WO2025255682A1 - Extracellular matrix-based hydrogel formulation, methods of making and uses thereof - Google Patents

Abstract

This disclosure relates to the field of extracellular matrix-based hydrogels, and in particular, to hydrogel formulations comprising a decellularized extracellular matrix (dECM) and surface-modified cellulose nanofibers (CNFs) for precise tissue modeling and methods of making and uses thereof.

Description

EXTRACELLULAR MATRIX-BASED HYDROGEL FORMULATION, METHODS OF MAKING AND USES THEREOF

CROSS-REFERENCE TO RELATED APPLICATIONS

[0001] The present application claims the benefit of priority from U.S. provisional patent application No. 63/659,461 filed on June 13, 2024, the contents of which are incorporated herein by reference in their entirety.

FIELD

[0002] The present disclosure relates to the field of hydrogels, and in particular, to hydrogel formulations and methods of making and uses thereof.

BACKGROUND

[0003] The behavior and fate of cells are constantly changing and depend on both the intrinsic biology (such as genetic composition, cytoskeletal contractility, secretion, and various signaling pathways) and extrinsic interactions with the immediate surroundings [1], [2], The extracellular microenvironment includes extracellular matrix (ECM), neighboring cells, and a variety of biochemical substances like hormones, growth factors, and cytokines [3], While the impact of biochemical signals is well-established, the understanding of how the ECM's biophysical properties influence cellular outcomes remains to be more precisely defined[4]. Recent studies discovered that the ECM serves a role beyond being a passive physical support structure, interacting continuously with cells [5], [6], ECM influences cell migration, proliferation, differentiation, and apoptosis, through a complex interplay of mechanical and topographical stimuli[7], [8], [9], The physical attributes of the ECM, including its stiffness, topography, and organization, exert significant influence over cell behavior[10], [11], Cells possess the ability to sense and respond to these physical cues through mechanotransduction pathways, leading to the aforementioned effects on cell biology[12], [13], [14], For instance, a rigid ECM can promote cell proliferation and spreading, while a more compliant ECM may facilitate cell migration and differentiationfl 5], [16], [17],

[0004] To model the interplay between cells and ECM in experimental systems, hydrogels are a material of choice. Hydrogels are gel-like networks composed of polymeric chains that can absorb and retain large quantities of water[18]. Hydrogels act as biomimetic platforms to recreate physiological environments for studying cell behavior, tissue regeneration, and therapeutic interventions. Hydrogels offer advantages such as biocompatibility, tunable mechanical properties, controlled release of bioactive molecules, and the ability to encapsulate cells. Hydrogels can be categorized into: (I) Synthetic Hydrogels: these hydrogels are synthesized from chemically modified polymers like polyethylene glycol (PEG), poly(acrylic acid) (PAA), or poly (vinyl alcohol) (PVA) and exhibit extraordinary properties for long-term applications. (II) Natural Hydrogels: derived from naturally occurring polymers such as alginate, collagen, hyaluronic acid, chitosan, or gelatin[19], [20], These natural hydrogels often exhibit excellent biocompatibility, bioactivity, and cell attachment. However, these natural hydrogels are prone to degradation and batch-to-batch variation limiting their usage. (Ill) Composite Hydrogels: which consist of both natural and synthetic hydrogels, offer some properties of both natural and synthetic hydrogels and have attracted attention for modeling many tissues in health and disease[20],

[0005] Among hydrogels, fibrillar hydrogels possess distinct advantages for modeling complex tissues compared to non-fibrillar hydrogels. Fibrillar hydrogels can be designed so that they emulate the fibrous structure found in native tissues, enabling them to accurately represent tissue architecture at the microscale level [21 ]. This is a particular issue for ECM in tissues where the fibrous arrangement plays a fundamental role in tissue function and mechanical integrity [22], By adjusting parameters such as the polymer composition, crosslinking density, and fiber alignment, hydrogels can be fabricated to replicate tissue-specific stiffness, elasticity, and viscoelastic behavior[23], [24], This allows the investigation of how cells interact with and respond to tissuespecific mechanical cues, providing insights into tissue development, cell-ECM interactions, and disease progression. The fibrous structure of hydrogels provides physical cues that guide cell alignment and migration[25]. Cells seeded within fibrillar hydrogels tend to align along the fibers, closely resembling the natural orientation observed in native tissues [22], [26], This feature is particularly relevant for tissues such as skeletal muscle, where aligned cell orientation is optimal for proper contractile function, or for nerve tissues, where directed axonal growth is optimal for proper neural signaling.

[0006] dECM hydrogels have attracted considerable attention for the development of in vitro models, due to their ability to closely replicate the native environment of various tissues. These materials excel in capturing the precise composition of targeted tissues, providing an optimal microenvironment for cell-to-cell and cell-to-ECM interactions. Their composition not only closely matches the physical structure of the native tissue but also offers tissue-specific biochemical cues, useful for guiding cell behavior and function. However, despite these significant advantages, dECM hydrogels typically fall short in terms of mechanical strength and tunability compared to synthetic hydrogels. To address the mechanical limitations of dECM hydrogels, several strategies such as chemical cross-linking or blending with other hydrogels have been explored[27]. The use of chemical crosslinking agents is a widely investigated approach, employing chemicals like l-ethyl-3-(3-dimethylaminopropyl) carbodiimide, glutaraldehyde, and genipin[27]. These agents are used to react with specific functional groups within the hydrogels. However, chemical cross-linking can lead to issues such as cytotoxicity to cells, the consumption of essential functional groups needed for proper cell adhesion, and potential disruption of the secondary and tertiary structures of proteins[27], [28], Another strategy is the incorporation of synthetic polymers, which can significantly enhance mechanical strength and tunability. However, the introduction of synthetic polymers might disrupt the physical cross-linking inherent in dECM hydrogels, potentially affecting the formation of fibers within the hydrogel matrix. Additionally, reinforcing dECM hydrogels with nano-scale fillers such as nanoparticles or nanofibers has shown promise in improving both the tensile strength and the compressive resistance of the hydrogel matrices [29], [30],

[0007] Nanoparticles, due to their high specific surface area, have been used in tuning the mechanical properties of hydrogels, even at low concentrations! 31 1. Various types of nanoparticles like graphene, carbon nanotubes, nanoclay, magnetic, and cellulose nanoparticles reinforce hydrogel matrices by acting as reversible crosslinkers, linking polymer chains for improved stress distribution, stiffness, extensibility, and toughness [32], Encapsulated within the hydrogel network, they blend a hydrogel’s properties with their own unique features. When modified on their surface functional groups for active participation in crosslinking, nanoparticles further integrate into the network, enhancing both mechanical and functional properties [31], [32], [33], Cellulose nanoparticles (including cellulose nanofibers, CNFs) are notable for their low cytotoxicity as well as for possessing unique properties that make them excellent for reinforcing hydrogels, enhancing mechanical strength, and introducing new functionalities[34], [35], CNFs are distinguished by their fibrillar and flexible nature and are recognized as economical materials for reinforcing hydrogels [36], These cellulose nanoparticles can undergo surface functionalization to facilitate specific chemical interactions with hydrogel networks[37]. For instance, the introduction of aldehyde groups to the surfaces of cellulose nanoparticles through chemical modification or functionalization serves multiple purposes in hydrogel synthesis[37], [38], Aldehyde groups act as crosslinking agents, facilitating the formation of three- dimensional hydrogel networks by reacting with amino groups in polymers or biomolecules. This crosslinking process enhances the mechanical properties of hydrogels, making them more robust and resilient. CNFs, with their semi-crystalline structure and ability to form entangled networks, are particularly adept at creating hydrogels with favorable mechanical properties [38],

[0008] The background herein is included solely to explain the context of the disclosure. This is not to be taken as an admission that any of the material referred to was published, known, or part of the common general knowledge as of the priority date.

SUMMARY

[0009] Developing hydrogel formulations for precise tissue modeling is beneficial for advancing tissue engineering and regenerative medicine applications. A novel approach that combines decellularized extracellular matrix (dECM) with surface- modified cellulose nanofiber materials, such as for example aldehyde-modified cellulose nanofiber materials (aCNFs) to create tunable nanofibrillar composite hydrogels is reported herein. Primary human lung fibroblasts were incorporated into the hydrogels to model fibroblast-to-myofibroblast transition that occurs in periods of wound repair, regeneration, and fibrosis. The hydrogels of the present disclosure demonstrate mechanical properties and structural characteristics closely mimicking the native lung tissue microenvironment in both healthy and fibrotic states. The incorporation of dECM facilitates in situ cell-matrix interactions, providing physiologically relevant extracellular cues that influence cell proliferation, morphology, and function. Notably, increasing the mechanical properties of the hydrogels led to differential expression levels of a-Smooth Muscle Actin (a-SMA) and changes in Yes-Associated Protein (YAP) nuclear localization, indicators of fibroblast- to-myofibroblast transition and mechanotransduction, respectively. Using a TRITC- dextran probe it was demonstrated that the presence of aCNF cross-links did not appreciably alter either the shape of the release curve or the time required to reach equilibrium, indicating that bulk incorporation of aCNF preserved overall macromolecule permeability of the dECM network while providing the desired mechanical reinforcement and supporting the use of the hydrogel formulations of the present dislosure for applications related to bioactive molecule release in tissues.

[0010] The innovative hydrogel formulation presented herein helps bridge the gap between 2D in vitro and 3D in situ cell and tissue microenvironments, enabling tuning of both biochemical and mechanical properties. The integration of dECM with surface-modified cellulose nanofiber materials, such as aldehyde-modified cellulose nanofiber materials, coupled with detailed analysis of cell behavior, provides a foundation to pursue 3D hydrogel models for defining the mechanisms that govern for example fibroblast-to-myofibroblast transition, a process relevant in wound repair, regeneration, and fibrosis.

[0011] The hydrogels of the present disclosure showed improved mechanical properties compared to hydrogels without the surface-modified CNF, such as increased storage modulus, loss modulus, and compression modulus, reduced hydrolytic degradation, and good macromolecule permeability, while viscoelastic properties of the hydrogel remain substantially unchanged. Further hydrogels of the present disclosure showed increased mechanical stiffness compared to corresponding hydrogels modified with surface modified cellulose nanocrystals, which showed a minimal increase in hydrogel stiffness. This highlights the unexpected advantages associated with the use of surface-modified cellulose nanofibers.

[0012] The mechanical properties of developed hydrogels in this disclosure can be tuned by varying the concentration of the surface-modified cellulose nanofibers, for example aCNFs.

[0013] Therefore, the present disclosure includes a composite hydrogel comprising a decellularized extracellular matrix (dECM) and surface-modified cellulose nanofibers (CNFs).

[0014] In some embodiments, the hydrogel of the disclosure is used in tissue modelling such as 2D and 3D cell culture, tissue engineering, as bioinks for bioprinting, analytical applications such as microfluidic platforms and gel contraction assays, and drug screening and delivery.

[0015] The present disclosure also includes a method of preparing the composite gel of the disclosure.

[0016] Other features and advantages of the present disclosure will become apparent from the following detailed description. It should be understood, however, that the detailed description and the specific examples, while indicating embodiments of the disclosure, are given by way of illustration only and the scope of the claims should not be limited by these embodiments, but should be given the broadest interpretation consistent with the description as a whole.

BRIEF DESCRIPTION OF THE DRAWINGS

[0017] Certain embodiments of the disclosure will now be described in greater detail with reference to the attached drawings in which:

[0018] FIGURE 1 shows the impact of dECM-based hydrogels on their rheological properties in exemplary embodiments of the disclosure. A) storage stimulus; B) loss modulus; C) compression modulus; and D) the ratio of storage modulus (G¹) and loss modulus (G") - for hydrogels with 0 % (dECM), 0.5 % w/v (RdECM 0.5%, exemplary) and 1 % w/v (RdECM 1.0%, exemplary) of surface- modified cellulose nanofiber (CNF).

[0019] FIGURE 2 shows (A) SEM images of dECM-based hydrogels and (B) Fiber diameter; C) Pore size; and D) Pore area) analysis of hydrogel morphology and structures for hydrogels with 0 %, 0.5 % w/v (exemplary) and 1 % w/v (exemplary) of surface-modified CNF.

[0020] FIGURE 3 shows the response of human lung fibroblasts (HLF)s to 3D Culture in dECM-Based Hydrogels in exemplary embodiments of the disclosure. A)- B) Rate of Contraction on day 3 and day 7; C) metabolic activity; and D) aSMA protein expression - for HLF with hydrogels with 0 %, 0.5 % w/v (exemplary) and 1 % w/v (exemplary) of surface-modified CNF.

