A method for producing leather using cell culture on a honeycomb macroporous polymer scaffold

Abstract

The present disclosure relates to a method for producing leather, comprising culturing cells on a honeycomb macroporous polymer scaffold. The present disclosure also relates to leather obtained by this method.

Description

Detailed Description of the Invention

[0001] [Technical field] The present disclosure relates to a method for producing leather comprising culturing cells on a honeycomb macroporous polymer scaffold. The present disclosure also relates to leather obtained by such a method.

[0002] BACKGROUND OF THEINVENTION Leather was ranked the number one most resource-intensive material used in the fashion industry, far ahead of cotton or polyester. While animal leather is prized for its durability and beauty, it faces multiple environmental issues, including intensive use of natural resources, animal welfare, and concerns over working conditions. Furthermore, for many applications, leather needs to be free of defaults such as scratches or moth damage. As traditional leather production relies primarily on animal husbandry, up to 80% of leather is discarded due to not meeting quality standards.

[0003] With brands under immense pressure to become more ethical and sustainable, there is a need for sustainable alternatives to animal leather that are based on cell biology.

[0004] Synthetic leathers have been developed to address some of these concerns, but lack the qualities of natural leather. Previous attempts to produce artificial leathers have been described. For example, EP1589098 describes a method for growing fibroblasts seeded on a three-dimensional bioactive scaffold. Such scaffolds can be made from collagen waste ("split") from the tanning process, particles of pure collagen, particles of collagen waste, or synthetic scaffolds (e.g. made of polymers such as HYAFF). WO2017 / 184967 describes the culture of stem cells or keratinocytes on collagen or PET membranes with a microporosity of 4 μm. However, these culture conditions are not optimal for cell proliferation and collagen production.

[0005] The tanning process degrades most of the cellular, molecular and extracellular matrix components. The only components that survive storage and tanning are the collagen and elastin fibers present in the final leather and secreted by the fibroblasts (Sharphouse, JH Leather Technician's Handbook. Leather Producer's Association. p. 104. ISBN 0-9502285-1-6). These collagen and elastin fibers are responsible for the mechanical properties of leather. For this reason, inducing the secretion of extracellular matrix (especially collagen and elastin) by the network of fibroblasts growing on the scaffold is a crucial step in leather development.

[0006] Thus, there remains a need to develop cell culture methods that promote cell proliferation as well as the production of collagen, which is essential for the production of high quality leather.

[0007] Summary of the Invention The present inventors have developed a new method for producing high-quality leather by culturing fibroblasts on honeycomb macroporous polymer scaffolds in vitro, which allows efficient high cell attachment, cell culture and proliferation over long periods of time with minimal stress and damage to the cells to provide a collagen-rich extracellular matrix that is tanned and transformed into a leather-like biofabricated material, resulting in increased collagen production by fibroblasts.

[0008] In particular, we have fabricated a scaffold with homogeneous yet randomly distributed honeycomb pores with uniform distances between the pores, generating a well-organized structure. Interestingly, this scaffold is capable of supporting cell viability with high extracellular matrix secretion compared to other macroporous sponge scaffolds.

[0009] In the membrane scaffolds of the present disclosure, the honeycomb structure of open pores at the surface and aligned finger-glove-like pores extending deep inside, in addition to the interconnected pores, direct multicellular organization and further enhance the deposition of fibrillar collagen. Such highly organized membrane scaffolds provide rigidity and a suitable microenvironment for cell colonization and development, and deposition of extracellular matrix (ECM) fibers.

[0010] This honeycomb macroporous scaffold, also referred to herein as a membrane scaffold, is particularly useful for obtaining a texture suitable for providing a leather-like product after the tanning process.

[0011] The present invention relates to a method for producing leather comprising the steps of: a) culturing in vitro fibroblasts on a honeycomb porous polymer scaffold to obtain a tissue, wherein the surface of said scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and the wall thickness of said pores is less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm, b) tanning said tissue, thereby forming said leather. In a preferred embodiment, said scaffold comprises polyester, preferably biodegradable polyester and / or bio-derived polyester, more preferably in a concentration of 1-50% (w / w), preferably 5-20% (w / w). In a particular embodiment, the macropores are interconnected with micropores, and preferably the micropores have a diameter of less than 20 μm, preferably less than 10 μm. According to the present disclosure, preferably the scaffold has a tensile strength of 0.1-10 MPa, 0.1-8 MPa, 0.1-5 MPa, 0.1-2 MPa, 0.1-1 MPa, preferably 0.1-0.8 MPa, more preferably 0.2-0.7 MPa, and / or a Young's modulus of 5-1 MPa, preferably 5-800 kPa, 5-700 kPa, 5-600 kPa, 5-500 kPa, 5-400 kPa, 5-300 kPa, 5-200 kPa, preferably 5-100 kPa, 10-80 kPa, 40-70 kPa, 20-60 kPa, more preferably 30-50 kPa. In a preferred embodiment, said scaffold is obtained by non-solvent induced phase separation (NIPS).

[0012] In one particular embodiment, the scaffold is an asymmetric scaffold comprising macropores which are homogeneously but not regularly distributed throughout the thickness of the scaffold, in particular presenting a first face comprising surface-opening macropores with a diameter of 100-280 μm, preferably 110-225 μm, a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm, and a second symmetrical face comprising only nanopores with a diameter of less than 10 nm, preferably less than 5 nm. In another particular embodiment, the scaffold is a symmetrical scaffold presenting two faces containing macropores with a diameter of 100-280 μm, preferably 110-225 μm, a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm.

[0013] In a preferred embodiment, bioactive molecules are grafted onto the surface of the scaffold, preferably said bioactive molecules are collagen, glucids or glucid-derived molecules such as, for example, glycosaminoglycans.

[0014] In another aspect, the present disclosure relates to tissues obtainable by the culture step a) of the method of the present invention, and to tanned leather obtainable by said method.

[0015] Finally, the disclosure relates to the use of a macroporous polymer scaffold for producing leather, preferably by tanning tissue obtained from fibroblasts cultured on said macroporous polymer scaffold, wherein the surface of said scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm.

[0016] [Drawing legend] Figure 1: Overview of the fabrication of polyester asymmetric membrane scaffolds by the NIPS process.

[0017] Figure 2: Overview of the fabrication of symmetric membrane scaffolds by physical and / or chemical methods to remove the asymmetric scaffold smooth surface.

[0018] Figure 3: Scanning electron microscopy images of the porous surface (a), the interior of the macropores (b), the cross-section (c), and the smooth surface (d) of a biodegradable polyester membrane scaffold. The porous surface (a) shows the presence of honeycomb-like macropores on top of the porous surface, the micropores connecting the macropores (b), the thinness of the smooth surface opposite the porous surface, and the absence of macropores and micropores on the smooth surface. Scale bars: 400 μm (a, c, and d), 200 μm (b).

[0019] Figure 4: Scanning electron micrographs of polyester sponge seeded with fibroblasts, fixed and dried. The polyester sponge scaffold is lined with cell bodies over its entire surface. Scale bars: 400 μm (a and b), 50 μm (c).

[0020] Figure 5: Scanning electron micrographs of fibroblast-seeded polyester membranes. Cells colonize the scaffold, depositing abundant extracellular matrix fibers. Scale bars: 400 μm (a and b) and 50 μm (c).

[0021] Figure 6: Proliferation of fibroblasts cultured on polyester sponge (circles, solid line) and membrane (squares, dashed line) scaffolds at various time points.

[0022] Figure 7: Tissue production by fibroblasts seeded on polyester membrane scaffolds 1 week (a) and 3 weeks (b) after seeding, as observed by fluorescence microscopy. Cell-derived collagen areas were observed by fluorescence microscopy using anti-collagen I antibody (Sigma, part number C2456). Cell nuclei were stained with DAPI (blue) and collagen I (green). Scale bar: 500 μm.

[0023] Figure 8: Tanning process with tanning / re-tanning process of tissue samples to obtain tanned leather. Steps a) tanning and re-tanning process are brought into the tanning drum, b) surface image of tanned leather under high-power microscope, scale bar: 2 mm, c) final result of obtained tanned leather.

[0024] Figure 9: Schematic representation (right) of open pore boundaries at the polyester sponge (top) and membrane (bottom) scaffold surfaces based on scanning electron microscopy images (left).

[0025] Figure 10: Distribution of surface porosity (%) of macropores at the surface of sponge (grey) and membrane (black) scaffolds. The median ± MAD% surface porosity of membrane and sponge scaffolds was 66.11 ± 8.53 and 21.49 ± 8.65, respectively. p-value = 0.0011655 by Wilcoxon rank sum test. ** p<0.01.

[0026] Figure 11: Pore wall thickness between open macropores in sponge (grey) and membrane (black) scaffolds, calculated by subtracting the distance between the centers of two pores by the sum of their radii. The median ± MAD wall thickness for membrane and sponge scaffolds was 29.37 ± 12.40 and 70.15 ± 26.60, respectively. p-value by Wilcoxon rank sum test = **** p<0.0001.

[0027] Figure 12: Representative results of traction mechanical tests performed on polyester scaffolds according to ISO 3376:2020 standard using an Instron 34SC-1 system with a 50N load cell (CAT.NO.: 2530-50N) and manual clamping grips (CAT.NO.: 2710-203). a) Measured stress / strain traction curve of polyester membrane scaffolds, b) Measurement of the elastic limit of the specimen by measuring Young's modulus, and c) Measurement of the tear load of the specimen. The speed was set and controlled at 100 mm / min during the test. Strength and displacement measurements were recorded. Stress was calculated by the initial thickness and width of the specimen. Elongation was calculated by the initial length and observed displacement.

[0028] Figure 13: Orientation of biomaterial fibers (cells and extracellular matrix) at the surface of in vitro skin. Orientation of biomaterial field lines at the surface of in vitro skin obtained by scanning electron microscopy (A and B): Scale bar: 100 μm. Orientation histogram (C) showing the frequency of biomaterial oriented regions at the surface of the in vitro skin sample.

[0029] Figure 14: Thermograms of DSC analysis of samples: macroporous scaffold (FSC), in vitro skin (FP) and aldehyde-tanned in vitro skin leather (CF-1).

[0030] FIG. 15: Resolution of the peaks and determination of the denaturation temperatures of the different species present in the sample: onset temperature of the initiation of the transition (T onset ), the denaturation temperature (Td) corresponding to the peak, and the offset temperature at the end of the transition (T offset ).

[0031] Detailed Description of the Invention The inventors have developed a method to improve cell growth and collagen secretion and produce high-quality leather using an in vitro culture process of fibroblasts on honeycomb macroporous polymer scaffolds. Such a method can impart the stiffness, mechanical strength, and abrasion resistance seen in leather. In particular, the inventors have fabricated a honeycomb porous polymer scaffold with a uniform pore distribution on its surface, resulting in a highly organized structure with uniform pore-to-pore distance.

[0032] Interestingly, this scaffold is capable of maintaining cell viability with high extracellular matrix secretion compared to sponge macroporous scaffolds. The secretion of large amounts of extracellular matrix by cells cultured on this scaffold allows for the production of high-quality leather.

[0033] The present disclosure relates to a method for producing leather comprising the steps of: a) culturing fibroblasts in vitro on a honeycomb porous polymer scaffold to obtain tissue, wherein the surface of the scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%; b) tanning said tissue, thereby forming said leather.

[0034] In this disclosure, the term "scaffold" refers to a three-dimensional support that provides physical and structural support to cells and allows tissue formation.

[0035] According to the method of the present disclosure, the surface of the scaffold contains macropores with a diameter of 100 to 280 μm, preferably 110 to 225 μm, into which cells can enter, providing a biological environment favorable for the proliferation and function of fibroblasts.

[0036] For the purposes of this disclosure, the term "pore diameter" refers to the average pore diameter measured by averaging the diameters of 5-30 pores measured using a scanning electron microscope (SEM) with ImageJ software (Rasband, WS, ImageJ, US National Institutes of Health, Bethesda, Maryland, USA, https: / / imagej.nih.gov / ij / , 1997-2018). In a preferred embodiment, the pore diameter is measured on one surface of the scaffold.

[0037] In order to maintain the mechanical structure of the scaffold while providing a suitable biological environment for cell growth, the surface of the honeycomb macroporous scaffold according to the present disclosure has a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%.

[0038] By analyzing representative images of the microstructure of the scaffold obtained by scanning electron microscopy (SEM), the surface pore area distribution on the surface of the scaffold can be determined. The surface pore area distribution means the ratio of the pore surface area to the surface area of ​​the whole scaffold. The surface porosity (surface pore area distribution) of the prepared membrane was calculated by the following formula:

[0039]

number

[0040] Here, A t is the total area of ​​the image, and A p is the areal porosity at distance z, and h is the height of the image. The surface pore area distribution corresponds to the percentage of pores determined by the electron microscope image at a selected magnification. The image can be analyzed using appropriate software that can measure the dimensions of the pore cross-sections visible on the scaffold surface and the surface pore area distribution. The surface area porosity can be measured as described in Example D.2 of the present application.

[0041] According to one particular embodiment, the macroporous scaffold preferably has a total porosity in the range of 65% to 98%, in particular 70% to 98%, 75% to 98%, more preferably 80% to 95%.

[0042] The "degree of porosity", "void fraction" or "total porosity" is different from the surface pore area distribution mentioned above. In fact, in the present invention, the terms "degree of porosity", "void fraction" or "total porosity" refer to the ratio of the volume of pores to the volume of material, measured according to gravimetric methods (Guarino V, et al. 2008 Sep;29(27):3662-70). The volume of the sample is calculated from the measurements of the sample dimensions. The mass of the sample can be measured to estimate the density of the sample. The density (ρSc) of each sample is the average value obtained from four samples. The porosity of the scaffold was then calculated using the following formula: Porosity=(1-(ρSc / ρPpoly)):100 where ρPpoly is the density of the polymer, based on the manufacturer's database.

