# A Recombinant Elastic Peptide Rescues Elasticity From a Self-Assembled Dermal Sheet Model Treated With Ascorbic Acid

Chloé Chave, Jorgan Guard, Thibault Massias, Aurore Berthier, Julie

Leignadier, Jérôme Sohier, Romain Debret

► To cite this version:

Chloé Chave, Jorgan Guard, Thibault Massias, Aurore Berthier, Julie Leignadier, et al.. A Recombinant Elastic Peptide Rescues Elasticity From a Self-Assembled Dermal Sheet Model Treated With Ascorbic Acid. Experimental Dermatology, 2026, 35 (2), 10.1111/exd.70205. hal-05534902

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Submitted on 3 Mar 2026

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**A recombinant elastic peptide rescues elasticity from a self-assembled dermal sheet model treated with ascorbic acid.**

Chloé Chave<fnref n="1" />,<fnref n="2" />, Jorgan Guard<fnref n="1" />, Thibault Massias<fnref n="1" />, Aurore Berthier<fnref n="1" />, Julie Leignadier<fnref n="2" />, Jérôme
Sohier<fnref n="1" />, Romain Debret<fnref n="1" />*

*Corresponding author :

Romain DEBRET
LBTI UMR5305 CNRS/UCBL
8, avenue Rockefeller
69008 Lyon, FRANCE

Email : romain.debret@cnrs.fr

**Ethical statement:**

Abdominal adult skin explants were collected from the Biological Resource Center GCS/CTC
hosted by Hospices Civils de Lyon (agreement number: AC-2019-3476, 2nd July 2020).

**Contributions:**

C.C.: investigation (equal); methodology (supporting); writing – original draft (equal). J.G.: investigation (equal). T.M.: investigation (supporting). A.B.: investigation (supporting). J.L.: funding acquisition. J.S.: methodology (supporting); writing – review & editing. R.D.: conceptualization; project administration; methodology (lead); writing – original draft (equal).

<footnote n="1">¹ Laboratoire de biologie tissulaire et ingénierie thérapeutique, UMR5305 CNRS/Université
Claude Bernard Lyon 1, Lyon, France.</footnote>

<footnote n="2">² Lucas Meyer Cosmetics by Clariant, Toulouse, France.</footnote>
**Abstract:**

The extracellular matrix (ECM) plays a pivotal role in determining the structure and function of the skin. Collagen and elastin, in particular, are responsible for providing tensile strength and elasticity, respectively. However, imbalances in the components of the extracellular matrix (ECM) during tissue engineering often result in the deterioration of the mechanical properties and physiological relevance of dermal substitutes, in part due to the detrimental effect of ascorbic acid (AA) on elastic fiber biosynthesis. The objective of this study is to investigate the potential of a synthetic elastic protein (SEP) to improve ECM remodelling and to restore the equilibrium between collagen and elastin in reconstructed dermal tissues.

Primary fibroblasts monolayers were treated with increasing concentrations of SEP in the presence or not of AA without affecting cytotoxicity. Western blot and immunofluorescence analyses showed that in the presence of AA -which typically reduced elastin synthesis- SEP improved elastic fiber formation without affecting type I collagen assembly. Additionally, 3D dermal substitutes treated with SEP and AA were analysed at the ultrastructural scale showing a specific colocalization of SEP with fibrillin-rich fibrils. Finally, dynamic mechanical analyses were conducted to measure mechanical properties of decellularized ECM. In the presence of SEP, we observed an increase of 46% and 40% of elastic and Young's moduli respectively.

By restoring ECM integrity, SEP is emerging as a promising biomimetic tool for the development of 3D skin substitutes. Its application has the potential to improve the physiological relevance and therapeutic value of engineered skin tissues.

**Keywords:**

Fibroblasts, tissue engineering, type I collagen, elastin, elasticity
# 1. Introduction:

The extracellular matrix (ECM) plays a pivotal role in tissue architecture, development, and functionality. It is a dynamic and complex network composed of fibrillar structural proteins, glycosaminoglycans, and other biomolecules that provide mechanical support and regulate cell behaviour<sup>1,2</sup>. The ECM is particularly important in maintaining the structure and mechanical properties of connective tissues, such as the dermis in the skin, where it supports vital cellular functions such as migration, proliferation, and differentiation<sup>3,4</sup>. Disruptions in its composition, remodelling, or mechanical properties have been associated with a range of pathological conditions, including, fibrosis, and impaired wound healing<sup>5,6</sup>.

Although cell 2D monolayers have been widely used, they are unable to recreate the architectural complexity and mechanical signals of the ECM *in vivo*<sup>7</sup>. In contrast, 3D models offer a more realistic representation of the interactions between cells and the ECM, making it possible to study cell behaviour more accurately<sup>8,9</sup>. For example, various 3D skin models have been developed, including those based on collagen gels seeded with fibroblasts<sup>10</sup>, and self-assembled dermal models made with human fibroblasts before keratinocyte seeding<sup>11,12</sup>. However, without considering mechanical properties like stiffness and elasticity, the true functional relevance of these models remains questionable. This underlines the need for integrating biomechanical assessments to better mimic the native ECM and its influence on processes like wound healing and tissue remodelling. These mechanical features are usually addressed by the measurement of the Young's modulus, which ranges from 10 to 40 kPa, depending on the anatomical location and hydration level<sup>13–15</sup>. In the dermis, collagen and elastin are key components that provide tensile strength and elasticity to withstand mechanical stress and adapt to deformation. Lysyl oxidases (LOX) are essential enzymes for the formation and stabilization of collagen and elastic fibers. They catalyse the oxidative deamination of lysine residues, enabling crosslinking between amine groups to form mature fibers<sup>16</sup>.