[0021] FIGURE 4 shows the A) Morphology of HLFs Cultured on the surface of dECM Hydrogels with Mechanical Properties (B) Cell area; C) Roundness; and D) Elongation) Representing Soft (dECM) and Stiff Environments (dECM and 1 % w/v (exemplary) of surface modified CNF) for Cells in exemplary embodiments of the disclosure.

[0022] FIGURE 5A-D shows a Single Cell Analysis of HLFs Cultured on Soft dECM Hydrogels and Reinforced dECM Hydrogels with exemplary 1% w/v surface- modified CNF Representing Stiff/Fibrotic Environments. A) Soft; B) Stiff; C) aSMA protein expression; and D) metabolic activity.

[0023] FIGURE 6 A-C shows the YAP Localization Differences in HLFs Cultured on Soft (dECM) vs. Stiff (exemplary RdECM 1.0% - surface-modified CNF) Hydrogels.

[0024] FIGURE 7 shows the exemplary preparation and proposed chemistry of exemplary aldehyde-modified cellulose nanofiber (aCNF)-reinforced lung dECM hydrogels. (A) Fresh porcine lung is minced, subjected to sequential physical agitation and detergent/enzymatic washes to remove cellular material, briefly dehydrated, lyophilized, and cryomilled to a powder. The powder (35 mg mL ¹) is solubilized for 36 h at RT in an acidic pepsin solution (2.45 mg mL ¹) and then neutralized on ice (pH ~ 7.2) to yield a clear pre-gel dECM solution. Chilled aldehyde-functionalized CNFs are dispersed into the pre-gel at 0.3 % or 0.6 % w/v with gentle vortexing. Two workflow options are used for cell introduction: (i) the aCNF-dECM mixture is cast into well-plates, incubated 2 h at 37 °C to gel, and cells are then seeded on the surface; (ii) cells are mixed into the aCNF-dECM pre-gel, gently pipetted for homogeneity, cast, and gelled to encapsulate the cells in 3-D. (B) Macroscopic appearance of fully gelled constructs in culture medium: native dECM (left), reinforced dECM containing 0.3 % aCNF (center) and 0.6 % aCNF (right), demonstrating uniform, self-supporting discs. (C) Proposed reinforcement mechanism: surface aldehyde groups on aCNFs react with e-amino groups of collagen within the dECM, forming dynamic Schiff-base imine linkages that tether nanofibers to the protein network and create an interpenetrating, mechanically robust matrix while preserving nanoscale fibrillarity.

[0025] FIGURE 8 shows exemplary aldehyde-functionalized cellulose nanofibers (aCNF) reinforcement, which stiffens lung-derived dECM hydrogels and improves their mechanical stability while preserving viscoelastic behavior. (A) Storage modulus (G') rises sequentially from native dECM to exemplary RdECM 0.3 % and RdECM 0.6 % hydrogels; (B) loss modulus (G") shows a proportional increase; (C) compression modulus tests reveal the corresponding increase in elastic modulus (E); (D) normalized stress-relaxation curves recorded at 10 % compressive strain display similar decay profdes for all formulations; (E) the half-relaxation time (t!4) extracted from (D) remains statistically unchanged, confirming that aCNF addition preserves intrinsic viscoelasticity; (F) equilibrium swelling ratio after 48 h immersion in culture medium decreases with increasing aCNF content, indicating a denser hydrogel network; (G) residual mass after 15 days in medium demonstrates reduced hydrolytic degradation in reinforced gels; (H) cumulative release of 70 kDa FITC-dextran over 30 h is attenuated by the tighter RdECM mesh, at the time point of 30h, top line is dECM, middle line is exemplary RdECM 0.3 % w/v and bottom line is exemplary RdECM 0.6 % w/v; (I-K) scanning electron micrographs highlight the progressive transition from a loose fibrillar matrix in dECM (I) to increasingly interpenetrating collagen-aCNF networks in exemplary RdECM 0.3 % (J) and exemplary RdECM 0.6 % (K) (scale bar = 50 pm). Each data point represents a single hydrogel puck (n = 4 for all other tests). One-way ANOVA with Tukey post-hoc analysis was applied; *p < 0.05 vs. dECM, fp < 0.05 between RdECM 0.3 % and 0.6 %.

[0026] FIGURE 9 shows exemplary stiff aCNF-reinforced hydrogels up- regulate myofibroblastic a-SMA without markedly altering cell or nuclear morphology. (A) Representative confocal microscopy images of human lung fibroblasts cultured for 72 h on native dECM (“Soft”) or exemplary RdECM 0.6 % (“Stiff’) hydrogels. Nuclei are stained with DAPI (blue), F-actin with phalloidin (green), and a-smooth-muscle actin (a-SMA) with monoclonal antibody (red); merged images highlight greater a-SMA fibre formation on stiff matrices (scale bar = 100 pm). (B) Normalised integrated a-SMA fluorescence intensity for four independent donors shows a significant increase on stiff hydrogels (mean ± SD; one-way ANOVA with Tukey post-hoc, *p < 0.05). (C) Cell spread area, (D) nuclear projected area, (E) cell eccentricity, and (F) nuclear eccentricity were extracted from masked phalloidin and DAPI channels; no statistically significant differences were detected between groups, although a trend toward reduced spread area and slightly higher elongation on the soft substrate was observed (n > 40-80 cells per donor, four donors). These data indicate that while covalent stiffening of the dECM matrix potentiates myofibroblastic a-SMA expression, overall cell and nuclear morphology remain largely conserved. [0027] FIGURE 10 shows that exemplary stiff aCNF -reinforced hydrogels drive nuclear translocation of the mechanosensitive transcriptional co-activator YAP. (A) Representative confocal microscopy images of primary human lung fibroblasts cultured for 72 h on native dECM (“Soft”) versus exemplary RdECM 0.6 % (“Stiff’) hydrogels. Nuclei are stained with DAPI (blue); YAP is immunolabelled (Red); phalloidin-stained F-actin outlines cell bodies (green); merged images highlight the pronounced translocation of YAP within nuclei on the stiff substrate (scale bar = 50 pm). (B) Quantification of YAP localization expressed as the nuclear-to-cytoplasmic fluorescence-intensity ratio. Each symbol represents the average of > 40-60 cells from an individual donor (three donors). An unpaired two-tailed t-test revealed a significant increase in nuclear YAP on stiff hydrogels, confirming effective transmission of the higher matrix stiffness to the Hippo- YAP pathway.

[0028] FIGURE 11 shows that exemplary matrix stiffness rather than integrin inhibition governs 3-D fibroblast-mediated contraction of lung dECM hydrogels. (A) Experimental timeline: human lung fibroblasts (three independent donors) were encapsulated in native dECM (“Soft”) or exemplary RdECM 0.6 % (“Stiff’) hydrogels (1 x 10⁶ cells mL ¹). cultured for seven days, and exposed on day 4 to the av -integrin antagonist CWHM-12 (1 pM) or vehicle. (B) Macroscopic top-view images of representative hydrogel pucks at day 7 illustrate the pronounced bulk contraction of Soft constructs and the resistance of Stiff constructs, with minimal visual impact of drug treatment (scale bar = 12 mm). (C-E) Quantification of gel contraction expressed as the final-to-initial radius ratio (R/Ro) for donors 1-3, respectively (mean ± SD). Across all donors, Soft hydrogels contracted significantly more than Stiff hydrogels (one-way ANOVA, *p < 0.05 versus Stiff). CWHM-12 did not appreciably alter contraction of either matrix, except in donor 1 where drug addition modestly reduced Soft-gel contraction (fp < 0.05 versus Soft vehicle).

[0029] FIGURE 12 shows that exemplary integrin blockade partially counteracts stiffness-induced myofibroblast activation in 3-D dECM hydrogels. (A) SUM-intensity projections of confocal z-stacks acquired on day 7 from fibroblasts encapsulated in native dECM (Soft), Soft treated with CWHM-12 (1 pM, added on day 4), exemplary RdECM 0.6 % (Stiff), or Stiff + CWHM-12. Nuclei (DAPI, blue), F-actin (phalloidin, green), and a-smooth-muscle actin (a-SMA, red) are shown individually and merged; robust a-SMA fibers are evident in Stiff gels and are reduced by drug treatment (scale bar = 100 gm). (B) Fold-change in integrated a-SMA fluorescence normalized to the Soft control for three independent donors (means from > 20 - 60 cells). Stiff gels significantly up-regulate a-SMA versus Soft, whereas CWHM-12 lowers a-SMA on both matrices, with donorspecific magnitude: moderate reduction (donor 1), return to Soft level (donor 2), and partial reduction (donor 3). (C) Volume-rendered 3-D views (nucleus + F-actin) illustrate increased cellular packing density and altered morphology in contracted Soft constructs compared with Stiff counterparts. One-way ANOVA with Tukey post-hoc test: *p < 0.05 versus Soft; fp < 0.05 versus corresponding vehicle control.

[0030] FIGURE 13 shows exemplary donor-dependent modulation of fibroblast and nuclear morphology by matrix stiffness and av-integrin blockade. SUM-intensity projections of confocal z-stacks (acquired and processed as described in Fig. 12) were analysed to obtain single-cell two-dimensional morphometries for three independent donors after 7 days of culture in native dECM (Soft), Soft + CWHM-12 (1 pM. added on day 4), exemplary RdECM 0.6 % (Stiff) and Stiff + CWHM-12. (A) Projected cell spread area shows a modest reduction on Stiff relative to Soft in donors 1 and 3, whereas donor 2 exhibits a slight increase, highlighting inter-donor variability; the drug produces marginal additional changes. (B) Projected nuclear area mirrors the cell-area trend, with a general but not universal decrease on Stiff and subtle drug effects. (C) Cell eccentricity (1 - minor/major axis) reveals small stiffness-dependent shifts towards elongation in some donors, again influenced idiosyncratically by CWHM-12. (D) Nuclear eccentricity remains largely unchanged across conditions. Symbols represent individual cells (> 20- 60 per donor per condition); horizontal lines denote donor means ± SD. One-way ANOVA per donor with Tukey post-hoc testing detected no consistent group-wide significance, indicating that morphological responses to covalent stiffening and integrin inhibition are highly donor-specific.

[0031] FIGURE 14 shows comparison of mechanical properties of dECM- based hydrogels reinforced with aldehyde-functionalized cellulose nanocrystals (aCNC) or nanofibers (aCNF, exemplary). (A) Storage modulus (G'), (B) loss modulus (G"), and (C) compression modulus of native lung-derived dECM hydrogels and hydrogels reinforced with either aldehyde-modified CNCs (aCNC, 0.5 % and 1.0 % w/v) or CNFs (aCNF, 0.5 % and 1.0 % w/v). Mechanical testing was performed at 37 °C using small -amplitude oscillatory rheology and unconfined compression. Incorporation of aCNFs significantly increased all three mechanical parameters in a concentration-dependent manner, while aCNCs had a minimal effect on hydrogel stiffness. Data are presented as individual values with mean ± SD. Statistical analysis by one-way ANOVA with Tukey's post-hoc test; p-values are indicated above brackets.

DETAILED DESCRIPTION

I, Definitions

[0032] Unless otherwise indicated, the definitions and embodiments described in this and other sections are intended to be applicable to all embodiments and aspects of the present disclosure herein described for which they are suitable as would be understood by a person skilled in the art. It is also to be understood that the terminology used herein is for the purpose of describing particular aspects only and is not intended to be limiting.

[0033] In understanding the scope of the present disclosure, the term “comprising” and its derivatives, as used herein, are intended to be open ended terms that specify the presence of the stated features, elements, components, groups, integers, and/or steps, but do not exclude the presence of other unstated features, elements, components, groups, integers and/or steps. The foregoing also applies to words having similar meanings such as the terms, “including”, “having” and their derivatives. The term “consisting” and its derivatives, as used herein, are intended to be closed terms that specify the presence of the stated features, elements, components, groups, integers, and/or steps, but exclude the presence of other unstated features, elements, components, groups, integers and/or steps. The term “consisting essentially of’, as used herein, is intended to specify the presence of the stated features, elements, components, groups, integers, and/or steps as well as those that do not materially affect the basic and novel characteristic(s) of features, elements, components, groups, integers, and/or steps.

[0034] Terms of degree such as “substantially”, “about” and “approximately” as used herein mean a reasonable amount of deviation of the modified term such that the end result is not significantly changed. These terms of degree should be construed as including a deviation of at least ±5% of the modified term if this deviation would not negate the meaning of the word it modifies. In addition, all ranges given herein include the end of the ranges and also any intermediate range points, whether explicitly stated or not.

