Field of the disclosure

[0001] The present disclosure relates to hydrogels, and in particular, extracellular matrix hydrogels obtained from decellularised tissue.

Background of the disclosure

[0002] The present application claims priority from Australian provisional application number 2022902125 filed on 28 July 2022, the entire contents of which are incorporated herein by reference.

[0003] Any discussion of the prior art throughout the specification should in no way be considered as an admission that such prior art is widely known or forms part of the common general knowledge in the field.

[0004] Recent studies have highlighted the significance of organoid technology in understanding human development and diseases (R. Heremans, Z. Jan, D. Timmerman, H. Vankelecom, Organoids of the Female Reproductive Tract: Innovative Tools to Study Desired to Unwelcome Processes. Front Cell Dev Biol 9, 661472 (2021); M. Hofer, M. P. Lutolf, Engineering organoids. Nat Rev Mater 10.1038/S41578-021 -00279-y, 1-19 (2021)). Organoids have provided us with an opportunity to study human developmental processes, such as early embryonic development and implantation, that were not investigated in detail due to ethical restrictions (H. Kagawa et al., Human blastoids model blastocyst development and implantation. Nature 601 , 600-605 (2022)). Patient-derived organoids are used in clinical practice for matching the right drug to the right patient. For example, rectal organoids are currently used to identify patients that would benefit from cystic fibrosis transmembrane conductance regulator-modulating drugs (J. F. Dekkers et al., Characterizing responses to CFTR- modulating drugs using rectal organoids derived from subjects with cystic fibrosis. Sci Transl Med 8, 344ra384 (2016)).

[0005] Human endometrial organoids have been developed from healthy and disease tissue biopsies. Endometrial glandular epithelial cells embedded in a small drop of Matrigel supplemented with activators or inhibitors of WNT, EGF, FGF, BMP, TGFb, and ROCK signaling in the culture medium allow the development of endometrial organoids from single cells (M. Y. Turco et al., Long-term, hormone-responsive organoid cultures of human endometrium in a chemically defined medium. Nat Cell Biol 19, 568-577 (2017); M. Boretto et al., Development of organoids from mouse and human endometrium showing endometrial epithelium physiology and long-term expandability. Development 144, 1775-1786 (2017)). Similar to human endometrial tissue in vivo, endometrial organoids respond to ovarian hormones and emulate cell type-specific changes that occur during the different phases of menstrual cycle. These organoids can be maintained in culture for a long time (~12 months) and cryopreserved without any apparent loss of their typical features. Organoids representing several different endometrial pathologies have also been developed, and they have already provided insights into the origins of different subtypes of endometrial cancer and revealed new biomarkers for the risk stratification of endometrial cancer patients.

[0006] The extracellular matrix (ECM) is an acellular component that not only acts as a physical scaffold for cellular organisation but also provides important biochemical and biomechanical signals affecting many basic cellular processes required for normal tissue development, differentiation and homeostasis (M. W. Pickup, J. K. Mouw, V. M. Weaver, The extracellular matrix modulates the hallmarks of cancer. EMBO Rep 15, 1243-1253 (2014); Y. Brown, S. Hua, P. S. Tanwar, Extracellular matrix-mediated regulation of cancer stem cells and chemoresistance. Int J Biochem Cell Biol 109, 90- 104 (2019)). ECM extracted from tissues by decellularization can retain the biochemical complexity, nanostructure and bio-inductive properties of the native tissue ECM and has been shown to support the regeneration of functional tissues (S. F. Badylak, D. O. Freytes, T. W. Gilbert, Extracellular matrix as a biological scaffold material: Structure and function. Acta Biomater 5, 1-13 (2009); L. T. Saldin, M. C. Cramer, S. S. Velankar, L. J. White, S. F. Badylak, Extracellular matrix hydrogels from decellularized tissues: Structure and function. Acta Biomater 49, 1-15 (2017)). ECM extracted from decellularized tissues therefore represents an excellent natural material for regenerative therapies.

[0007] Currently, organoids are cultured in commercially available ECM derived from murine Engelbreth-Holm-Swarm (EHS) sarcoma (trade name Matrigel). The EHS tumor is an embryonal carcinoma derived from parietal endoderm that secretes an excessive amount of basement membrane extracellular matrix (e.g., collagen IV, laminin-1 , nidogen-1 , and perlecan) (S. Futaki et al., Molecular basis of constitutive production of basement membrane components. Gene expression profiles of Engelbreth-Holm-Swarm tumor and F9 embryonal carcinoma cells. J Biol Chem 278, 50691-50701 (2003); R. W. Orkin et al., A murine tumor producing a matrix of basement membrane. J Exp Med 145, 204-220 (1977)). Both biomechanical and biochemical properties of the EHS tumor-derived ECM are quite different from the normal tissue ECM.

[0008] In this context, it is an object of the present invention to overcome or ameliorate at least one of the disadvantages of the prior art, or to provide a useful alternative.

Summary of the disclosure

[0009] In work leading to the present disclosure, the inventor surprisingly found that extracellular matrix hydrogel obtained from native healthy tissue can support organoid growth and organogenesis in vivo. Using human and bovine endometrial tissue, the inventors developed hydrogels which supported the growth of endometrial, colon and lung organoids. Organoids grown on these hydrogels were proteomically more similar to native tissue than organoids grown in Matrigel.

[0010] In one aspect, the present disclosure provides an extracellular matrix hydrogel obtained from a decellularized healthy tissue.

[0011] The tissue is preferably an endometrial tissue.

[0012] In another aspect, the present disclosure provides an extracellular matrix hydrogel obtained from a decellularized endometrial tissue.

[0013] In some examples, the endometrial tissue has been separated from muscle tissue. In some examples, the endometrial tissue has been separated from myometrium tissue (smooth muscle of the uterus).

[0014] In some examples, the endometrial tissue is not cancerous. The endometrial tissue may be obtained from a biopsy or a uterus. [0015] The tissue may be obtained from a human or bovine subject.

[0016] The tissue may be decellularized by treating the tissue with a surfactant selected from sodium dodecyl sulfate (SDS) and sodium deoxycholate (SDC). In some examples, the surfactant is present at a concentration of between about 1% and 4%. In some examples, the surfactant is present at a concentration of about 4%. In some examples, the surfactant is SDC.

[0017] The hydrogel may be supplemented with a cross-linker. The cross-linker may be genipin or N- (3-Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC).

[0018] In some examples, the hydrogel is manually shaped or moulded. For example, the hydrogel may be in the form of a tube or lumen.

[0019] In some examples, the hydrogel is supplemented with laminin.

[0020] The hydrogel may be obtained by: lyophilizing or freeze-drying the decellularized tissue sample to form a frozen sample; crushing, grinding or milling the frozen sample to form a powder; treating the powder with an acid and/or protease to form a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0021] In some examples, the hydrogel has a pH of between about 5 and 9, such as between about 6 and 8.

[0022] In another aspect, the present disclosure provides a scaffold formed from a hydrogel of the invention.

[0023] The scaffold may be porous, and may be formed by ice templating in which the hydrogel is frozen and lyophilised to produce a porous scaffold.

[0024] The scaffold may be used in cell culture and/or in tissue regeneration.

[0025] In another aspect, the present disclosure provides an organoid obtained by culturing cells on a hydrogel of the invention.

[0026] In some examples, the organoid is an endometrial organoid. In some examples, the cultured cells are endometrial cells.

[0027] In some examples, the organoid is an organoid of a tubular organ. The tubular organ may be a colon, a lung or a gastrointestinal organ. In some examples, the organoid is a human organoid.

[0028] In another aspect, the present disclosure provides a method of promoting repair of a wound comprising applying to the wound a hydrogel of the invention, a scaffold of the invention or an organoid of the invention.

[0029] The wound may be an endometrial wound, a uterine wound, a fallopian wound, an ovarian wound, a vaginal wound, a colon wound, a lung wound, a wound of a gastrointestinal organ, a flesh wound, a liver wound or a cardiac wound.

[0030] In another aspect, the present disclosure provides a method of promoting tissue regeneration comprising applying to the tissue a hydrogel of the invention, a scaffold of the invention or an organoid of the invention. [0031] In another aspect, the present disclosure provides a method of treating an endometrial disorder in a subject comprising administering to the subject a hydrogel of the invention, a scaffold of the invention or an organoid of the invention.

[0032] In some examples, the endometrial disorder is endometriosis, adenomyosis, Asherman’s syndrome or endometrial atrophy.

[0033] In another aspect, the present disclosure provides a method of producing an extracellular matrix from a tissue sample, the method comprising: treating the tissue sample with a surfactant to produce a decellularized tissue sample; rinsing the decellularized tissue sample with an aqueous solution to produce the extracellular matrix.

[0034] In another aspect, the present disclosure provides a method of producing an extracellular matrix hydrogel from a tissue sample, the method comprising: treating the tissue sample with a surfactant to produce a decellularized tissue sample; rinsing the decellularized tissue sample with an aqueous solution; lyophilizing or freeze-drying the decellularized tissue sample to form a frozen sample; crushing, grinding or milling the frozen sample to form a powder; and treating the powder with an acid and/or protease to form a hydrogel.

[0035] In some examples, the method further comprises supplementing the hydrogel with a crosslinker. The crosslinker may be genipin or N-(3-Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC).

[0036] The method may further comprise shaping or molding the hydrogel.

[0037] In some examples, the method further comprises supplementing the hydrogel with laminin.

[0038] In some examples, said treating the powder comprises: treating the powder with an acid and/or protease to form a pre-gel solution; and neutralizing the pre-gel solution with a base to form a hydrogel.

[0039] In some examples, the powder is treated with a protease. The powder may be treated with a protease under acidic conditions. The protease may be pepsin. The acid may be hydrochloric acid or acetic acid.

[0040] In some examples, the surfactant is SDS or SDC. The surfactant may be present at a concentration of between about 1% and 4%. In some examples, the surfactant is SDS present at a concentration of between about 1% and 4%, or SDC present at a concentration of about 4%.

[0041] In some examples, the hydrogel has a pH of between about 5 and 9, such as a pH of between about 6 and 8.

[0042] In some examples, the tissue sample is an endometrial tissue sample. In some examples, the endometrial tissue sample has been separated from myometrium tissue. In some examples, the endometrial tissue is not cancerous. In some examples, the tissue sample is obtained from a human or bovine subject. [0043] In some examples, the method further comprises manually shaping or moulding the hydrogel.

Brief description of the drawings

[0044] Figure 1. Decellularization protocol and assessment for endometrial tissue. (A) Overview of the steps involved in endometrial hydrogel preparation from the bovine endometrium. (B) Histological images of native and decellularized endometrial tissue sections (n=4) from the three protocols to check for the decellularization efficiency. Yellow arrowheads denote the presence of endometrial glands in the stromal compartment, which were also present in the decellularized tissue samples. SEM highlights remarkable preservation of both luminal (le) and glandular (eg) epithelial compartments in the decellularized tissues. SEM imaging of the lyophilized endometrium of P1 , P2 and P3 (n=3 biological replicates per group) showed a three-dimensional network of long ECM protein fibres without any intervening cellular material. (C) DNA quantification in native and decellularized endometrium (n=6 biological replicates per group) was normalized to the initial dry weight of each sample; ****p = <0.0001 , unpaired t test. Gel electrophoresis of DNA extracted from the fresh and decellularized endometrium for P1 , P2 and P3. Dry weight (gram) in native and decellularized endometrium tissue (n=4 biological replicates per group); *p = <0.05, unpaired Student’s t-test. SDS quantification showing P1 and P2 sample (n=3 biological replicates per group) after decellularization; Data represent mean ± SEM pg SDS per mg of dry tissue. (D) Immunostaining for fibronectin and hydroxyproline in the native and decellularized endometrium tissue (n=4 biological replicates per group), eg, glandular; le, luminal; s, stroma. Scale bars, 100 pm unless indicated otherwise.

[0045] Figure 2. Biochemical characterization of endometrium-derived extracellular matrix. (A, B) Confocal Raman micro-spectroscopic analysis of the native and decellularized endometrial tissue sections (n=4 replicates per group) represented as (A) high-resolution confocal Raman maps and (B) corresponding Raman spectra of the tissue sections. Raman signatures of the corresponding components of phenylalanine (Phe), glycosaminoglycan (GAG), asymmetric carbon-carbon stretching mode (C-C Asy), amide III, and collagen are labelled, while spectral intensities across samples have been normalized to the C-C asymmetric stretching mode. Superimposed spectra are shown at the top, and separated spectra are shown at the bottom. (C-F) Quantification of soluble collagen, insoluble collagen, hydroxyproline, and sulfated GAG (sGAG) in the native and decellularized endometrial tissue (n=3 replicates per group). Soluble and insoluble collagen, hydroxyproline and sGAG contents in the native and decellularized endometrium were normalized to the initial dry weight of the sample; ****p = <0.00001 ; ***p = <0.0001 ; **p = <0.001 ; * p = <0.05, unpaired Student’s t-test. (G) Fourier transform infrared spectroscopy (FTIR) spectra highlighting transmittance peaks of collagen amide A (~3300cm- 1), amide B (~3100cm-1), amide I (~1650cm-1), amide II (~1550cm-1), and amide III (~1200cm-1) in decellularized tissues belonging to all three groups. Scale bars, 100 pm unless indicated otherwise.

