<figure><img src="image_1.png" /></figure>

# Engineered micro-structured biomimetic material for modelling the outer blood-retinal barrier

Chloé Dujardin<sup>a,*</sup> <sup>†</sup>, Walter Habeler<sup>b,c</sup> <sup>‡</sup>, Paola Aprile<sup>a,§</sup>, Alessandra Dellaquila<sup>a,§</sup>, Christelle Monville<sup>b</sup>, Didier Letourneur<sup>a,1</sup>, Teresa Simon-Yarza<sup>a,**,1 ‖</sup>

## ARTICLE INFO

**Keywords:**

Tissue engineering

In vitro modelling

Outer blood-retinal barrier

Polysaccharide hydrogel

Freeze-drying

Pre-vascularization

## ABSTRACT

The outer blood-retinal barrier (oBRB) is compromised in several retinal pathologies, such as age-related macular degeneration affecting over 200 million people worldwide. This 200–350 µm thick tissue includes the retinal pigment epithelium (RPE), the Bruch's membrane, and the vascularized choroid supplying the outer retina. Degeneration of the RPE and/or choroid leads to photoreceptor loss and, ultimately, blindness. Current *in vitro* co-culture oBRB models developed to better understand the diseases and to propose therapeutic alternatives are often simplistic, focusing on 2D cultures, or face limitations including non-physiological dimensions or low throughput.

This study presents an innovative scaffold-driven approach to model the oBRB using a polysaccharide membrane engineered by freeze-drying. Our specific protocol allowed to mimic the oBRB structure, within physiological dimensions, generating a non-porous surface to culture the hiPSC-derived RPE monolayer, and an internal 3D porous structure for the choroidal network. Results showed that the inner porous structure promoted physiological self-organization of endothelial cells and pericytes. Our single-piece functional material allowed the cultivation of both RPE and choroidal compartments in close proximity, favoring cellular interactions, while maintaining them in their designated locations. This cyto-compatible, easy-to-use, and off-the-shelf membrane, produced at large amounts and low costs, provides a physiologically relevant biomaterial for oBRB tissue modelling.

## 1. Introduction

The outer blood-retinal barrier (oBRB) is disrupted in many retinal pathologies, notably in age-related macular degeneration affecting about 200 million individuals worldwide, with a prevalence increasing due to the global aging of the population [1]. The oBRB is around 200–350 µm thick [2], and is composed of i) the retinal pigment epithelium (RPE), notably responsible for the barrier functions and crucial for the proper functioning of the photoreceptors, ii) the Bruch's membrane, an acellular collagenous membrane, and iii) the choroid, the vascular layer supplying the outer retina [3,4]. In several retinal pathologies, atrophy of the RPE, along with degenerative alterations of the choroid, leads to a loss of photoreceptors and ultimately, to blindness

[5]. The pathological mechanisms underlying these diseases are mostly misunderstood, as the interactions between the different cell types within the oBRB remain poorly characterized [4]. To better understand the tissue and its alterations, physiologically relevant human oBRB models integrating both the RPE and the choroid are required. Only few co-culture models have been reported so far [5]. More than half of them remained simplistic, with 2D cultures of RPE and endothelial cells on opposite sides of a membrane [6–13]. To produce more relevant vascularized 3D models [14], the choroidal vascularization has notably been modelled in 3D hydrogels using organoids [15,16], microfluidic chips [17–20], and bioprinting [21,22]. Yet, these techniques come with some limitations, such as non-physiological dimensions, the low throughput due to the intricate designs required for microfluidic

<footnote><sup>a</sup> Université Paris Cité, Université Sorbonne Paris Nord, Laboratory for Vascular Translational Science (LVTS), INSERM U1148, 75018, Paris, France</footnote>

<footnote><sup>b</sup> Université Paris-Saclay, Univ Evry, INSERM, I-Stem, UMR861, Corbeil-Essonnes, 91100, France</footnote>

<footnote><sup>c</sup> I-Stem, CECS, Corbeil-Essonnes, 91100, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>** Corresponding author.</footnote>

<footnote>E-mail addresses: chloe.dujardin@inserm.fr (C. Dujardin), teresa.simon-yarza@inserm.fr (T. Simon-Yarza).</footnote>

<footnote n="1">¹ Equal contribution.</footnote>
systems, or the extended printing time, which is further constrained by
the need to match the ink composition with specific cell types in
bio-printed systems.

Various materials have been investigated to model the oBRB, notably decellularized tissues, synthetic materials and natural materials [5]. The superiority of the latter relies on their biocompatibility, biodegradability, mechanical properties, and availability, in particular for bacterial and plant-based materials. In this context, polysaccharides, as an important class of natural macromolecular materials, highly abundant in nature, have been widely studied for tissue engineering applications [23]. Notably, hydrogels can be easily synthesized from polysaccharides, by self-gelation, ionic or covalent cross-linking [23].

Amongst the different techniques to tailor the structure of hydrogels, freeze-drying (FD) appears as an interesting approach to cope with the current limitations of oBRB models. The FD process consists in freezing the samples, and then lowering the pressure to remove the ice water by sublimation, based on the water phase transition properties. Often used in the food and pharmaceutical industries [24,25], FD has also been used in materials science to generate porous scaffolds, notably within hydrogels. The rationale behind this technique is that during freezing, the water gets separated from the polymer and the growth of ice crystals causes the concentration of the polymeric chains. As the ice sublimates, the polymeric chains remain stacked, leading to a porous structure, with the size of the pores being directly proportional to the size of the ice crystals [26]. By controlling the ice crystal growth, it could thus be possible to tune the shape and size of the pores. Various polysaccharide hydrogels such as agarose, chitosan, alginate or carboxymethyl cellulose, have been successfully freeze-dried. Yet, in most studies, the samples were frozen in a conventional freezer before drying, leading to an uncontrolled crystallization, and thus an ungoverned porosity [27–31]. The tailoring of the FD process to closely mimic biological structures thus remains largely unexplored.

Our group has focused on hydrogels made of pullulan and dextran, two biocompatible and biodegradable polysaccharides, covalently cross-linked and freeze-dried [32,33], for various applications such as vascular, bone, and hepatic modelling [34–36]. By combining these two polymers, the resulting hydrogel beneficiates from the mechanical strength of dextran and the flexibility of pullulan [37,38]. These materials can be submitted to UV- or gamma-irradiations for sterilization, making them suitable for *in vitro* culture and *in vivo* applications. Recent works unravelled the mechanisms of ice crystals formation in the pullulan-dextran hydrogels, highlighting that the porous structure is strongly conditioned by the nucleation of ice [39,40]. Yet, the influence of the different freezing parameters on the ice crystals growth, and thus on the resulting porous structure, is still uncharted.

Here, we propose a scaffold-driven approach to model the oBRB, based on a 3D structured hydrogel pullulan-dextran membrane, micro-engineered by FD. The FD process was specifically developed to meet with the oBRB structural requirements within a single-piece material at physiological dimensions, with a non-porous side to support the RPE and an inner 3D porous network for the choroidal vascularization. Addition of a second FD step following the coating of the material with extracellular matrix proteins further improved the structural and biological properties of the membrane while enabling a stable and lasting coating. Moreover, FD allowed for long-term storage of our material and thus provided a ready-to-use cell culture support, which can be produced and used at high-throughput. By co-cultivating human induced pluripotent stem cell (hiPSC)-derived RPE cells on top of the membrane, and primary retinal endothelial cells (ECs) supported by primary retinal pericytes (PCs) within the porous network, we successfully obtained a physiologically relevant oBRB 3D model. Our single-piece design allows for the cultivation of the cell types in close proximity, favoring cellular interactions, while maintaining them in their designated locations. Our approach offers easy-to-use, off-the-shelf, sterilizable and cytocompatible single-piece membrane for oBRB tissue modelling.

## 2. Results and discussion

### 2.1. *Fabrication of a porous structured membrane*

The hydrogel was fabricated by injection molding and *in situ* cross-linking of pullulan/dextran aqueous solution with trisodium trimetaphosphate (STMP) as cross-linker at a concentration of 2.25 %. Our patented protocol [32] relies on aqueous solutions and thus prevents against synthesis-induced cytotoxicity. The manufacturing technique enables a control of the membrane thickness while producing membranes at high throughput (each 50 cm² mold leading to around 30–40 membranes of 1 cm diameter). The spacer was tailored to reach a final thickness between 200 and 250 µm to mimic the thickness of the native oBRB tissue [41]. A patterned Teflon stamp was applied on one side of the mold, imprinting the pattern onto one surface of the hydrogel, which enabled a clear distinction between its two sides (Supp. Fig. 1).

To generate the porous microstructure of the hydrogel mimicking the oBRB architecture, we freeze-dried the samples using a customized protocol and with the patterned side facing upwards during FD. By using a programmable freeze-dryer with cooling shelves, it was possible to precisely set the freezing rates and temperatures, leading to a better control of the crystal growth and thus of the pore size (Supp. Fig. 2) [39]. Notably, a rapid cooling is known to prevent water crystallization [42], and thus pore formation. We used this principle to generate the non-porous smooth surface intended for the culture of the RPE monolayer. We successfully obtained this smooth surface on the side in contact with the shelves by freezing the samples on a thermally conductive iron tray at a rate of $-1 \, ^\circ C \cdot min^{-1}$ to reach $-40 \, ^\circ C$ (condition: Metal no annealing in Fig. 1a). With this controlled freezing, the surface porosity, defined as the pore area relative to the total surface area, of this smooth side was evaluated at only 2.2 % (Fig. 1b). The average pore radius was around $1.36 \pm 0.2 \, \mu m$, which is too small for the RPE cells to penetrate the structure upon seeding, allowing their confinement on the smooth side as intended [3]. When frozen in an uncontrolled manner in a $-20 \, ^\circ C$ freezer, as it is often done in the literature [27–31], the membranes did not present a low-porosity smooth side (Supp. Fig. 2), showing the importance of the directional fast freezing rate. Interestingly, as the cold slowly spreads within the material (from the shelves to the top surface), crystals are formed in the hydrogel, leading to pores in the thickness and on the top surface of the membrane (condition: Metal no annealing in Fig. 1a). Notably, the patterned side, facing upwards during the FD, had a porosity of 7.7 % in average (Fig. 1b) with larger pores which would allow vascular cell penetration in the structure upon seeding on this side (Fig. 1a).