[0035] As used in this disclosure, the singular forms “a”, “an” and “the” include plural references unless the content clearly dictates otherwise.

[0036] In embodiments comprising an “additional” or “second” component, the second component as used herein is chemically different from the other components or first component. A “third” component is different from the other, first, and second components, and further enumerated or “additional” components are similarly different.

[0037] The term “and/or” as used herein means that the listed items are present, or used, individually or in combination. In effect, this term means that “at least one of’ or “one or more” of the listed items is used or present.

[0038] The abbreviation, “e.g.” is derived from the Latin exempli gratia and is used herein to indicate a non-limiting example. Thus, the abbreviation “e.g.” is synonymous with the term “for example.” The word “or” is intended to include “and” unless the context clearly indicates otherwise.

[0039] The term “surface-modified cellulose nanofiber” refers to cellulose nanofiber which is modified to contain appropriate reactive functional groups on the surface of said cellulose nanofiber.

[0040] The term “cellulose nanofiber” or “CNF” refers to cellulose fibrils with a high aspect ratio (length to width ratio, width of the CNF is greater than or equal to 20 nm and length of greater than or equal to 1 micron), as compared to cellulose nanocrystals (CNC), which are rod-like particles with low aspect ratio (width of the CNC is less than 20 nm and length of between 100 to 250 nm). The aspect ratio can be measured by any method known in the art, such as dynamic laser scattering, transmission electron microscopy (TEM), and particle shape analysis software.

[0041] The term “composite hydrogel of the disclosure” or “composite hydrogel of the present disclosure”, and variations thereof, refers to a hydrogel that contains dECM and surface-modified CNF as defined in the disclosure.

[0042] It will be understood that any component defined herein as being included may be explicitly excluded by way of proviso or negative limitation, such as any specific compounds or method steps, whether implicitly or explicitly defined herein.

II. Composite Hydrogels of the Disclosure and Uses Thereof

[0043] The present disclosure includes a composite hydrogel comprising a decellularized extracellular matrix (dECM) and surface-modified cellulose nanofibers (CNFs).

[0044] In some embodiments, the CNFs used in the composite hydrogel of the disclosure have a width of greater than or equal to 20 nm and length of greater than or equal to 1 micron. In some embodiments, the CNFs have a width of about 30 nm to about 60 nm and a length of about 1 micron to about 200 microns.

[0045] In some embodiments, the surface-modified CNFs comprise one or more functional groups on a surface of the CNFs, wherein the groups are selected from ehyde Jt_u ald ^H ), methacrylate ( ° ), carboxylate .- n ° , carboxylic acid

O H , vinyl thiol , alkyne azide ^N . hydrazide ^H and the like.

[0046] In some embodiments, the surface-modified CNFs comprise methacrylate functional groups on the surface of the CNFs. In some embodiments, the surface-modified CNFs comprise aldehyde functional groups on the surface of the CNFs.

[0047] In some embodiments, modifying the surface of the CNFs by introducing functional groups on the surface of the cellulose allows the CNFs to be uniformly dispersed in the composite gel of the present disclosure. In some embodiments, the functional groups on the surface of the CNFs act as crosslinkers in the hydrogel by reacting with amino groups in polymers or biomolecules present in the dECM. Thus, surface-modified CNFs facilitate the formation of three-dimensional hydrogel network.

[0048] In some embodiments, the surface-modified CNFs are present in the hydrogel of the disclosure in an amount of about 0.1 % w/v to about 2 % w/v. In some embodiments, the surface-modified CNFs are present in the hydrogel of the disclosure in an amount of about 0.3 % w/v, about 0.5 % w/v, about 0.6 % w/v, or about 1 % w/v, and values therebetween.

[0049] In some embodiments, the surface-modified CNFs are present in the hydrogel of the disclosure in an amount of about 0.1 % w/v to about 0.5 % w/v to mimic healthy tissues. In some embodiments, the surface-modified CNFs are present in the hydrogel of the disclosure in an amount of about 0.6 % w/v to about 2 % w/v to mimic diseased tissues.

[0050] In some embodiments, the dECM is a digested dECM.

[0051] In some embodiments, the dECM comprises ECM from a mammalian tissue, such as human or porcine tissue and the like. In some embodiments, the dECM comprises ECM from one or more of lung, liver, brain skin, kidney, or heart or the like. In some embodiments, the dECM comprises ECM from lung tissue.

[0052] In some embodiments, the dECM is present in the hydrogel of the disclosure in an amount of about 1 % w/v to about 3 % w/v. In some embodiments, the dECM is present in the hydrogel of the disclosure in an amount of about 1 % w/v, about 1.25 % w/v, or about 1.5 % w/v and values therebetween.

[0053] In some embodiments, the weight ratio of the dECM to the surface- modified CNFs is about 30:1 to about 1:2. In some embodiments, the weight ratio of the dECM to the surface-modified CNFs is about 5:1, about 2.5:1, or about 1.2:1.

[0054] In some embodiments, the composite hydrogel of the disclosure further comprises additives such as, but not limited to, one or more of cytokines, cells, cell culture media, and bioactive growth factors and the like.

[0055] In some embodiments, the hydrogel of the disclosure is seeded with one or more cells or populations of cells. In some embodiments, the cells are selected from one or more of vascular cells, lung cells, cardiac cells, muscle cells, neural cells, endocrine cells, paracrine cells bone cells, and dermal cells, and the like. In some embodiments, the cells are selected from one or more of fibroblasts, endothelial cells, epithelial cells, pericytes, osteocytes, and neurocytes, and the like.

[0056] In some embodiments, the functional groups on the surface of the CNFs are further functionalized and/or conjugated to bioactive molecules. In some embodiments the bioactive molecules are therapeutic agents. In some embodiments, the bioactive molecules are selected from antibodies, peptides, sugars, carbohydrates, nucleic acids, and small molecule drugs.

[0057] In some embodiments, the composite hydrogel of the disclosure is a fibrous hydrogel.

[0058] In some embodiments, the composite hydrogel of the disclosure possesses improved mechanical properties, such as increased storage modulus, loss modulus, and compression modulus, compared to mechanical properties of the same hydrogel but without the surface-modified CNF.

[0059] In some embodiments, the surface-modified CNF reinforces the composite hydrogel of the disclosure. In some embodiments, the composite hydrogel of the disclosure is reinforced while the physical structures of the hydrogel, such as fiber’s dimension and porosity, redistribution of internal stress and viscoelastic properties remain substantially unchanged compared to the same hydrogel but without the surface-modified CNFs.

[0060] In some embodiments, the composite hydrogel of the disclosure has a compression moduli of greater than or equal to 7kPa. In some embodiments, the composite hydrogel of the disclosure has compression moduli of about 7kPa to about 25kPa. In some embodiments, the composite hydrogel of the disclosure has compression moduli of about 8 kPa, about lOkPa, about 13kPa, about 16kPa, or about 20kPa, and values therebetween.

[0061] In some embodiments, increasing the concentration of the CNFs in the composite hydrogel increases the compression moduli of the hydrogel, thus producing a stiffer hydrogel. Thus, in some embodiments, the mechanical properties of the composite hydrogel of the disclosure are tuned by adjusting the concentration of the CNF to mimic various stages of healthy and diseased tissues, while preserving fibrous structure that resembles native tissues.

[0062] In some embodiments, the composite hydrogel of the disclosure has a reduced hydrolytic degradation, compared to a hydrolytic degradation of the same hydrogel but without the surface-modified CNF of the present disclosure. In some embodiments, the composite hydrogel of the disclosure has a hydrolytic degradation of less than 10% after 15 days. In some embodiments, the composite hydrogel of the disclosure has a hydrolytic degradation of between about 0% to about 10% after 15 days.

[0063] In some embodiments, the composite hydrogel of the disclosure has similar macromolecule permeability, compared to a macromolecule permeability of the same hydrogel but without the surface-modified CNFs of the present disclosure. In some embodiments, the macromolecule permeability of the composite hydrogel of the disclosure is within +/- 5% macromolecule permeability, compared to a macromolecule permeability of the same hydrogel but without the surface-modified CNFs of the present disclosure.

[0064] The present disclosure includes a use of the composite hydrogels of the disclosure in the following applications: tissue modelling such as 2D and 3D cell culture and the like, tissue engineering, as bioinks for bioprinting, analytical applications such as microfluidic platforms, gel contraction assays and the like, and drug screening and delivery.

[0065] In some embodiments, the composite hydrogels of the disclosure are used to model healthy and diseased tissues.

[0066] In some embodiments, the composite hydrogels of the disclosure are used to form a 3D model for fibroblast-to-myofibroblast transition, which occurs in wound repair, regeneration, and fibrosis.

[0067] In some embodiments, the composite hydrogels of the disclosure are used to model pulmonary fibrosis.

II, Methods of Preparing the Composite Hydrogels of the Disclosure

[0068] The disclosure also includes a method of preparing the composite hydrogels of the disclosure. In some embodiments, the method comprises combining a decellularized extracellular matrix (dECM) and a surface-modified cellulose nanofibers (CNFs) to form the composite hydrogels.

[0069] To form the dECM, a tissue is decellularized by any method known in the art, for example by a method described in M. Dabaghi et al. , “A Robust Protocol for Decellularized Human Lung Bioink Generation Amenable to 2D and 3D Lung Cell Culture,” Cells, vol. 10, no. 6, p. 1538, Jun. 2021. [0070] In some embodiments, the dECM obtained by decellularization is in the form of an aqueous suspension. In some embodiments, the dECM obtained by decellularization is in the form of a powder, obtained by drying the dECM aqueous suspension by for example lyophilization and then grinding to obtain fine powder.

[0071] In some embodiments, the dECM is further digested to obtain a digested dECM, by any method known in the art, for example by enzymatic digestion with pepsin under acidic conditions. In some embodiments, when the dECM is in the form of a powder, said powder is solubilized prior to or simultaneously with the digestion step. The resulting dECM is in the form of a digested dECM liquid solution.

[0072] In some embodiments, the dECM is present in the digested dECM liquid solution in an amount of about 20 mg/ml to about 30 mg/ml.

[0073] In some embodiments, the dECM undergoes one or more purification steps throughout the steps of preparing the digested dECM solution.

[0074] In some embodiments, the dECM is in the form of a liquid solution. In some embodiments, the dECM liquid solution comprises a buffer. In some embodiments, the buffer is phosphate-buffered saline (PBS). In some embodiments, the pH of the dECM liquid solution is from about 7.2 to about 7.4.

[0075] In some embodiments, the surface-modified CNFs comprise hydroxyl groups on the surface of CNFs.

[0076] Surface-modified CNFs are either commercially available or is prepared using any method known in the art. For example, hydroxyl groups on a surface of CNFs are oxidized to obtain reactive functional groups including carboxyl or aldehyde, which are optionally further modified using known reactants to obtain other functional reactive groups. In some embodiments, a portion of the functional groups on the surface of the CNF are further functionalized and/or conjugated to bioactive molecules.

[0077] In some embodiments, the resulting surface-modified CNFs are in the form of an aqueous suspension. In some embodiments, the aqueous suspension of the surface-modified CNFs comprises a buffer. In some embodiments, the buffer is Hank's Balanced Salt Solution (HBSS).

[0078] In some embodiments, the surface-modified CNFs are used as a concentrated suspension obtained from preparations as described above. [0079] In some embodiments, the surface-modified CNFs undergo one or more purification steps throughout the preparation steps.

[0080] In some embodiments, the step of combining the dECM solution with the surface-modified CNF is carried out at a temperature of about 0°C. In some embodiments, both the dECM and the surface-modified CNFs are cooled to about 0°C prior to being combined.

[0081] In some embodiments, the method of preparing the composite hydrogels of the disclosure further comprises seeding a population of cells on the hydrogel of the disclosure. In this embodiment, 5000 cells/cm² to 50,000 cells/cm² are seeded on the hydrogel of the disclosure.

[0082] In some embodiments, the method of preparing the composite hydrogels of the disclosure further comprises encapsulating a population of cells in the hydrogel of the disclosure, while the cells are added at the step of combining the dECM solution with the surface-modified CNF. In this embodiment, 0.5*10⁶ cells/ml to 4*10⁶ cells/ml can be encapsulated in the hydrogel of the disclosure.

[0083] In some embodiments, the method of preparing the composite hydrogels of the disclosure further comprises adding additives such as, but not limited to, one or more of cytokines, cells, cell culture media, and bioactive growth factors and the like.

[0084] The present disclosure also includes the following embodiments:

1. A hydrogel composition comprising decellularized extracellular matrix (dECM) and cellulose-based nanomaterials.

2. The hydrogel composition of embodiment 1, wherein the cellulose-based nanomaterials comprise and cellulose nanofibers (CNFs).

