# Hydrogels from Wharton's jelly as alternative to conventional extracellular matrix-based constructs

Anais Lavrand, Lorinne Adam, Alexis Da Rocha, Flora Lemaire, Capucine

Loth, Adrien Baldit, Maxime Vassaux, Laurence van Gulick, Abdelilah

Beljebbar, Pedro Augusto, et al.

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## ► To cite this version:

Anais Lavrand, Lorinne Adam, Alexis Da Rocha, Flora Lemaire, Capucine Loth, et al. Hydrogels from Wharton's jelly as alternative to conventional extracellular matrix-based constructs. *International Journal of Biological Macromolecules*, 2025, 328, pp.147552. 10.1016/j.ijbiomac.2025.147552. hal-05296711

HAL Id: hal-05296711

https://hal.sorbonne-universite.fr/hal-05296711v1

Submitted on 3 Oct 2025

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Distributed under a Creative Commons Attribution 4.0 International License
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# Hydrogels from Wharton's jelly as alternative to conventional extracellular matrix-based constructs

Anais Lavrand<sup>a</sup>, Lorinne Adam<sup>a</sup>, Alexis Da Rocha<sup>b</sup>, Flora Lemaire<sup>a</sup>, Capucine Loth<sup>c</sup>, Adrien Baldit<sup>b</sup>, Maxime Vasseaux<sup>d</sup>, Laurence Van Gulick<sup>e</sup>, Abdelilah Beljebbar<sup>e</sup>, Pedro Augusto<sup>f</sup>, Alexandre Berquand<sup>g</sup>, Chrystelle Salameh<sup>h</sup>, Nadine Nassif<sup>h</sup>, Fouzia Boulmedais<sup>c</sup>, Cédric Mauprivez<sup>a,i</sup>, Esteban Brenet<sup>a,j</sup>, Halima Kerdjoudj<sup>a,*</sup>

## ARTICLE INFO

**Keywords:**

Wharton's jelly

Hydrogel

Extracellular matrix

Physicochemical characterization

## ABSTRACT

Extracellular matrix (ECM)-based hydrogels are increasingly used in regenerative medicine; however, animal-derived sources such as rat tail tendon ECM (RT-ECM)—the current gold standard—face limitations for clinical translation. We propose hydrogels derived from decellularized human Wharton's jelly (WJ-ECM) as a bioactive and sustainable alternative. Compared to RT-ECM, WJ-ECM hydrogels exhibit a distinct biochemical composition, combining type I/III collagen and glycosaminoglycans, and form thinner fibrils. Upon exposure to culture medium, they display reduced syneresis. Rheological and confined compression tests reveal lower stiffness and decreased permeability. Functionally, WJ-ECM hydrogels demonstrate superior hemostatic properties, reduce intracellular ROS accumulation in neutrophils, and promote an anti-inflammatory phenotype in macrophages. *In vivo* implantation confirms their biocompatibility with minimal adverse effects. These results highlight the potential of WJ-ECM hydrogels as a clinically relevant biomaterial for regenerative medicine and cell culture applications.

## 1. Introduction

Cells are intimately connected to their surrounding extracellular matrix (ECM). Natural polymers such as collagen and hyaluronic acid contain ECM-like components and can be used to generate hydrogels or scaffolds that regulate cell behavior. [1,2] However, fully replicating the biomolecular and biophysical cell-instructive properties of the ECM—a complex network of proteins, glycosaminoglycans, and associated growth factors—remains a significant challenge. Biologics-based materials, particularly those derived from human sources, hold great potential for regenerative medicine applications. Tissue decellularization

allows for the generation of multiphasic hydrogels and scaffolds that closely mimic the native ECM. [3,4] While animal-derived ECM materials have been widely used in regenerative medicine, concerns regarding potential immune responses (e.g., reactions to bovine collagen, proteoglycan link proteins, glycan antigens, and α-galactosyl epitopes), pathogen transmission, batch-to-batch variability, and religious or regulatory restrictions have driven a shift toward human-derived ECM. Human-derived ECM offers several advantages, including greater clinical translatability, improved biocompatibility, and ease of use. [5] For example, ECM derived from human dermal fibroblasts has been employed as a substrate for cell culture and is

<footnote><sup>a</sup> University of Reims Champagne Ardenne, Reims, France</footnote>

<footnote><sup>b</sup> University of Lorraine, CNRS UMR 7239 LEM3, Metz, France</footnote>

<footnote><sup>c</sup> University of Strasbourg, CNRS Institut Charles Sadron, Strasbourg, France</footnote>

<footnote><sup>d</sup> Université de Rennes, CNRS, Institut de Physique de Rennes (IPR) – UMR 6251, Rennes, France</footnote>

<footnote><sup>e</sup> University of Reims Champagne Ardenne, UR BIOSPECTRE, Reims, France</footnote>

<footnote><sup>f</sup> Université Paris-Saclay, CentraleSupélec, Laboratoire de Génie des Procédés et Matériaux, Centre Européen de Biotechnologie et de Bioéconomie, Pomacle, France</footnote>

<footnote><sup>g</sup> University of Reims Champagne Ardenne, UMR CNRS 7369 MEDyC, Reims, France</footnote>

<footnote><sup>h</sup> Laboratoire de Chimie de la Matière Condensée de Paris (LCMCP), CNRS, Sorbonne Université, Collège de France, Paris, F-75005, France</footnote>

<footnote><sup>i</sup> Pôle de santé buccodentaire, CHU de Reims, Reims, France</footnote>

<footnote><sup>j</sup> Pôle tête et cou, Oto rhino laryngologie, CHU de Reims, Reims, France</footnote>

<footnote>* Corresponding author at: Université de Reims Champagne Ardenne, 1 avenue du Marechal Juin, 51100, Reims, France. E-mail address: halima.kerdjoudj@univ-reims.fr (H. Kerdjoudj).</footnote>
increasingly used in tissue repair, including applications in vascular and cardiac valve engineering. [6,7] Although fibroblast-derived ECMs exhibit biomolecular cell-instructive properties, their thickness is typically limited to 30 µm, [8] and their application in regenerative medicine requires extensive *in vitro* culture periods. Given the global birth rate of approximately 140 million babies per year [(World Fertility Report, 2024)], perinatal tissues present a valuable opportunity for the development of biologics, as they are readily accessible from both a technical and ethical standpoint. Perinatal tissues have been successfully used as allografts in various medical fields, including ocular surface reconstruction, burn and ulcer treatments, and craniofacial reconstruction. [9,10] While the clinical application of fetal membranes is well established, the use of umbilical cord tissue remains relatively novel in this context. [11] The human umbilical cord, a crucial structure for fetal growth and development, measures approximately 50–60 cm in length at full term. This connective tissue contains blood vessels embedded within Wharton's jelly (WJ), a specialized mucoid connective tissue arranged in a helical fashion around the blood vessels. WJ is now widely recognized as an optimal ECM, providing mechanical, physical, and biochemical support. [9,11] It is rich in fibrillar and non-fibrillar collagens, proteoglycans, glycosaminoglycans, immunomodulatory cytokines, and growth factors. [12,13] WJ-based biologics are among the few immune-privileged connective tissues, [14] making them attractive for homologous defect repair. Due to its exceptionally bioactive composition, decellularized WJ matrix has been combined with poly-caprolactone to improve the biocompatibility and host integration of electrospun scaffolds. [15] Clinical applications of WJ-derived biologics are increasingly reported in medical literature, showing promising outcomes in functional recovery and pain management. [16] Translating WJ-derived ECM macromolecules into bioactive hydrogels opens new avenues for stimulating a range of biological processes essential for tissue regeneration. For instance, umbilical cord-derived hydrogel has shown potential for treating degenerative intervertebral discs. [17] Similarly, injectable decellularized WJ hydrogels containing exosomes have demonstrated efficacy in promoting meniscal tear healing by enhancing meniscal chondrogenesis, preserving articular cartilage, and preventing secondary cartilage degeneration. [18]

Despite the growing interest in WJ-derived ECM hydrogels for regenerative medicine applications, their comprehensive characterization remains incomplete. The understanding of molecular self-assembly is a crucial factor for the fabrication of biomaterials. This study presents the development of a WJ-derived hydrogel without additional chemical crosslinking and provides a comparative assessment of its structural, physicochemical, mechanical, and biological properties relative to those of rat-tail-tendon ECM—the most commonly used preclinical ECM source. Our findings help bridge the gap between the widespread availability of umbilical cord-derived tissues and the design of functional ECM-based hydrogels for cell culture and tissue engineering applications.

## 2. Material and methods

### 2.1. Extracellular matrix extraction and characterization

Wharton's Jelly (WJ) was extracted from the umbilical cord and decellularized as previously described [13] then freeze-dried. Rat tail tendon (RT) was extracted, washed several times in PBS 1× and freeze-dried without further treatment (Fig. 1). The isolation of extracellular matrix (ECM) components was performed by incubating 10 mg/mL of freeze-dried tissues in HCl 0.1 N with 1 mg/mL pepsin at 4 °C for 24 h for RT-ECM and 48 h for WJ-ECM. The digested suspension was centrifuged at 300g, 4 °C for 5 min and the undigested matrix was removed. The pH was adjusted to 7.2 with NaOH 2 N, buffered with 10 % of the volume in NaCl 9 % and kept at 4 °C or freeze-dried. Hydrogel was formed by incubating the buffered ECM digests at 37 °C for 40 min. To form sponges, WJ- and RT-ECM hydrogels were frozen at -80 °C for 24 h followed by lyophilization for 24 h (COSMOS -80, Cryotec, France).

