# Biohybrid microstructured hydrogels obtained via in situ extracellular matrix deposition and decellularization using supercritical CO2

Vanessa Morais Lima, Albane Carré, Emmanuelle Poque, Maria-Dimitra Chiotelli, Natan Wiele, Christelle Harscoat-Schiavo, Raphaëlle Savoire, Teresa Simon-Yarza

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Vanessa Morais Lima, Albane Carré, Emmanuelle Poque, Maria-Dimitra Chiotelli, Natan Wiele, et al.. Biohybrid microstructured hydrogels obtained via in situ extracellular matrix deposition and decellularization using supercritical CO2. Biofabrication, 2025, 17 (3), pp.035030. 10.1088/1758-5090/adebb4. hal-05172652

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# Biohybrid microstructured hydrogels obtained via *in situ* extracellular matrix deposition and decellularization using supercritical CO₂

To cite this article: Vanessa Morais Lima *et al* 2025 *Biofabrication* 17 035030

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PAPER

# Biohybrid microstructured hydrogels obtained via *in situ* extracellular matrix deposition and decellularization using supercritical CO₂

Vanessa Morais Lima¹ ID, Albane Carré¹ ID, Emmanuelle Poque² ID, Maria-Dimitra Chiotelli¹ ID, Natan Wiele² ID, Christelle Harscoat-Schiavo² ID, Raphaëlle Savoire² ID and Teresa Simon-Yarza¹,* ID

¹ Université Paris Cité, Université Sorbonne Paris Nord, LVTS, Inserm U 1148 Paris, France

² Université de Bordeaux, CNRS, Bordeaux INP, CBMN, UMR 5248, F-33600 Pessac, France

* Author to whom any correspondence should be addressed.

E-mail: teresa.simon-yarza@inserm.fr and vanessa.morais-lima@inserm.fr

**Keywords:** extracellular matrix, biohybrid material, scaffold, polysaccharide hydrogel, supercritical CO₂, decellularization

Supplementary material for this article is available online

## Abstract

In recent decades, our understanding of biomaterials has shifted from seeing them simply as physical supports for cells or drug delivery platforms to recognizing their active and dynamic role in tissue repair, guided by their physicochemical, mechanical, and biological properties. Biologically derived materials such as the decellularized extracellular matrix (dECM) offer the advantage of replicating the biomolecular cellular environment and have been proposed for tissue regeneration. However, their use as scaffolds is hindered by poor mechanical properties and limited tunability of physical features. Herein, we fabricated a bioinspired hybrid hydrogel by integrating a chemically cross-linked microporous polysaccharide scaffold with native ECM directly secreted by cells. First, the scaffold synthesis and culture conditions were optimized to enhance ECM deposition by fibroblasts. To obtain an acellular scaffold, decellularization using supercritical CO₂ was performed and compared to a conventional method, demonstrating its superiority in ensuring efficient decellularization while preserving an enriched ECM lining the surface of the pores and preventing scaffold damage. The biohybrid hydrogel was characterized by a very low amount of DNA (<5 ng DNA mg⁻¹) and a network of highly interconnected pores covered by an abundant ECM including collagen I, collagen IV, fibronectin, elastin and laminin. This work presents a new versatile strategy that can be adapted to various tissues to engineer biomimetic microstructured materials overcoming the limitations associated with polymer-based and dECM-based strategies when used independently.

## 1. Introduction

In the past decades the design of biomaterials, as physical supports for cells or as drug delivery platforms, has evolved towards an active and dynamic vision. Currently, the development of biomaterials aims at mimicking the extracellular matrix (ECM) that is known to play a major role in tissue repair [1] and sets the basis to establish the ideal characteristics of the scaffolds for tissue engineering and regenerative medicine. By controlling their physical, chemical and mechanical properties, it is possible to enhance their contribution to reparative process, through

the promotion of cell proliferation, differentiation, and migration, immunomodulatory responses, and angiogenesis, amongst others [2, 3].

Numerous strategies to develop biomaterials to mimic the ECM are reported. The most common ones consist in the incorporation or grafting of one or several components of the ECM into synthetic materials, or directly using them to form the constructs. Faced to these simplistic approaches, there are studies in which the whole ECM is used, in general after subjecting it to decellularization processes [4]. These strategies present some advantages but also limitations. Working with only a few components of
the ECM facilitates the preparation and characterization of the scaffolds, but fails to recapitulate the properties of the native ECM. Besides, the use of whole decellularized ECM (dECM) is a more comprehensive strategy that presents major concerns too. The limited number of human donors leads to the use of animal dECM, with associated immunological risks and also results in differences in the final matrix properties compared to the native one due to inter-species variability. After decellularization, the physical-chemical and mechanical properties of the matrix are often compromised making manipulation complicated and overall, providing cells with signals that can induce disturbed cell behavior [4]. Moreover, complete revascularization and recellularization of whole decellularized organs remain unreachable [5]. Thus, dECM is frequently processed as a powder that can be used as a gel or even as an ink for 3D bioprinting. In this case, protein cross-linking can overcome the lack of mechanical integrity [4].