3. The hydrogel composition of embodiment 1 or 2, wherein the CNFs comprise surface functionalized CNFs.

4. The hydrogel composition of embodiment 1 or 2, wherein the surface modifications of the CNFs comprise vinyl, thiol, alkyne, azide, hydrazide modifications.

5. The hydrogel composition of any one of embodiments claims 1 to 3, wherein the CNFs comprise carboxylated aldehyde-modified CNFs (aCNFs). 6. The hydrogel composition of embodiment 1 or 2, wherein the CNFs comprise cellulose nanofiber methacrylate (CNF-MA).

7. The hydrogel composition of any one of embodiments 1 to 4, wherein the aldehyde and carboxylate modifications i) act as crosslinking agents, facilitating the formation of three-dimensional hydrogel networks when reacting with groups present in polymers or biomolecules, and ii) allows for bioconjugation and functionalization, enabling the attachment of bioactive molecules and facilitating controlled release of encapsulated substances.

8. The hydrogel composition of any one of embodiments 1 to 4, wherein the concentration of cellulose nanomaterials can be adjusted to produce a range of mechanical properties.

9. The hydrogel composition of embodiment 1, wherein the dECM comprises ECM from a mammalian tissue.

10. The hydrogel composition of embodiment 1, wherein the dECM comprises ECM from lung, liver, brain skin, kidney, heart.

11. The hydrogel composition of embodiment 1, wherein the dECM can be functionalized with methacrylic anhydride to generate methacrylated dECM (MA- dECM). The formation of photocrosslinkable MA-dECM and CNC-MA/CNF-MA on demand.

12. The hydrogel composition of embodiment 1 wherein additives can be incorporated into the hydrogel, comprising but not limited to cytokines, cells, cell culture media, bioactive growth factors.

13. The hydrogel of any one of embodiments 1 to 12 that can be used for applications comprising: microfluidic platforms, tissue engineering, drug delivery, bioprinting, bioinks, gel contraction assays, wound healing, drug screening, 2D and 3D cell culture.

EXAMPLES

[0085] The following non-limiting examples are illustrative of the present disclosure:

Example 1 : Methods

[0086] Tissue Decellularization for Isolation of ECM: Lung tissue decellularization process involves several steps, as reported in previous study [39], Fresh pig lung (pig lungs were generously donated by Jamie Waldron Butchers, Hamilton, ON, Canada) tissues were chopped and immediately frozen in liquid nitrogen, then stored at -80°C before decellularization. The decellularization began by soaking the tissues in 0.1% Triton X-100 (Sigma Aldrich, Canada, Catalog number: T9284-500ml) in a 4-liter beaker with continuous stirring for 30 minutes. The tissues were then filtered through a mesh strainer to remove the liquid, followed by a 30-minute wash in phosphate-buffered saline (PBS) to remove any remaining Triton X-100. The tissues were subsequently treated with 2% sodium deoxy cholate (SDC, Sigma Aldrich, Canada, Catalog number: D6750-100G) and left overnight at 4°C. The next day, the SDC was removed, and the tissues were washed for 24 hours in PBS with an antibiotic- antimycotic (ThermoFisher, Canada, Catalog number: 15240062) solution, including at least one change of PBS. After the antibiotic-antimycotic wash, the tissues were agitated in sterilized deionized (DI) water for two hours, then washed for 30 minutes in IM sodium chloride. Following this, tissues were rinsed in DI water and treated with 1% Triton X-100. A final wash in PBS with antibiotic-antimycotic solution lasted 3 days. After these washes, the tissues were agitated in sterilized DI water for 30 minutes, strained, and stored at -80°C for lyophilization using a Benchtop Freeze Dryer (FreeZone 2.5 Liter -84C, Labconco). Post-freeze-drying, the tissues were ground into fine dECM powders in the presence of liquid nitrogen. These powders were then stored at -20°C for future experimental purposes.

[0087] Surface Modification of CNFs with Aldehyde Groups: In a roundbottom flask, a dispersion of CNFs (2 g of CNFs; Cellulose Lab, New Brunswick, Canada, Product Number: CNF-Slurry-SMC, Width: nominal width 30-80 nm; Length: up to several hundred micron, surface group: hydroxyl) was prepared in DI water forming a 1.0 wt % CNF suspension containing 0.2 mM TEMPO (Sigma Aldrich, Canada, Catalog number: 214000-5G) and 2 mM sodium bromide (Sigma Aldrich, Canada, Catalog number: 310506-100G). To initiate the reaction, 6 mL sodium hypochlorite (containing 10-15 % chlorine; product # 425044, Sigma Aldrich) dissolved in 50 mL of DI water and then was added to the flask. The resulting suspension was stirred at room temperature at a constant rpm for 24 hours, while maintaining the pH at ~10 using 1 M NaOH. The reaction was quenched by introducing 1 mL of ethylene glycol (Sigma Aldrich, Canada, Catalog number: 324558-100ML). The obtained product underwent dialysis through 14 water changes against DI water using a cellulose membrane (12 - 14 kDa cutoff; Sigma Aldrich, Canada, Catalog number: D9652-100FT). To further modify the reactants, a surface oxidation process was conducted using sodium periodate (NaI04; Sigma Aldrich, Canada, Catalog number: 311448-100G) to yield surface-modified CNFs with aldehyde functional groups. The NaI04 and suspension were mixed at a mass ratio of 1:1. The flask was covered with aluminum foil to protect the NaI04 from photodegradation. The resulting mixture was continuously stirred at room temperature for 24 hours. The oxidation reaction was then halted by the addition of ethylene glycol (1 mL). Subsequently, the aCNFs suspension underwent dialysis against DI water with 14 water changes. Finally, the suspension was concentrated using evaporation of excess water.

[0088] dECM-Based Hydrogels reinforced with aCNFs: To digest and solubilize dECM powder, dECM powder with a concentration of 22 mg/mL was added to an acidic solution containing pepsin (Sigma Aldrich, Canada, Catalog number: P6887-5G) with a concentration of 1 mg/mL. As pepsin enzymatic activity is optimum at pH of 2, the pH of the solution was adjusted and monitored, ensuring it remained within the range of 2 to 3. If the pH exceeded 3, 1 N HC1 was added to lower it accordingly. Stirring was maintained at room temperature (RT) for a duration of three days. After 24 hours, the pH was measured and adjusted if necessary. Once the digestion process was complete, the dissolved dECM underwent centrifugation at approximately 4000 g for 15 minutes. The resulting supernatant, containing the dissolved dECM, was passed through a series of cell strainers with mesh sizes of 200 pm, 100 pm, 60 pm, and 30 pm and transferred to conical tubes, while the nondissolved materials were discarded. To cool down, the tubes containing the dissolved dECM were placed on ice alongside an equal volume of lOx PBS that was also kept on ice. To reach a final concentration of 20 mg/mL of digested dECM solution, lOx PBS was added to the dissolved dECM, representing 10% of its volume. The pH of the dissolved dECM was subsequently adjusted to approximately 7.2-7.4 using 10 N and 1 N NaOH. Throughout the process, all steps were performed on ice. For the preparation of dissolved dECM with various concentrations, ice-cold PBS at a pH of 7.2-7.4 was added to the stock of dissolved dECM (concentration = 20 mg/mL).

[0089] To create reinforced dECM-based hydrogels containing aCNFs, aCNF suspensions were buffered with 10X Hank's Balanced Salt Solution (HBSS) and then integrated at concentrations of 0.5% and 1% with neutralized digested dECM precursors. These hydrogels, enhanced with aCNFs, were categorized as RdECM (Reinforced dECM) 0.5% and RdECM 1.0%, indicating the respective aCNF concentration within the hydrogels. Each hydrogel formulation consistently incorporated 12.5 mg/mL of digested dECM, while the aCNF content varied, being present at concentrations of 0%, 0.5%, or 1.0% by weight.

[0090] Hydrogel Rheology Measurements: The rheological properties of all hydrogels at different concentrations were evaluated using a DHR Controlled Stress Single Head CMT rheometer (HR20, TA instruments, New Castle, DE, USA), employing a parallel plate geometry. Oscillation frequency sweeps were utilized to measure the storage (G¹) and loss (G") modulus of the samples, with angular frequencies ranging from 0. 1 to 100 rad/s and a shear strain fixed at 0.6%. To derive a consolidated G' and G" value for each sample, the values from the plateau region were averaged, specifically between 1 and 10 rad/s. The compressive moduli of all hydrogels were determined by compressing them to 100 pm (equivalent to 10% strain) at a steady linear speed of 5 pm/s. The reported values were calculated from the slope found in the linear portion of the strain versus stress curve. For the generation of materials for rheology measurements, the hydrogel solutions were added to polydimethylsiloxane (PDMS) molds and gelled in an incubator. To fabricate PDMS molds, the PDMS monomer and curing agent were combined in a 10: 1 ratio, degassed in a desiccator for approximately 30 minutes, and then cured overnight at 65 °C. The PDMS molds were rinsed with 70% ethanol, dried in a biosafety cabinet, and autoclaved prior to the addition of dECM hydrogels. Then, the hydrogel pregel solutions were added to PDMS molds and gelled overnight in an incubator before conducing rheological measurements.

[0091] Cell Culture: Human lung fibroblasts (HLFs) derived from lung tissue samples donated for medical research (Hamilton Integrated Research Ethics Board - HiREB - 5305-T) were initially isolated and cultured in DMEM (ThermoFisher, Canada, Catalog number: 11965118) supplemented with 10% fetal bovine serum (FBS, Wisent Inc., Saint-Jean-Baptiste, Canada, product number: 080-450) and 1 % penicillinstreptomycin (Gibco, United States, Catalog number: 15140122). Custom PDMS molds (12 mm in diameter with the height of 4 mm) were used for shaping dECM hydrogel pucks. Concentrated HLFs were suspended in precursor dECM hydrogel solutions at varying concentrations of aCNFs to achieve a final cell density of 1 x 10⁶ cells/mL when encapsulating cells within hydrogels. The cells were gently and thoroughly mixed with the precursor dECM hydrogel solutions and poured into the PDMS disks. After incubating at 37 °C for two hours to allow the formation of dECM hydrogels, DMEM media was added to each disk containing the hydrogels, and the samples were incubated overnight. The following day, the dECM hydrogels containing HLFs were detached from the PDMS disks and transferred to well plates, where they were submerged in DMEM. For culturing cells on hydrogels, the final cell density was 1 x 10⁵ cells/cm² Cells were regularly tested with My coStrip™ - Mycoplasma Detection Kit (Invivogen, CA, USA, Product Number: REP-MYS-20).

[0092] To prepare substrates with a soft interface for cell culture, PDMS with a base-to-curing-agent ratio of 70: 1 was mixed, degassed, and added to cell culture flasks. These flasks were then placed on a leveled platform and left undisturbed for at least 72 hours to allow the PDMS to cure fully. To enhance curing, the flasks were subsequently placed in an oven at 75 °C for another 72 hours. After curing, the flasks were washed with isopropanol for 2 hours on a well-plate shaker to remove any uncured or loose PDMS, followed by three thorough washes DI water. The flasks were then air-dried at RT and sterilized under ultraviolet light in a biosafety cabinet for at least one hour.

[0093] A series of surface modifications were performed to improve cell adhesion and growth. Initially, a 0.1 mg/mL Poly-D-Lysine solution (Sigma Aldrich, Canada, Catalog number: A-003-E) was added to each flask and incubated at RT for 2 hours, followed by two washes with autoclaved DI water. This was followed by the addition of a 0.4% glutaraldehyde solution (Electron Microscopy Sciences, Hatfield, PA, USA, with catalog number 16220) to each flask for 30 minutes, then washed three times with autoclaved DI water. After drying thoroughly in a biosafety cabinet, a collagen solution (0.1 mg/mL, Advanced Biomatrix, San Diego, CA, United States; product number: 5056-20ML) was added to each flask and stored at 4 °C for at least 24 hours. Before cell seeding, the collagen solution was removed, and the surface was thoroughly washed three times with PBS.