Thus, controlled and directional FD enabled us to generate a non-porous surface, suitable for the culture of a monolayered epithelium, and a porous surface, on opposite sides of a thin hydrogel membrane. In the following, we will refer to the patterned porous side as the “rough” side, and to the non-porous side facing the freeze-dryer shelves as the “smooth” side.

Yet, with these FD parameters, the porous structure was composed of small rounded pores (Fig. 1a, 1st row, last column) and characterised by a low inner porosity (1.73 %; Fig. 1c, first column), defined as the total pore volume relative to the total hydrogel volume, which was poorly adapted to the formation of the pre-vascularization. Indeed, as the choroidal vascularization is composed of blood vessels with a minimal diameter around 10 µm and up to 100 µm [43], the objective was to obtain an inner porous network with elongated pores with a minimal width of 10 µm. To improve the inner porosity, we then optimized the freezing protocol to obtain larger ice crystals and thus larger pores within the membrane.

We first analyzed the impact of the FD support material on the porosity by comparing hydrogels freeze-dried on a metallic support (iron tray) or on plastic (polystyrene Petri dishes), the thermal conductivity of polystyrene being lower than the one of iron. The freezing program remained the same (cooling at $-1 \, ^\circ C \cdot min^{-1}$ to $-40 \, ^\circ C$ followed
<figure><img src="image_2.png" /><figcaption>**Fig. 1. Optimization of the freeze-drying (FD) process.** (a) Representative images of scanning electron microscopy (SEM, 1st and 2nd columns) conducted on dried samples, and of confocal microscopy (3rd column), of the middle slice of hydrated membranes, for 4 different FD protocols. Scale bars: 200 µm. (b–c) For the 4 protocols, quantification of: (b) the porosity of the rough and smooth sides of dried membranes from SEM images and (c) the inner porosity of hydrated membranes from confocal imaging. The surface porosity is defined as the pore area relative to the total surface area, while the inner porosity is defined as the total pore volume relative to the total volume. All membranes here were cross-linked with a 2.25 % STMP solution. Graphs represent the mean with the standard deviation. Statistics: (b–c) Brown-Forsythe & Welch ANOVA test with Dunnett's T3 multiple comparison test, on each data set separately. (ns, not significant; *, p-value <0.05; **, p-value <0.001; ***, p-value <0.0001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>

by 2 h at -40 °C). The low porosity of the smooth side was maintained when switching to plastic (Fig. 1a, no annealing conditions), with a 2.2 % porosity on metal to 0.9 % on plastic (non-significant, Fig. 1b). Confocal imaging showed that the shape of the inner pores changed from small and round pores to elongated pores (Fig. 1a), which led to an increase of the global inner porosity, from 1.73 % on metal to 6.6 % on plastic (p-value <0.001, Fig. 1c). The overall porosity of the rough side,

calculated from SEM images, was similar between the two supports with an average of 7.65 % on metal versus 5.89 % on plastic (non-significant, Fig. 1b). Interestingly, the number of pores per mm² drastically decreased on plastic, from an average of 16,900 on metal to 1,700 on plastic (p-value <0.0001, *Supp Figure 3a*). This suggests that the pore size on the rough side increased on plastic, which is confirmed by the increase in maximal pore radius and the mean pore radius from 10.3 µm
to 51.5 µm, and from 1.3 µm to 2.6 µm respectively (p-value <0.0001 in both cases; **Supp Figure 3b-c**), which would favor vascular cell penetration in the porous network. To understand how the support material impacts the membrane porosity, we analyzed the freezing recordings of the two conditions, obtained with probes placed on the samples during FD (**Supp Figure 4a**). Surprisingly, we evidenced that the effective freezing speed was not affected by the material conductivity, being similar between the metal and plastic conditions (–0.81 °C·min⁻¹ vs. –0.80 °C·min⁻¹ respectively, **Supp Figure 4b**). The ice nucleation temperature was also similar between the two conditions (–8.83 °C and –8.63 °C respectively, **Supp Figure 4c**). In contrast, the duration of the ice nucleation and crystallization significantly increased when the FD was conducted on plastic compared to metal (6.1 min vs. 2.4 min, respectively; p-value <0.001; **Supp Figure 4d**). The longer the ice nucleation, the larger the crystals become, leading to larger pores in the resulting scaffold [26]. By freeze-drying the samples on plastic, we thus obtained a higher inner porosity, with elongated inner pores, and larger pores on the rough side, both factors being important for an efficient pre-vascularization. Yet, preliminary cell seeding assays demonstrated that the inner porosity was still not sufficient to allow optimal cell penetration (data not shown).

To further increase the inner pore size in the membrane, we added an annealing step of 2 h after the initial freezing, and we compared two annealing temperatures: –10 °C and –20 °C. The annealing consists in

raising the temperature of a frozen sample, remaining below its melting temperature, to increase the rate of crystal growth by reducing the solution viscosity, thus leading to larger pores [44,45]. The surface porosities remained similar upon addition of an annealing step (**Fig. 1b**). The inner porosity significantly increased from 6.6 % to 12.2 % with an annealing at –20 °C and reached 19.9 % with a –10 °C annealing (p-values <0.001; **Fig. 1c**), and the pores maintained their elongated shape while becoming larger in width (**Fig. 1a**). For each path of connected pores within the inner structure, we calculated from confocal images the longest shortest path (LSP), which is the shortest route to connect the two most distant points. The median LSP increased from 19.1 µm to 35.0 µm (p-value = 0.008, **) with an annealing at –20 °C, and reached 36.5 µm (p-value = 0.024, *) with an annealing at –10 °C, suggesting that the annealing induced an increase of the pore connectivity [46]. As the highest inner porosity characterized by longer, wider and interconnected pores was obtained with the annealing at –10 °C, this step was kept for the following experiments.

Moreover, to ease the membrane handling during 3D cell culture, we increased the membrane stiffness by increasing the STMP cross-linker concentration from 2.25 % to 3 %, leading to a higher density of covalent bonds between the polymer chains [47]. The Young modulus effectively increased from 7.0 kPa to 12.0 kPa (**Supp Figure 5a**). Importantly, this change in cross-link density did not alter the surface porosities (**Supp Figure 5b,d**). Although a small decrease of the inner

<figure><img src="image_3.png" /><figcaption>**Fig. 2.** Effect on the second freeze-drying on the membrane structure. (a) Representative images of scanning electron microscopy (SEM, 1st, 2nd and 3rd rows) conducted on dried samples, and of confocal microscopy (4th row), of the middle slice of hydrated samples, on membranes after the first and second freeze-dryings (1-FD and 2-FD respectively). The 5th row represents a 3D reconstruction of the cross-section of hydrated membranes from confocal stack imaging, with the smooth side located on top. Scale bars: 200 µm. (b) Surface porosity of the rough and smooth sides, calculated from SEM imaging of dried membranes after the 1-FD and 2-FD. The surface porosity is defined as the pore area relative to the total surface area. (c) Inner porosity, calculated from confocal stack images of hydrated membranes after the 1-FD and 2-FD. The inner porosity is defined as the total pore volume relative to the total volume. (d) Percentage of useful pores in the inner porosity of hydrated membranes after the 1-FD and 2-FD, calculated from confocal imaging. Useful pores are defined as pores with a minimal width of 10 µm. All membranes analyzed here were cross-linked with a 3 % STMP solution. Graphs represent the mean with the standard deviation. Statistics: (b) and (d) Welch's *t*-test; (c) Mann-Whitney test. (ns, not significant; **, p-value <0.01, ***, p-value <0.0001).</figcaption></figure>
porosity was observed from 19.6 % to 16.6 % (p-value = 0.029, **Supp Figure 5c**), the pores maintained the elongated shape (**Supp Figure 5d**). All the following experiments were conducted on membranes cross-linked with 3 % STMP.

## 2.2. Improvement of the membrane porosity with a second freeze-drying

To enhance cellular adhesion on the membrane, the latter was coated with a collagen I solution after the first FD step (1-FD). The membranes were then further freeze-dried to concentrate the collagen on the external and porous surfaces of the membranes and to increase the stability of the coating, based on previous studies from our group [34]. Addition of a second FD step (2-FD) also proved advantageous to the hydrogel structure in several respects (Fig. 2). Firstly, no alteration was observed on the smooth side as shown by SEM imaging (Fig. 2a) and the smooth side porosity was similar between the 1-FD and 2-FD samples (0.44 % and 0.26 % respectively, non-significant, Fig. 2b). This was allowed by the pattern imprinted on the membrane allowing to maintain the polarity of the membrane during the 2-FD: the smooth side was once again placed facing the cooling FD shelves, and the fast-freezing rate of -1 °C/min prevented the formation of ice crystals on this surface as during the 1-FD.

In contrast, the rough side porosity increased by two-fold, from 4.9 % to 10.5 % (p-value <0.01, Fig. 2b). The inner porosity measured from confocal imaging (Fig. 2a), followed the same trend with a two-fold increased, from 16.6 % to 38.8 % (p-value <0.0001, Fig. 2c). We quantified the percentage of useful pores, defined as the pores with the width higher than 10 µm, the minimal width of the choroidal capillaries [48]. 37.3 % of the pores are considered as useful after the 1-FD. This percentage reached 62.6 % after the 2-FD (p-value <0.0001, Fig. 2d). Moreover, the longest shortest path median increased between the two FD, from 36.5 µm to 62.0 µm (p-value = 0.0006,**), suggesting that the 2-FD induces an increase of the pore connectivity, which would favor the membrane pre-vascularization. This increase in pore size during the 2-FD might be explained by the crystallization of the collagen coating solution during the freezing step. Since the pores were filled with collagen solution, as it froze, the volume of the pores increased as the volume of ice water is higher than the volume of liquid water, leading to larger pores once the membrane is dried. Moreover, the 2-FD program was the same as the first one, and thus contained an annealing step, which further increased the size of the pores as demonstrated in section 2.1. It is also important to note that new pores were also generated during the second freeze-drying, due to the formation of new crystals in the polymeric matrix.