In the development of 3D dermal models, ascorbic acid (AA) has been widely used for its role in enhancing collagen synthesis, as it is an essential cofactor for prolyl and lysyl hydroxylases, enzymes critical for collagen stability and maturation in addition to LOX activity<sup>17–19</sup>. However, although AA is effective in stimulating collagen production, it has a negative effect on elastin production<sup>20,21</sup>. Consequently, its systematic use in 3D dermal models can lead to an imbalance between collagen and elastin synthesis, affecting the global mechanical behaviour of the ECM. While collagen enhances the tensile strength and structural integrity of the dermis, the decrease in elastin can compromise the tissue's ability to withstand mechanical stress and deformation<sup>22–25</sup>.

In this study, we aim to address the imbalance between collagen and elastin in 3D dermal models by introducing a Synthetic Elastin Protein (SEP), a recombinant biomimetic protein produced in our laboratory<sup>26</sup>. Unlike traditional methods, SEP closely recapitulates biophysical and biological behaviour of endogenous tropoelastin and offers a promising approach to restoring elastin levels and enhancing the mechanical properties of reconstructed dermal sheets. By integrating SEP, we seek to improve elasticity and stiffness, ensuring these models better reflect the *in vivo* characteristics of the skin to support therapeutic applications.
**2. Materials and Method:**

**2.1. Synthetic Elastin Protein (SEP) production**

The SEP used in this study was produced as described by Boëté et al. (2022) with a primary sequence
as follows:

CDYKDDDDK{(VGVAPGVGVLPG)₆ AAAKAAAKAAK}₆(VGVAPGVGVLPG)₆ GGACLGKACGRKRK²⁶.

In short, SEP is a recombinant protein engineered to mimic native elastin, containing alternating
hydrophobic and lysine-rich crosslinking domains. The protein was expressed in E. coli and purified
using inverse transition cycling (ITC) to ensure purity. The final product was stored in 0.9 % NaCl at -
20 °C until utilisation.

**2.2. Cell culture**

Normal human dermal fibroblasts (NHDF) were obtained from woman abdominal biopsies ranging
from 20 to 45 years of age. Abdominal adult skin explants were collected from the Biological Resource
Center GCS/CTC (AC-2019-3476, 2nd July 2020) hosted by Hospices Civils de Lyon, in France, or from
Promocell (Lot number #C2511). The cells were cultured in Dulbecco's Modified Eagle Medium
(DMEM, Gibco-Invitrogen, Carlsbad, USA #41966-029) supplemented with 10 % (v/v) heat inactivated
fetal calf serum (Gibco-invitrogen) and antibiotics (100 U/mL penicillin and 100 µg/mL streptomycin,
Gibco, #15070-063). Cells were maintained at 37 °C and 5 % CO2 in a humid atmosphere with medium
renewal 3 times a week. The various studies were carried out using sub-confluent cells between the
second and the tenth passage. Cells were regularly tested for mycoplasma contamination using the
MycoAlert Mycoplasma detection kit (Lonza #LT07-118, Basel, Switzerland)

**2.3. Reconstruction of a human 3D dermal substitute**

Human dermal tissues were generated following the tissue engineering self-assembly approach. A
filter paper anchor (Whatman™, Life Sciences) previously cut to the appropriate dimensions (20 mm
external diameter and 13 mm internal diameter) was placed at the bottom of each well in a 12-well
plate (Thermo Fisher Scientific, #150628). The anchors were washed with PBS, Dulbecco's Phosphate
Buffered Saline (Sigma-Aldrich #D8537). Primary human dermal fibroblasts were seeded at a
concentration of 15 000 cells/cm² in the 12-well plates. After 1 week of culture, cells were treated or
not with SEP. Treatment was applied every other day for one week. Dermal sheets were cultured in
culture medium supplemented with 50 µg/mL AA (Sigma-Aldrich # A4034-100G). The medium was
changed 3 times a week, with a final culture time of 4 weeks. At the end of the incubation period, the
dermal sheets were washed and then incubated for 5 min at 37 °C with 10 mM EDTA (Sigma, #E9884-
500g) to facilitate their detachment.

**2.4. Cell viability**

Cytotoxicity was assessed using tetrazolium salt reduction, or MTT (Sigma-Aldrich #M2128-500MG),
as an endpoint. MTT was reduced by intracellular dehydrogenases in viable cells, leading to the
formation of purple formazan crystals that were insoluble in aqueous solutions. These crystals were
then dissolved in an organic solvent, and the absorbance of the resulting solution is measured.
Fibroblasts were seeded at a density of 5 000 cells/cm² and treated or not with different concentrations
of SEP. MTT solution (5 mg/mL) was added according to the supplier's recommendations. Optical
density was measured at a wavelength of 570 nm with subtraction of the second filter at 650 nm using
an AMR-100/T microplate reader (Allsheng, China).