[0094] Immunoblots of HLFs protein extracts from hydrogels: DMEM was gently aspirated from each well using a pipette, to ensure the hydrogels stayed intact and undamaged. Subsequently, the samples underwent three washes with PBS, each lasting 5 minutes. For the digestion of hydrogels and extraction of proteins, 0.5 mL of 1 mg/mL collagenase (Cedarlane, Canada, Product Number: 7415) was added to the dECM hydrogels, and 1.5 mg/mL to the RdECM hydrogels. The dECM hydrogels were incubated for 1 hour, and the RdECM hydrogels for 1.5 hours, with all incubations conducted at 37 °C. After incubation, an additional 0.5 mL of FBS-enriched DMEM was introduced to each well to deactivate the collagenase. The hydrogels were then manually disrupted by pipetting them up and down, and the contents transferred to 2 mL Eppendorf tubes. These were centrifuged at 4 °C and 4000 rpm for 5 minutes. The supernatant was then discarded, and the samples were placed on ice for 20 minutes, followed by a wash with cold IX PBS. To prepare the lysis buffer solution, 800 pL of RIPA lysis buffer (ThermoFisher, Canada, Catalog number: 89900) was mixed with 100 pL of Protease Inhibitor Cocktail (Sigma Aldrich, Canada, Catalog number: P2714) and 100 pL of PhosSTOP (Sigma Aldrich, Canada, Catalog number: PHOSS-RO). Each sample’s pellet received 100 pL of this lysis buffer, then underwent brief ice-sonication using an ultrasonic homogenizer in short bursts. The samples were left to incubate on ice for another 20 minutes. For cell lysate collection, the solution was centrifuged in a microcentrifuge at 4 °C and 16,000 g for 20 minutes. The supernatant was then transferred to a new tube and stored at -80°C for downstream immunoblots.

[0095] The total protein concentration of the samples was measured using the Pierce™ Bicinchoninic Acid (BCA, ThermoFisher, Canada, Product Number: 23240), following the guidelines provided by the manufacturer. To achieve normalization of these protein concentrations, suitable amounts of PBS were added to each sample. The total cell lysate protein was combined with IX Laemmli Sample Buffer (Bio-Rad Laboratories, product number 1610747). This mixture was then electrophoresed on 4- 20% Mini-PROTEAN TGX stain-free precast gels (Bio-Rad Laboratories, product number 4568093) and the proteins were transferred onto a PVDF membrane using the reagents from the Transfer-Blot Turbo RTA Transfer Kit (Bio-Rad Laboratories, product number 1704272). The membranes were subjected to blocking at room temperature for one hour using a 5% Casein solution (Bio-Rad Laboratories, product number 1706404) in IX Tris Buffered Saline with TWEEN® 20. For protein detection, Clarity Western ECL Substrate (Bio-Rad Laboratories, product number 1705061) was used, and the results were captured using a ChemiDoc MP Imaging System. Image acquisition was set to auto-exposure.

[0096] Hydrogel Staining and Microscopy of HLF: To assess the viability of hydrogels formed under different conditions, the LIVE/DEAD™ Cell Imaging Kit (ThermoFisher, Mississauga, Ontario, with catalog number R37601) was employed. For nuclei staining, the NucBlue™ Live ReadyProbes™ Reagent (Hoechst 33342) from Invitrogen™ was used following the manufacturer's instructions. During the fixation process, three washes with PBS were performed between each staining step and then the samples were incubated with a 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA, with catalog number 15712) for 15 minutes. After fixation, samples were washed with PBS for at least one time. To ensure proper permeabilization, the fixed samples were treated with 0.5% Triton X-100 for 15 minutes. To minimize non-specific staining, a 3% bovine serum albumin (BSA, Wisent Inc., Saint-Jean-Baptiste, Canada, product number: 800-095-EG) solution was used supplemented with 0.05 % Triton X-100 and the samples were incubated for 2 hours. For antibody staining, primary antibodies were incubated with the samples in 1% BSA supplemented with 0.05 % Triton X-100 at 4 °C for 24 hours, followed by 2-hour incubation at RT with secondary antibodies, Hoechst, and phalloidin (Texas Red™-X Phalloidin, ThermoFisher, catalog number T7471) in 1% BSA supplemented with 0.05 % Triton X-100. The fixed cells were stained using a monoclonal anti-a-smooth muscle actin antibody (aSMA, Abeam, Canda, Product Number: ab7817; Clone: 1A4 or Sigma Aldrich, catalog number A2547; Clone: 1A4). Widefield imaging of the cells was conducted using a ThermoFisher EVOS M7000, ZEISS Celldiscoverer 7, or Nikon AIR Inverted Confocal microscopes equipped. For a-SMA expression analysis, the mean gray value within the regions of interest was calculated using ImageJ[40] or Fiji[41] software. Results and Discussion

[0097] Mechanical Properties of Composite Nanofibrillar dECM-based Hydrogels: In this work, CNFs functionalized with aldehyde groups, named as aCNFs, were mixed into a pregel dECM solution at two concentrations and allowed to gel overnight in an incubator at 37 °C. Pristine CNFs are prone to aggregation in an aqueous solution, which limits their ability to disperse uniformly and interact with the components of the dECM hydrogel solution. To address this challenge, the primary hydroxyl groups of CNFs were oxidized to carboxyl groups via TEMPO oxidation, followed by sodium periodate oxidation to introduce aldehyde groups onto the surface of CNFs[40], [41], Covalently crosslinked dECM-based hydrogels were successfully formed through the Schiff-base reach on[42]. The improvement in the mechanical properties of hydrogels was evidenced by the increased storage modulus, loss modulus, and compression modulus, all correlating with the concentration of aCNFs (FIGURE 1).

[0098] As the concentration of aCNFs increases, there is a notable increase in G', with RdECM 1.0 % showing the highest values (FIGURE 1A). While not wishing to be bound by theory, the increase in the storage modulus is likely due to the high aspect ratio and substantial surface area of aCNFs, which provide more points of interaction and crosslinking within the hydrogel network. Similar to the storage modulus, the loss modulus (G”) was also increased by increasing the concentration of aCNFs (FIGURE IB). While not wishing to be bound by theory, the observed increases in G” at higher concentrations of aCNFs could be attributed to the denser packing and entanglement of nanofibers, potentially resulting in greater Schiff-base cross- linking[45]. The compression modulus follows a similar trend to G' and G”, with RdECM 1.0 % showing the highest values (FIGURE 1C). Despite the significant enhancements in both the storage modulus (G¹) and loss modulus (G") with increasing concentrations of aCNF, the ratio of G7G" remained relatively unchanged (FIGURE ID). This consistent ratio value suggests that the addition of aCNF influences both the elastic and viscous components of the hydrogel network proportionally. The unaltered G7G" ratio indicates that while the hydrogels become stiffer and more capable of dissipating mechanical energy due to the denser crosslinking and increased interactions facilitated by aCNF, their fundamental viscoelastic behavior — characterized by the interplay between elasticity and viscosity — does not shift markedly. This observation highlights the ability of aCNF to enhance mechanical properties such as stiffness and energy dissipation without disrupting the inherent viscoelastic balance of the dECM hydrogels, important for maintaining predictable behavior under mechanical stress and deformation similar to dECM hydrogels without any reinforcement.

[0099] The mechanism(s) responsible for the improvement in mechanical properties could be multiple: The functionalizing with aldehyde groups on CNF surfaces will enable reactions with primary amine groups in the ECM proteins [46], There is also a possibility that the mechanical properties of these hydrogels was further enhanced by multiple interactions. For instance, the aCNFs became entwined, adding to the gel's stability. Additionally, the presence of hydrogen bonding interactions between the hydroxyl groups on the aCNFs with the amine groups of ECM proteins is likely to further enhance the hydrogel mechanical properties. Lastly, electrostatic interactions wherein the negatively charged carboxyl groups on aCNFs attract to the positively charged amine groups found in ECM proteins may improve mechanical stability.

[00100] The stiffness (Young’s modulus) of healthy lung tissue is typically around 3.7 ± 1.3 kPa[45], although this value can vary significantly, with a range from 0.5 to 9 kPa according to different reports [46], In contrast, fibrotic lung tissue displays a significantly higher stiffness before decellularization, averaging at 18.9 ± 11.1 kPa[45], The range of values for fibrotic tissue is notably broader, extending to approximately 100 kPa[46], Using these studies, the stiffness of lung tissues fall within the range of 1-5 kPa for healthy tissue and typically exceed 10 kPa for fibrotic tissues[45], [46], [47], [48], After decellularization, dECM hydrogels derived from non-IPF and IPF lungs were reported to have stiffness values of 1.1 ± 0.2 kPa and 6.8 ± 2.8 kPa, respectively, which indicates a significant reduction in stiffness compared to their intact conditions [45], Using these values, dECM hydrogels with a compression modulus of 4.27 ± 2.10 kPa are considered as a normal or soft environment for HLFs. Meanwhile, both RdECM 0.5% and RdECM 1.0% hydrogels, with compression moduli of 8.52 ± 3.15 kPa and 21.35 ± 2.02 kPa respectively, are representative of a stiff or fibrotic environment for HLFs. [00101] Morphological Analysis of Hydrogels: dECM hydrogels are rich in collagen, a key element in the formation of fibrous structure of these hydrogels [49], Collagen, the most abundant protein in the ECM, has a unique ability to form fibrillar networks due to its distinct triple-helix structure, enabling the self-assembly of long, thin fibers integral to the fibrous scaffold of the hydrogel[49], [50], Fibrous hydrogels more closely mimic the natural arrangement and physical interactions found in tissues, which can be important for certain types of cellular responses and for the study of specific mechanotransduction pathways. In this work, composite nanofibrillar lung dECM hydrogels were developed featuring tunable mechanical properties, representing both healthy and fibrotic lungs, while preserving fibrous structures that resemble native lung tissue. The hydrogels were designed such that their physical structures such as fiber’s dimension and porosity remained largely unchanged, even as the stiffness of the hydrogels increased.

[00102] SEM analysis visually confirmed the presence of nanofiber formation in all dECM-based hydrogels FIGURE 2A). SEM imaging allowed for the quantification of three key parameters: fiber diameter, pore size, and pore area (FIGURE 2B, C, and D), following previously established custom image analysis algorithm [39], A slight reduction in fiber diameter of dECM-based hydrogels was observed with the increase in aCNFs content (FIGURE 2B). Specifically, RdECM 1.0% hydrogels exhibited fibers with smaller diameters of 71.8 ± 3.2 nm, in contrast to dECM hydrogels which had diameters of 81.7 ± 5.3 nm. Despite these differences in fiber diameters, the pore sizes and areas within RdECM hydrogels showed no significant difference compared to dECM hydrogels (FIGURE 2C and D). Overall, the incorporation of aCNFs to reinforce dECM hydrogels did not alter the physical structures of the hydrogels at great scale, which could be a factor in how cells interact with these hydrogels.

[00103] Activation of HLF Cultured Within Composite Nanofibrillar dECM Hydrogels: Myofibroblasts are cells that play a crucial role in wound healing and tissue repair by producing and remodeling the ECM[51], [52], [53], However, their persistent activation is a hallmark of pathological fibrosis. In fibrotic diseases, fibroblasts transform into myofibroblasts in response to various stimuli, including mechanical stress, cytokines, and growth factors. This transition is characterized by the de novo expression of a-SMA, which is incorporated into intracellular stress fibers. These stress fibers are contractile structures that increase the cell's ability to contract and exert force on the surrounding ECM[54], [55], The result is a stiffening of the tissue and excessive deposition of ECM components, which is detrimental in chronic conditions and leads to the disruption of tissue architecture and function[56],

[00104] To investigate the viability and activation of HLF within the composite nanofibrillar dECM hydrogels, in vitro experiments were conducted using cells encapsulated in three-dimensional puck structures. Specifically, HLF cells were encapsulated within these composite nanofibrillar dECM hydrogels and cultured for 7 days to assess how changes in the hydrogel's stiffness affect the activation of the HLF cells. Initially, the viability of human lung fibroblasts (HLFs) embedded in the composite nanofibrillar dECM hydrogels was assessed using a Live & Dead assay. All hydrogels maintained cell viability at both Day 3 and Day 7. However, the cellular morphology varied depending on the type of dECM hydrogel and evolved over time. On Day 3, cells in the softer dECM hydrogels predominantly exhibited an elongated morphology, suggesting a more favorable environment for cell spreading. In contrast, cells within the stiffer RdECM hydrogels displayed a mix of elongated and rounded morphologies, with the majority of cells in the RdECM 1.0% hydrogels adopting a rounded form, indicative of a more restrictive mechanical environment. By Day 7, a greater number of cells in the softer dECM hydrogels had maintained or assumed an elongated morphology, while some cells began to show rounded characteristics. A similar trend was observed in both RdECM hydrogel formulations, indicating a consistent response to the hydrogel's stiffness over time. This variability in cell shape over time and between hydrogel formulations highlights the impact of microenvironment stiffness on cellular behavior and adaptation.