[0046] Figure 3. Mechanical properties of endometrium-derived extracellular matrix hydrogels. (A-C) Particle size analysis of the decellularized endometrial powder of P1 , P2 and P3 (n=4 biological replicates per group) by dynamic light scattering. (D) Gross and SEM images of endometrium-derived ECM hydrogels from the three protocols at concentrations of 10 mg/mL. (E) Turbidimetric gelation kinetics of the decellularized endometrium ECM hydrogel (n=3 biological replicates per group) at a concentration 10 mg/mL generated using three different protocols (P1 , P2 and P3). Water uptake capacity (F) and degradation (G) analysis of the three hydrogels (n=3 biological replicates per group). (H-L) Rheological properties of endometrial ECM hydrogels. Storage modulus, loss modulus, complex viscosity, and oscillation stress of hydrogels and matrigel (n=3 biological replicates per group); ****p = <0.00001 ; ***p = <0.0001 ; **p = <0.001 , one-way ANOVA for multiple group comparisons. (M) A representative gross image of the mouse uterine arm before and after decellularization. (Na-d) A device comprised a tuberculin syringe, a cotton bud, and a pipette tip to develop tubular structures from hydrogels and soft materials. A gross image of agarose tube developed using this device. (Ne-I) Gross images of tubular structures developed from different materials using our device. Scale bars, 100 pm unless indicated otherwise.

[0047] Figure 4. Proteomic profiling of decellularized endometrium. (A) Schematic representation of the proteomics workflow. Proteins were extracted from the respective P1 , P2 and P3 decellularized endometrium samples, digested and labelled with 4plex iTRAQ and combined (n = 4 pooled from each treatment group). To reduce sample complexity, pooled peptides were fractionated by high pH fractionation prior to LC-MS/MS analysis. (B) Hierarchical clustering analysis of 1141 proteins differentially expressed in decellularized endometrium ECM comparing between the three protocols P1 , P2 and P3. Proteomic analysis detected 25 and 28 ECM proteins (shown in the box) that were upregulated in the P3 group than in the P1 and P2 groups, respectively. (C) Dot plot represents the ECM protein expression profile for the core matrisome (encompassing ECM glycoproteins, collagens, and proteoglycans) and matrisome-associated (ECM-affiliated proteins, ECM regulators, and secreted factors) achieved from the three different methods (P1 , P2 and P3). The colour of each circle represents protein expression level, and the circle size indicates the relative value of the readout measurement across all conditions. (D) Western blot validating the expression of laminin, fibronectin and collagen I in endometrium decellularized using three protocols (n=3 biological replicates per group). GAPDH expression used as a loading control across the samples. ***p = <0.0001 ; **p = <0.001 , *p = <0.05, one-way ANOVA for multiple group comparisons. (E,F,G) Quantification of western blot band intensities relative to loading control (GAPDH).

[0048] Figure 5. Endometrium extracellular matrix-derived hydrogels support human and mouse organoid cultures. (Aa-d) Brightfield images of mouse endometrial organoids in Matrigel and endometrial hydrogels (P1 , P2, P3). (Ae) Compared to round-shaped organoids in Matrigel, organoids in P1 hydrogel showed budding (At), tubular (Ag) and glandular features (Ah). (Ai) The organoid forming efficiency of mouse endometrial organoids in Matrigel, P1 , P2 and P3 hydrogel; ****p = <0.00001 , two-way ANOVA with multiple comparison test. (Aj) Percentages of organoid numbers representing round, tubular and glandular-shaped organoids in Matrigel versus P1 ; ****p = <0.00001 , two-way ANOVA with multiple comparison test. (Ba-h) H&E and co-immunostaining for Foxa2, Ck8 and Ki67 of organoids cultured in Matrigel and P1 hydrogel. Percentages of Foxa2+ (Bi) and Ki67+ (Bj) positive cells in organoids grown in Matrigel versus P1 hydrogel. (Bk) Images of mouse endometrial organoid growth in P1 hydrogel supplemented with increased concentrations of laminin (L) (100P1 :0L, 75P1 :25L, 50P1 :50L, 25P1 :75L and 0P1 :100L). (Bl, Bm) Percentages of organoid formation efficiency and the number of round and branched-shaped organoids in P1 hydrogel containing different laminin concentrations; ****p = <0.00001 ; ***p = <0.0001 , two-way ANOVA with multiple comparison test. (C, D) Brightfield images and organoid formation efficiency of human endometrial cancer cells, Ishikawa, (Ca-d), normal endometrium (Ce-h), endometrial cancer (Ci-I), human colon (Cm-p) and mouse lung (Cq-t) organoids in P1 , P2, P3 hydrogel and Matrigel. ****p = <0.00001 , ***p = <0.0001 ; **p = <0.001 , *p = <0.05, two-way ANOVA with multiple comparison test. (E) Gross images of normal human endometrium before and after decellularization. (F) Brightfield images of human endometrial organoids cultured in human endometrial P3 hydrogel and Matrigel. (G) Organoid formation efficiency in human endometrial P3 hydrogel versus matrigel. (H) Venn diagram highlighting the number of common and differentially expressed proteins in the native tissue versus organoids grown in the P3 hydrogel and Matrigel. (I, J) Heatmap and violin plot analysis of 1300 proteins commonly identified between the three conditions. The MS intensities were normalized to Z-score. (K) A line plot analysis displaying that organoids cultured in the P3 hydrogel had a similar protein expression profile with the native human tissue from which they are derived than the organoid grown in the Matrigel. (L) Ingenuity pathway analysis (IPA) highlighted the major signaling pathways were either downregulated or missing in the organoids cultured in Matrigel relative to the P3 hydrogel organoids and the native tissue. Z-scores were used to predict activation (z-score > 2; orange) or inhibition (z-score < - 2; blue) of each function. Scale bars, 100 pm unless indicated otherwise. (M) Heat map depicting similarities between human and bovine hydrogels.

[0049] Figure 6. Generation of extracellular matrix hydrogel-derived porous scaffolds for culturing human endometrial cells. (A) Schematic workflow for fabricating porous scaffolds from bovine P3 hydrogel by using lyophilization procedure. (B) The gross view of the P3 scaffolds generated for 96-, 24- and 12-well culture plates can be stored at room temperature. (C-E) SEM and H&E images confirmed the honeycomb-like porous appearance of scaffolds. (F, G) Gross images of scaffolds seeded with endometrial cells (Ishikawa). (H-l) Histology analysis showed that these scaffolds support the growth of organoids. (J) Immunolocalization of phospho-histone 3 (PH3, a marker of proliferating cells) and cytokeratin 8 (CK8, a maker of epithelial cells) detected the presence of proliferating epithelial cells in these scaffolds. Scale bars, 100 pm unless indicated otherwise.

[0050] Figure 7. Gross images of mouse endometrial tissue highlighting distinct luminal and glandular compartments. Bars: 100 pm.

[0051] Figure 8. MS chromatogram traces for laminin isolated from the bovine endometrium. Single MS chromatogram of peptide from laminin at m/z of (A) 415.23 Da, (B) 637 Da, (C) 610.65 Da and (D) 444.25 Da, respectively, of bovine endometrial-derived laminin versus laminin purchased from Sigma (standard control). Retention time (RT) is a measure of the time taken for a solute to pass through a chromatography column. It is calculated as the time from injection to detection.

[0052] Figure 9. Gross and H&E-stained images of cultured primary human endometrial cells on the P3 hydrogel-derived scaffolds indicating that normal patient cells can grow on these scaffolds. Bars: 100 pm. Detailed description

Definitions

[0053] In the context of this specification, the terms "a" and "an" are used herein to refer to one or to more than one (i.e. to at least one) of the grammatical object of the article. By way of example, "an element" means one element or more than one element.

[0054] The term "about" is understood to refer to a range of +/- 10%, preferably +/- 5% or +/- 1% or, more preferably, +/- 0.1%.

[0055] The terms "comprise", "comprises", "comprised" or "comprising", "including" or "having" and the like in the present specification and claims are used in an inclusive sense, ie, to specify the presence of the stated features but not preclude the presence of additional or further features.

[0056] The term “stem cell” refers to a subset of progenitor cells that have the capacity or potential, under particular circumstances, to differentiate into a more specialised cell type, and which retains the capacity, under certain circumstances, to proliferate without substantially differentiating. In that regard, stem cells are generally capable of self-renewal.

[0057] Where numerical ranges are used to describe certain embodiments of the present disclosure, it will be understood that each range should be considered to encompass subranges therein. For example, the description of a range such as from 1 to 6 should be considered to include subranges such as from 1 to 5, from 2 to 4, from 2 to 6 and so on. Likewise, the description of a range of between 1 and 6 should be considered to include subranges such as between 2 and 5, between 1 and 3, between 3 and 6 and so on.

[0058] Unless specifically defined otherwise, all technical and scientific terms used herein shall be taken to have the same meaning as commonly understood by one of ordinary skill in the art (for example, in cell culture, biochemistry, protein chemistry and biochemistry).

Decellularisation

[0059] The present disclosure provides ECM obtained from a decellularized tissue sample. In some examples, the present disclosure provides an ECM hydrogel obtained from a decellularized tissue. The tissue sample may comprise endometrial cells, gastrointestinal cells, skin cells, lymph cells, lung cells, hepatocytes, liver cells, pancreatic cells, colon cells, cardiac cells, muscle cells, oesophageal cells, tracheal cells, bladder cells, muscle cells, arterial cells, nerve cells, umbilical cord cells, vaginal cells or stomach cells. The cells may, for example, be human, bovine, porcine, avian, rat, goat or murine cells. The tissue is preferably healthy tissue in that it is not cancerous or otherwise diseased.

[0060] To prepare an ECM or ECM hydrogel, the tissue sample may be cut into pieces and washed in water, optionally comprising an antibiotic such as penicillin/streptomycin, gentamycin, kanamycin, nystatin, primocin, normocin and/or amphotericin. The tissue sample and water may be agitated or stirred at room temperature for between 1 hour and 48 hours, such as between about 5 hours and 48 hours, or between about 5 hours and 24 hours, or between about 7 hours and 24 hours, or between about 10 hours and 22 hours, or between about 10 hours and 20 hours. Preferably the water is replaced more than once during the washing such as every 3 to 4 hours. [0061] The washed tissue is preferably decellularized by treating the tissue with a surfactant. The surfactant may be, for example, Tween, Triton-X, sodium dodecyl sulfate (SDS) or sodium deoxycholate (SDC). Preferably, the surfactant is SDC. The surfactant may be present at a concentration of between about 0.1% and 10%, such as between about 0.2% and 9%, or between about 0.3% or 8%, or between about 0.4% and 7%, or between about 0.5% and 6%, or between about 0.6% and 5%, or between about 0.7% and 4.5%, or between about 0.8% and 4%, or between about 0.9% and 4%, or between about 1% and 4% such as about 1%, about 1.5%, about 2%, about 2.5%, about 3%, about 3.5% or about 4%. In some examples the surfactant is SDS present at a concentration of about 1%, or about 1 .5%, about 2%, about 2.5%, about 3%, about 3.5% or about 4%. In some examples, the surfactant is SDC present at a concentration of about 1%, or about 1 .5%, about 2%, about 2.5%, about 3%, about 3.5% or about 4%.

[0062] The tissue sample may be treated with the surfactant at room temperature for between about 1 day and 10 days, or until the tissue appears translucent. In some examples, the tissue is incubated for about 1 day, about 2 days, about 3 days, about 4 days, about 5 days, about 6 days, about 7 days, about 8 days, about 9 days or about 10 days. Preferably, the tissue sample is treated for about 4 or 5 days. The surfactant may be replaced with fresh surfactant one or more times during the treatment. For example, the surfactant may be replaced with fresh surfactant every 3 to 4 hours. The tissue is preferably agitated or stirred during the surfactant treatment.

[0063] Following treatment with the surfactant, the tissue may be rinsed with water for between about 1 hour and 48 hours, such between about 12 hours and 24 hours, and then treated with a DNase. The DNase may be present in a salt solution such as 1 M sodium chloride.

[0064] Following treatment with the surfactant, and optionally with a DNase, the tissue is preferably washed with water one or more times to remove residual surfactant. The decellularized tissue may be washed with water for between 1 hour and 1 week, such as between about 6 hours and 6 days, or between about 12 hours and 5 days, or between about 1 day and 4 days, or between about 1 day and 3 days, or between about 1 day and 2 days, such as for about 1 day or about 2 days. After washing in water, the surfactant is preferably absent or present at less than about 1 pg per mg of dry tissue, such as between about 0.9 pg and 0.01 pg, or between about 0.8 pg and 0.01 pg, or between about 0.7 pg and 0.01 pg, or between about 0.6 pg and 0.01 pg, or between about 0.5 pg and 0.01 pg, or between about 0.4 pg and 0.01 pg, or between about 0.3 pg and 0.01 pg, or between about 0.2 pg and 0.01 pg, or between about 0.2 pg and 0.05 pg or between about 0.2 pg and 0.1 pg per mg of dry tissue.