### 2.2. Protein and collagen quantifications

Total protein content was determined at absorbance (λ) 280 nm using Nanodrop® (Thermo Fisher Scientific, USA). WJ- and RT-ECM digests (10 µL) as well as 4-hydroxyproline standards were dried in thermoresistant, Teflon capped microtubes and resuspended in HCl (100 µL, 6 N) under anoxia in a globe box. The tubes were tightly closed and heated to 110 °C overnight. Then, NaOH (101 µL, 6 N) were added and the neutralized hydrolysate (10 µL) were processed for AccQTag (Waters) labeling according to the supplier's recommendations, directly in total recovery sampler tubes. Amino acids were analyzed by UPLC-MS on a Acquity Premier system equipped with a QDa mass detector (all from Waters), using a Cortecs C18 column (15 cm × 2.1 mm, 1.6 µm particles, also from Waters) and a 5-min eluting gradient, ranging from 1 % to 14 % acetonitrile in water equilibrated with 0.1 % formic acid. 4-hydroxyproline was monitored based on the mass detector signal recorded in single ion monitoring focusing at 302.2 Da, with 10 V cone voltage. Calibration curves and sample concentration calculations were all managed in Empower based on peak areas. After data extraction, actual sample contents were corrected based on sample dilutions and volumes used in the analysis.

### 2.3. DNA quantification

DNA quantification was performed on WJ- and RT-ECM digests using MasterPure® Complete DNA & RNA Purification kit (Biosearch Technology, UK) according to the manufacturer instruction. The quantity of extracted DNA was assessed by measuring the absorbance (λ<sub>260</sub> Nanodrop, Thermo Scientific, Rockford, IL, USA) with λ<sub>260</sub>/λ<sub>280</sub> ratio for all measured samples comprised between 1.8 and 2.

<figure><img src="image_4.png" /><figcaption>Fig. 1. Methodological procedure for ECM extraction.</figcaption></figure>
## 2.4. Sulfated and non-sulfated glycosaminoglycans (GAGs) quantification

Sulfated and non-sulfated GAGs quantification was performed on WJ- and RT-ECM extracts using Blyscan and Purple-Jelley kits (Biocolor, UK), respectively according to the manufacturer instruction. Absor-bances were measured respectively at λ₆₅₆ and λ₆₅₅ with a standard made with bovine tracheal chondroitin 4-sulfate for sulfated GAG dosage and hyaluronan for hyaluronic acid dosage.

## 2.5. Rheology and transmittance

To follow the gelation kinetics, the increase in the storage modulus (G') was assessed using ElastoSens® rheometer (Rheolution®, Canada) in soft mode for 40 min at 37 °C. The absorbance of digests and hydrogels was assessed using a FLUOstar Omega microplate reader (BMG Labtech, France). Briefly, 150 µL of digests were placed in 96-wells plate and the λ₃₀₀ to λ₇₀₀ was measured. The absorbance measurement was recorded a second time after incubating the plate at 37 °C for 40 min, determining the absorbance (A) of the formed hydrogels. Transmittance was calculated with the formula:

$$Transmittance (\%) = 10^{2-A}$$

## 2.6. In vitro fibrillogenesis degree and gel fraction

The efficiency of hydrogel's networks formation was evaluated using the in vitro fibrillogenesis degree (FD) and the gel fraction and measurements. FD representing the percentage of collagen involved in the gel network was determined. For that, the concentration of collagen was first assessed in the digest. The hydrogel was centrifuged at 14000g for 30 min and the concentration of collagen in the supernatant was performed using o Sircol kit protocol (Biocolor, UK). The absorbance was measured at λ₆₅₆ and the standard curve was made with bovine collagen in acetic acid provided by the kit. Fibrillation degree was calculated with the formula:

$$FD (\%) = \left( 1 - \frac{[collagen]_{supernatant}}{[collagen]_{digest}} \right) \times 100$$

The percentage of the gel fraction was obtained as follow. Hydrogels were freeze-dried, weighed and then rehydrated for 15 min in PBS 1 ×. The hydrated samples were weighed again, and the gel fraction was calculated with the formula:

$$Gel fraction (\%) = \frac{m_h}{m_0} \times 100$$

Where m_h refers to the mass of hydrated hydrogel, while m_0 represents the freeze-dried hydrogel.

## 2.7. Single scan polarization-resolved second harmonic generation (SS-pSHG)

WJ- and RT-ECM hydrogels were imaged using two-photon excitation laser scanning confocal microscopy with SS-pSHG (LSM 710-NLO, Carl Zeiss SAS, Germany) coupled with a CHAMELEON femtosecond Titanium-Sapphire Laser (Coherent, USA).

## 2.8. Scanning electron microscopy (SEM)

The freeze-dried hydrogels were also coated with a gold-palladium film using ion sputtering (JEOL JFC 1100, France) for observations with SEM (evo 10, Zeiss, Germany) with an accelerating voltage of 5 kV.

## 2.9. Atomic force microscopy (AFM)

AFM imaging was performed on air-dried hydrogels. Briefly, a drop of digest was deposited onto a freshly cleaved mica disk incubated at 37 °C for 40 min then dried in air-dryer oven at 37 °C (GelAir Dryer, BIORAD, USA) for 24 h. Samples were imaged in PeakForce QNM mode using a MultiMode 8 (Bruker, Billerica USA) using MPP-12120-10 TAP150A probes, calibrated on mica prior to the experiment and having the following characteristics: resonance frequency ranging from 135 to 159 kHz and Spring constant of 5 N/M (nominal value). The images were processed using Nanoscope Analysis v. 9.2 (Bruker, Billerica USA).

## 2.10. Transmission electron microscopy (TEM)

TEM was performed on resin embedded hydrogels following the standard preparation procedure. WJ- and RT-ECM hydrogels were fixed in glutaraldehyde 2.5 % for 48 h and washed in PBS 1×. samples were cut into pieces of 1 mm and immersed in osmic acid in the dark and at room temperature for 1 h then dehydrated in solutions with gradually increasing concentration of ethanol for 15 min and in propylene oxide for 15 min. Samples were placed in a mix of epoxy resin (Epon®) and propylene oxide (v/v) overnight at room temperature. Samples were embedded in pure epoxy resin for 48 h at 60 °C. ultra-thin sections (70–80 nm-thick slices) were performed using an ultra-microtome (Ultracut-E Ultramicrotome, Leica, France). Specimens were contrasted using UAR-EMS uranyl acetate stain (LFG Distribution, France) and observed with a TEM (JEM-2100F, JEOL, France) operating with an accelerating voltage of 70 kV.

## 2.11. Water holding capacity (WHC)

WJ- and RT-ECM of hydrogels were prepared in homemade cylinder molds and weighed, air-dried at 37 °C for 3 h then weighed. The WHC was determined using the following formula:

$$WHC (\%) = \left( 1 - \frac{m}{m_0} \right) \times 100$$

Where m refers to the air-dried hydrogel, while m_0 represents the initial hydrogel.

## 2.12. Deswelling and syneresis

WJ- and RT-ECM hydrogels (formed from 150 µL in a 96-well plate) were weighed and placed in 1 mL of DMEM without phenol red supplemented with decomplemented fetal bovine serum (FBS 10 %, v/v) and penicillin/streptomycin/amphotericin cocktail (PSA 1 %, v/v) at 37 °C for a week. The hydrogels were weighted every day and the deswelling ratio was calculated with the formula:

$$Deswelling (\%) = \frac{m_d}{m_0} \times 100$$

Where m_d refers to the deswelled hydrogel, while m_0 represents the non-deswelled hydrogel.

The protein content in the supernatant was measured every day by Nanodrop® at λ₂₉₀.

WJ- and RT-ECM hydrogels, formed in Eppendorf, were weighted and centrifuged at 1500g for 10 min at 20 °C. The released water was removed and the remaining hydrogels were weighted. Syneresis was determined following the formula:

$$Syneresis (\%) = \left( 1 - \frac{m}{m_0} \right) \times 100$$

Where m refers to the centrifuged hydrogel, while m_0 represents the non-centrifuged hydrogel.
## 2.13. Wettability

A drop of demineralized water (1.5 µL) was deposited on the surface of the air-dried hydrogels and the water contact angle was recorded at room temperature using a Drop Shape Analyzer-DSA25 (Kruss-scientific, Germany). High-resolution images were captured at a rate of 25 images/ s using the DSA25's camera and drop Shape Analyzer software was used to analyze the contact angles, while the absorption time was quantified using ImageJ version 2.9.

## 2.14. Gravimetric swelling

WJ- and RT-ECM sponges were weighted and soaked in PBS for 15 min at room temperature. The gravimetric swelling was determined after weighting the hydrated sponges using the following equations:

$$ \text{Swelling ratio (\%)} = \frac{m - m_0}{m} \times 100 $$

Where m refers to the mass of sponge with PBS, while m₀ represents the mass of dried sponge.

## 2.15. Total porosity

WJ- and RT-ECM sponges were weighted (m₀) and soaked in absolute ethanol for 15 min at room temperature. The total porosity was determined after weighting the ethanol-sponges using the following equations:

$$ \text{Porosity (\%)} = \frac{m - m_0}{m} \times 100 $$

Where m refers to the mass of sponge with ethanol, while m₀ represents the mass of dried sponge.

## 2.16. Differential scanning calorimetry

The temperature resistance of WJ- and RT-ECM hydrogels (5 mg) was performed with a TA Q-20 machine (TA Instruments, Germany). The heating rate was set at 5 °C/min and the temperature ranged from 20 °C to 80 °C.