**2.5. Human 3D dermal substitute viability**
Dermal substitutes were washed with PBS and incubated for 30 min at 37 °C in DMEM supplemented with 4 mM calcein-AM (Sigma #148504-34-1) and 2 mM propidium iodide (Sigma #25535-16-4). After washing twice with PBS, the dermal sheets were observed under a fluorescence microscope (EVOS-FL, ThermoFisher, Scientific).

## 2.6. Western blotting

Fibroblasts were grown at high density (50 000 cells/cm²) in 6-well plates (Thermo Fisher Scientific #140675) in complete medium. Once confluence was reached, cells were treated with 50 µg/mL AA, SEP (500 µg/mL) or both for 48 h. After the incubation period, complete medium was replaced with serum-free medium. After 48 h, conditioned media and total matrix proteins were harvested for soluble type I collagen and elastin analyses, respectively. The total matrix protein concentration was determined using a Pierce BCA Assay Kit (Thermo Fisher Scientific, #23225) according to the manufacturer's recommendations. The absorbance at 560 nm was measured. The proteins were denatured for 5 min at 95 °C in load buffer (62.5 mM Tris-HCl pH=6.8, 0.01 % w/v bromophenol blue, 10 % v/v glycerol, 2 % w/v SDS and 50 mM DTT), separated by denaturing polyacrylamide gel electrophoresis (SDS-PAGE) using prefabricated Mini-ProTEAN TGX gels (#4561094, Bio-Rad). Migration was performed using a Mini Protean III system (Bio-Rad) containing migration buffer (25 mM Tris-HCl, 192 mM glycine and 0.1% w/v SDS) at a constant voltage of 200 V. The proteins were then electroblotted onto polyvinylidene difluoride (PVDF) membranes using the Transblot Turbo Transfer Pack (Bio-Rad #1704158) combined with a transfer device. The PVDF membranes were then blocked for 1 h with tris-buffered saline (TBS) containing 0.1 % Tween 20 and 3 % bovine serum albumin (BSA). The primary antibodies were diluted in TBS containing 0.1 % Tween and BSA (1%) and incubated overnight at 4 °C: rabbit polyclonal anti-type I collagen (Novotec, #20111, 1:1000); mouse monoclonal anti-elastin (Merk, #MAB2503, 1:1000); rabbit polyclonal anti-FLAG (Sigma, #F7425, 1:1000). The membranes were incubated for 1 h at room temperature with goat anti-rabbit HRP conjugate (Bio-Rad #170-6516) or goat anti-mouse HRP conjugate (BioRad #170-6515) diluted 1:10000 in TBS, 0.1 % Tween 20. The membranes were then incubated for 5 min with ECL reagent (Thermo Fisher Scientific, #34580), and the antigens detected by chemiluminescence. The signal was recorded using the iBrightTH CL750 Imaging System (Invitrogen). Protein signals were evaluated using ImageJ software.

## 2.7. Indirect immunofluorescence staining

Fibroblasts (50000/cm²) were seeded onto microscope slides in culture medium. After 72 h, 50 µg/mL AA, 500 µg/mL SEP, or both, were added for a further 48 h. For β-Aminopropionitrile (βAPN, Sigma, #A3134-5G) treatment, fibroblasts were incubated with either SEP alone or SEP combined with 350 µM βAPN, and the cells were then fixed with 4 % formaldehyde for 10 min. Saturation with 5 % BSA was carried out and then different primary antibodies were incubated overnight at 4 °C: rabbit anti-lysyl oxidase-like 1 (Home-made, 1:100); rabbit polyclonal anti-fibrillin 1 (Sigma, #HPA021057, 1:100); rabbit polyclonal anti-fibronectin 1 (Abcam, #ab23750, 1:100); mouse monoclonal anti-elastin (Merk, #MAB2503, 1:100); rabbit polyclonal anti-type I collagen (Novotec, #20111, 1:100); rabbit polyclonal anti-FLAG (Sigma, #F7425, 1:100). The cells were then incubated for 1 h at room temperature with Alexa Fluor 488-conjugated IgG or Alexa 546-conjugated IgG secondary antibodies (1:1000) (Invitrogen, #A11034 #A11030). The nuclei were labelled with 2 µg/mL 4',6-diaminido-2-phenylindole (DAPI) and the cells were then mounted with PermaFluor (Thermo Fisher Scientific, #TA-030-FM). Images were acquired with a fluorescence microscope (EVOS-FL) and ImageJ software. For fluorescence quantification, images were acquired using a Cytation 1 multimode reader and GenV software (Agilent BioTek). Four wells per condition were analyzed and a total of nine images per well were captured, each separated by a distance of 700 µm. The mean gray value of each image was
extracted, and cell count was assessed for each field using ImageJ software to normalize fluorescence intensity to cell number.