[00105] The rate of contraction is higher in soft environments compared to stiff environments. This is due to the greater ability of cells to manipulate and remodel the matrix in a softer environment[57]. In soft environments, cells can exert traction forces more effectively, leading to significant tissue contraction even without the extensive presence of collagen fibrils. Conversely, in stiff environments, while collagen fibrils are important for transferring and maintaining tension, the overall ability of the tissue to contract is reduced due to the greater mechanical resistance of the environment[57]. In this study, a similar pattern of behavior in the contraction of dECM hydrogels was observed as depicted in FIGURE 3A and B. Specifically, the softer dECM hydrogels demonstrated the highest rate of contraction, with a marked decrease in diameter observed from Day 3 to Day 7 (FIGURE 3A and B). Conversely, composite nanofibrillar RdECM 0.5% and RdECM 1.0% hydrogels showed less contraction compared to the softer dECM hydrogels, suggesting a stiffer cellular environment (FIGURE 3A and B). This variation highlights the influence of microenvironment stiffness on cellular contraction dynamics, with softer matrices facilitating greater contraction due to less mechanical resistance.

[00106] The metabolic activity of HLFs embedded within the composite nanofibrillar hydrogels was quantitatively assessed at Day 3 and Day 7 using a WST-1 assay, the results of which are depicted in FIGURE 3C. On Day 3, RdECM 0.5% hydrogels demonstrated significantly higher metabolic activity compared to the control dECM, whereas RdECM 1.0% showed elevated but not statistically significant differences. By Day 7, the metabolic activities in both RdECM 0.5% and RdECM 1.0% reduced substantially, aligning closely with that of the dECM hydrogels. Notably, while the metabolic activity within the dECM hydrogels remained stable from Day 3 to Day 7, a significant reduction was observed in both RdECM formulations. The initial increase in metabolic activity observed in RdECM 0.5% on Day 3 could be attributed to the enhanced mechanical properties provided by the added aCNF. This observation is in line with findings from the previously published studies, where cells displayed increased metabolic activity in stiffer environments [58], [59], In addition, aSMA protein expression was significantly increased for HLFs cultured within RdECM 0.5 % and RdECM 1.0% hydrogels compared to soft dECM hydrogels (FIGURE 3D). The elevation of aSMA protein indicates fibroblast-to-myofibroblast transition by increasing the stiffness of hydrogels. The reinforced composite nanofibrillar dECM hydrogels (RdECM 0.5% and RdECM 1.0%) exhibited similar aSMA protein expression.

[00107] Activation of HLF Cultured on the Surface of Composite Nanofibrillar dECM Hydrogels: Next, HLFs were cultured on the surface of the composite nanofibrillar dECM hydrogels to analyze their morphology and response to increased stiffness (FIGURE 4). Since both RdECM 0.5% and RdECM 1.0% hydrogels demonstrated the capability to activate fibroblasts, this portion of the study was focused exclusively on the stiffest variant, RdECM 1.0%. Additionally, cells used for culture on the surface of the hydrogels were initially grown and expanded on soft substrates (E ~ 1.5 kPa[60]) for at least two passages to reset their mechanical memory [61] (see Methods). To initiate the analysis of cell morphology, cells were stained with phalloidin for highlighting F-actin fibers, enabling the creation of cell masks as shown in FIGURE 4A. Cells seeded on the softer dECM hydrogel exhibited greater spreading and elongation, with a larger projected area, compared to those on the stiffer RdECM 1.0% hydrogels (FIGURE 4A). This observation was quantitatively confirmed, showing a significant reduction in the area of cells seeded on stiff RdECM 1.0% hydrogels compared to those cultured on soft dECM hydrogels (FIGURE 4B). Furthermore, cells appeared rounder and less elongated, or exhibited a reduced aspect ratio (AR), when seeded on stiff RdECM 1.0% hydrogels, in contrast to those on soft dECM hydrogels (FIGURE 4C and D). Collectively, these findings show a notable alteration in HLF morphology when grown on stiffer substrates, a condition that mimics the fibrous structure characteristic of their native microenvironment.

[00108] It is well-documented that cells cultured on stiff substrates exhibit an elongated morphology with increased projected cell areas compared to those cultured on soft substrates [62], [63], However, when cultured within 3D environments, these cells often display different morphological behaviors. A notable reversal in cellular morphology has been observed when transitioning from 2D to 3D cultures: cells encapsulated in stiff environments became rounder and underwent significant reductions in volume compared to those in softer environments [64], [65], This phenomenon underscores the influence of mechanical environments on cellular behavior. Moreover, even when fibrous and non-fibrous hydrogels shared similar chemical compositions, cells exhibited distinct morphological and functional behaviors [23], [66], In particular, cells cultured on stiff, fibrous hydrogels — mimicking the structure of native tissues or collagen — tended to be smaller and rounder compared to their counterparts in soft, fibrous hydrogels [23], [67], The findings with composite nanofibrillar dECM-based hydrogels of the present disclosure corroborated these observations. Additionally, consistent morphologies were observed in cells cultured in both 2D and 3D, suggesting that the presence of fibrous structures in the composite nanofibrillar dECM-based hydrogels of the present disclosure provide a more tissue-like 3D environment even in 2D cultures. [00109] Next, the activation of fibroblasts into myofibroblasts was explored on these hydrogels. Similar to the findings from the 3D cell culture of encapsulated HLFs, aS MA protein expression of HLFs was upregulated in response to the stiffer ECM, as illustrated in FIGURE 5 A and B. Quantification of aSMA protein expression revealed an approximate 1.7-fold increase upon seeding cells on the stiffer ECM (FIGURE 5C). Additionally, the metabolic activity of these cells was examined as seen in FIGURE 5D. Here, no significant difference in metabolic activity was observed between the two hydrogel groups.

[00110] Nuclear Localization of YAP in HLF Cultured on Reinforced dECM hydrogels: YAP is a transcription co-activator and a central player in the Hippo signaling pathway, a pathway that regulates organ size, cell proliferation, apoptosis, and stem cell self-renewal[12], [13], The Hippo pathway is sensitive to mechanical cues from the environment, such as stiffness and cell density. When the Hippo pathway signaling is downregulated or disrupted, YAP and TAZ are not phosphorylated by LATS1/2 kinases, which allows them to translocate into the nucleus. Inside the nucleus, YAP and TAZ interact with TEAD transcription factors and other regulatory proteins to modulate the expression of genes that promote cell proliferation and inhibit apoptosis. [12], [68], [69], The localization and activity of YAP are significantly influenced by mechanical signals. For instance, in a stiff microenvironment, YAP localizes in the nucleus, promoting the expression of genes that lead to cell proliferation and survival[70],

[00111] In this study, YAP was utilized as an additional biomarker to verily the success of the reinforcement strategy of the present disclosure in increasing the stiffness of dECM-based hydrogels and effectively transmitting mechanical cues to cells. For this purpose, cells were initially grown and expanded on soft substrates for two passages prior to seeding them on hydrogels as described above. A low cell density of 10 x io⁴ cells/cm² was used to minimize cell-to-cell interactions, as such interactions have been reported to potentially influence YAP activity[71], [72], An increase in YAP nuclear localization was observed in cells seeded on the stiffer RdECM 1.0% hydrogels, compared to those on the softer dECM hydrogels (FIGURE 6).

[00112] In conclusion, the present study demonstrates the development and application of tunable nanofibrillar hydrogels, integrating dECM with aCNFs to closely mimic the mechanical and structural characteristics of native lung tissue. The hydrogels' adjustable mechanical properties, achieved through the modulation of nanomaterial concentration, provide a versatile platform for modeling various stages of healthy and diseased tissues. This work demonstrates enhanced mechanical properties of the composite nanofibrillar hydrogels, such as increased storage, loss, and compression moduli with the incorporation of aCNFs. These improvements will aid in simulating the stiffened tissue environment characteristic of fibrotic diseases like pulmonary fibrosis. The study also highlights the hydrogels' ability to support cell viability and proliferation, as demonstrated by the high metabolic rates of HLFs cultured within and on these materials. Furthermore, the differential expression aSMA and the localization of YAP in response to the hydrogels' varying stiffness underscore utility for studying myofibroblast activation and mechanotransduction, key processes in fibrosis. The observed changes in cell morphology, such as reduced size and altered shape, provide clear evidence that the composite nanofibrillar dECM hydrogels with tunable mechanical properties effectively influence cellular responses. This research paves the way for more accurate and physiologically relevant models of pulmonary fibrosis, offering significant potential for the fields of disease modeling, drug screening, and ultimately, regenerative medicine.

Example 2

Tissue Decellularization for Isolation of ECM

[00113] Lung tissue decellularization process involves several steps, as reported in previous study[39] and shown in Figure 7A. Fresh pig lung (pig lungs were generously donated by Jamie Waldron Butchers, Hamilton, ON, Canada) tissues were chopped and immediately frozen in liquid nitrogen, then stored at -80°C before decellularization. The decellularization began by soaking the tissues in 0.1% Triton X-100 (Sigma Aldrich, Canada, Catalog number: T9284-500ml) in a 4-liter beaker with continuous stirring for 30 minutes. The tissues were then filtered through a mesh strainer to remove the liquid, followed by a 30-minute wash in phosphate-buffered saline (PBS) to remove any remaining Triton X-100. The tissues were subsequently treated with 2% sodium deoxycholate (SDC, Sigma Aldrich, Canada, Catalog number: D6750-100G) and left overnight at 4°C. The next day, the SDC was removed, and the tissues were washed for 24 hours in PBS with an antibiotic-antimycotic (ThermoFisher, Canada, Catalog number: 15240062) solution, including at least one change of PBS. After the antibiotic- antimycotic wash, the tissues were agitated in sterilized deionized (DI) water for two hours, then washed for 30 minutes in IM sodium chloride. Following this, tissues were rinsed in DI water and treated with 1% Triton X-100. A final wash in PBS with antibiotic- antimycotic solution lasted 3 days. After these washes, the tissues were agitated in sterilized DI water for 30 minutes, strained, and stored at -80°C for lyophilization using a Benchtop Freeze Dryer (FreeZone 2.5 Liter -84C, Labconco). Post-freeze-drying, the tissues were ground into fine dECM powders in the presence of liquid nitrogen. These powders were then stored at -20°C for future experimental purposes.

Surface Modification of CNFs with Aldehyde Groups

[00114] In a round-bottom flask, a dispersion of CNFs (2 g of CNFs; Cellulose Lab, New Brunswick, Canada, Product Number: CNF-Sluny-SMC, Width: width 30-80 nm; Length: up to several hundred micron, surface group: hydroxyl) was prepared in DI water forming a 1.0 wt % CNF suspension containing 0.2 mM TEMPO (Sigma Aldrich, Canada, Catalog number: 214000-5G) and 2 mM sodium bromide (Sigma Aldrich, Canada, Catalog number: 310506-100G). To initiate the reaction, 6 mL sodium hypochlorite (containing 10-15 % chlorine; product # 425044, Sigma Aldrich) dissolved in 50 mL of DI water and then was added to the flask. The resulting suspension was stirred at room temperature at a constant rpm for 24 hours, while maintaining the pH at ~10 using 1 M NaOH. The reaction was quenched by introducing 1 mL of ethylene glycol (Sigma Aldrich, Canada, Catalog number: 324558-100ML). The obtained product underwent dialysis through 14 water changes against DI water using a cellulose membrane (12 - 14 kDa cutoff; Sigma Aldrich, Canada, Catalog number: D9652- 100FT). To further modify the reactants, a surface oxidation process was conducted using sodium periodate (NaIO4; Sigma Aldrich, Canada, Catalog number: 311448-100G) to yield surface-modified CNFs with aldehyde functional groups. The NaIO4 and suspension were mixed at a mass ratio of 1 : 1. The flask was covered with aluminum foil to protect the NaIO4 from photodegradation. The resulting mixture was continuously stirred at room temperature for 24 hours. The oxidation reaction was then halted by the addition of ethylene glycol (1 mL). Subsequently, the aCNFs suspension underwent dialysis against DI water with 14 water changes. Finally, the suspension was concentrated using evaporation of excess water. dECM-Based Hydrogels reinforced with aCNFs

[00115] To digest and solubilize dECM powder, dECM powder with a concentration of 35 mg/mL was added to an acidic solution containing pepsin (Sigma Aldrich, Canada, Catalog number: P6887-5G) with a concentration of 2.45 mg/mL (dECM to pepsin ratio was 10:0.75) as depicted in Figure 7A. As pepsin enzymatic activity is optimum at pH of 2, the pH of the solution was adjusted and monitored, ensuring it remained within the range of 2 to 3. If the pH exceeded 3, 1 N HC1 was added to lower it accordingly. Stirring was maintained at room temperature (RT) for a duration of three days. After 24 hours, the pH was measured and adjusted if necessary. Once the digestion process was complete, the dissolved dECM underwent centrifugation at approximately 4000 g for 5 minutes. The resulting supernatant, containing the dissolved dECM, was passed through a series of cell strainers with mesh sizes of 200 pm, 100 pm, 60 pm, and 30 pm and transferred to conical tubes, while the non-dissolved materials were discarded. To cool down, the tubes containing the dissolved dECM were placed on ice alongside an equal volume of lOx PBS that was also kept on ice. To reach a final concentration of 30 mg/mL of digested dECM solution, lOx PBS was added to the dissolved dECM, representing 10% of its volume. The pH of the dissolved dECM was subsequently adjusted to approximately 7.2-7.4 using 10 N and 1 N NaOH. Throughout the process, all steps were performed on ice. For the preparation of dissolved dECM with various concentrations, ice-cold PBS at a pH of 7.2-7.4 was added to the stock of dissolved dECM (concentration = 30 mg/mL). The entire process is shown in Figure 7A.