## 2.17. Molecular dynamics simulations

We perform molecular dynamics simulations of the collagen microfibril using LAMMPS. [19] The interactions between the atoms are described and parameterized using the CHARMM36m potential, [20] which is facilitated by the use of the CHARMM-GUI. [21] The initial structure of the type I collagen microfibril is generated using ColBuilder. [22] The structure is based on the microfibril crystallographic structure obtained by Orgel et al. [23] using X-ray fiber diffraction on native rat tail tendon in situ. The structure is further refined using additional high-resolution structures of short collagen-like peptides from X-ray crystallography and mass spectrometry data. The initial structure of the type III collagen microfibril is obtained from the type I structure and homology modelling performed using the I-Tasser server. [24] Homology modeling is performed to replace the α-2 chain by a third α-1 chain. The hydration of the microfibril is performed adding 12,600 water molecules to the system. The water molecules are inserted at random locations such that no atom in the water molecule is found less than 1.2 Å away from any other atom in the system. Thirty-two chlorine ions are also inserted to render the system charge neutral. The hydrated and neutralized molecular model is then relaxed at constant volume and at a constant temperature of 308 K (NVT ensemble) until the total energy of the system is converged. For the least equilibrated system, that is the type III microfibril, convergence of the energy is reached after approximately 1 ns. The complete validation of the hydration procedure was

published elsewhere. [25] Once equilibrated and relaxed, the dynamics of the molecular model of the microfibrils are simulated at a constant pressure of 1 atm and a constant temperature of 308 K. The pressure is only applied on the lateral surfaces of the microfibril model, while the long direction of the model is maintained at a constant length of 67 nm which is known to be preserved experimentally.

## 2.18. Confined compression test

WJ- and RT-ECM hydrogels in cylindrical shape (formed from 1.2 mL in a 48-well plate) were subjected to confined compression in a 10 mm diameter chamber with two porous end-plates using a universal tensile/ compression test machine (Zwick 0.5) within close physiological conditions with samples immersed in NaCl 0.9 % at 37 °C. The 0.1 N load was applied with a controlled velocity of 0.02 mm/s and held for 30 min. The time displacement curves were fitted using the model developed by Selvadurai and Suvorov, [26] leading to the characterization of the parameters through an optimization loop using the Constrained Optimization By Linear Approximation (COBYLA) method from the Python Scipy library. Young's modulus (E), Poisson's ratio (ν) and the permeability (k) values were extracted.

## 2.19. Flow properties

The flow properties of WJ- and RT-ECM hydrogels were assessed in a rheometer with controlled stress (Thermo Scientific HAAKE MARS iQ) using a cross-hatched plate-plate geometry (35 mm of diameter, 1 mm gap). The temperature was maintained constant at 37 °C using a Peltier system. The rheological evaluation was carried out with hydrogels, which were first placed in the rheometer and maintained at rest for 180 s before shearing. After resting, the samples were sheared from 0.01 to 100 s⁻¹ with a log increasing stepwise protocol to guarantee steady-state shear conditions, while the shear stress and apparent viscosity were measured.

WJ- and RT-ECM extracts were stained with ponceau red and Evans blue, respectively and were transferred in syringes with 27G needle. The solution was manually extruded on a silicone sheet to take pictures.

## 2.20. Raman spectroscopy

Raman spectra of WJ-ECM and RT-ECM were acquired using a near-infrared Raman spectrometer (Labram HR 800, Horiba Jobin Yvon S.A. S., France). The system consisted of a microscope (Olympus BX41, France) coupled to a spectrometer equipped with a 600 groove/mm diffraction grating. A motorized xy-stage (Marzhauser, Germany) enabled automated sample scanning with a spatial resolution of 1.5 µm. The excitation source (532 nm) was provided by laser diode (Laser Quantum, Germany) delivering 25 mW of laser power on the sample. The laser beam was focused on the sample with a 100 x objective (NA 0.90, Olympus, France). The backscattered light was collected by the objective and transmitted to the spectrometer equipped with a Pelletier-cooled charge-coupled device (CCD) detector. Each spectrum was acquired with a 10 s integration time in the 1000–1800 cm⁻¹ spectral region, with a spectral resolution of 4 cm⁻¹. Data processing was performed in MATLAB (MathWorks, MA, USA). Preprocessing steps included instrument response correction, wavenumber calibration, fluorescence background subtraction, cosmic ray removal, and baseline correction using a fifth-order polynomial fit. The spectra were smoothed using a seven-point Savitzky-Golay filter, followed by normalization via Standard Normal Variate (SNV) transformation.

## 2.21. Hemocompatibility

WJ- and RT-ECM hydrogels (formed from 300 µL in a 24-well plate) were exposed to UV for 20 min and incubated in the presence of peripheral blood (200 µL) and PBS 1 × (1.8 mL) for 4 h at 37 °C. Negative
and positive controls corresponding to PBS-diluted blood and water diluted blood, respectively were used. Absorbance of released hemoglobin was measured at λ540nm and the hemolysis was determined as following:

$$ \text{Hemolysis (\%)} = \frac{A - A_1}{A_2 - A_1} \times 100 $$

Where A refers to the absorbance of hemoglobin in the presence of the hydrogels, while A₁ and A₂ represent the absorbance of hemoglobin in PBS and distilled water respectively.

UV-exposed hydrogels were incubated in the presence of peripheral blood (50 µL) and CaCl₂ (5 µL, 0.2 M) at 37 °C. A mixture of peripheral blood and CaCl₂ in the absence of hydrogel was used as control. After 5 min of incubation, distilled water (2 mL) was added. Absorbance of released hemoglobin was measured at λ540nm and the blood coagulation index (BCI) was determined as following:

$$ \text{BCI}(\%) = \frac{A}{A_1} \times 100 $$

Where A refers to the absorbance of hemoglobin in the presence of the hydrogels, while A₁ represents the absorbance of hemoglobin in blood/ CaCl₂ mixture.

The hydrogels were fixed with glutaraldehyde 2.5 %, dehydrated, coated with a gold-palladium film using ion sputtering and imaged by SEM (evo 10, Zeiss, Germany).

The kinetic of coagulation was also assessed by Elastosens® rheometer macro-holder. Briefly, sponges (60 mg) were placed in the macro-holder and a mixture of peripheral blood (1 mL) and CaCl₂ (100 µL, 0.2 M) was added. Placed at 37 °C, the coagulation kinetic was assessed for 2 h. The resulting clot was placed on absorbing sheet for 10 min before being imaged.

## 2.22. Inflammation

### 2.22.1. Neutrophil activation

Human neutrophils were isolated from peripheral blood with Polymorphprep (ProteoGenix, France). WJ- and RT-ECM hydrogels (formed from 300 µL in a 96-well plate) were exposed to UV for 20 min. Neutrophils (i.e. 1 × 10⁶) were added to each well and incubated for 2 h at 37 °C and 5 % CO₂ in the presence of RPMI-Glutamax® (Gibco, USA), autologous serum (2.5 %, v/v) and penicillin/streptomycin/amphotericin cocktail (PSA, 1 %, v/v), with or without lipopolysaccharide (LPS, 10 ng/mL, Sigma Aldrich). The cell culture medium was harvested and stored at -20 °C until analysis.

### 2.22.2. Intracellular reactive oxygen species (ROS) measurement

After 2 h of contact with hydrogels, Neutrophils were washed in PBS 1× and intracellular ROS were labeled using Muse® Oxidative Stress Kit (Luminex, USA) and analyzed by flow cytometry with the LSRFortessa™ system (BD Biosciences, France).

### 2.22.3. Macrophage polarization

U937 cell line M0 differentiation was obtained by cultivating 10⁶ cells per well in 12-well plate with RPMI-Glutamax® (Gibco, USA), FBS (0.5 %, v/v), PSA (1 %, v/v), phorbol 12-myristate 13-acetate (PMA, 25 nM, Thermo Fisher) and Vitamin D3 (20 nM, Sigma Aldrich) at 37 °C overnight. Mediums were then removed and replaced by RPMI-Glutamax® with FBS (5 %, v/v), PSA (1 %, v/v), PMA (25 nM) and Vitamin D3 (20 nM) for 24 h at 37 °C. Mediums were changed with the same mix in the absence of PMA. U937 were incubated 48 h at 37 °C to obtain M0 macrophages. M0 were seeded on UV-sterilized WJ- and RT-ECM hydrogels, and incubated at 37 °C for 48 h and 5 % CO₂. The cell culture media were then collected and stored at -80 °C until analysis.

### 2.22.4. ELISA tests

Quantification of cytokines was performed by ELISA. IL-8 proteins dosage was performed on the neutrophil's culture media according to the IL-8 Duoset® ELISA kit (R&D systems, Germany) instructions. IL-1β, TNF-α, IL-10, VEGF, IL1-ra and TGF-β production was assessed on the U937 culture media according to IL-1β, TNF-α, IL-10, VEGF, IL1-ra and TGF-β Duoset® ELISA kits (R&D systems, Germany). Absorbances were measured at λ450nm with a λ570nm correction. Cytokine concentration was calculated according to the corresponding standard curve.

## 2.23. Live dead staining

Human dermal fibroblasts were seeded at 10⁴ cells/cm² on UV-decontaminated WJ-and RT-ECM hydrogels and cultured for 4, 7 and 11 days in DMEM-Glutamax® (Gibco, USA) supplemented with FBS (5 %, v/v) and PSA (1 %, v/v). Live/Dead staining was performed on adhered fibroblasts using calcein/propidium iodide according to the Live Dead Cell Viability Assay kit for 3D and 2D cell culture (Merck, USA) protocol. Images were acquired immediately after staining with the Axiovert 200 M (Zeiss, Germany) microscope.