Overall, the final optimized protocol with two freeze-drying steps conducted on plastic and with an annealing step at -10 °C led to an increase of the inner porosity by 2000 % (from 1.73 % to 38.8 %) compared to original setup. We obtained a micro-structured hydrogel membrane, with a physiological thickness [41], with a smooth side characterized by a low porosity intended for the culture of the RPE monolayer and a porous network composed of elongated and interconnected pores, with diameters within the range of the native tissue [43]. Moreover, in previously published oBRB co-culture models [6,9, 17,20], the distance between the RPE and the choroid was often too large compared to the native Bruch's membrane (around 7 µm [49]), due to the thickness of the material separating the two layers, which can interfere with cellular interactions. By our original single-material approach integrating the two compartments within a thin membrane, the distance between the smooth side and the first pores was reduced (Fig. 2, cross-sections), decreasing the distance between the two cell compartments. Noteworthy, future developments could focus on further increasing the inner porosity as it has been reported in the literature that the ratio of vessel lumen area versus stromal area could reach 73 % in the choroid [41].

## 2.3. Collagen-coated membrane properties for in vitro experiments

As previously mentioned, to enhance cellular adhesion, the pores and surfaces of the membrane were coated with collagen I between the 1-FD and the 2-FD. Immunostaining against collagen I on hydrated membranes showed that the membrane was efficiently coated on the two surfaces (Cross-section, Fig. 3a) and in the pores (Middle slice, Fig. 3a), with an average pore surface coverage of 67 %. Collagen quantification, performed with a kit detecting both soluble and cross-linked collagen, showed that an average of 50.6 µg of protein was incorporated per milligram of dry membrane. Collagen content was stable at 37 °C upon hydration (after 3 h and 8 days; Fig. 3b). Additionally, second harmonic generation (SHG) imaging demonstrated that the coated collagen was fibrillar and remained stable upon hydration for up to 14 days (Fig. 3c).

We then performed physical characterizations of our membrane, often poorly investigated in previous 3D oBRB models [5].

Firstly, if the interactions between the RPE and the choroid are not yet fully understood, the molecular exchanges between the two layers are well described in the literature [3,48]. Notably, the largest macromolecule exchanged between the RPE and the choroid is the vitamin A with a molecular weight of 75 kDa [50]. To be suitable for the modelling of the oBRB, the scaffold thus needs to be permeable to molecules up to 75 kDa. To assess the membrane permeability, we measured the exclusion limit of the collagen-coated membrane by analyzing the diffusion coefficient of FITC-Dextran of various molecular weights (MW) across the membrane. The exclusion limit was calculated at 133 kDa (Fig. 3d), making our membrane suitable for oBRB modelling in terms of permeability.

Secondly, the stability of the membrane in cell culture conditions was also analyzed over time. Results showed that the membrane mass was stable over the span of one month, enabling long-term cell culture experiments (Fig. 3e).

Lastly, the mechanical properties of the membrane were measured by nano-indentation on the smooth side of 4 samples for 22 days (Fig. 3f). The samples showed a similar Young modulus, around 12 kPa, with no significant difference with time. If it has become increasingly evident that the mechanical properties of a biomaterial have an important impact on cellular behaviour and tissue functionality [51], comparing our results to the properties of the native tissue is challenging since their evaluation differs greatly in methods between the different studies [52]. However, the elasticity of the choroid tissue and of the sclera, measured by indentation have been reported around 20 kPa [52, 53], which is in the range of the elasticity of our hydrogel membrane, around 12 kPa. Together, these results suggest that the membrane was suitable for the modelling of the oBRB in terms of permeability and elasticity and for long-term *in vitro* culture experiments.

## 2.4. Formation of a functional RPE on the smooth side

To model the RPE, several models have relied on the ARPE-19 cell line [6,8,13,17,19,20], whose limitations are numerous, including lack of or limited pigmentation and variable barrier functions [5]. Since the first differentiation protocols published in the late 2000's [54], hiPSC-derived RPE are considered as one of the most relevant cell type to model the RPE. Here, RPE cells derived from hiPSC were seeded on the smooth side of collagen-coated membranes rehydrated in a laminin solution to improve RPE cell adhesion (Fig. 4a). The cells adhered to the smooth surface and did not penetrate the porous structure, being exclusively located on the smooth side (Supp Figure 6a). The average mortality of the RPE cells on the membrane on D4 was similar to the one of RPE cells cultivated on a laminin-coated 24-well plate used as control (Supp Figure 6b). Moreover, image analysis of the cells after 7 days (Supp Figure 6c) confirmed the excellent cytocompatibility of the membrane, with only 0.79 ± 0.45 % cell mortality (n = 10). SEM imaging showed that the RPE cells formed a monolayer on D14 and displayed the typical RPE cobblestone shape (Fig. 4b). Moreover, imaging
<figure><img src="image_4.png" /><figcaption>**Fig. 3. Characterization of the collagen-coated membrane.** (a) Immunostaining against collagen I performed on collagen-coated membrane. Representative images of the middle slice of the hydrogel (on top) and of the cross-section (on bottom) obtained from the 3D reconstruction of a confocal z-stack. On the cross-section, the smooth side is located on top. The membrane is imaged by fluorescent microscopy thanks to the addition of FITC-Dextran during the hydrogel synthesis. Scale bars: 200 µm. (b) Collagen quantification performed on uncoated dried membranes ("membrane" condition), or coated membranes ("membrane + collagen") either dried or hydrated for 3 h or 8 days. (c) Visualization of collagen fibers by SHG imaging. Collagen-coated double freeze-dried membranes were rehydrated in cell culture conditions at 37 °C for 1, 7 or 14 days and second harmonic imaging was conducted to assess collagen fiber formation. A large field of view of the hydrogel is presented (left column) along with a zoom on one pore (right column). In the zoomed image, the pore outline is highlighted by a white dotted line. Scale bars: 100 µm for the large field images and 70 µm for the zooms. (d) Permeability assay performed on collagen-coated membranes to evaluate their exclusion limit (n = 6 for each molecular weight). (e) Evolution of the mass of collagen-coated membranes hydrated at 37 °C in DMEM through time to assess their stability (n = 5). (f) Young modulus of the smooth side of collagen-coated membrane measured by nanoindentation. Nine samples were tested at different timepoints, with several measurements per timepoints. The samples were hydrated in DMEM and kept at 37 °C between measurements. All experiments were conducted on collagen-coated 2-FD 3 % STMP membranes. Graphs represent the mean with the standard deviation. Statistics: (b) Brown-Forsythe & Welch ANOVA test with Dunnett's T3 multiple comparison test. (f) Nested One-Way ANOVA with Sidak's multiple comparison test between each consecutive timepoint. (ns, not significant; ****, p-value <0.0001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>

of the RPE-seeded membrane with a high-resolution microscope showed that the cells presented a weak pigmentation at D7 which got stronger by D14, highlighting that the cells developed a pigmentation through time (Fig. 4c and Supp. Fig. 7). Similarly, the cells developed tight junctions, important for the barrier function of the epithelium (ZO-1, Fig. 4d-e),

with low expression of ZO-1 at D7 but homogeneous staining over the entire epithelium by D14 (Supp Figure 8a). The formation of the barrier was confirmed by transepithelial electrical resistance measurements (TEER) conducted over 3 weeks of culture and which showed that the TEER increased over the first 11 days and then stabilized suggesting the
<figure><img src="image_5.png" /><figcaption>**Fig. 4.** Retinal pigment epithelium on the smooth side of collagen-coated membranes. (a) Schematic representation of the RPE seeded membrane. Created with BioRender. (b) Cobblestone morphology of the RPE monolayer seeded on the membrane for 14 days, imaged by scanning electron microscopy. Scale bar: 50 µm. (c) Pigmentation of the RPE cells, after 14 days of culture, visible on bright-field microscopy. Scale bar: 100 µm. (d) Representative confocal images of RPE immunostainings allowing to evaluate the expression of RPE typical markers after 14 days of culture. Scale bars: 100 µm for the first three columns and 25 µm for the fourth column. (e) Cross-section reconstructions of samples stained against ZO-1, Ezrin and MERTK (from top to bottom) after 14 days of culture. Scale bars: 25 µm. (f–g) Concentration of (f) pigment epithelium-derived factor (PEDF) and (g) vascular endothelial growth factor (VEGF) measured by ELISA assays in the supernatant of RPE cells cultivated on the smooth side. The control medium was analyzed along all samples and the resulting value was subtracted from all data points. Graphs represent the mean with the standard deviation. Statistics: Brown-Forsythe & Welch ANOVA test with Dunnett's T3 multiple comparison test (ns, not significant; **, p-value <0.01). All experiments were conducted on collagen coated 2-FD 3 % STMP membranes. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>

formation of a stable epithelial barrier (Supp Figure 8b). Even if the TEER measurements are known to be set-up and experimenter-dependent, it is still worth noting that the value measured at maturation was comparable to those reported in studies using human explanted RPE cells [55] and hiPSC-derived RPE cells [56]. Immunostainings at 14 days of culture confirmed the expression by the RPE cells of ezrin, marker of the microvilli, of MITF, a critical transcription factor for RPE development and function, along with the expression of markers of the visual cycle (RPE65 and CRALBP), of POS phagocytosis (MERTK)

and of transepithelial transport (BEST-1) (Fig. 4d). Cross-section reconstructions of the ezrin and MERTK stainings demonstrated that those two markers were properly located on the apical side of the RPE sheet, suggesting the proper polarization of the RPE cells, whereas ZO-1 is located between the cells as expected (Fig. 4e). Finally, if numerous RPE cells were still proliferative on D14, as shown by the positive Ki67 staining, most of the cells became quiescent at D21 as expected for further use (Supp. Fig. 9). Only 0.55 ± 0.37 % of the cells displayed a Ki67 positive staining at D21 (n = 5).
ELISA assays were performed on the culture supernatants at different time points to quantify the secretion of two typical growth factors: pigment epithelium-derived factor (PEDF) and vascular endothelial growth factor (VEGF). RPE cells cultivated on the membranes produced PEDF, with a concentration increasing through time as the cells matured and reaching 3.1 ng mL⁻¹ on D14 corresponding to a secretion of 9.3 ng·cm⁻² (Fig. 4f). Wide range of PEDF secretions have been previously reported in the literature, ranging from 1.3 ng·cm⁻² to 184 ng·cm⁻² for different cell types such as explanted RPE or hiPSC-derived RPE [11, 57–59]. Results showed that RPE cells also produced VEGF, with a concentration being stable throughout the second week in culture and measured at 96 pg mL⁻¹ after 2 weeks corresponding to a secretion of 290 pg·cm⁻² (Fig. 4g). As for PEDF, a wide range of VEGF secretions have been reported in the literature, from 100 pg·cm⁻² to 7 ng·cm⁻² [11, 55, 57, 59, 60]. For both PEDF and VEGF, the secretions measured in this study fell within the range of values reported in the literature.