The possibility to obtain cell-derived dECM keeping its 3D organization has been explored in several ways [6]. In particular, coating materials prepared in the laboratory with ECM directly secreted by cells has opened new perspectives for the obtainment of biomimetic biomaterials [7, 8]. This strategy, called cell-deposited matrix (CDM), has been mainly used to study the role of cells and biochemical stimuli on the synthesis of ECM, as well as the impact of this CDM on the behavior of cells seeded on this biohybrid material [9–11]. Nevertheless, the use of CDM approach to develop hybrid material-based therapies is an innovative path barely explored until now, but with great potential for tissue engineering applications [8]. The advantages of this strategy are derived from the possibility of engineering the 3D scaffolds, by controlling physical parameters, such as the morphology, internal microarchitecture and mechanical properties, and by regulating the chemical composition through bioengineering strategies like culturing ECM secreting cells. In this context, fibroblasts are the main cells responsible for the production of ECM in normal tissue repair [12]. On the other hand, hydrogels are one of the most commonly used materials for tissue engineering, due to their physical, chemical and mechanical properties mimicking those of the ECM. Applying CDM to porous hydrogels is a promising approach for advancing the manufacturing of materials that replicate the chemical, physical and mechanical properties of native ECM.

In this work, our aim was to obtain a biomimetic scaffold by promoting ECM in situ deposition on a chemically crosslinked porous hydrogel. Polysaccharide-based scaffolds made of pullulan and dextran that mimic the glycosaminoglycans of the ECM, and previously reported for several tissue engineering purposes [13–20], were formulated to

support fibroblast 3D cell culture and ECM deposition. Two methods to incorporate gelatin or collagen type I into the composition were developed (figure 1(I)). A first freeze-drying (FD) step was applied for pore formation. After collagen or gelatin coating, a second freeze-drying step (2FD) allowed the production of off-the-shelf porous scaffolds. In total, five formulations were compared in terms of physical and mechanical properties. Then, their ability to support fibroblasts cell culture up to 21 days was evaluated based on several parameters, including cell proliferation and phenotype within the pores, metabolic activity and by assessing the deposition and distribution of several proteins of the ECM (figure 1(II)). To remove cell components while preserving the ECM, a conventional method based on detergent and enzymatic treatment was compared to our protocol using supercritical carbon dioxide (scCO₂) (figure 1(III)), a fluid domain of CO₂ that allows excellent penetration to remove cell debris with reduced physical damage of the structures.

## 2. Methods

### 2.1. Preparation of the scaffolds

Pullulan-dextran (PuD) hydrogels were prepared following a previously reported protocol by mixing a pullulan (MW 200 kDa; Hayashibara Co.) and dextran (MW 500 kDa; Pharmacosmos) (75:25 w/w) in deionized water at a final concentration of 30% (w/w) and in the presence of sodium chloride (NaCl) (21% w/w of the polymer solution) [14, 15]. For confocal analysis purpose, 1% fluorescein isothiocyanate-labeled dextran 500 kDa (TdB Consultancy, Uppsala, Sweden) was added to the polymer solutions before crosslinking. PuDGel was prepared by adding gelatin 5% (w/w) directly to the polysaccharide solution followed by chemical crosslinking using 3% (w/v) sodium trimetaphosphate (STMP, Sigma) under alkaline conditions (1 M NaOH). The hydrogels (PuD and PuDGel) were casted between two glass plates separated with a 1 mm spacer before incubation at 50 °C for 20 min. The resulting non-porous gels were punched into 5 mm diameter scaffolds (16.5 mm diameter in the case of scaffolds intended for mechanical testing) and bathed in 10X PBS to lower the pH to 7.4 then in distilled water overnight to remove the excess reagents and salts. PuD and PuDGel were freeze-dried to obtain **PuD-FD** and **PuDGel-FD** respectively. Conductivity of the scaffolds prior freeze-drying was measured to assure that the ionic strength remained the same between the different batches using conductivity meter 145 A+ (thermo Scientific Orion, USA). Therefore, the scaffolds were freeze-dried on a petri-dish with enough volume of NaCl 0.025% (w/v) to cover all the hydrogels following an established protocol [21]. After a controlled freezing
<figure><img src="image_6.png" /></figure>

step under atmospheric pressure, primary drying occurred at -20 °C at 0.010 mbar followed by a second drying at 30 °C. The freeze-drying system consisted of a freeze-dryer (Cryotec Pilote, Lunerl Viel, France) connected to a vacuum pump (E2M28, BOC Edwards, Crawley, UK).

PuD-FD were rehydrated in 0.1X PBS then freeze-
dried a second time to obtain PuD-2FD scaffolds.

PuDGel-2FD and PuDColl-2FD were prepared
by rehydrating PuD-FD scaffolds in a 1% (w/v)

gelatin solution (type B, bovine, Sigma) or 1 mg ml⁻¹
collagen solution (type 1, bovine, in HCl 0.01 M,
Collagen Solutions) via syringe pumping [14]. In
brief, the scaffolds were placed inside a syringe with
either gelatin or collagen solution. Then, the piston
of the syringe was placed, and the tip was blocked
with a locker. Coating was achieved by applying neg-
ative pressure. The proteins were left to deposit for
2 h at room temperature (RT) before being washed,
in PBS for collagen neutralization then in 0.1X PBS
for fibril formation, and in 0.1X PBS for the gelatin coated scaffolds. Samples of pristine PuD-FD, PuD-FD coated by gelatin and PuD-FD coated by collagen, were lyophilized for a second time leading to the double freeze-dried PuD gelatin PuDGel-2FD and the double freeze-dried PuD collagen I PuDColl-2FD.

