Method for producing leather using cell culture on cellular macroporous polymer scaffolds

Abstract

The present disclosure relates to a method for producing leather comprising the step of culturing cells on a cellular macroporous polymer scaffold. The present disclosure also relates to leathers obtainable by this process.

Description

Technical Field

[0001] The present disclosure relates to a method for producing leather, comprising the step of culturing cells on a honeycomb macroporous polymer scaffold. The present disclosure also relates to leather obtainable by the method. Background Art

[0002] Leather is the most resource-intensive material used in the fashion industry, far more than cotton or polyester. While animal leather is favored for its durability and beauty, it presents multiple environmental challenges: intensive use of natural resources, animal welfare and working conditions. In addition, for many applications, leather needs to be free of defects such as scratches or insect bites. As traditional leather production relies mainly on cattle farming, up to 80% of raw hides are discarded because they do not meet quality standards.

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

[0004] Although synthetic leather is developed to solve some of these problems, it lacks the quality of natural leather. Attempts to make engineered leather have been described previously. For example, EP1589098 describes a method for growing fibroblasts on a three-dimensional bioactive scaffold. The scaffold can be made of collagen waste from a tanning process ("split"), pure collagen microparticles, collagen waste particles or synthetic scaffolds (e.g., made of polymers such as HYAFF). WO2017 / 184967 describes culturing stem cells or keratinocytes on collagen or PET films with 4 μm micropores. However, these culture conditions are not optimal for cell growth and collagen production.

[0005] Tanning step decomposes most cells, molecules and extracellular matrix components. The only components that resist storage and tanning are collagen fibers and elastin fibers, which are present in the leather in the final state and secreted by fibroblasts (Sharphouse, JH Leather Technician's Handbook.Leather Producer's Association.p.104.ISBN0-9502285-1-6). These collagen and elastin fibers are responsible for the mechanical properties of leather. For this reason, the secretion of the network-induced extracellular matrix (collagen, elastin etc.) of the support by fibroblast proliferation is a key step in leather development.

[0006] Therefore, there is still a need to develop cell culture methods that promote cell proliferation and production of collagen necessary for high-quality leather. Summary of the invention

[0007] The inventors have developed a new method for producing high-quality leather by culturing fibroblasts in vitro on a honeycomb macroporous polymer scaffold, which has effective high cell attachment, cell culture and long-term proliferation while minimizing stress and damage to the cells, and allows fibroblasts to produce larger amounts of collagen, thereby producing a collagen-rich extracellular matrix to be tanned and converted into a leather-like biomanufacturing material.

[0008] In particular, the inventors produced scaffolds with uniform but irregular honeycomb pore distribution, resulting in a well-organized structure with uniform pore spacing. Interestingly, this scaffold can maintain cell viability and is accompanied by higher extracellular matrix secretion compared to other sponge-like macroporous scaffolds.

[0009] According to the present invention, honeycomb-shaped holes are formed on the surface of the membrane scaffold, and neatly arranged glove-shaped holes extend deep therein, and the interconnected holes can guide multicellular tissue and increase the deposition of fibrous collagen. The well-organized membrane scaffold increases stiffness and provides a suitable microenvironment for cell colonization and development and the deposition of extracellular matrix (ECM) fibers.

[0010] Such a honeycomb macroporous scaffold, also referred to herein as a membrane scaffold, is particularly useful for obtaining a tissue that is particularly suitable for producing a leather-like technical product after a tanning step.

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

[0012] In a specific embodiment, the support is an asymmetric support comprising macropores uniformly but irregularly distributed throughout the thickness of the support, in particular, it has a first face and a second opposing face, the first face comprising macropores with a diameter of 100-280 μm, preferably 110-225 μm, open at the surface, and 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 the second opposing face only comprises nanopores with a diameter of less than 10 nm, preferably 5 nm. In another specific embodiment, the support is a symmetrical support containing macropores, which has two faces, wherein the faces contain macropores with a diameter of 100μm-280μm, preferably 110μm-225μm, wherein the surface pore area distribution is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and the pore wall thickness is less than 70μm, preferably less than 60μm, more preferably 2μm-50μm.

[0013] In a preferred embodiment, bioactive molecules are grafted onto the surface of the scaffold. Preferably, the bioactive molecules are collagen, carbohydrates, or carbohydrate-derived molecules, such as glycosaminoglycans.

[0014] In another aspect, the present disclosure relates to a tissue obtainable by the culturing step a) of the method of the present invention, and to a tanned leather obtainable by the method as described above.

[0015] Finally, the present disclosure relates to the use of a macroporous polymer scaffold for the production of leather, preferably by tanning a tissue obtained from a fibroblast cell culture on said macroporous 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. BRIEF DESCRIPTION OF THE DRAWINGS

[0016] Figure 1 : Schematic diagram of the method for producing polyester asymmetric membrane scaffolds using NIPS technology.

[0017] Figure 2 : Schematic diagram of producing a symmetric membrane scaffold by removing the smooth surface of the asymmetric scaffold by physical and / or chemical methods.

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

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

[0020] Figure 5 : Scanning electron micrographs of polyester films seeded with fibroblasts. Cells colonize scaffolds rich in extracellular matrix fiber deposition. Scale bars: 400 μm (a and b) and 50 μm (c).

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

[0022] Figure 7 : Tissues generated by fibroblasts seeded into polyester film scaffolds after 1 week (a) and 3 weeks (b) of seeding and observed by fluorescence microscopy. Collagen secretion of cells was observed by fluorescence microscopy using anti-collagen I antibody (Sigma, ref. C2456). Cell nuclei were stained with DAPI (blue) and collagen I (green). Scale bar: 500 μm.

[0023] Figure 8 : Tanning process of tanning tissue samples to obtain tanned leather using tanning / retanning processes: a) Tanning and retanning processes in a tanning drum, b) Surface image of tanned leather obtained by optical microscopy, scale bar: 2 mm. c) Final product of the tanned leather obtained.

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

[0025] Fig.10 : Distribution of surface porosity percentage (%) of macropores on 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. Wilcoxon rank sum test p-value = 0.0011655, **p < 0.01.

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

[0027] Fig.12 : Representative results of traction mechanical testing of polyester stents using an Instron 34SC-1 system equipped with a 50N load cell (CAT.NO:2530-50N) and a manual fastening clamp (CAT.NO.2710-203) in accordance with ISO3376:2020. a) Record the stress / strain traction curve of the polyester film stent, b) Determine the elastic limit of the sample to measure the Young's modulus, and c) Determine the tear load of the sample. The speed was applied during the test and set to 100mm / min. Record the strength and displacement measurements. The stress was calculated using the initial thickness and width of the sample. The elongation was calculated using the initial length and the observed displacement.

[0028] Fig.13 : Directivity of biomaterial fibers (cells and extracellular matrix) on the surface of in vitro skin. Directional field lines of biomaterial on the surface of in vitro skin obtained by scanning electron microscopy (A and B), scale bar: 100 μm. Directivity histogram showing the frequency of the biomaterial directional field on the surface of in vitro skin samples (C).

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

[0030] Fig.15 : Decomposition of peaks and determination of denaturation temperatures of different substances in the sample: Onset temperature (T onset ), corresponding to the denaturation temperature of the peak (Td) and the shift temperature at the end of the transition (T offset ). DETAILED DESCRIPTION

[0031] The inventors have developed a method for producing high-quality leather using an in vitro culture step of fibroblasts on a honeycomb macroporous polymer scaffold to improve cell growth and collagen secretion, providing leather with hardness, mechanical strength and wear resistance. In particular, the inventors prepared a honeycomb macroporous polymer scaffold with uniformly distributed pores on the surface of the scaffold, forming a well-organized structure with uniform pore spacing.

[0032] Interestingly, compared with the sponge macroporous scaffold, the scaffold can maintain cell viability and simultaneously have high secretion of extracellular matrix. The high secretion of extracellular matrix by cell culture on the scaffold can enable the production of high-quality leather.

[0033] The present disclosure relates to a method for producing leather comprising the following steps:

[0034] a) culturing fibroblasts in vitro on a honeycomb macroporous polymer scaffold to obtain tissue, wherein the surface of the scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, and a surface pore area distribution of 40%-95%, preferably 50%-95%,

[0035] 55%-95%, 60%-95%, 70%-95%, more preferably 75-95%,

[0036] b) tanning the tissue, thereby forming the leather.

[0037] According to the present disclosure, the term "scaffold" refers to a three-dimensional support that provides physical and structural support for cells and enables tissue formation.

[0038] According to the method of the present disclosure, the surface of the scaffold comprises macropores with a diameter of 100 μm-280 μm, preferably 110 μm-225 μm, into which cells can penetrate, thereby conferring a biological environment conducive to the proliferation and function of fibroblasts.

[0039] For the purpose of this disclosure, the term "pore size" refers to the average of 5-30 pore diameters measured by using ImageJ software (Rasband, WS, ImageJ, US National Institutes of Health, Bethesda, Maryland, USA, https: / / imagej.nih.gov / ij / , 1997-2018), the average of pore diameters measured with a scanning electron microscope (SEM). In a preferred embodiment, the pore size is measured on one surface of the scaffold.

[0040] In order to maintain the mechanical structure of the scaffold while providing a suitable biological environment for cell proliferation, the surface pore area distribution of the honeycomb macroporous scaffold surface according to the present invention is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, and more preferably 75%-95%.

[0041] The surface pore area distribution at the scaffold surface can be determined by analyzing representative images of the scaffold microstructure obtained by scanning electron microscopy (SEM). The surface pore area distribution refers to the ratio of the pore surface area to the total scaffold surface area. The surface porosity (surface pore area distribution) of the manufactured membrane is calculated by the following equation:

[0042]

[0043] Among them: A t is the total area of ​​the image, A p is the area porosity at distance z, and h is the height of the image. The surface pore area distribution corresponds to the percentage of pores identified by the electron microscope image at a selected magnification. The image can be analyzed using appropriate software that can measure the size of the pore portion visible on the surface of the support and the surface pore area distribution. The surface area porosity can be determined as described in Example D.2 of the present application.

[0044] According to a specific embodiment, the total porosity of the macroporous scaffold is preferably 65%-98%, in particular 70%-98%, 75%-98%, and more preferably 80%-95%.

[0045] "Porosity", "void fraction" or "total porosity" is different from the surface pore area distribution as described above. In fact, for the purpose of the present invention, the term "porosity" or "void fraction" or "total porosity" refers to the percentage of pore volume relative to the volume of the material, as measured according to the weight method (Guarino V, et al. 2008 Sep; 29 (27): 3662-70). The volume of the sample is calculated from the measured value of the sample size. Measuring the mass of the sample can deduce the sample density. For each type of sample, the density (ρSc) is the average value obtained from 4 samples. Then, the porosity of the scaffold is calculated using the following formula:

[0046] Porosity = (1-(ρSc / ρPoly)): 100

[0047] where ρPoly is the density of the polymer according to the manufacturer's database.

[0048] The scaffold according to the present disclosure is a honeycomb macroporous scaffold (also referred to herein as a membrane scaffold), which is a well-organized structure that is uniform in pore pitch, ie, the walls separating the pores have the same thickness.

[0049] In a preferred embodiment, on the surface of the honeycomb macroporous scaffold according to the present disclosure, the pore distance corresponding to the pore wall thickness is less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm.

[0050] As used herein, the hole wall thickness refers to the distance between the two opposite walls of the hole. The hole wall thickness can be measured by any method known in the art. In a specific embodiment, the hole wall thickness can be measured from an SEM image, for example, by calculating the distance between the holes, as described in Haeri, Morteza & Haeri, Mohammad. (2015); Journal of Open Research Software; 3.10.5334 / jors.bn and Example D.2 of the present application. The hole distance corresponds to the average surface area between the holes, which can be calculated by calculating the distance between the centers of two adjacent holes minus the sum of the radii of each hole. In another embodiment, the hole wall thickness can also be calculated by modeling the geometry of the stent surface as described in Example D.2 of the present application. The method includes modeling the geometry of the surface and transposing the hole surface area to a circle, the average radius of the circle being determined by the average value of the distance between the pixel points on the line and the center of the area.

[0051] The honeycomb macroporous support according to the present disclosure can be designed to have a thickness suitable for leather synthesis. In particular, the thickness of the support can be 0.1-2.5 mm, in particular 0.3-2 mm.

[0052] The thickness of the sample can be measured according to ISO 2589:2016 standard method. The thickness is measured using a digital thickness gauge J-40-L manufactured by Checkline that complies with ISO 2589:2016. 30 measurements are taken on the sample and the average is taken.

[0053] In specific embodiments, the tensile strength of the stent according to the present disclosure is 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, and more preferably 0.2-0.7 MPa.

[0054] In another specific embodiment, the Young's modulus of the stent according to the present disclosure is 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.