[0065] In native tissue, the different ECM components provide structural support and transmit functional signals to resident cells. It may, therefore, be desirable to maintain the native ECM proteins after decellularization in order to replicate the native tissue ECM microenvironment and signaling. In that context, the decellularized tissue described herein preferably comprises soluble collagen at a concentration of between about 5 pg/mg and 40 pg/mg, such as between about 5 pg/mg and 35 pg/mg, or between about 5 pg/mg and 30 pg/mg or between about 10 pg/mg and 30 pg/mg. In some examples, insoluble collagen is present in the decellularized tissue of the present disclosure at a concentration of between about 10 pg/mg and 40 pg/mg, such as between about 15 pg/mg and 35 pg/mg, or between about 15 pg/mg and 30 pg/mg, or between about 15 pg/mg and 20 pg/mg, or between about 25 pg/mg and 30 pg/mg, or between about 30 pg/mg and 35 pg/mg. In some examples, hydroxyproline is present in the decellularized tissue of the present disclosure at a concentration of between about 0.05 pg/mg and 0.3 pg/mg, or between about 0.1 pg/mg and 0.3 pg/mg, or between about 0.1 pg/mg and 0.25 pg/mg, or between about 0.3 pg/mg and 0.6 pg/mg, or between about 0.3 pg/mg and 0.5 pg/mg. In some examples, sulphated glycosaminoglycans (sGAG) are present in the decellularized tissue of the present disclosure at a concentration of between about 1 pg/mg and 3 pg/mg, or between about 1 .5 pg/mg and 2.5 pg/mg.

Hydrogels

[0066] The decellularized tissue may then be lyophilised or freeze-dried and milled to form a powder. The powder may comprise particles having an average diameter of between about 300 nm and 700 nm, such as between about 350 nm and 650 nm, or between about 350 nm and 600 nm, or between about 400 nm and 600 nm, or between about 400 nm and 575 nm, or between about 425 nm and 575 nm.

[0067] The powder may be digested with an acid protease such as pepsin and/or an acid such as hydrochloric acid or acetic acid. For example, the powder may be treated with a protease such as pepsin and/or an acid such as hydrochloric acid or acetic acid for between about 1 hour and 100 hours, or between about 10 hours and 100 hours, or between about 24 hours and 100 hours, or between about 48 hours and 84 hours, or between about 60 hours and 84 hours, such as about 72 hours to produce a pre-gel solution. The pre-gel solution may be used directly in the formation of a scaffold or neutralised with a base such as sodium hydroxide to have a pH of between about 7 and 8 such as about 7.5. Accordingly, it will be understood that neutralising in this context refers to increasing the pH so as to bring it closer to 7 but not necessarily precisely to 7. In some examples, the pre-gel may be neutralised to form a hydrogel having a pH of between about 5 and 9, such as between about 5.5 and 9, or between about 5.5 and 8.5, or between about 6 and 8.5, or between about 6 and 8, or between about 6.5 and 8, or between about 6.5 and 7.5, such as about 7. Following neutralisation, the pre-gel may be maintained at a cool temperature such as less than 10°C (eg, about 4°C) or a gel may be formed by bringing the pre-gel to room temperature. In some applications, for example, the pre-gel is neutralised and stored in a refrigerator.

[0068] In some examples, the decellularized ECM powder is digested with pepsin and HCI for between about 12 hours and 100 hours, such as between about 24 hours and 84 hours, or between about 36 hours and 84 hours, or between about 48 hours and 84 hours, or between about 60 hours and 84 hours, such as about 66 hours or about 72 hours or about 78 hours. The digested samples may then be equilibrated in a buffer such as PBS to produce a pre-gel. The pre-gel may be neutralised with, for example, NaOH. In some examples, the decellularized ECM powder is dissolved in acetic acid, followed by neutralisation with NaOH. The solubilized ECM powder may have a pH of between about 1 and 6, such as between about 2 and 6 or between about 3 and 6 or between about 3 and 5, such as about 3.5 or about 4 or about 4.5.

[0069] Alternative methods for ECM digestion include an extraction process to solubilize and form an ECM hydrogel from soft tissue. Proteins and glycoproteins may be extracted using a homogenization process involving pestle and mortar or high-speed shear mixed within a high salt buffer that physically disrupts the ECM particles and collagen fiber structure at physiologic pH. Homogenization may involve a dispase enzymatic step that cleaves fibronectin, collagen IV, and collagen I and digests the ECM, a urea extraction step which further disrupts the non-covalent bonding and increases the solubility of the ECM proteins, and centrifugation that removes residual non-soluble ECM components. The resulting solubilized extracts form an ECM hydrogel when increasing the temperature of the extract to about 37°C or by decreasing the pH to about 4 using an acid such as acetic acid or HCI.

[0070] The hydrogel may have a water uptake capacity of about 30% or greater after 3 hours hydration time, such as between about 30% and 100% or between about 40% and 100%. In some examples, the hydrogel has a water uptake capacity of about 50% or greater after about 4 hours hydration time. The hydrogel may have a storage modulus G’ at 37°C of between about 100 Pa and 600 Pa, such as between about 200 Pa and 600 Pa, or between about 300 Pa and 600 Pa, or between about 300 Pa and 500 Pa, or between about 300 Pa and 400 Pa. The hydrogel may have a complex viscosity at 37°C of between about 100 Pa.S and 700 Pa.S, such as between about 200 Pa.S and 600 Pa.S, or between about 300 Pa.S and 600 Pa.S, or between about 300 Pa.S and 500 Pa.S, or between about 300 Pa.S and 400 Pa.S. The hydrogel may have an oscillation stress at 37°C of between about 1 Pa and 6 Pa, such as between about 2 Pa and 6 Pa, or between about 3 Pa and 6 Pa, or between about 3 Pa and 5 Pa, or between about 3 Pa and 4 Pa.

[0071] In some examples, the present disclosure provides an extracellular matrix obtained from decellularized healthy endometrial tissue. In some examples, the present disclosure provides an extracellular matrix obtained from decellularized endometrial tissue, wherein the endometrial tissue is not cancerous. In some examples, the present disclosure provides an extracellular matrix obtained by treating endometrial tissue with a surfactant selected from sodium dodecyl sulfate and sodium deoxycholate, wherein the endometrial tissue is not cancerous. In some examples, the present disclosure provides an extracellular matrix obtained by treating endometrial tissue with sodium deoxycholate, wherein the endometrial tissue is not cancerous, and wherein the sodium deoxycholate is present at a concentration of about 4%. In some examples, the present disclosure provides an extracellular matrix obtained by treating endometrial tissue with sodium deoxycholate for about 4 or 5 days, wherein the endometrial tissue is not cancerous, and wherein the sodium deoxycholate is present at a concentration of about 4%.

[0072] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating a tissue sample with a surfactant to produce a decellularized tissue sample; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0073] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with a surfactant to produce a decellularized tissue sample; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0074] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with a surfactant selected from sodium dodecyl sulfate and sodium deoxycholate to produce a decellularized tissue sample; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0075] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate to produce a decellularized tissue sample; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0076] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0077] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0078] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; washing the decellularized tissue sample with water; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0079] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; washing the decellularized tissue sample with water; treating the decellularized tissue sample with a DNase; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0080] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; washing the decellularized tissue sample with water; treating the decellularized tissue sample with a DNase; washing the decellularized tissue sample water; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0081] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; washing the decellularized tissue sample with water; treating the decellularized tissue sample with a DNase; washing the decellularized tissue sample water; lyophilising and milling the decellularized tissue sample to produce a powder; digesting the powder with pepsin and hydrochloric acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel. [0082] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate for about 4 or 5 days to produce a decellularized tissue sample, wherein the sodium deoxycholate is present at a concentration of about 4%; washing the decellularized tissue sample with water for between about 12 hours and 24 hours; treating the decellularized tissue sample with a DNase; washing the decellularized tissue sample water for about 1 or 2 days; lyophilising and milling the decellularized tissue sample to produce a powder; digesting the powder with pepsin and hydrochloric acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0083] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with a surfactant to produce a decellularized tissue sample, wherein the endometrial tissue has been separated from myometrium tissue; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0084] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate to produce a decellularized tissue sample, wherein the endometrial tissue has been separated from myometrium tissue; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[0085] In some examples, the present disclosure provides a method of producing an extracellular matrix scaffold from a tissue sample, the method comprising: treating the tissue sample with a surfactant to produce a decellularized tissue sample; rinsing the decellularized tissue sample with an aqueous solution to produce the extracellular matrix; lyophilizing or freeze-drying the decellularized tissue sample to form a frozen sample; crushing, grinding or milling the frozen sample to form a powder; and treating the powder with an acid to form a pre-gel; freezing the pre-gel to form a scaffold.

[0086] In some examples, the present disclosure provides a method of producing an extracellular matrix hydrogel comprising: treating an endometrial tissue sample with sodium deoxycholate to produce a decellularized tissue sample, wherein the endometrial tissue has been separated from myometrium tissue, and wherein the sodium deoxycholate is present at a concentration of about 4%; lyophilising and milling the decellularized tissue sample to produce a powder; treating the powder with an acid to produce a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

Organoids and uses

[0087] Organoids are self-organising three-dimensional in vitro tissue models that recapitulate many of the physiologically relevant features of the normal or diseased tissue from which they are derived. Organoids can capture different stages of tumourigenesis and cancer genomic subtypes, and they can be used to uncover biological changes underlying metastatic progression. Importantly, organoid cultures can be subjected to a wide range of functional assays, and they can be rapidly tested for their sensitivity to novel drugs, drug combinations, and drugs given in defined temporal sequences. Organoid model systems can be expanded to contain stromal or immune cells, with the latter particularly pertinent to the development and in vitro testing of new immunotherapies (Pauli, C., et al., (2017) Cancer Discov, 7(5): p. 462-477; Drost, J. and H. Clevers, (2018) Nat Rev Cancer, 18(7): p, 407-418). Organoids also have applications in the study of organogenesis, disease modelling, toxicity assays, and regenerative medicine. For example, genome editing of organoid cultures can allow evaluation of patient-specific mutations found in certain cancers. Other applications include organoid co-culture with pathogenic bacteria or viruses to study infections, the use of organoids to propagate intracellular pathogens, or to produce biological products.

[0088] In some examples, the present disclosure provides organoids obtained by culturing cells in the presence of an ECM or a hydrogel of the present disclosure. The cells may be incubated in a solution comprising about 50% medium and about 50% hydrogel, or about 40% medium and about 60% hydrogel, or about 30% medium and about 70% hydrogel, or about 20% medium and about 80% hydrogel, or about 10% medium and about 90% hydrogel. The cells may be endometrial cells, gastrointestinal cells, skin cells, lymph cells, lung cells, hepatocytes, liver cells, pancreatic cells, colon cells, cardiac cells, muscle cells, oesophageal cells, tracheal cells, bladder cells, muscle cells, arterial cells, nerve cells, umbilical cord cells, vaginal cells or stomach cells. The organoid may be a pancreatic organoid, an endometrial organoid, a gastrointestinal organoid, a muscle organoid, a heart organoid, a lung organoid, a colon organoid, a uterine organoid, a skin organoid, an ovarian organoid, a fallopian organoid, a liver organoid or a muscle organoid. Other cell and organoid types will be apparent to the skilled addressee.

[0089] The organoid may be used, for example, in drug screening, modelling a disorder, designing a regenerative therapy or studying disease progression. For example, the organoid may be used to study endometrial pathologies such as endometriosis, which involves the growth of endometrial tissue outside the uterine cavity, adenomyosis, the presence of endometrial glands and stroma within the myometrium, Asherman’s syndrome, the presence of intrauterine and/or intracervical adhesions, or endometrial atrophy, characterised by a nonproliferating endometrium. [0090] The histoarchitecture of the uterus is relatively similar across species from viviparous lizards to primates and includes three major layers: an outer covering, the serosa; a middle muscular layer, the myometrium; and an inner mucosa layer, the endometrium (J. E. Girling, The reptilian oviduct: a review of structure and function and directions for future research. J Exp Zoo! 293, 141-170 (2002)); (A. T. Major, M. A. Estermann, Z. Y. Roly, C. A. Smith, An evo-devo perspective of the female reproductive tract. Biol Reprod 10.1093/biolre/ioab166 (2021)). The endometrium is a complex layer that lies nearest to the uterine lumen and is further subdivided into epithelial and stromal compartments (J. Goad, Y. A. Ko, M. Kumar, S. M. Syed, P. S. Tanwar, Differential Wnt signaling activity limits epithelial gland development to the anti-mesometrial side of the mouse uterus. Dev Biol 423, 138-151 (2017)). The epithelium contains luminal and glandular epithelium. The endometrium is a highly hormonally responsive tissue which undergoes remodeling primarily due to cyclical changes in ovarian hormones (A. M. Kelleher, F. J. DeMayo, T. E. Spencer, Uterine Glands: Developmental Biology and Functional Roles in Pregnancy. Endocr Rev 40, 1424-1445 (2019)). The endometrium undergoes expansion under the influence of estrogen during the proliferative phase of the menstrual cycle and it acquires a more secretory phenotype during the second half of the cycle due to the rising levels of progesterone to support the embryonic development (H. O. D. Critchley, J. A. Maybin, G. M. Armstrong, A. R. W. Williams, Physiology of the Endometrium and Regulation of Menstruation. Physiol Rev 100, 1149-1179 (2020)). If embryonic implantation and subsequent pregnancy are not established, then the endometrium undergoes breakdown and sloughs off during menstruation. Endometrial repair ensues immediately after menses leading to the scarless repair of the endometrium and the tissue get ready for the next menstrual cycle. The process of endometrial repair is led by the endometrial stem/progenitor cells located toward the base of endometrial glands in the basal layer of the endometrium (S. M. Syed et al., Endometrial Axin2(+) Cells Drive Epithelial Homeostasis, Regeneration, and Cancer following Oncogenic Transformation. Cell Stem Cell 26, 64-80 e13 (2020)). In most organ systems, active Wnt signaling marks stem/progenitor cells that are responsible for their maintenance and repair (H. Clevers, K. M. Loh, R. Nusse, Stem cell signaling. An integral program for tissue renewal and regeneration: Wnt signaling and stem cell control. Science 346, 1248012 (2014)). High Wnt signaling activity is observed in the uterine glandular epithelium, and the loss of Wnt signaling in uterus the compromises its development and functions (B. A. Parr, A. P. McMahon, Sexually dimorphic development of the mammalian reproductive tract requires Wnt-7a. Nature 395, 707-710 (1998)). Cell lineage tracing studies in mice have identified Lgr5 and Axin2, two well-known Wnt signaling targets, as a marker of a subset of glandular epithelial cells that act as endometrial stem/progenitor cells (R. Seishima et al., Neonatal Wnt-dependent Lgr5 positive stem cells are essential for uterine gland development. Nat Commun 10, 5378 (2019)). Under high Wnt culture conditions, endometrial stem/progenitor cells form endometrial organoids. Endometrial organoids have been developed from normal and abnormal human endometrium. These organoids replicate many endometrial physiological functions, including histological and secretory changes associated with cyclical hormonal variations, and have already provided unique insights into the pathogenesis of human endometrial diseases, such as endometriosis, adenomyosis, and endometrial cancer. The ECM hydrogels of the present disclosure support the growth of both human and mouse endometrial organoids, which was comparable to their growth in Matrigel. The proteome of organoids cultured in the ECM hydrogel of the present disclosure was more similar to the native tissue from which they were derived than Matrigel-grown organoids.