## 2.2. Characterization of the scaffolds

**Swelling behavior.** Scaffolds kinetics of hydration were determined by monitoring their mass swelling ratio over time. All scaffolds were weighed in a dry state ($W_{dry}$) before being hydrated 48 h in PBS under mid rotation at 37 °C. Hydrogels weights were monitored during the hydration ($W_{swollen}$) at different time points (2 min, 5 min, 10 min, 30 min, 1 h, 24 h and 48 h). Before each measurement, the excess of PBS was removed using absorbent paper. Mass swelling ratio was calculated using the following formula:

$$ \text{Mass Swelling ratio} = \frac{W_{\text{swollen}} - W_{\text{dry}}}{W_{\text{dry}}} $$

where $W_{swollen}$ is the mass of the hydrogel in the swollen state and $W_{dry}$ is the mass in the dry state respectively. For each condition nine samples were evaluated, and results were expressed as mean values ± SD.

**In vitro enzymatic degradation.** After 48 h of hydration in PBS, the excess of solvent was removed using absorbent paper and scaffolds were weighed ($W_0$) and incubated in an enzymatic solution of 10% (v/v) pullulanase (Sigma, E2412) and 5% (v/v) dextranase (Sigma, D0443) in PBS for 48 h at 37 °C. Scaffolds were weighed ($W_d$) at different time points until no remaining mass was observed. For each time point, the percentage of residual mass was calculated according to the following formula:

$$ \text{Residual mass (\%)} = \frac{W_d}{W_0} \times 100 $$

where $W_d$ is the mass of the scaffold at each degradation time whereas $W_0$ is the initial mass. For each condition, a total of nine samples were evaluated and results were expressed as mean values ± SD.

**Porosity.** Surface and inner structuration of the dried polysaccharide-based scaffolds were visualized using a TeneoVS scanning electron microscope (SEM), with a BSE detector under low vacuum mode (40 Pa) at a 20 kV acceleration voltage (ImagoSeine core facility of the Institut Jacques Monod).

**Mechanical testing.** Shear storage modulus G' was measured with the ElastoSens™ Bio lab equipment (Rheolution). In order to fit the device sample holder,

16.5 mm diameter and 1 mm height scaffolds were prepared following the steps described previously. These scaffolds were then hydrated in PBS for 2 d at RT or at physiological temperature (37 °C) to assess the impact of cell culture conditions at physiological temperature. For the measurements, the scaffolds were placed in contact with the silicon membrane of the sample holder. Shear storage modulus (G') was measured using a sequence of 1 min with a low-amplitude vibrating stimulation every 20 s.

Similar measurements were performed on hydrogels which were cultured with 3T3 fibroblast cells, and on culture control hydrogels. Therefore, Balb/c 3T3 fibroblasts were cultured for 7 d within the hydrogels under physiological conditions (5% CO₂, 37 °C) prior to measurements. Control scaffolds without cells were cultured under the same conditions. For each condition, nine samples were evaluated and results were expressed as mean values ± SD.

## 2.3. In vitro studies

**Fibroblasts culture.** Mouse Balb/c 3T3 clone A31 fibroblasts (ATCC, CCL-163™) were cultured in T75 flasks (Corning®) using high glucose Dulbecco's Modified Eagle Medium (4.5 g l⁻¹ glucose), supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin (Sigma). Culture medium was changed every two days and cell splitting was done at 80% confluence using trypsin solution (Gibco).

**Seeding of fibroblasts in the scaffolds.** All scaffolds were treated with UV-C light for 2 h before cell seeding. Homogeneous cell seeding of Balb/c 3T3 was achieved at a cell concentration of 1 × 10⁶ or 5 × 10⁵ cells per scaffold. The freeze-dried scaffolds at the dry state were placed in a 10 ml syringe and hydrated with cell solution by negative pumping ten times slowly. Cell-loaded hydrogels were incubated in 24-well plates at 37 °C 5% CO₂ for 1 h before the addition of 1 ml of culture medium as previously described. For larger scaffolds used in mechanical testing at day 7, cells were drop-seeded with 300 µl of 10 000 cells/µl solution, on top of the scaffold. Samples were cultured up to 21 d changing medium every day, then kept at -20 °C until further use.

**Cell metabolic activity.** Metabolic activity was quantified at several time points (1, 4, 7, 10, and 14 d) using the in vitro Toxicology Assay Kit (Resazurin-based, TOX8, Sigma-Aldrich). Cell-loaded scaffolds were incubated with 0.5 ml of 10% resazurin in fresh culture medium for 3 h at 37 °C, 5% CO₂. 100 µl of the resulting solution was transferred to a 96-well plate and fluorescence was measured using Infinite M200 Pro, TECAN® plate microplate reader
(560Ex/590Em). All samples were run in triplicate in three independent experiments.