## 2.8. Transmission electronic microscopy

Human dermal tissue was generated as described above and treated with 50 µg/mL SEP. After 4 weeks, dermal substitutes were fixed with 2 % glutaraldehyde for 15 min at 4 °C. The cells were then fixed with 2 % glutaraldehyde in sodium cacodylate buffer. Sample preparation, including dehydration, embedding, and ultrathin sectioning, was carried out by Ciqle, a specialized imaging. Co-labelling was performed with gold beads of two different sizes (5 nm for fibrillin-1 and 15 nm for the FLAG tag).

## 2.9. Decellularization of 3D dermal substitute

Once the incubation period was over, the dermal sheet culture medium was removed and washed 3 times with PBS. In order to eliminate the fibroblasts, the plates containing the dermal sheets were frozen at -80 °C for 30 min and then immersed directly in a water bath at 37 °C for 5 min, then rinsed with PBS. The different procedures were repeated three times. After the last wash, the dermal layers were stored at 4 °C in PBS.

## 2.10. Dynamic mechanical analysis (DMA)

The mechanical properties of dermal sheets treated or not with different concentrations of SEP (50 and 200 µg/mL) were analysed by uniaxial stretch with a dynamic mechanical analyser (DMA 242 E Artemis, NETZSCH, Germany). Following decellularization, the dermal sheets were dried overnight at room temperature. After drying, the dermal sheets were cut to the desired size (6 mm long by 4 mm wide) using a scalpel and placed into the DMA jaws. To rehydrate the samples, a tank containing 65 mL of PBS was inserted into the system, immersing the dermal sheets at a constant 22 °C temperature. Following a two-minute rehydration period, measurements were initiated at a frequency of 1 Hz for a 10 min duration, with the objective of measuring the storage modulus, which represents the elastic component of the material. To determine the Young's modulus of the samples, an increasing strain up to 200 µm was applied and the stress was recorded as a function of strain. The Young's modulus was calculated from the slope of the linear portion of the stress-strain curve, which corresponds to the elastic behaviour of the material. The thickness of dermal sheets was determined by confocal microscopy (Zeiss Confocal 880 NLO AiryScan) and a value of 15 µm was found for one tested sample in each condition. Hence a representative thickness of 15 µm was used for all tested samples.

## 2.11. Statistical Analysis

The statistical tests were performed using GraphPad Prism software. The normality and homogeneity of the variances were tested before carrying out the statistical tests. Kruskal-Wallis or one-way ANOVA were used to compare different groups between each other. For multiple comparisons a Dunn's or Tukey *post hoc* were used. Statistically significant values are p<0.05 *; p<0.01 ** and p<0.001 *** and are expressed as mean ± SD.
**3. Results:**

**3.1. Evaluation of the cytotoxicity and morphological effects of SEP on fibroblasts**

The cytotoxicity of SEP was assessed using the reduction of tetrazolium salt (MTT assay) to determine cell viability after 24, 48, and 72 h of treatment with varying concentrations of SEP (10–500 µg/mL) (Figure 1A). No significant reduction in cell viability was observed across the different conditions and time points, indicating that SEP did not induce cytotoxicity in fibroblasts. Interestingly, at 500 µg/mL after 24 h, statistically significant increase in metabolic activity was detected, suggesting a transient stimulatory effect of SEP at higher concentrations.

In light of the absence of toxicity, the potential impact of SEP on fibroblast morphology was evaluated after 72 h (Figure 1B). Bright field microscopy confirmed that no visible morphological alterations occurred at any tested concentration, further supporting the biocompatibility of SEP with this cell type.

**3.2. Effects of SEP and AA on ECM component synthesis and maturation by fibroblasts**

Previous findings have demonstrated that AA increases the production of type I collagen at the expense of elastin<sup>20</sup>. Based on this, we sought to investigate the impact of SEP and AA on the ability of fibroblasts to synthesize and mature extracellular matrix (ECM) components. Western blot analysis was performed to quantify type I collagen and elastin secretion under these different treatments separately or combined (Figures 2A and 2B).

Regarding type I collagen (Figure 2A), the immature form did not show significant difference between the considered treatments (Figure 2B). AA increased the production of the cleaved forms as expected compared to the untreated condition or in presence of SEP (Figure 2C, 2D). In contrast, SEP did not influence type I procollagen cleavage (Figure 2C), but induced a 50% decrease of the mature form (Figure 2D). This indicated that SEP did not alter the main effect of AA on type I collagen synthesis, but may slightly modify AA-induced collagen fibrillogenesis.

Elastin detection (Figure 2E) exhibited a distinct response pattern; as previously described, AA treatment reduced elastin production by fibroblasts (Figure 2F). While SEP did not influence the global type I collagen levels, its structural similarity to elastin led to an increase in the intensity of the band corresponding to elastin (Figure 2F). However, when fibroblasts were treated simultaneously with AA and SEP, no significant reduction in elastin intensity was observed when compared to SEP treatment alone. No significant effect of AA was observed when looking at anti-FLAG tag present on the SEP sequence (Figure 2G). This suggested that the presence of AA had no impact on the beneficial effect of SEP on elastin secretion, nor on the quantity of soluble SEP.