[0091] In some examples, the present disclosure provides an organoid that may be grafted into a subject to promote repair of an injured organ. In some examples, the present disclosure provides a method of repairing a damaged organ comprising applying an organoid of the present disclosure to the damaged organ. In some examples, the present disclosure provides a method of repairing a damaged organ comprising applying a hydrogel of the present disclosure to the damaged organ. In some examples, the present disclosure provides a method of repairing a damaged organ comprising applying an organoid and a hydrogel of the present disclosure to the damaged organ.

[0092] The hydrogel may be delivered directly or indirectly to the site of an injury. For example, minimally invasive delivery may involve injection with a pre-gel viscous fluid using a catheter or syringe. The pre-gel may then polymerise at physiologic temperature into a hydrogel which conforms to the shape of the injury or defect. Alternatively, the hydrogel may be delivered directly to the site of the injury or defect.

[0093] The tissue damage to be repaired may be caused by an injury such as a burn, a disease or a lesion. The damaged organ may be a female reproductive organ such as the vagina, endometrium, uterus, fallopian tube, cervix etc. For example, the organ damage may be due to cancer such as endometrial or ovarian cancer, or a lesion caused by treating (e.g., excising) the cancer. Endometrial cancer is a heterogenic constellation of diseases for which etiopathogenesis has historically been dichotomized into two groups based on clinico-histological characteristics. Type I tumors were postulated to be estrogen-mediated, well-to-moderately differentiated endometrioid lesions on a background of juxtaposed hyperplasia in younger women. Type II endometrial cancer referred to poorly differentiated tumors of endometrioid or non-endometrioid histology arising in a milieu of endometrial atrophy and were thought to be estrogen-independent. Lynch syndrome can result in endometrial cancers of both types as well as a multitude of extra-uterine malignancies.

[0094] In some examples, the present disclosure provides a method of treating an endometrial or gynaecological disorder or infertility in a subject, the method comprising administering to the subject a hydrogel and/or an organoid of the present disclosure. The disorder may, for example, be endometriosis, adenomyosis, Asherman’s syndrome or endometrial atrophy. In some examples, the infertility is absolute uterine factor infertility. In some examples, the infertility is Mayer-Rokitansky- Kuster-Hauser syndrome, uterine hypoplasia or uterine malformation. In some examples, the infertility is caused by hysterectomy, for example, due to malignant uterine tumor, benign diseases (including leiomyoma and adenomyosis), postpartum hemorrhage, and loss of fertility due to intrauterine adhesions (Asherman’s syndrome).

[0095] In some examples, the organ is a heart. The hydrogel and/or organoid of the present disclosure may be used to treat a heart disorder such as ischemic injury. In some examples, the organ is a brain. The hydrogel and/or organoid of the present disclosure may be used to treat a brain disorder such as traumatic brain injury. [0096] The hydrogel of the present disclosure may be shaped or moulded to suit a particular purpose, such as a medical use. For example, the hydrogel may be moulded into a tubular shape for use in repairing a tubular organ. As described further below, a tubular structure may be formed using a syringe and a sterile cotton bud. The hydrogel may be supplemented with a cross-linker such as genipin or N-(3-Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC) to enhance the stability of the hydrogel and assist in shaping or moulding the hydrogel. Those skilled in the art will understand that other cross-linkers may be used to assist in shaping or moulding the hydrogel.

[0097] In some examples, the hydrogel of the present disclosure is absorbed into, adsorbed onto, or otherwise dispersed onto or into a biocompatible substrate. Non-limiting examples of a biocompatible substrate include: a mesh, a non-woven, decellularized tissue, a polymer composition, a polymeric structure, a cell growth scaffold, an implant, an orthopedic implant, and intraocular lens, sutures, intravascular implants, stents, and transplants. The compositions described herein may be applied to or incorporated into, by any suitable method, a non-woven material, such as a bandage, a suture, an implant, such as a ceramic, metal, or polymeric implant, for example a prosthesis, artificial or otherwise-modified vessel, a valve, an intraocular lens, or a tissue implant. In some examples, the hydrogel is coated onto a biocompatible structural material, such as a metal, an inorganic calcium compound such as calcium hydroxide, calcium phosphate or calcium carbonate, or a ceramic composition.

Items of the present disclosure

[0098] Set forth below are non-limiting Items of the present disclosure.

[0099] Item 1 . An extracellular matrix hydrogel obtained from a decellularized healthy tissue.

[00100] Item 2. The hydrogel of Item 1 wherein the tissue is an endometrial tissue.

[00101] Item 3. An extracellular matrix hydrogel obtained from a decellularized endometrial tissue.

[00102] Item 4. The hydrogel of Item 2 or Item 3 wherein the endometrial tissue has been separated from myometrium tissue.

[00103] Item 5. The hydrogel of any one of Items 2 to 4 wherein the endometrial tissue is not cancerous.

[00104] Item 6. The hydrogel of any one of Items 1 to 5 wherein the tissue is obtained from a human or bovine subject.

[00105] Item 7. The hydrogel of any one of Items 1 to 6 wherein the tissue is decellularized by treating the tissue with a surfactant selected from sodium dodecyl sulfate (SDS) and sodium deoxycholate (SDC).

[00106] Item 8. The hydrogel of Item 7 wherein the surfactant is present at a concentration of between about 1% and 4%.

[00107] Item 9. The hydrogel of Item 8 wherein the surfactant is present at a concentration of about 4%.

[00108] Item 10. The hydrogel of any one of Items 7 to 9 wherein the surfactant is SDC. [00109] Item 11. The hydrogel of any one of Items 1 to 10 wherein the hydrogel is supplemented with a cross-linker.

[00110] Item 12. The hydrogel of Item 11 wherein the cross-linker is genipin or N-(3- Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC).

[00111] Item 13. The hydrogel of any one of Items 1 to 12 wherein the hydrogel is manually shaped or moulded.

[00112] Item 14. The hydrogel of Item 13 wherein the hydrogel is in the form of a tube or lumen.

[00113] Item 15. The hydrogel of any one of Items 1 to 14 wherein the hydrogel is supplemented with laminin.

[00114] Item 16. The hydrogel of any one of Items 1 to 15 wherein the hydrogel is obtained by: lyophilizing or freeze-drying the decellularized tissue sample to form a frozen sample; crushing, grinding or milling the frozen sample to form a powder; treating the powder with an acid and/or protease to form a pre-gel; and neutralising the pre-gel with a base to produce a hydrogel.

[00115] Item 17. The hydrogel of any one of Items 1 to 16 wherein the hydrogel has a pH of between about 5 and 9.

[00116] Item 18. The hydrogel of any one of Items 1 to 16 wherein the hydrogel has a pH of between about 6 and 8.

[00117] Item 19. A scaffold formed from a hydrogel of any one of Items 1 to 18.

[00118] Item 20. The scaffold of Item 19 wherein the scaffold is porous.

[00119] Item 21. The scaffold of Item 19 or Item 20 wherein the scaffold is formed by ice templating in which the hydrogel is frozen and lyophilised to produce a porous scaffold.

[00120] Item 22. The scaffold of any one of Items 19 to 21 for use in cell culture.

[00121] Item 23. The scaffold of any one of Items 19 to 21 for use in tissue regeneration.

[00122] Item 24. An organoid obtained by culturing cells on a hydrogel of any one of Items 1 to 18.

[00123] Item 25. The organoid of Item 24 wherein the organoid is an endometrial organoid.

[00124] Item 26. The organoid of Item 24 or Item 25 wherein the cells are endometrial cells.

[00125] Item 27. The organoid of Item 24 wherein the organoid is an organoid of a tubular organ.

[00126] Item 28. The organoid of Item 27 wherein the tubular organ is a colon, a lung or a gastrointestinal organ.

[00127] Item 29. A method of promoting repair of a wound comprising applying to the wound a hydrogel of any one of Items 1 to 18, a scaffold of any one of Items 19 to 23 or an organoid of any one of Items 24 to 28.

[00128] Item 30. The method of Item 29 wherein the wound is an endometrial wound, a uterine wound, a fallopian wound, an ovarian wound, a vaginal wound, a colon wound, a lung wound, a wound of a gastrointestinal organ, a flesh wound, a liver wound or a cardiac wound. [00129] Item 31. A method of promoting tissue regeneration comprising applying to the tissue a hydrogel of any one of Items 1 to 18, a scaffold of any one of Items 19 to 23 or an organoid of any one of Items 24 to 28.

[00130] Item 32. A method of treating an endometrial disorder in a subject comprising administering to the subject a hydrogel of any one of Items 1 to 18, a scaffold of any one of Items 19 to 23 or an organoid of any one of Items 24 to 28.

[00131] Item 33. The method of Item 32 wherein the endometrial disorder is endometriosis, adenomyosis, Asherman’s syndrome or endometrial atrophy.

[00132] Item 34. A method of producing an extracellular matrix from a tissue sample, the method comprising: treating the tissue sample with a surfactant to produce a decellularized tissue sample; and rinsing the decellularized tissue sample with an aqueous solution to produce the extracellular matrix.

[00133] Item 35. A method of producing an extracellular matrix hydrogel from a tissue sample, the method comprising: treating the tissue sample with a surfactant to produce a decellularized tissue sample; rinsing the decellularized tissue sample with an aqueous solution; lyophilizing or freeze-drying the decellularized tissue sample to form a frozen sample; crushing, grinding or milling the frozen sample to form a powder; and treating the powder with an acid and/or protease to form a hydrogel.

[00134] Item 36. The method of Item 35 further comprising supplementing the hydrogel with a crosslinker.

[00135] Item 37. The method of Item 36 wherein the cross-linker is genipin or N-(3- Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC).

[00136] Item 38. The method of any one of Items 35 to 37 further comprising shaping or molding the hydrogel.

[00137] Item 39. The method of any one of Items 35 to 38 further comprising supplementing the hydrogel with laminin.

[00138] Item 40. The method of any one of Items 35 to 39 wherein treating the powder comprises: treating the powder with an acid and/or protease to form a pre-gel solution; and neutralising the pre-gel solution with a base to form a hydrogel.

[00139] Item 41. The method of any one of Items 35 to 40 wherein the powder is treated with a protease.

[00140] Item 42. The method of Item 41 wherein the powder is treated with a protease under acidic conditions.

[00141] Item 43. The method of any one of Items 35 to 42 wherein the protease is pepsin.

[00142] Item 44. The method of any one of Items 35 to 43 wherein the acid is hydrochloric acid or acetic acid.

[00143] Item 45. The method of any one of Items 35 to 44 wherein the hydrogel has a pH of between about 5 and 9. [00144] Item 46. The method of any one of Items 35 to 45 wherein the hydrogel has a pH of between about 6 and 8.

[00145] Item 47. The method of any one of Items 34 to 46 wherein the surfactant is SDS or SDC.

[00146] Item 48. The method of any one of Items 34 to 47 wherein the surfactant is present at a concentration of between about 1% and 4%.

[00147] Item 49. The method of any one of Items 34 to 48 wherein the surfactant is SDS present at a concentration of between about 1% and 4%, or SDC present at a concentration of about 4%.

[00148] Item 50. The method of any one of Items 34 to 49 wherein the tissue sample is an endometrial tissue sample.

[00149] Item 51 . The method of any one of Items 34 to 50 wherein the endometrial tissue sample has been separated from myometrium tissue.

[00150] Item 52. The method of any one of Items 34 to 51 wherein the endometrial tissue is not cancerous.

[00151] Item 53. The method of any one of Items 34 to 52 wherein the tissue sample is obtained from a human or bovine subject.

[00152] Item 54. The method of any one of Items 35 to 53 further comprising manually shaping or moulding the hydrogel.