We then analyzed the production and remodelling of extracellular matrix (ECM) by the RPE on the membranes. Immunostainings revealed the presence of collagen I, laminin, collagen IV, and fibronectin (Fig. 5a), main components of the RPE basement membrane in the Bruch's membrane [61]. Noteworthy, immunostaining could not distinguish the coated collagen I and coated laminin from the proteins produced in situ by the cells. Quantitative measurements, as ELISA as- says, could be performed to measure the quantity of laminin and/or collagen I through time compared to unseeded coated membranes as

control, to see if the protein quantity increases through time suggesting that the RPE cells produce these proteins. Multiplex assays evidenced the secretion of MMP-1 (a type I collagenase) and MMP-2 (a type IV collagenase), suggesting an ECM remodelling by the RPE (Fig. 5b-c). Interestingly, a decrease in MMP-1 through time was observed, while the secretion of MMP-2 increased over the first week. This suggests an initial remodelling of the coated collagen I followed by a remodelling of the collagen IV produced by the cells. MMP-3, responsible of the degradation of multiple ECM proteins, was detected in lower concen- trations, and was similar for the different time points (35.6 ± 27.5 pg mL⁻¹ at D1; 17.9 ± 11.6 pg mL⁻¹ at D3 and 28.5 ± 21.5 pg mL⁻¹ at D7; *n* = 3 for each). Only few studies have reported the quantification of MMPs concentrations in RPE culture. Most of them were conducted on the ARPE-19 cell line cultivated on culture plates, and reported con- centrations of MMP-1, MMP-2 and MMP-3 around 9 ng mL⁻¹, 4 ng mL⁻¹ and 84 pg mL⁻¹ respectively which are in the range of those reported here [62–64]. Together, these results showed that the RPE produced and remodeled ECM proteins, forming a basement membrane resembling the native one in the Bruch's membrane.

Multiplex assays were conducted to assess the secretion of inflam-
matory markers. CCL-5/RANTES, IL-1α, IL-1β or TNF-α, all secreted by
RPE cells in response to inflammatory signals [65], were not detected in
the RPE supernatants over the first week in culture (Supp. Table 1). IL-6,
IL-8 and GM-CSG were detected at concentrations lower than those re-
ported in the literature, notably for hiPSC-RPE cultivated on laminin

<figure><img src="image_6.png" /><figcaption>**Fig. 5.** Extracellular matrix proteins production and remodelling by the RPE cells on the smooth side. (a) 3D reconstruction of confocal stacks of the RPE monolayer on the smooth side. Samples were stained against extracellular proteins (visible in red) after 7 days of culture. Cell nuclei are visible in blue (DAPI) and the hydrogel in green (FITC-Dextran). Each stack measures 500*500 µm² and 50 µm in thickness. (b–c) Concentration of (b) matrix metalloproteinase-1 (MMP-1) and (c) MMP-2 measure by Luminex assay in the supernatant of RPE cells cultivated on the smooth side. The control medium was analyzed along all samples and the resulting value was subtracted from all data points. Graphs represent the mean with the standard deviation. Statistics: Brown-Forsythe & Welch ANOVA test with Dunnett's T3 multiple comparison test (ns, not significant; *, p-value <0.05; **, p-value <0.01). All experiments were conducted on collagen coated 2-FD 3 % STMP membranes. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>
521 coated plates [66]. Indeed, Hytti et al. reported a basal secretion of IL-6 and IL-8 at concentrations around 20 ng mL⁻¹ and 12 ng mL⁻¹ respectively while we observed secretion below 10 pg mL⁻¹ and 40 pg mL⁻¹ respectively (Supp. Table 1) [66]. CCL-2, a chemokine basally produced by RPE cells, was the most secreted inflammatory marker, amongst those analyzed, with a concentration increasing through time (from 1,378 pg mL⁻¹ on D1 to 1,649 pg mL⁻¹ on D3 (p-value = 0.04, *), to 1,920 pg mL⁻¹ on D7 (p-value = 0.0026, **, RM one-way ANOVA)). Nevertheless, these values remained inferior to levels reported in the literature, that were around 100 ng mL⁻¹ [66,67]. MMP-9, associated with an inflammatory behavior, was undetected in the cell media. Thus, the RPE cells exhibited low levels of inflammatory markers when cultivated on the membranes, compared to previous models reported in the literature.

Overall, we obtained on the smooth side of the membrane a func-
tional and polarized RPE monolayer, comparable to the native tissue,
exhibiting all main RPE markers, secreting typical growth factors and
producing a basement membrane with a composition similar to the
native Bruch's membrane. The cytocompatibility of our membrane has
been confirmed by the low mortality of the RPE cells and the low
expression of inflammatory markers.

2.5. Endothelialization and stabilization by pericytes

Human umbilical vascular endothelial cells (HUVECs) have been
abundantly used in vascularized models and notably for choroidal
vasculature [6-8,13,17,19,20], a choice due in part to the easy access
compared to other primary EC type. However, a study conducted in the

<figure><img src="image_7.png" /><figcaption>**Fig. 6.** Pre-vascularization of the inner porous structure of collagen-coated membranes. (a) Schematic representation of the endothelial cells (ECs in yellow) seeded in the membrane. Created with BioRender. (b) Representative confocal image of the inner porosity of membrane seeded with ECs and stained against CD31. Scale bar: 200 µm. (c) Zoom on the pore highlighted by the dotted frame in image (b). Scale bar: 100 µm. (d) Schematic representation of the endothelial cells and pericytes (ECs in yellow and PCs in red) seeded in the membrane. Created with BioRender. (e) Representative confocal image of the inner porosity of membrane seeded with EC + PC and stained against CD31 and NG2. The cells were seeded to a ratio of 1:5 PC:EC. Scale bar: 200 µm. (f) Zoom on the pore highlighted by the dotted frame in image (e). Scale bar: 100 µm. (g-i) Concentration in the supernatant on day 7 of (g) epidermal growth factor (EGF), (h) hepatocyte growth factor (HGF) and (i) CD31 for membranes seeded with ECs or with ECs and PCs. Measures were obtained by Luminex assay. (j) Nitric oxide concentration in the supernatant at day 7 in the EC or EC + PC conditions. All experiments were conducted on collagen-coated 2-FD 3 % STMP membranes. For all assays, the control medium was analyzed along all samples and the resulting value was subtracted from all data points. Graphs represent the mean with the standard deviation. Statistics: Welch t-tests (*, p-value <0.05; **, p-value <0.01; ***, p-value <0.0001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>
early 2010's reported significant differences in the gene expression of HUVECs compared to ocular vascular ECs, suggesting that HUVECs are not appropriate to model ocular vascularization [68]. As a more physiological source, primary human retinal microvascular endothelial cells (HMRVECs) were recently used in oBRB models [10,12,18]. Although HMRVECs are explanted from the inner blood-retinal barrier, and are thus not choroidal, they are known to possess features that make them more relevant for choroidal modelling [4,68].

HRMVECs were thus seeded in the porous structure of the membrane to mimic the choroidal pre-vascularization (Fig. 6a). The ECs entered the inner porosity (Supp Figure 10.a), and, as for the RPE, spatial confinement was achieved as the cells remained in the porous structure, throughout the entire thickness without crossing to the smooth side (Supp Figure 10.a). Viability assay demonstrated a high survival rate on D3 (Supp Figure 10.b) and on D6 (images not shown). Image analysis showed an average EC mortality of 5.6 ± 2.4 % on D3, and of 10.1 ± 4.7 % on D6 (non-significant, Supp Figure 10.c). In the porous network, the ECs adhered to the pores (Fig. 6.b-c and Supp Figure 10.d), and expressed EC markers such as CD31 (Fig. 6.b-c) and VE-Cadherin (Supp Figure 10.e). The porous structure of our membrane is thus suitable for the culture of an endothelial network.

Yet, it is now well established that addition of support cells is key to induce robust and stable vascularization [69]. In this perspective, fibroblasts, mesenchymal stem cells and/or pericytes (PCs) have been included in some oBRB models to support the formation of the vascular network [17,18,20–22]. We decided to add PCs in our vascular network. Indeed, used only in one model [22], PCs are crucial for blood vessel maturation and stabilization [69,70]. They surround the ECs in the perivascular space, and their coverage is particularly high in the choroid, reaching an average of 11 %, with the highest coverage observed in the capillaries [71].

To account for the choroidal cell complexity, we thus added human primary retinal PCs to the vascular compartment [48]. ECs and PCs were seeded at the same time on the rough side of hydrated collagen-coated membranes (Fig. 6d). Different ratios were tested (1:2, 1:5, 1:10 PCs: ECs) and the 1:5 ratio resulted in the best cellular organization (data not shown). At this ratio, the cells entered the porous structure, and self-organized with the ECs on the inside of the lumen and with the PCs on the outside, as shown by the CD31 and NG2 stainings (Fig. 6e-f). Analysis of the immunostaining images showed that the average coverage of the endothelium by the PCs was of 14.3 ± 2.7 % (n = 7 samples, at least 8 pores analyzed by sample), with low variation between the different pores and the different samples. This value is in the range of the coverage observed in the native tissue (around 11 %) [71].