## 2.24. In vivo biocompatibility

A subcutaneous implantation model was used in this study. Eight-week-old male Wistar rats (n = 4) were anesthetized by inhalation of isoflurane (isuflu-vet, Netherland) at 5 % for induction and 2.5 % for maintenance. Four hydrogels were separately implanted subcutaneously in each animal and the animals were daily followed by weight survey. After 56 days of implantation, rats were euthanized by cervical dislocation procedure preceded by over-inhalation of isoflurane. Samples were harvested, fixed in formol, and embedded in paraffin. 3 µm-thick sections were deparaffined and stained with hematoxylin-eosin-safran or picosirius red (DiaPath, Italy). Immunohistochemistry was performed on 3 µm-thick sections, using CD68 antibody (AB125212, dilution 1/1000, ABCAM, Netherland) and revealed according to the Mouse and Rabbit Specific HRP/DAB Detection IHC kit (AB64264, ABCAM, Italy) protocol. Images were acquired using an OLYMPUS VS 120 scanner with polarized light for picosirius red stain observation. The acquired images were analyzed using QPath software focused on different histological parameters to assess the biocompatibility of hydrogels.

## 2.25. Statistical analysis

Statistical analyses were conducted using GraphPad Prism 10. The physicochemical characterizations were performed on four independent samples per condition. Inflammatory studies were performed on 6-independent donors of blood and on three independent hydrogels per donor. Results were presented with histograms (mean ± SEM), and statistical analysis was conducted using t-test, Mann & Whitney test and two-way ANOVA test. Results with p-value <0.05 were considered statistically significant.

## 2.26. Ethical statement

All human samples used to design the hydrogels/scaffolds and used for primary dermal fibroblasts as well as blood were obtained from informed patients with written consent and in accordance with legal and ethical guidelines (Article R 1243-47). All procedures were conducted with the appropriate authorization and registration number (DC-2014-2262) issued by the French National "Cellule de Bioéthique". Animal experiments were carried out in accordance with the 3R rule and in compliance with the animal care and use committee of the University of Reims Champagne-Ardenne (2021111219002699).
## 3. Results and discussion

In the field of regenerative medicine, human-derived hydrogels have recently gained prominence due to their high content of bioactive domains and growth factors. [5] The umbilical cord, with its intrinsic regenerative properties, [12,13] holds significant potential to overcome many limitations associated with xenogeneic collagen sources, making it highly suitable for clinical translation. In this study, we developed hydrogels derived from human Wharton's Jelly extracellular matrix (WJ-ECM) and compared their structural, physical, mechanical, and biological properties with those of hydrogels obtained from rat tail tendon-derived ECM (RT-ECM). Rat tail tendon collagen is widely used in tissue engineering because it is rich in type I collagen, facilitating comparison with the literature. [27,28] Moreover, hydrogels derived from rat tail collagen exhibit physico-chemical properties comparable to those of clinical-grade collagen derived hydrogels. [29,30] Unlike rat tail tendon, which is free of DNA contamination, WJ tissue contains DNA at a concentration of 72.7 ± 25.4 ng/µL—below the threshold established by Crapo et al. [31] To eliminate residual DNA, WJ tissue was decellularized using a detergent-enzymatic treatment. [12,13] This protocol effectively preserves key growth factors, including basic fibroblast growth factor, vascular endothelial growth factor, hepatocyte growth factor, and transforming growth factor-beta. While rat tail tendon dissolves in acidic solutions, decellularized WJ requires enzymatic solubilization. To ensure a standardized comparison, both tissues were solubilized using pepsin digestion, which preserves collagen ultrastructure by selectively cleaving telopeptides—regions where the three α-chains do not form a stable triple helix. [32] The removal of terminal collagen peptides also reduces the immunogenicity of the resulting hydrogels. [33] To characterize ECM composition, we quantified DNA, glycosaminoglycanes (GAGs), total protein, and collagen (hydroxyproline) contents (Table 1, Fig. SI-1).

WJ-ECM: Wharton's Jelly extracellular matrix, RT-ECM: Rat tail extracellular matrix, Hyp: Hydroxyproline, sGAGs: Sulfated glycosaminoglycanes, nsGAGs: Non-sulfated glycosaminoglycanes.

DAPI staining does not reveal nuclear structures in either extract, and DNA quantification confirms levels below the detection limit of the kit used (12 ng/µL), further validating compliance with Crapo's recommendations. [31] While the total protein content is significantly lower in WJ-ECM compared to RT-ECM (p < 0.001, t-test), no significant difference in collagen content (~ 4 mg/mL) is observed between both extracts (Fig. SI-1 A). Since pepsin digestion breaks down proteins into smaller peptides, variations in ECM protein composition may arise due to differential degradation patterns. Notably, sulfated and non-sulfated GAG content is 2.7- and 4-fold higher, respectively, in WJ-ECM compared to RT-ECM (p = 0.02, t-test, Fig. SI-1B). Given the differences in concentration ranges (collagen in mg/mL vs. GAGs in µg/mL), we assume that both extracts are primarily composed of collagen. SDS-PAGE analysis confirms that WJ- and RT-ECM solutions contain α1 and α2 collagen chains, as well as their dimer (β chain), exhibiting a pattern consistent with type I collagen. However, WJ-ECM also displays an additional band between the α1 and α2 chains (Fig. SI-1C), corresponding to type III collagen. [34,35] Type I collagen is a heterotrimer composed of two [α-1(I)] chains and one α-2(I) chain, whereas type III collagen is a homotrimer consisting of three identical [α1(III)] chains. MALDI TOF/TOF mass spectrometry confirms the presence of type I and

<table><caption>**Table 1**

Biochemical characterization of Wharton's jelly and rat tail tendon extracts.</caption><thead><tr><th></th><th>Total protein (mg/mL)</th><th>Hyp (mg/mL)</th><th>sGAGs (µg/mL)</th><th>nsGAGs (µg/mL)</th></tr></thead><tbody><tr><td>WJ-ECM</td><td>4.13 ± 0.03</td><td>2.94 ± 0.04</td><td>17.57±5.50</td><td>9.06 ± 0.62</td></tr><tr><td>RT-ECM</td><td>3.64 ±1.09</td><td>3.37 ± 0.20</td><td>6.29 ± 0.58</td><td>2.04 ± 0.26</td></tr></tbody></table>

III collagens in WJ-ECM, along with trace amounts of type V collagen (Tables SI-1 and SI-2). The ratio of type I / III collagen in WJ-ECM is approximately 2. Circular dichroism (CD) spectroscopy used to analyze the collagen folding, shows that both extracts exhibit characteristic sinusoidal CD spectra, confirming the presence of stable triple-helical collagen structures. WJ-ECM display a negative peak at 200 nm, a positive peak at 224 nm, and a crossover point at 215 nm, while RT-ECM exhibit a negative peak at 198 nm, a positive peak at 225 nm, and a crossover point at 218 nm (Fig. SI-1D and E). Differences in peak intensity may reflect the co-expression of type I and III collagens. [36] Upon thermal denaturation, CD spectra demonstrate progressive alterations of the two-lobe pattern, indicating a gradual disruption of native collagen triple-helical structures. The disappearance of positive peaks, characteristic of random polypeptide chain conformations, confirms collagen unfolding.

### 3.1. Characterization of the hydrogels

The WJ- and RT-ECM extracts form fibrillogenesis initiation buffers (FIBs) when re-equilibrated to a physiological pH (7.4), salt concentration, and kept at a low temperature 4 °C. The most abundant minerals in interstitial fluid are Na⁺ (~140 mM) and Cl⁻ (~100 mM), making up ~ 80 % of the total osmolarity (~300 mM). [37] In this study, salt concentration was equilibrated using 150 mM (i.e. 0.9 %) NaCl solution, commonly referred to as physiological saline. Collagen undergoes a lower critical solution temperature-like transition, forming a hydrogel when the temperature increases from 25 °C to 37 °C (Fig. 2A). [35] This transition facilitates in vitro fibrillogenesis at body temperature. The FIBs are incubated at 37 °C, allowing the collagen fibrils self-assembly. Noticed that collagen fibrils self-assembly is also obtained at different FIBs storage conditions (i.e. 4 °C, -20 °C, freeze-dried), suggesting an easy storage of the FIB solutions. Based on its composition, it is expected that WJ-ECM hydrogel results from the heterotypic fibrils (i.e. co-assembly of type I and type III collagen) while the RT-ECM hydrogel is formed by the homotypic fibrils assembly. Indeed, the multi-scaled characterization of both hydrogels reveals major differences. Macroscopically, WJ-ECM hydrogel is semi-transparent while the RT-ECM hydrogel is opalescent. When the WJ-ECM hydrogel is covered on a cardboard, the text of “BIOS” on the cardboard is clearly observed, indicating its high transparency (Fig. 2B). The light transmission data, assessed in the wavelength range of 300-700 nm, shows an average transmittance of the WJ-ECM hydrogel at 650 nm higher than that of RT-ECM with a transmittance, i.e. > 90 % versus 40 %, respectively (Fig. SI-2 A).