Examples

Results

Decellularisation of endometrial tissue

[00153] Bovine endometrium bears many histological and biochemical similarities to human endometrium (M. A. Abedal-Majed, A. S. Cupp, Livestock animals to study infertility in women. Anim Front 9, 28-33 (2019); C. A. Gray et al., Developmental biology of uterine glands. Biol Reprod 65, 1311-1323 (2001)). Additionally, the whole bovine uterus is readily available from abattoirs. Tissue extraction, decellularization and hydrogel formation was performed using bovine uterus samples broadly in five stages: 1) Surgical isolation of the endometrium; 2) Removal of cellular and nuclear material (decellularization); 3) Lyophilization or freeze-drying; 4) Low temperature milling to make powder; and 5) Digestion, pH neutralisation and gelation (Fig. 1A). For decellularization, three different detergent (i.e., surfactant) treatments were used, namely 4% sodium dodecyl sulfate (SDS; hereafter referred as P1), 1% SDS (P2), and 4% sodium deoxycholate (SDC; P3) (Fig. 1 B). Surfactant treatments led to white translucency of tissues while maintaining their gross appearance and size (Fig. 1A). The decellularized tissues obtained from the three protocols were then evaluated using histology and scanning electron microscopy (SEM) to check for the decellularization efficiency before progressing to the next step (Fig. 1 Ba-i). Examination of hematoxylin and eosin (H&E) stained native and decellularized endometrial tissue sections revealed the loss of nuclear material (hematoxylin) with no apparent loss in tissue morphology (Fig. 1 Ba-f). [00154] Endometrial glands are normally present in the stromal compartment and these glands were also readily identifiable in the decellularized tissue samples (yellow arrowheads; Fig. 1 Bd-f). SEM analysis confirmed the preservation of both luminal (le) and glandular (eg) epithelial compartments in the decellularised tissues (Fig. 1 Bg-i). Next, the decellularized endometrium was lyophilized (Fig. 1A, 1 B). Scanning electron microscopy (SEM) imaging of lyophilized endometrium revealed the presence of a three-dimensional network of long ECM protein fibers without intervening cellular material (Fig. 1 Bj-o). The amount of DNA in the fresh endometrium tissue was significantly reduced after decellularization (p<0.000; Fig. 1Ca), which was further confirmed with gel electrophoresis methodology, revealing a very low content of residual DNA in the decellularized endometrium (Fig 1Cb). Dry weight of the decellularized endometrium was significantly reduced compared to the fresh native tissue (p<0.05; Fig 1Cc). Residual SDS content in the P1 and P2 group was low, indicating a successful elimination of cytotoxic SDS in the samples (Fig 1Cd). To monitor if the decellularized endometrium preserve essential ECM proteins, such as fibronectin and collagen/hydroxyproline, immunostaining was performed and confirmed the presence of fibronectin and hydroxyproline in the decellularised endometrium was comparable to the native tissue (Fig 1 D).

Biochemical characterisation of endometrium-derived ECM

[00155] The different ECM components provide structural support and transmit functional signals to resident cells (M. W. Pickup, J. K. Mouw, V. M. Weaver, The extracellular matrix modulates the hallmarks of cancer. EMBO Rep 15, 1243-1253 (2014)). Confocal Raman micro-spectroscopy was used to visualize the spatial distribution of different components in extensive detail and compare the spectral profiles between native and decellularized endometrium from the three different decellularization procedures (P1 , P2, P3; Fig 2A, 2B). Confocal Raman images showed remarkable similarity in the spatial distribution of ECM proteins in native and decellularized tissues (Fig. 2A). The Raman spectra across all samples detected prominent peaks at 1004 cm ¹ (Phe:phenylalanine), 1061 cm ¹ (GAG:glycosaminoglycan), 1135 cm ¹ (C-C Asy: asymmetric carbon-carbon stretching mode), 1298 cm ¹ (Amide III), and 1450 cm ¹ (collagen) (Fig 2B). Among three different decellularization protocols, we observed a higher degree of peak intensities for GAG and Amide III, which represents the collagen secondary structures, in P3 compared to other groups (Fig. 2B).

[00156] To validate the Raman micro-spectroscopy data, analyses were performed for selected ECM proteins including collagens (both soluble and insoluble type), sulfated glycosaminoglycans (sGAG) and hydroxyproline in native and decellularized endometrial tissues (Fig. 2C-F). Significant enrichment of both soluble and insoluble forms of collagen was observed in decellularized tissues compared to the native tissue (Fig. 2C, 2D). Hydroxyproline is a major component of collagen and involved in collagen biosynthesis, stability and strength (M. E. Grant, D. J. Prockop, The biosynthesis of collagen. 3. N Engl J Med 286, 291-300 (1972)). Hydroxyproline concentration was increased in decellularised tissues than native controls in the P3 group (Fig. 2E). In P1 and P2 groups, hydroxyproline concentration was either unchanged or decreased relative to their respective controls (Fig. 2E). The amount of sulfated glycosaminoglycans was relatively stable across the samples except a slight decrease in decellularised tissue samples belonging to the P3 group (Fig. 2F). To further characterize the molecular structure and detect the functional groups, such as amide bonds and sugars present in the decellularized tissue samples, Fourier-transform infrared spectroscopy (FTIR) was employed. The IR spectra clearly showed transmittance bands corresponding to amide A (~3300 cm ¹ ), amide B (~3100 cm ¹ ), amide I (~1650 cm ¹ ), amide II (~1550 cm ¹ ), and amide III (~1200 cm ¹ ) stretching and bending modes consistent with the presence of peptide backbone of proteins in decellularized tissues in all three samples (Fig. 2G), suggesting that the chemical integrity of ECM proteins is not compromised by the decellularization protocols.

Mechanical properties of endometrium-derived ECM hydrogel

[00157] To prepare ECM hydrogels, lyophilized decellularized bovine endometrium was cryomilled into a fine powder to ensure uniform digestion during the gelation step (Fig. 1A). The diameter of particles in ECM powder was measured by dynamic light scattering. The particle size distribution yields an average diameter of approximately 420.2 ± 10.27 nm for P1 ECM (Fig 3A), 568.3 ± 7.01 nm for P2 ECM (Fig 3B) and 420.2 ± 3.45 nm for P3 ECM (Fig 3C). Hydrogels were then generated from ECM powders representing three different protocols (P1 , P2 and P3) at 10 mg/ml concentration (Fig. 3D). All three hydrogels solidified at physiological pH and temperature (Fig. 3D). SEM imaging of solidified ECM hydrogels revealed distinct interconnected fibrillary organization akin to collagen type 1 hydrogels (Fig. 3D). The gelation kinetics of three hydrogels was assessed by turbidimetric analysis. This technique measures the increase in turbidity, and thus absorbance, observed during the assembly of collagen fibrils (J. Fernandez-Perez, M. Ahearne, The impact of decellularization methods on extracellular matrix derived hydrogels. Sci Rep 9, 14933 (2019)). All three hydrogels underwent gelation after a lag phase (Fig. 3E). The P3 hydrogel became more turbid reaching 90% of gelation in 15 minutes than P1 and P2 hydrogels (Fig. 3E). Hydrogels are polymeric and hydrophilic materials that have three dimensional structures with ability to entrap a large amount water (E. M. Ahmed, Hydrogel: Preparation, characterization, and applications: A review. J Adv Res 6, 105-121 (2015)). The stability of the hydrogels was tested by examining their ability to absorb water and undergo degradation in the presence of a reducing agent, dithiothreitol (DTT) (Fig. 3F, 3G). The water uptake capacity of P3 hydrogels was lower compared to P1 and P2 hydrogels (Fig. 3F). Consistently, the degradation resistance of P3 hydrogel was 3-fold higher than P1 and P2 hydrogels (Fig. 3G).

[00158] To investigate the mechanical properties of endometrial hydrogels, rheological assessments were performed using a parallel plate rheometer. The storage modulus (G’; represents elastic behaviour of material when deformed) and loss modulus (G”; reflects viscous behaviour of material when deformed) of hydrogels was measured with increasing temperature from pre-gelling (5°C) to physiological temperature (37°C) (Fig. 3H). The P3 hydrogel had a higher storage and loss modulus than P1 and P2 hydrogels and Matrigel (Fig. 3H, 3I), suggesting that P3 hydrogel has higher viscoelasticity and stronger mechanical strength relative to other gels. To determine which hydrogel has mechanical properties equivalent to the endometrial tissue, storage modulus, complex viscosity, and oscillation stress of the three endometrial hydrogels, Matrigel, and decellularized bovine endometrial tissues was compared (Fig. 3J-L). The assessments of these parameters revealed that the mechanical properties of P3 hydrogels are more closely related to the decellularized endometrial tissue (Fig. 3J-M), suggesting P3 hydrogel might provide a more natural and supportive environment for the growth of endometrial organoids. [00159] The ability of the hydrogels to develop tube-like structures to mimic the shape and size of a decellularized mouse uterus was also tested (Fig. 3M, 3N). A simple setup of a tuberculin syringe and a sterile cotton bud to develop these tubal structures was used (Fig. 3Na). Agarose was first used to show that a tubal structure with a patented lumen can be easily developed using the device (Fig. 3Nb- d). Similar structures were then developed using endometrial hydrogels and Matrigel (Fig. 3Ne-l). Consistent with their mechanical properties (Fig. 3G-L), Matrigel and P1/P2 hydrogel were too soft to form a proper tube-like structure (Fig. 3Ne-g). Tubal structures made from these materials were solid and looked rough from the outside (Fig. 3Ne-g). In comparison, the P3 hydrogel-derived tubes were smooth from the outside and grossly appeared similar to the decellularized mouse uterus (Fig. 3M, 3Nh). To further improve the strength and stability of P3 hydrogels, two non-cytotoxic crosslinkers were used, genipin and N-(3-Dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC), that are known to crosslink adjacent collagen microfibrils (H. W. Sung, W. H. Chang, C. Y. Ma, M. H. Lee, Crosslinking of biological tissues using genipin and/or carbodiimide. J Biomed Mater Res A 64, 427- 438 (2003)). The addition of genipin, EDC, and their combination led to a stable patented lumen formation in P3 hydrogel-derived tubal structures (Fig. 3Ni-l). Therefore, these tubes appear architecturally comparable to the mouse uterus. A dark blue coloration typically appears in hydrogels after the addition of genipin (H. W. Sung, W. H. Chang, C. Y. Ma, M. H. Lee, Crosslinking of biological tissues using genipin and/or carbodiimide. J Biomed Mater Res A 64, 427-438 (2003)).

Proteomic profiling of decellularized endometrium

[00160] An in-depth characterization of ECM proteins present in the decellularized bovine endometrium was also performed (Fig. 4A). The 4plex iTRAQ labeling system containing four pooled samples from each protocol (P1 , P2, P3) was used, enabling a comparison of the three protocols in the same LC-MS/MS condition, thereby reducing experimental variation (Fig 4A) (M. F. B. Jamaluddin, P. Nahar, P. S. Tanwar, Proteomic Characterization of the Extracellular Matrix of Human Uterine Fibroids. Endocrinology 159, 2656-2669 (2018)). The LC-MS/MS analysis revealed that 439 proteins were differentially expressed between P3 and P1/P2 groups (Fig. 4B). The total identified proteins were screened with an existing ECM protein database (Matrisome) to facilitate identification of ECM and ECM-associated proteins (A. Naba et al., The matrisome: in silico definition and in vivo characterization by proteomics of normal and tumor extracellular matrices. Mol Cell Proteomics 11 , M111.014647 (2012)). This step-by-step process enabled the matrisome signature enriched in the decellularized endometrium matrix achieved from the three different methods (P1 , P2 and P3) to be defined and to analyze differences in ECM protein levels between them. Matrisome analysis led to the identification of ECM proteins that were then sorted into matrisome subcategories within the core matrisome (ECM glycoproteins, collagens, and proteoglycans) and matrisome-associated proteins (ECM-affiliated proteins, ECM regulators and secreted factors) (Fig 4C). Altogether, 54% ECM proteins from three treatment groups were detected as core matrisomal proteins which contained 49% ECM glycoproteins, 46% collagens, and 5% proteoglycans in the decellularized endometrium (Fig 4C). The remaining 46% ECM proteins were classified as matrisome-associated proteins that comprised of 25% ECM-affiliated proteins, 44% ECM regulators, and 31% secreted factors (Fig 4C).

[00161] The comparative analysis of matrisome revealed differences in the composition of P1 , P2, and P3 (Fig. 4B). Compared to the P3 group, 25 and 28 ECM proteins were decreased in the P1 , and P2 groups, respectively (Fig 4C). Many of these proteins (e.g., laminin subunit alpha 2: LAMA2) are the regulators of the physiological functions of the endometrium. The expression of three key ECM proteins (LAMA1 : laminin subunit alpha 1 , fibronectin, collagen 1) was validated in P1 , P2, and P3 samples using Western blotting (Fig. 4D). Laminin was only detectable in the endometrium decellularized using 4% SDC (P3) but not with 1% (P2) or 4% (P1) SDS (Fig. 4D, 4E). Laminins are a major component of Matrigel. Fibronectin and collagen 1 protein expression were comparable in P2 and P3 but decreased in the P1 group (Fig. 4D, 4F, 4G).