[0043] The scaffolds according to the present disclosure are honeycomb macroporous scaffolds (also referred to herein as membrane scaffolds), which are highly organized structures with uniform distances between pores, i.e., the walls separating the pores have the same thickness.

[0044] In a preferred embodiment, on the surface of the honeycomb macroporous scaffold according to the present disclosure, the distance between pores, corresponding to the width of the pore walls, is less than 70 μm, preferably less than 60 μm, more preferably between 2 and 50 μm.

[0045] As used herein, the width of a pore wall refers to the distance between two opposing walls of a pore. The width of a pore wall can be measured by any method known in the art. In a particular embodiment, the width of a pore wall can be measured from an SEM image by calculating the distance between the pores, as described, for example, in Haeri, Morteza & Haeri, Mohammad. (2015); Journal of Open Research Software; 3.10.5334 / jors.bn and in Example D.2. of the present application. The distance between the pores corresponds to the average surface area between the pores, which can be calculated by subtracting the distance between the centers of two adjacent pores by the sum of the radii of each pore. In another embodiment, the width of the pore wall can also be calculated by modeling the shape of the surface of the scaffold, as described in Example D.2. of the present application. This method consists of modeling the shape of the surface and transposing the pore surface area to a circle with a mean radius determined by the average distance of the pixelated points on a line from the center of such area.

[0046] The honeycomb macroporous scaffold according to the present disclosure may be designed to exhibit a thickness compatible with the synthesis of leather, in particular said scaffold may have a thickness of 0.1 to 2.5 mm, in particular 0.3 to 2 mm.

[0047] The thickness of the samples may be measured according to the ISO 2589:2016 standard. The thickness is measured with a Checkline digital thickness instrument J-40-L in accordance with ISO 2589:2016. Thirty measurements were taken, distributed and averaged for each sample.

[0048] In a particular embodiment, the scaffold according to the present disclosure has a tensile strength of 0.1-10 MPa, 0.1-8 MPa, 0.1-5 MPa, 0.1-2 MPa, 0.1-1 MPa, preferably 0.1-0.8 MPa, more preferably 0.2-0.7 MPa.

[0049] In another specific embodiment, the scaffold according to the present disclosure has a Young's modulus of 5-1 MPa, preferably 5-800 kPa, 5-700 kPa, 5-600 kPa, 5-500 kPa, 5-400 kPa, 5-300 kPa, 5-200 kPa, preferably 5-100 kPa, 10-80 kPa, 40-70 kPa, 20-60 kPa, more preferably 30-50 kPa.

[0050] Physical and mechanical testing of the scaffolds, namely the measurement of tensile strength and Young's modulus, was carried out according to the ISO 3376:2020 standard. The environmental conditions are ambient temperature and humidity. An Instron 34SC-1 system is used to carry out the tests. The system is equipped with an Instron 50N load cell (CAT.NO.: 2530-50N) and an Instron manual clamping grip (CAT.NO.: 2710-203). A speed is imposed during the test and is set to 100mm / min. Strength and displacement measurements are recorded. The applied stress during the test is calculated by the following formula:

[0051]

number

[0052] where F is the traction force applied by the system to the surface of the tensile specimen; S TS is the minimum surface area of ​​the tensile specimen, and W and T TS are the width and thickness of the tensile specimen.

[0053] The elongation is calculated from the initial length, the displacement, and the displacement on the traction axis.

[0054] The Young's modulus is calculated using the above parameters as described in Example D.3. (FIG. 12) herein using the following formula:

[0055]

number

[0056] E Mem is the measured Young's modulus of the membrane scaffold, σ is the measured tensile stress, and ε is the axial strain in the linear elastic region of the membrane scaffold.

[0057] Tensile strength is the maximum stress that a membrane scaffold can withstand while being stretched or pulled before breaking, and can be measured as described in Example D.3. (Figure 12) of the present application.

[0058] According to one particular embodiment, said polymeric support does not have a fibrous structure.

[0059] Polymers according to the present disclosure that can be used in the scaffold are polyesters derived from poly(α-hydroxy acids).

[0060] Non-limiting examples of polyesters derived from poly(α-hydroxy acids), particularly bio-derived and / or biodegradable polyesters, that can be used to form the scaffold include, for example, polylactic acid (PLA), polyglycolic acid (PGA), copolymers of lactic acid and glycolic acid (PLGA), poly(3-hydroxybutyrate), poly(4-hydroxybutyrate), poly(3-hydroxyvalerate), poly(5-hydroxyvalerate), poly(3-hydroxypropionate), poly(3-hydroxyhexanoate), poly(3-hydroxyoct ... polycaprolactone; homopolymers and copolymers of poly(butylene succinate) and poly(butylene adipate), such as poly(butylene succinate-co-butylene adipate) (PBSA) and poly(butylene succinate) (PBS); and polyhydroxy esters of 3-, 4-, 5- and 6-hydroxyalkanoic acids, and mixtures thereof, that are polyhydroxyalkanoates (PHAs) and their derivatives.

[0061] In a preferred embodiment, the scaffold comprises 1-50% (w / w), preferably 5-20% (w / w) of polyester polymer.

[0062] Polyesters have interesting properties from a mechanical point of view and are compatible with cell culture. They can be bio-based since they can be produced from agricultural waste and / or are completely biodegradable. However, polyesters are hydrophobic and impermeable to oxygen, which does not allow the diffusion of molecules after reticulation.

[0063] Thus, to facilitate gas exchange, the oxygen-impermeable polymer scaffold may further comprise nano- and micropores with a diameter of less than 20 μm, particularly between 0.1 μm and 10 μm, more particularly between 2 μm and 8 μm.

[0064] The scaffold, which is composed of an oxygen impermeable polymer, such as a polyester polymer, and contains micropores, can be an asymmetric scaffold or a symmetric scaffold.

[0065] Honeycomb macroporous polymer scaffolds according to the present disclosure, also referred to herein as micro / macroporous membrane scaffolds, can be achieved by phase inversion (e.g., non-solvent induced phase separation (NIPS)), where the exchange of solvent and non-solvent controls the pore formation. The outflow of solvent and intrusion of non-solvent between the network chain polymers in the presence of macroporogen allows the formation of micro- and macro-pores in the polymer, forming dual porosity in the scaffolds according to the present disclosure.

[0066] Phase inversion is a method well known in the art and includes, for example, non-solvent induced phase separation (NIPS), thermally induced phase separation (TIPS), vapor induced phase separation (VIPS), and polymerization induced phase separation (PIPS).

[0067] TIPS is a method in which a polymer solution is formed at high temperature using a high boiling point solvent and then cooled to induce phase separation and solidification of the polymer. A microporous membrane is obtained after extraction of the diluent.

[0068] In the VIPS process, a cast film of polymer and solvent is exposed to an atmosphere of vapor of a non-solvent molecule (usually water). The vapor penetrates the film, causing precipitation of the polymer, which ultimately results in a symmetric porous membrane without a dense skin. Since the thermodynamic properties of the casting solutions in the NIPS and VIPS processes are similar, the VIPS process should provide membranes with similar morphology to those produced using the NIPS process.

[0069] PIPS is a phase separation that occurs in a multi-component mixture induced by the polymerization of one or more components. The molecular weight of the reactive components increases, causing one or more components to become immiscible with each other, resulting in spontaneous phase separation. The morphology of the final phase-separated structure is generally random, since the occurrence and course of phase separation are stochastic.

[0070] In particular, the scaffold can be fabricated by the following steps: preparing a solution of the polymer, comprising the polymer and at least one solvent for the polymer, as described above; adding a solid porogen to the prepared polymer solution under uniform stirring; pouring the solution comprising the polymer and porogen onto a solid support; introducing the polymer solution into a non-solvent solution (e.g., ethanol and water); and finally immersing the porogen / polymer complex into a porogen solvent to dissolve the porogen (Figure 1).

[0071] According to the above-mentioned method, the solid substrate onto which the polymer solution is deposited can be made of glass, metal, or plastic that is resistant to the solvent, such as, for example, polytetrafluoroethylene (Teflon®), nylon 6,6, or poly(ethylene terephthalate).

[0072] Examples of suitable porogens include inorganic salt crystals, such as sodium chloride crystals or potassium chloride crystals, as well as amorphous materials, such as poly(ethylene glycol) (PEG), polyvinylpyrrolidone, sucrose crystals, gelatin spheres, and paraffin spheres.

[0073] In particular, the concentration of the porogen in the polymer solution may in particular be from 10% to 98% by weight, more in particular from 30% to 95% by weight, relative to the weight of the polymer.

[0074] The solvent for the polymer can be easily selected by the person skilled in the art and can be selected from the following non-limiting examples: N,N-dimethylformamide, N-methylpyrrolidone, ethyl lactate, ethyl acetate, dimethylacetamide, dimethylsulfoxide, chloroform and mixtures thereof, and the non-solvent can in particular be water.

[0075] The formation of pores of different sizes is observed on the side in contact with the non-solvent and the side in contact with the substrate due to different rates of solidification. When the polymer solution is in direct contact with the non-solvent, a smooth nanoporous surface is directly formed, whereas when the non-solvent slowly diffuses between the substrate and the collodion, the solvent slowly bleeds out and polymer-rich and polymer-poor phases are formed.

[0076] After solidification, the polymer-rich phase forms the solid mass of the scaffold, while the polymer-poor phase forms an interconnected network of pores of different sizes that open at the surface and extend deep into the scaffold. The honeycomb macroporous polymer scaffold obtained by the NIPS process is an asymmetric scaffold with a network of interconnected macropores and micropores. The honeycomb macroporous scaffold presents a smooth surface and a porous surface, the porous surface containing macropores with a diameter of 100-280 μm, in particular 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably less than 2-50 μm, the pores extending deep from the porous surface to the smooth surface.

[0077] By asymmetric honeycomb macroporous polymer scaffold is meant a scaffold in which macropores with a diameter of 100-280 μm, in particular 110-225 μm, are not present throughout the thickness of the scaffold. In particular, said asymmetric scaffold presents a first face (porous face) containing macropores with a diameter of 100-280 μm, in particular 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95% and a pore wall thickness of preferably less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm, and a second symmetric face (smooth face) containing only nanopores with a diameter of less than 10 nm, more precisely less than 5 nm.

[0078] In another particular embodiment, said honeycomb macroporous polymer scaffold is a symmetric scaffold.

[0079] By symmetric scaffold is intended a scaffold having a diameter of 100-280 μm, in particular 110-225 μm, with macropores uniformly distributed throughout the thickness of the scaffold with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of preferably less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm; In particular, the scaffold has two macroporous faces, both of which contain macropores with a diameter in particular of 100-280 μm, in particular of 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm.

[0080] A symmetric honeycomb macroporous scaffold can be achieved by any method known to one skilled in the art. Non-limiting examples can be a) physical abrasion of the smooth surface and / or b) chemical dissolution of the smooth surface using a polyester polymer solvent (Figure 2).

[0081] In particular, polyester symmetric membrane scaffolds can be manufactured by the following steps: creation of a polymeric asymmetric scaffold as described above, physical abrasion of the smooth surface and / or chemical dissolution using a polyester polymer solvent, and finally immersion of the scaffold in water to remove the physical / chemical agents.

[0082] In order to functionalize the surface of the scaffold designed as above and to enhance the cytocompatibility of the scaffold, bioactive molecules can be grafted onto the surface of the scaffold.The bioactive molecules can be selected from polysaccharides such as cellulose, pectin, pullulan, keratan, hyaluronic acid, chondroitin sulfate, chitosan and heparin; proteins such as fibrinogen and collagen, especially soluble collagen; peptides known for their cell adhesion ability, for example, containing sequences such as arginine-glycine-aspartic acid (RGD) or arginine-glycine-aspartic acid-serine (RGDS); and mixtures thereof.These bioactive molecules can be grafted by any method known in the art.

[0083] According to a particular embodiment, the bioactive molecule is collagen. Indeed, collagen specifically interacts with integrin receptors expressed on the surface of skin-derived cells, allowing the adhesion of the cells. To graft collagen onto the surface of the scaffold, in a particular embodiment, the scaffold can be treated with plasma generated in air or oxygen, which adds electronegative silanol groups. Collagen is grafted onto the surface of the scaffold by immersing the scaffold in a solution containing collagen, followed by rinsing, preferably with phosphate-buffered saline (PBS).

[0084] According to a preferred embodiment, said bioactive molecules may be glucides and / or glucide-derived molecules, such as glycosaminoglycans (e.g. heparin sulfate, heparin, chondroitin sulfate, dermatan sulfate, keratan sulfate, hyaluronic acid), which play a fundamental role in the organization and homeostasis of the extracellular matrix due to their unique biophysicochemical properties (viscoelasticity, high water-holding capacity, ability to interact specifically or non-specifically with various proteins).

[0085] To graft glucides and / or glucide-derived molecules (e.g., glycosaminoglycans) onto the surface of the scaffold, the scaffold may first be pretreated to provide a positive charge, preferably positive amine groups, on the surface of the scaffold before being functionalized with negatively charged molecules. The positive amine groups may be added by treating the scaffold with plasma generated in oxygen or air and immersing the scaffold in a polylysine solution, by treating the scaffold with plasma generated in nitrogen, or by aminolysis reaction.

[0086] Once the surface of the scaffold has been treated to be positively charged, the scaffold can be functionalized with glucids and / or glucid-derived molecules by immersing the scaffold in a solution containing the glucids and / or glucid-derived molecules and then rinsing, for example, with ultrapure water.