To assess the global organization of ECM components, immunostaining was conducted for type I collagen, elastin, and the SEP FLAG tag (Figure 2H). Fluorescence quantification mostly confirmed the observations made by Western blot analyses. Treatment with AA increased type I collagen assembly (Figure 2H a-b and 2I) but inhibited the formation of elastic fibers (Figure 2H e-f and 2J). On the contrary, SEP strongly stimulated the deposition of elastic fibers (Figure 2H e,g and 2J). However, the fluorescent signal did not discriminate elastin from SEP and the observable increase was rather due to the fibrillar incorporation of the SEP. Even though it was unexpected, SEP also improved the type I collagen assembly (Figure 2H a, c and 2I), indicating that the decrease of type I collagen mature form observed in Western blot had no negative consequence on collagen fibrillogenesis. Notably, the combined treatment with AA and SEP led to a weak but significant reduction of elastic fibers deposition compared to treatment with SEP alone (Figure 2H g-h and 2J), while no significant difference was
observed on SEP deposition as evidenced by the FLAG-tag labelling (Figure 2H k-l and 2K). Since AA did not affect the positive effect of SEP on the soluble form as demonstrated by Western blot, this suggested that AA may partially hinder the SEP-induced elastin assembly.

Additional analyses examined other key ECM proteins, including fibrillin-1, fibronectin, and lysyl oxidases (Figure S1), showed no observable changes in their secretion or assembly following the co-administration of SEP and AA.

To assess whether SEP integration into the ECM depends on lysyl oxidase activity, we treated post-confluent fibroblast monolayers with β-aminopropionitrile (βAPN), a specific inhibitor of lysyl oxidases (Figure 2L). In untreated conditions (Figure 2L a-d), a sparse network of elastic fibers was observed, reflecting baseline fiber formation. Treatment with SEP (Figure 2L e-h) demonstrated an association of SEP with elastin fibrillar deposits. However, when lysyl oxidase activity was inhibited (Figure 2L i-l), no association of the SEP with preformed elastic fibers was observed, thus confirming that lysyl oxidase activity was required for the association of SEP with elastin in the extracellular compartment.

All together, these 2D *in vitro* data suggested that the addition of SEP could address the challenge of elastin production during the fabrication of 3D dermal substitutes, enabling the restoration of baseline elastin levels in these models.

### 3.3. Impact of SEP on the morphology and ultrastructure of 3D dermal constructs

To evaluate whether SEP could attenuate the negative impact of AA on elastin production, we developed a 3D dermal substitute model. Dermal sheets were cultured for one month with AA, and SEP was applied at various concentrations during the second week. Macroscopic observations revealed concentration-dependent differences (Figure 3A). At a concentration of 50 µg/mL, only slight visible changes were observed, while at 200 µg/mL the extracellular matrix appeared denser, with increased whiteness and opacity. To further quantify these macroscopic differences, surface plots (Figure 3A, d,e, f) were generated based on grayscale intensity values from the macroscopic images. The corresponding line profiles (Figure 3A, g, h, i) provided a detailed representation of grayscale intensity across the dermal substitutes. The results showed more pronounced and notably broader peaks and troughs at 50 µg/mL and 200 µg/mL. The calculation of the aera under the curve (sum of gray level averages per pixel row), reflecting marked variations in surface texture, indicated a 30 % and 95 % increase toward whiter values for 50 and 200 µg/mL respectively. These analyses supported the visual observation of increased matrix density and structural organization at higher SEP concentrations. This likely reflected enhanced extracellular matrix deposition and structural reorganization. These observations tended to confirm that SEP integrates into the ECM of the dermal constructs. This integration hintde at a broader impact of SEP on the functional properties of the ECM, opening perspectives on how it may influence tissue mechanics.

The macroscopic differences were corroborated by phase contrast microscopy analyses performed after one month of culture (Figure 3B). At a concentration of 50 µg/mL, arrows highlight the initial presence of SEP within the ECM, visible as small dot-like structures. At 200 µg/mL, SEP accumulation became more evident, with a clear enhancement in its incorporation. These observations highlighted the dose-dependent effect of SEP treatment on ECM structural properties.

Fibrillin-1 is a structural glycoprotein that plays a crucial role in the formation of the microfibrillar scaffold, which is essential for the assembly of elastic fibers. These microfibrils provide a model for tropoelastin deposition and are essential for the organisation and stabilisation of elastic fibers. To
confirm whether SEP integrated with the elastic fibers, the ultrastructure of the dermal substitute was examined using transmission electron microscopy (TEM). Co-immunolabelling was performed with gold beads of two different sizes (5 nm for fibrillin-1 and 15 nm for the FLAG tag) to facilitate the detection and localisation of these components within the matrix. TEM analyses provided further evidence of SEP integration into elastic fibers within the dermal substitutes (Figure 3C). Images at different scales revealed consistent co-localisation of SEP with fibrillin-rich microfibrils, while no association was observed at collagen fibers level. These results provided definitive evidence of SEP's incorporation into elastic fibers and reinforced a potential role as an ECM structural component of treated dermal sheets.