[0055] According to ISO 3376:2020 standard, physical and mechanical tests are performed on the bracket, i.e., tensile strength and Young's modulus are determined. Environmental conditions are ambient temperature and humidity. An Instron 34SC-1 system is used to perform the test. It is equipped with an Instron 50N force sensor (CAT.NO.: 2530-50N) and an Instron manual fastening clamp (CAT.NO.: 2710-203). The speed is applied during the test and set to 100 mm / min. Strength and displacement measurements are recorded. The stress applied during the test is calculated by the following equation:

[0056]

[0057] Where: F is the traction force applied by the system on the surface of the tensile specimen; S TS is the minimum surface area of ​​the tensile specimen, W and T TS are the width and thickness of the tensile specimen.

[0058] The elongation is calculated using the initial length and displacement as well as the displacement on the pulling axis.

[0059] Young's modulus is calculated as described in Example D.3. Fig.12 ) with the above parameters using the following equation:

[0060]

[0061] E Mem is the Young's modulus of the membrane scaffold under test, σ is the measured tensile stress, and ε is the axial strain in the linear elastic region of the membrane scaffold.

[0062] The tensile strength is the maximum stress that a membrane support can withstand when stretched or pulled before breaking, and can be measured as described in Example D.3. of the present application ( Fig.12 ).

[0063] According to a particular embodiment, the polymer support does not have a fibrous structure.

[0064] A polymer according to the present disclosure that can be used in a stent is a polyester derived poly(alpha-hydroxy acid).

[0065] Non-limiting examples of polyester-derived poly(α-hydroxy acids), particularly bioderived and / or biodegradable polyesters, that can be used to form the scaffold are homopolymers and copolymers of hydroxy acids, such as polylactic acid (PLA), polyglycolic acid (PGA), poly(lactic acid and glycolic acid) (PLGA), poly(3-hydroxybutyrate), poly(4-hydroxybutyrate), poly(3-hydroxyvalerate), poly(5-hydroxyvalerate), poly(3-hydroxypropionate), poly(3 -hydroxycaproate), poly(3-hydroxyoctanoate), poly(3-hydroxyoctadecanoate); polycaprolactone; homopolymers and copolymers of poly(butylene succinate) and poly(butylene adipate), such as poly(butylene succinate-co-butylene adipate) (PBSA), poly(butylene succinate) (PBS); and polyhydroxyalkanoates (PHA) and their derivatives, polyhydroxy esters of 3-, 4-, 5- and 6-hydroxyalkanoic acids and mixtures thereof.

[0066] In a preferred embodiment, the scaffold comprises 1-50% (w / w), preferably 5-20% (w / w), of a polyester polymer.

[0067] From a mechanical point of view, polyesters have interesting properties, they are compatible with cell culture. They can be of biological origin, as they can be produced from agricultural waste and / or can be fully biodegradable. However, polyesters are hydrophobic and impermeable to oxygen and do not allow molecular diffusion after reticulation.

[0068] Therefore, to facilitate gas exchange, the oxygen-impermeable polymer stent may further comprise nanopores and micropores having a diameter less than 20 μm, particularly 0.1 μm to 10 μm, and more particularly 2 to 8 μm.

[0069] The stent composed of an oxygen-impermeable polymer (such as a polyester polymer) and comprising micropores may be an asymmetric stent or a symmetric stent.

[0070] Honeycomb macroporous polymer scaffolds, also referred to herein as microporous / macroporous membrane scaffolds according to the present disclosure, can be achieved by phase inversion (e.g., non-solvent induced phase separation (NIPS)), in which the exchange between solvent and non-solvent controls the formation of pores. In the presence of a macroporous porogen, the escape of solvent and the entry of non-solvent will generate micropores and macropores in the polymer between the network chain polymers to allow the creation of dual porosity in the scaffold according to the present disclosure.

[0071] Phase inversion is a method well known in the art and includes, for example, non-solvent induced phase separation (NIPS), thermal induced separation (TIPS), vapor induced phase separation (VIPS) and polymerization induced phase separation (PIPS).

[0072] TIPS is a method in which a polymer solution is formed at high temperature with a high boiling point solvent and cooled to induce phase separation and polymer solidification. A microporous membrane is obtained after the diluent is extracted.

[0073] In the VIPS process, a cast film consisting of a polymer and a solvent is exposed to a vapor atmosphere of a non-solvent molecule (typically water). As the vapor penetrates into the film, the polymer precipitates, which ultimately forms a symmetrical porous membrane without a dense surface layer. The thermodynamic properties of the casting solutions in the NIPS and VIPS processes are almost similar, indicating that the VIPS process should produce membranes with similar morphology to those produced using the NIPS process.

[0074] The PIPS process is a phase separation that occurs in a multi-component mixture induced by the polymerization of one or more components. The increase in molecular weight of the reactive components makes one or more components immiscible with each other, resulting in spontaneous phase separation. Due to the randomness of the phase separation initiation and process, the morphology of the final phase separation structure is usually random.

[0075] In particular, the scaffold can be prepared by the following steps: preparing a solution of the polymer, which contains 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 containing the polymer and the porogen on a solid support; and introducing the polymer solution into a non-solvent solution (e.g., ethanol and water), and finally immersing the porogen / polymer composite into a porogenic solvent to dissolve the porogen ( Figure 1 ).

[0076] According to the above method, the solid substrate on which the polymer solution is deposited can be made in particular of glass, metal or solvent-resistant plastic, such as polytetrafluoroethylene. Nylon 6,6 or polyethylene terephthalate.

[0077] Examples of suitable porogens include inorganic salt crystals, such as sodium chloride crystals or potassium chloride, and amorphous substances such as poly(ethylene glycol) (PEG), polyvinyl pyrrolidone, sucrose crystals, gelatin spheres, and paraffin spheres.

[0078] In particular, the concentration of the porogen in the polymer solution may be specifically 10 wt% to 98 wt%, more specifically 30 wt% to 95 wt%, relative to the weight of the polymer.

[0079] The solvent for the polymer can be easily selected by a person skilled in the art, and non-limiting examples can be selected from: N,N-dimethylformamide, N-methylpyrrolidone, ethyl lactate, ethyl acetate, dimethylacetamide, dimethyl sulfoxide, chloroform and mixtures thereof, and the non-solvent can be especially water.

[0080] Due to the different condensation kinetics on the surface in contact with the non-solvent and on the surface in contact with the substrate, the formation of pores of different sizes is observed. The direct contact of the polymer solution with the non-solvent will directly lead to a smooth nanoporous surface, while the slow diffusion of the non-solvent between the substrate and the collodion will lead to a slow separation of the solvent, producing a polymer-rich phase and a polymer-poor phase. After condensation, the polymer-rich phase forms a solid mass of the scaffold, and the polymer-poor phase forms an interconnected network of pores of different sizes, which open on the surface of the scaffold and extend deep into the scaffold. The honeycomb macroporous polymer scaffold obtained by the NIPS process is an asymmetric scaffold with an interconnected network of macropores and micropores. The honeycomb macroporous support has a smooth surface and a porous surface, wherein the porous surface contains macropores opening on the surface, whose diameter is 100μm-280μm, especially 110μm-225μm, and whose surface pore area distribution is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75%-95%, and the pore wall thickness is less than 70μm, preferably less than 60μm, more preferably 2μm-50μm, wherein the pores extend from the porous surface depth to the smooth surface.

[0081] Asymmetric honeycomb macroporous polymer scaffold means that there are no macropores with a diameter of 100 μm-280 μm, especially 110 μm-225 μm, throughout the thickness of the scaffold. In particular, the asymmetric scaffold has a first face (porous face) and a second opposite face (smooth face), the first face comprising macropores with a diameter of 100 μm-280 μm, especially 110 μm-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 preferably a pore wall thickness of less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm, and the second opposite face (smooth face) only comprises nanopores with a diameter of less than 10 nm, more precisely 5 nm.

[0082] In another specific embodiment, the cellular macroporous polymer scaffold is a symmetrical scaffold.

[0083] A symmetrical scaffold means a scaffold with macropores uniformly distributed throughout the thickness of the scaffold, wherein the macropore diameter is 100-280 μm, especially 110-225 μm, the surface pore area distribution is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75-95%, and the pore wall thickness is preferably less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm, and in particular wherein the scaffold has two macroporous faces, in particular wherein both faces contain macropores with a diameter of especially 100-280 μm, especially 110-225 μm, and the surface pore area distribution is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75-95%, and the pore wall thickness is less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm.

[0084] Symmetrical honeycomb macroporous scaffolds may be achieved by any method known to those skilled in the art. Non-limiting examples may be: a) physical abrasion of the smooth surface, and / or b) chemical dissolution of the smooth surface using a polyester polymer solvent ( Figure 2 ).

[0085] In particular, the polyester symmetric membrane scaffold can be produced by the steps of fabrication of the polymer asymmetric scaffold as described above, physical abrasion and / or chemical dissolution of the smooth surface using a polyester polymer solvent, and finally immersing the scaffold in water to remove the physical / chemical agents.

[0086] In order to functionalize the surface of the scaffold designed as described above and increase the cell compatibility 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, keratin, hyaluronic acid, chondroitin sulfate, chitosan and heparin; proteins such as fibrinogen and collagen, especially soluble collagen; peptides such as peptides known for their cell adhesion ability, for example, peptides containing arginine-glycine-aspartic acid (RGD) or arginine-glycine-aspartic acid-serine (RGDS) sequences; and mixtures thereof. The bioactive molecules can be grafted by any method known in the art.

[0087] According to a specific embodiment, the bioactive molecule is collagen. In fact, collagen allows cell adhesion because collagen specifically interacts with integrin receptors expressed on the surface of skin-derived cells. In order to graft collagen to the surface of the scaffold, in a specific embodiment, the scaffold can be treated with a plasma generated in air or oxygen to add electronegative silanol groups. Then, the collagen is grafted on the surface of the scaffold by immersing the scaffold in a solution containing collagen and then rinsing, preferably with a phosphate buffered saline solution (PBS).

[0088] According to a preferred embodiment, the bioactive molecule can be a carbohydrate and / or carbohydrate-derived molecule, such as glycosaminoglycans (e.g., heparin sulfate, heparin, chondroitin sulfate, dermatan sulfate, keratan sulfate, hyaluronic acid). Due to its unique biophysical and chemical properties (viscoelasticity, high water retention capacity, ability to interact specifically or nonspecifically with various proteins), glycosaminoglycans play an important role in extracellular matrix organization and homeostasis.

[0089] In order to graft sugars and / or sugar-derived molecules (e.g., glycosaminoglycans) on the surface of the scaffold, the scaffold can first be pretreated to bring positive charges, preferably positive amine groups, to the surface of the scaffold, and then functionalized with negatively charged molecules. The positive amine groups can be added by treating the scaffold with a plasma generated in oxygen or air and immersing the scaffold in a polylysine solution, by treating the scaffold with a plasma generated in azo, or by an aminolysis reaction. When the surface of the scaffold has been treated to be positively charged, the scaffold can be functionalized with sugars and / or sugar-derived molecules by immersing the scaffold in a solution containing the sugars and / or sugar-derived molecules, followed by rinsing, for example, with ultrapure water.

[0090] According to a preferred embodiment, the positive amine groups are added by aminolysis of the polymer using a solution of at least one aliphatic α,ω-diamine. The aminolysis reaction can be carried out in particular by immersing the polymer stent in a solution comprising an aliphatic α,ω-diamine, preferably by immersing the stent in a 1,6-hexanediamine solution in propanol or water and mixtures thereof, followed by rinsing, for example, with ultrapure water.

[0091] According to the methods of the present disclosure, fibroblasts are seeded onto a polymer scaffold as described above and cultured under appropriate growth conditions, particularly in a culture medium that can proliferate and induce secretion of extracellular matrix, such as different types of collagen and elastin fibers.

[0092] Fibroblasts according to the present disclosure are cells that synthesize extracellular matrix (such as collagen, elastin, glycoprotein and non-proteoglycan and other extracellular matrix components). Fibroblasts can be mammalian or non-mammalian fibroblasts. Cells can be a combination of single cell types or cell types. Since the only components that resist storage and tanning processes are collagen and elastin fibers secreted by fibroblasts, in a preferred embodiment, fibroblasts are the only cells cultivated on a support according to the method disclosed herein.

[0093] Many culture media are commercially available and are well known to those skilled in the art. The culture media may be a basic culture medium that contains, in particular, inorganic salts, amino acids, vitamins and carbon sources necessary for the cells and a buffer system for adjusting the pH. The basic culture media that can be used in the method of the present invention include, for example but not limited to, MEM culture media, DMEM / F12 culture media, DMEM culture media RPMI culture media, Ham's F12 culture media, IMDM culture media and KnockOut TM 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 antifungal agents.