[0087] According to a preferred embodiment, the positive amine groups are added by aminolysis of the polymer with a solution of at least one aliphatic α,ω-diamine. The aminolysis reaction can in particular be carried out by immersing the polymer scaffold in a solution containing an aliphatic α,ω-diamine, preferably in a 1,6-hexanediamine solution in propanol or water and mixtures thereof, followed by rinsing, for example, with ultrapure water.

[0088] According to the methods of the present disclosure, fibroblasts are seeded onto the polymer scaffold as described above and cultured under appropriate growth conditions, in particular in a medium that allows the induction of proliferation and secretion of extracellular matrix, such as different types of collagen and elastin fibers.

[0089] Fibroblasts according to the present disclosure are cells that synthesize extracellular matrix such as collagen, elastin, glycoproteins and non-proteoglycan polysaccharides, among other extracellular matrix components. Fibroblasts may be mammalian or non-mammalian fibroblasts. The cells may be a single cell type or a combination of multiple cell types. In a preferred embodiment, fibroblasts are the only cells cultured on the scaffold according to the method of the present disclosure, since collagen and elastin fibers secreted by fibroblasts are the only components that survive the storage and tanning process.

[0090] Many culture media are commercially available and well known to those skilled in the art. The culture medium may be a minimal medium, which contains, among other things, mineral salts, amino acids, vitamins, a carbon source essential for the cells, and a buffer system for adjusting the pH. Basal media that can be used in the method according to the invention include, but are not limited to, for example, MEM medium, DMEM / F12 medium, DMEM medium, RPMI medium, Ham's F12 medium, IMDM medium and KnockOut® DMEM medium (Life Technologies). Depending on the medium used, it may be necessary or desirable to add glutamine, ascorbic acid, growth factors, one or more antibiotics such as streptomycin, penicillin, and / or antimycotin.

[0091] In another embodiment, the present disclosure also relates to a tissue (also called in vitro skin) obtained by cell culture step a) as described above, preferably such tissue comprising fibroblasts cultured on a honeycomb macroporous polymer scaffold as described above comprising macropores with a diameter of 100-280 μm, preferably 110-225 μm, with a surface pore area distribution of 40%-95% and a pore wall thickness of less than 70 μm.

[0092] In one particular embodiment, the scaffold comprises a polyester, preferably a biodegradable polyester and / or a partially or fully bio-based polyester, preferably in a concentration of 1-50% (w / w), preferably 5-20% (w / w).

[0093] In a preferred embodiment, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer in which the macropores are interconnected with micropores, preferably with a diameter of the micropores less than 20 μm, more preferably less than 10 μm.

[0094] In a preferred embodiment, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer having a tensile strength of 0.1-10 MPa, 0.1-8 MPa, 0.1-5 MPa, 0.1-2 MPa, 0.1-1 MPa, preferably 0.1-0.8 MPa, more preferably 0.2-0.7 MPa.

[0095] In another preferred embodiment, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer having a Young's modulus of 5-1 MPa, preferably 5-800 kPa, 5-700 kPa, 5-600 kPa, 5-500 kPa, 5-400 kPa, 5-300 kPa, 5-200 kPa, preferably 5-100 kPa, 10-80 kPa, 40-70 kPa, 20-60 kPa, more preferably 30-50 kPa.

[0096] In another embodiment, the tissue comprises fibroblasts cultured on the asymmetric or symmetric scaffold described above.

[0097] In another preferred embodiment, said tissue comprises fibroblasts cultured on said scaffold obtained by said Non-solvent Induced Phase Separation (NIPS).

[0098] In a preferred embodiment, said tissue is obtained after at least 1 week, preferably 2 weeks, 3 weeks or 4 weeks of culture.

[0099] Culturing fibroblasts on scaffolds such as those described above can yield tissues containing a densely organized meshwork of cells and extracellular matrix, preferably with cell / extracellular matrix grooves of 200-400 μm as observed by light and / or scanning electron microscopy.

[0100] The tissue (in vitro skin) according to the present disclosure has at least one of the following characteristics: the tissue contains at least 50 μg / mg of collagen per mg of dry tissue, preferably at least 60, 70, 80, 90 or 100 μg / mg, more preferably 50-300 μg / mg, even more preferably 60-250 μg / mg, the amount of collagen being preferably measured by the hydroxyproline assay as defined in Example E.1.1 of the present application; the tissue comprises an elastin mass in a proportion of at least 0.3% (w / w), preferably at least 0.4% (w / w), of the total dry mass of the tissue, the mass of elastin being preferably measured by the elastin assay defined in Example E.2.1 (elastin content), in particular by measuring the elastin concentration in the supernatant of dried tissue digested with oxalic acid; - the tissue comprises a total fatty acid mass of less than 1% (w / w), preferably less than 0.1 or 0.01% (w / w), of the total dry mass of the tissue, preferably as determined by gas chromatography-mass spectrometry as defined in Example E.6.1 of the present application; - the tissue comprises glycosaminoglycans (GAGs) corresponding to the sum of sulfated GAGs and hyaluronic acid (HA) of less than 400 ng / mg, preferably less than 300 ng / mg, more preferably less than 200 ng / mg per total wet mass of the tissue, preferably less than 200 ng / mg HA per total wet mass of the tissue, preferably less than 150, less than 100 ng / mg HA per total wet mass of the tissue and / or less than 300 ng / mg sulfated GAGs per total wet mass of the tissue, preferably less than 200 ng ECS / mg sulfated GAGs per total wet mass of the tissue. In a preferred embodiment, sulfated GAGs are measured by staining with dimethylene blue as described in Example E.7.1 and / or hyaluronic acid is measured by addition of ethanol acetate saturated with cetylpyridinium as described in Example E.7.1. - the tissue is a material that is isotropic at the surface, preferably analyzed by 2D Fast Fourier Transform (FFT) observed by SEM, as described in Example E.3, Directional Analysis of Biomaterial Fibers in In Vitro Skin, In particular, the tissue shows the absence of representative peaks with a consistent variation of a few percent, as shown in Figure 13C, said tissue contains less than 0.3 ppm, preferably less than 0.2 ppm, of total extractable heavy metals, preferably as measured by the heavy metal content of Example E.4.1.

[0101] the tissue comprises less than 2 mg / g, preferably less than 1 mg / g, of free amines per mass of dry tissue, preferably as determined by the free amino group determination of Example E.5.1, - the tissue has at least two denaturation temperatures (Td), preferably three Td, determined by a Differential Scanning Calorimetry (DSC) assay, preferably measured by a wet Differential Scanning Calorimetry (DSC) assay as defined in Example F1.1, more preferably measured in the range of 5-100°C at a constant rate of 5°C / min.

[0102] The tissue obtained by the cell culture process described above is then tanned by any method known in the art to create chemical bonds between the elastin and collagen fibers and transform it into leather.

[0103] The types of tanning processes used in tanning leather include chrome tanning using chromium(III) sulfate tanning agent, tanning using aluminum salts, aldehydes, and organic compounds, vegetable tanning using tannins, and tanning using low molecular weight phenolic or acrylic polymers (hereinafter referred to as "polymer tanning").

[0104] In another aspect, the present disclosure relates to tanned leather obtained by the method as described above. As used herein, the term "leather" or "synthetic leather" refers to a material obtained by tanning or chemical treatment of animal hide or tissue containing collagen, elastin and other components of the extracellular matrix. According to the present disclosure, said tissue is obtained by an in vitro culture process of fibroblasts and contains a dense mesh of cells and extracellular matrix components.

[0105] The tanned leather obtained by the method as described above (in vitro skin leather) has at least one of the following properties: said leather has at least two denaturation temperatures (Td), preferably three Td, determined by a Differential Scanning Calorimetry (DSC) assay, said denaturation temperatures being preferably measured by a wet Differential Scanning Calorimetry (DSC) assay as defined in Example F.1.1, more preferably in the range of 5-100°C at a constant rate of 5°C / min; said leather has a total porosity, preferably between 30 and 60%, measured by mercury intrusion porosimetry as described in Example F.2.1 Mercury Intrusion Porosimetry; The leather preferably has a water vapor permeability of between 0.5 and 5.5 mg / cm, measured at 23°C in the presence of 50% relative humidity, as described in Example F.3.1 Water vapor permeability 2 It has a water vapor permeability of .h.

[0106] The disclosure also relates to the use of a macroporous scaffold as described above for producing leather, preferably by tanning tissue obtained from the culture of fibroblasts on said honeycomb macroporous polymer scaffold, more preferably the surface of said scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, with a surface pore area distribution of 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%. In a preferred embodiment, the surface of said scaffold has a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm. In one particular embodiment, said scaffold is a polyester scaffold, preferably a honeycomb polyester macroporous scaffold. In another particular embodiment, said scaffold is an asymmetric or symmetric scaffold. Embodiments of the present invention are illustrated in the following specific examples, which are intended to be illustrative and not limiting.

[0107] [Example] A. Fabrication of asymmetric honeycomb polyester porous scaffolds using NIPS 1- Preparation of macroporous honeycomb polyester scaffolds (membrane scaffolds) The asymmetric polyester membrane is prepared according to the NIPS method. ポリマー / v 溶媒 ) are obtained by dissolving the selected polyester in the selected solvent with or without the selected porogen in different mass ratios at 70 °C for 2 h until complete dissolution. After removing air bubbles, the solution was cast on a glass plate and spread to form a film. The thickness of the cast film was limited by seven paper wedges on two lengths of the glass plate. The glass plate with the polymer film was immediately immersed in a Milli-Q water bath at room temperature (i.e., 20–22 °C). After solidification, the formed film was washed thoroughly with Milli-Q water and stored at room temperature in a desiccator under vacuum (Figure 1).

[0108] 2- Morphological characterization of polyester membrane scaffolds Polyester membranes were prepared by the NIPS process using polymer solutions with various polymer contents. Most of the membranes had two distinct faces that appeared different to the naked eye, indicating the presence of an asymmetric structure. Optical and SEM observations allowed a more detailed investigation of the morphology of the two faces (Figure 3).

[0109] These observations confirm the asymmetric structure of the membrane with a smooth upper surface (Fig. 3d) and a macroporous lower surface (Fig. 3a, b), with pores extending deep from the porous surface to several nanometers into the smooth surface (Fig. 3c).

[0110] As expected, in the NIPS method, the solidification of the polyester polymer occurred when a film of the polymer solution was immersed in a bath of water, which is a non-solvent for the selected polyester polymer. During this process, the polymer solvent, which is miscible with the non-solvent, was gradually removed, which led to the gelation, then vitrification and crystallization of the polyester polymer. More precisely, the top surface of the film was in direct contact with water, while the bottom surface was in contact with the substrate. Therefore, the solidification phenomenon occurred faster on the top surface than on the bottom surface. On the top surface, a polymer-concentrated gel was formed due to the rapid outflow of the solvent, and a smooth surface was formed after the solidification of the film. As the diffusion of the non-solvent proceeded through the polymer gel, phase separation occurred from a continuous polymer-rich phase to a dispersed polymer-poor phase, which became increasingly pronounced from the top to the bottom of the polymer film. The phase separation proceeded over time, and eventually the polyester-rich phase solidified, and the pores filled with the non-solvent were formed by the fusion of droplets of the polymer-poor phase. In this way, the NIPS method allows the production of asymmetric membranes that contain glove finger shaped macropores that are open on one side and extend through the depth of the membrane, and on the other side form a smooth surface without macro / micropores (Figure 3c) (see Al Tawil et al., European Polymer Journal. 105, 370-88, 2018).

[0111] 3. Effect of adding macroporogen We were interested in asymmetric membranes with surface-opening macropores with sufficiently large diameters (~140 μm) and suitable morphology to accommodate fibroblasts and their ECM. We therefore added porogens to better control the pore size and interconnections between the macropores. Such membranes showed surface-opening macropores with an increase in the number and diameter of macro- and micropores upon addition of selected macroporogens.

[0112] Interestingly, observation of the films by SEM (Figure 3) revealed that the addition of macroporogen led to the formation of a dual pore structure with a network of interconnected macropores and micropores, in addition to the classical increase in porosity that is attributed to the extraction of porogen during the solidification step of the film. Under these conditions, a highly organized honeycomb-like macropore structure (Figure 3a) was observed, and in addition, a small number of micropores (Figure 3b) were observed in the walls of the macropores.

[0113] 4. Surface modification of polyester membrane The polyester membrane was functionalized with glucid-derived molecules after aminolysis reaction. The development of red Ponceau color after aminolysis and brilliant green color after immobilization of glucid-derived molecules indicates the successful surface modification of the polyester membrane surface. All the coloring was uniform on all sample surfaces. Moreover, untreated scaffolds did not retain the coloring, which means that the surface modification is a key step for immobilizing HA and retaining cells.

[0114] B. Cell behavior and tissue production in asymmetric honeycomb polyester porous scaffolds.

[0115] 1. Cell behavior on polyester sponges and membranes Cell behavior is stimulated by biochemical (soluble factors), physical cell-cell or cell-ECM interactions, and mechanical stimulation of the substrate (scaffold). As a result, cells adhere, proliferate, differentiate, become dormant, or transform. Here, we highlight the influence of scaffold strength in combination with morphostructural parameters on cell behavior.

[0116] Fibroblasts were seeded on PDMS and polyester sponges obtained by solvent casting and particle leaching (SCPL) with calibrated sodium chloride (Ref. S5886-5KG, Sigma) or organic macropores, and on polyester membranes obtained by NIPS with selected macropores. These scaffolds were functionalized with glucid-derived molecules, and fibroblast seeding and culture were maintained for up to 5 weeks.