**Staining and confocal imaging.** At 7, 14 and 21 d of culture, cell-loaded or decellularized hydrogels were rinsed with PBS, fixed with paraformaldehyde 4% for 1 h at RT, rinsed three times with PBS, permeabilized with 0.1% (v/v) Triton X100 for 45 min at RT followed by overnight incubation of primary antibodies for ECM proteins (Collagen IV ab6586, Collagen I ab270993, Fibronectin ab268020, Laminin PA1-16 730, and Elastin bs-1756 R, 1:100) at 4 °C. After rinsing 3 times with PBS, the samples were incubated with Phalloidin Alexa Fluor 633 (1:500, Invitrogen A22284), DAPI 5 mg ml⁻¹ (1:1000), and secondary antibody Goat anti-rabbit Alexa Fluor 555 (1:300, Abcam ab150078) 0.5 mg ml⁻¹ for 2 h at RT. Images and 3D stacks were obtained using an inverted Zeiss LSM 780 confocal microscope fitted with 10× objectives (CRI—U1149 Imaging facility) over up to 300 µm in the scaffold depth. Images were reconstructed and analyzed using Fiji and FluoRender. Three independent samples (biological replicates) with three replicate samples (three different scaffolds, technical replicates) were used per condition. 3D stacks images analysis was performed using Fiji. To assess cell adhesion, we quantified the number of pores filled by cells relative to the total number of pores in four fields per z-stack across ten different stacks, with data collected from at least three independent experiments. ECM protein quantification was performed by calculating the ECM protein-covered area relative to the total area per z-stack. The z-stack image acquisition was carried out using confocal microscopy, capturing a total depth of approximately 150 µm per hydrogel, with optical sections acquired at 20 µm intervals. Image analysis was conducted across two independent experiments, with a minimum of two hydrogel constructs analyzed per experimental group.

## 2.4. Decellularization

**Supercritical CO₂ apparatus.** The experimental setup is schematized in figure S1. The apparatus consists in a quick-opening 50 ml stainless steel reactor, 3 cm internal diameter (ROR reactor, Autoclave-Maxitech, France) placed in an oven (Binder GmbH, Tuttlingen, Germany). Inside the reactor, the assembly is made of a 1.55 cm length stirring bar, a 1.5 cm high glass spacer with a metallic grid overtopped by a paper filter. The CO₂ is fed into the reactor by a syringe pump (Teledyne ISCO 260D, Serlabo, France). The pressure and the solution temperature are monitored by pressure sensor (400 bar, ASCO, Lucé, France) and a K-type thermocouple (1 °C accuracy, Watlow, France), respectively. A heated metering valve (Autoclave Engineer, Wexford, Ireland) allows controlled depressurization rate. A second reactor

with the same assembly of stirring bar, glass spacer, grid and paper filter but without CO₂ introduction was also placed in the oven.

**Supercritical CO₂ procedure.** Decellularization experiments were performed on collagen coated hydrogel after 14 d of cell culture. Three samples to decellularize were placed in the reactor, on top of the paper filter, and immersed 15 ml 0.5% SDS. Stirring was set at 500 rpm and temperature regulated at 45 °C. The reactor was pressurized by introducing scCO₂ until 170 bar pressure was reached. Then, the system was left in these conditions (45 °C, 170 bar) for 90 min. Finally, pressure was released and treated samples could be recovered. Using the second reactor, all tests using scCO₂ had a control called 'Air + 0.5%SDS' with the same conditions (solvent composition, stirring, temperature, duration) but without scCO₂. 3 samples were used for each control experiment. After experiments, the samples were washed three times for 1 h with PBS solution at RT under agitation, then stored at -20 °C until further analysis.

**Standard decellularization protocol.** Standard decellularization was performed on collagen coated hydrogel after 14 d of cell culture following the protocol described by Silva and colleagues [22]. The process involved sequential incubation steps, with approximately 60 mg of hydrogel per 1 ml of reagent volume, under agitation at RT unless otherwise specified. Hydrogels were thawed and washed with PBS. Samples were incubated for 18 h in a hypotonic buffer (10 mM Tris-HCl, 0.1% EDTA, pH 7.8) then washed three times in PBS, with each wash lasting 1 h. Subsequent incubation was performed for 24 h in a detergent solution containing 0.2% SDS and 10 mM Tris-HCl (pH 7.8). The samples were washed three times using a hypotonic wash buffer (10 mM Tris-HCl, pH 7.8), with each wash lasting 20 min. The samples were incubated for 3 h at 37 °C in a DNase solution (50 U ml⁻¹ DNase I in 10 mM Tris-HCl, pH 7.8). Residual detergent and DNase were removed by washing the samples three times in PBS (20 min per wash). Finally, the decellularized samples were frozen at -20 °C for further analysis.

## 3. DNA quantification

DNA quantification was performed on collagen coated hydrogel after 14 d of cell culture decellularized or not. For DNA quantification, half of the scaffold was weighed and degraded in 10% (v/v) pullulanase + 5% (v/v) dextranase in PBS for 2 h at 37 °C under agitation. For cell digestion, a solution containing 1 mg ml⁻¹ proteinase K, 75 mM NaCl, 25 mM EDTA, was added and digestion was allowed to carry on for 2.5 h at 55 °C. The following steps were
done at RT. Saturated NaCl (6 M) was added to stop
reaction. Tubes were vortexed for 30 sec then centri-
fuged for 5 min at 15 000 rpm. The supernatant was
transferred into a new 1.5 ml Eppendorf tube with
equal volume of isopropanol, vortexed for 30 sec then
centrifuged at 15 000 rpm for 15 min. The super-
natant was discarded and the pellet was washed twice
by resuspension in 70% ethanol, vortexed 30 s and
centrifugated for 15 min at 15 000 rpm. After dry-
ing for 2 h in the fume hood, the pellet was resuspen-
ded in 40 µl pure water for DNA quantification using
the Quant-iT PicoGreen dsDNA assay kit (Invitrogen,
ThermoFisher scientific, P7589) following the sup-
plier protocol. Two replicate samples from three inde-
pendent experiments were used per condition.