[0094] On the other hand, the present disclosure also relates to a tissue (also referred to as in vitro skin) obtainable by the cell culture step a) as described above, preferably, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer scaffold as described above, the scaffold 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.

[0095] In a particular embodiment, the scaffold comprises a polyester, preferably a biodegradable and / or partially or fully bioderived polyester, preferably in a concentration of 1-50% (w / w), preferably 5-20% (w / w).

[0096] In a preferred embodiment, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer having macropores interconnected with micropores, preferably having a diameter of less than 20 μm, preferably less than 10 μm.

[0097] 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.

[0098] In another preferred embodiment, the tissue comprises fibroblasts cultured on a honeycomb macroporous polymer, and 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, and more preferably 30-50 kPa.

[0099] In another embodiment, the tissue comprises fibroblasts cultured on an asymmetric scaffold or a symmetric scaffold as described above.

[0100] In another preferred embodiment, the tissue comprises fibroblasts cultured on the scaffold obtained by non-solvent induced phase separation method (NIPS).

[0101] In a preferred embodiment, the tissue is obtained after at least one week in culture, preferably two, three or four weeks.

[0102] As described above, culturing fibroblasts on scaffolds can result in tissues comprising a densely organized network of cells and extracellular matrix, preferably with cell / extracellular matrix grooves of 200-400 μm as observed under photon and / or scanning electron microscopy.

[0103] The tissue (in vitro skin) according to the present disclosure has at least one of the following properties:

[0104] - said tissue comprises at least 50 μg / mg, preferably at least 60, 70, 80, 90 or 100 μg / mg, more preferably 50-300 μg / mg, even more preferably 60-250 μg / mg of collagen per mg of dry tissue. The amount of collagen is preferably measured by the hydroxyproline assay as defined in Example E.1.1 of the present application.

[0105] - said tissue comprises an elastin content percentage of at least 0.3% (w / w), preferably at least 0.4% (w / w), based on the total dry mass of the tissue. The elastin content is preferably measured by the elastin assay (elastin content) as defined in Example E.2.1, in particular by determining the elastin concentration in the supernatant of oxalic acid digested dry tissue.

[0106] - said tissue comprises less than 1% (w / w), preferably less than 0.1 or 0.01% (w / w) of total fatty acids in the total dry mass of the tissue, preferably measured by gas chromatography-mass spectrometry as defined in Example E.6.1 of the present application,

[0107] - the tissue contains less than 400ng / mg, preferably 300ng / mg, more preferably 200ng / mg of total glycosaminoglycans (GAGs) per unit of total tissue wet weight, corresponding to the sum of sulfated GAGs and hyaluronic acid (HA), preferably the tissue contains less than 200ng / mg HA per unit of total tissue wet weight, preferably less than 150, 100ng / mg HA per unit of total tissue wet weight and / or less than 300ng / mg of sulfated GAGs per unit of total tissue wet weight, preferably 200ng ECS / mg of sulfated GAGs per unit of total tissue wet weight. In a preferred embodiment, sulfated GAGs are determined by staining with dimethylene blue as described in Example E 7.1, and / or hyaluronic acid is measured by adding acetate-saturated ethanol and cetylpyridine as described in Example E 7.1.

[0108] - The tissue is an isotropic material on the surface, preferably analyzed by 2D Fast Fourier Transform (FFT) of SEM observation, as described in Example E.3 Orientation analysis of biomaterial fibers in in vitro skin. In particular, the tissue does not show representative peaks, percentage changes and Fig.13 The same as shown in C.

[0109] - said tissue contains less than 0.3, preferably less than 0.2 ppm of extractable and total heavy metals, preferably measured as determined in Example E.4.1.

[0110] - the tissue contains less than 2 mg / g, preferably less than 1 mg / g, free amines per unit mass of dry tissue, preferably measured according to the quantification of free amino groups in Example E.5.1,

[0111] - said tissue has at least two denaturation temperatures (Td), preferably three Td, determined by differential scanning calorimetry (DSC). Said denaturation temperatures are preferably determined by wet differential scanning calorimetry (DSC) as defined in Example F1.1, more preferably measured in the range of 5-100°C at a constant speed of 5°C / min.

[0112] The tissue obtained by the above cell culture step is then tanned by any method known in the art to create chemical bonds between the elastin fibers and collagen, thereby converting it into leather.

[0113] Various tanning processes can be used to tan leather, including chrome and chromium (III) sulfate tanning agents, tanning using aluminum salts, aldehydes, organic compounds, vegetable tanning using tannic acid, and tanning using low molecular weight phenolic and acrylic polymers (hereinafter referred to as "polymer tanning").

[0114] In another aspect, the present disclosure relates to tanned leather obtainable by the method as described above. As used herein, the term "leather" or "synthetic leather" refers to a material obtained by tanning or chemically treating animal skins or tissues, which contain collagen, elastin and other components of the extracellular matrix. According to the present disclosure, the tissue is obtained by an in vitro culture step of fibroblasts and contains a dense network of cells and extracellular matrix components.

[0115] The tanned leather (in vitro leather) obtainable by the above process has at least one of the following properties:

[0116] - the leather has at least two degeneration temperatures (Td), preferably three Td, determined by differential scanning calorimetry (DSC). The degeneration temperatures are preferably determined by wet differential scanning calorimetry (DSC) as defined in Example F.1.1, more preferably measured in the range of 5-100°C at a constant speed of 5°C / min.

[0117] - the leather has a total porosity of 30% to 60%, preferably measured by mercury intrusion porosimetry as described in Example F.2.1 Mercury intrusion porosimetry;

[0118] - The water vapor permeability of the leather is 0.5-5.5 mg / cm 2 .h, preferably measured at 23° C. and 50% relative humidity, as described in Example F.3.1 Water vapour permeability.

[0119] The present disclosure also relates to the use of a macroporous scaffold as described above for producing leather, preferably by tanning a tissue obtained from a fibroblast culture on the honeycomb macroporous polymer scaffold, more preferably, wherein the surface of the scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, and the surface pore area distribution on the surface is 40%-95%, preferably 50%-95%, 55%-95%, 60%-95%, 70%-95%, more preferably 75-95%. In a preferred embodiment, the pore wall thickness of the surface of the scaffold is less than 70 μm, preferably less than 60 μm, more preferably 2-50 μm. In a specific embodiment, the scaffold is a polyester scaffold, preferably a honeycomb polyester macroporous scaffold. In another specific embodiment, the scaffold is an asymmetric or symmetric scaffold.

[0120] Embodiments of the present invention are described in the following specific examples, which are intended to be illustrative only and are not to be construed as limiting.

[0121] Example

[0122] A. Asymmetric honeycomb-shaped polyester macroporous scaffolds obtained using NIPS

[0123] 1 - Fabrication of macroporous honeycomb polyester scaffolds (membrane scaffolds)

[0124] Asymmetric polyester membranes were prepared according to the NIPS process. 聚合物 / v 溶剂 ) is obtained by dissolving or selecting a polyester (with or without a selected porogen) in a selected solvent at different mass ratios, heating at 70°C for 2 hours to ensure complete dissolution. After the bubbles are eliminated, the solution is poured onto a glass plate and spread to form a film. The thickness of the cast film is limited by 7 paper wedges on the two long sides of the glass plate. Then, the glass plate carrying the polymer film is immediately immersed in a Milli-Q water bath at room temperature (i.e., 20-22°C). After condensation, the formed film is thoroughly washed with Milli-Q water and stored in a desiccator under vacuum at room temperature ( Figure 1 ).

[0125] 2-Morphological characteristics of polyester film scaffolds

[0126] Polyester films were prepared using the NIPS process using polymer solutions with various polymer contents. For most films, the two faces appeared different in appearance to the naked eye, indicating the presence of an asymmetric structure. Observation by optical microscopy and SEM allowed a more detailed examination of the morphology of the two faces ( Figure 3 ).

[0127] These observations confirm the asymmetric structure of the membrane, which has a smooth top surface ( Figure 3 d) and the bottom of the macropore ( Figure 3 a, b), the macroporous bottom surface has pores of several nanometers extending deep from the porous surface to the smooth surface ( Figure 3 c).

[0128] As expected, using the NIPS process, when a film of a polymer solution is immersed in a water bath as a non-solvent for the selected polyester polymer, coagulation of the polyester polymer occurs. During this process, the polymer solvent miscible with the non-solvent is gradually removed, thereby causing the gelation of the polyester polymer, followed by its vitrification and crystallization. More precisely, the top side of the film is in direct contact with water, while the bottom side is in contact with the substrate. Thus, the coagulation phenomenon occurs faster on the top side than on the bottom side. On the top side, the rapid detachment of the solvent leads to the formation of a polymer concentrated gel, which subsequently produces a smooth surface after the film solidifies. The progressive diffusion of the non-solvent through the polymer gel leads to phase separation into a continuous polymer-rich phase and a dispersed polymer-poor phase, which becomes increasingly apparent from the top side to the bottom side of the polymer film. Over time, this eventually leads to the solidification of the polyester-rich phase, and pores filled with the non-solvent are generated by the coalescence of droplets of the polymer-poor phase. Thus, the NIPS process can produce asymmetric membranes containing macropores in the form of glove fingers, opening on one face and extending deep into the membrane to form a smooth face without macro / micropores on the other side ( Figure 3 c) (see al tawil et al., European polymer journal. 105, 370-88, 2018).

[0129] 3. The role of adding macroporous porogen

[0130] The inventors are interested in asymmetric membranes with surface macropores that are large enough in diameter (about 140 μm) and morphologically suitable for accommodating fibroblasts and their ECM. Therefore, the inventors added porogens to better control the pore size and interconnection between macropores. When the selected macroporogen is added, the membrane shows open macropores on the surface, and the number and diameter of macropores and micropores are increased.

[0131] Interestingly, through SEM ( Figure 3 ) showed that, in addition to the typical increase in porosity attributable to the extraction of the porogen during the membrane coagulation step, the addition of the macroporous porogen led to the formation of a dual pore structure with an interconnected network of macropores and micropores. Under these conditions, a well-organized honeycomb macroporous structure ( Figure 3 a) and a small amount of micropores in the macropore wall ( Figure 3 b).

[0132] 4. Surface modification of polyester film

[0133] The polyester film was functionalized with the carbohydrate-derived molecules after the aminolysis reaction. The surface modification of the polyester film surface was successful, as it was stained with red carmine after aminolysis and with bright green after the carbohydrate-derived molecules were immobilized. All coloration was uniform on all surfaces of the sample. In addition, the untreated scaffolds did not retain coloration, which means that surface modification is a key step to fix HA and accommodate cells.

[0134] B. Cell behavior and tissue generation in asymmetric honeycomb polyester macroporous scaffolds.

[0135] 1. Cell Behavior on Polyester Sponges and Membranes

[0136] Cell behavior is stimulated by biochemical (soluble factors), physical cell-cell or cell-ECM interactions and matrix (scaffold) mechanical stimulation. As a result, cells can adhere, proliferate, differentiate, dormant or transform. Here, the inventors emphasize the influence of scaffold stiffness combined with morphological-structural parameters on cell behavior.

[0137] 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, as well as polyester membranes obtained by NIPS with selected macropores. These scaffolds were functionalized with carbohydrate-derived molecules, seeded with fibroblasts, and maintained in culture for up to 5 weeks.

[0138] From the first hour of inoculation, cells adhered to the PDMS and polyester sponge surfaces and began to proliferate. SEM images of polyester sponge ( Figure 4 ) showed that the entire sponge surface was covered with a cell layer: cell bodies (round cells) were detected on the entire surface of the sponge and could be detected at different culture times (3 weeks and 5 weeks). The observed cell density was surprising compared to the expected results. Since the cells are biologically active (vitality and proliferation), the ECM deposition rate is low (Dekker et al., Tissue Engineering of Cartilage and Bone: Novartis Foundation Symposium 24. Volume 249. 2003). This result prompted us to hypothesize that there may be a crosstalk between the morphology and mechanical properties of the scaffold and the observed cell behavior.

[0139] Fibroblasts exhibit a different behavior on polyester membranes obtained with the NIPS process. As in the case of sponges, cells begin to adhere from the first hour after inoculation, they proliferate and colonize the entire membrane surface. Very interestingly, the SEM images ( Figure 5 ) showed that compared with polyester sponge, it had fewer cell bodies and a large number of extracellular fibers distributed in the macropores and micropores.

[0140] From a morphological point of view, the PDMS and polyester sponge scaffolds exhibited a non-uniform structure with low surface porosity and high wall thickness between the macropores. For the polyester membrane, the uniform but irregular distribution of honeycomb pores produced a well-organized structure with high surface porosity and low wall thickness between the macropores ( Fig. 9 ). Blocks of material (polyester) exist between the voids in the sponge at irregular thicknesses, changing the mechanical properties, while the thin pore walls in the membrane create a less rigid material with varying degrees of elasticity.