**3.4. Decellularization efficiency and effect on type I collagen organization**

In order to characterize the contribution of SEP to the mechano-structural properties of the ECM, a prerequisite was the decellularization of dermal substitutes to avoid the influence of cells' tensile forces that impact the global tissue behaviour. To evaluate the efficiency of the decellularization process, we performed live/dead assays in the absence of decellularization (Figure 4 a-c), and after decellularization (Figure 4 g-i), in the presence or not of SEP. The results clearly demonstrate that decellularization has induced total cell death (red cells) compared to the non-decellularized control where all cells were alive (green cells). In the same conditions, we investigated the effect of decellularization on the structural organization of the type I collagen, representing the global ECM structure. Anti-type I collagen labelling revealed that decellularization exhibited a dense, well-defined collagen fibrillar network similar to that of the non-decellularized control matrix (Figure 4 d-f and j-l), suggesting that the structural integrity of the ECM was preserved. Moreover, SEP treatments did not affect neither the cell viability, nor the collagen organization during the decellularization process.

**3.5. Dynamic mechanical analysis of 3D dermal sheets**

Dynamic mechanical analysis (DMA) was conducted to evaluate the effect of SEP on the elasticity of the reconstructed dermal model. After confirming that SEP integrated into elastic fibers and that the dermal substitutes maintained their structural integrity in a decellularized state, we evaluated whether this integration improves the mechanical properties of the ECM. The elastic moduli (E') were measured over 10 min stretching period at a frequency of 1 Hz (Table 1). In these conditions, the treatment with 50 µg/mL SEP did not significantly modify the elastic modulus, while a 200 µg/mL SEP treatment led to a significant increase in elasticity from 15 kPa to 22 kPa (+ 46 %).

This increase in elasticity was further corroborated by the determination of the Young's moduli, calculated from the strain-stress relationship applied to the samples (Table 1). A significant increase of the Young's modulus was observed following treatment with 200 µg/mL SEP, increasing from 5 kPa to 7 kPa (+ 40 %).
**4. Discussion:**

Many studies have successfully developed advanced dermal substitutes incorporating AA to promote collagen production, but these often neglect the critical role of elastic fibers<sup>27-29</sup>. In this study, we have demonstrated that SEP significantly increases both the elastic modulus (E') and the Young's modulus from self-assembled dermal sheets. The decrease of skin Young's modulus coupled to an increase of the ratio of dissipated energy have been associated to ageing and defective elastic fibers in the cutis laxa syndrome *in vivo*<sup>30</sup>. Unfortunately, it was not possible to calculate the ratio of dissipated energy due to very low values of the loss modulus (E") in our conditions (not shown). However, the increase in both Young's and elastic moduli indicates that the dermal sheets are both stiffer and more deformable in the presence of SEP. Recent studies have reported the use of a new sodium derivative of AA, alone or in combination with other antioxidants, in topical formulations. This compound is said to maintain the same positive effects on collagen synthesis as commercial versions of AA, while preserving the integrity of elastic fibers. The compound even showed a tendency to increase elastin biosynthesis in skin explants<sup>31,32</sup>, and in a cohort of volunteers when applied with serum formulation twice a day during three months<sup>33</sup>. However, there is currently no evidence of any effect on the mechanical properties and, consequently, functionality of the elastin-rich fibrillar deposits observed.

The measurement of mechanical properties is affected by exacerbated cell contraction in self-assembled dermal substitutes<sup>34</sup>. In order to focus on ECM characterization and eliminate the traction effect of cells in our experimental conditions, we used protocols adapted from the literature to obtain a decellularized matrix<sup>35-38</sup>. Despite the implementation of various washes, the presence of chromatin, marked by propidium iodide, was still evident in the Live/Dead assay. This suggests that the nuclei may be resistant to decellularization treatments. However, TEM images indicate that the plasma membranes have disappeared (Figure S2), which prevented the cell-matrix contact as expected. To ensure the integrity of the ECM following decellularization, the matrix was first evaluated microscopically, where no apparent disruptions in its overall structure were observed. However, ultrastructural analysis using TEM (Figure S2) revealed damages in the collagen fibril network, indicating that the decellularization process compromised its ultrastructural organization. This highlights the importance of conducting detailed analyses beyond standard microscopy to fully assess the impact of decellularization on the ECM. Similar observations have been reported previously showing heterogenous collagen density at the sites of decellularization<sup>39</sup>, which could be minimized by the use of cryoprotectants<sup>40</sup>. In turn, the modifications at the ultrastructural scale may diminish the mechanical properties of the ECM, and the effects of the SEP may also be under-estimated in these conditions. Additionally, decellularization may influence other functional aspects of the ECM beyond mechanical properties, notably in the case of recellularization for tissue reconstruction purpose, emphasizing the need for comprehensive assessments.

The ultrastructural analysis of non-decellularized dermal substitutes treated with SEP provided critical insights into its integration into elastic fibers. The observed colocalization of SEP with fibrillin-1-rich fibrils is a determining point of our study regarding the roles of microfibrils in elastin polymerization and further elastic fiber assembly. Fibrillin-1, a major structural glycoprotein of microfibrils, serves as a scaffold for tropoelastin deposition, facilitating elastic fiber maturation by lysyl oxidase crosslinking activity and mechanical resilience<sup>41-45</sup>. The biomimetic incorporation of SEP into elastic fibers by a lysyl oxidase dependent activity, since it is counteracted by the βAPN inhibitor, reinforces its potential to mimic the natural function of elastin, contributing to ECM integrity and elasticity.