[0117] From the first hour after seeding, cells attached and started to proliferate on both the PDMS and polyester sponge surfaces. SEM images of the polyester sponge (Figure 4) showed that the entire sponge surface was lined with a cell layer. Cell bodies (rounded shaped cells) were detectable all over the sponge surface, even at different culture periods (3 and 5 weeks). The observed cell density was surprising compared to the expected results. The cells were biologically active (viable and proliferative), so there was less ECM deposition (Dekker et al.,Tissue Engineering of Cartilage and Bone: Novartis Foundation Symposium 24.Volume 249.2003). This result suggested that there was likely a crosstalk between the morphological and mechanical properties of the scaffold and the observed cell behavior.

[0118] Fibroblasts behaved differently on the polyester membrane obtained by the NIPS method. Similar to the sponge condition, the cells attached and proliferated from 1 h after seeding, colonizing the entire membrane surface. Very interestingly, the SEM images (Figure 5) showed fewer cell bodies compared to the polyester sponge and a prominent spread of extracellular fibers in the macro- and micropores.

[0119] From a morphological point of view, the PDMS and polyester sponge scaffolds showed a heterogeneous structure with low surface porosity and high wall thickness between the macropores. In the polyester membrane, the homogeneous but irregular distribution of honeycomb pores results in a highly organized structure with high surface porosity and thin wall thickness between the macropores (Figure 9). The irregularly thick mass of material (polyester) present in the voids of the sponge changes the mechanical properties, while the thin pore walls of the membrane give rise to a less stiff material with different degrees of elasticity.

[0120] The strength of the scaffold affects the cytoskeleton organization and therefore the morphology and therefore the behavior of the cells. Various studies have shown that stiffer substrates generally result in more rounded cell shapes and tightly distributed, whereas softer substrates promote cell spreading (Humphrey JD, Dufresne ER, Schwartz MA. Nat Rev Mol Cell Biol. 2014 Dec; 15(12): 802-12). It is not surprising that these changes in cell morphology are accompanied by changes in cell behavior, including viability, proliferation and / or ECM deposition. The inventors suggest that the presence of thick walls between the sponge pores makes the material too stiff to induce colony formation, and furthermore, causes colonies to form in very narrow paths and close to each other.

[0121] To quantitatively assess cell proliferation on either polyester sponges or membranes, we tracked cell proliferation using the Cell Proliferation Reagent WST-1 assay (Sigma, part number C5015944001), which is used for non-radioactive, spectrophotometric quantification of cell proliferation and viability of cell populations. As shown in Figure 6, fibroblasts on polyester membranes (Figure 6, dashed rectangles) have a growth curve with a gentle slope indicating an early stage of cell adaptation (<10 days), followed by a stage of cell proliferation (10-15 days), and a final stage of steady cell proliferation (>15 days), which we speculate correlates with the increase in ECM secretion observed in Figure 5. On the other hand, the growth curve of fibroblasts on sponges (Figure 6, circles and solid lines) shows an upward trend, indicating continued cell proliferation even after 4 weeks of culture. The sustained proliferation rate after 3 weeks of culture (Figure 6, circles and solid line) reflects a higher cell proliferation rate and can be associated with the less secretion and deposition of ECM observed on the surface of cultured and fixed polyester sponges and observed by scanning electron microscopy (Figure 4). On the other hand, the stabilization of cell proliferation after 2 weeks of culturing fibroblasts on polyester honeycomb membrane scaffolds is consistent with the less amount of cell bodies with high secretion of ECM observed by scanning electron microscopy (Figure 5). This antagonistic relationship between cell proliferation and ECM secretion is well explained by the antagonistic relationship of cell proliferation versus cell specialization (Cooper GM. The Cell: A Molecular Approach. 2nd edition. Sunderland (MA):Sinauer Associates; 2000).

[0122] 2. Tissue formation in asymmetric honeycomb polyester porous scaffolds Micropatterned surfaces, the alignment of ECM fibers, cells, and interconnected 3D pore structures are important microstructural features. It has been proven that micropatterned structures induce multicellular tissues and fibrous collagen deposition, and also affect the alignment and shape of individual cells (Gilchrist, CL, Ruch, DS, Little, D. & Guilak, F. Biomaterials 35, 10015-10024 (2014)). Furthermore, aligned cues were sufficient to induce cell shape, alignment, adhesion, and deposition of fibrous collagen matrix. In this case, the structure of the polyester sponge is defined as a micropatterned structure (pores follow the geometry of the porous body), while the pores of the polyester membrane are an ordered honeycomb structure at the surface and an ordered globe structure deep inside. Orderly aligned cells are more likely to differentiate, meaning that the cells are active and secrete more ECM (Du, Y., et al. Biomaterials 218, 119334 (2019)).

[0123] The relationship between structures (macro- and micro-scale) regulates cell behavior and encourages cells to secrete ECM when growing in the polyester membrane, as shown in Figure 5. The deposition of collagen fibers in the pores and their attachment to the pore walls definitely improves the mechanical properties of the construct.

[0124] To characterize cell proliferation, cell distribution, ECM expression, and ECM fiber distribution in the cultured asymmetric honeycomb polyester membrane scaffold constructs, fibroblasts were seeded onto the polyester membrane and cultured for 1 and 3 weeks, after which they were fixed with 3.7% PFA in PBS 1X and stained with anti-collagen 1 specific antibody (Sigma, product code C2456) in combination with a cell nucleus staining reagent (DAPI) as shown in Figure 7.

[0125] As shown in Figure 7a, membrane constructs cultured for 1 week show that cell bodies proliferate within the macropore structure of the membrane, but the amount of secreted collagen-1 is low and collagen-1 fibers are absent. This result is consistent with the observation that cells proliferate during the first 2 weeks of culture on membrane scaffolds, as shown in Figure 6 (circles, full lines).

[0126] In contrast, membrane constructs cultured for 3 weeks displayed tissue containing a dense organized mesh of cells and collagen-1 (Figure 7b), in particular 200-400 μm cell / collagen-1 grooves reminiscent of the interpapillary processes protruding from the papillary dermis at the interface between the dermis and epidermis in animals (Q. Zeng, L. K. Macri, A. Prasad, R. A. Clark, D. I. Zeugolis, C. Hanley, Y. Garcia, A. Pandit, D. I. Leavesley, D. Stupar, M. L. Fernandez, C. Fan, Z. Upton, 6.20 Skin Tissue Engineering☆, Editor(s): Paul Ducheyne, Comprehensive Biomaterials II, Elsevier, 2017, Pages 334-382). This result is consistent with the observation that fibroblasts cultured on membrane scaffolds do not proliferate but enter a phase of ECM-secreting cell specialization, as shown in FIG. 6 (square shapes, dashed line).

[0127] C. Tanning and Fabrication of Cultured Leather on Asymmetric Honeycomb Polyester Porous Scaffolds.

[0128] The constructs obtained after culturing fibroblasts on asymmetric honeycomb polyester macroporous scaffolds as described above were tanned and re-tanned using methods known in the art to generate chemical bonds between the elastin and collagen fibers and transform them into leather. The cultured polyester sponges and membranes were tanned and re-tanned in a laboratory tanning drum following the above-mentioned protocol used for traditional animal leather (Figure 8a).

[0129] Unfortunately, polyester sponges cultured with fibroblasts for 4–5 weeks became brittle and fragmented during the tanning and retanning process. On the other hand, polyester membrane structures seeded for 4 or 5 weeks maintained their structural integrity after the tanning and retanning treatments, resulting in leather-like craft products, as shown in Figure 8c. Indeed, as shown in Figure 8a, an asymmetric honeycomb polyester macroporous scaffold cultured for 4 weeks, tanned and retanned, exhibited the leather grain motive characteristic of full-grain leather when observed under a low-magnification optical microscope (Figure 8b).

[0130] D. Geometric and Mechanical Comparison of Polyester Sponges and Membranes 1.Geometric From the morphology point of view, the polyester sponge showed heterogeneously distributed porosity (Figure 9, top), low surface porosity, and high wall thickness between macropores, whereas the polyester membrane showed a highly organized structure with homogeneous but irregular distribution of honeycomb-structured macropores, high surface porosity, and thin wall thickness between macropores (Figure 9, bottom).

[0131] 2. Surface Morphological Analysis of Porous Scaffolds: Sponges and Membranes Scanning electron microscope (SEM) images at specific magnifications were used to measure the surface porosity, pore size and surface pore area distribution of the membranes. The porous surface of the scaffolds was imaged by SEM, and the corrected magnification was used to standardize the image segmentation using ImageJ software (Rasband, WS, ImageJ, US National Institutes of Health, Bethesda, Maryland, USA, https: / / imagej.nih.gov / ij / , 1997-2018). The raw images (gray / black images) from the outer surface of the porous scaffolds were binarized between 0 and 255 pixels, with low brightness pixels assumed as pore regions and high brightness pixels assumed as material-containing boundaries, respectively. An appropriate function was used to remove high-frequency and low-frequency noise caused by the binarization of the pores from the plan view of the background (Figure 9).

[0132] For automated measurements of surface pore area, circularity, and Feret pore size, each macropore on the surface area of ​​the hollow sponges or membranes was digitized in the binarized images using the "Analyze Particles" function in ImageJ. From these measurements, surface porosity, surface pore area distribution, and Feret pore size were obtained for a statistically representative number of sponge and membrane scaffolds.

[0133] The surface porosity of the prepared membrane was calculated by the following formula:

[0134]

number

[0135] where At is the total area of ​​the image, Ap is the areal porosity at distance z, and h is the height of the image.

[0136] Surface pore area distribution (pore number / area) is defined by the distribution of the number of labeled pores on a selected surface. Feret pore size is the longest distance between any two points along the boundary of a pore, also known as the maximum caliper or maximum diameter.

[0137] Additional analysis of the sponge and membrane scaffold structure was performed using the ND plugin in ImageJ as described in Haeri, Morteza & Haeri, Mohammad. (2015). Journal of Open Research Software. 3. 10.5334 / jors.bn. ND was used to calculate the distance between pores and to determine the thickness of the walls between the pores. The distance between the pores, i.e. the average surface area between the pores, was determined by dividing the distance between the centers of three adjacent pores by the sum of the pore radii. The pore centers were determined digitally by determining the center of the geometric pore boundary corresponding to the pore. The calculation of the wall distance between the pores was determined from the distance between the centers of the pores corrected for the area of ​​the pores (Haeri, Morteza & Haeri, Mohammad. (2015). Journal of Open Research Software. 3.10.5334 / jors.bn).

[0138] As shown in Figure 10, the surface porosity of the membrane scaffolds is 21.49 ± 8.65 compared to 66.11 ± 8.53 for the sponge scaffolds, which is 3 times higher than that observed for the sponge scaffolds. Furthermore, as shown in Figure 11, the median ± MAD mean pore wall thickness of the membrane scaffolds is 29.37 ± 12.40 compared to 70.15 ± 26.60 for the sponge scaffolds, which is 2.4 times lower than that observed for the sponge scaffolds.

[0139] The low interconnected porosity, plus the presence of material masses with irregular thickness between the sponge pores, alters the mechanical properties and the intercellular communication, respectively. A high interconnected porosity, like that of a membrane, allows intercellular communication and coordinates the deposition of ECM fibers. Furthermore, the thin pore walls of the membrane result in a less stiff material with different mechanical properties. The stiffness of the scaffold affects the cytoskeleton organization and therefore also the cell morphology and thus the behavior. Various studies (Yeung T,et al:CellMotil Cytoskeleton 2005;60(1):24-34;Gaudet C,et al.Biophys J 2003;85(5):3329-35) have shown that stiffer scaffolds with higher porosity generally induce a more rounded cell shape with close distribution, while softer substrates promote cell spreading. It is not surprising that such changes in cell morphology are accompanied by changes in cell behavior, including ECM deposition. The presence of rigid walls between the sponge pores may make the sponge a more rigid material and may also induce cell aggregation and colonization of the scaffold in very narrow pathways and close to each other. Under these conditions, cells form junctions that allow survival, while secretory activity is minimized and metabolism is low while maintaining a high proliferation rate. Thus, the behavior of cells on polyester sponges and membranes is closely correlated with the morpho-mechanical parameters of the scaffold (porosity, pores, pore wall thickness). This correlation directly controls the properties of the tanned construct.

[0140] (3. Mechanical Analysis of Asymmetric Honeycomb Polyester Porous Scaffolds) Several polyester membrane scaffold samples were tested for their mechanical properties. Various parameters such as thickness, and different polymer and porogen blends and combinations were tested. Each combination was tested in the wet state.

[0141] To prepare standard tensile specimens, each specimen was cut with a press knife as specified in ISO 2419-2012. The environmental conditions were room temperature and humidity. The samples were shaped like dog bones. The specimens were 10 mm wide and 50 mm long. The thickness was dependent on the specimen preparation.

[0142] The thickness of each sample was measured according to the ISO 2589:2016 standard. Thickness is measured with a Checkline J-40-L digital thickness gauge in accordance with ISO 2589:2016. Measurements were taken in triplicate, dispersed and averaged for each sample.

[0143] Physical and mechanical testing of the scaffolds, namely the measurement of tensile strength and Young's modulus, was carried out according to the ISO 3376:2020 standard. The environmental conditions are ambient temperature and humidity. An Instron 34SC-1 system is used to carry out the tests. The system is equipped with an Instron 50N load cell (CAT.NO.: 2530-50N) and an Instron manual clamping grip (CAT.NO.: 2710-203). A speed is imposed during the test and is set to 100mm / min. Strength and displacement measurements are recorded. The stress applied during the test is calculated by the following formula:

[0144]

number

[0145] where F is the traction force applied by the system to the surface of the tensile specimen; S TS is the minimum surface area of ​​the tensile specimen, and W and T TS are the width and thickness of the tensile specimen.