4. Statistics

Data were expressed as mean ± standard deviation. Statistical significance was calculated using one-way or two-way ANOVA followed by Tukey *post hoc* test with multiple comparisons using GraphPad® Prism software. Values of *p* < 0.05 were considered statistically significant.

5. Results

**5.1. Material fabrication and characterization**

Five different formulations composed of pullulan and
dextran crosslinked with STMP were obtained as
dried scaffolds following a single (FD) or a second
freeze-drying (2FD) step (figure 1 (I). These formu-
lations have been identified as **PuD-FD** (no coating,
single FD), **PuDGel-FD** (gelatin added to the initial
polymers solution before cross-linking, single FD),
**PuD-2FD** (no coating, double FD), **PuDGel-2FD**
(gelatin coating after the first freeze-drying, double
FD), **PuDColl-2FD** (collagen coating after the first
freeze-drying, double FD).

Scanning electron microscopy observations of the
surfaces and cross-sections of the scaffolds in the dry
state revealed differences in porosity (figure 2(A)).
In all cases, one surface appeared to be less por-
ous, with a complete absence of visible pores in the
case of PuDGel-FD. The other face presented pores
that, in the case of PuDColl-2FD, appeared to be
less numerous but larger (>250 µm width) than for
the other materials. Regarding the internal poros-
ity, cross-section images confirmed the presence of a
homogeneous interconnected porous network within
the entire thickness of the PUD matrix, regardless of
the composition. In the case of PuDGel-FD, the size of
the pores near the non-porous surface appeared smal-
ler (<200 µm), whereas near the other face they were
less numerous but larger (>300 µm).

Differences between single FD and double FD
were also found in the swelling kinetics, with slower

swellings in the 2FD formulations, that was more pro-
nounced when proteins were incorporated via this
method. After 5 min, the swelling of PuD-FD and
PuDGel-FD almost reached a plateau with a swelling
ratio of around 12. At that same time, PuD-2FD swell-
ing ratio was near 11, and PuDGel-2FD and PuDColl-
2FD showed an average ratio of 9 and 10, respect-
ively. The presence of the proteins also caused a delay
on the enzymatic degradation using pullulanase and
dextranase (figure 2(C)). Complete degradation of
PuD-FD, PuD-2FD and PuDGel-FD was observed
after 100 min, 140 min and 150 min respectively,
whereas PuDGel-2FD and PuDColl-2FD degraded
after 220 min and 300 min. Thus, delay in the degrad-
ation kinetics caused by the incorporation of proteins
was greater than the one caused by a second freeze-
drying step.

Average shear storage modulus (G') of all the scaffolds remained between 1.7 and 3.5 kPa (figure 2(d)), within the range of those reported for several soft tissues, including brain, lung, kidney cortex, liver, thyroid, pancreas and cardiac muscle [23]. The impact of the temperature (25 °C vs 37 °C) was greater in the single freeze-dried scaffolds, with significantly lower values at 37 °C (average G' of 3.2 kPa vs 2.2 kPa for PuD-FD and 2.2 kPa vs 1.7 kPa for PuDGel-FD). In the absence of cells, the incorporation of proteins was associated to a decrease of G', both at 25 °C and 37 °C. Unexpectedly, the addition of a second freeze-drying step led a G' that was independent of the temperature in the case of PuD-2FD and PuDColl-2FD, and a significant increase for PuDGel-2FD scaffolds at 37 °C. We also checked possible differences in the G' due to the presence of cells and the culture medium within the scaffolds. When cultured with cells (1 × 10⁶ fibroblasts/samples; 7 d of culture), we observed significant differences only in the samples containing gelatin. Indeed, both conditions PuDGel-FD and PuDGel-2FD showed a significant increase of G', suggesting a remodeling of these scaffolds by cells.

**5.2. 3D culture of fibroblasts in the scaffolds**

To further investigate the effect of cells, fibroblasts
were seeded homogenously within the entire thick-
ness of the five different scaffolds via negative pressure
induced seeding, as previously described [13, 14]. Cell
behavior was evaluated through cell metabolic activ-
ity over 14 d culture. When cultured on scaffolds
without protein (PuD-FD and PuD-2FD), the meta-
bolic activity decreased progressively as soon as from
day 3 (figure 3(A)). This reduced metabolic activ-
ity was lower after 7 d, then values remained stable
until day 14. A similar trend was observed for scaf-
folds in which gelatin was added before cross-linking
(PuDGel-FD), with a reduction until day 10 and then
a slight increase at day 14. In contrast, when the pro-
tein was deposited on the surface of the pores after a
<figure><img src="image_7.png" /></figure>

first freeze-drying step (PuDGel-2FD and PuDColl-2FD), the activity of the cells was maintained and increased over the 14 d, both for gelatin and collagen coating. The obtained results motivated the exclusion of uncoated and PuDGel-FD formulations for ECM deposition studies, since the organization in spheres and the reduced metabolic activity could result in low ECM deposition on the surface of the pores. Thus, only double-freeze dried samples were retained for *in situ* ECM deposition by fibroblasts.