[0141] Since the scaffold stiffness affects the cytoskeleton organization, it also affects the cell morphology and thus the behavior. Various studies have shown that a harder matrix usually induces a rounder cell shape, which is tightly distributed, while a soft matrix promotes cell spreading (Humphrey JD, Dufresne ER, Schwartz MA. Nat Rev Mol Cell Biol. 2014 Dec; 15 (12): 802-12). These changes in cell morphology are accompanied by changes in cell behavior (including vitality, proliferation and / or ECM deposition), which is not surprising. The inventors believe that the presence of thick walls between the sponge pores makes it harder, thereby inducing cell aggregation and colonization on the scaffold in a very narrow manner, close to each other.

[0142] To quantitatively assess cell proliferation in polyester sponges or membranes, the inventors used the cell proliferation reagent WST-1 assay to track cell proliferation, which is a non-radioactive, spectrophotometric quantification of cell proliferation and cell population viability (Ref. 5015944001, Sigma). Figure 6 As shown, fibroblasts on polyester membrane ( Figure 6 The proliferation curve of the 24-hectare WT ( , square dotted line) has a gentle slope, indicating an initial stage of cell adaptation (<10 days), followed by a cell proliferation stage (10-15 days) and a final stage of stable cell growth (>15 days), which the inventors speculate is related to Figure 5 On the other hand, the proliferation curve of fibroblasts on the sponge ( Figure 6 , circles and solid lines) had an upward trend, indicating that cells continued to proliferate even after 4 weeks of culture. The maintenance of the proliferation rate after 3 weeks of culture ( Figure 6 , circles and solid lines) reflect a higher rate of cell proliferation and are associated with lower secretion and deposition of ECM on the surface of immobilized polyester sponges in culture and as observed by scanning electron microscopy ( Figure 4 On the other hand, after culturing fibroblasts on polyester honeycomb membrane scaffolds for two weeks, the stability of cell proliferation was consistent with the small number of cell bodies accompanied by high secretion of ECM observed by scanning electron microscopy ( Figure 5 ). This antagonism between cell proliferation and ECM secretion can be well described by the antagonism of cell proliferation on cell specialization (Cooper GM. The Cell: A Molecular Approach. 2nd edition. Sunderland (MA): Sinauer Associates; 2000).

[0143] 2. Tissue Formation on Asymmetric Honeycomb Polyester Macroporous Scaffolds

[0144] Micropatterned surfaces, arrangement of ECM fibers, cells and interconnected 3D pore structures are important microstructural features. It has been shown that micropatterned structures guide multicellular tissues and fibrous collagen deposition, and also affect the arrangement and shape of single cells (Gilchrist, CL, Ruch, DS, Little, D. & Guilak, F. Biomaterials 35, 10015–10024 (2014)). In addition, the arranged clues are sufficient to guide cell shape, arrangement, adhesion and fibrous collagen matrix deposition. In this case, the structure of polyester sponge is defined as micropatterned structure (pores follow the geometry of porogens), while the pores of polyester film have an ordered honeycomb structure on the surface and an arranged glove-like structure at depth. Orderly arranged cells are more likely to differentiate, which means that cells are active and secrete more ECM (Du, Y., et al. Biomaterials 218, 119334 (2019)).

[0145] When proliferating in polyester membranes, the relationship between structures (macro and micro) modulates cell behavior and drives cells to secrete ECM, e.g. Figure 5 The deposition of collagen fibers in the pores and the attachment of collagen fibers to the pore walls will certainly improve the mechanical properties of the structure.

[0146] To characterize cell proliferation, cell distribution, ECM expression, and ECM fiber distribution in cultured asymmetric honeycomb polyester membrane scaffold constructs, polyester membranes were seeded with fibroblasts and cultured for 1 and 3 weeks, fixed with 3.7% PFA in PBS 1X, and stained with an anti-collagen 1 specific antibody (Sigma, ref. C2456) combined with a cell-nucleus staining reagent (DAPI), as described above. Figure 7 shown.

[0147] like Figure 7 As shown in a, membrane constructs cultured for 1 week showed proliferation of cell bodies into the membrane macroporous structure, low secretion of collagen-1 and absence of collagen-1 fibers. This result is consistent with the observation of cell proliferation during the first two weeks of culture in the membrane scaffold, as shown in Figure 6 As shown (circle, solid line).

[0148] In contrast, membrane constructs cultured for 3 weeks showed organization of a dense network of cells and collagen-1 ( Figure 7b), especially with 200-400 μm cell / collagen-1 grooves, reminiscent of reticular ridges, which protrude from the papillary dermis at the interface of animal dermis and epidermis (Q. Zeng, LK Macri, A. Prasad, RA F Clark, DI Zeugolis, C. Hanley, Y. Garcia, A. Pandit, DI Leavesley, D. Stupar, ML 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 and enter the ECM secreting cell specialization stage, such as Figure 6 As shown (square, dotted line).

[0149] C. Tanning and leather production of cultured asymmetric honeycomb polyester macroporous scaffolds.

[0150] The construct obtained after culturing fibroblasts on the asymmetric honeycomb polyester macroporous scaffold as described above was tanned and retanned using methods known in the art to create chemical bonds between the fibers of elastin and collagen, thereby converting it into leather. The cultured polyester sponges and membranes were tanned and retanned in an experimental tanning drum according to the protocol described for traditional animal leather ( Figure 8 a).

[0151] Unfortunately, polyester sponges cultured with fibroblasts for 4-5 weeks became brittle and broke apart during the tanning and retanning steps. In contrast, polyester film constructs inoculated for 4 or 5 weeks maintained their structural integrity after the tanning and retanning steps and became leather-like processed products, such as Figure 8 c. In fact, Figure 8 As shown in a, the asymmetric honeycomb polyester macroporous scaffolds cultured for 4 weeks, tanned and retanned, showed the leather grain texture pattern characteristics of full-grain leather when observed under a low-magnification optical microscope ( Figure 8 b).

[0152] D. Geometric and Mechanical Comparisons between Polyester Sponge and Membrane

[0153] 1. Geometry

[0154] From a morphological point of view, polyester sponge exhibits non-uniformly dispersed porosity ( Fig. 9, above), with low surface porosity and high wall thickness between macropores. On the other hand, the polyester membrane showed a uniform but irregular honeycomb macropore distribution, resulting in a well-organized structure with high surface porosity and low wall thickness between macropores ( Fig. 9 ,Down).

[0155] 2. Surface morphology analysis of porous scaffolds: sponges and membranes

[0156] Scanning electron microscope (SEM) images of specific magnifications were used to measure the surface porosity, pore size, and surface pore area distribution of the membrane. The porous surface of the scaffold was imaged by SEM, and the segmentation of the image was standardized with the corrected magnification using ImageJ software (Rasband, WS, ImageJ, US National Institutes of Health, Bethesda, Maryland, USA, https: / / imagej.nih.gov / ij / , 1997-2018). The original image (gray / black image) of the outer surface of the porous scaffold was binarized to 0-255 pixels, where low-brightness and high-brightness pixels were assumed to be the pore area and the boundary of the containing material, respectively. The high-frequency and low-frequency noise generated by the pore binarization from the background image was eliminated using an appropriate function ( Fig. 9 ).

[0157] In the binary images, each macropore on the surface area of ​​the hollow sponge or membrane was digitized using the "Analyze Particles" function of ImageJ to automatically measure the surface area, circularity, and Feret pore size. Through these measurements, statistically representative surface porosity, surface pore area distribution, and Feret pore size of sponges and membrane scaffolds were obtained.

[0158] The surface porosity of the fabricated membranes was calculated by the following equation:

[0159]

[0160] Where: At is the total area of ​​the image, Ap is the area porosity at distance z, and h is the height of the image.

[0161] The surface pore area distribution (number of pores / area) is defined by the distribution of the number of marked pores on a selected surface. The Feret pore diameter is the longest distance between any two points along the pore boundary, also known as the maximum card diameter or maximum diameter.

[0162] As described in Haeri, Morteza & Haeri, Mohammad. (2015). Journal of Open Research Software. 3.10.5334 / jors.bn, other analyses of sponge and membrane scaffold structures were performed using the ND plugin in ImageJ. Journal of Open Research Software. 3.10.5334 / jors.bn. ND is used to calculate the pore distance and determine the pore-pore wall thickness. The pore distance, i.e., the average surface area between the pores is obtained by calculating the distance between the centers of 3 adjacent pores minus the sum of the radius of each pore. The center of mass of the pore is digitally determined by determining the center of the geometric pore boundary corresponding to the pore. The calculation of the wall distance between the pores is obtained by the distance of the pore center of mass corrected by the pore area (Haeri, Morteza & Haeri, Mohammad. (2015). Journal of Open Research Software. 3.10.5334 / jors.bn).

[0163] like Fig.10 As shown in Figure 2, the surface porosity of the membrane scaffolds was 3 times higher than that observed for the sponge scaffolds, with the median ± MAD of the surface porosity of the membrane scaffolds being 66.11 ± 8.53 and that of the sponge scaffolds being 21.49 ± 8.65. Fig.11 As shown, the pore wall thickness of the membrane scaffolds was 2.4 times lower than that observed in the sponge scaffolds, with the mean wall thickness median ± MAD of 29.37 ± 12.40 for the membrane scaffolds compared to 70.15 ± 26.60 for the sponge scaffolds.

[0164] In addition to low interconnected porosity, the presence of irregular thickness of material blocks between the voids in the sponge changes the mechanical properties and cell-to-cell communication, respectively. For example, high interconnected porosity in the membrane enables cell communication and arranges the deposition of ECM fibers. In addition, thin pore walls in the membrane will result in a lower stiffness material with different mechanical properties. Since the scaffold stiffness affects the cytoskeletal organization, it also affects the cell morphology and therefore the behavior. Various studies (Yeung T, et al. Cell Motil Cytoskeleton 2005; 60(1): 24-34; Gaudet C, et al. Biophys J 2003; 85(5): 3329-35) have shown that stiffer scaffolds with high porosity generally induce rounder cell shapes with tight distribution, while soft matrices promote cell spreading. It is not surprising that these changes in cell morphology are accompanied by changes in cell behavior, including ECM deposition. The presence of rigid walls between the sponge pores makes it more rigid as a material and can induce cells to aggregate and colonize the scaffold in a very narrow manner, close to each other. Under these conditions, cells will establish connections that allow them to survive while minimizing secretory activity and having a low metabolism while maintaining a high proliferation rate. In this way, the cell behavior on polyester sponges and membranes is closely related to the morphological-mechanical parameters of the scaffold (porosity, pores, pore wall thickness). This correlation directly controls the structural properties after tanning.

[0165] 3. Mechanical Analysis of Asymmetric Honeycomb Polyester Macroporous Scaffolds

[0166] Several polyester film scaffold samples were tested for mechanical properties. Different parameters such as thickness and different polymer / porogen formulations and combinations were tested. Wet tests were performed on each combination.

[0167] Each sample was cut with a press cutter to create a standard tensile specimen as specified in ISO 2419-2012. Environmental conditions were ambient temperature and humidity. The sample shape resembled a dog bone. The test section was 10 mm wide and 50 mm long. The thickness depended on the sample preparation.

[0168] The thickness of each sample was measured according to ISO 2589:2016. The thickness was measured using a digital thickness gauge J-40-L manufactured by Checkline that complies with ISO2589:2016. Three measurements were taken for each sample and the average value was taken.

[0169] The stent was physically and mechanically tested to measure tensile strength and Young's modulus according to ISO 3376:2020. The Instron 34SC-1 system was used for testing. It was equipped with an Instron 50N load cell (CAT.NO.: 2530-50N) and an Instron manual clamp (CAT.NO.: 2710-203). The speed was applied during the test and set to 100 mm / min. Strength and displacement measurements were recorded.

[0170] The stress applied during the test is calculated using the following equation:

[0171]

[0172] Where: F is the traction force applied by the system on 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.

[0173] The elongation is calculated using the initial length and displacement of the pulling axis. Representative results of mechanical tests on polyester supports according to ISO 3376:2020 are shown in Fig.12 The original stress / strain curve is obtained and displayed in Fig.12 a. The elastic limit is defined as the lowest stress point that distinguishes measurable permanent plastic deformation from elastic deformation, such as Fig.12 As shown in b.

[0174] The Young's modulus of the membrane scaffold was measured using the following equation:

[0175]

[0176] E Mem is the Young's modulus of the membrane support under test, σ is the measured tensile stress, and ε is the axial strain in the linear elastic region of the membrane support.

[0177] The tensile strength is the maximum stress that a membrane scaffold can withstand when stretched or pulled before breaking, e.g. Fig.12 As shown in c.

[0178] The data obtained from these measurements are: Young's modulus, elastic limit and tensile strength. These values ​​will be used to characterize the mechanical properties of the asymmetric honeycomb polyester macroporous scaffolds.