[0146] The elongation is calculated from the initial length, the displacement, and the displacement on the traction axis.

[0147] Representative results of traction mechanical tests performed on polyester scaffolds according to the ISO 3376:2020 standard are shown in Figure 12. Raw stress / strain curves were obtained and displayed in Figure 12a. The elastic limit was defined as the lowest stress point at which permanent plastic deformation was measured and distinguishable from elastic deformation, as shown in Figure 12b.

[0148] The Young's modulus of the membrane scaffold was calculated using the following equation:

[0149]

number

[0150] E Mem is the measured Young's modulus of the membrane scaffold, σ is the measured tensile stress, and ε is the axial strain in the linear elastic region of the membrane scaffold.

[0151] Tensile strength is the maximum stress that a membrane scaffold can withstand while being stretched or pulled before breaking, as shown in FIG. 12c.

[0152] The data obtained from these measurements are: Young's modulus, elastic limit and tensile strength. These values ​​are used to evaluate the mechanical properties of the asymmetric honeycomb polyester porous scaffold.

[0153] Young's modulus of the membrane scaffold was measured as 40.80 ± 14.21 kPa, mean ± standard deviation, and the tensile strength of the membrane scaffold was measured as 0.4244 ± 0.1603 MPa, mean ± standard deviation.

[0154] E. Characteristics of tissue (in-vitro skin) obtained by culturing fibroblasts on a membrane scaffold 1. Collagen content 1.1 Hydroxyproline assay protocol: In-vitro skin obtained by culturing fibroblasts on a membrane porous scaffold for 4 weeks and rabbit skin (shaved rabbit skin (dermis / epidermis)) were cut with a cutting tool with a diameter of 11 mm and dried using an oven (time: 16 h; T ° C: 50 ° C). The samples were weighed on a precision balance. The samples were mechanically ground and 100 μL of hydrolysate was collected for the determination and subjected to the measurement of hydroxyproline present in the samples.

[0155] Hydroxyproline concentrations in samples were measured according to the protocol provided by the manufacturer (Reference: MAK-357, Sigma Aldrich). A standard curve was obtained and linear regression of the standard curve yielded the equation of the curve and the regression coefficient R2.

[0156] The concentration of hydrolyzed hydroxyproline (μg / μL) was calculated using the following formula: Hydrolyzed hydroxyproline = (B / V) x D B: amount of hydrolyzed hydroxyproline (μg) calculated from the standard curve, V: amount of hydrolyzed sample added to the well (μL), D: dilution factor of the hydrolyzed sample.

[0157] The concentration of the hydrolysate can be converted to collagen equivalents by multiplying by the factor 7.49 (RB Neuman and MA Logan. J. Biol. Chem. 184, 299, (1950)).

[0158] 1.2 Results To evaluate collagen secretion in in-vitro skin, samples were seeded and cultured for 4 weeks in the presence of collagen activator. Hydroxyproline quantification was performed weekly during the 4-week culture according to the protocol described above. The collagen content of in-vitro skin after 4 weeks of culture was measured by hydroxyproline assay to be 106.53 μg / mg, compared to 69.03 μg / mg for rabbit skin.

[0159] 1.3 Conclusion The analysis of hydroxyproline in biological tissues is the standard method for measuring the content of collagen extracted from different tissues (GFCaetanoa, M. Fronzab et al.Pharmaceutical biology,vol.54,NO.11,2555-2559,(2016)). However, this method poses difficulties related to the extraction of collagen from the analyzed tissue. Thus, the measurement of collagen concentration by hydroxyproline assay in dry animal skin varies between 5 and 30% of the total dry matter (GFCaetanoa, M.Fronzab et al.11,2555-2559,(2016)). Caetano et al. used a hydroxyproline assay to measure collagen in rat skin (GF Caetanoa, M. Fronzab et al. Pharmaceutical biology, vol. 54, NO. 11, 2555-2559, (2016)). This hydroxyproline assay was normalized to the mass of dry skin before extraction and assay. The study showed a ratio of approximately 6 μg / mg hydroxyproline per mg dry skin. This corresponds to a mass ratio of 45 μg / mg collagen per mg dry skin, or 4.5% in the skin analyzed.

[0160] Parsons et al. also measured collagen in mouse skin using a hydroxyproline assay (Parsons KK, Maeda N, Yamauchi M, Banes AJ, Koller BH. Am J Physiol Endocrinol Metab. 2006 Jun;290). The hydroxyproline assay was normalized to the mass of dry skin before extraction and assay. The study showed a ratio of approximately 28 μg / mg hydroxyproline per mg dry skin. This corresponds to a mass ratio of 210 μg / mg collagen per mg dry skin, or 21% of the skin analyzed.

[0161] Taking these results into account, we have prepared in-vitro skin whose collagen composition is consistent with the results of extraction and hydroxyproline assays found in the literature.

[0162] Indeed, in our results, the amount of collagen was 106.53 μg per mg of in-vitro skin (dry), which corresponds to a mass ratio of 10.6% in the analyzed sample, which is within the range of results obtained in animal skin by this quantification process. A summary of the results obtained in the scientific literature and in this study is shown in Table 1.

[0163] [Table 1]

[0164] 2. Elastin content The mechanical properties of skin are determined by its composition. Elastin, which represents only a few percent of the dry mass, is responsible for the mechanical response to small stresses. In particular, it determines the rate of contraction after deformation, which contributes to the sensory imprint left on the skin after mechanical stress.

[0165] The inventors extracted and measured the total elastin content of in-vitro skin, and by comparing these results with studies in mammalian skin, the inventors showed that the elastin content extracted from the in-vitro skin samples was in the range of mammalian skin.

[0166] 2.1 Elastin measurement protocol In-vitro skin obtained by culturing fibroblasts for 4 weeks on membranous macroporous scaffolds with a circular (19 mm diameter) or square (17 mm side) shape was collected and washed with physiological saline (DPBS, Thermo, product number 10010), then dried in a drying oven and weighed on a precision balance. The dried material was cut with a scalpel and digested with oxalic acid at high temperature, and then the debris was removed by centrifugation. The supernatant was collected for analysis. 100 μL of the supernatant was collected and the elastin concentration in the supernatant was measured using the Fastin Elastin Assay Kit (Biocolor LTD., product number F2000). A standard curve (elastin provided in the kit) was obtained. Linear regression of the standard curve gave the equation of the curve and the regression coefficient R2.

[0167] 2.2 Results -Proofreading of protocols To verify the efficiency of elastin extraction using the protocol under the present extraction conditions, we performed sequential extraction tests on three 17 mm square in-vitro skin samples and measured the amount of elastin present in each extract (Table 2).

[0168] [Table 2]

[0169] Considering the standard deviation of the fifth and subsequent extractions (#extraction5, Table 2), we determined that four consecutive extractions are effective for extracting elastin from in-vitro skin using the protocol described above.

[0170] To assess the portion of the non-specific signal that may be related to the unseeded scaffolds, we performed a similar extraction and quantification of elastin on unseeded membrane scaffolds. We measured an absorbance equivalent to 1.42 μg of elastin. This allowed us to determine the threshold value corresponding to the non-specific signal or background reading. This value is very close to that obtained in the fifth extraction in the previous experiment. This confirmed that four extractions were efficient to extract the total amount of elastin present in the sample using the extraction method defined above.

[0171] -Elastin quantification To assess the total elastin of the two in-vitro skin samples, samples with a diameter of 19 mm were dried and then weighed. For these two samples, a dry mass of 50.1 mg and 51.4 mg was measured. Four successive elastin extractions were performed, resulting in a total volume of 2 mL containing the elastin extract. Non-specific signals were subtracted based on the previous measurements. In-vitro skin samples #1 and #2 contained 232.55 and 209.69 μg of elastin, respectively, corresponding to 0.45% and 0.42% of the dry mass (Table 3).

[0172] [Table 3]

[0173] Turner et al. measured elastin in porcine skin samples taken from different sites using a method equivalent to that described herein. The amount of elastin was normalized to the dry mass. The measurements reported in this study ranged from 2.45 to 11.30 μg / mg, i.e., 0.24% to 1.1% by mass (Turner NJ, Pezzone D, Badylak SF. Tissue Eng Part C Methods. 2015 Apr;21(4):373-84).

[0174] Considering the measurements carried out within the framework of this study, we have created an in-vitro skin whose elastin composition coincides with the results of extraction and quantification found in the literature. Indeed, using this experimental quantification method, we measured a mass fraction of elastin of 4.35 μg / mg for dry in-vitro skin. This corresponded to a mass ratio of 0.43%, whereas the literature recalls measurements located between 0.24% and 1.1%. A summary of the results obtained in the study by Turner et al. and those obtained in the present study is shown in Table 4.

[0175] [Table 4]

[0176] 3. Orientational analysis of biomaterial fibers in in-vitro skin Two-dimensional fast Fourier transform (FFT) analysis of the alignment of biomaterial networks observed by SEM of in vitro skin was performed using the Directionality plugin of ImageJ (ImageJ Directionality PlugIn. https: / / imagej.net / plugins / directionality) following the method previously described by Hamley et al. (Hamley, Introduction to Soft Matter: Polymers, Colloids, Amphiphiles and Liquid Crystals, John Wiley & Sons Ltd, Chichester, UK 2000). The Directionality plugin calculates spatial frequencies in the image based on a set of radial directions. This method produced a normalized histogram revealing the amount of fibers present between 0° and 180° with a bin size of 1° as detailed in Deravi et al. (Deravi, et al. (2017), Macromolecular Materials and Engineering. 302). Representative images of the surface of the analyzed in vitro skin specimens showing histograms of the frequency distribution of biomaterial fiber orientation vectors and fiber directionality are shown in Figure 13.

[0177] The results of directional histogram analysis of the biomaterial orientation at the surface of in vitro skin show a variation of a few percent and no representative peaks, consistent with an isotropic material (Figure 13C).

[0178] In contrast, animal skin is an anisotropic material, with mechanical and structural weaknesses in specific directions relative to the axis of symmetry of the animal from which the skin originates. The isotropic surface of in vitro skin leather is explained by its method of manufacture, which allows a high degree of control over the parameters that affect cell growth, in contrast to the growth of animal leather. This difference from conventional animal leather simplifies the use of the leather and does not require special consideration regarding the orientation of the cut edge of the leather surface.

[0179] 4.Heavy metal content The excessive presence of hazardous substances in clothing materials can pose a threat to consumer health, especially when the products are intended for children and / or come into direct contact with human skin. Heavy metals are one such hazardous substance, and therefore threshold values ​​for heavy metal content are set in standards applied in various jurisdictions. These standards aim to limit consumer exposure to heavy metals and ensure a product life cycle compatible with environmental concerns.

[0180] 4.1 Materials and Methods Metal determination was performed in in vitro skin (size: 26x43cm) by an external standards laboratory (contract research organization, CRO) Eurofins BLC (UK). Measurement of heavy metal content in in vitro skin was performed according to international standards: a) total heavy metal content in digested samples and b) content of heavy metals extractable by artificial acid sweat. Table 5 shows the permissible limits of total and extractable heavy metal content in commercial leather.

[0181] Sampling was carried out in accordance with BS EN ISO 2418-2017, which specifies the method of sampling (location of the leather piece), labelling and marking of samples to ensure identification. This applies to all types of mammalian leather, regardless of the tannery used.

[0182] Screening for metals was performed by atomic absorption spectrometry (AAS) with prior microwave digestion of the leather in accordance with BS EM ISO 17072-2:2019 and CPSC-CH-E1001-08.3. The detection limits for heavy metals tested by these methods are 0.1 ppm or mg / kg. Where appropriate, samples were conditioned and tested at 23°C and 50% relative humidity (standard atmosphere) in accordance with BS EN ISO 2419-2012.

[0183] Quantification of extractable metals was performed using atomic absorption spectrometry (AAS) in accordance with BS EN ISO 17072-1:2019. This method is capable of measuring extractable metals in leather and is not specific to the compound or oxidation state of the metal. This method is particularly suitable for the measurement of extractable chromium in chrome tannage comparisons. Where necessary, samples were conditioned and tested in accordance with BS EN ISO 2419-2012 at 23°C and 50% relative humidity (standard atmosphere). The detection limit for heavy metals tested using these methods is 0.1 ppm or mg / kg.

[0184] The concentration of a chemical per unit mass in a solid is usually measured in units of mass of chemical (milligrams, mg or micrograms, μg) per total mass (kilograms, kg), i.e. mg / kg or μg / kg. These concentrations are sometimes expressed in parts per million (ppm) or parts per billion (ppb): 1 ppm = 1 mg / kg, 1 ppb = 1 μg / kg. A measurement of 6 mg / kg is equivalent to 6 ppm or 6,000 ppb or 6,000 μg / kg.

[0185] [Table 5]

[0186] 4.2 Results The extractable and total heavy metal contents in the in vitro skin were quantified, and the results are shown in Table 6 (extractable content) and Table 7 (total content), respectively.

[0187] [Table 6]

[0188] [Table 7]

[0189] 4.3 Discussion The physical properties of leather can vary depending on the method of production and the structure of the chemicals used in its production. The pH value of leather can vary depending on the pH value of the chemicals used in its production, the additives used in the final stages of the leather manufacturing process and the proportions in which they are used.

[0190] The total amount of mineral matter in leather can be determined by the ash method and should not exceed 2.5%. In addition, depending on the nutritional status of the animal, up to 0.5% mineral matter may be present in natural skin. When the mineral content of in vitro skin was examined, the amount of heavy metals was negligible, with only copper and lead present above 0.1 ppm or mg / kg. The presence of copper may be related to the use of cell culture media, however, it was difficult to determine the origin of the lead traces. In our investigation, all values ​​were found to be below the limit values ​​for heavy metal content in commercial leather.