Multiplex assays were conducted on the supernatants of mono- (EC) and co-cultured (EC + PC) samples to assess secretion and/or uptake of angiogenic proteins. Results on D7 showed a decrease of the epidermal growth factor (EGF) concentration in the co-culture condition (Fig. 6g). As EGF is involved in PC proliferation and recruitment [72–74], this decrease in EGF levels strongly suggests an effective recruitment of the PC to the endothelium in agreement with the obtained confocal images. Moreover, a significant increase of the hepatocyte growth factor (HGF) concentration was observed in the co-culture condition compared to the EC monoculture (Fig. 6h). As HGF is basally secreted by PCs to protect them against apoptosis and to strengthen EC junctions [75], this result highlights that PCs were functional when cultivated with ECs in the porous structure. Moreover, the secretion of the interleukin IL-6, basally secreted by PCs, notably in angiogenesis, was increased in the EC + PC condition compared to the EC monoculture with a concentration of 4, $151 \pm 431 \text{ pg mL}^{-1}$ versus $456 \pm 540 \text{ pg mL}^{-1}$ (p-value <0.0001), similarly to the literature [76,77], and confirming the functionality of PCs in co-culture. Additionally, the concentration of secreted CD31 decreased in co-culture on D7 (Fig. 6i). Given that CD31 is a trans-membrane protein involved in EC cell-cell junctions, its secretion implies either an activation of the endothelium or an endothelial cell mortality. In both cases, the decrease of secreted CD31 in the co-culture

condition suggests a stabilization of the endothelium by the PCs. Moreover, we observed in the EC + PC condition on D7 a slight increase of the secretion of nitric oxide (Fig. 6j), important in the regulation of vasodilation, inflammation and angiogenesis [78], and continuously secreted in the retinal vasculature [79]. Our results are consistent with two previous studies that observed an increase of HGF secretion along with a decrease of EGF and CD31 upon addition of PCs [76,80]. It is interesting to note that these two studies were conducted in 2D, with either no direct contact between the two cell types, or with the ECs cultured on top of a PC monolayer. As far as we know, this is the first time the interactions between ECs and PCs were analyzed in a 3D EC-PC retinal physiological organization.

We then analyzed the production of ECM proteins in the EC and EC + PC conditions by performing immunostainings against collagen I and collagen IV on D5 of co-culture. Results showed that both proteins are produced in both conditions (Fig. 7a). The presence of collagen IV is particularly important for this model as it is the main component of the basement membrane in the choroid [81,82]. Interestingly, the zoom images in the last column highlight that the produced collagen IV was not observed in the same location in both conditions (Fig. 7a). Indeed, the cells directly adhered to the side of the pores in the EC monoculture, and the collagen was thus mainly located on the surface of the pores. In contrast, in the EC + PC condition, the collagen was mostly observed within the lumen of the pores between the ECs and the PCs, highlighting the mimicry of the 3D physiological organization of the two cell types. We also analyzed how the PCs affected the ECM remodelling. Levels of MMP-1 did not significantly change between the different timepoints (D1, D3 and D7) when ECs were cultivated alone (Fig. 7b). In contrast, in co-culture with PCs, the MMP-1 concentration increased through time and reached on D7 a concentration of $121.8 \pm 21.6 \text{ ng mL}^{-1}$, significantly higher than in mono-culture (11.9 ± 5.7 ng mL$^{-1}$, p-value <0.0001, Fig. 7b). These data suggest an increased collagen I remodelling in presence of PCs. The concentrations of MMP-2 were similar for both conditions at each timepoint, suggesting a similar remodelling of the collagen IV in both conditions, with yet a higher variability in the EC monoculture (Fig. 7c). A higher MMP-3 concentration on D7 in the EC + PC conditions compared to EC alone ($46.1 \pm 18.6 \text{ pg mL}^{-1}$ versus $15.5 \pm 10.1 \text{ pg mL}^{-1}$; p-value = 0.04) also indicated an increased ECM remodelling in presence of PCs.

By simultaneously adding the ECs and PCs in the porous network, we thus obtained a more stable pre-vascularization, with physiological organization and coverage of the ECs by the PCs, and with production of the key ECM proteins of the native choroidal tissue. We thus selected this condition for the modelling of the entire tissue with both the RPE monolayer and the vascular compartment.

## 2.6. A complete and functional oBRB model on the hydrogel membrane

Once we established the culture protocols and characterized each cellular compartment, we assembled the RPE, ECs and PCs to model the entire oBRB. We first seeded the RPE cells on the smooth side of collagen-coated membrane rehydrated in a laminin solution. On day 8 of RPE culture, the samples were flipped and ECs and PCs were seeded to a 5:1 ratio on the rough porous side (Fig. 8a).

Immunostaining and confocal imaging were conducted on D7 of co-culture (corresponding to D15 for the RPE cells). Despite the 8 days of membrane hydration for RPE culture, the collagen coating stability (Fig. 3b-c) allowed the ECs to adhere to the pores, self-organize with the PCs and express EC markers as CD31 (Fig. 8b). Moreover, we assessed the expression of typical markers by the RPE cells in tri-culture. Immunostainings showed that they expressed ezrin (Supp Figure 11a) and ZO-1 tight junctions (Supp Figure 11b). TEER measurements showed that the barrier functions of the RPE were not altered in the tri-culture with ECs and PCs (Supp Figure 11c).

3D reconstruction of the oBRB model showed a distinct location of the two cell layers, with the RPE on the smooth side, and the ECs and PCs
<figure><img src="image_8.png" /><figcaption>**Fig. 7.** Extracellular matrix proteins production and remodelling by the vascular cells in the inner porous structure of collagen coated membranes. (a) 3D reconstruction of the inner porosity of samples cultivated with either endothelial cells (EC, 1st row) or with endothelial cells and pericytes (EC + PC, 2nd row). The samples were stained after 5 days of culture against collagen I (in red, first column) and collagen IV (in magenta, 2nd and 3rd columns). The last column displays zooms of the pores highlighted by the dotted frame in the 2nd column images. Cell nuclei are visible in blue (DAPI) and the membrane in green (FITC-Dextran). Each 3D reconstruction measures 1 mm². Vertical blue scale bar (in the 2 first columns): 300 µm. Scale bar in the zooms (3rd column): 100 µm. (b–c) Evolution of (b) the matrix metalloproteinase-1 (MMP-1) and (c) MMP-2 concentrations over a week for the EC and EC + PC conditions. Measurements were done by Multiplex assays. For all assays, the control medium was analyzed along all samples and the resulting value was subtracted from all data points. All experiments were conducted on collagen-coated 2-FD 3 % STMP membranes. Graphs represent the mean with the standard deviation. Statistics: Two-way ANOVA with Tukey's multiple comparison tests (*, p-value <0.05; **, p-value <0.01; ***, p-value <0.0001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>

within the inner porosity, thanks to a thin non-porous layer separating
the two compartments in our membrane (Fig. 8b and Supp Figure 11a).
Quantification of the distance between the RPE layer and the closest
vascular cells showed that only 24.6 ± 5.8 µm separate the two cell
compartments (measured on n = 20 samples). This represents a signif-
icant improvement compared to several previous models which strug-
gled in maintaining a stable long-term co-culture. For instance, Chung *et*

al. added a 300 µm thick blank hydrogel between the RPE and the EC
network to avoid vessel regression, potentially altering cellular com-
munications. Cell migration from one side to the other was also reported
in several oBRB models [9,13,20]. Moreover, the analysis of the mem-
brane diameter and thickness over time showed that the membrane was
stable for at least 16 days in culture, without any degradation or
shrinkage (Supp. Fig. 12). This constitutes an advantage of our material
<figure><img src="image_9.png" /><figcaption>**Fig. 8.** Complete oBRB modelling on the collagen-coated membrane. (a) Schematic representation of the oBRB tissue model indicating the localization of both the epithelium and the vascularization cells. Created with BioRender. (b) 3D reconstruction of part of a membrane cultivated with RPE cells on the smooth side (visible in blue) and with endothelial cells (in yellow) and pericytes (in red) in the porous structure (in green) for 7 days. For proper visualization of the pre-vascularization, the channel representing the hydrogel was removed in the two images on the left. The entire thickness of the hydrogel is visible in the top left and bottom images while the top right image shows a section of the hydrogel to see the seeded inner porous network. In all images, the smooth side is located on top. Vertical blue scale bar (thickness) = 300 µm. (c–j) Concentrations in the supernatant at D7 of (c) CD31; (d) angiopoietin-2 (Ang-2); (e) epidermal growth factor (EGF); (f) matrix metalloproteinase 2 (MMP-2); (g) vascular endothelial growth factor (VEGF); (h) pigment epithelial growth factor (PEDF); (i) hepatocyte growth factor (HGF) and (j) matrix metalloproteinase-1 (MMP-1). Three conditions were analyzed: either RPE cells alone (RPE - control n°1), either the pre-vascularization compartment alone (EC + PC – control n°2) or the two together to form the entire oBRB (RPE + EC + PC). Measures were done by multiplex assays. The control medium was analyzed along all samples and the resulting value was subtracted from all data points. All experiments were conducted on collagen-coated 2-FD 3 % STMP membranes. Graphs represent the mean with the standard deviation. Statistics: (c, d, h, i) Welch t-test; (e, f, g, j) Brown-Forsythe & Welch ANOVA test with Dunnett's T3 multiple comparison test (ns, not significant; *, p-value <0.05; **, p-value <0.01; ***, p-value <0.001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)</figcaption></figure>

compared to ECM-based hydrogels, such as collagen or fibrin, that are
used in several oBRB co-culture models, but that are known to shrink
due to forces applied by the cells. Such materials often require chemical
modifications [83–85] or addition of another polymer, such as poly-
saccharides [83,86], to prevent rapid shrinkage.