Typical in vitro fibrillogenesis proceeds through an initial lag phase, rapid growth of collagen fibrils, and a plateau phase during which the collagen fibrils mature and stabilize. The rate of assembly of fibrils, monitored by measuring the turbidity, shows sigmoidal curves with sharper slopes and shorter lag period and time of half gelation (t₁/₂) for RT-ECM hydrogel in comparison to WJ-ECM one. Specifically, the WJ-ECM hydrogel displays gelation ~ 7 min and t₁/₂ of ~ 19 min after incubation at 37 °C, while the RT-ECM hydrogel exhibits gelation time of few seconds and t₁/₂ of ~ 12 min (Fig. SI-2B). The rheological characteristics of the hydrogels showing that storage modulus (G') arose increasing later in WJ-ECM hydrogel (~ 16 min) versus RT-ECM (~ 7 min). The maximum G' is 129 ± 21 Pa for WJ-ECM hydrogel and 160 ± 9 Pa for RT-ECM hydrogel (p < 0.001, t-test, Fig. 2C). Taken together, these results reveal a longer nucleation phase in WJ-ECM compared to RT-ECM hydrogels.

Collagen hydrogels is considered as biphasic, [38] consisting of a loosely connected network of collagen fibrils filled with a large excess of interstitial fluid. The *in vitro* fibrillogenesis degree (FD), the gel fraction and the water holding capacity (WHC) of hydrogels are also assessed and the results show that the WJ-ECM hydrogel exhibited a lower FD in comparison with RT-ECM hydrogel (82 ± 1.15 and 85 ± 0.2 %, p < 0.0001, t-test, Fig. 2D) while the gel fraction is close in both hydrogels (i.
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Characterization of Wharton's jelly and rat tail tendon hydrogels. A: Methodological procedure for hydrogel formation. B: Macroscopic views of WJ-ECM (left red box) and RT-ECM (right blue box) hydrogels, showing a transparent WJ-ECM hydrogel. C: Evolution of storage modulus (G') of WJ-ECM (red dots) and RT-ECM (blue dots) hydrogels at 37 °C, showing a slower self-assembly along with a softer WJ-ECM hydrogel. D: *In vitro* fibrillogenesis degree (FD) and E: Water holding capacity (WHC) of WJ-ECM and RT-ECM hydrogels (n = 4, t-test). F: Kinetic of deswelling (syneresis) and G: Protein release after two (full bars) and six days (dashed bars) of incubation of WJ-ECM (red bars) and RT-ECM (blue bars) hydrogels in culture medium, revealing a lower deswelling of WJ-ECM hydrogel. H: Water contact angle on WJ-ECM and RT-ECM hydrogels. I: Centrifugation-assisted syneresis and J: macroscopic views of hydrogels after centrifugation of WJ-ECM (red arrow) and RT-ECM (blue arrow) hydrogels, confirming a higher syneresis in RT-ECM hydrogels. K: Differential scanning calorimetric profile of WJ-ECM and RT-ECM hydrogels. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

e. 2.09 ± 0.24 for WJ-ECM and 1.83 ± 0.18 for RT-ECM hydrogel, p = 0.37, t-test, Fig. SI-2C). Interestingly, the WHC is around 97 % for both hydrogels (Fig. 2E), suggesting the absence of differences in the ultrastructure (nanopores) and the capillary forces within the two hydrogels. [39,40]

For clinical applications, when a hydrogel is implanted into a tissue defect, it is important that it does not swell excessively, as prolonged swelling may cause increased pressure on the surrounding tissue. [41] However, collagen hydrogels in a physiological environment can exhibit deswelling (i.e. syneresis) and degradation behavior. [42,43] In this study, the mass loss of WJ-ECM and RT-ECM hydrogels was assessed at 37 °C in two different media, i.e. DMEM and PBS (Fig. SI-2D). The mass of the hydrogels decreases sharply after 24 h of incubation in both media, reaching equilibrium after 48 h (Fig. 2F and Fig. SI-2D). Although not visible to the naked eye (Fig. SI-2E), the mass loss rate of the WJ-ECM hydrogel is significantly lower than that of the RT-ECM hydrogel, with values of 57 ± 3 % and 80 ± 2 % of their initial

weight at 48 h, respectively (p < 0.0001, two-way ANOVA). The mass loss of the hydrogels may be attributed to degradation, deswelling, or the release of un-crosslinked proteins (i.e., pepsin) that leach out from the hydrogels. To assess potential degradation, protein content in the surrounding media was measured. The results reveal the presence of proteins, but only in trace amounts (~ 0.2 % of the total protein content in the hydrogels, Fig. 2G, SI-2F and SI-2G). MALDI TOF/TOF mass spectrometry reveals the presence of collagens in WJ-ECM and RT-ECM supernatants. Interestingly, a release of fibronectin is observed in WJ-ECM supernatant. (Table SI-3 and SI-4). These released ECM components are not bound to the hydrogel network, and thus, are not contributing toward the integrity of the hydrogels. These findings suggest that the hydrogel mass loss is primarily due to deswelling (i.e. syneresis upon aging without external stimuli [43]) rather than degradation, supporting the suitability of WJ-ECM hydrogels for cell culture applications. Syneresis is the process of contraction in hydrogel volume accompanied by expulsion of the water that was initially entrapped by
the self-assembled network. Enhanced hydrophobicity of the hydrogel results in increase in hydrophobicity of the self-assembled fibers followed by expulsion of water from the network. [43] Contact angle analysis, used to determine the hydrophilic properties of hydrogels, shows that the WJ-ECM hydrogel has a more hydrophilic surface compared to the RT-ECM hydrogel, with values of 29 ± 1.44° and 69 ± 2.28°, respectively (p < 0.0001, t-test, Fig. 2H). To confirm the higher syneresis in RT-ECM hydrogel, we next examined the differences in the hydrogel behavior following centrifugation (250 g). The syneresis is lower in WJ-ECM hydrogel compared to RT-ECM hydrogel (0.22 ± 0.04 % vs 25 ± 8 %, p = 0.02, Fig. 2I and J), indicating a higher hydrophobicity of the self-assembled homotypic collagen fibrils versus heterotypic collagen fibrils. DSC profiles, which show the thermal behavior of hydrogels, indicate that WJ- and RT-ECM hydrogels exhibit an endothermic peak, indicative of ECM denaturation primarily due to the desorption of bound water from the hydrogels. The melting temperature is recorded at around 51 °C for both hydrogels (Fig. 2K).

We above reported a higher GAG content in the WJ-ECM extract compared to the RT-ECM extract. The collagen/hyaluronic acid and collagen/GAG ratios in the WJ-ECM extract are approximately 11 and 21, respectively, which are lower than those in the RT-ECM extract (31 and 95, respectively). Thus, the lower FD and the longer duration of the nucleation phase observed in WJ-ECM hydrogels could be attributed to the higher GAG content in the WJ-ECM extract. These findings also suggest that the hydrogen bonding interactions between collagen fibrils and hyaluronic acid are stronger in WJ-ECM hydrogels compared to RT-ECM hydrogels. [44,45] This raises the question of the role of GAGs in the WJ-ECM FIBs during hydrogel formation. To investigate this, the gelation process of WJ- and RT-ECM FIBs was carried out in the presence of collagenase (which promotes collagen degradation) and hyaluronidase (which promotes GAG degradation). The treated FIBs were then placed in individual tubes and gently inverted after 40, 60, 90, and 120 min of incubation at 37 °C (Fig. SI-3). As expected, in the presence of collagenase, WJ- and RT-ECM FIBs lose their structure and flow down, confirming the absence of collagen self-assembly. However, in the presence of hyaluronidase, the RT-ECM hydrogel is able to maintain its structure after 40 min of incubation at 37 °C, whereas the WJ-ECM self-standing hydrogel is observed after 120 min. These findings suggest that hyaluronidase digestion generates smaller GAG fragments, delaying fibril precipitation, indicating the presence of large GAGs in WJ-ECM FIBs. [45]

The content in GAGs and the presence of type III collagen are known to influence the number density and diameter of collagen fibrils. [45,46] It is well established that when the nucleation phase is prolonged, the resulting collagen fibrils have smaller diameters, whereas a short nucleation phase produces larger fibrils. [47] In this study, the size of the collagen fibrils was assessed using single-scan polarization-resolved second harmonic generation (SS-pSHG), transmission electron microscopy (TEM) and atomic force microscopy (AFM). The SS-pSHG signal, indicating noncentrosymmetric packing of collagen triple helices within the hydrogels, reveals uniform distribution of thin heterotypic WJ-ECM collagen fibrils, whereas an increased backscattered SHG signal is observed in the homotypic RT-ECM collagen fibrils (Fig. 3A), consistent with the above-macroscale observations. At the fibril scale, TEM and AFM microscopies reveal that the diameter of the collagen fibrils in the WJ-ECM hydrogel is three times smaller than in the RT-ECM hydrogel (p < 0.0001, t-test, Fig. 3B-D). AFM, one of the few techniques that can assess D-band periodicity [48] shows that the D-banding distance of collagen in the WJ- and RT-ECM hydrogels is 55 ± 5 nm and 68 ± 1 nm, respectively (Fig. 3E). These results may be partially explained by the presence of GAGs, which coat the fibrils and promote the formation of highly hydrated collagen fibrils. The lower hydration level in RT-ECM may lead to fibril fusion and to the formation of thicker ‘fibrils’. [42] It well known that type III collagen interact axially with type I collagen. In addition, the WJ-ECM extract consist of atelocollagen molecules, in which both the N-terminal and C-terminal telopeptides are removed.