[0191] Today, consumer demands are moving towards materials that are free of hazardous substances and have a reduced environmental footprint. This necessitates the production of leather products that contain little or no heavy metals, especially in products that come into contact with human skin. In this study, the total heavy metal contents in the in vitro skin and the extractable heavy metal contents in the skin using artificial sweat were found to be below the limit values ​​for commercial leather (Table 5).

[0192] This makes in vitro skin a promising alternative for use in commercial leather production due to its low heavy metal content. In conclusion, it is expected that in the future the trend towards environmentally friendly products will have a greater impact on leather producers and the proposed limits will be lowered further. Therefore, the leather industry, especially the processing of natural leather, needs to prepare for these new limits and future transformations. In this context, an alternative such as that proposed in this study is a reliable approach that does not sacrifice the quality of the products manufactured using animal skins.

[0193] 5. Quantitative determination of free amino groups 5.1 Materials and Methods Proteins were extracted from in vitro skin by ultrasonication (Sonicator Q700 Qsonica) (power 50%, 14 s). Proteins were stained by adding 0.5 mL of 0.01% (w / v) TNBSA solution (Thermo Scientific, product number TS-28997) to 0.5 mL of each sample, and further incubated at 37°C for 2 hours. The reaction was stopped by adding 0.5 mL of 10% SDS (sodium bicarbonate buffer (0.1 M, pH 8.5)) solution and 0.25 mL of 1 N hydrochloric acid solution.

[0194] The absorbance of the solution was measured at 335 nm. The quantitative number of amines contained in the samples was determined by comparison with a calibration curve established with glycine solution (10 mg / mL, serially diluted to produce a standard range).

[0195] 5.2 Results Glycine was used to construct a standard curve by measuring the absorbance at 335 nm for given concentrations, which corresponds to the production of a colored compound after reaction with TNBSA (Table 8).

[0196] [Table 8]

[0197] [Table 9]

[0198] 5.3 Discussion The intensive use of dyeing and tanning processes in consumer goods such as leather, clothing and toys can, under certain conditions, lead to the formation of total / free amino groups. For this reason, the European Parliament has recently proposed an amendment 19 of Council Directive 76 / 769 / EEC on the restriction of certain substances. eme The limits set by this law, currently under review, are 6207 mg / L for total amines and 75 mg / L for free amines. The industrial method CEN ISO TS 17234, which is based on DIN 53316, is capable of detecting 20 of the substances listed in the decree.

[0199] The free amine contents of the membrane scaffold and in vitro skin were 12.9 mg / L and 25.9 mg / L, respectively, which are significantly below the permissible limit for free amines set by the Directive (75 mg / L). These results are very promising and in line with green technologies.

[0200] To compare the free amine content of in vitro skin and mammalian skin, a bibliographic analysis of the scientific literature on the subject was performed. Soomro et al. quantified the free amine content of human skin using a new method with pre-column derivatization (chromatography) using trifluoroacetylacetone and isobutyl chloroformate as the mobile phase (Soomro SA (2014), Journal of chromatography). This method was applied to the analysis of amines in human skin samples after hydrolysis. In human skin, total free amines were quantified using gas chromatography to be about 0.9 mg / g human skin.

[0201] The total free amine content of the in vitro skin was 25.6 mg / L (0.75 mg / gin vitro skin), which is comparable to the results of mammalian skin reported in the literature. Therefore, it is possible to use a variety of tanning agents commonly used in the traditional leather industry for the tanning of in vitro skin.

[0202] 6. Measurement of fatty acid content 6.1 Materials and Methods Fatty acid quantification was performed by an external standards laboratory (Contract Research Organisation) Eurofins BLC (UK). Samples were received and processed in accordance with ISO 2418:20179, which specifies the location of the laboratory samples on the leather pieces and how they should be labelled and marked for future identification. Samples were reconditioned and tested at 23°C ± 2°C and 50% relative humidity as specified in the standard reference atmosphere requirements of ISO 2419:2012 (for leather).

[0203] Quantification of in vitro skin fatty acids was performed by BLC by a) fat extraction by supercritical phase chromatography (SFC)4 followed by b) coupling with GC-MS analysis according to the protocol described in GB / T 9722-200611.

[0204] Internal standards were selected to measure the main components of fatty acids, volatile reactants and other impurities. Hydrogen / nitrogen was used as the carrier gas and diatomaceous earth (0.18mm-0.25mm) was used as the stationary phase. The flow rate was determined according to the detection limits of different fatty acids. The length of the column was 2m and the temperature was maintained at 80°C during the experiment. The process is detailed in ISO 12966-4:201512.

[0205] Sampling was performed according to ISO 2418-20179, which specifies the method of sampling (position on the leather pieces), labelling and marking of samples to ensure their identification. This standard is applicable to all types of mammalian leather, regardless of the type of tanning.

[0206] In vitro skin (26x43cm) or rabbit skin samples were washed with 1X DPBS saline (Fisher, 12037539), sealed with a vacuum device, and sent to a contract research laboratory (CRO).

[0207] Gas chromatography-mass spectrometry (GC-MS) is a sensitive method used for the identification and quantification of fatty acids. Before detection, the molecules of the carbon chain must be modified. Therefore, fatty acids are often modified by derivatization methods with acids (most often hydrochloric acid) to form fatty acid methyl esters (FAMEs). During the GC stage, the samples are separated according to the different FAMEs. These FAMEs undergo strong fragmentation by electron impact ionization in the mass spectrometer, resulting in the formation of many fragments. For quantification, the largest fragments are selected, showing high abundance and maximum specificity for individual fatty acid types.

[0208] 6.2 Results The total fatty acid content in the in vitro skin was quantified using the method described above, and the results are shown in Table 10.

[0209] [Table 10]

[0210] In this study, the measured in vitro skin fatty acid content was low, below the assay detection limit of <10 mg / kg.

[0211] 6.3 Conclusion According to various published results, the total fatty acid content of animal skin is about 4%. With the use of bath feed, the total fatty acid content increases to about 15% (Mihai AL; Metrology Promoting Harmonization&Standardization in Food&Nutrition; 3 Imeko Foods). The presence of fatty acids in leather makes it soft and supple. It also prevents the leather from wearing and breaking and increases its waterproofing. The total fatty acid content of in vitro skin was quantified. The results were found to be less than 10mg / kg, which corresponds to 0.001%. This is lower than the 4% fatty acid content of conventional animal leather.

[0212] There are also product lines that require soft and supple leather, such as gloves and other products that come into contact with the consumer's skin. The softness and suppleness of the leather can be adjusted during the bath feed process, where the individual fibers are evenly coated with bath feed that gives them special properties (Alaskan, A; Polymer Bulletin-Volume 79, doi.org / 10.1007 / s00289-021-03579-Z).

[0213] The low fat content of the in vitro skins allows them to be precisely tuned in the subsequent bath-feeding process to suit a wide range of product applications. In particular, the low fat content of the in vitro skins makes the degreasing step unnecessary and eliminates the risk of fat bleeding and condensation staining on the surface of the in vitro skins, a common drawback of conventional leather.

[0214] 7. Measurement of sulfated GAG and hyaluronic acid (HA) content Glycosaminoglycans (GAGs), including sulfated GAGs and hyaluronic acid, are responsible for the compressive strength of skin. However, the tanning process requires their removal. The water-retaining capacity of GAGs prevents the circulation of tanning agents within the collagen network. Therefore, to open this network, the GAGs must be removed.

[0215] 7.1. Hyaluronic Acid and Sulfated Glycosaminoglycan Assays Samples of in vitro skin (containing a collagen secretion inducer: 19 mm diameter disk) or rabbit skin (170.1 mg wet mass) were washed with physiological saline (DPBS, Thermo, product number 10010), weighed, cut with a scalpel, and digested with proteinase K at 55°C under stirring, then ground in a Potter mill and centrifuged to remove debris, and the supernatant was collected for analysis.

[0216] The sulfated glycosaminoglycan content was measured by simultaneously staining and precipitating sulfated GAGs with dimethylmethylene blue, dissociating the GAG-dye complex, and quantifying the released dye according to the protocol provided by the manufacturer (Biocolor LTD., No. B1000). A standard curve (chondroitin sulfate included in the kit) was determined, and linear regression was performed on the standard curve to obtain the equation of the curve and the regression coefficient R2.

[0217] The hyaluronic acid content was determined by precipitation of total GAGs by addition of ethanol saturated with acetic acid, resuspension of HA by addition of cetylpyridinium, purification of free HA by repeating the previous step, and resuspension of HA in distilled water. The concentration of hyaluronic acid present in the samples was determined according to the protocol provided by the manufacturer. A standard curve (chondroitin sulfate included in the kit) was obtained, and linear regression to the standard curve gave the equation of the curve and the regression coefficient R2.

[0218] 7.2 Results - Measurement of sulfated GAG content To evaluate the GAG ​​concentration in the three in vitro skin samples, 19 mm diameter samples were dehydrated and weighed. The samples were then cut, digested with Proteinase K, and pulverized. The amount of GAG was measured according to the instructions in the Blyscan kit (B1000). A standard curve was generated with chondroitin sulfate standards. Table 11 shows the chondroitin sulfate equivalent (μg CSE) concentrations in the skin.

[0219] [Table 11]

[0220] To assess the portion of nonspecific signal that may be related to unseeded scaffolds, the same extraction protocol and measurement of sulfated GAG concentration was performed on three unseeded scaffolds: the absorbance corresponding to 0.44 μg chondroitin sulfate equivalents (CSE) was measured and a threshold corresponding to the nonspecific signal was determined and subtracted from each measurement.

[0221] - Determination of HA content To assess the hyaluronic acid (HA) content in in vitro skin samples, the same samples were analyzed using the Biocolor Purple-Jelley kit (H1000). A standard curve was established with HA standards and the concentrations measured in in vitro skin are reported in Table 12. The nonspecific signal was 0.04 μg HA, which corresponds to the signal of the unseeded membrane scaffold.

[0222] [Table 12]

[0223] -A test to measure the amount of GAG and HA per mg of dry mass To report measurements of GAGs and HA relative to the dry mass of the samples, a similar procedure was performed after oven drying (50°C, 6 h) of in vitro skin samples of the same dimensions. However, the total mass of sulfated GAGs after subtraction of nonspecific signals was 0.04 μg for ECS and 0.01 μg for AH. This corresponds to a density of 1.3 ng ECS / mg sulfated GAGs and 0.3 ng / mg HA, much lower than the wet mass measurements, whereas a higher density would have been expected. Thus, the drying procedure appears to interfere with the quantification of GAGs in in vitro skin.

[0224] 7.3 Discussion To assess the amount of GAGs in in vitro skin, quantification techniques for sulfated GAGs and HA were applied. The mass densities of GAGs in these skins were measured: an average density of sulfated GAGs of 87.9 ± 2.2 ng ECS / mg (chondroitin sulfate equivalent) and an average density of HA of 57.86 ± 4.9 ng / mg were obtained. This density refers to the wet mass, as drying interferes with the measurements. This may be due to extraction problems after drying, e.g., collapse of the membrane scaffold around the GAGs or degradation during drying.

[0225] In a study by Armstrong and Bell, hyaluronan in rabbit skin was measured to be 551 ng / mg wet skin (Armstrong SE, Bell DR. Anal Biochem 2002;308(2):255-64). This result is 1.67 higher than the presently published result, but is within the same order of magnitude. This difference may be explained by physiological changes, e.g., a decrease in the amount of hyaluronan in the skin with age or site (Barbosa I,et al.Glycobiology 2003;13(9):647-53;Templeton DM.Connect Tissue Res 1988;17(1):23-32). Varma et al. reported that the distribution of GAGs in the skin of various animals, including rats, is essentially half HA, the other half chondroitin sulfate GAGs, and a small proportion of heparin (Varma RS, Karger; 1982; Fabianek, Herp and Pigman. Comparative Biochemistry and Physiology. Volume 14, Issue 1, January 1965, Pages 21-28). Fabianek et al. showed that the amount of chondroitin sulfate (non-HA GAG) in rabbit skin was 319 μg / g dry mass, using a conversion factor of 3:1 to wet mass. The inventors found that when this correction was applied, the concentration of chondroitin sulfate (non-HA GAG) was 106 ng / mg wet mass (Fabianek, Comparative Biochemistry and Physiology. Volume 14, Issue 1, January 1965, Pages 21-28). This value is 5.47 times smaller than that found in control rabbit skin measured in this study. This difference may be explained by factors related to sample processing, as well as differences in extraction and GAG measurement techniques.

[0226] In the in vitro skin, the total amount of GAGs, which corresponds to the sum of sulfated GAGs (chondroitin sulfate) and HA, was 145.8ng / mg, whereas in the control rabbit skin measured in this study, it was 883.9ng / mg. Compared to the analyzed rabbit skin, there was 5.2 times less HA and 6.7 times less sulfated GAGs. However, when comparing these results with those of Armstrong and Bell and Fabianek et al., there is a difference of minus 17% in sulfated GAGs and 9.5 times less HA between rabbit skin and in vitro skin.

[0227] Mass levels of sulfated GAGs of 87.9 ± 2.2 ng / mg and HA levels of 57.86 ± 4.9 ng / mg were measured in skin in vitro. These values ​​are lower than those measured in rabbit skin, but the concentration of sulfated GAGs (chondroitin sulfate) is close to the value found in the literature of 106 ng / mg in rabbit skin.