[0179] The average value±standard deviation of the Young's modulus of the membrane scaffold was measured to be 40.80±14.21 kPa, and the average±standard deviation of the tensile strength of the membrane scaffold was measured to be 0.4244±0.1603 MPa.

[0180] E. Characterization of tissue obtained by culturing fibroblasts on membrane scaffolds (in vitro skin).

[0181] 1. Collagen content

[0182] 1.1 Hydroxyproline determination protocol:

[0183] In vitro skin and rabbit skin (shaved rabbit skin (dermis / epidermis)) obtained from 4-week fibroblast culture on membrane macroporous scaffolds were cut with a 11 mm diameter cutting tool and oven dried (time: 16 h; T°C: 50°C). The samples were weighed in a precision balance. The samples were mechanically pulverized and 100 μL of hydrolyzate was recovered for determination of the dose of hydroxyproline present in the samples.

[0184] The concentration of hydroxyproline in the sample was determined according to the protocol provided by the manufacturer (ref. MAK-357, Sigma Aldrich). A standard curve was obtained, and the curve equation and regression coefficient R2 were obtained by linear regression of the standard curve.

[0185] The concentration of hydrolyzed hydroxyproline (μg / μL) was calculated as follows:

[0186] Hydrolyzed hydroxyproline = (B / V) x D

[0187] Wherein B: Hydroxyproline hydrolysis amount calculated from the standard curve (μg); V: Volume of sample hydrolysis solution added to the well (μL); D: Dilution multiple of the sample after hydrolysis.

[0188] Therefore, the hydrolyzed concentration can be converted to its collagen equivalent by multiplying by a factor of 7.49 (RB Neuman and MA Logan. J. Biol. Chem. 184, 299, (1950)).

[0189] 1.2 Results

[0190] To evaluate collagen secretion in ex vivo skin, samples were plated and cultured for four weeks under collagen activation. Hydroxyproline quantification was performed weekly during the four weeks of culture according to a previously described protocol. The amount of collagen in ex vivo skin after 4 weeks of culture determined by the hydroxyproline assay was 106.53 μg / mg, while for rabbit skin it was 69.03 μg / mg.

[0191] 1.3 Conclusion

[0192] The analysis of hydroxyproline in biological tissues is a reference method for determining the content of collagen extracted from different tissues (GF Caetanoa, M. Fronzab et al. Pharmaceutical biology, vol. 54, NO. 11, 2555–2559, (2016)). However, this method has the difficulty of extracting collagen from the analyzed tissue. Therefore, the collagen concentration in dry animal skin measured by the hydroxyproline assay is 5-30% of the total dry mass (GF Caetanoa, M. Fronzabet al. Pharmaceutical biology, vol. 54, NO. 11, 2555–2559, (2016)).

[0193] Caetano et al. used the hydroxyproline assay to measure collagen in rat skin (GF Caetanoa, M. Fronzab et al. Pharmaceutical biology, Vol. 54, No. 11, 2555–2559, (2016)). The hydroxyproline assay has been standardized according to the mass of dry skin before extraction and assay. The study showed a ratio of about 6 μg / mg hydroxyproline per mg dry skin, which represents a mass ratio of 45 μg / mg collagen per mg dry skin or 4.5% of the skin analyzed.

[0194] Parsons et al. also used the hydroxyproline assay to measure collagen in mouse skin (Parsons KK, Maeda N, Yamauchi M, Banes AJ, Koller BH. Am J Physiol Endocrinol Metab. 2006 Jun; 290). The hydroxyproline assay was standardized according to the mass of dry skin before extraction and assay. The study showed a ratio of about 28 μg / mg hydroxyproline per mg dry skin, which represents 210 μg / mg collagen per mg mouse dry skin, a mass ratio of 21% in the analyzed skin.

[0195] Taking these results into account, the inventors produced in vitro skin with collagen composition consistent with extraction and hydroxyproline assay results found in the literature.

[0196] In fact, the present results show a collagen level of 106.53 μg collagen per mg in vitro skin (dry), a mass ratio of 10.6% in the analyzed sample, which is within the range of results obtained in animal skin by this quantitative method. A summary of the results obtained in the scientific literature and those obtained in this study is shown in Table 1.

[0197]

[0198] Table 1: Collagen extraction rates from animal tissues and skin in vitro as determined by the hydroxyproline assay.

[0199] 2. Elastin content

[0200] The mechanical properties of the skin are determined by its composition. Elastin, which accounts for only a few percent of the dry mass, is responsible for the mechanical response to low stresses. In particular, it prompts contraction after deformation, which helps the skin to leave a sensory impression after mechanical stress.

[0201] The inventors extracted and measured the total elastin content of in vitro skin. By comparing these results with studies on mammalian skin, the inventors found that the elastin content extracted from in vitro skin samples was within the range of mammalian skin.

[0202] 2.1 Elastin assay protocol

[0203] In vitro skin obtained by culturing fibroblasts for 4 weeks on membrane macroporous scaffolds of circular (19 mm diameter) or square (17 mm side) geometry was collected, washed with physiological saline solution (DPBS, Thermo ref. 10010), dried in a drying oven and weighed with a precision balance. The dried material was cut with a scalpel and digested with oxalic acid at high temperature and centrifuged to remove debris. The supernatant was collected for analysis. 100 μL of the supernatant was recovered and the elastin concentration in the supernatant was determined using the Fastin elastin assay kit (Biocolor LTD. ref. F2000). A standard curve (elastin provided in the kit) was obtained. The curve equation and regression coefficient R2 were obtained by linear regression of the standard curve.

[0204] 2.2 Results

[0205] - Solution calibration

[0206] To verify the efficiency of extracting elastin using this protocol under the present extraction conditions, the inventors conducted consecutive extraction tests on three 17 mm square in vitro skin samples and determined the amount of elastin present in each of these extracts (Table 2).

[0207]

[0208] Table 2: Average elastin amounts measured in three consecutive in vitro skin extractions.

[0209] Taking into account the standard deviation starting from the fifth extraction (#Extraction 5, Table 2), the inventors determined that four consecutive extractions using the aforementioned protocol could effectively extract elastin from skin in vitro.

[0210] To evaluate the non-specific signal fraction that could be associated with the unseeded scaffolds, the inventors performed the same extraction and elastin assay on unseeded membrane scaffolds. The inventors measured an absorbance equivalent to 1.42 μg elastin equivalent. This result allowed the determination of the threshold corresponding to the non-specific signal or the reading background. This value was very close to the value obtained in the fifth extraction of the previous experiment, confirming that four extractions were effective in extracting all the elastin present in the sample using the previously defined extraction method.

[0211] - Quantification of elastin

[0212] To evaluate the total elastin in two samples of in vitro skin, samples of 19 mm diameter were dried and then weighed. The dry matter mass measured for these two samples was 50.1 mg and 51.4 mg. Four consecutive extractions of elastin were performed with a total volume of 2 mL containing elastin extracts. Nonspecific signals were subtracted from the measured values ​​of the previously performed assays. The in vitro skin samples numbered #1 and #2 contained 232.55 μg and 209.69 μg of elastin, respectively, which were 0.45% and 0.42% of their dry mass (Table 3).

[0213]

[0214]

[0215] Table 3: Average weight of elastin measured on two samples of in vitro skin.

[0216] Turner et al. measured elastin in pig skin samples from different locations using a method similar to that described herein. The amount of elastin was standardized to dry mass. The measured values ​​reported in this study ranged from 2.45-11.30 μg / mg, i.e., mass percentages of 0.24% and 1.1% (Turner NJ, Pezzone D, Badylak SF. Tissue Eng Part C Methods. 2015 Apr; 21(4): 373-84).

[0217] Taking into account the measurements carried out within the framework of this work, the inventors produced an in vitro skin in which the elastin composition was consistent with the extraction and quantification results found in the literature. In fact, using this experimental quantitative method, the inventors measured a mass proportion of elastin in dry in vitro skin of 4.35 μg / mg, corresponding to a mass ratio of 0.43%, while the values ​​measured in the literature range from 0.24% to 1.1%. The results obtained in the study of Turner et al. and summarized by the present study are shown in

[0218] Table 4.

[0219]

[0220] Table 4: Determination of elastin extraction yield and quantification from animal tissues and non-tanned in vitro skin materials.

[0221] 3. Analysis of the Orientation of Biomaterial Fibers in Skin in Vitro

[0222] According to the method previously described in Hamley et al. (Hamley, Introduction to Soft Matter: Polymers, Colloids, Amphiphiles and Liquid Crystals, John Wiley & Sons Ltd, Chichester, UK 2000), the image was oriented using the ImageJ Directionality PlugIn. https: / / imagej.net / plugins / directionality ) 2D Fast Fourier Transform (FFT) analysis that measures the arrangement of biomaterial networks observed by SEM in in vitro skin. The Directionality plugin calculates the spatial frequencies in the image based on a set of radial directions. The method produces a normalized histogram that reveals the number of fibers present in the 0°-180° 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 samples show the direction vectors of the biomaterial fibers and a histogram of the fiber directionality frequency distribution is shown in Fig.13 middle.

[0223] The results of the directional histogram analysis of the biomaterial orientation in the in vitro skin surface showed that there was no representative peak and the percentage of variation was consistent with that of an isotropic material ( Fig.13 C).

[0224] Animal hides, on the contrary, are anisotropic materials, with mechanical and structural weaknesses in specific directions relative to the animal's axis of symmetry from which the skin originates. The isotropic surface of in vitro skin leathers is explained by their production method, which, in contrast to the growth of animal hides, allows a high degree of control over the parameters that influence cell growth. This difference from conventional animal hides simplifies the use of the hides, which do not require special consideration of the direction of cuts on their surface.

[0225] 4. Heavy metal content

[0226] The presence of excessive amounts of hazardous substances in clothing materials may pose a threat to consumer health, especially if the product is intended for children and / or comes into direct contact with human skin. As heavy metals are one of these hazardous substances, threshold values ​​for heavy metal content are set in the standards applicable in different jurisdictions. These standards are intended to limit consumer exposure to heavy metals and ensure that the product life cycle is compatible with environmental issues.

[0227] 4.1 Materials and methods

[0228] Metal quantification was performed on in vitro skin (size: 26×43 cm) by an external reference research laboratory (Contract Research Organization, CRO) Eurofins BLC (UK). The determination of heavy metal content in in vitro skin was performed in accordance with international standards, measuring: a) total heavy metal content on digested samples, and b) heavy metal content extracted with artificial acid sweat solution. Table 5 shows the limits of total heavy metal content and extractable heavy metal content for acceptable materials of commercial leather.

[0229] Sampling is carried out in accordance with BS EN ISO 2418-2017, which specifies the method of sampling (location of leather pieces), labelling and marking of samples to ensure identification. It applies to all types of leather of mammalian origin, regardless of the method of tanning.

[0230] Metal screening was performed by atomic absorption spectroscopy (AAS) on pre-microwave digestion of leather according to BS EM ISO 17072-2:2019 and CPSC-CH-E1001-08.3. The detection limit for heavy metals tested using these methods is 0.1 ppm or mg / kg. Where necessary, samples were conditioned and tested at 23°C and 50% relative humidity (standard atmospheric pressure) according to BS EN ISO 2419-2012.

[0231] The determination of extractable metals is carried out by atomic absorption spectrometry (AAS) according to BS EN ISO 17072-1:2019, using extraction with an artificial acid sweat solution as a preliminary step. This method can determine the extractable metals in leather; it is not specific to the compound or the oxidation state of the metal. The method is particularly suitable for the determination of extractable chromium in chrome-tanned leather. Where necessary, samples are conditioned and tested at 23°C and 50% relative humidity (standard atmospheric pressure) according to BS EN ISO 2419-2012. The detection limit for heavy metals tested using these methods is 0.1 ppm or mg / kg.

[0232] The concentration of a chemical per unit mass in a solid is usually measured in milligrams (mg) or micrograms (μg) per unit mass of the total mass (kilograms (kg)), i.e. mg / kg or μg / kg. Sometimes these concentrations are expressed in parts per million (ppm) or parts per billion (ppb): 1 ppm = 1 mg / kg and 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.

[0233]

[0234]

[0235] Table 5: Extractable content limits of heavy metals in commercially available leather (total content limits), expressed in parts per million (ppm).

[0236] 4.2 Results

[0237] Extractable and total heavy metal content was quantified in in vitro skin. The results are shown in Tables 6 (extractable content) and 7 (total content), respectively.

[0238]

[0239] Table 6: Results of in vitro determination of extractable heavy metals in skin.

[0240]

[0241] Table 7: Results of in vitro determination of total heavy metals in skin

[0242] 4.3 Discussion

[0243] The physical properties of leather can vary depending on the manufacturing method and the structure of the chemicals used in the manufacturing process. The pH of leather can vary depending on the pH of the chemicals used in its manufacturing, the additives used in the final stage of the leather manufacturing process, and the proportions in which they are used.