However, it has been reported that N-terminal cleavage in type III collagen is slow. [49] Therefore, the presence of the N-terminus may hinder the radial packing of collagen fibrils, thereby affecting their shape and diameter. [49]

The intermolecular distances between tropocollagen molecules are closely linked to water content. [25] Water is crucial in mediating the stability and self-assembly of the collagen. To further investigate and explore the potential impact of collagen hydration on *in vitro* fibrillogenesis, molecular dynamics simulations were performed (Fig. SI 4 A). Type I and type III collagen microfibrils were simulated under ambient lateral pressure (1 atm) and physiological temperature, and several structural parameters were sampled over a 0.5 ns trajectory. A quantitative comparison of structure, dynamics, hydrogen bonding, and internal stress between type I and type III collagen fibrils is presented in Figs. 3F to 3I, respectively. Focusing on the radial distribution function of alpha-carbon atoms (gcc)—both part of the protein backbone—the signal is observed between 10 Å and 20 Å, indicating intermolecular rather than intramolecular distances (Fig. 3F). Distinct peaks corresponding to the spacing between tropocollagen molecules within the fibril are observed at 12 Å, 15 Å, and 18 Å for type I collagen, consistent with experimentally determined distances. [50] In contrast, the type III collagen fibrils exhibit broader and less defined peaks, especially the first and third ones, shifted to 13 Å and 19 Å, respectively. These observations suggest a less ordered and less densely packed arrangement of tropocollagen molecules in type III fibrils. Further insight is gained from the radial distribution of water oxygen atoms around alpha-carbon atoms, which reflects the organization of water molecules surrounding the fibrils (Fig. 3G). Type I collagen showed a sharp peak at ~4 Å and a broader peak at ~5.5 Å, corresponding to the two-layered hydration shell previously described. [51] For type III collagen fibrils, the second peak is notably broader and less pronounced, suggesting a less structured and likely less dense hydration shell. This difference may explain the increased intermolecular distances seen in type III fibrils.

To assess fibril dynamics, the root mean squared displacement (RMSD) of protein backbone atoms was calculated at 1 ps intervals, along with the number of hydrogen bonds formed between water and tropocollagen molecules (Fig. 3H and I). Type III tropocollagen molecules exhibit reduced mobility and a higher number of hydrogen bonds with surrounding water molecules. Overall, these comparative simulations support the hypothesis that type III collagen fibrils exhibit looser and less ordered molecular packing than type I collagen fibrils. This structural difference likely allows more space for water-tropocollagen interactions in type III fibrils, which accounts for the higher hydrogen bond count, increases water-mediated drag, and reduces molecular mobility. Thus, the molecular dynamics simulations suggest a reduction in water-collagen interactions in homotypic fibrils as compared to heterotypic fibrils, leading to less axial packing.

Fibrils with a larger diameter improve the rigidity of the hydrogel, while smaller diameter fibrils favor interactions with ECM components, reducing plastic deformation and increasing the elasticity of the hydrogel. [36,52] As described earlier, both WJ-ECM and RT-ECM are biphasic, with a randomly oriented network of homo (RT-ECM) or heterotypic (RT-ECM) collagen fibrils filled with a large excess of water. According to biphasic theory, under compressive loads, the applied stress is initially supported by fluid pressure before being transferred to the fibril network. The rate of this transfer is determined by the permeability and stiffness of the collagen matrix. [38] As expected, the elastic modulus (E) of the WJ-ECM hydrogel (with thinner collagen fibrils) is lower compared to the RT-ECM hydrogel (with larger fibrils) with E = 1.09 ± 0.05 kPa for WJ-ECM and 1.40 ± 0.02 kPa for RT-ECM (p = 0.02, Mann-Whitney test, Fig. 4J and SI-4). Rheological testing to determine G' and confined compression to determine E of both hydrogels show that the mechanical properties from the independent methods are in very good agreement (WJ-ECM < RT-ECM), albeit in different magnitude orders (hundreds of Pa to kPa). Furthermore, when fully swollen hydrogels undergo deformations that affect local volume
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**Fig. 3.** Ultra-structural and mechanical characterization of Wharton's jelly and rat tail tendon hydrogels. A, B and C: Single scan polarized-resolved second harmonic generation, transmission electron microscopy and atomic force microscopy images of WJ-ECM (upper red rectangle) and RT-ECM (lower blue rectangle) hydrogels, respectively, confirming differences in collagen fiber self-assembly (red and blue arrows indicate WJ- and RT-derived collagen fibrils, scale bars = 50 µm, 0.2 µm and 1 µm). D and E: Diameter of fibrils and D-bending periodicity, showing that WJ-ECM hydrogel results from thinner fibrils auto-assembly (n > 50, t-test). F-I: Molecular dynamics simulation of the relaxed structure of the type I and type III collagen microfibril. F: Radial distribution function of alpha-carbons in the protein backbone, focusing on the peaks between 10 Å and 20 Å associated with tropocollagen distances. G: Radial distribution function of water oxygen atoms around alpha-carbons. H: Root mean-squared displacement of the atom in the protein backbone for 2 ps. I: Hydrogen bond count between water molecules and the protein. J and K: Elastic modulus (E) and permeability (k) of WJ-ECM and RT-ECM hydrogels determined by confined compression tests (n = 5, t-test). L: Macroscopic views, showing the injectability of WJ-ECM (left) and RT-ECM (right) FIBs. M: Flow behavior of WJ-ECM and RT-ECM hydrogels at 37 °C, revealing a pseudoplastic behavior of hydrogels. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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**Fig. 4.** Characterization of Wharton's jelly and rat tail tendon freeze-dried sponges. A: Methodological procedure for sponge preparation. B: Scanning electron microscopy and C: x ray nano-tomography images of the WJ-ECM (left red box) and RT-ECM (right blue box) sponges (scale bars = 100 and 500 µm), showing porous scaffolds. D: Thickness, E: porosity and F: gravimetric swelling ratio results, indicating absorption superiority of the WJ-ECM sponge (n = 5, t-test). G: Raman spectra of WJ-ECM (red line) and RT-ECM (blue line) sponges. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

conservation and pore deformations, such as compression (Fig. SI-4B-C), water is forced through the pores. The WJ-ECM hydrogel exhibits lower fluid permeability (k), more than 8-times lower than that of the RT-ECM hydrogel ($k = 0.34 \times 10^{-11} \pm 0.04 \times 10^{-11} \text{ m}^4/\text{Ns}$ for WJ-ECM and $2.87 \times 10^{-11} \pm 0.49 \times 10^{-11} \text{ m}^4/\text{Ns}$ for RT-ECM; $p = 0.007$,

Mann-Whitney test, Fig. 4K). The permeability of the network, which enables fluid flow, is commonly controlled by the porosity. Although the WHC results show no significant differences in nanopore characteristics between the two hydrogels (Fig. 2E), the data suggest that the pores in the RT-ECM hydrogel are smaller, which may contribute to its increased
stiffness. Poisson's ratio, which quantifies the compressibility of the hydrogels, is also lower for the WJ-ECM hydrogel compared to the RT-ECM hydrogel ($\nu = 0.05 \pm 0.02$ for WJ-ECM and $0.21 \pm 0.01$ for RT-ECM; $p = 0.007$, Mann–Whitney test, Fig. SI-4D). These findings align with previous literature [53,54] which demonstrated that Poisson's ratio values for knee cartilage are related to the collagen/GAG ratio. Injectable or extrudable hydrogels have been gaining popularity, particularly in minimally invasive surgeries, [55] which are widely performed across various surgical specialties. The FIBs were loaded into syringes and injected into different shapes, as depicted in Fig. 3L. The shear-thinning properties of the hydrogels (i.e., the ability of the hydrogel to decrease in apparent viscosity with increasing shear rate, as when flowing through a nozzle) were also assessed. The results highlight that both hydrogels exhibit pseudoplastic behavior, as evidenced by a gradual decrease in apparent viscosity as the shear rate increased (Fig. 3M), which is an expected behavior for biopolymer suspensions and hydrogels. Contrary to the quasi-static assays, when WJ-ECM hydrogel behaves slightly weaker than the RT-ECM hydrogel (Figs. 2C, 3J), both gels present similar behavior under high deformation flow (Fig. 3M). Additional experiments were performed to determine whether collagen extracted from WJ, like that from RT, can exhibit lyotropic properties and thereby mimic the fibrillar geometries of native ECMs [29] as the collagen hierarchical microstructure in bone has been shown to play a role in bone regeneration. [30] At high concentrations beyond the liquid crystal threshold (see SI methods), birefringent nematic phases were indeed observed (Fig. SI-5). Notably, pure collagen typically gives rise to cholesteric phases at such concentration; the observed difference may be attributed to the presence of type III collagen and the removal of telopeptides in the WJ-ECM preparation. Further work will be dedicated to establish a complete phase diagram and characterize the conditions governing this self-organization. These findings strengthen the potential of Wharton's jelly-derived collagen as a versatile component for the design of various biomimetic structural biomaterials.