Endometrium ECM-derived hydrogels support human and mouse organoid cultures

[00162] Organoids were developed from both human and mouse cells in endometrial hydrogels (P1 , P2, P3) and Matrigel (Fig. 5A-C). Mouse endometrial cells showed robust growth and developed spherical shape organoids in Matrigel (Fig. 5Aa). An equal number of mouse endometrial cells was then plated in an identical culture medium in the ECM hydrogels (Fig. 5Ab-d). Organoid forming efficiency, organoid shape, and organoid size was comparable between P3 hydrogel and Matrigel (Fig. 5Ad, i). Organoids formed in the P1 hydrogel were not spherical and appeared to have both luminal and glandular compartments (Fig. 5Ab, 5Ae-j, 5B), which were akin to the mouse endometrial tissue (Fig. 7). To validate if endometrial organoids developed in the P1 hydrogel have a distinct glandular compartment, the expression of Foxa2, a known marker of uterine glands, was examined (Fig. 5B). Foxa2+ cells were interspersed with Foxa2- cells in the spherical shaped mouse endometrial organoids grown in the Matrigel (Fig. 5Ba-d). In contrast, Foxa2+ cells were restricted to the glandular projections coming out of the Foxa2- central body of the organoids developed in the P1 hydrogel (Fig. 5Be-h). The number of Foxa2+ and Ki67+ (a marker of proliferating cells) cells were comparable in organoids grown in both Matrigel and P1 hydrogel (Fig. 5Bi, Bj).

[00163] Although organoids cultured in the P1 hydrogel closely mimic the architectural arrangement of endometrial epithelial cells of the native tissue, fewer organoids were developed in the P1 hydrogel than in Matrigel and P3 (Fig. 5Ai). Since the biochemical analysis revealed a lower laminin expression in the P1 hydrogel relative to P3 (Fig. 4D), the P1 hydrogel was supplemented with laminin (Fig. 5Bk- m). Laminin was isolated from the bovine endometrium and added it to the P1 hydrogel (Fig. 5Bk, 8). Supplementation of laminin to the P1 hydrogel increased the number of organoids in a concentration dependent manner (Fig. 5Bk, I). These organoids also progressively acquired a round morphology and become similar to organoids grown in Matrigel and P3 hydrogel (Fig. 5Bm).

[00164] Similar experiments were then performed using human cells to explore if ECM hydrogels support human organoid cultures (Fig. 5C). Immortalized endometrial cancer cells (Ishikawa) and primary normal and cancerous patient-derived endometrial cells were cultured in ECM hydrogels and Matrigel (Fig. 5C). Robust organoid development was present in both P3 and Matrigel (Fig. 5Cc, d, g, h, k, I, 5D). The ability of ECM hydrogels to support the growth of organoids from other tubular organs, such as the colon and lung was also tested (Fig. 5Cm-t, 5D). Seeding of primary human colon and mouse lung cells led to the robust development of organoids in the P3 hydrogel and Matrigel (Fig. 5Cm-t, 5D). Similar to mouse endometrium (Fig. 5A), some organoid growth was also observed in the P1 hydrogels, indicating that endometrium-derived ECM can also support the growth of organoids from other tubular organs. [00165] Investigations were then performed into whether ECM hydrogel derived from the normal human endometrium would also be applicable for culturing endometrial organoids (Fig. 5E-G). Human endometrium was decellularized and hydrogels were similar to the bovine endometrial-derived P3 hydrogel were developed (Fig. 1 , 5E). Similar to the bovine P3 hydrogel, human P3 hydrogel supported the growth of normal human endometrial organoids, which was comparable to the growth observed in the Matrigel (Fig 5F, 5G).

Proteomic differences in human endometrial organoids cultured in P3 hydrogel and Matrigel

[00166] Studies in multiple organs have established that signals emanating from the ECM proteins drive the pathogenesis of neighbouring epithelial cells (M. W. Pickup, J. K. Mouw, V. M. Weaver, The extracellular matrix modulates the hallmarks of cancer. EMBO Rep 15, 1243-1253 (2014)). The matrisome of cancer tissues significantly differs from the matrisome of their normal counterparts (A. Naba et al., The matrisome: in silico definition and in vivo characterization by proteomics of normal and tumor extracellular matrices. Mol Cell Proteomics 11 , M111 .014647 (2012)). The molecular differences in organoids cultured in the P3 hydrogel and Matrigel was examined. Specifically, the proteome of normal human endometrial organoids cultured in the bovine P3 hydrogel and Matrigel was analysed (Fig. 5H-L). The proteome of human endometrial organoids cultured in the P3 hydrogel was more closely related to the proteome of the native patient endometrial tissue than the organoids grown in the Matrigel (Fig. 5H, 5I). 1300 proteins were commonly identified between the tissue and organoids (Fig. 5H). MS intensities of these proteins were normalized to the Z-score (Fig. 5H, 5J). The proteins in the tissue, P3 hydrogel organoids, and Matrigel organoids were ranked based on the Z-score (Fig. 5J, 5K). This analysis revealed that the P3 hydrogel organoids are more proteomically similar to the native endometrial tissue than the Matrigel-grown organoids (Fig. 5J, 5K). Ingenuity pathway analysis (IPA) revealed that some of the major signaling pathways were either downregulated or missing in the organoids cultured in Matrigel relative to the P3 hydrogel organoids and the native tissue (Fig. 5L). For example, The Integrin-Linked Kinase (ILK) pathway and RHOA signaling were detected in both the native tissue and P3 hydrogel organoids but not in the Matrigel-grown organoids (Fig. 5L). Both signaling pathways are regulators of endometrial functions and fertility and their dysregulation leads to the pathogenesis of endometriosis and endometrial cancer.

Generation of ECM hydrogel-derived porous scaffolds for culturing human endometrial cells

[00167] Porous scaffolds are three-dimensional polymeric materials with interconnected pores that are widely used in tissue engineering (G. Lutzweiler, A. Ndreu Halili, N. Engin Vrana, The Overview of Porous, Bioactive Scaffolds as Instructive Biomaterials for Tissue Regeneration and Their Clinical Translation. Pharmaceutics 12 (2020)). Tests were performed into whether P3 hydrogel is biocompatible for fabricating porous scaffolds for supporting organoid growth and for future applications in regenerative therapies. The ice templating technique was used in which P3 hydrogel was progressively frozen so that biomaterial particles started concentrating around the growing ice crystals and once the sample was fully frozen, ice crystals were removed by lyophilization, leaving a desired porous biomaterial (Fig. 6A) (H. Joukhdar et al., Ice Templating Soft Matter: Fundamental Principles and Fabrication Approaches to Tailor Pore Structure and Morphology and Their Biomedical Applications. Adv Mater 33, e2100091 (2021)). Using this technique, scaffolds of required shapes and sizes can be developed and stored at room temperature (Fig. 6B). SEM and H&E staining confirmed a honeycomb-like porous appearance of these scaffolds (Fig. 6C-E). Next, endometrial cancer cells (Ishikawa) were seeded onto P3 scaffolds and assessed their growth after 12 days of the culture period. Both grossly and histologically, the three-dimensional growth of endometrial cells was clearly observed throughout the P3 scaffold (Fig 6F, 6G). Immunolocalization of phospho-histone 3 (PH3), a nuclear mitotic marker, and cytokeratin 8 (CK8), a marker of epithelial cells, revealed the presence of PH3+ CK8+ proliferating epithelial cells in these scaffolds (Fig 6J), suggesting that endometrial cells are actively dividing to form these organoids. Primary human endometrial cells were also cultured on these scaffolds to show that normal patient-derived cells are also able to grow on these scaffolds (Fig. 9).

Materials and methods

Endometrial tissue collection and decell ularisation

[00168] Grossly and histopathologically normal female reproductive tracts of cows (age 12-18 months) were collected from an abattoir and transported immediately to the laboratory in Dulbecco’s modified Eagle’s medium (Sigma, USA). Upon arrival, the tissues were washed briefly with phosphate-buffered saline (PBS) containing 1% penicillin/streptomycin and amphotericin to remove excess blood and reduce initial microbial load. The uterus was carefully removed, and the rest of the tissues (broad ligament, Fallopian tubes and ovaries) were discarded. Next, the endometrium was surgically separated from the myometrium. The endometrium was then cut into two-centimetre pieces and placed in Milli-Q water (containing 1% penicillin/streptomycin and amphotericin) under constant agitation on a magnetic stirrer for overnight at room temperature. Fresh Milli-Q water was replaced in every 3-4 hours interval. Each batch of endometrial tissue contained four pooled animals being generated for decellularization treatment in this study. After 24-hour wash with Milli-Q water, each batch of endometrial tissues (n=4 pooled) were treated with the respective surfactants for decellularization: Protocol 1 (P1) - 4% sodium dodecyl sulfate (SDS) (Sigma), Protocol 2 (P2) - 1% SDS, and Protocol 3 (P3) - 4% sodium deoxycholate (SDC) solution (Sigma). Each of the protocols were performed under constant agitation (stir plate), with fresh solution changed every 3-4 hours at room temperature until all the tissue pieces become translucent (usually after 4-5 days). After surfactant treatment, tissues were extensively rinsed with water overnight, followed by 2000kU DNase (Sigma) in 1 M NaCI for three hours at room temperature. Decellularized tissues were then washed in Milli-Q water for additional two days, with multiple water replacement to remove potentially cytotoxic residues of SDS/SDC, prior to lyophilization and milling into powders.

[00169] Normal human premenopausal endometrium and endometrial cancer tissue samples were collected from patient undergoing surgery by following guidelines approved by Institutional Human Research Ethics Committee at the University of Newcastle and the Hunter New England Human Research Ethics Committee. The human endometrium was processed similar to the bovine tissue and was decellularized using protocol 3 (4% SDC).

Histology, immunofluorescence (IF) and immunohistochemistry (IHC)

[00170] Decellularized and native endometrial tissues from each protocol were fixed for 20 hours in neutral buffered formalin solution (Sigma) at 4°C (n=4/each). Hematoxylin and eosin staining was performed using the standard protocol to visualize the nuclei and cytoplasm. Immunostainings were performed as described (S. M. Syed et al., Endometrial Axin2(+) Cells Drive Epithelial Homeostasis, Regeneration, and Cancer following Oncogenic Transformation. Cell Stem Cell 26, 64-80 e13 (2020)). The primary antibodies were used for this study were rabbit anti-fibronectin (1 :500, ab32419, Abeam USA), rabbit anti-hydroxyproline (1 :200, 73812, Cell Signaling Technology USA), rabbit anti-Ki67 (1 :400, 15580, Abeam) (44), rabbit anti-Foxa2 (1 :100, 108422, Abeam) and rat anti-cytokeratin 8 (1 :250, TROMA-1 , Developmental Studies Hybridoma Bank) were used. The following secondary antibodies were used in this study Alexa 488/594 conjugated anti-rabbit/rat IgG (1 :250; Jackson ImmunoResearch Labs) for IF and Biotin or HRP conjugated anti-rabbit IgG (1 :250; Jackson ImmunoResearch Labs) for IHC.

Scanning electron microscopy (SEM)

[00171] Three decellularized and native endometrium tissues from each treatment were characterized by SEM analysis. All tissues were fixed in 2% glutaraldehyde (Sigma) overnight and used for SEM imaging. Briefly, the fixed tissues were passed through increasing concentrations of ethanol starting from 10% to 100% ethanol at 30-minute intervals and left in 100% ethanol at least overnight for dehydration. Next the dehydrated tissues were critical point dried with liquid carbon dioxide in a critical- point dryer (Leica CPD300) and then attached to an aluminium stub using double-sided carbon tape for gold-sputter coating (SPI-Module Gold Sputter Coater, Structure Probe). Finally, images were acquired using Carl Zeiss Sigma VP FE-SEM (Zeiss) microscope.

Residual DNA and SDS quantification of decellularized tissue

[00172] Total DNA was extracted from the native (n=6) and decellularized endometrium tissue (n=6) using QIAamp DNA Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. The purity and yield of the DNA in the extracting solution was measured by NanoDrop™ 2000/2000c spectrophotometer (Thermo Fisher Scientific, USA) at optical densities at 260 nm and 280 nm. To evaluate the size of the remaining DNA fragments in the decellularized endometrium for all treatment groups (n=6), including native endometrium (n=6), gel electrophoresis was performed using 1.5% agarose gel in tris-acetate-EDTA buffer containing 0.1% ethidium bromide. 12 pL from each extracted DNA sample and 2.4 pl of loading dye were loaded into each well. HyperLadder™ 100 bp was used as a reference. After running the gel (80 V; 70 min), the DNA was visualized and photographed using LAS-3000 Imager (Fujifilm, Japan). Decellularized and lyophilized endometrium tissue from the two SDS protocol (P1 and P2) were analyzed in triplicate for SDS contents using a colorimetric based SDS Detection and Estimation kit (G-Biosciences, USA). The absorbance was determined by Thermo Scientific Helios y Gamma Spectrophotometer and the concentrations from a standard curve were used to measure the SDS quantity in the tissues.