[0244] The total amount of inorganic matter in raw hides can be determined by the ash method and should not exceed 2.5%. In addition, 0.5% of minerals can be present in natural raw hides, depending on the nutritional status of the animal. When examining the mineral content of skin in vitro, it was found that the amount of heavy metals was negligible, with only copper and lead present greater than 0.1 ppm or mg / kg. The presence of copper may be related to the cell culture medium used, while the source of the lead traces is difficult to determine. We found that all values ​​in the study were below the limit values ​​for heavy metal content in commercial leather.

[0245] Nowadays, consumer demand is shifting towards materials that do not contain harmful substances and have less impact on the environment. Therefore, there is a need to produce leather products that contain little or no heavy metals, especially in products that come into contact with human skin. This study found that the total heavy metal content in in vitro skin and extractable skin using artificial sweat solution was lower than the limit value for commercial leather (Table 5).

[0246] The low heavy metal content of in vitro skin makes it a viable alternative for commercial leather manufacturing. In conclusion, it is expected that in the future, the demand trend towards environmentally friendly products will have a greater impact on leather producers and the proposed limit values ​​will be reduced. Therefore, the leather industry, especially the processing of natural leather, must be prepared for these new limit values ​​and future transitions. In this context, alternatives such as the one proposed in this study are responsible solutions that do not sacrifice the quality of products manufactured using animal skins.

[0247] 5. Quantification of free amino groups

[0248] 5.1 Materials and methods

[0249] Proteins were extracted from in vitro skin by sonication (Sonicator Q700 Qsonica) (50% power, 14 s). Proteins were stained by adding 0.5 mL of 0.01% (w / v) TNBSA solution (Thermo Scientific. ref. TS-28997) to 0.5 mL of each sample and incubating 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 HCl solution.

[0250] The absorbance of the solution was measured at 335 nm. The quantification of the amines contained in the sample was determined by comparison with a calibration curve established with glycine solution (10 mg / mL, serially diluted to generate a standard range).

[0251] 5.2 Results

[0252] A calibration line was constructed with glycine by measuring the absorbance at 335 nm, which corresponds to the chromogenic compound formed after reaction with TNBSA at a given concentration (Table 8).

[0253] Concentration (mg / L) 0 5 10 15 20 25 30 Absorbance at 335 nm 0,2221 0,2970 0,3785 0,4303 0,4879 0,5453 0,5945

[0254] Table 8: Absorbance values ​​at 335 nm as a function of glycine concentration as standard (mg / L).

[0255]

[0256] Table 9: Values ​​of free amine content in in vitro skin and non-seeded membrane scaffolds obtained by interpolation of the calibration line.

[0257] 5.3 Discussion

[0258] Under certain conditions, the extensive use of dyeing and tanning in consumer products such as leather, clothing and toys can lead to the formation of total / free amino groups. For this reason, the European Parliament recently adopted the 19th amendment to Council Directive 76 / 769 / EEC on the restriction of certain substances. The legislation currently under review sets limits of 6207 mg / L for total amines and 75 mg / L for free amines. There is an industrial method CEN ISO TS17234 based on DIN 53316 that can detect the 20 substances listed in the legislation.

[0259] The free amine levels of the membrane scaffold and in vitro skin were 12.9 and 25.9 mg / L, respectively, which are well below the acceptable limit of free amines stipulated by the directive (75 mg / L). These results are very encouraging and in line with green technology.

[0260] To compare the free amine content of in vitro skin and mammalian skin, a literature analysis of the scientific literature on this topic was performed. Soomro et al. used a new method to quantify the free amine content of human skin by pre-column derivatization (chromatography) with trifluoroacetylacetone and isobutyl chloroformate as mobile phase (Soomro SA (2014), Journal of chromatography). This method was used to analyze amines in hydrolyzed human skin samples. In human skin, total free amines were quantified at approximately 0.9 mg / g human skin using gas chromatography.

[0261] The total free amine content in the in vitro skin was 25.6 mg / L (0.75 mg / g in vitro skin). This result is comparable to that found in the literature for mammalian skin. Therefore, various tanning agents commonly used in the traditional leather industry can be used to tan the in vitro raw hides.

[0262] 6. Determination of fatty acid content

[0263] 6.1 Materials and methods

[0264] Fatty acid quantification was performed by an external reference research laboratory (Contract Research Organization, CRO) Eurofins BLC (UK). Samples were received and processed according to ISO 2418:20179, which specifies the location of laboratory samples in the leather pieces and the method of marking and labeling laboratory specimens for future identification. Samples were reconditioned and tested at 23°C ± 2°C and 50% relative humidity in accordance with the standard reference environment requirements of ISO 2419:2012 (leather).

[0265] Quantification of skin fatty acids in vitro was performed by BLC coupling: a) fat extraction by supercritical chromatography (SFC)4 followed by b) GC-MS analysis according to the protocol described in GB / T9722-200611.

[0266] Internal standards were selected to determine the main components of fatty acids, volatile reactants and other impurities. Hydrogen / nitrogen was used as carrier gas, and diatomaceous earth (0.18 mm to 0.25 mm) was used as stationary phase. The flow rate was determined according to the detection limit of different fatty acids. The length of the column was set to 2 m, and the temperature was maintained at 80 ° C throughout the experiment. The method is detailed in ISO 12966-4:201512.

[0267] Sampling is carried out according to ISO 2418-20179, which specifies the sampling method (location of the leather pieces), sample labelling and marking to ensure identification. It applies to all types of leather of mammalian origin, regardless of the tanning method used.

[0268] In vitro skin (26 x 43 cm) or rabbit skin samples were washed with 1X DPBS saline (Fisher, 12037539), sealed in a vacuum machine, and sent to a reference research laboratory (CRO).

[0269] Gas chromatography-mass spectrometry (GC-MS) is a sensitive method for identifying and quantifying fatty acid content. In order to detect them, the carbon chain molecules need to be modified in advance. Therefore, fatty acids are usually modified by derivatization methods using acids (most commonly hydrochloric acid) to form fatty acid methyl esters (FAME). In the GC phase, the samples are separated according to different FAME. These FAME are then strongly fragmented by electron impact ionization in the mass spectrometer and many fragments are formed. For quantitative purposes, the largest fragments that show high abundance and highest specificity for a single fatty acid type are selected.

[0270] 6.2 Results

[0271] Total fatty acid content was quantified in in vitro skin using the method described above. The results are shown in Table 10.

[0272]

[0273] Table 10: Quantification of fatty acid content in in vitro skin samples

[0274] In this study, the measured values ​​of fatty acid levels in skin in vitro were low and below the detection limit of the measured values ​​<10 mg / kg.

[0275] 6.3 Conclusion

[0276] Various published results indicate that the total fatty acid content of animal skin is about 4%. After using a bath lotion, the total fatty acid content increases to about 15% (Mihai AL; Metrology Promoting Harmonization & Standardization in Food & Nutrition; 3Imeko Foods). The fatty acids present in leather make it soft and flexible. It also prevents the leather from wearing and makes it waterproof. The total fatty acid content was quantified in in vitro skin. It was found to be less than 10 mg / kg, equivalent to 0.001%. Compared to the 4% fatty acids of conventional animal skin, its content is low.

[0277] Some product lines require soft and flexible leather, such as gloves and other products that come into contact with the consumer's skin. The softness and flexibility of the leather can be adjusted during the bath treatment process, during which the bath evenly coats the individual fibers and gives them specific properties (Alaskan, A; Polymer Bulletin-Volume 79, doi.org / 10.1007 / s00289-021-03579-Z).

[0278] Due to the low fat content of the in vitro skin, the fat content can be precisely adjusted in the subsequent bath feeding steps to make it suitable for various uses in various products. In addition, the low oil content of the in vitro skin makes it unnecessary to perform a degreasing step and avoids the risk of oil spillage and condensed stains on the surface of the in vitro skin, which are common defects in conventional raw hides.

[0279] 7. Determination of sulfated GAG and hyaluronic acid (HA) content

[0280] Glycosaminoglycans (GAGs), including sulfated GAGs and hyaluronic acid, are responsible for the compressive strength of the skin. However, the tanning process requires the removal of GAGs. Their water-retaining capacity prevents the circulation of tanning agents in the collagen network; therefore, GAGs must be removed to open up this network.

[0281] 7.1. Determination of hyaluronic acid and sulfated glycosaminoglycan content

[0282] In vitro skin (with collagen secretion inducer: 19 mm diameter disc) or rabbit skin (170.1 mg wet weight) samples were washed with physiological saline solution (DPBS, Thermo ref. 10010), weighed, cut with a scalpel and digested with proteinase K at 55°C under stirring, then ground with a ceramic mill and centrifuged to remove debris and collect the supernatant for analysis.

[0283] The content of sulfated glycosaminoglycans was determined by simultaneous staining and precipitation of sulfated GAGs with dimethylmethylene blue, dissociation of the GAG-dye complex and quantification of the released dye according to the protocol provided by the manufacturer (Biocolor LTD. ref. B1000). A standard curve (chondroitin sulfate provided in the kit) was obtained and the curve equation and regression coefficient R2 were obtained by linear regression of the standard curve.

[0284] The hyaluronic acid content was determined by adding acetate-saturated ethanol to precipitate total GAGs, adding cetylpyridine to resuspend HA, repeating the previous steps to purify HA separation, and resuspending HA in distilled water. The concentration of hyaluronic acid present in the sample was determined according to the protocol provided by the manufacturer. A standard curve (chondroitin sulfate provided in the kit) was obtained, and the curve equation and regression coefficient R2 were obtained by linear regression of the standard curve.

[0285] 7.2 Results

[0286] - Determination of sulfated GAG content

[0287] To evaluate the concentration of GAGs in the three in vitro skin samples, 19 mm diameter samples were weighed after drainage. Then, the samples were cut, digested with proteinase K treatment and ground. The amount of GAGs was measured according to the instructions of the Blyscan kit (B1000). A standard curve was established using a chondroitin sulfate standard. The concentrations measured in the skin are reported in micrograms of chondroitin sulfate equivalents (μg CSE) in Table 11.

[0288]

[0289] Table 11: Average mass of sulfated GAGs measured in three in vitro skin samples. * The sample volume used for quantification of GAGs in rabbit skin was 10 μL instead of 100 μL.

[0290] To assess the fraction of nonspecific signal that could be related to the unseeded scaffolds, the same extraction protocol was used on the three unseeded scaffolds and the concentration of sulfated GAGs was determined. The absorbance corresponding to 0.44 μg chondroitin sulfate equivalent (CSE) was measured and a threshold corresponding to the nonspecific signal could be determined, which was subtracted from each measurement.

[0291] - Determination of HA content

[0292] To assess the hyaluronic acid (HA) content in the in vitro skin samples, the same samples were analyzed using the Biocolor Purple-Jelley kit (H1000). A standard curve was established using HA standards, and the concentrations measured in the in vitro skin are shown in Table 12. The nonspecific signal was 0.04 μg HA, corresponding to the signal of the unseeded membrane scaffold.

[0293]

[0294] Table 12: Average HA mass measured in three samples of in vitro skin * The sample volume used for HA quantification in rabbit skin was 5 μL instead of 10 μL.

[0295] - Test for determination of the amount of GAG and HA per mg dry mass

[0296] To report the measured values ​​of GAG and HA as the dry quality , the same procedure was also performed on in vitro skin samples of the same size after oven drying (50 °C, 6 h). However, after subtracting nonspecific signals, the total mass of sulfated GAGs was 0.04 μg ECS, while the total mass of AH was 0.01 μg. This corresponds to a density of 1.3 ng ECS / mg sulfated GAGs and 0.3 ng / mg HA, which is much lower than the wet weight determination, while the expected density should be higher. Therefore, the drying process seems to interfere with the quantification of GAGs in in vitro skin.

[0297] 7.3 Discussion

[0298] The amount of GAGs in the skin in vitro was assessed using the sulfated GAG and HA quantification technique. The mass density of the GAGs in these skins was measured: the average density of sulfated GAGs was 87.9 ± 2.2 ng ECS ​​ / mg (chondroitin sulfate equivalents), and the average density of HA was 57.86 ± 4.9 ng / mg. This density is referred to as wet mass, because drying interferes with the measurement. This can be attributed to extraction problems after drying, for example due to collapse of the membrane scaffold around the GAGs, or even due to degradation during drying.