## 3.2. From hydrogels to 3D porous sponges

The formed hydrogels exhibited high water content and a very limited porous structure at the nanoscale. To create macropores for biomedical devices, freeze-drying processing was employed (Fig. 4A), providing high-quality and shelf-stable collagen-based sponges under mild conditions. [56] This process provides high porosity with low shrinkage. Scanning electron microscopy (SEM) and X ray tomography examinations of the WJ-ECM and RT-ECM sponges (Figs. 4B-C) reveal a porous structure with thicknesses of $2.03 \pm 0.2$ mm and $1.3 \pm 0.1$ mm, respectively ($p < 0.01$, Fig. 4D), which allows for greater water retention and enhanced cell infiltration. For biomedical applications, the porosity and swelling properties of sponges are considered critical features. [56] The porosity of the WJ-ECM and RT-ECM sponges is found to be approximately $91.8 \pm 1.9\%$ and $89.15 \pm 2.9\%$, respectively (Fig. 4E). While the gel fraction is the same in both hydrogels (Fig. 2D), our results indicate that the WJ-ECM sponges absorb twice as much water as the RT-ECM sponges, with gravimetric swelling ratios of $18 \pm 3$ and $11 \pm 0.3$, respectively ($p = 0.02$, t-test, Fig. 4F). It is interesting to notice this higher interaction of WJ-ECM with water is already demonstrated by the low hydrogel syneresis (Fig. 2I). The higher water absorption capacity of WJ-ECM sponges may be attributed to their biochemical composition, including GAGs and type III collagen content, as well as the presence of thinner collagen fibers that provide a larger surface area for water interaction. Raman spectroscopy was conducted to identify similarities between the WJ-ECM and RT-ECM spectra. Data for the main vibration bands are compiled in Table SI-5. As shown in Fig. 4G, the Raman spectra of RT-ECM and WJ-ECM exhibit the main characteristic bands of type I collagen. [57] The WJ-ECM Raman spectrum displays a prominent band at $1668 \text{ cm}^{-1}$, attributed to the amide I region. This band is primarily associated with peptide carbonyl

stretching vibrations, with additional contributions from C—N stretching and N—H in-plane bending. In contrast, the amide I band in RT-ECM is shifted to $1655 \text{ cm}^{-1}$, indicating a difference in the secondary structure or molecular organization of collagen. Such shifts in amide I frequencies are typically correlated with variations in collagen fibril packing, hydrogen bonding interactions, or hydration states. The amide III bands at $1275 \text{ cm}^{-1}$ and $1247 \text{ cm}^{-1}$ arise from N—H bending and C—N stretching vibrations. The intensity ratio between these two bands is known to be sensitive to mechanical stress and protein conformation. [58] Therefore, variations in this ratio between WJ-ECM and RT-ECM may indicate differences in collagen cross-linking, fibrillar alignment, and mechanical properties, [59] as previously described. A Raman band at $1448 \text{ cm}^{-1}$ is observed in both spectra, corresponding to CH₂ bending and deformation vibrations of collagen. The distinct Raman peak at $1004 \text{ cm}^{-1}$, attributed to the C—C stretching vibration of phenylalanine, appears more intense in the WJ-ECM spectrum compared to RT-ECM, confirming variations in collagen composition. Bands at $1610 \text{ cm}^{-1}$, $1586 \text{ cm}^{-1}$, and $1555 \text{ cm}^{-1}$ are also observed in WJ-ECM, corresponding to C=C stretching of aromatic amino acid residues (i.e., C=C aromatic ring vibrations of phenylalanine and tyrosine, phenylalanine and tryptophan, and tryptophan and tyrosine, respectively). Additional Raman bands at $1099 \text{ cm}^{-1}$ and $1129 \text{ cm}^{-1}$, attributed to C-OH vibrations, [60] are exclusively detected in the WJ-ECM spectrum. Furthermore, in line with the biochemical analyses, the $1034 \text{ cm}^{-1}$ band, assigned to S=O stretching vibrations of GAGs, [61] is more intense in WJ-ECM compared to RT-ECM. Notably, the carboxylate group signal at $1342 \text{ cm}^{-1}$ varies in intensity, indicating possible differences in HA content. [62] Compared to WJ-ECM, the Raman spectrum of RT-ECM exhibits unique bands at $1744 \text{ cm}^{-1}$ and $1301 \text{ cm}^{-1}$, characteristic of lipid and ester content, likely due to the absence of Triton treatment in tendon rat samples. In conclusion, the Raman data reveal structural and molecular differences between RT-ECM and WJ-ECM, particularly in collagen organization, aromatic amino acid composition, and GAGs content.

Most collagen sponges possess outstanding pro-coagulant properties due to their excellent porous structure and hygroscopic properties. Collagen sponges with a total porosity of up to 90 % and a water absorption capacity of 10-times their weight are expected to exhibit strong hemostatic effects. [63] Given the WJ-ECM sponge's porosity ($> 95\%$) and water absorption capacity (18 times its own weight), it is expected to be an ideal hemostatic material. The hemocompatibility of WJ-ECM was also explored here. Freeze-drying can increase the concentration of salts within the sponge, potentially leading to hemolysis. However, the results show that the hemolytic rate of both WJ-ECM and RT-ECM sponges is below 5 % (Fig. 5A), the threshold set by ASTM F 756–2000, indicating favorable blood compatibility. The hemostatic sponge facilitates blood clot formation, promoting blood cell adhesion, platelet activation, and the transformation of fibrinogen into fibrin, as observed in SEM (Fig. 5B). Macroscopically, WJ-ECM sponges are more effective in absorbing blood moisture compared to RT-ECM sponges (Fig. 5C). The blood coagulation index (BCI) of the ECM sponges was also determined. The WJ-ECM sponge has a significantly lower BCI compared to the RT-ECM sponge (two-times lower, $p < 0.001$, Mann Whitney test, Fig. 5D), suggesting that the WJ-ECM sponge is more efficient for coagulation, likely due to the larger amount of type III collagen. [46] Monitoring coagulation by measuring the evolution of the G' of blood samples in contact with the ECM sponges show that the WJ-ECM sponge results in a shorter reaction time (the time at which the storage modulus of the clot begins increasing) of 200 s versus 700 s for the RT-ECM sponge. Additionally, the WJ-ECM sponge produces clots that are six- times stiffer than those formed with the RT-ECM sponge ($p < 0.0001$, Mann Whitney test, Fig. 6E). In conclusion, the WJ-ECM sponge, with its superior hemostatic properties, could be an effective strategy for promoting wound healing and tissue repair, improving the prospects for in vitro and in vivo applications.
<figure><img src="image_20.png" /><figcaption>**Fig. 5.** Hemostatic properties of Wharton's jelly and rat tail tendon freeze-dried sponges. A: Methodological procedure for sponge hemocompatibility testing. B: Hemocompatibility results. C: scanning electron microscopy and D: macroscopic views of blood clot within WJ-ECM (left red boxes) and RT-ECM (right blue boxes) sponges (scale bars = 10 µm, *n* = 6, Mann Whitney). E: Blood coagulation index, F: Kinetic of coagulation and G: storage modulus of the clots, indicating a higher pro-coagulant property of WJ-ECM hydrogel (n = 6, Mann Whitney test). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

### 3.3. *In vitro* innate immune responses and *in vivo* biocompatibility

Although the collagen is known for its low immunogenicity, [33] hydrogels derived from ECMs can be recognized by neutrophils as damage-associated molecular patterns (DAMPs), potentially leading to the activation of immune cells (i.e., leukocytes) and triggering excessive inflammation and fibrotic encapsulation. [64] The innate immune response is a biological network responsible for protecting the host from foreign threats. Cytocompatibility of the hydrogels was first evaluated according to ISO/EN 10993 part 5 guidelines. The results show that fibroblasts viability remains above the 70 % threshold, which is considered the indicator of non-cytotoxic behavior (Figs. SI-6 A and B). Furthermore, dermal fibroblasts adhere and proliferate on the both hydrogels, demonstrating the suitability of WJ-ECM hydrogels as a cell culture platform (Fig. 6A). Neutrophils, known as ‘leader’ cells in host defense, can generate significant amounts of reactive oxygen species (ROS), leading to a vicious cycle of inflammation and oxidative stress. In this study, we investigated the neutrophil respiratory burst response (i.e., intracellular ROS accumulation) to WJ-ECM and RT-ECM hydrogels as well as the IL-8 production (Fig. 6B). To focus on neutrophil responses to the hydrogels, the results were normalized to the basal production of ROS in neutrophils, thus controlling for individual variability in neutrophil responses. Flow cytometry reveals a decrease in intracellular ROS accumulation upon contact with WJ-ECM hydrogel, whereas a significant increase (~30-time increase; *p* < 0.02, Mann Whitney test, Fig. 6C) in ROS accumulation is observed with RT-ECM hydrogel.

Additionally, a significant increase in IL-8 production is noted in both hydrogels compared to basal neutrophil levels (Fig. 6D). We previously demonstrated that the Wharton's jelly matrix can mitigate the effects of lipopolysaccharide (LPS) on neutrophils. [13] Thus, the synergistic effect of hydrogels and the LPS inflammatory stimulus was further assessed (Fig. 6B). In contrast to RT-ECM hydrogel, a minor cumulative effect of LPS and WJ-ECM hydrogel on ROS accumulation is observed (Fig. 6C). However, IL-8 production is lower in the presence of WJ-ECM hydrogel compared to RT-ECM hydrogel (Fig. 6D), suggesting that WJ-ECM hydrogel, as Wharton's jelly matrix, [13] can exert an anti-inflammatory effect.