The next step was to evaluate the capacity of the double FD scaffolds, PuDGel-2FD and PuDColl-2FD, to promote cell penetration and ECM deposition within the interconnected pores of the scaffolds. To study cell adhesion and proliferation into the hydrogels throughout time, staining of actin cytoskeleton of the cells was done using phalloidin. Confocal microscopy imaging of cells cultured in PuD-2FD, PuDGel-2FD, and PuDColl-2FD was done at different time points up to 21 d of culture (figures 3(B) and S2).
<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /><figcaption>**Figure 3.** Fibroblasts 3D culture in the scaffolds. (A). Metabolic activity as assessed by Resazurin assay. (B). Cell morphology within the pores of the scaffolds assessed by actin and nuclei staining. (C). Quantification of the % of pores loaded with cells based on image analysis. Scale bar = 300 µm. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001, \*\*\*\*p < 0.0001.</figcaption></figure>

Regarding cell morphology and organization, PuD-2FD uncoated scaffolds led to cell assembly in aggregates. After 7 d, these cells clusters had rounded and discoidal shapes, and after 14 d and 21 d cells had

proliferated. In the case of coated scaffolds, either with gelatin or collagen, cells started by adhering on the edges of the pores at day 7, and then proliferated until complete lining of the pores. In the case
<figure><img src="image_11.png" /></figure>

of gelatin, we could observe after 21 d more empty pores compared to collagen (PuDColl-2FD). In most of the pores containing cells, the whole pore volume was filled with cells. Based on confocal images, the percentage of pores loaded with cells at 7 d, 14 d and 21 d was quantified (figure 3(C)). Results showed that for all the time points, the percentage of pores loaded with cells was significantly higher for PuDColl-2FD scaffolds than for others. For all the times under study, the PuDColl-2FD showed the highest percentage of pores containing cells, with values that ranged from 74% to 84% at 7 d and 21 d respectively, whereas the maximum value for PuDGel-2FD remained below 66% at 21 d, indicating that collagen coated pores provided a better support for fibroblast adhesion and growth.

### 5.3. ECM in situ deposition

We then assessed the deposition of proteins of the ECM onto the pores of the scaffolds by the fibroblasts at different time points. In particular, the presence and distribution of collagen I and IV, fibronectin, laminin, and elastin after 7 d, 14 d and 21 d, was

evaluated by immunostaining and confocal imaging (figure 4). After only 7 d of cell culture, it was possible to observe a fibroblast-derived matrix enriched in all the studied proteins. The ECM proteins in the PuD-2FD scaffolds after 7 d appeared in aggregates. At 14 d, the distribution followed the shape of the pores. After 21 d, we observed a global decrease of the amount of protein. The presence of the ECM proteins in the pores of gelatin and collagen coated scaffolds was more homogenously distributed within the pores from day 7 and increased with time, being detectable throughout the entire 3D porous network from day 14. Based on image analysis, we quantified the area covered by the ECM proteins demonstrating an increased secretion of ECM when fibroblasts were cultured on the PuDColl-2FD matrices (figure 5).

Based on these results, PuDColl-2FD scaffold appeared as the best candidate to support fibroblast cell culture for ECM production on the pores of polysaccharide scaffolds. Regarding the time of culture, 14 d of cell culture was identified as the best compromise considering cell culture time and protein secretion. Thus, the formulation PuDColl-2FD after
<figure><img src="image_12.png" /></figure>

14 d of fibroblasts 3D culture was retained to perform the next step, i.e. decellularization using scCO₂ to obtain a biohybrid material.

### 5.4. Decellularization using scCO₂

Decellularization was assessed on collagen coated hydrogels after 14 d of cell culture. Several parameters on scCO₂ were evaluated, including temperature, pressure, time of exposure and the addition of co-solvents to decellularize the scaffolds. Their impact on the scaffold physical properties, the decellularization capacity, and the preservation of the deposited ECM was assessed (results not shown). From these experiments, the following conditions were fixed for our study: 170 bar, 45 °C, 90 min. The addition of SDS as surfactant at a very low concentration (0.5%) was found necessary for DNA efficient removal. The

decellularization efficiency of this method was compared to the treatment of the material under the same conditions but without applying scCO₂ (thereafter referred as Air + 0.5% SDS) and to the use of a detergent/enzymatic-based method (named standard method, SDS St) as previously reported [22] that is representative of the most frequently used methods for tissue decellularization [4]. The superiority of the scCO₂ protocol was demonstrated with an almost complete removal of DNA content (figure 6(A)). Indeed, residual DNA in the treated samples was near 1 ng of DNA per mg of scaffold, whereas values using the standard protocol were found near 55 ng mg⁻¹ of scaffold. To note, the Air + 0.5% SDS group that was treated under the same conditions used for scCO₂ but without scCO₂, thus under atmospheric pressure, led to values comparable to those of the cell
<figure><img src="image_13.png" /></figure>

loaded control samples, demonstrating the efficacy of the scCO₂. Confocal 3D images of the immunostained samples confirmed these results showing the near complete absence of nuclear DNA, with almost no DAPI positive staining in the samples treated with scCO₂ (figure 6(B)). Confocal images also revealed that this decellularization high efficiency was accompanied by the preservation of the ECM proteins within the pores of the scaffold (figures 6(B) and S3). Indeed, the scCO₂ decellularization also resulted in the enhanced preservation of the ECM as compared to the standard decellularization treatment, as demonstrated by the presence of higher amounts of all the proteins (figure 6(C)).