[0299] A study conducted by Armstrong and Bell measured HA 551 ng / mg wet skin in rabbit skin (Armstrong SE, Bell DR. Anal Biochem 2002; 308(2): 255-64). This result is 1.67 greater than the result presented herein, but of the same order of magnitude. This difference may be explained by physiological changes, such as the amount of HA in the skin decreases with age and location (Barbosa I, et al. Glycobiology 2003; 13(9): 647–53; Templeton DM. Connect Tissue Res 1988; 17(1): 23-32). According to Varma et al., the distribution of GAG in the skin of different animals (including rat skin) is basically half HA, the other half chondroitin sulfate GAG, and a small amount 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. demonstrated that the amount of chondroitin sulfate (non-HA GAG) in rabbit skin is 319 μg / g dry weight, and the conversion factor to wet weight is 3:1. Using this correction, the inventors found that the concentration of chondroitin sulfate (non-HA GAG) is 106 ng / mg wet weight (Fabianek, Comparative Biochemistry and Physiology. Volume 14, Issue 1, January 1965, Pages 21-28). This value is 5.47 times smaller than the control rabbit skin measured in this study. This discrepancy may be explained by factors related to sample processing and differences in extraction and GAG determination techniques.

[0300] The amount of total GAGs in the in vitro skin, corresponding to the sum of sulfated GAGs (chondroitin sulfate) and HA, was 145.8 ng / mg, compared to 883.9 ng / mg measured in the control rabbit skin in this study. HA and sulfated GAGs were 5.2-fold and 6.7-fold lower, respectively, compared to the rabbit skin analyzed. However, comparing these results with those of Armstrong and Bell and Fabianek et al., it was found that the difference in sulfated GAGs between rabbit skin and in vitro skin was -17%, and HA was reduced by 9.5-fold.

[0301] The mass level of sulfated GAGs measured in the in vitro skin was 87.9±2.2 ng / mg and the HA level was 57.86±4.9 ng / mg. Although these values ​​are lower than those measured in rabbit skin, the concentration of sulfated GAGs (chondroitin sulfate) is close to the concentration of 106 ng / mg found in the literature for rabbit skin.

[0302] F. In vitro leather characterization

[0303] 1. Differential Scanning Calorimetry (DSC)

[0304] The property commonly used to characterize collagen (whether natural, structurally modified or modified by tanning chemistry) is hydrothermal stability: this is defined as the effect of moisture and heat on the integrity of the material, usually expressed as a denaturation transition (ADCovington, RA Hancock, IA Ioannidis, J. Soc. Leather Technologists Chemists 73, 1989) 1:8). Depending on the processing (tanning), the value of this parameter is an important physical parameter for characterizing raw hides and their typical uses. Covalent bonds, especially those generated during the tanning process by crosslinking of tanning agents with collagen fibers, increase the size of the cooperative unit through intermolecular and intramolecular crosslinking and increase the denaturation temperature (Td) (S. Menashi, A. Finch, PJ Gardner, DA L Edward, Biochem. Biophys. Acta 144 (1976) 623: 625). Denaturation is defined as a transition from a triple helix to a randomly wound form, which occurs in the domains between crosslinks (M. Komanowsky, J. Am. Leather Chemist Assoc. 86 (8) (1991) 269: 28). The bonds that stabilize the supercoil are hydrogen bonds, hydrophobic bonds, van der Waals bonds, and interactions between residues with opposite charges on the side chains. The non-random distribution of ionizable and hydrophobic side chains along the repeating unit leads to the appearance of charged and hydrophobic blocks, which contribute to the stability of the structure through electrostatic and hydrophobic interactions (GS Young, Stud. Conserv. 43 (2) (1998) 65: 79).

[0305] This increase in Td varies with the nature and type of reactive groups involved. Therefore, the method of tanning the leather directly affects its shrinkage. Traditionally, the Td of leather tanned with vegetable agents fluctuates around 70°C, while the Td of chrome-tanned leather fluctuates around 80-90°C. For leather tanned with synthetic tanning agents, Td is 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).

[0306] The hydrothermal stability of collagen in skin and leather is characterized by the shrinkage of the material when heated in water at a specified temperature. Differential scanning calorimetry (DSC) allows the study of the enthalpy changes associated with collagen denaturation, where shrinkage is a macroscopic manifestation. This work aims to analyze the hydrothermal stability of leather obtained according to the method of the present invention. Before tanning, uninoculated macroporous membrane scaffolds and in vitro skin were used as reference materials. Tanned leathers of in vitro skin according to the present invention were tested with two different methods and aldehyde tanning to compare the effects of the tanning methods on the denaturation temperature (Td) of the tanned leathers. The results of the analysis and measurements show that the tanned leathers of in vitro skin have a hydrothermal stability close to that of tanned leathers of animal origin.

[0307] 1.1 Differential Scanning Calorimetry (DSC) Protocol

[0308] The measurements were performed using a 2014 Polyma DSC calorimeter (Netzsch). Temperature and heat flux were calibrated with indium (melting point: 156.6 °C and latent heat of fusion: 28.45 J / g). The samples were weighed (3 mg), immersed in water for several hours and then sealed in an aluminum crucible. The thermal changes were measured relative to the observed values ​​of the reference sample, purged by a 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 rate at which heat must be applied to keep the two tests at the same temperature. When the test sample undergoes an endothermic process, the energy input required to maintain the temperature increases and a signal is given on the rising temperature curve.

[0309] 1.2 Results

[0310] Differential scanning calorimetry (DSC) is used to determine the denaturation temperature (Td) of the sample. This denaturation temperature is obtained by analyzing the variation interval of the transition temperature (onset and offset), where the peak is assimilated to the denaturation temperature (Td). The last method is more suitable for the complex systems (scaffolds and biomaterials) of the present invention (Carsote, C., Badea, E.Herit Sci 7, 48 (2019)).

[0311] The measurements were performed using a Netzsch DSC 2014 Polyma calorimeter at a constant rate of 5°C / min in the range of 10-95°C. Fig.14 Shown are the DSC curves of different samples: macroporous scaffold (FSC), in vitro skin (FP), aldehyde tanned leather (CF-1). Fig.15 Decomposition of the peaks showing the different species present in the sample and determination of the denaturation temperature: the onset temperature (T onset ), corresponding to the denaturation temperature of the peak (Td) and the shift temperature at the end of the transition (T offset ).

[0312] Looking closely at the thermograms obtained for the different samples, these can be decomposed into two (for FSC samples) and three (for FP, CF-1 and CF-2 samples) peaks / deconvolutions with intervals corresponding to different denaturation temperatures occurring in this complex system. The denaturation temperatures for each sample and each of these peaks / deconvolutions are shown in Table 13. The results show a minimum recorded Td of 44.2°C, corresponding to the first peak / deconvolution of the unseeded macroporous scaffold (FSC), with almost no detectable denaturation enthalpy. The first peak / deconvolution of in vitro skin (FP) increases to 52.6°C, and in terms of enthalpy, this minority of peaks can correspond to collagen that has not been modified by tanning, with values ​​close to those of non-chemically modified collagen obtained by Carsote et al. (Carsote, C., Badea, E.Herit Sci 7, 48 (2019)).

[0313] The Td of the second peak / deconvolution of the unseeded macroporous scaffold (FSC) rises to 77.4 °C, which is the largest in terms of enthalpy and should correspond to the collapse of the geometry of the unseeded scaffold at high temperatures. The denaturation temperatures are almost similar for the second peak / deconvolution of in vitro skin (FP) rawhide and aldehyde tanned (CF-1) leather, which are 72.1 and 71.8, respectively. The second peak / deconvolution with a higher enthalpy than the first peak should correspond to more or less chemically modified collagen fibers and biomaterials entangled on the scaffold, which form a more heat-resistant composite.

[0314] The highest denaturation temperatures recorded for in vitro skin (FP) and aldehyde tanned (CF-1) leather are 83.5 and 82.9, respectively. The third and last peak with the highest enthalpy corresponds to the collagen fibers chemically bound by the tanning agent and the biomaterial, which are more or less chemically modified and entangled in the scaffold, thus forming a composite material resistant to high temperatures as a whole. The presence of collagen and chemically modified biomaterials makes the composite material resistant to thermal denaturation and maintains the geometry of the composite at about 83°C, compared to 77.4°C for the unseeded macroporous scaffold.

[0315]

[0316] Table 13: Denaturation temperatures measured by DSC

[0317] These results show the enthalpy change associated with collagen denaturation and the temperature at which this phenomenon occurs in leather tanning ( Fig.14 and Table 13). Covalent bonds, especially those generated during the tanning process, stabilize collagen fiber bonds and increase their denaturation temperature (Td). In particular, this stabilization effect brought about by tanning was observed when the Td of the first peak / deconvolution of in vitro skin (FP) increased from 52.6°C before tanning to 59.3°C of aldehyde tanned leather (CF-1). Regarding the second and third peaks / deconvolutions, the inventors observed that the formation of a composite material of mixed collagen, biomaterial and scaffold had a significant effect on the denaturation temperature (Td) compared to tanning. In fact, these second and third peaks / deconvolutions Td were similar between in vitro skin and aldehyde (CF-1) tanned leather.

[0318] These results suggest that the effects of tanning agents on Td in animal hides can be present in in vitro skin-tanned leathers.

[0319] Larsen et al. measured the Td of different tanned hides, including Mimosa and Sumach (Larsen, René & Vest, M. & Nielsen, K.. (1993), Journal of the Society of Leather Technologists and Chemists. 77. 151-156). They compared the Td of untanned hides with the Td of these two leathers. Their results showed that the Td increased significantly after tanning, with Mimosa and Sumach being 75.3 and 79.3°C, respectively, and untanned hides being 56.5°C. In addition, they reported the inhomogeneity of the measurements between the two leather samples, and this could be explained by the organization of the collagen fibers, which occurred in a tanning-dependent manner.

[0320] As mentioned above, the thermograms obtained for different samples of the present invention are complex because they consist of two or three different peaks / deconvolutions. Similar analysis of the decomposition of DSC thermograms has 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 the composite system show that there are endothermic fluctuations in the degradation and rearrangement regions of the polymer chains. This behavior can be estimated by estimating the onset denaturation temperature (T onset ), peak temperature (Td) and shift denaturation temperature (T offset ) to determine the denaturation region of the system. These temperatures reflect, in turn, the onset of denaturation, which corresponds to the reorganization of chains of different natures, the optimal deformation of the chains and the stability after deformation. This phenomenon is similar to the observations obtained with the samples of this study.

[0321] In each thermogram of FP and FC-1, three fluctuations are clearly visible, affected by the stability before and after denaturation. This behavior is not detectable in the scaffold (FSC), which, unlike the other samples, is a non-complex system that does not contain biomaterials. In the in vitro cultured skin (FP), the complex system consists of a macroporous scaffold (FSC) and biomaterials containing cells with an extracellular matrix (ECM) rich in collagen fibers. Tanned leather (CF-1 and CF-2) constitutes a complex ternary system containing a scaffold (FSC), biomaterials, especially cross-linked collagen fibers, and a tanning agent. Therefore, as Fig.15 As shown in the decomposition of the curve in , the denaturation area corresponding to the material state is defined. From these decompositions, it can be seen that the leather of the present invention shows similar denaturation characteristics, and the denaturation temperature range varies from 55.8 to 88.7°C depending on the aldehyde tanning method. The average Td (72.2°C) characteristics of the leather of the present invention are very close to the range of new aldehyde tanned leather (76.6°C; 87°C) (Bai, Xue & Jinming, Chang & Chen, Yi & Fan, Haojun & Shi, Bi. (2013)). Journal of the American Leather Chemists Association. 108. 404-410).

[0322] Another important factor that strongly influences the Td value is the water content of the material. Therefore, the lower the water content, the higher the Td. The reason for this phenomenon is that during the drying process, thermal stability is enhanced by forming strong intermolecular and intramolecular bonds and ionic bonds between acidic and basic groups. Typically, shrinkage phenomena are measured in aqueous media, and for this reason, the measurement of Td is carried out in excess water (A. Finch, DA Ledward, Biochim. Biophys. Acta 278 (1972) 433-439).

[0323] In this study, the inventors have retained these classical conditions in the implementation of the measurements. Therefore, considering these conditions, the analysis by DSC is very useful for evaluating the thermal stability of the leather and allows comparison between the two tanning methods used.

[0324] In order to make a reference between the measurements obtained by DSC and the temperature resistance measurements usually used in tanneries, the inventors used a shrinkage temperature tester (Tr) specially designed for leather. The device consists of a beaker filled with water, in which the leather sample is immersed under mechanical tension. The leather sample is then subjected to a series of temperature changes (20-100°C) using a heating plate placed under the system. The shrinkage of the sample at a given temperature (Tr) is recorded by the displacement measured by a thousandths of a meter (Borletti). The shrinkage temperature (Tr) of the in vitro skin (FP) and aldehyde-tanned leather (CF-1) was measured and shown in Table 14. The results show that the Tr of the in vitro skin (FP) fluctuates around 65°C, which corresponds to the temperature interval between the first peak / deconvolution and the second peak / deconvolution obtained by DSC ( Fig.14 , Table 13, FP). On the other hand, the Tr of the aldehyde-tanned leather fluctuates around 75 °C, which corresponds to the temperature interval between the second peak / deconvolution and the third peak / deconvolution obtained by DSC ( Fig.14 , Table 13, CF-1).