RAMAN micro-spectroscopy

[00173] Confocal Raman analysis of biological samples (before and after decellularization) were performed using a correlative Raman-AFM microscope (alpha300 RA, Witec.GmbH) system, coupled to a 785 nm laser source, and calibrated to the 520 cm ¹ silicon standard peak. A laser power of 50 mW was maintained throughout the measurements, using a Zeiss EC Epiplan-Neofluar Die 100x I NA 0.9 objective. A grating with a groove density of 300 per mm was selected to achieve a spectral coverage of -100 cm ¹ to 3200 cm ¹ . Data acquisition parameters include 10 accumulations per point and 10 s integration time. In a hyperspectral Raman imaging platform, the automated confocal microscope system offers an acquisition of full Raman spectrum at each pixel. For the measurements, a tissue section/extracellular matrix were transferred to a silicon substrate, which was surface functionalized with a proprietary hydrogel. Raman maps on 80 pm x 40 pm tissue sections were obtained under similar conditions (100 scan lines, each with 200 points) to generate a 20,000-pixel image with a pixel resolution of 400 nm.

Fourier-transformed infrared (FTIR) analysis

[00174] The identification of characteristic functional groups in ECM was carried out by analyzing the vibrational spectra of the samples using a Frontier MIR/NIR spectrometer from Perkin Elmer. For measurements, samples were mixed with KBr, used as a carrier, in a ratio of 1 :200 mg (sample: KBr), pressed into a thin pellet for spectral measurement in the transmittance mode within the range of 4000 cm ¹ - 500 cm ¹ and 64 scans were measured at a resolution of 2 cm ¹ for each sample.

ECM quantification

[00175] Sulfated glycosaminoglycans (GAGs, Blyscan™ Glycosaminoglycan Assay kit, Biocolor, UK), insoluble collagen (Sircol™ Insoluble Collagen Assay kit, Biocolor), soluble collagen (Sircol™ Soluble Collagen Assay kit, Biocolor), hydroxyproline (Hydroxyproline Assay kit, Sigma) were quantified from the native (n=3) and decellularized endometrium tissues (n=3/each protocol). The absorbance was determined by Varioskan LUX multimode microplate reader Version 1 .00.38 (Thermo Fisher Scientific). Samples were run in triplicate and averaged.

Laminin isolation

[00176] Laminin isolation from the bovine endometrium tissue samples was performed as described (H. K. Kleinman, Preparation of basement membrane components from EHS tumors. Curr Protoc Cell Biol Chapter 10, Unit 10 12 (2001)). Briefly, 100 g of bovine endometrium (n=4 pooled) was placed in 200 mL of 3.4 M NaCI buffer and homogenize with an electric homogenizer until dispersed. Homogenate were centrifuged for 15 minutes at 8000 x g, 4 °C to pellet. The pellet was resuspended and homogenize in 0.5 M NaCI buffer and then stirred overnight at 4 °C. Homogenate were centrifuged for 15 minutes at 8000 x g, 4 °C to obtain supernatant. Ammonium sulfate was added to the 0.5 M NaCI supernatant to 30% saturation with vigorous stirring for 1 hour. Sample was then centrifuged for 15 minutes at 8000 x g, 4 °C to separate laminin-1 pellet. Laminin pellets were dissolved in 300 mL TBS and dialyze for 2 hours against 2 liters TBS and dialysis process was repeated twice against 2 liters TBS each time. Dialysis bag was then emptied to measure the volume of sample and addition of NaCI to a final concentration of 1.7 M. The supernatant was further dialyzed against TBS for three changes and the amount of laminin is determined by the Lowry assay.

Lyophilisation and particle size distribution measurements

[00177] The decellularized endometrium tissue samples were lyophilized for 72 hours (Flexi-Dry™ Microprocessor Freeze-Dryer, USA) and then milled into fine powders using 6970EFM Freezer Mill (SPEX, USA). These powders can be stored at -20 °C for future use. The average diameter of the particles (nanometer) was measured by dynamic light scattering at 25 °C using Zetasizer Nano ZS90 (wavelength = 632.8 nm). All measurements were performed in triplicate and were averaged to obtain the final value. Fabrication ofdecellularized ECM scaffolds

[00178] One gram of decellularized ECM powder prepared using P3 protocol was dissolved in 100 mL of 1% acetic acid. The homogeneous solution was then transferred to 12-well culture plate, frozen at -80 °C for 8 hours, and then lyophilized for 48 hours to harvest the scaffolds. To eliminate acetic acid, the scaffolds were immersed in 1% w/v NaOH for 30 minutes, flushed with distilled water and lyophilized to create the final scaffolds.

Proteomic digestion and ITRAQ labeling

[00179] The lyophilized and cryomilled ECM powder (n=4 pooled) from the respective three protocols was resuspended in lysis buffer and homogenized with a BeadBug homogenizer (Benchmark Scientific, USA) to create a homogenous ECM suspension. The ECM suspension was dissolved in 8 M Urea and reduced with 10 mM dithiothreitol (Sigma) at 37 °C for 1 hour, followed by alkylation with 20 mM iodoacetamide (Sigma) in the dark at room temperature for 1 hour. Protein samples were then digested with trypsin/Lys-C (Promega, USA) at 1 : 25 ratio to protein and incubated at 37 °C for 3 hours. Samples were then diluted to < 0.75 M by the addition of 50 mM triethylammonium bicarbonate, pH 7.8 (Sigma) and digested overnight at 37 °C. The following day, peptide solutions were acidified with 1% trifluoroacetic acid (TFA) (Sigma), desalted using the solid phase extraction (SPE) columns (Oasis PRIME HLB, Waters, Australia), and eluted with gradient 60, 80, 100 acetonitrile, 0.1 % TFA. Peptide concentrations were measured using Qubit 2.0 Fluorometer assay (Invitrogen, USA). Each iTRAQ reagent (AB SCIEX, USA) was dissolved in 70 pL of ethanol and added to the respective peptide mixture for labelling. 100 pg of each sample was labeled as (P1 : 4% SDS)-114, (P2: 1% SDS)- 115 and (P3: 4% SDC)-116. Additionally, the samples were multiplexed and vacuum dried.

Off-line high pH reversed-phase fractionation

[00180] iTRAQ-labeled peptides were fractionated by high pH fractionation using a Dionex UltiMate 3000 capLC system (Dionex, USA) equipped with Acquity UPLC® M-Class CSH™ C18 130A 1.7 pm, 300 pm x 100 mm column, with the following mobile phases: 2% Acn in 98% HPLC water (solvent A) and 98% Acn in 2% HPLC water (solvent B); both solvents were adjusted to pH 10 with ammonium formate. Peptides were separated using a 50 min gradient from 5% to 95% solvent B at a flow rate of 5 pL/min. The elution was monitored by absorbance at 214 nm, and the column was operated at 25 °C. A total of 56 fractions were collected into a deep 96-well LoBind plate (Eppendorf). Based on the high pH fractionation spectra, 39 out of the 56 fractions were concatenated into 13 fractions by combining fractions. Each pooled fraction was concentrated by vacuum centrifugation and reconstituted in 10 pL of 2% Acn and 0.1% TFA.

Proteomic LC/MS-MS analysis

[00181] The resulting peptide mixtures were analyzed by electrospray liquid chromatography mass spectrometry (LC MS/MS) (M. F. B. Jamaluddin, P. Nahar, P. S. Tanwar, Proteomic Characterization of the Extracellular Matrix of Human Uterine Fibroids. Endocrinology 159, 2656-2669 (2018)). Dionex Ultimate 3000 nanoLC system (Thermo Fisher Scientific) was connected to an Orbitrap Exploris 480 mass spectrometer (Thermo Fisher Scientific, Germany). 4.5pL of peptides were delivered to the trap column (Acclaim PepMapTM100, 100 pm x 2 cm, nanoViper fitting C18, 5 pm, 100A°, Thermo Fisher Scientific, USA) using solvent A (0.1% formic acid in HPLC water) and separated on the analytical column (EASY-Spray Column, PepMap, 75 pm x 25 cm, nanoviper fitting, C18, 2 pm, 100A°, Thermo Fisher Scientific, USA) using a 90 min linear gradient from 2% to 40% solvent B (0.1% formic acid in acetonitrile) at a flow rate of 300 nL/min. The MS was operated in data-dependent acquisition mode, the mass-to-charge (m/z) range for the acquisition of MS1 spectra was 360-1500 m/z at an Orbitrap full MS scan with a resolution of 60,000, RF lens is 40%, automatic gain control (AGC) target value: 1e6, with a maximum injection time of 50 ms. In the MS2, the top 20 peptide precursors were selected for HCD fragmentation using a normalized collision energy of 36.0 at a resolution of 15,000 with maximum ion injection time of 120 ms and AGC target value set at 2e5. A 1 .2 m/z isolation window was used for MS/MS scans. Dynamic exclusion was set for 15 seconds. Spectra were analyzed using Proteome Discoverer (PD) software v.2.5 (Thermo Fisher Scientific) and searched against bovine protein sequences downloaded on 12/07/2020 from UniProtKB containing 1 ,783 entries. The parameters were set as follows: trypsin digestion; two missed cleavages; 6 minimum peptide length; fixed modifications of cysteine (carbamidomethylation) (+57.021 Da); variable modifications of methionine oxidation (+15.995 Da), lysine acetylation (+42.011 Da), lysine methylation (+14.016 Da) and iTRAQ4plex labelling of lysine (+144.102 Da). A precursor mass tolerance was set to 10 ppm and fragmentation mass tolerance was 0.02 Da. All other parameters were used as default settings. A fixed false discovery rate (FDR) threshold was set at < 1% to report peptide and protein identifications. The data was then mapped using an in silico matrisome list to categorize and annotate all identified proteins (A. Naba et al., The matrisome: in silico definition and in vivo characterization by proteomics of normal and tumor extracellular matrices. Mol Cell Proteomics 11 , M111.014647 (2012)). The raw mass spectrometry data have been deposited in the public repository MassIVE using the identifier: MSV000089054, available at

Analyses of iTRAQ data was visualized in heat maps and dot plots using ProHits-viz (J. D. R. Knight et al., ProHits-viz: a suite of web tools for visualizing interaction proteomics data. Nat Methods 14, 645- 646 (2017)) and PD software, respectively. For proteomic analysis of normal human endometrial organoids cultured in the bovine P3 hydrogel and Matrigel, protein abundances were normalized by z- score transformation for heatmap analysis, line plot and violin plot. Venn diagram plotter was used to highlight the number of shared and differentially expressed proteins between the groups.

Western blotting analysis

[00182] The lyophilized and cryomilled ECM powders (n=4 pooled) from the respective three protocols were resuspended in RIPA buffer and homogenized with a handheld sonicator (Benchmark Scientific, USA) to create a homogenous ECM suspension (M. F. B. Jamaluddin, P. B. Nagendra, P. Nahar, C. Oldmeadow, P. S. Tanwar, Proteomic Analysis Identifies Tenascin-C Expression Is Upregulated in Uterine Fibroids. Reprod Sci 26, 476-486 (2019)). Protein extracts (30 pg) were subjected to SDS- PAGE (4-20%, Mini-PROTEAN TGX™ Gels, Bio-Rad, Hercules, CA). SDS-PAGE was used to separate protein homogenates, which were transferred to a nitrocellulose membrane (NC45; 0.45 pM; Sigma). Membranes were blocked for 1 hour in 5% milk (weight/volume ratio) in Tris-buffered saline containing 0.1% Tween-20 (TBST), followed by incubation with rabbit anti-collagen (dilution 1 :1000, AB745, Millipore USA), anti-fibronectin (dilution 1 :1000, ab32419, Abeam USA), anti-laminin (dilution 1 :1000, L9393, Sigma, St. Louis, MO, USA), or anti-GAPDH (dilution 1 :5000, #5174, Cell Signaling Technology USA) overnight at 4 °C. After three washes in 1 x TBST, membranes were incubated with secondary antibody (Jackson ImmunoResearch Laboratories) for 1 hour at room temperature. Membranes were thoroughly washed three times in 1 x TBST before imaging. Reactive protein was detected using an LAS-3000 imager (Fujifilm). Densitometric analysis of the Western blot was performed using Imaged plugin software (NIH), and all proteins were quantified relative to the loading control.

Gelation

[00183] For gelation (P. A. Link, R. A. Pouliot, N. S. Mikhaiel, B. M. Young, R. L. Heise, Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture. J Vis Exp 10.3791/55094 (2017)), the ECM powder was digested at 10 mg/mL in pepsin (1 mg/mL) in 0.1 M HCI (Sigma, St. Louis, MO, USA) for 72 hours at room temperature under constant stirring. After 72 hours, the pepsin-solubilized samples were equilibrated to cytocompatible salinity by adding 10% 10X PBS for 2 hours at room temperature, in constant rotation. Pre-gel solution was then neutralized to pH 7.5 by addition of 10 M NaOH and thoroughly mixed. Neutralized pre-gel solution was stored in aliquots at -20 °C for prolonged storage until use.

Degradation and water uptake capacity assays

[00184] ECM endometrium gels generated from three protocols were prepared and equivalent amounts (100mg) of room temperature dried gels were weighed (Wo) (R. Yang et al., Injectable adaptive self-healing hyaluronic acid/poly (gamma-glutamic acid) hydrogel for cutaneous wound healing. Acta Biomater 127, 102-115 (2021)). Gels were then incubated in 1 mL PBS containing 1 M dithiothreitol (DTT) at 37 °C. At each predetermined time point, the PBS were removed, and the remaining gels were weighed (W). The percentage of degradation was calculated by: Weight loss (%) = W/Wo x 100%. The experiments were performed in triplicates. For water uptake capacity assessment, ECM endometrium gel from three protocols were freeze-dried and equivalent (100 mg) dry weights (Wi) were recorded before being re-immersed in deionized water. At predetermined intervals of 3, 8, 11 and 24 h, the swollen gels were dabbed dry with Kimwipes to eliminate surface water and the hydrated weights (Ws) were recorded at each time point. The equilibrium water uptake capacity was calculated as follows: Water uptake capacity (%) = (W _s -W)/W x 100% (X. Chen et al., Development of rhamnose-rich hydrogels based on sulfated xylorhamno-uronic acid toward wound healing applications. Biomater Sci 7, 3497-3509 (2019)). The experiments were performed in triplicates.