## 6. Discussion

Synthesis conditions of the five formulations led to dried porous scaffolds with a thickness between 1 mm and 1.3 mm (figure 2(A)). As observed by SEM, pores on the surface of the materials were large enough to allow cell penetration within the material that could colonize them through the interconnected network of

pores. During the FD process, the scaffolds are placed on shelves that are cooled-down until complete freezing of the material. This means that the freezing of the samples takes place from the bottom to the top with a faster ice-crystal formation on the surface in contact with the shelf that leads to less crystal formation and smaller crystal size, and in consequence to less and smaller pores on this surface compared to the upper surface where water crystal growth is slower [24]. Another consequence of this different crystal growth kinetics, was the gradient in the sizes of the pores that was observed on the SEM images of the cross-sections. This gradient was more evident in the samples that underwent one FD step (PuD-FD and PuDGel-FD). In PuDGel-FD, the direct incorporation of protein in the polysaccharide network was associated to an obvious gradient of pores size with a complete absence of pores on one of the faces. On the other hand, the second FD step, contributed to maintain a more homogenous pore size, with more abundant pores. SEM also revealed the presence of precipitates on the surface of PuDGel-2FD and PuDColl-2FD corresponding to gelatin or collagen. The absence of
precipitates on the matrix of polysaccharides in the PuDGel-FD confirmed the possible cross-linking of the gelatin to the polysaccharide chains through the STMP which was used here as a cross-linker for the polysaccharide chains. This can be explained by the presence of hydroxyl groups in the amino acids of gelatin that in basic conditions could react with the STMP cross-linker in the same way as polysaccharides. This mechanism has been recently proposed to obtain gelatin nanoparticles for applications in the food industry [25]. Although STMP has been mainly used in the past to cross-link polysaccharides, these results confirm its potential to be applied for gelatin-based hydrogels. In the 2FD method, the free proteins in solution were deposited on the surface of the already crosslinked polysaccharide matrix. A similar protocol used in a recent study demonstrated that the interaction between the polysaccharide matrix and the proteins after the second freeze-drying step is sufficiently strong to prevent protein leaching for up to 14 d under cell culture conditions [24].

Swelling of the materials was fast, with stable values reached between 10 and 30 min after hydration. The swelling ratio slightly differed between the 5 formulations. While the addition of a second FD reduced the swelling, the surface coating was impacted when protein was added after the first FD, reducing the swelling kinetics but reaching similar values as the control without coating after 30 min. A plausible explanation can be a lower material hydrophilicity due to the presence of the proteins that present hydrophilic and hydrophobic domains and are less soluble than the polysaccharides used in this work that are characterized by high density of hydroxyl groups [26]. The degradation kinetics was also slower for the samples containing proteins, but clear differences were found depending on the protein loading method, with a prolonged degradation time of the scaffolds in which the proteins were added after the first FD step. This was expected since the enzymes used in this study, namely pullulanase and dextranase, hydrolyze α-1,6 or α-1,4-glycosidic branch chain linkages in pullulan and α-1,6-glycosidic linkage in dextran polymers, but have no action on peptide bonds [27, 28]. It is important to point out that this *in vitro* enzymatic degradation does not reflect the *in vivo* resorption, where other physiological enzymes are implicated and that depends also on the inflammatory environment of the implant.

Regarding the shear stress modulus ($G'$) of the scaffolds, all values were found between 1500 Pa and 3500 Pa, similar to the values described in the literature for soft human tissues [23]. When comparing the different formulations, a trend was observed, with lower values after protein incorporation, suggesting that the contribution of crosslinked polysaccharide networks to the storage modulus was higher than in the case of collagen and gelatin that seemed to

facilitate the elastic deformation of the scaffolds independently of the fabrication method. When $G'$ was measured at 25 °C and at 37 °C, a significant reduction with increasing temperature was observed in single FD formulations, whereas double FD showed a mechanical behavior independent of temperature that can be advantageous for its application. When evaluating the impact of cell culture ($1 \times 10^6$ fibroblasts/scaffold) on the mechanical properties, an increase was observed only in the gelatin containing samples, that was less pronounced when gelatin was added after the first FD step. Taken together, these results indicate that incorporating the proteins after the first FD and performing a second FD is advantageous since it results in samples that are mechanically more stable, with $G'$ values that will remain almost constant after cell culture.