To further assess cellular functionality and investigate cellular in-
teractions within the oBRB model, we compared three conditions by
multiplex assays: (1) RPE cells alone on the smooth side (RPE), (2) ECs
and PCs cultivated in the porous network (EC + PC), or (3) the complete
oBRB model (RPE + EC + PC). Firstly, results showed similar levels of
secreted CD31 in the tri-culture condition compared to the EC + PC condition (Fig. 8c), suggesting that the RPE did not disrupt the endothelial stabilization induced by the PCs (Fig. 6i). Secondly, similar levels of angiopoietin-2 (Ang-2) were measured in the EC + PC and the tri-culture condition (Fig. 8d). As Ang-2 is basically produced by ECs, its secretion in our model suggests EC integrity. Interestingly, the levels reported here are 4–5 times higher than those previously reported in 2D co-culture of HRMVEC and HRMPC [76,80]. As Ang-2 is known to promote angiogenesis and vascular tube formation in vitro [87], the 3D structure of our membrane seems to increase the functionality of our vascular network compared to 2D co-culture. Moreover, the RPE cells responded as PCs (Fig. 6g) to the EGF in the culture medium (Fig. 8e), known to increase glucose transport by the RPE [88]. In tri-culture, the concentration of EGF further decreased, suggesting a proper function of the RPE and PC that both consume it. Concerning the ECM remodelling, the concentration of MMP-2 in the tri-culture condition was similar to the one in the EC + PC conditions and higher than in the RPE condition. This is likely due to the stronger remodelling of the vascular compartment on D7 of tri-culture, compared to the RPE compartment cultivated on the membrane for 8 additional days.

Interestingly, the analysis of several soluble factors suggested the presence of interactions between the two compartments of the tri-culture model. Firstly, quantification of the VEGF concentration showed that the RPE cells produced VEGF in monoculture, while EC and PC consumed the VEGF in the culture medium (Fig. 8f), to levels similar to those reported in the literature [76]. When cultured together, we observed a regulation of the VEGF concentration as the measured tri-culture value was higher than the expected one (corresponding to the addition of the two monoculture controls) (Fig. 8f). This suggests either a decrease in the EC + PC uptake in tri-culture or a higher secretion by the RPE in presence of the vascular cells, as it has been reported in the literature for RPE-EC co-culture [8,89]. Secondly, results showed a higher PEDF concentration in the oBRB model compared to the RPE monoculture (Fig. 8g). This suggests an increased PEDF secretion by the RPE in co-culture, and thus an increased activity of the RPE. This result is consistent with previous observations in RPE-EC co-cultures with various cells lines (primary RPE/primary ECs [9], and hESC-derived RPE/HRMVECs [10]). If our results are in the same range as those of Shokoohmand et al., the PEDF concentrations observed by Skottman et al. using hESC-derived RPE and HRMVEC are around 10 times higher than the PEDF concentrations observed in our study [9,10]. Yet, since the RPE cells are known to increase their PEDF secretion as they mature, these differences can probably be explained by the difference in the culture duration of the RPE cells (6 weeks for Skottman et al. vs. 3 weeks for Shokoohmand et al. and vs. 2 weeks in our case). Moreover, a decrease in the HGF concentration was observed in the oBRB model compared to the EC + PC condition (Fig. 8h). To the best of our knowledge, no previous co-culture oBRB model has reported measurements of HGF. Yet, this trend could be explained by a reduced secretion of HGF by the PCs in presence of the epithelium, or more probably by a consumption of HGF by the RPE cells, known to respond to HGF in vitro via the c-Met receptor [90,91]. Additionally, the MMP-1 concentration in the oBRB model with the tri-culture was similar to the RPE control but was significantly lower than in the EC + PC control, suggesting a reduced collagen remodelling of the vascular compartment in presence of the RPE (Fig. 8i). To the best of our knowledge, no previous co-culture oBRB model has reported measurements of the MMP-1 concentration. Interestingly, Song et al. reported a decreased expression of MMP-1 measured by single-cell RNA sequencing in both ECs and fibroblasts when cultivated in 3D in presence of RPE, which is in line with our results [22]. Lastly, we did not observe an increase in inflammatory marker secretion in the tri-culture conditions (Supp. Table 2). As in the control conditions, RANTES and TNF-α were undetectable in the oBRB model (Supp. Table 2). Concentrations of IL-1β and IL-6 were similar in tri-culture and in the EC + PC condition, with low concentrations in the RPE control (Supp. Table 2). MCP-1, GM-CSF, IL-1α and IL-8 were

secreted in the three conditions (Supp. Table 2). To analyse the interactions between the RPE and EC + PC layers, we calculated the expected concentrations if no regulation was observed by adding the value of the RPE control to the value of the EC + PC control (3rd column in the Supp. Table 2). Interestingly, the tri-culture value (4th column in the Supp. Table 2) was lower than the range of the expected concentration for all 4 inflammatory markers, suggesting a regulation of MCP-1, GM-CSF, IL-1α and IL-8 in the tri-culture condition.

### 3. Conclusion

Overall, our work provides a new approach to model the oBRB based on a single-piece membrane with a micro-structure generated by FD to mimic the structural features of the tissue. We thus obtained a membrane with a smooth non-porous side suitable for the culture of a functional and polarized RPE, expressing the main RPE markers and tight junctions, and secreting typical growth factors. Optimization of the FD protocol allowed to generate an inner porous structure composed of elongated and interconnected pores, with a diameter suitable for the culture of vascular cells and in range with the native choroidal vessels. The inner structure facilitated the 3D self-organization of PCs around the endothelium, to stabilize the latter and better model the choroidal vessels characterized by a high PC coverage. Each cell type produced ECM proteins to form basement membrane resembling the native ones. Further characterization of the membrane also assessed its suitability its terms of permeability and elasticity to model the oBRB. Finally, the tri-culture model encompassing the RPE, the ECs and the PCs showed that our membrane allowed for a proper cell compartmentation of the two cell layers, with functional inter-cellular interactions. This is noticeable by the short distance between the 2 compartments, only 2–3 times higher than the native Bruch's membrane. Thus, our approach offers an easy-to-use, off-the shelf and cytocompatible membrane for the modeling of a pre-vascularized oBRB tissue.

Our model could be easily adapted to the establishment of oBRB disease models, notably by using cells derived from patients, and to the screening of pharmaceutical compounds. If we used primary vascular cells in this study, the consolidation of protocols that have been recently reported to differentiate hiPSC into choroidal ECs and PCs, will offer an interesting alternative to primary cells for future developments [92]. We could also envision in future developments adding other cell types in the porous network, such as fibroblasts, melanocytes, or immune cells, to even better mimic the native tissue. Additionally, for such in vitro developments, we could add an active perfusion through the porous network, particularly since the blood flow is particularly high in the choriocapillaris [93].

Previous results in our team have demonstrated the potential of pullulan/dextran hydrogels to be used as grafts with excellent in vivo performances in small and large animals [36,94,95]. Future efforts will be devoted to assess our oBRB membrane as a potential transplant in oBRB pathologies. Latest therapeutic strategies have focused on the replacement of the degenerative RPE layer, by implantation of a RPE sheet grown in vitro. Yet, a degeneration of the choroidal vascularization is often observed in oBRB pathologies. A biodegradable and biocompatible implant integrating both the RPE and the choroid could thus improve clinical outcomes [4]. In this context, the pre-vascularization of the graft with ECs and PCs would benefit the RPE implantation, by improving graft integration along with implanted cell survival [69,96,97].

Finally, the manufacturing method established here provides flexibility to adjust material properties, including the thickness and the size of the membrane, without altering the synthesis duration, or to tailor the membrane stiffness. Mastering the FD process allows adapting the porosity to match other tissues characteristics. Thus, we can envision modelling other types of vascularized epithelial tissues.
# 4. Materials and methods

## 4.1. Injection mold fabrication

Injection molds were custom-fabricated from two Plexiglas slides (10 cm * 5 cm each). The first slide was drilled with one hole in the center to insert the injection syringe, and two holes on opposite corners to allow the excess of solution to drain out the mold. The other slide was covered with a patterned material (208G, Technion Europe, France) that will be imprinted on one side of the hydrogel to distinguish the two sides. The two slides were separated with a 130 µm spacer (216 GP, Techniflon Europe, France) and the mold was tightly closed by clamps.

## 4.2. Hydrogel membrane preparation

The polysaccharide solution was prepared as previously reported [33]. Briefly, pullulan (Mw = 200 kDa, Hayashibara, Japan), dextran (Mw = 500 kDa, Pharmacosmos, Denmark) and NaCl (S9888, Sigma-Aldrich) were dissolved in deionized (DI) water, to a final concentration of 22.5 % (w:v), 7.5 % (w:v) and 35 % (w:v) respectively. When needed, Fluorescein Isothiocyanate (FITC)-labelled dextran (FD-500, TdB Labs) was added to the solution to a concentration of 0.5 mg mL⁻¹ to make the hydrogel fluorescent. The solution was stirred for 1 h at room temperature (RT, 300 rpm) and then left with no agitation for 24 h at RT for air bubble removal.

The cross-linking of the membrane was performed according to a protocol patented by our laboratory [32]. First, a solution of either 225 mg mL⁻¹ (2.25 % condition) or 300 mg mL⁻¹ (3 % condition) of trisodium trimetaphosphate (STMP, Na₃P₃O₉, T5508, Sigma) was prepared in DI water and stirred for 15 min for complete dissolution. In the meantime, to activate the hydroxyl groups of the polysaccharide chains, NaOH (1 mL, 10 M in DI water, 221465, Sigma) was added to the polysaccharide solution (10 g) which was then stirred for 15 min at RT (200 rpm). The STMP solution (1 mL) was then added to the activated solution and rapidly mixed (400 rpm). Once homogenized, the solution was injected in the injection mold and cross-linked for 20 min at 50 °C. Once fully cross-linked, the membrane was peeled from the mold and neutralized in PBS 10X (2 baths of 30 min). To wash the excess of salts, the hydrogel was then rinsed five times for 30 min in DI water. It was finally incubated overnight in PBS 0.1X.