[0228] F. In vitro skin characterization 1. Differential Scanning Calorimetry (DSC) A property commonly used to characterize collagen, whether native, structurally modified, or chemically modified by tanning, is its hydrothermal stability. It is defined as the effect of moist heat on the integrity of the material, usually defined in terms of denaturation transitions (AD Covington, RA Hancock, IA Ioannidis, J. Soc. Leather Technologists Chemists 73, 1989) 1:8). The value of this parameter, which depends on the processing (tanning), is an important physical parameter in the characterization of leather and its typical use. The covalent bonds generated by crosslinking of collagen fibers with tanning agents, especially during the tanning process, increase the size of the covalent units by intermolecular and intramolecular crosslinks and increase the denaturation temperature (Td) (S. Menashi, A. Finch, PJ Gardner, DALedward, Biochem. Biophys. Acta 144 (1976) 623:625). Denaturation is defined as the transition from triple helix to randomly bent conformations that occurs in the domains between the crosslinks (M. Komanowsky, J. Am. Leather Chemist Assoc. 86(8)(1991) 269:28). The bonds that stabilize the coiled coil are hydrogen bonds, hydrophobic bonds, van der Waals bonds, and interactions between oppositely charged residues in the side chains. The nonrandom distribution of ionizable hydrophobic side chains along the repeat unit results in charged and hydrophobic patches that contribute to the stabilization of the structure through electrostatic and hydrophobic interactions (GS Young, Stud. Conserv. 43(2)(1998) 65:79).

[0229] This increase in Td depends on the nature and type of reactive groups involved. Thus, the method of tanning has a direct effect on the shrinkability of leather. Traditionally, vegetable tanned leathers have a Td of around 70 ° C, while chrome tanned leathers have a Td of around 80-90 ° C. Their synthetic tanned counterparts have a Td of about 75 ° C (R. Komsa-Penkova, R. Koynova, G. Kostov, BG Tenchov, Biochem. Biophys. Acta 129 (1996) 171: 181; C. Chahine, Changes in hydrothermal stability of leather and parchment with deterioration: a DSC study).

[0230] The hydrothermal stability of collagen contained in skin and leather is characterized by the shrinkage of the material when heated in water at a constant temperature. Differential scanning calorimetry (DSC) makes it possible to study the enthalpy changes associated with the denaturation of collagen, the shrinkage being its macroscopic manifestation. The aim of this study is to analyze the hydrothermal stability of leather obtained according to the method of the invention. As reference materials before tanning, unseeded membrane macroporous scaffolds and in vitro skin were used. To compare the effect of the tanning method on the denaturation temperature (Td) of the tanned leather, the in vitro skin according to the invention by two different methods, tanned leather, aldehydes were tested. The results of the analysis and measurements showed that the in vitro skin tanned leather has a hydrothermal stability close to that of tanned leather of animal origin.

[0231] 1.1 Differential Scanning Calorimetry (DSC) Protocol The measurements were performed using a 2014 Polymer DSC calorimeter (Netzsch). Temperature and heat flux were calibrated with indium (melting point: 156.6 °C, latent heat of fusion: 28.45 J / g). The samples were weighed (3 mg) and immersed in water for several hours before being sealed in an aluminum crucible. The heat change was measured relative to that observed in a reference sample while purging with nitrogen atmosphere. The two trays (reference and sample) were heated separately at a constant rate of 5 °C / min in the range of 10-95 °C. The instrument records the heating rate required to keep both tests at the same temperature. When the test sample undergoes an endothermic process, the energy input required to maintain the temperature increases, resulting in a signal on the temperature rise curve.

[0232] 1.2 Results Differential scanning calorimetry (DSC) was used to measure the denaturation temperature (Td) of the samples, which is obtained by the analysis of the intervals of variation of the transition temperatures (onset and offset) with a peak similar to the denaturation temperature (Td). This last method is more adapted to the current complex systems (scaffolds and biomaterials) (Carsote, C., Badea, E. Herit Sci 7, 48 (2019)).

[0233] The measurements were performed with a Netzsch DSC 2014 Polymer Calorimeter at a constant rate of 5 °C / min in the range of 10-95 °C. Figure 14 shows the DSC curves of the different samples: macroporous scaffold (FSC), in vitro skin (FP) and in vitro skin leather tanned with aldehydes (CF-1). Figure 15 shows the resolution of the peaks and the different types of denaturation temperatures present in the samples: onset temperature of the transition initiation (T onset ), the denaturation temperature (Td) corresponding to the peak, and the offset temperature of the end of the transition (T offset ), indicates the decision.

[0234] Looking closely at the thermograms obtained for the different samples, they can be divided into two (for the FSC sample) and three (for the FP, CF-1, and CF-2 samples) peaks / deconvolutions with intervals corresponding to the different denaturation temperatures occurring in this complex system. The denaturation temperatures of each sample and their respective peaks / deconvolutions are shown in Table 13. The results show that the lowest recorded Td of 44.2 °C corresponds to the first peak / deconvolution of the unseeded macroporous scaffold (FSC), with an almost undetectable denaturation enthalpy. The first peak / deconvolution of the in vitro skin (FP) increases to 52.6 °C, and in terms of enthalpy, this minority of peaks could correspond to collagen not modified by tanning, with a value close to the value of 54.2 °C obtained by Carsote et al. for non-chemically modified collagen (Carsote, C., Badea, E. Herit Sci 7, 48 (2019)).

[0235] The Td of the second peak / deconvolution of the unseeded macroporous scaffold (FSC) increased to 77.4 °C. This peak is the largest in terms of enthalpy and should correspond to the collapse of the structure of the unseeded scaffold at high temperatures. The second peak / deconvolution of the in vitro skin leather (FP) and aldehyde-tanned leather (CF-1) have almost the same denaturation temperatures, 72.1 °C and 71.8 °C, respectively. This second peak / deconvolution has a higher enthalpy than the first peak / deconvolution and should correspond to collagen fibers, some chemically modified, entangled within the scaffold, forming a more heat-resistant composite, and biomaterials.

[0236] The highest denaturation temperatures recorded for in vitro skin (FP) and aldehyde-tanned leather (CF-1) are 83.5 and 82.9, respectively. This third and last peak with the highest enthalpy is believed to correspond to collagen fibers chemically bonded by the tanning agent and to some extent chemically modified biomaterials intertwined in the scaffold. The presence of collagen and chemically modified biomaterials makes the composite resistant to thermal denaturation and the structure of the composite is maintained up to about 83 °C compared to 77.4 °C for the unseeded macroporous scaffold.

[0237] [Table 13]

[0238] These results show the collagen denaturation of tanned leather and the change in enthalpy related to the temperature at which the phenomenon occurs (Figure 14 and Table 13). In particular, the covalent bonds generated during tanning stabilize the collagen fiber bonds and increase their denaturation temperature (Td). This stabilization by tanning is observed especially when it increases the Td of the first peak / deconvolution of the untanned in vitro skin (FP) at 52.6°C to 59.3°C for the aldehyde-tanned comparison (CF-1). Regarding the second and third peaks / deconvolution, we observed that the formation of a complex of mixed collagen, biomaterial and scaffold has a dominant effect on the denaturation temperature (Td) compared to tanning. Indeed, the Td of these second and third peaks / deconvolution are similar between the in vitro skin and the aldehyde-tanned leather (CF-1).

[0239] These results indicate that the effect of tanning agents on the Td of animal leathers is also observed in in vitro skin-tanned leathers.

[0240] Larsen et al. measured the Td of various tanned leathers, including mimosa and sumac leathers (Larsen, Rene & Vest, M. & Nielsen, K.. (1993), Journal of Society of Leather Technologists and Chemists. 77. 151-156). They compared the Td of untanned leathers with that of these two leathers. The results showed that the Td increased significantly after tanning, being 75.3°C and 79.3°C for mimosa and sumac, respectively, compared with 56.5°C for untanned leather. Furthermore, heterogeneity of the measurements between the two leather samples was reported, which was explained by the tanning-dependent organization of collagen fibers.

[0241] As mentioned before, the thermograms obtained for the different samples of the present invention are complex, since they consist of two or three different peaks / deconvolutions. Similar analyses of the decomposition of DSC thermograms have been reported in thermogravimetric studies of copolymers or composite systems (Luo K, Wang L, Chen X, Zeng X, Zhou S, Zhang P, Li J. Polymers (Basel). 2020 Nov 9; 12 (11): 2631; Athanasoulia, Ioanna-Georgia and Tarantili, Petroula A. Pure and Applied Chemistry, vol. 89, no. 1, 2017, pp. 141-152). The DSC curves corresponding to this composite system showed endothermic fluctuations located in the region of decomposition and rearrangement of the polymer chains. This behavior allowed the determination of the onset temperature of denaturation (T onset ), peak temperature (Td), and offset temperature of denaturation (T offset ) allowed us to define the denaturation region of the system. These temperatures successively reflect the onset of denaturation, the optimization of the deformation, and the post-deformation stability of the chains, which correspond to the rearrangement of the different chains. This phenomenon is similar to the observations made with the samples in this study.

[0242] In each thermogram of FP and FC-1, three fluctuations are clearly visible, bounded by the stability before and after denaturation. This behavior is not detected in the case of the non-composite scaffold (FSC) that does not contain biological material, unlike the other samples. In the case of the cultured in vitro skin (FP), the composite system consists of a macroporous scaffold (FSC) and a biological material that contains cells together with an extracellular matrix (ECM) rich in collagen fibers. The tanned leathers (CF-1 and CF-2) constitute a complex ternary system that includes the scaffold (FSC), the biological material (especially the cross-linked collagen fibers) and the tanning agent. From this, a denaturation region corresponding to the state of the material is defined, as shown by the decomposition of the curves in Figure 15. From these decompositions, the leathers of the present invention show similar denaturation characteristics, with a denaturation temperature range that varies between 55.8 and 88.7 °C in the aldehyde tanning method. The average Td (72.2°C) properties of the inventive comparison are still very close to the range of new aldehyde-tanned leathers (76.6; 87°C) (Bai, Xue & Jinming, Chang & Chen, Yi & Fan, Haojun & Shi, Bi. (2013). Journal of the American Leather Chemists Association. 108. 404-410).

[0243] Another important factor that strongly influences the Td value is the moisture content of the material. Thus, the lower the moisture content, the higher the Td. This phenomenon is due to the formation of strong inter- and intra-molecular bonds, as well as ionic bonds, between acidic and basic groups during drying, which promotes thermal stabilization. Classically, the phenomenon of shrinkage is measured in aqueous media, which is why Td measurements are performed with an excess of water (A. Finch, DA Edward, Biochim. Biophys. Acta 278 (1972) 433-439).

[0244] In the present study, we maintained these classical conditions in realizing the measurements. Therefore, taking these conditions into account, the DSC analysis is very useful to evaluate the thermal stability of leather and allows a comparison of the two tanning methods adopted.

[0245] To obtain a correlation between the measurements obtained with DSC and the heat resistance measurements commonly used in the tanning industry, we used a shrinkage temperature tester (Tr) specially designed for leather. This device consists of a beaker filled with water, in which a leather sample is immersed under mechanical tension. The leather sample is then subjected to a range of temperatures (20-100 °C) with a heating plate placed under the system. The shrinkage of the sample at a given temperature (Tr) is recorded by the displacement measured with a percentage dial gauge (Borletti). The shrinkage temperatures (Tr) of the in vitro skin (FP) and the aldehyde-tanned leather (CF-1) were measured and are shown in Table 14. As a result, the Tr of the in vitro skin (FP) oscillates around 65 °C, which corresponds to the temperature between the first and second peaks / deconvolution obtained with DSC (Figure 14, Table 13, FP). On the other hand, the Tr of aldehyde-tanned leather oscillates around 75°C, which corresponds to the temperature between the second and third peaks obtained by DSC (Figure 14, Table 13, CF-1).

[0246] [Table 14]

[0247] Measurement of shrinkage temperature (Tr) using a tester is less accurate than that by DSC (measurement of thickness of thin samples and displacement by observer). Nevertheless, this method is commonly used in the tanning industry and provides a useful framework for comparing DSC measurements with measurements by tanners and craftsmen (GS Young, Stud. Conserv. 43(2)(1998)65:79). The results show that the aldehyde-tanned leather of the present invention has a thermal stability very close to that of animal leathers freshly tanned by the same method.

[0248] 2. Mercury Intrusion Porosimetry 2.1 Materials and Methods Mercury porosity measurements were performed using a micrometer porosimeter (AutoPore IV 9500, Micrometrics) with a maximum applied pressure of 413 MPa. The samples used for this measurement were the scaffold (FSC) and the in vitro skin aldehyde-tanned leather (CF-1).

[0249] Measurements were performed at 24°C. FSC samples were pre-dried at 50°C for 24 hours and CF-1 samples at 80°C for 24 hours. Equilibration time for each pressure point was 10 seconds. Blank correction was performed before measurements. Samples were cut into 3 cm x 2 cm pieces (approximately 0.6 g) and first degassed and then filled with low pressure distilled mercury. The mercury-filled penetrometer was removed from the low pressure port, weighed and then placed in the high pressure port. Mercury was introduced into the pores of the sample with increasing pressure. Using the recorded pressure and the volume of mercury introduced into the pores, the porosity was calculated.

[0250] 2.2 Results -In vitro porosimetry properties of skin leather The total porosity (ε) calculated from the MIP test is shown in Table 15.

[0251] [Table 15]

[0252] The scaffold (FSC) has a high percentage of total porosity (ε) (87.21%) according to the data extracted from the MIP analysis shown in Table 15. The structural properties of the scaffold according to the present disclosure show high porosity with an average pore size of 20 μm in average diameter, compatible with the seeding and infiltration of animal cells (Al Tawil, E., et al. European Polymer Journal).