The biomimetic properties of SEP distinguish it from elastin-derived peptides, which primarily activate signalling pathways and have limited functional integration into the ECM<sup>46</sup>. SEP mimics the biological role of tropoelastin, offering structural and mechanical advantages. It is noteworthy that a transient
increase in metabolic activity was observed at the highest concentration (500 µg/mL) after 24 h. This effect may be linked to the presence of the VGVAPG sequence within SEP, a motif known to promote cellular activity and proliferation, as previously observed with elastin-derived peptides on fibroblasts ⁴⁷. We assume that a small amount of SEP would have been degraded by proteases present in the medium to generate such active peptides. However, this increase was no longer detectable after 48 h, potentially because fibroblasts recognised SEP as a structural protein and incorporated it into the ECM structure that protected it from further degradation. This is consistent with our prior findings which have demonstrated that SEP remained quite inert in other cell types, such as HUVECs and SMCs ⁴⁸. Microscopic analysis revealed no discernible fibroblast morphological alterations across all tested concentrations, indicating that SEP did not disrupt cellular functions or structural integrity. This observation is consistent with a structural role within the ECM as observed with tropoelastin, which contributes to matrix assembly without influencing cellular signalling ⁴⁹. Together, these properties make SEP particularly promising for applications in cell therapy and tissue engineering, where maintaining cell viability, structural integrity, and mechanical functionality is critical.

Finally, the treatments with AA and SEP reveal an intriguing antagonistic interaction regarding the secretion of type I collagen and tropoelastin soluble forms. While SEP alone decreased the mature form of type I collagen, it enhanced tropoelastin. In contrast, AA stimulates type I collagen synthesis while reducing elastin production, thereby disrupting the mechanical balance of the ECM as previously demonstrated ²⁰. The co-administration of SEP together with AA restored elastin release levels to a baseline similar to untreated conditions, and preserved beneficial effects on type I collagen fibrillogenesis. This interaction may result from competition between SEP and AA during critical processes, such as lysyl oxidase-mediated crosslinking or elastic fiber assembly. We showed that, although AA reduces elastic fiber formation, it does not affect lysyl oxidase distribution, which remains similar in the presence or absence of the vitamin (Figure S1). This highlights that AA exerts opposing effects on two major ECM components – elastin and collagen ⁵⁰ – without directly altering lysyl oxidase expression or distribution. As suggested by Davidson et al. and Faris et al., AA decreases elastin mRNA stability while it stabilizes type I collagen mRNA ⁵¹,⁵². Hence, lysyl oxidase activity may be more reflective of enhanced collagen metabolism induced by AA rather than a direct effect on enzymatic activity.

**5. Conclusion:**

In conclusion, this study proves that SEP displays intrinsic characteristics for enhancing 3D dermal models by adjusting their mechanical properties and improving their physiological relevance. Combined treatments with SEP and AA in equivalent dermal models highlight the potential to optimize the balance between collagen and elastin, further refining the functionality of engineered skin tissues. This dual strategy represents a significant advancement in tissue engineering and regenerative medicine. However, the molecular mechanism allowing SEP to enhance mechanical properties remains an intriguing area of investigation. While it is clear that SEP deposition is dependent of lysyl oxidase activity, it is not yet known whether it facilitates the neoformation of elastic fibers, or if it predominantly incorporates into pre-existing structures. Further studies are needed to clarify these mechanisms and to explore potential interactions between SEP and other ECM components.
**Acknowledgements:**

We acknowledge the contribution of SFR Santé Lyon-Est (UAR3453 CNRS, US7 Inserm, UCBL) facility: CIQLE (a LyMIC member), especially Elisabeth Errazuriz for her help in transmission electron microscopy acquisition and immunogold sample preparation. We also acknowledge the contribution of SFR Biosciences (Université Claude Bernard Lyon 1, CNRS UAR3444, Inserm US8, ENS de Lyon) ANIRA ImmOs - Seahorse platform, especially Laurence Canaple, for assistance with the Cytation 1.

**Conflicts of Interest:**

The authors declare that they have no particular conflict of interest.

**Data Availability Statement:**

The authors confirm that the data supporting the findings of this study are available within the article and its supplementary materials.
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<caption>**Table 1. Mechanical properties of reconstructed dermal substitutes treated with SEP.**</caption>

Comparison of mean values (± standard deviation) for the elastic modulus (E') measured using dynamic mechanical analysis (DMA) at 1 Hz for 10 min and Young's modulus determined through strain-sweep analysis under untreated, SEP 50 µg/mL, and SEP 200 µg/mL conditions. Statistical analysis was performed using the Kruskal-Wallis test compared to the untreated group. \*p < 0.05; \*\*p < 0.01.

