferentiated while differences in vascularization were marginal. Co- and tri-cultures containing BM-MSC or HNF contained a basal cell layer positive for pan-cytokeratin, ciliated cells, mucus-producing goblet cells, tight junctions and epithelial cytoskeleton comparable to native tissue. Although ciliation amount in HNF-tri-cultures was comparable to tri-cultures with BM-MSC, and HNF-tri-cultures were also found to be positive for epithelial cell marker expression, epithelial morphology observed in PAS reaction and immunohistochemistry more closely resembled the native state in BM-MSC-tri-cultures with regard to cell arrangement, cell marker localization and layer thickness. Moreover, vascularization in tri-cultures with BM-MSC associated with a significantly higher number of branching points compared to all other cultures. Although differences in area and length of the vascular-like structures were only found to be statistically significant for BM-MSC-tri-cultures in comparison to co-cultures with ASC or HNF, an overall trend towards higher values in BM-MSC-tri-cultures was observed.

Interestingly, tri-culture reduced ciliation in comparison to the corresponding co-culture. The presence of a third cell type in the tri-culture system resulting in higher consumption of nutrients and growth factors might be a possible reason. Increasing the media exchange frequency or extension of the ALI period could therefore be strategies to increase ciliation in tri-cultures. However, this was not tested in this study in order to keep all parameters constant. In contrast, this effect was not generally discovered for vascularization. Based on the mucociliary differentiation observed in the co-culture of HUVEC and HRE (see Supplementary Figure S1 and Supplementary Figure S5), a negative influence of HUVEC on the ciliation is not indicated. Positive α-tubulin staining inside BM-MSC found in BM-MSC-tri- and co-cultures can be explained by detection of intracellular microtubuli as part of the cytoskeleton, since α-tubulin acts as marker for microtubuli not only in cilia (Zhou et al., 2017).

Most studies dealing with airway epithelial co-cultures use fibroblasts as supporting cell type (Hiemstra et al., 2019). Their ability to promote epithelial cell differentiation has been confirmed in several studies (Albers et al., 2015; Abs et al., 2019). In our first study that established the tri-culture system based on HNEC, we therefore used HNF as supporting cell type. Nevertheless, access to fibroblasts from respiratory origin might be limited for tissue engineering applications as it requires painful surgery and can be associated with high donor variability depending on the patient´s medical history (Law et al., 2020; Luengen et al., 2020). MSC provide high potential for autologous cell use as they can be obtained in high numbers through rather minimal invasive procedures (Mushahary et al., 2018). Furthermore, Taşkiran et al., who compared differences in molecular signatures of BM-MSC and dermal fibroblasts, recommend the use of BM-MSC for developmental and differentiation investigations (Taşkiran and Karaosmanoğlu, 2019). Although MSC and fibroblasts share features in tissue remodeling, angiogenesis and wound healing, fibroblasts bear the risk of hyperproliferation and fibrosis while MSC are known to have anti-fibrotic properties (Zupan, 2021). Taken together, these investigations support our findings regarding the ability of BM-MSC to promote in vitro differentiation of respiratory epithelial cells. Although co-cultures with ASC exhibited a low degree of cilia and mucus formation, the corresponding tri-cultures lacked signs of complete mucociliary differentiation. This correlated with the general reduction of mucociliary differentiation in tri-cultures when compared to co-cultures. However, epithelial cell layers in ASC-co-cultures were unusually thick and deficient in cell polarization and a basal cell layer, showing signs of epithelial hyperproliferation. These results go in line with observations made by Kobayashi et al., who found mucin5AC and β-tubulin expression of tracheal epithelial cells in vitro to be reduced in co-cultures with ASC compared to co-cultures with fibroblasts. Moreover, the authors could show an enhanced epithelial proliferation in ASC-co-cultures (Kobayashi et al., 2010).

With respect to vascularization, both MSC and fibroblasts are known to have pro-angiogenic properties (Zupan, 2021). Several investigations could prove the ability of both cell types to promote formation of vascular-like structures in vitro (Rao et al., 2012; Freiman et al., 2016; Lee et al., 2016). In a previous study published by our group, we found both fibroblasts and MSC of different origins to be suitable as supporting cell types for vascularization in vitro, with BM-MSC showing a widespread network of smaller vascular-like structures and more branching points in co-culture with HUVEC compared to the other cell types (Kniebs et al., 2020). This matches our results, although the effect was only visible in BM-MSC-tri-cultures and not as strong in co-cultures. Lumen formation observed for all tri-cultures (see Supplementary Figure S6) indicates presence of essential ECM components known to be synthesized by supporting cells (Newman et al., 2011).