The hydrogel was punched into 12 mm diameter discs, placed in metallic or polystyrene dishes (351029, Corning) with the patterned side on top (referred as the rough side; the side facing the shelves is referred as the smooth side) and loaded in the freeze-dryer (Pilot -80 freeze dryer, Cryotec®, France). The samples were all first frozen at a speed of -1 °C·min⁻¹ to reach -40 °C. Then, either the -40 °C temperature was maintained for 2 h, or an annealing step was added by increasing the temperature to -20 °C or -10 °C for 2 h before going back to -40 °C. The drying program was consistent between the conditions: 1) decreasing the pressure to 0.05 mbar at a constant temperature of -40 °C for 2 h, 2) increasing the temperature to -5 °C at a rate of 0.2 °C·min⁻¹, 3) maintaining the temperature of -5 °C and the pressure of 0.05 mbar for 8 h, 4) increasing the temperature to 10 °C at a rate of 0.2 °C·min⁻¹, 5) maintaining these parameters for 3 h, and 6) increasing the temperature to 30 °C and decreasing the pressure to 0.001 mbar for 1 h.

To study the effect of freezing the hydrogels in a -20 °C freezer, in an uncontrolled manner, instead of in the programmable freeze-dryer, the 130 µm spacer of the molds was replaced by a 1 mm spacer. The membranes were then synthesized as previously described, frozen on a metallic tray in a -20 °C freezer or in the freeze-dryer as control, loaded in the cold freeze-dryer (-40 °C), and dried using the program described above.

A schematic representation of the membrane preparation is presented in the Supp. Fig. 13.

## 4.3. Collagen coating

To enable cellular adhesion, the freeze-dried membranes were coated with collagen, as previously described [34]. Briefly, highly purified bovine corium acid solubilized collagen (FS22024, Collagen Solutions, United Kingdom) was diluted in 0.01 M HCl to obtain a final concentration of 1 mg mL⁻¹. The samples were placed in a 20 mL sealed syringe with the collagen solution. The collagen penetration within the porous structure was obtained by vacuum impregnation, by flushing the solution back and forth with the syringe. The samples were then incubated for 2 h at RT, to favor collagen deposition. To neutralize the collagen, a buffer, composed of 6 mg mL⁻¹ of NaOH, 37 mg mL⁻¹ of NaHCO₃ and 47 mg mL⁻¹ of HEPES, was added in the syringe (350 µL of buffer for 6 mL of collagen solution). The solution was rapidly homogenized and incubated for 20 min at 37 °C. The samples were then placed in the FD support, with the patterned side up, and freeze-dried a second time (2-FD) using the same program as for the first FD (1-FD).

## 4.4. Microscopy imaging

Bright-field imaging was conducted using the DMIL LED microscope (Leica) equipped with the DMC 2900 camera (Leica). A high-resolution bright field microscope (Nanozoomer S20, Hamamatsu) was used to measure the diameter of the samples through time and to evaluate the RPE cell pigmentation. Confocal acquisitions were performed with a TCS SP8 Leica microscope, equipped with a 25 × water immersion objective (HCX IRAPO, 25 × /0.95, Leica). Scanning electron microscopy images were acquired using the JSM-IT100 InTouchScope™ Scanning Electron Microscope (SEM, JEOL Ltd.) in low-vacuum mode (30 Pa).

Microscopy images were analyzed using the Fiji ImageJ software (Java, version 1.8.0), the Icy software (Institut Pasteur and France Bioimaging), the NDP.view software (Hamamatsu Photonics) and MATLAB (R2023a).

## 4.5. Membrane characterization

### 4.5.1. Surface porosity

Scanning electron images were acquired on each side of freeze-dried membranes ($n \ge 6$ for each condition). A MATLAB program adapted from a published code was used to quantify the surface porosity [98], and is presented in the Supplementary Materials.

### 4.5.2. Inner porosity

Membranes were hydrated in PBS 48 h before imaging. For each sample ($n \ge 6$ for each condition), a z-stack of the entire thickness of the membrane was acquired with an image every 10 µm, over a 1.6*1.6 cm² area. The inner porosity was then quantified using 3D Object counter plugin of the Fiji ImageJ software. The resulting list of the volume of each pore was used to compute the inner porosity with the following formula:

$$ \text{Inner Porosity (\%)} = \frac{\text{Total volume of the pores}}{\text{Total volume of the hydrogel}} \times 100 $$

The plugin can also compute the bounding box of each pores, which was used to calculate the percentage of useful pores, defined as the percentage of pores having at least a 10 µm width, the minimal size of the capillaries in the oBRB [48]. To analyse the connectivity of the pores, the same images were processed using the Skeletonize (2D/3D) and the Analyse Skeleton (2D/3D) plugins to measure the longest shortest path (LSP) for each porous route.

### 4.5.3. In vitro stability

To assess the in vitro stability of the membrane, samples ($n = 5$) were hydrated in PBS and incubated at 37 °C for a month. The mass of the samples was measured every few days, after removing the excess of PBS with an absorbent paper, and normalized by the mass at day 1 after
hydration.

### 4.5.4. Mechanical properties of the smooth side

The Young modulus of the smooth side of hydrated collagen-coated membranes (n = 9) was studied through time by nanoindentation (PIUMA Nanoindenter, Optics11Life, Netherlands). Spherical probes with 0.5 N m⁻¹ of stiffness and a tip radius of 100 µm were used. Four samples were hydrated in PBS at 37 °C and measured on days 1, 4, 8, 15 or 22 after hydration. Nanoindentation measurements were performed in immersion in PBS at RT. Each specimen was analyzed by a matrix scan of 3 × 3 points with displacements of 150 µm between two measures. An indentation of 7 µm lasting for 5 s was conducted at each point, in indentation control mode. The probe being adhesive to the surface, the Young modulus was calculated using the JKR model, with a Poisson's ratio of 0.5, a maximum load of 80 % and a maximum load for contact point of 20 %. The data analysis was performed using the DataViewer V2 software.

### 4.5.5. Permeability

Samples were hydrated in PBS 48 h prior to analysis and were then mounted in Cell Crown inserts (CellCrown™24NX, Scaffex, Finland) in a 24 well-plate. A solution of FITC-Dextran (20 kDa, 40 kDa, 70 kDa or 150 kDa, TdB Labs) at 5 mg mL⁻¹ in PBS was placed in the bottom chamber while PBS was placed in the upper chamber (n = 5 per condition). Every half-hour for 4 h, a small volume was withdrawn from the upper chamber and diluted in PBS in a 96-well plate. Standards of FITC-Dextran were prepared in PBS for each molecular weight. The fluorescence of the standards and the samples were then measured with fluorescent plate reader (Varioskan LUX, ThermoScientific). The diffusion coefficient across the membrane for each molecular weight was calculated from the following formula (adapted from Lee et al. [99]):

$$ \ln\left(\frac{\alpha_1}{\alpha_2 - C_2(t)}\right) = D^*\beta^*(t - t_0) $$

Where $α_1 = N/V - C_2^0$; $α_2 = N/V$ and $β = (A^*V) / (h^* (V_1 + A^*h/2) * (V_2 + A^*h/2))$

With: N = quantity of FITC-Dextran in the system (in mg); $V_1$ = Volume in the upper chamber (in cm³); $V_2$ = Volume in the lower chamber (in cm³); A = the membrane total area (in cm²); h = the thickness of the membrane (in cm) and V = the total volume = $V_1+V_2+A^*h$.

The diffusion coefficient was then plotted against the logarithm of the molecular mass to obtain the exclusion limit of the membrane.

### 4.5.6. Collagen coating imaging

First, immunostainings were conducted on collagen-coated samples and controls, hydrated for 24 h in PBS at 37 °C, fixed with 10 % formalin (F0046, DiaPath, Italy) for 1 h at 4 °C and stained against collagen I. Incubation was performed overnight at 4 °C for the primary antibody, and at RT for 2 h in the dark for the secondary antibody. A list of primary and secondary antibodies with their dilutions is reported in Table 3 in the Supplementary materials. Z-stacks of the membranes were acquired by confocal microscopy. From the confocal images, the total collagen coverage of the pores was computed with the ImageJ software by comparing the surface of collagen and the total surface of pores. The presence of collagen fibrils was assessed by bi-photon induced second harmonic generation (SHG) using a confocal bi-photon microscope equipped with an 810 nm excitation source and a 405/10 nm bandpass, on non-FITC labelled collagen-coated membranes at different timepoints after hydration in PBS.

### 4.5.7. Collagen coating quantification

The collagen content was quantified in dried and hydrated

membranes using the Sircol™ Soluble collagen assay kit (S1000, BioColor Ltd, United Kingdom), (n ≥ 6 for each condition). The samples were first weighted in their dry state and then, if needed, hydrated at 37 °C in PBS. On the day of the assay, the membrane were first degraded in a solution of pullulanase (10 % v:v, E2412, Sigma-Aldrich) and dextranase (5 % v:v, D0443, Sigma-Aldrich) in PBS for 2 h at 37 °C. The quantity of collagen in the supernatants was measured as per manufacturer's instruction. Reading of the absorbance was done at 556 nm (Varioskan LUX, ThermoScientific™) and the collagen amount was computed thanks to the standard curve and normalized versus the sample initial dry weight.

## 4.6. Retinal pigment epithelium (RPE) cell seeding

### 4.6.1. RPE cells differentiation

Human iPSC (PC56c2) obtained from Phenocell (France) was differentiated into RPE cells using a published protocol [100]. Once differentiated, the cells were frozen at passage 1 in CryoStor® CS10 cell cryopreservation media (C2874, Sigma-Aldrich) for long term storage.

### 4.6.2. RPE cell culture

RPE cells were thawed two weeks prior any experiment, in a T25 flask coated with L7™ hPSC matrix as per manufacturer's instruction (FP-5020, Lonza), with 4 million cells per flask. The cells were incubated at 37 °C.5 % CO₂ in RPE media: DMEM (high glucose, GlutaMAX™ Supplement, pyruvate; 31966047; Gibco, ThermoFischer) + 4 % Knockout serum (KSR, 10828010, Gibco, ThermoFischer) + 1X non-essential amino-acids (MEM NEAA 100X, 11140035, ThermoFischer) + 1X Antibiotic-Antimycotic (15240062, Gibco, Thermo-Fischer) and 0.1 % 2-mercaptoethanol (50 mM, 31350010, Gibco, ThermoFischer).

### 4.6.3. RPE cell seeding on the smooth side of collagen-coated membranes

Prior to cell seedings, the membranes were exposed to UV-C light for 2 h for sterilization, and then rehydrated in a laminin 521 solution (CTG, Biolamina) at 20 µg mL⁻¹ (diluted in DPBS with calcium and Magnesium (14040091, Gibco, ThermoFischer)) for 2 h at 37 °C. Each membrane was then mounted in a 24-well plate Cell Crown, with the smooth side placed on top.