In this study, we investigated the immunomodulatory effects of WJ-ECM and RT-ECM hydrogels on macrophage secretion. Following biomaterial implantation, monocytes are recruited to the site and differentiate into macrophages, which then respond to the implant. Macrophages are essential in the regenerative process through the secretion of various mediators. Over the past few years, two sub-populations of macrophages have been identified: (i) the M1 phenotype, which promotes a pro-inflammatory state, and (ii) the M2 phenotype, which promotes wound healing and tissue remodeling. [64] Here, U927-differentiated macrophages (M0) were cultured on both hydrogels. Macrophages adhere on both hydrogels, as shown in fig. SI-6C. Cytokine production was analyzed at 48 h (Fig. 6E). The data were normalized to DNA content and the basal production of M0 macrophages (Fig. 6F). A reduction in IL-1β production and an increase in IL-1ra (IL-1β antagonist) production are observed on both hydrogels. However,
<figure><img src="image_21.png" /><figcaption>**Fig. 6.** Biocompatibility and innate neutrophil and macrophage responses behavior in the presence of Wharton's jelly and rat tail tendon derived hydrogels. A: Epifluorescent microscopy images of dermal fibroblasts live/dead staining visualized after 4, 7 and 11 days, revealing the fibroblast proliferation over the time onto the WJ-ECM (upper red box) and RT-ECM (lower blue box) hydrogels (scale bars = 100 µm). B: Methodological procedure for evaluating the neutrophil response to hydrogels. C and D: Intracellular accumulation of ROS in neutrophils and IL-8 production by neutrophils in the presence of WJ-ECM and RT-ECM hydrogels without and with co-stimulation with LPS. Results were normalized to the basal response of neutrophils (n = 6, Wilcoxon-Mann-Whitney test). E: Methodological procedure for evaluating the macrophage response to hydrogels. F: Production of mediators by M0 U937-differentiated macrophage in the presence of WJ-ECM and RT-ECM hydrogels without and with co-stimulation with LPS. Results were normalized to the basal response of M0 macrophages (n = 6, Wilcoxon-Mann-Whitney test). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

macrophages cultured on the WJ-ECM hydrogel produce lower level of TNF-α, whereas no significant change on TNF-α production is observed on the RT-ECM hydrogel. Interestingly, IL-10, a potent anti-inflammatory mediator, is over produced on the WJ-ECM hydrogel but down produced on the RT-ECM hydrogel. In contrast, VEGF and TGF-β are down produced on the WJ-ECM hydrogel, while a slight

increase in the production of these factors is observed on the RT-ECM hydrogel. Taken together, these results suggest that WJ-ECM hydrogel reduces inflammation without paracrine stimulating fibroblast and endothelial cell recruitment, while RT-ECM hydrogel has a largely inert effect on macrophages.

Despite the low immunogenicity of umbilical cord tissue, [14] there
is always a risk of inflammatory responses following hydrogel trans-
plantation. The host's immune response to implanted biomaterials in-
volves a complex inflammatory cascade, which is followed by cellular
recruitment, collagen deposition, and the potential formation of cystic
cavities that wall off the foreign biomaterials. In this study, WJ-ECM and
RT-ECM hydrogels were implanted into the subcutaneous space of
immunocompetent rats to assess the immune reaction over time.
Immunocompetent rats were selected to ensure that immune responses
could be thoroughly evaluated. The results describe histological exam-
inations of the host's response for four weeks post-implantation, during
the chronic inflammation phase. All animals recovered well, with no
signs of pain after surgery. Healing of the skin incisions made for sub-
cutaneous graft implantation proceeds without any complications or
visible signs of inflammation or edema. Overall, the hydrogels are not
fully resorbed, and no significant inflammatory reactions or rigidifica-
tion are observed. Hematoxylin and eosin (H&E) staining of the WJ-ECM
(xenogenic) and RT-ECM (allogenic) hydrogels does not show the
presence of granulomatous tissue or fibrotic capsules (Figs. 7A-B). The
presence of CD68-positive cells (macrophages) within the hydrogels
indicates that the hydrogels were gradually being resorbed without
causing tissue necrosis (Fig. 7C). To confirm the absence of fibrotic
capsules, sections were stained with picrosirius red following standard
histological procedures. In combination with polarization microscopy,
picrosirius red staining is commonly used to study fibrosis. [65] Bright-
field images (Figs. 7E and F) reveal large amounts of collagen deposi-
tion, appearing as red-stained areas. However, polarized light

microscopy (Fig. 7E' and F') shows a red-orange birefringence, indica-
tive of random type I collagen fibrils. The absence of greenish-yellow
birefringence characteristic of type III collagen suggests that most
collagen fibrils does not form the typical complex networks of type I and
type III collagen. WJ-ECM and RT-ECM hydrogels appear black due to a
lack of birefringence. The observed cavitations within the hydrogels are
likely a result of ECM resorption mediated by macrophages. These
findings suggest that both WJ-ECM and RT-ECM hydrogels meet the
basic safety requirements for implantable hydrogels, demonstrating
limited adverse effects in the subcutaneous implantation model.

4. Conclusion

In summary, a sustainable and renewable WJ-ECM hydrogel was
developed and compared to RT-ECM. Our results revealed key structural
and functional differences between the two hydrogels. RT-ECM hydro-
gels, characterized by a shorter fibrillogenesis nucleation phase, formed
through homotypic type I collagen self-assembly. In contrast, WJ-ECM
hydrogels resulted from the self-assembly of heterotypic type I and III
collagen along with glycosaminoglycans, leading to thinner collagen
fibrils and a shorter D-banding pattern. Although both hydrogels
exhibited similar water-holding capacity, WJ-ECM hydrogels showed
lower deswelling and syneresis upon exposure to culture medium.
Rheological and mechanical characterizations demonstrated that WJ-
ECM hydrogels were softer and less permeable than RT-ECM hydro-
gels. Functionally, WJ-ECM hydrogels exhibited superior hemostatic

<figure><img src="image_22.png" /><figcaption>**Fig. 7.** Subcutaneous biocompatibility of Wharton's jelly and rat tail tendon derived hydrogels. Left panel showed WJ-ECM hydrogel and right panel showed RT-ECM hydrogel. Hematoxylin-eosin-Safran (HES) staining of paraffin-embedded explant at lower (A and B) and higher (A' and B') magnifications (scale bars = 150 and 50 µm). Higher magnifications correspond to dashed red and blue rectangles. CD68 immunohistochemical staining of the tissue in contact with the hydrogels (scale bars = 50 µm). Picrosirius red staining in bright field (E and F) and polarized (E' and F') microscopy (scale bar = 100). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
properties, reduced intracellular ROS accumulation in neutrophils, and
modulated inflammation by decreasing proinflammatory mediator
release while enhancing anti-inflammatory profile. *In vivo*, WJ-ECM
hydrogels elicited minimal adverse effects in a subcutaneous implanta-
tion model, reinforcing their potential as safe and effective biomaterials
for regenerative medicine. Thus, these findings provide valuable insights
into the development of biocompatible, human-derived ECM hydrogels
that could serve as viable alternatives to non-human ECM sources,
particularly for cell culture platform, drug delivery systems and regen-
erative medicine applications.

CRediT authorship contribution statement

**Anais Lavrand:** Writing – review & editing, Methodology, Formal analysis, Data curation. **Lorinne Adam:** Writing – review & editing, Methodology, Conceptualization. **Alexis Da Rocha:** Writing – review & editing, Formal analysis, Conceptualization. **Flora Lemaire:** Writing – review & editing, Data curation, Conceptualization. **Capucine Loth:** Writing – review & editing, Data curation, Conceptualization. **Adrien Baldit:** Writing – review & editing, Funding acquisition, Conceptualization. **Maxime Vasseaux:** Writing – review & editing, Investigation, Formal analysis, Data curation, Conceptualization. **Laurence Van Gulick:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Abdelilah Beljebbar:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Pedro Augusto:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Alexandre Berquand:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Chrystelle Salameh:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Nadine Nassif:** Writing – review & editing, Formal analysis, Data curation, Conceptualization. **Fouzia Boulmedais:** Writing – review & editing, Data curation, Conceptualization. **Cédric Mauprivez:** Writing – review & editing, Project administration, Formal analysis, Data curation, Conceptualization. **Esteban Brenet:** Writing – review & editing, Data curation, Conceptualization. **Halima Kerdjoudj:** Writing – original draft, Validation, Supervision, Resources, Project administration, Conceptualization.

Declaration of competing interest

The authors declare that they have no conflict of interest.

Acknowledgements

Institut Carnot MICA "DERIDAGE" and Agence nationale pour la
Recherche "HyCareMat" (ANR-23-51CE-0039-01) projects are
acknowledged for financial support. A. Lavrand was supported by a
doctoral fellowship of Grand Reims and Institut Carnot MICA.
Communauté urbaine du Grand Reims, Département de la Marne,
Région Grand Est and European Union (FEDER Champagne-Ardenne
2014-2020, FEDER Grand Est 2021-2027) are acknowledged for their
financial support to the Chair of Biotechnology of CentraleSupélec and
the Centre Européen de Biotechnologie et de Bioéconomie. Authors are
grateful to the staff of Reims Maternity Hospital for providing umbilical
cords and to the staff of pediatric surgery department of Reims hospital
for providing foreskin explants and to the "pole de medicine bucco-
dentaire" for providing human blood. The authors acknowledge Dr. J. B.
Vincourt from IMoPA, (UL-UMR 7365 CNRS) and the proteomics plat-
form of IBSLor, for amino acid and proteomics analysis, the staff of the
Core PICT (Dr. Christine Terryn and L. Wortham) for SEM and TEM
imagings, URCACyt (Dr. Sandra Audonnet) for flow cytometry analysis,
URCANim from university of Reims Champagne Ardenne, MécaRhéo
and MicroMat (UL - CNRS UMR 7239), and corresponding equipment
availability. Authors also thank Pr. Camille Boulagnon and Nicole
Bouland for histology technical support.

**Appendix A. Supplementary data**

Supplementary data to this article can be found online at https://doi.
org/10.1016/j.ijbiomac.2025.147552.

Data availability

Data will be made available on request.

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