Biochemical and mechanical assessment of ECM hydrogels

[00185] The turbidity of the decellularized ECM gels (P1 , P2 and P3) were assessed using spectrophotometry by measuring the absorbance of as prepared gels as a function of time using a Varioskan LUX microplate reader. 100 pL of the ECM gels were pipetted into 96-well plate and absorbance at 450 nm was measured at 37 °C for 30 minutes at 1 minute interval. Mechanical assessment of the hydrogels and bovine tissues was performed on a TA Instruments Dynamic Hybrid Rheometer (DHR-3, TA Instruments). For rheological testing of non-gelled ECM, 50 pL of the ECM solution was placed in between the 8mm parallel plate geometries. For the pre-gelled ECM hydrogels 50 pL hydrogel was solidified at 37°C in a 24-well plate and then scooped out intact before putting between the 8mm parallel plate geometries. However, for bovine tissues, an 8 mm punch biopsy of the samples was used. Shear rheology was measured by subjecting the digested (non-gelled) ECM, pregelled ECM or bovine endometrium to a controlled strain with a continuous oscillation and with an oscillation frequency of 1 rad/sec, an oscillation strain ranging from 0.2% to 2.0% and an axial force of 0.02N. For a subset of samples (non-gelled ECM), this was accompanied by a temperate ramp from 5°C-37°C and rheological profile of gelation was monitored in real time. All other samples were measured at 37°C. Storage modulus was determined as the mean value within the linear viscoelastic range. Data obtained from three biological repeats for each experimental setup.

Hollow cylinder generation

[00186] All preparations of agarose cylinders were performed under sterile conditions. A simple assembly of one ml tuberculin syringe and cotton buds was used to create these hollow cylinders. Different material extracts (including P3 hydrogel, Matrigel, P3 hydrogel + EDC, P3 hydrogel + genipin, and P3 hydrogel + EDC + genipin) were loaded slowly into the syringe barrel by a sterile transfer pipette, and cotton bud was pushed into the internal middle of the syringe. The hollow agarose cylinders in the syringe were incubated for 30 minutes at 37 °C. After 30 minutes, the pipette tip was then removed, and the plunger was used to extrude the now solidified agarose cylinder and cotton bud. The agarose was finally pushed off from the cotton bud and placed in a Petri dish for further testing.

Mouse and human organoids

[00187] Organoids from mouse endometrium (S. M. Syed et al., Endometrial Axin2(+) Cells Drive Epithelial Homeostasis, Regeneration, and Cancer following Oncogenic Transformation. Cell Stem Cell 26, 64-80 e13 (2020)), human endometrium (H. C. Fitzgerald, P. Dhakal, S. K. Behura, D. J. Schust, T. E. Spencer, Self-renewing endometrial epithelial organoids of the human uterus. Proc Natl Acad Sci U S A 116, 23132-23142 (2019)), human endometrial cancer (M. Boretto et al., Patient-derived organoids from endometrial disease capture clinical heterogeneity and are amenable to drug screening. Nat Cell Biol 21 , 1041-1051 (2019)), mouse lung (C. E. Barkauskas et al., Lung organoids: current uses and future promise. Development 144, 986-997 (2017)), human colon (J. Bruce, G. E. Kaiko, S. Keely, Isolation and In Vitro Culture of Human Gut Progenitor Cells. Methods Mol Biol 2029, 49-62 (2019)), and Ishikawa cells (endometrial cancer cells) (J. Goad, Y. A. Ko, M. Kumar, M. F. B. Jamaluddin, P. S. Tanwar, Oestrogen fuels the growth of endometrial hyperplastic lesions initiated by overactive Wnt/beta-catenin signalling. Carcinogenesis 39, 1105-1116 (2018)) were prepared.

[00188] The uterine horns of adult female mice were collected and washed with Mg2+ and Ca2+ free Hank’s Balanced Salt Solution (HBSS). Uterine epithelium was isolated as described previously (Syed et al., 2020 Cell Stem Cell). Briefly, both uterine arms were slit open and then cut longitudinally into 3- 4 mm pieces. These tissue fragments were then incubated in Pronase/DNase enzyme solution [10mg/ml Pronase (Sigma-Aldrich) from Streptomyces griseus and 0.5 mg/ml DNase I (Sigma-Aldrich)] for 13-14 hours at 4 °C on a shaker. After incubation, enzymatic digestion was stopped by adding 10% v/v fetal bovine serum (FBS) (Bovogen), and then the whole mixture with cells were passed through 70 mm cell strainer to collect uterine epithelial cells. Cells were then washed with DMEM-F12 media containing 5% FBS, 1% L-glutamine (Sigma-Aldrich) and 1% penicillin-streptomycin (Thermo Fisher Scientific) by centrifugation at 1500 rpm for 5 minutes. After washing, cells were incubated for 2-3 hours on a cell culture plate with above-mentioned FBS containing DMEM-F12 media for differential attachment.

[00189] After differential attachment, uterine epithelial cells (15000 cells/well) were resuspended in 30% media and 70% Matrigel/P1 hydrogel/P2 hydrogel/P3 hydrogel and was placed as 50 ml droplets in each well of a 24-well cell culture plate. For laminin rescue experiment, same number of cells were mixed with 30% media and 70% mixture of laminin and P1 hydrogel with different ratios. Then 50 ml droplets were placed in each well of a 24-well cell culture plate. All droplets were incubated at 37 °C for 20 minutes to solidify and then overlaid with mouse uterine organoid culture medium as described previously (Syed et al., 2020 Cell Stem Cell). The uterine organoid culture medium contained 25% Wnt3A-R-spondin3-noggin conditioned media (WRN-CM) and 75% Advanced DMEM-F12, supplemented with 1% Glutamax (Thermo Fisher Scientific), 1% HEPES (Sigma-Aldrich #H0887), 1% penicillin-streptomycin (Thermo Fisher Scientific) and additional growth factors such as 2% B27 (Thermo Fisher Scientific), 1% N2 (Thermo Fisher Scientific), 1% insulin-transferrin-selenium (ITS) (Sig ma-Ald rich), 0.2% Primocin (Jomar Life Research #ant-pm-1), 50 ng/ml mouse EGF (Sigma- Aldrich), 50ng/ml human FGF10 (Peprotech), 1 mM nicotinamide (Sigma-Aldrich), 0.5 mM A83-01 (TGFb/Alk inhibitor) (Tocris). 10 mM ROCK inhibitor (Y-27632; 10 mM; TOCRIS) was added for the first 3-days and then the concentration of ROCK inhibitor was reduced to half (5mM) for rest of the culture period. Media was changed every 2-3 days and after 12 days organoids were harvested for further processing.

[00190] For human endometrial epithelium isolation, freshly collected endometrium was washed with PBS containing 1% penicillin/streptomycin and amphotericin to remove excess blood. Then the tissue was minced into fine pieces and incubated with Accumax (Thermo Fisher Scientific #00-4666-56) for 3- 3.5 hours at room temperature on a shaker. After incubation the digested solution was directly transferred to double volume of TrypLE express (Thermo Fisher Scientific #12604021) solution. The mixture was then incubated at 37 °C water bath for 20-25 minutes and continuously agitated in every 5 minutes. After second round of enzymatic digestion, 10% v/v fetal bovine serum (FBS) (Bovogen) was added, and the whole mixture with cells were passed through 70 mm cell strainer to collect epithelial cells. Then the cells were propagated by 2-dimensional culture on a matrigel coated cell culture plate in Advanced DMEM-F12 media containing 5% FBS, 1% Glutamax (Thermo Fisher Scientific), 1% HEPES (Sig ma-Ald rich), 1% penicillin-streptomycin (Thermo Fisher Scientific) and additional growth factors such as 50 ng/ml human EGF, 10 mM ROCK inhibitor (Y-27632; 10 mM; TOCRIS) and 0.2% Primocin (Jomar Life Research). After 70% confluence was achieved, epithelial cells were detached by trypsinization and centrifuged at 1500 rpm for 5 minutes to collect the epithelial cells. For organoid culture, epithelial cells (15000 cells/well) were resuspended in 30% media and 70% Matrigel/P1 hydrogel/P2 hydrogel/P3 hydrogel/human P3 hydrogel and was placed as 50 ml droplets in each well of a 24-well cell culture plate. All droplets were incubated at 37 °C for 20 minutes to solidify and then overlaid with human uterine organoid culture medium containing 25% Wnt3A-R-spondin3-noggin conditioned media (WRN-CM) and 75% Advanced DMEM-F12, supplemented with 1% Glutamax (Thermo Fisher Scientific), 1% HEPES (Sigma), 1% penicillin-streptomycin (Thermo Fisher Scientific) and additional growth factors such as 2% B27 (Thermo Fisher Scientific), 1% N2 (Thermo Fisher Scientific), 1% insulin-transferrin-selenium (ITS) (Sigma-Aldrich), 0.2% Primocin (Jomar Life Research), 50 ng/ml human EGF (Sigma-Aldrich), 100 ng/ml human FGF10 (Peprotech), 1.25 mM N- Acetyl-L-cysteine (Sigma-Aldrich), 1 mM nicotinamide (Sigma-Aldrich), 2 nM Estrogen (Sigma #E8875), 0.5 mM A83-01 (TGFb/Alk inhibitor) (Tocris). 10 mM ROCK inhibitor (Y-27632; 10 mM; TOCRIS) was added for the first 3-days and then the concentration of ROCK inhibitor was reduced to half (5mM) for rest of the culture period. Additionally, only for benign endometrial organoid culture 1 mM Dexamethasone was added in the medium. Media was changed every 2-3 days and after 12 days organoids were harvested for further processing.

[00191] Organoid proliferation was assessed using Presto Blue (A13261 , Thermo Fisher Scientific) and CellTiter-Glo Luminescent 3D cell viability assay following manufacturer’s protocols. Organoids were fixed in 4% (w/v) paraformaldehyde at room temperature for 1 hour before embedding into paraffin blocks.

Human endometrial cell culture in decellularized ECM scaffolds

[00192] Decellularized bovine endometrial scaffolds were cut horizontally into thin slices. These scaffolds were then placed in UV for 1 hour and washed three times with the culture medium for 5 minutes each. Scaffolds were soaked overnight in the culture medium in a cell culture incubator at 37°C. The following day, these scaffolds were taken out from the incubator and transferred into a 24- well cell culture plate. ~1 million primary human endometrial cells or Ishikawa cells were reconstituted in 100 pL of the culture medium and then seeded on the scaffolds. After 30 minutes of incubation, 600 pL of the culture medium was added to the scaffolds in their respective wells. The culture medium was changed every 3 ^rd day and the scaffolds were harvested on the 8 ^th and 14 ^th day and imaged under stereoscope before histological analysis.

Microscopy and image acquisition

[00193] Fluorescence and brightfield images were acquired using Olympus BX43 fluorescence microscope with Olympus objectives (x4, NA = 0.16; x10, NA = 0.46; x20, NA = 0.75; x40, NA = 0.95) or Olympus DP72 CCD (charge-coupled device) with colour camera using cellSens software (Olympus). Gross images were acquired by using SMZ25 stereoscope equipped with Nikon DS-Fi2 camera and Nikon P2-SHR Plan Apo 2X objective (NA = 0.312). Images consisted of 15-30 stacks with variable z-spacing and were captured and merged using NIS-Elements software. Organoid images during the culture period were captured by JuLiTM Stage Real-Time Cell History Recorder (NanoEnTek, South Korea). All imaging was done at room temperature except organoid imaging during the culture period was done at 37 °C.

Statistical analysis

[00194] Image analysis and quantification was done using Imaged software (NIH) and statistical analyses were performed on GraphPad Prism (v.7.04). All data is presented as the mean ± standard error. All tests were performed in triplicate (n=3) unless otherwise specified. Statistical analyses were done using one-way ANOVA, two-way ANOVA and unpaired t test. Data are presented as mean ± SEM. Significance was determined where P is values less than 0.05. For the proteomics data analysis, comparison between two groups was performed using ANOVA (Individual proteins) method to calculate the p-values and p < 0.05 indicate as significant difference. For pathway analysis, z-scores were used to predict activation (z-score > 2; orange) or inhibition (z-score < - 2) of each function. The Raw data obtained from the confocal Raman analysis measurements were pre-processed in Project Five data evaluation software from Witec to account for autofluorescence and cosmic ray anomalies during data acquisition. Autofluorescence was removed through baseline correction of the raw data by using a shape function, while cosmic ray noise was removed by an automated detection algorithm embedded in the Project Five software. The multiple components present in each tissue section were isolated with a “True component analysis” wizard, which identifies individual component based on the corresponding spectral signature. The spectral region of interest from 800 cm ¹ to 1550 cm ¹ was selected while plotting Raman maps of the individual components. Subsequently, these individual Raman maps for discrete components are integrated to generate a combined map and corresponding Raman spectra of the samples.

[00195] It will be appreciated by those skilled in the art that the present disclosure may be embodied in many other forms.