[0325] sample Shrinkage temperature (Tr) FP 65℃ CF-1 75℃

[0326] Table 14: Shrinkage temperature (Tr) of in vitro skin and in vitro skin tanned leathers.

[0327] The shrinkage temperature (Tr) measurements obtained using the tester are less accurate than those obtained by DSC (thin sample thickness and displacement measurements by an observer). Nevertheless, this method is more commonly used in tanneries and provides a useful model for comparing DSC measurements with those of tanneries and traders (GS Young, Stud. Conserv. 43(2)(1998) 65:79). The results show that, using the same method, the aldehyde-tanned leather of the present invention has a thermal stability very close to that of newly tanned animal leather.

[0328] 2. Mercury Intrusion Porosimetry

[0329] 2.1 Materials and methods

[0330] Mercury intrusion porosimetry measurements were performed with a microporometer (AutoPore IV 9500, Micrometrics) with a maximum applied pressure of 413 MPa. The samples used for the measurements were: scaffold (FSC) and aldehyde-tanned leather of in vitro skin (CF-1).

[0331] The measurement was carried out at 24°C. The FSC sample was pre-dried at 50°C for 24h and the CF-1 sample was pre-dried at 80°C for 24h. The equilibrium time for each pressure point was set to 10 seconds. A blank correction was used before measurement. The sample was cut into 3cm×2cm pieces (about 0.6g) and first degassed, and then injected with low-pressure distilled mercury. The mercury-filled injection gauge was taken out from the low-pressure port, weighed, and then placed in the high-pressure port. As the pressure increases, mercury enters the pores of the sample. The recorded pressure and the volume of mercury entering the pores are used to calculate the porosity.

[0332] 2.2 Results

[0333] - In vitro leather porosity characteristics

[0334] The total porosity (ε) calculated from the MIP test is listed in Table 15.

[0335] ε(%) FSC 87,21 FP 14,78 CF-1 51,63 Untanned cowhide 59,07 sheepskin 57,26

[0336] Table 15: Total porosity of scaffolds (FSC), ex vivo skin (FP) and aldehyde tanned leather (CF-1) compared to untanned bovine and chrome tanned sheepskin.

[0337] According to the data extracted from the MIP analysis shown in Table 15, the scaffold (FSC) has a high percentage of total porosity (ε) (87.21%). The design of the structural features of the scaffold according to the present disclosure shows that it has high porosity, and the average pore size is compatible with animal cell seeding and penetration, with an average diameter of 20 μm (Al Tawil, E., et al. (2018). European Polymer Journal).

[0338] The total porosity in in vitro skin (FP) dropped sharply to 14.78%. This drop is due to the occupation of the pore volume by biological substances: cells and extracellular matrix (ECM) components secreted by cells. This biological substance is the basic component of in vitro skin new tissue, and it is present in tanned rawhide and conventional animal leather (He, Xiu et al. (2019). Journal-American Leather Chemists Association. 114. 41-47).

[0339] Tanning is the step of converting conventional or external skin into leather by producing chemical bridges between the collagen fibers of ECM. Due to the cross-linking of this network, a space called macropore, mesopore and nanopore is produced between the collagen chains. This porosity of leather is a preferred characteristic and is the reason for its breathing phenomenon. With the leather (CF-1) tanned with aldehyde, the total porosity is reduced to 51.63%. These results are very similar to the results obtained with chrome tanned sheep leather (Table 15).

[0340] 2.3 Discussion

[0341] The porosity properties of a material are one of its fundamental characteristics, they are selection parameters that influence the framework of its use and the type of products in which it is used. For leather, the porosity of different types of leather determines its final use: decoration, clothing, leather goods, furniture, etc. It can also be used as an indicator for variations in its manufacturing process, especially during the retanning and finishing stages. In this report, the porosity properties of leather obtained from the in vivo skin of the present disclosure were measured using mercury intrusion porosimetry (MIP). The results show that the scaffold has sufficient porosity to be used for animal cell culture. The growth of cells within the scaffold allows the cells to colonize within the scaffold and deposit biological substances, which are essential components of the in vitro skin. The in vivo skin leather has a total porosity comparable to that obtained with conventional animal leather.

[0342] 3. Water vapor permeability

[0343] 3.1 Materials and methods

[0344] -Material

[0345] Water permeability measurements were performed using a permeameter-diffusion meter (IGAsorp, Hiden Isochema). It consists of two compartments, one upstream of the sample and one downstream of the sample. During the measurement, permeates (water molecules) were introduced into the upstream compartment, while the downstream compartment was purged with dry gas (N2 BIP, Air Products). Detection was performed using the water measurement method. During the measurement, a mirror hygrometer was used to measure water-rich gas, and the probe 1311DR was from General Eastern (Elcowa-Mulhouse).

[0346] The experimental setup used for the measurements includes:

[0347] - a permeation cell for placing the sample to be tested, placed in a closed container at a constant temperature of 23°C and consisting of two compartments, one made of stainless steel and the other made of Plexiglas;

[0348] - Feed upstream of the sample, where a purge gas (technical grade nitrogen) circulates in the first step and the permeate then permeates in the second step;

[0349] - A feeder downstream of the membrane, in which BIP type nitrogen (gas) circulates, used as carrier gas for the mirror hygrometer.

[0350] The measurement was carried out at 23°C and 50% relative humidity. The sample used for the measurement was aldehyde tanned leather (CF-1). The active surface area of ​​the sample was 3.6 cm 2 .

[0351] - Experimental setup

[0352] The differential permeation measurement is performed in two consecutive steps. In the first step, called the cleaning step, as much water and residual molecules as possible are removed from the sample, the gas chamber and the gas circuit. Then, nitrogen is purged into the upstream and downstream compartments until a stable low water content is obtained. In the second step, the measurement step, water vapor at 50% relative humidity is introduced into the compartment. The permeate then diffuses through the sample and is carried by the carrier gas downstream of the sample to the detector (mirror hygrometer). The changes in the flow rate are recorded in real time from the changes in the dew point temperature measured directly.

[0353] - Flux and permeation calculations

[0354] With flow rate f = 9.3 cm 3 / s and total pressure pt = 1atm to supply dry nitrogen (vapor pressure p in The receiving compartment of )(BIP nitrogen) is not completely anhydrous and its vapor pressure varies from p in to p out (vapor pressure at the compartment inlet and outlet) increases. The flux J(L,t) is given by:

[0355] J(L, t) = f.(pout-pin) / SRTr Equation 1

[0356] R is the ideal gas constant, S is the exposed surface, and Tr is the temperature at the time of measurement (K).

[0357] The permeability coefficient P is determined by:

[0358] P = Jst.L / Δa Equation 2

[0359] Among them J st is the steady-state flux, L is the sample thickness, and Δ is the activity difference across the sample.

[0360] 3.2 Results

[0361] The water vapor permeability was measured using an osmotic diffusometer at 23°C and 50% relative humidity. The measurement results recorded in real time are shown in Table 16. The permeability coefficient of the in vitro skin leather (CF-1) tanned with aldehyde was 3.47 mg / cm 2 .h. Representative values ​​for cowhide have also been added to this table (cowhide: Radwag wagi elektroniks - testing laboratory. https: / / radwag.com / pl / ).

[0362] sample Thickness(μm) <![CDATA[Water vapor permeability (mg / cm 2 .h)]]> CF-1 455,2±0,7 3,47 Cowhide leather 230 0,7-5,3

[0363] Table 16: Water vapour permeability of different samples recorded at 23°C and 50% relative humidity.

[0364] 3.3 Discussion

[0365] There are different methods to measure the water vapor permeability of leather, the most widely used is the measurement method according to ISO 14268:2013 (Leather-Physical and mechanical tests-Determination of water vapor permeability (ISO / FDIS14268:2012).2013). Water vapor permeability depends on many operations during the leather manufacturing process, including tanning and finishing. Leather is manufactured in the laboratory by combining scaffolds and animal cells.

[0366] The leather (CF-1) was produced using an aldehyde tanning process. For CF-1, water vapor permeation measurements were recorded as 3.47 mg / cm 2.h. It is known that leather is generally considered to have good water vapor permeability (WVP), with a permeability higher than 20 and a water vapor permeability value higher than 0.8 (Leather international. 18 September 2001). This result shows that the leather has good water vapor permeability consistent with conventional leather. Different types of animal leather have been analyzed for water vapor permeability (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 / Smieshoweski et al. measured the water vapor permeability of different raw hides using standard methods and methods developed in their laboratory. The different types of leather selected for their study showed that for samples with a thickness of about 1.2 mm, the water vapor permeability ranged from 0.3 to 4.9 mg / cm 2 On the other hand, research published by Radwag Testing Laboratories shows that cowhide leather used in the footwear industry has a 0.7 mg / cm 2 .h-5.3mg / cm 2 .h water vapor permeability, which depends on the manufacturing method: wrinkled, pressed or untreated.

[0367] Based on these results, the inventors observed that the in vitro skin leather has a water vapor permeability comparable to that of conventional leather. The recorded values ​​are very close to the values ​​published by measuring leather samples of animal origin. It is important to note that this test was carried out on unfinished tanned leather with an average thickness of 0.45 mm; it has been shown that as the thickness of the leather decreases, the water vapor permeability increases significantly (Smiechowski K., et al. Journal of the Society of Leather Technologists and Chemists, ISSN 0144-0322, Vol. 98, Num. 6, 2014).

[0368] Considering these results, measuring the water vapor permeability of in vitro leathers of different thicknesses, tanning methods and potential processing levels will allow us to discover important manufacturing factors that modify the water vapor permeability of in vitro leathers.

Claims

1. A method for producing leather comprising the following steps: i) culturing fibroblasts in vitro on honeycomb macroporous polymer scaffolds to obtain tissue, The surface of the stent comprises 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, ii) tanning the tissue to form the leather.

2. The method of claim 1, wherein the scaffold comprises a polyester, preferably a biodegradable and / or bioderived polyester.

3. The method of claim 2, wherein the scaffold comprises polyester, preferably biodegradable and / or bioderived polyester, in a concentration of 1-50% (w / w), preferably 5-20% (w / w).

4. The method according to any one of claims 1-3, wherein the macropores are interconnected with micropores, preferably the micropores have a diameter of less than 20 μm, preferably less than 10 μm.

5. The method of any one of claims 1-4, wherein the tensile strength of the stent is 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.

6. The method of any one of claims 1-5, wherein the Young's modulus of the scaffold is 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, and more preferably 30-50 kPa.

7. The method according to any one of claims 1 to 6, wherein the scaffold is obtained by a non-solvent induced phase separation method (NIPS).

8. A method according to any one of claims 1-7, wherein the support is an asymmetric support comprising macropores, and the macropores are uniformly but irregularly distributed throughout the thickness of the support, in particular, the support has a first face and a second opposing face, the first face comprising macropores with a diameter of 100 μm-280 μm, preferably 110 μm-225 μm, opening to the surface, and 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, and the second opposing face comprises only nanopores with a diameter of less than 10 nm, preferably 5 nm.

9. The method according to any one of claims 1-7, wherein the scaffold is a symmetrical scaffold containing macropores, which has two faces, each of which contains macropores with a diameter of 100 μm-280 μm, preferably 110 μm-225 μm, and 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.

10. The method according to any one of claims 1 to 9, wherein bioactive molecules are grafted onto the surface of the scaffold.

11. The method of claim 10, wherein the bioactive molecule is collagen, a carbohydrate or a carbohydrate-derived molecule such as a glycosaminoglycan.

12. Tissue obtainable after step a) of the method according to any one of claims 1-11, preferably comprising fibroblasts cultured on a macroporous scaffold as defined in any one of claims 1-11.

13. The tissue according to claim 12, having at least two, preferably three, denaturation temperatures (Td), preferably at least three Td, determined by differential scanning calorimetry (DSC).

14. The tissue according to any one of claims 12 or 13, which is isotropic at the surface of the tissue.

15. The tissue according to any one of claims 12 to 14, comprising less than 1% (w / w) total fatty acids mass percentage in the total dry mass of the tissue.

16. Tanned leather obtainable by the method according to any one of claims 1 to 11.

17. Tanned leather according to claim 16, having at least two, preferably three, denaturation temperatures (Td) determined by differential scanning calorimetry (DSC).

18. Tanned leather according to claim 16 or 17, having a total porosity of 30-60%.

19. Use of a macroporous polymer scaffold for producing leather, preferably by tanning tissue obtained from a fibroblast cell culture on the macroporous scaffold, wherein the surface of the scaffold comprises macropores with a diameter of 100-280 μm, preferably 110-225 μm, and 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.

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