<table><thead><tr><th></th><th>E' (kPa)<br>mean ± SD</th><th>Adjusted<br>p-value</th><th>Young Modulus<br>(kPa) mean ± SD</th><th>Adjusted<br>p-value</th></tr></thead><tbody><tr><td>Untreated<br>(n=25)</td><td>15.61 ± 4.3</td><td></td><td>5.09 ± 1.9</td><td></td></tr><tr><td>SEP 50 µg/mL<br>(n=22)</td><td>15.78 ± 5.4</td><td>ns</td><td>5.17 ± 2.1</td><td>ns</td></tr><tr><td>SEP 200 µg/mL<br>(n=7)</td><td>23.04 ± 2.7</td><td>\*\*; 0.004</td><td>7.11 ± 1.9</td><td>\*; 0.038</td></tr></tbody></table>
A

<figure><img src="image_2.png" /></figure>

B

<figure><img src="image_3.png" /><figcaption>**Figure 1.** Evaluation of the cytotoxicity and morphological effects of SEP on fibroblasts.</figcaption></figure>

(A) Cell viability was assessed by MTT assay after treatment with different concentrations of SEP (10–500 µg/mL) during 24, 48, and 72 h. DMSO (5%) was used as a negative control. Results are expressed as a percentage of the untreated control group (mean ± SD; n=3; Kruskal-Wallis test; * p<0.05; *** p<0.001).
(B) Representative bright field images of fibroblasts treated with SEP for 72 h at the indicated concentrations (scale bars = 400 µm).
<figure><img src="image_4.png" /><figcaption>Figure 2. Effects of SEP and AA on ECM component synthesis and maturation by fibroblasts.</figcaption></figure>

Western blot analysis of type I collagen (A) and elastin and FLAG tagged SEP (E) in fibroblasts treated with AA, SEP (500 µg/mL), or their combination. Quantification of western blots (B) immature collagen, (C) pro collagen cleaved forms, (D) Mature collagen, (F) Elastin and (G) SEP. Mean ± SD; n=3; ANOVA one-way test; * p<0.05; ** p<0.01; *** p<0.001. (H) Immunofluorescence images showing the distribution of type I collagen (a-d), elastin (e-h), and FLAG-tagged SEP (i-l) under different treatments (scale bars = 150 µm). Fluorescence quantification of (I) type I collagen, (J) elastin and (K) SEP. Data are displayed as violin plots showing the median and interquartile range; n=36; Welch's ANOVA, post-hoc Dunn's comparisons, not significant (ns); * p<0.05; ** p<0.01; *** p<0.001; **** p<0.0001. (L) Immunofluorescence images showing the localization of nuclei (a, e, i),
elastin (b, f, j), SEP (c, g, k), and the merged channels (d, h, l) for fibroblasts untreated (a-d), treated with SEP (e-h), or treated with SEP and βAPN (i-l). Scale bars = 40 µm.
A

Untreated

<figure><img src="image_5.png" /></figure>

a

50

<figure><img src="image_6.png" /></figure>

b

200

<figure><img src="image_7.png" /></figure>

c

<figure><img src="image_8.png" /></figure>

d

e

f

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

f

<figure><img src="image_10.png" /></figure>

B

Untreated

<figure><img src="image_11.png" /></figure>

a

<figure><img src="image_12.png" /></figure>

b

<figure><img src="image_13.png" /></figure>

c

C

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /><figcaption>Figure 3. Impact of SEP on the morphology and ultrastructure of 3D dermal constructs.</figcaption></figure>

(A) Macroscopic images of dermal substitutes cultured with SEP concentrations of 0, 50, and 200 µg/mL (a, b, c, respectively) (scale bars = 3 mm). Surface plots (d, e, f) corresponding to the macroscopic images (a, b, c) represent grayscale intensity values ranging from 0 to 255 based on
pixel distribution. Line profiles (g, h, i) display grayscale intensity values as a function of pixel distance for images (a, b, c), respectively. (B) Representative phase-contrast microscopy images of the extracellular matrix from reconstructed dermal substitutes treated with SEP at 0, 50, and 200 µg/mL (a,b,c, respectively). The dot-shaped structures that correspond to the SEP deposit are indicated by white arrows (scale bars = 100 µm). (C) Transmission electron microscopy (TEM) images of immunogold co-labelling of fibrillin-1 (5 nm beads) and SEP (15 nm beads) in the extracellular matrix after treatment with SEP at 50 µg/mL Scale bars: 200 nm (a), 100 nm (b), and 50 nm (c).
<figure><img src="image_17.png" /><figcaption>Figure 4. Effects of decellularization on cell viability and type I collagen organization.</figcaption></figure>

Live/dead staining of reconstructed dermal substitutes before (a-c) and after decellularization (g-i).
Green fluorescence (calcein-AM) indicates live cells, while red fluorescence (propidium iodide) marks dead cells. Immunofluorescence staining for type I collagen before (d-f) and after decellularization (j-l) under different SEP treatment conditions. Scale bars = 100 µm.
Supplementary Information:

<figure><img src="image_18.png" /><figcaption>**Figure S1** Immunofluorescence images showing the distribution of fibrillin 1 (a-d), fibronectin (e-h), and lysyl oxidase like 1 (i-l) under different treatments (scale bars = 100 µm).</figcaption></figure>

<figure><img src="image_19.png" /><figcaption>**Figure S2** Transmission electron microscopy (TEM) images of non-decellularized (a) and decellularized (b) MEC. Scale bars: 1µm.</figcaption></figure>