[00116] To create reinforced dECM-based hydrogels containing aCNFs, aCNF suspensions were buffered with 10X Hank's Balanced Salt Solution (HBSS) and then integrated at concentrations of 0.3% and 0.6% with neutralized digested dECM precursors. These hydrogels, enhanced with aCNFs, were categorized as RdECM (Reinforced dECM) 0.3% and RdECM 0.6%, indicating the respective aCNF concentration within the hydrogels. Each hydrogel formulation consistently incorporated 15.0 mg/mL of digested dECM, while the aCNF content varied, being present at concentrations of 0%, 0.3%, or 0.6% by weight.

Hydrogel Rheology Measurements

[00117] To obtain discs with uniform geometry for mechanical testing, poly dimethylsiloxane (PDMS) molds were fabricated first. The PDMS pre-polymer and curing agent were mixed at a 10 : 1 (w/w) ratio, degassed in a desiccator for ~30 min, poured into 3D-printed molds, and cured overnight at 65 °C. After curing, the PDMS molds were rinsed with 70 % ethanol, dried in a biosafety cabinet, and autoclaved. Pregel dECM, RdECM 0.3 %, or RdECM 0.6 % solutions were dispensed into the sterile PDMS molds and allowed to gel overnight at 37 °C. All hydrogels were then immersed in complete culture medium and incubated for at least 24 h at 37 °C to ensure full swelling equilibrium before any rheological measurements were performed. Mechanical characterization was carried out on a DHR-HR20 controlled-stress rheometer (TA Instruments, New Castle, DE, USA) equipped with a 8 mm parallel-plate geometry maintained at 37 °C. Oscillatory frequency sweeps (0.1-100 rad s ') were performed at a constant shear strain of 0.6 % to determine the storage (G') and loss (G") moduli. Plateau values were obtained by averaging data between 1 and 10 rad s '. Unconfined compression tests were conducted by displacing the upper plate at 5 pm s ' until a deformation of 100 pm (~ 10 % strain) was reached; the compressive modulus (E) was calculated from the linear region of the resulting stress-strain curve. Stress-relaxation measurements: immediately after the oscillatory and compression tests, a single 10 % shear-strain step was applied within 1 s, and the decay in shear stress was recorded continuously for 180 s (3 min). The relaxation modulus at each time point was obtained by dividing the instantaneous shear stress by the applied strain and was normalized to the value at t = 1 s for direct comparison among formulations.

Hydrogel swelling and degradation assays

[00118] Hydrogels for swelling and degradation studies were prepared exactly as described for rheological testing. Briefly, sterile PDMS moulds (10 : 1 base : curingagent, cured overnight at 65 °C, ethanol-rinsed, autoclaved) were filled with pre-gel solutions of dECM, RdECM 0.3 %, or RdECM 0.6 % and incubated overnight at 37 °C to obtain disc-shaped gels of identical initial height (~4 mm) and radius (6mm). Immediately after gelation, each disc was placed in a 12-well plate containing 2 mL of complete culture medium pre-warmed to 37 °C, and photographed (top view) with a digital camera (time = 0 h). Plates were then incubated at 37 °C, 5 % CO2. After 48 h, gels were imaged again under identical conditions. Radii at 0 h (Ro) and 48 h (FUx) were measured in ImageJ and the swelling ratio (Q) was calculated as R48/R0. Three to four discs per formulation were analysed, and Q is reported as mean ± SD. Disc-shaped hydrogels (dECM, RdECM 0.3 %, RdECM 0.6 %; n = 3 per formulation) were released from the PDMS moulds and transferred to 12-well plates containing 2 mL sterile Hank’s Physiological Salt Solution (HPSS). After 1 h at 37 °C (to leach unbound solutes), 3 replicates from each condition were removed, frozen (-80 °C), lyophilised for 48 h, and weighed to ±0. 1 mg to obtain the baseline dry mass Mo. The remaining discs were placed in 2 mL HBSS and incubated at 37 °C, 5 % CO2 for 15 days; HBSS was replaced every 2 days. On day 15, gels were rinsed twice in HPSS to remove degraded proteins, frozen, lyophilised to constant weight, and weighed (Mis). The extent of degradation was expressed as percentage mass remaining: 100

[00119] Individual values from the four replicates per formulation were averaged and are reported as mean ± SD.

TRITC-dextran release assay

[00120] Macromolecular diffusion was quantified with 70 kDa TRITC-dextran (25 mg mL ¹ x 5 mL; Chondrex Inc., Redmond, WA, USA). Ice-cold pre-gel suspensions of dECM, RdECM 0.3 %, and RdECM 0.6 % were prepared at a constant dECM concentration of 15 mg mL ¹ (1.5 % w/v). The TRITC-dextran stock (25 mg mL ¹) was diluted directly into each pre-gel to give a final dextran concentration of 1 mg mL ¹. and the mixtures were gently vortexed (5 s, low speed) for homogeneity. Hydrogels (n = 4 discs per formulation) were cast in sterile PDMS molds and gelled overnight at 37 °C. To equalise dextran concentration in hydrogels, discs were transferred to 12-well plates containing 2 mL HPSS supplemented with 1 mg mL ¹ TRITC-dextran and incubated for 24 h at 37 °C, 5 % CO2. After equilibration, gels were moved to fresh wells with 3 mL dextran-free HPSS (t = 0 h). At 0, 1, 2,3, 4, 6, 8, 12, 24, and 30 h, 100 pL of the release medium was removed and replaced with pre-warmed HPSS to maintain sink conditions. Fluorescence was read in a black 96-well plate (Z exc = 555 nm, Z_em = 580 nm) on a microplate reader. Concentrations were obtained from a TRITC-dextran standard curve (0-100 pg mL ¹ HPSS). The cumulative mass released at each time point was expressed as a percentage of the total dextran content per disc . Results are presented as mean ± SD (n = 4). Cell Culture

[00121] Human lung fibroblasts (HLFs) derived from lung tissue samples donated for medical research (Hamilton Integrated Research Ethics Board - HiREB - 5305-T) were initially isolated and cultured in DMEM (ThermoFisher, Canada, Catalog number: 11965118) supplemented with 10% fetal bovine serum (FBS, Wisent Inc., Saint-Jean- Baptiste, Canada, product number: 080-450) and 1% penicillin-streptomycin (Gibco, United States, Catalog number: 15140122). Custom PDMS molds (12 mm in diameter with the height of 4 mm) were used for shaping dECM hydrogel pucks. Concentrated HLFs were suspended in precursor dECM hydrogel solutions at varying concentrations of aCNFs to achieve a final cell density of 1 * 10⁶ cells/mL when encapsulating cells within hydrogels. The cells were gently and thoroughly mixed with the precursor dECM hydrogel solutions and poured into the PDMS disks. After incubating at 37 °C for two hours to allow the formation of dECM hydrogels, DMEM media was added to each disk containing the hydrogels, and the samples were incubated overnight. The following day, the dECM hydrogels containing HLFs were detached from the PDMS disks and transferred to well plates, where they were submerged in DMEM. For culturing cells on hydrogels, the final cell density was 5 * 10⁴ cells/cm² Cells were regularly tested with My coStrip™ - Mycoplasma Detection Kit (Invivogen, CA, USA, Product Number: REP- MYS-20).

Immunoblots of HLFs Protein Extracts from Hydrogels

[00122] DMEM was gently aspirated from each well using a pipette, to ensure the hydrogels stayed intact and undamaged. Subsequently, the samples underwent three washes with PBS, each lasting 5 minutes. For the digestion of hydrogels and extraction of proteins, 0.5 mL of 1 mg/mL collagenase (Cedarlane, Canada, Product Number: 7415) was added to the dECM hydrogels, and 1.5 mg/mL to the RdECM hydrogels. The dECM hydrogels were incubated for 1 hour, and the RdECM hydrogels for 1.5 hours, with all incubations conducted at 37 °C. After incubation, an additional 0.5 mL of FBS-enriched DMEM was introduced to each well to deactivate the collagenase. The hydrogels were then manually disrupted by pipetting them up and down, and the contents transferred to 2 mL Eppendorf tubes. These were centrifuged at 4 °C and 4000 rpm for 5 minutes. The supernatant was then discarded, and the samples were placed on ice for 20 minutes, followed by a wash with cold IX PBS. To prepare the lysis buffer solution, 800 pL of RIPA lysis buffer (ThermoFisher, Canada, Catalog number: 89900) was mixed with 100 pL of Protease Inhibitor Cocktail (Sigma Aldrich, Canada, Catalog number: P2714) and 100 pL of PhosSTOP (Sigma Aldrich, Canada, Catalog number: PHOSS-RO). Each sample’s pellet received 100 pL of this lysis buffer, then underwent brief ice-sonication using an ultrasonic homogenizer in short bursts. The samples were left to incubate on ice for another 20 minutes. For cell lysate collection, the solution was centrifuged in a microcentrifuge at 4 °C and 16,000 g for 20 minutes. The supernatant was then transferred to a new tube and stored at -80°C for downstream immunoblots.

[00123] The total protein concentration of the samples was measured using the Pierce™ Bicinchoninic Acid (BCA, ThermoFisher, Canada, Product Number: 23240), following the guidelines provided by the manufacturer. To achieve normalization of these protein concentrations, suitable amounts of PBS were added to each sample. The total cell lysate protein was combined with IX Laemmli Sample Buffer (Bio-Rad Laboratories, product number 1610747). This mixture was then electrophoresed on 4- 20% Mini-PROTEAN TGX stain-free precast gels (Bio-Rad Laboratories, product number 4568093) and the proteins were transferred onto a PVDF membrane using the reagents from the Transfer-Blot Turbo RTA Transfer Kit (Bio-Rad Laboratories, product number 1704272). The membranes were subjected to blocking at room temperature for one hour using a 5% Casein solution (Bio-Rad Laboratories, product number 1706404) in IX Tris Buffered Saline with TWEEN® 20. For protein detection, Clarity Western ECL Substrate (Bio-Rad Laboratories, product number 1705061) was used, and the results were captured using a ChemiDoc MP Imaging System. Image acquisition was set to auto-exposure.

Hydrogel Staining and Microscopy of HLF

[00124] To assess the viability of hydrogels formed under different conditions, the LIVE/DEAD™ Cell Imaging Kit (ThermoFisher, Mississauga, Ontario, with catalog number R37601) was employed. For nuclei staining, the NucBlue™ Live ReadyProbes™ Reagent (Hoechst 33342) from Invitrogen™ was used following the manufacturer's instructions. During the fixation process, three washes with PBS were performed between each staining step and then the samples were incubated with a 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA, with catalog number 15712) for 15 minutes for cells seeded on hydrogels and 30 minutes for cells encapsulated inside hydrogels. After fixation, samples were washed with PBS for at least one time. To ensure proper permeabilization, the fixed samples were treated with 0.2% Triton X-100 for 10 minutes for cells seeded on hydrogels and 20 minutes for cells encapsulated inside hydrogels. To minimize non-specific staining, a 3% bovine serum albumin (BSA, Wisent Inc., Saint-Jean-Baptiste, Canada, product number: 800-095-EG) solution was used and the samples were incubated for 24 hours. For antibody staining, primary antibodies were incubated with the samples in 3% BSA supplemented with 0.05 % Triton X-100 at 4 °C for 24 hours, followed by 2-hour incubation at RT with secondary antibodies, Hoechst, and phalloidin (Texas Red™-X Phalloidin, ThermoFisher, catalog number T7471) in 3% BSA supplemented with 0.05 % Triton X-100. The fixed cells were stained using a monoclonal anti-a-smooth muscle actin antibody (aSMA, Abeam, Canda, Product Number: ab7817; Clone: 1A4 or Sigma Aldrich, catalog number A2547; Clone: 1 A4). Widefield imaging of the cells was conducted using a ThermoFisher EVOS M7000, ZEISS Celldiscoverer 7, or Nikon AIR Inverted Confocal microscopes equipped. For a-SMA expression analysis, the mean gray value within the regions of interest was calculated using ImageJ[40] or Fiji [41] software.