Significant differences of the cell metabolic activity depending on the scaffold were observed (figure 2(A)). As expected, and in line with previous results [16], the absence of proteins in PUD-FD and PUD-2FD was associated with a lack of cell adhesion leading to cell quiescence and a progressive reduction in cellular metabolic activity over time. Similarly, when gelatin was added before freeze-drying, the metabolic activity decreased until day 10 and then a slight increase was observed. Even when increasing the percentage of incorporated protein to 10% w/v, the cell metabolic activity followed a similar trend (results not shown). This may be due to the fact that the protein was entangled in the polysaccharide due to covalent cross-linking mediated by STMP and therefore its integrin-binding domains were not sufficiently accessible to the cells [29]. As the cells cannot adhere to the edges of the pores, they tended to form aggregates in the center of the pores. The fibroblasts arranged in aggregates, as observed in figure 3(B) (and figure S2), remained quiescent with a low metabolic activity. This hypothesis aligns with previous studies with fibroblasts in spheroids reported in the literature [30]. On the contrary, when the proteins, gelatin or collagen, where added after the first freeze-drying, the cells adhered to the surface of the pores and results suggest that they could migrate and proliferate. This is supported by an increased metabolic activity with time and the observation of more pores loaded with cells (figure 3). The obtained results motivated the exclusion of uncoated and PuDGel-FD formulations for ECM deposition studies, since the organization in spheres and the reduced metabolic activity could result in low ECM deposition on the surface of the pores.

The next step was to evaluate the capacity of the double FD scaffolds, PuDGel-2FD and PuDColl-2FD, to promote cell penetration and ECM deposition within the interconnected pores of the scaffolds, indicating that collagen coated pores
provided a better support for fibroblast adhesion and growth.

To decellularize tissues, several strategies have been proposed in the literature. Amongst them, chemical-based protocols and in particular protocols using detergents are the most used, frequently in combination with physical and enzymatic methods to improve their efficacy. In those protocols, detergents act by disrupting cell membranes and breaking covalent bonds, whereas enzymes such as DNAs cleave nucleotide bonds contributing to the removal of nuclear content. To reach acceptable decellularization levels, long incubation times and rinsing steps are needed, resulting in tedious processes, that last from hours to weeks, and that are associated to biochemical damage and the removal of proteins, including collagen and fibronectin [31]. As an alternative, methods based on scCO₂ have been proposed in the last years. The advantages of scCO₂ to decellularize ECM and tissues compared to other methods are several, such as the obtention of sterile virus-free acellular tissues in shorter times avoiding the use of potential toxic agents with better preservation of the proteins and secretome [32]. In this process, the ECM is perfused with supercritical CO₂, this is, above its critical point (Tc = 31.1 °C, Pc = 7.38 MPa) (figure 1). Under these conditions, CO₂ behaves both as a gas and as a liquid with excellent extraction properties [33]. Due to the low polarity of CO₂, it is associated to low amounts of co-solvents to remove polar compounds. Here, hydrogels immersed in 0.5% SDS solution under stirring at 500 rpm were treated with scCO₂ at 170 bar and 45 °C for 90 min. The choice of immersing the scaffolds was done to avoid structural damages of the material due to the perfusion of scCO₂ at high pressure and to material shrinkage caused by dehydration. These conditions and setup were proved efficient to remove almost all the DNA content as shown in figure 6. The residual DNA content in the scaffolds was comparable to that of other ECM-derived materials already commercialized and used in clinical applications [34]. To note, samples undergoing the same treatment in the absence of scCO₂ (Air + 0.5% SDS) presented levels of DNA comparable to non-treated samples. The standard protocol was partially effective but DNA was still detectable in all the analyzed samples (figures 6(A) and (B)). The porous structure of the scaffolds was preserved, as shown in confocal images in figure 6(B). The superiority of our method was also proved by the higher levels of proteins remaining on the pores of the scaffolds. Indeed, the standard treatment led to significantly lower levels of all the proteins under study. This is consistent with previously reported results with different types of tissues [35–37]. Interestingly, the scaffolds that underwent a treatment comparable to scCO₂ group but without

applying the supercritical fluid also showed a loss of collagen I, fibronectin, elastin and laminin, that was not accompanied by an efficient removal of DNA (figure 6). Under our optimized conditions, scCO₂ acted as a potent decellularization agent that not only did not cause ECM damage compared to standard methods but that had indeed a protective effect.

## 7. Conclusion

The development of hydrogels for tissue engineering that match the physical-chemical, mechanical and biochemical characteristics of native ECMs remains a challenge for the fabrication of biomaterials. We proposed an original and versatile approach based on *in situ* ECM deposition on polysaccharide scaffolds, mediated by fibroblasts, and followed by an optimized decellularization method using scCO₂. Previous works with polysaccharide scaffolds, show that we can easily tune the properties of polysaccharide-based scaffolds for different *in vitro* and *in vivo* applications, including vascular [13, 14], liver [15, 16], bone [18, 38], neural [17] tissues. Thus, the biofabrication method presented here could be applied to various polysaccharide scaffolds presenting different characteristics depending on the targeted tissue, in combination with allogenic tissue-specific cells to match not only the physical and mechanical properties but also the ECM composition while reducing the immunological risk.

## Data availability statement

The data cannot be made publicly available upon publication because they are not available in a format that is sufficiently accessible or reusable by other researchers. The data that support the findings of this study are available upon reasonable request from the authors.

## Acknowledgment

This project has received funding from the French Research Agency ANR under the project 'EXCALYBUR' (Grant No. ANR-20-CE18-0001-01) and from IdEx Université Paris Cité, ANR-18-IDEX-0001ANR-18-IDEX-0001 under the project 'POLCA'. 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 infrastructure (ANR-10-INBS-04) and GIS-IBiSA; and the support of the Région Île-de-France (Sesame).
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