Cells were detached using TrypLE™ enzyme (12604013; Gibco, ThermoFischer), and gathered in a Falcon tube for centrifugation (300 g for 5 min). The cells were resuspended, counted, and centrifuged a second time. RPE media was added to obtain a cellular concentration of 4 million mL⁻¹. 100 µL of cell suspension was then added on each membrane. After 1 h of incubation at 37 °C.5 % CO₂, 1 mL of RPE media is added in each well. The media was changed on the next day, and twice a week, for the duration of the experiment. The removed media was frozen at -20 °C in Eppendorf tubes (1.5 mL) for further analysis.

## 4.7. Pre-vascularization

Human primary retinal microvascular endothelial cells (HRMVECs, ACBRI 181) and human primary retinal pericytes (HRPCs, ACBRI 183) were purchased from Cell Systems (USA) at passage 3 and amplified per manufacturer's instructions. Cells were frozen at passage 7 and passage 5 for the ECs and PCs respectively in Cryostor media. Cells were thawed three days prior to any experiments in T75 flasks (1 million cells per flask) coated with a 20 µg mL⁻¹ collagen solution (PBS for 2 h at RT, FS22005, Collagen solutions, UK). Both ECs and PCs were incubated at 37 °C.5 % CO₂ in EGM complete media (CC-3162, Lonza, Switzerland). The media was changed the next day.

On the day of the seeding, the membranes were exposed to UV-C light for 2 h for sterilization, rehydrated in PBS for 2 h at 37 °C and then mounted in a 24-well plate Cell Crown, with the rough patterned side placed on top. Cells were detached from the T75 flask with TrypLE solution (3 min incubation at 37 °C), centrifuged at 300 g for 5 min, resuspended in EGM and counted. A second centrifuge was performed to
obtain a solution at a EC concentration of 3.5 million·mL⁻¹ in EGM media. In the case of the experiments with pericytes, PCs were added to the same solution at a final PC concentration of 700,000 cells·mL⁻¹ (1:5 PC:EC ratio). 100 µL of cellular solution was added per membrane. After 1 h incubation 37 °C·5 % CO₂, 1 mL of EGM media was added to each sample and placed back in the incubator. The media was changed on the next day, and twice a week, for the duration of the experiment. The removed media was frozen at -20 °C in Eppendorf tubes (1.5 mL) for further analysis.

## 4.8. oBRB modelling by tri-culture seeding

To model the oBRB in tri-culture, RPE were first seeded on the smooth side of the membranes as described above (Section 4.6.3). Additional membranes were coated with laminin and incubated in RPE media at 37 °C, without RPE cells, to serve as controls for the EC + PC seeding.

After 8 days of RPE culture, the EC and the PC were added by flipping the Cell Crown in each well and following the method described above (section 4.7). Once the EC/PC were added, the media was switched to co-culture media composed of 2/3 RPE media +1/3 EGM media +100 µg mL⁻¹ L-ascorbic acid (A4403, Sigma-Aldrich) + 10 ng mL⁻¹ vascular endothelial growth factor (Human VEGF-165, Animal-Free Recombinant Protein, PeproTech®, AF-100-20, Gibco, ThermoFischer). The media was changed on the next day, and twice a week, for the duration of the experiment. The removed media was frozen at -20 °C in Eppendorf tubes (1.5 mL) for further analysis.

## 4.9. Cell culture analysis

### 4.9.1. Cell viability

To assess cellular viability, a cytotoxicity assay, based on the measurement of lactate dehydrogenase (LDH) activity, was conducted (Cytotoxicity Detection Kit PLUS, 744.934.001, Roche, Switzerland) according to the manufacturer's instructions. Three samples were incubated in media supplemented with a lysis solution (10 % v:v) for 15 min, to obtain positive controls corresponding to the maximal LDH release. Media was used as blank. Supernatants were gathered and incubated in the reaction solution for 15 min. Readings were performed in absorbance at 490 nm and 690 nm with a microplate reader (Varioskan LUX, ThermoScientific™). The cytotoxicity rate was determined from the absorbance readings as follows:

$$ \text{Cytotoxicity} = \frac{A_{\text{sample}} - A_{\text{media}}}{A_{\text{CTL+}} - A_{\text{media}}} \times 100 $$

To assess cellular mortality of RPE or endothelial cells seeded on the membranes, samples were rinsed once with DPBS and incubated for 1 h at 37 °C·5 % CO₂ in a staining solution containing Hoechst 33342 (10 µg mL⁻¹, H3570, ThermoFischer) and Ethidium Homodimer-I (4 µM, 46043, Sigma) in media (either RPE media or EGM-2). After incubation, the samples were rinsed in DPBS and imaged with a confocal microscope.

### 4.9.2. Staining

At various timepoints, samples were fixed in 10 % formalin (F0046, DiaPath, Italy) for 1 h at 4 °C under agitation. Samples were rinsed with PBS (3 washs of 15 min) and permeabilized using a 0.1 % Triton solution in PBS for 1 h at RT under agitation followed by 3 PBS washs of 15 min. For immunostainings, the primary antibody solution in PBS was incubated at 4 °C overnight. After rinsing with PBS, the secondary antibody and DAPI (1:1,000, 4',6-diamidino-2-phenylindole dihydrochloride, 62247, Thermo Scientific™) were incubated for 2 h at RT. For actin staining, the samples were fixed, permeabilized and incubated with DAPI (1:1000) and Phalloidin-Tetramethylrhodamine B isothiocyanate (1:500, P1951, Sigma-Aldrich) for 2 h at RT in the dark. Samples were imaged by confocal microscopy. The list of the primary and secondary

antibodies is presented in the Supplementary Materials (Table 3) along with the corresponding dilutions.

### 4.9.3. ELISA and multiplex assays

ELISA assays were performed to measure the concentration of either VEGF (RAB0508, Sigma-Aldrich; $n \ge 3$ for each timepoint) or pigment epithelium growth factor (PEDF, CYT420, Sigma-Aldrich; $n = 6$ for each timepoint) in the culture supernatant. Moreover, multiplex assays were performed to measure angiogenic, inflammatory markers and metalloproteinases in the supernatants (FCSTM03-08, FCSTM07-04 and LXSAHM-07; R&D systems, Bio-Techne, USA). The samples were thawed overnight at 4 °C and the assays were conducted per manufacturer's instructions ($n \ge 4$ for each timepoint and for each condition). The experimental results were obtained either with a plate reader (Varioskan LUX, ThermoScientific™) for ELISA assays or suspension array system (BioPlex-200, Bio-Rad, USA). The multiplex data were analyzed with the Bio-Plex Manager software (Bio-Rad). For all ELISA and multiplex experiments, media was incubated in the empty plate well as control to account for growth factor degradation at 37 °C. The control media was analyzed along all samples and the resulting value was subtracted from all data points.

### 4.9.4. Nitric oxide assay

Quantification of nitric oxide was done by analysis of cell culture supernatant using a NO assay kit (ab272517, abcam, UK), as per manufacturer's instruction. Reading of the optical densities (Varioskan LUX, ThermoScientific™) was done at 540 nm and NO fold change was calculated by normalization of absorbance values compared to the controls.

### 4.9.5. Transepithelial electrical resistance (TEER) measurements

Measurements of the TEER was conducted on seeded scaffolds at different timepoints. The samples were placed in Cell-Crown with 200 µL media in the upper chamber and 600 µL in the lower chamber. The Millicell® ERS 3.0 Digital Voltohmometer was used following manufacturer's instructions. The measurements were conducted on $n = 10$ samples for the RPE monoculture and $n \ge 4$ for the tri-culture study, with 3 unseeded membranes as controls.

## 4.10. Statistical analysis

All the statistical analysis were performed using the Prism 10 software (GraphPad, USA). For each analysis, a normality test was performed to assess whether the data followed a normal or lognormal distribution. If the test revealed a lognormal distribution, a logarithm transformation was applied on the data prior to any statistical analysis. For each figure, the type of statistical test is indicated in the figure caption, results are shown as mean±standard deviation (SD).

## CRediT authorship contribution statement

**Chloé Dujardin:** Writing – original draft, Validation, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Walter Habeler:** Project administration, Methodology, Conceptualization. **Paola Aprile:** Methodology, Investigation, Conceptualization. **Alessandra Dellaquila:** Methodology. **Christelle Monville:** Writing – review & editing, Resources, Project administration, Funding acquisition, Conceptualization. **Didier Letourneur:** Writing – review & editing, Supervision, Resources, Project administration, Funding acquisition, Conceptualization. **Teresa Simon-Yarza:** Writing – review & editing, Supervision, Resources, Project administration, Funding acquisition, Conceptualization.

## Declaration of competing interest

The authors declare that they have no known competing financial
interests or personal relationship that could have appeared to influence
the work reported in this paper.

Acknowledgements

This project has received funding from the French 'Fondation pour la
Recherche Médicale' (Project: REP202210016100). I-Stem is part of the
Biotherapies Institute for Rare Diseases supported by the Association
Française contre les Myopathies-Téléthon. This work was in part funded
by the H2020-NMBP-HUBS-2018 TBMED project: Grant Agreement No.
814439. C. D. was funded by the AMX program of the Ecole Poly-
technique and the French ministry for Higher Education and Research.
The authors thank Dr. Olivier Goureau for his support and expertise
during this project. The authors gratefully acknowledge the Photonic
Imaging Platform IMA'CRI (Centre de recherche sur l'Inflammation -
CRI, INSERM) and Dr. Samira Benadda for their support and assistance
in this work. They also acknowledge the ImagoSeine core facility of the
Institut Jacques Monod, member of the France BioImaging infrastruc-
ture (ANR-10-INBS-04) and GIS-IBiSA; and the support of the Region Ile-
de-France (Sesame).

**Appendix A. Supplementary data**

Supplementary data to this article can be found online at https://doi.
org/10.1016/j.biomaterials.2025.123357.

Data availability

Data will be made available on request.

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