In this study, we analyzed levels of cytokines IL6, IL8, angiopoietin 1, angiopoietin 2, EGF, VEGF, FGF-b and PIGF known to be involved in vascularization (Lovett et al., 2009; Amirsadeghi et al., 2020; Sun et al., 2021). Rather high standard deviations observed in LEGENDPlex assays result from donor-dependent variation. Donor tissue was derived from patients with unknown medical history and was treated anonymously due to reasons of data protection. Therefore, it cannot be precluded that a patient´s medical history may influence the cell behavior in vitro. In general, primary cell differentiation is often connected to high donor-to-donor variability (Russell et al., 2018). Statistically significant differences in the LEGENDPlex assays were only found for VEGF and angiopoietin 2. VEGF is known to promote vasculogenesis by increasing cell proliferation, vascular permeability and vasodilatation (Melincovici et al., 2018). In addition, its expression was documented in a broad variety of tissues and cell types including respiratory epithelial cells (Melincovici et al., 2018; Luengen et al., 2020). As VEGF is supplemented in endothelial cell growth medium 2 (EGM2) with a concentration of 0.5 ng/ml and might additionally be present in the fetal calf serum (FCS) supplement as well, most cultures were found to consume VEGF, while HUVEC-co-cultures with ASC and HNF showed strong VEGF production. This might be a sign for still ongoing vascularization processes in these two cultures. In monocultures of BM-MSC, ASC and HNF, VEGF and angiopoietin 2 production was comparably low (see Supplementary Figure S7). The role of angiopoietin 2 in vascularization and angiogenesis is understood to be highly regulated and context-dependent as it can act as either agonist or antagonist to the potent pro-angiogenic growth factor angiopoietin 1 by binding to the same Tie2 receptor (Akwii et al., 2019). Among other parameters, increased VEGF concentration contributes to the pro-angiogenic activity of angiopoietin 2 (Akwii et al., 2019). Interestingly, we observed VEGF consumption and little angiopoietin 2 production in co-cultures with HRE. These findings support the concept of respiratory epithelial cells and endothelial cells promoting each other in differentiation (Eichmann and Simons, 2012; Luengen et al., 2020). Our data suggest that HRE additionally require VEGF for differentiation and that differentiating HRE also trigger formation of vascular structures. Combining the vascularization and differentiation outcomes with VEGF and angiopoietin 2 levels, it appears that supporting cells assist in providing the necessary growth factor levels, especially in the context of high VEGF-demand by both HUVEC and HRE.

Regarding growth factors IL6, IL8, angiopoietin 1, EGF, FGF-b and PIGF, differences between cultures were found to be non-significant. The observed trend towards higher PIGF levels in cultures containing HUVEC can be explained by the fetal origin of the cells. The protein belonging to the VEGF family was found to be involved in several physiological and pathological vascularization processes. Although its pro-angiogenic effects might be beneficial for vessel formation, increased PIGF levels are connected to pathological angiogenic processes (Shibuya, 2008). In order to avoid potential undesirable effects of PIGF and in addition enhance availability of autologous endothelial cells, HUVEC could be replaced in future experiments by adipose-derived microvascular endothelial cells (Freiman et al., 2018). As angiopoietin 1 is well-characterized with regard to its strong angiogenic effects, higher levels in co-cultures with HUVEC in comparison to tri-cultures support the assumption of upregulated vascularization processes in these cultures (Akwii et al., 2019). Interestingly, a significantly higher production of angiopoietin 1 in the HNF monoculture was not reflected in tri- and co-cultures revealing the importance of cell type interactions for changes in cytokine expression (see Supplementary Figure S7). Next to promoting vascularization, EGF is also known to be involved in mucociliary differentiation of respiratory epithelial cells (Cozens et al., 2018; Luengen et al., 2020). The same applies for FGF-b, which is known to participate in both epithelial repair and angiogenesis processes (Murakami and Simons, 2008; Guzy et al., 2015). Both IL6 and IL8 are pro-inflammatory cytokines, and although inflammation represents a necessary component in tissue regeneration and vascularization, de-regulation and high levels of both can contribute to chronic inflammation and cancer development (Eming et al., 2017; Kaur et al., 2020; Fousek et al., 2021). Altogether, the low interleukin levels observed in the tri-cultures are promising with regard to future applications of the mucosa model. However, the generally available data on cytokines required for successful mucociliary differentiation of HRE is insufficient beyond the cytokines VEGF, EGF and FGF-b. While well-defined and established cytokine panels for vascularization analysis exist, only little is known on the intrinsic factors that finally lead to a mucociliary phenotype. On the other hand, several pro-inflammatory factors are involved in pathological processes of the diseased airway epithelium: IL33, TSLP and TGF-β1 are known regulators in common airway diseases like asthma, allergic rhinitis, COPD or cystic fibrosis (Aschner and Downey, 2016; Stewart et al., 2018; Hong et al., 2020). The low levels of the epithelial “alarmins” IL33 and TSLP (Mitchell and O'Byrne, 2017), that have been found in our study (see Supplementary Figure S8) contribute to the promising results already observed with low IL6 and IL8 levels. And although TGF-β1 was detected in co-cultures with HUVEC and monocultures of supporting cell types (see Supplementary Figure S8), levels in HRE cultures were observed to be very low. Taken together, these results may in future qualify our model to be used as in vitro platform for studies on airway diseases.