[0253] In in vitro skin (FP), the total porosity decreased sharply, reaching 14.78%. This decrease is explained by the occupation of the pore volume by biological materials, such as cells and components of the extracellular matrix (ECM) secreted by cells. These biological materials are essential components of the neoformed tissue of in vitro skin and are present in tanned leather as well as in conventional animal leather (He, Xiu et al. Journal-American Leather Chemists Association. 114. 41-47).

[0254] Tanning is a process that transforms conventional or in vitro skin into leather by creating chemical crosslinks between the collagen fibers of the ECM. This network crosslinking results in the creation of spaces called macropores, mesopores, and nanopores between the collagen chains. This porosity of leather is a highly valued property and is responsible for its breathing phenomenon. The leather tanned with aldehydes (CF-1) showed a reduction in the total porosity to 51.63%. These results are very similar to those obtained with chrome-tanned sheepskin leather (Table 15).

[0255] 2.3 Discussion The porosimetry of a material is one of the essential characteristics of that material and is a parameter of choice that influences the type of product in which it is used and the framework of its use. In the case of leather, the porosimetry of different types of leather guides its final application in decoration, clothing, leather goods, furniture, etc. The porosimetry also indicates the changes during the manufacturing process, especially the retanning and finishing stages. In this report, mercury intrusion porosimetry (MIP) was used to measure the porosimetry properties of the leather obtained from the in vitro skin of the present disclosure. The results showed that the scaffold has sufficient porosity for the culture of animal cells. The proliferation of cells within the scaffold allowed the colonization of the scaffold by cells and the deposition of biological materials, which are essential components of in vitro skin. The leather of the in vivo skin showed a total porosity comparable to that of conventional animal leather.

[0256] 3.Water vapor permeability 3.1 Materials and Methods -material Permeability measurements were performed using a permeation diffusion meter (IGAsorp, Hiden Isochema). It consists of two compartments, upstream and downstream of the sample. During the measurement, the permeant (water molecules) is introduced into the upstream compartment, while the downstream compartment is swept with dry gas (N2 BIP, Air Products). Hydrometer detection is used. During the measurement, the moisture-enriched gas is measured with a mirror hygrometer, General Eastern probe 1311 DR (Elcowa - Mulhouse).

[0257] The experimental setup used for the measurements was as follows: It is thermostated at -23°C and consists of two compartments, one made of stainless steel and the other made of plexiglass; - a feed upstream of the sample, where the purge gas, industrial grade nitrogen, is circulated in the first step, and then the permeate is circulated in the second step; -Feeder downstream of the film, through which BIP type nitrogen (gas) is circulated as carrier gas for the mirror hygrometer.

[0258] The measurements were performed at 23°C and 50% relative humidity. The sample used in this study was aldehyde-tanned leather (CF-1). The active surface area of ​​the sample was 3.6 cm 2 It is.

[0259] -Experimental setup Differential transmission measurements are performed in two successive steps. The first step, called the purge step, removes as much moisture and residual molecules as possible from the sample, cell and gas circuit. Nitrogen is then swept through the upstream and downstream compartments until a stable low moisture content is obtained. In the second step, the measurement step, water vapor at 50% relative humidity is introduced into the cell. The water vapor diffuses through the sample and is carried by a carrier gas to a detector (mirror hygrometer) downstream of the sample. This change in flow is recorded in real time by direct measurement of the change in dew point temperature.

[0260] - Flow rate and infiltration calculations Flow velocity f=9.3cm 3 / s, and dry nitrogen (vapor pressure p in The receiving compartment, where BIP nitrogen is supplied, is not completely free of water and its vapor pressure is p in From p out (vapor pressure at the compartment inlet and outlet). The flow rate J(L,t) is given by:

[0261]

number

[0262] where R is the ideal gas constant, S is the exposed surface, and Tr is the temperature (K) at the time of measurement.

[0263] The hydraulic conductivity P is calculated from the following formula:

[0264]

number

[0265] J st is the steady-state flux, L is the thickness of the sample, and Δa is the difference in activity on either side of the sample.

[0266] 3.2 Results The water vapor transmission rate was measured using a permea-diffusiometer at 23°C and 50% relative humidity. The real-time measurements are shown in Table 16. The permeability coefficient of the aldehyde-tanned leather (CF-1) was 3.47 mg / cm 2 Representative values ​​for cowhide leather have also been added to this table (Cattle leather: Radwag wagi elektroniks-testing laboratory. https: / / radwag.com / pl / ).

[0267] [Table 16]

[0268] 3.3 Discussion There are various methods for measuring the water vapour permeability of leather, but the most widely used is the measurement according to ISO 14268:2013 (Leather - Physical and mechanical tests - Determination of water vapour permeability (ISO / FDIS 14268:2012) 2013). Water vapour permeability depends on many operations in the leather manufacturing process, including tanning and finishing. Leather is produced in the laboratory by combining scaffolds with animal cells.

[0269] Leather (CF-1) was produced using aldehyde tanning. The water vapor transmission rate of CF-1 was 3.47 mg / cm 2 .h was recorded. Generally, leather is considered to have good water vapor permeability (WVP) and is known to have a transmission factor of more than 20 and a water vapor permeability value of more than 0.8 (Leather international 18 September 2001). The present results show that this leather has good water vapor permeability as seen in conventional leathers. Water vapor permeability analysis of various types of animal leathers has been carried out (Skenderi Z., et al. Ergonomics 2013, June 12-15, 2013, Zadar, 9-14; Smiechowski K., et al. Journal of the Society of Leather Technologists and Chemists, ISSN 0144-0322, Vol. 98, Num. 6, 2014; Radwag wagi elektroniks-testing laboratory. https: / / radwag.com / pl / ). measured the water vapor permeability of different leathers using standard methods and methods developed in their laboratory. The various types of leather selected for their study had permeabilities ranging from 0.3 to 4.9 mg / cm for samples with a thickness of around 1.2 mm. 2On the other hand, a study published by the Radwag Testing Laboratory showed that cowhide leather used in the footwear industry has a water vapor permeability ranging from 0.7 to 5.3 mg / cm, depending on the manufacturing method: wrinkled, pressed, or untreated. 2 It varies between .h.

[0270] Based on these results, we observed that the in vitro skin leather has water vapor permeability comparable to conventional leather. The recorded values ​​are very close to published values ​​for measurements made on animal-derived leather samples. It is important to note that the tests were performed on unfinished tanned leather with an average thickness of 0.45 mm. Water vapor permeability has been shown to increase significantly as the thickness of the leather decreases (Smiechowski K., et al. Journal of the Society of Leather Technologists and Chemists, ISSN 0144-0322, Vol. 98, Num. 6, 2014).

[0271] Considering these results, measurements of the water vapor permeability of in vitro skin leathers with different thicknesses, tanning methods, and potential finishes will allow for discovering manufacturing factors important for altering the water vapor permeability of in vitro skin leathers. [Brief description of the drawings]

[0272] [Figure 1] Overview of the fabrication of polyester asymmetric membrane scaffolds by the NIPS process. [Diagram 2] Overview of symmetric membrane scaffold fabrication methods by physical and / or chemical methods to remove asymmetric scaffold smooth surfaces. [Diagram 3]Scanning electron microscopy images of the porous side (a), inside the macropores (b), cross-section (c), and smooth side (d) of a biodegradable polyester membrane scaffold. The porous side (a) shows the presence of honeycomb-like macropores on top of the porous side, micropores connecting the macropores (b), the thin thickness of the smooth side opposite the porous side, and the absence of macropores and micropores on the smooth side. Scale bars: 400 μm (a, c, and d), 200 μm (b). [Figure 4] Scanning electron micrographs of polyester sponge seeded with fibroblasts, fixed and dried, with cell bodies lining the entire surface of the polyester sponge scaffold. Scale bars: 400 μm (a and b), 50 μm (c). [Diagram 5] Scanning electron micrographs of fibroblast-seeded polyester membranes, showing that cells colonize the scaffold while depositing abundant extracellular matrix fibers. Scale bars: 400 μm (a and b) and 50 μm (c). [Figure 6] Proliferation of fibroblasts cultured on polyester sponge (circles, solid) and membrane (squares, dashed) scaffolds at various time points. [Figure 7] Tissue production by fibroblasts seeded on polyester membrane scaffolds 1 week (a) and 3 weeks (b) after seeding, as observed by fluorescence microscopy. Cell-derived collagen areas were observed by fluorescence microscopy using anti-collagen I antibody (Sigma, Cat. No. C2456). Cell nuclei were stained with DAPI (blue) and collagen I (green). Scale bar: 500 μm. [Figure 8] Tanning process involving tanning / re-tanning process of tissue samples to obtain tanned leather. Steps a) tanning and re-tanning process are brought into the tanning drum, b) surface image of tanned leather under high-end microscope, scale bar: 2mm, c) final result of obtained tanned leather. [Figure 9] Schematic representation (right) of open pore boundaries at the surfaces of polyester sponge (top) and membrane (bottom) scaffolds based on scanning electron microscopy images (left). [Figure 10]Distribution of macropore surface porosity (%) on the surface of sponge (grey) and membrane (black) scaffolds. For membrane and sponge scaffolds, median ± MAD% surface porosity was 66.11 ± 8.53 and 21.49 ± 8.65, respectively. p-value=0.0011655 by Wilcoxon rank sum test, **p<0.01. [Figure 11] Pore ​​wall thickness between open macropores in sponge (grey) and membrane (black) scaffolds, calculated by subtracting the distance between the centers of two pores by the sum of their radii. Median ± MAD wall thickness for membrane and sponge scaffolds was 29.37 ± 12.40 and 70.15 ± 26.60, respectively. p-value = ****p < 0.0001 by Wilcoxon rank sum test. [Figure 12] Representative results of traction mechanical tests performed on polyester scaffolds according to ISO 3376:2020 standard using an Instron 34SC-1 system with a 50N load cell (CAT.NO.:2530-50N) and manual clamping grips (CAT.NO.:2710-203). a) Stress / strain traction curves of polyester membrane scaffolds measured, b) Elastic limit of the specimens measured by measuring Young's modulus, c) Tear load of specimens measured. Speed ​​was set and controlled at 100mm / min during the test. Strength and displacement measurements were recorded. Stress was calculated by the initial thickness and width of the specimen. Elongation was calculated by the initial length and observed displacement. [Figure 13] Orientation of biomaterial fibers (cells and extracellular matrix) at the surface of in vitro skin. Orientation of biomaterial field lines at the surface of in vitro skin obtained by scanning electron microscopy (A and B): Scale bar: 100 μm. Orientation histogram showing the frequency of biomaterial oriented regions at the surface of the in vitro skin sample (C). [Figure 14] Thermograms of DSC analysis of samples: macroporous scaffold (FSC), in vitro skin (FP) and in vitro skin leather tanned with aldehydes (CF-1). [Figure 15]Resolution of the peaks and determination of the denaturation temperatures of the different species present in the sample: the onset temperature for the start of the transition (Tonset), the denaturation temperature corresponding to the peak (Td) and the offset temperature for the end of the transition (Toffset).

Claims

1. Leather manufacturing method including the following steps: i) In vitro, fibroblasts are cultured on a honeycomb porous polymer scaffold to obtain tissue. Here, the surface of the scaffolding includes macropores with a diameter of 100 to 280 μm, a surface pore area distribution of 40% to 95%, and a pore wall thickness of less than 70 μm, ii) The tissue is tanned to form the leather.

2. The method according to claim 1, wherein the scaffolding includes polyester.

3. The method according to claim 1 or 2, wherein the scaffolding comprises biodegradable polyester and / or bio-derived polyester.

4. The method according to claim 1, wherein the scaffolding contains polyester at a concentration of 1 to 50% (w / w).

5. The method according to claim 1, wherein the macropores are interconnected with the micropores.

6. The method according to claim 5, wherein the micropores have a diameter of less than 20 μm.

7. The method according to claim 1, wherein the scaffolding has a tensile strength of 0.1 to 10 MPa.

8. The method according to claim 1, wherein the scaffolding has a Young's modulus of 5 to 1 MPa.

9. The method according to claim 1, wherein the scaffold is obtained by non-solvent-induced phase separation (NIPS).

10. The scaffolding is an asymmetrical scaffolding that includes macropores that are uniform but not regularly distributed throughout the entire thickness of the scaffolding, A first surface containing macropores opening to the surface, with a diameter of 100 to 280 μm, a surface pore area distribution of 40% to 95%, and a pore wall thickness of less than 70 μm, The method according to claim 1, wherein a second plane of symmetry is exhibited, which contains only nanopores with a diameter of less than 10 nm.

11. The method according to claim 1, wherein the scaffold is a symmetrical scaffold having two surfaces containing macropores with a diameter of 100 to 280 μm, a surface pore area distribution of 40% to 95%, and a pore wall thickness of less than 70 μm.

12. The method according to claim 1, wherein a bioactive molecule is grafted onto the surface of the scaffold.

13. The method according to claim 12, wherein the bioactive molecule is collagen, gluside, or a gluside-derived molecule such as glycosaminoglycan.

14. A tissue obtained after step a) of the method according to claim 1, comprising fibroblasts cultured on the porous scaffold described in claim 1.

15. The tissue according to claim 14, having a denaturation temperature (Td) determined by at least two differential scanning calorimetry (DSC) assays.

16. The structure according to claim 14, wherein the structure is isotropic on the surface of the structure.

17. The tissue according to claim 14, wherein the proportion of the total fatty acid mass in the total dry mass of the tissue is less than 1% (w / w).

18. Tanned leather obtained by the method described in claim 1.

19. The tanned leather according to claim 18, having a denaturation temperature (Td) determined by at least two differential scanning calorimetry (DSC) assays.

20. The tanned leather according to claim 18, having a total porosity of 30-60%.

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