Results and Discussion

Mechanical Properties of Composite Nanofibrillar dECM-based Hydrogels

[00125] In this work, CNFs functionalized with aldehyde groups, named as aCNFs, were mixed into a pregel dECM solution at two concentrations and allowed to gel overnight in an incubator at 37 °C. Pristine CNFs are prone to aggregation in an aqueous solution, which limits their ability to disperse uniformly and interact with the components of the dECM hydrogel solution. To address this challenge, the primary hydroxyl groups of CNFs were oxidized to carboxyl groups via TEMPO oxidation, followed by sodium periodate oxidation to introduce aldehyde groups onto the surface of CNFs [42,43], Covalently crosslinked dECM-based hydrogels were successfully formed through the Schiff-base reaction (Figure 7B and C) [44], The improvement in the mechanical properties of hydrogels was evidenced by the increased storage modulus, loss modulus, and compression modulus, all correlating with the concentration of aCNFs (Figure 8).

[00126] As the concentration of aCNFs increases, there is a notable increase in G, with RdECM 0.6 % showing the highest values (Figure 8A-C). While not wishing to be bound by theory, the increase in the storage modulus is likely due to the high aspect ratio and substantial surface area of aCNFs, which provide more points of interaction and crosslinking within the hydrogel network. Similar to the storage modulus, the loss modulus (G”) was also increased by increasing the concentration of aCNFs (Figure 8B). The observed increases in G” at higher concentrations of aCNFs could be attributed to the denser packing and entanglement of nanofibers, potentially resulting in greater Schiff- base cross-linking[45], The compression modulus follows a similar trend to G and G”, with RdECM 0.6 % showing the highest values (Figure 8C).

[00127] Stress-relaxation experiments confirmed that covalent stiffening with aCNF preserved the intrinsic viscoelastic balance of the hydrogels. When a 10 % shear step was applied, all formulations displayed a rapid decay in shear stress, relaxing ~60 % of the initial value within the first 30 s and approaching a common plateau after 180 s (Figure 8D). The characteristic half-relaxation time iA) — the point at which the stress had fallen to 50 % of its peak — remained statistically unchanged across native dECM, RdECM 0.3 %, and RdECM 0.6 % (Figure 8E). Thus, while aCNF increased the absolute magnitudes of both storage and loss moduli, the time-dependent redistribution of internal stress was unaffected, indicating that reinforcement enhances stiffness without compromising the gels’ ability to dissipate mechanical energy on physiologically relevant timescales.

[00128] While not wishing to be bound by theory, the observed enhancement in mechanical properties is most plausibly attributed to the aldehyde functional groups introduced on the CNF surfaces, which form covalent Schiff-base linkages with primary amine groups in the ECM proteins [46], There is also a possibility that the mechanical properties of these hydrogels was further enhanced by multiple interactions. For instance, the aCNFs became entwined, adding to the gel's stability. Additionally, the presence of hydrogen bonding interactions between the hydroxyl groups on the aCNFs with the amine groups of ECM proteins is likely to further enhance the hydrogel mechanical properties. Lastly, electrostatic interactions wherein the negatively charged carboxyl groups on aCNFs attract to the positively charged amine groups found in ECM proteins may improve mechanical stability.

[00129] The stiffness (Y oung’s modulus) of healthy lung tissue is typically around 3.7 ± 1.3 kPa[47], although this value can vary significantly, with a range from 0.5 to 9 kPa according to different reports [48], In contrast, fibrotic lung tissue displays a significantly higher stiffness before decellularization, averaging at 18.9 ± 11.1 kPa[47], The range of values for fibrotic tissue is notably broader, extending to approximately 100 kPa[48], Using these studies, the stiffness of lung tissues fall within the range of 1-5 kPa for healthy tissue and typically exceed 10 kPa for fibrotic tissues[47-50]. After decellularization, dECM hydrogels derived from non-IPF and IPF lungs were reported to have stiffness values of 1.1 ± 0.2 kPa and 6.8 ± 2.8 kPa, respectively, which indicates a significant reduction in stiffness compared to their intact conditions [47], Using these values, dECM hydrogels with a compression modulus of 1.56 ± 0.47 kPa are considered as a normal or soft environment for HLFs. Meanwhile, both RdECM 0.3% and RdECM 0.6% hydrogels, with compression moduli of 8.60 ± 1.34 kPa and 16.61 ± 2.42 kPa respectively, are representative of a stiff or fibrotic environment for HLFs.

Physical Characterization of Composite dECM Hydrogels

[00130] dECM hydrogels are rich in collagen, a key element in the formation of fibrous structure of these hydrogels[51]. Collagen, the most abundant protein in the ECM, has a unique ability to form fibrillar networks due to its distinct triple-helix structure, enabling the self-assembly of long, thin fibers integral to the fibrous scaffold of the hydrogel [51,52], Fibrous hydrogels more closely mimic the natural arrangement and physical interactions found in tissues, which can be important for certain types of cellular responses and for the study of specific mechanotransduction pathways. In this work, composite nanofibrillar lung dECM hydrogels featuring tunable mechanical properties were developed, representing both healthy and fibrotic lungs, while preserving fibrous structures that resemble native lung tissue.

[00131] SEM images confirmed that all three formulations — native dECM, RdECM 0.3 %, and RdECM 0.6 % — retained a nanofibrillar architecture that is characteristic of collagen-rich lung ECM (Figure 8I-K). Qualitatively, fibers were continuous and interwoven in every sample, and no obvious disruption or densification was observed upon aCNF incorporation (Figure 8 I-K). Thus, covalent reinforcement preserved the morphological cues that cells encounter in the native matrix. Hydrogel hydration behavior changed markedly, however, once aCNFs were introduced. After 48 h in complete culture medium, native dECM discs swelled roughly 6.5 ± 0.6 % relative to their initial sizes, whereas RdECM 0.3 % and RdECM 0.6 % swelled only 1.07 ± 0.44 % and 1.80 ± 0.5 %, respectively (Figure 8F). Long-term stability mirrored the swelling trend. After 15 days in serum-containing medium (with complete medium exchange every 48 h), native dECM lost ~ 11% of its dry mass. In contrast, the dry mass of RdECM 0.3 % and RdECM 0.6 % did not significantly change after 15 days, demonstrating that even modest aCNF contents markedly slow hydrolytic fragmentation of the protein network, further confirming the formation of crosslinks in collagen networks. Macromolecular transport, assessed with 70 kDa TRITC-dextran, was likewise slightly impacted by reinforcement (Figure 8H). Release of the 70 kDa TRITC-dextran probe (hydrodynamic diameter ~ 10 nm) followed the same biphasic profile in all three hydrogel formulations: a short burst phase during the first few hours, followed by a gradual approach to a common plateau reached at roughly 12 h. The presence of aCNF cross-links did not appreciably alter either the shape of the release curve or the time required to reach equilibrium, indicating that bulk incorporation of aCNF preserved overall macromolecule permeability of the dECM network while providing the desired mechanical reinforcement.

Stiffness-driven Pro-fibrotic Activation of HLF Seeded on the Composite dECM Hydrogels

[00132] Myofibroblasts are cells that play a crucial role in wound healing and tissue repair by producing and remodeling the ECM[53-55], However, their persistent activation is a hallmark of pathological fibrosis. In fibrotic diseases, fibroblasts transform into myofibroblasts in response to various stimuli, including mechanical stress, cytokines, and growth factors. This transition is characterized by the de novo expression of a-SMA, which is incorporated into intracellular stress fibers. These stress fibers are contractile structures that increase the cell's ability to contract and exert force on the surrounding ECM[56,57], The result is a stiffening of the tissue and excessive deposition of ECM components, which is detrimental in chronic conditions and leads to the disruption of tissue architecture and function[58]. Primary human lung fibroblasts (HLF) from 3 to 4 IPF-free donors were plated at 5 x 10³ cells cm ² on either the compliant dECM hydrogel (~ 1.5 kPa) or the stiff aCNF -reinforced RdECM 0.6 % (~ 16 kPa) and analysed after 72 h (Figure 9 and Figure 10). The low cell seeding density minimised cell-to-cell contacts, isolating the effect of substrate mechanics. Confocal images revealed an increase in a- SMA expression for cells on the stiff matrix compared to cells seeded on soft native dECM hydrogels (Figure 9A). Quantification showed that every donor exhibited a stiffness-dependent increase in normalized integrated a-SMA intensity, though the foldchange ranged from ~1.4 to -3.9, underscoring inter-individual heterogeneity (Figure 9B). Similar to the findings from the use of classic polyacrylamide gels, the results of this study demonstrated that matrix rigidities above -10 kPa are sufficient to trigger a-SMA expression and myofibroblast differentiation in lung fibroblasts[59], Under identical culture conditions, YAP fluorescence was predominantly cytoplasmic on the soft native dECM substrate but shifted to the nucleus on the stiff RdECM 0.6 % (Figure 10A). Nuclear-to-cytoplasmic (N/C) YAP ratios averaged per donor were significantly higher on the stiff gels (Figure 10B), consistent with stiffness-mediated inhibition of Hippo signalling and activation of mechanotransducive gene programs[60]. Together with the a-SMA data, this confirms that the covalently reinforced hydrogel of the present disclosure transmits a pro-fibrotic mechanical signal that is sensed at both the cytoskeletal and transcriptional levels.

[00133] Fibroblast morphology is classically expected to scale with substrate rigidity in 2D culture — cells spread and elongate as elastic modulus rises into the tens of kilopascals because increased integrin tension promotes focal-adhesion growth and actin-myosin contractility [61,62], In the fibre-containing dECM system of the present disclosure this canonical trend behaved slightly different and inconsistent in some cases: cell and nuclear areas changed by only -10-15 % and the direction of change varied among the four donors, while eccentricity shifts were similarly small and donor-specific (Figure 9C-F). Such behaviour is not anomalous; several studies have shown that when rigidity cues are presented within, or atop, a collagenous fibre network the physical guidance and local compliance of the fibres often dominate over bulk modulus, leading to smaller, rounder, or morphology-invariant fibroblasts even on nominally stiff matrices [23,63-67], Optical -tweezer and micro rheology work further demonstrates that fibroblasts actively remodel peri-cellular collagen, creating micro-domains whose effective stiffness can diverge from the macroscopic value and thereby blur populationscale correlations between bulk modulus and cell shape[66]. Donor heterogeneity adds a second layer of complexity. Variations in integrin repertoire, cytoskeletal tension setpoints, and collagen-binding proteins have all been linked to divergent spreading responses on identical substrates[68,69]. For cell spread area, donor 1 increased its mean footprint on the stiff RdECM 0.6 %, whereas donor 2 showed a slight reduction (~ 7 %), donor 3 a pronounced reduction (~ 18 %), and donor 4 exhibited no detectable change. Cell eccentricity shifted only modestly: it decreased for donors 1, 3, and 4 but increased for donor 2, confirming that bulk stiffening did not drive a uniform gain in elongation. A similar pattern emerged for the nucleus. Nuclear area fell in donors 1-3 (by 6-12 %) yet rose in donor 4, while nuclear eccentricity decreased in donors 1 and 3 but increased in donors 2 and 4. Taken together, these data demonstrate that a jump from - 1.5 kPa to - 16 kPa can elicit diametrically opposite geometric adaptations in different donors, even though all donors converge on the same biochemical read-outs (a-SMA up-regulation and YAP nuclear localisation). Such morphological “decoupling” from stiffness is increasingly recognised in fibrous micro-environments. Baker et al. showed in a hyaluronic-acid fibre network that fibroblasts spread less on the stiffest formulation, attributing the effect to local fibre guidance and non-linear strain-stiffening that concentrate forces in a limited pericellular zone[23]. Wisdom et al. demonstrated that fibroblasts exploit fibre buckling to remodel their immediate surroundings, generating “mechanically insulated” niches whose apparent stiffness can be several-fold lower than the macroscopic value[70]. Collectively, these studies — and the present data — support the notion that in a 3-D-like fibrous setting, cells sense and manipulate the lattice at the micron scale; the emergent morphology therefore reflects a balance between intrinsic contractility, donor-specific integrin repertoire[68], and the local topology of collagen- aCNF filaments rather than the bulk modulus alone.

[00134] From a modelling standpoint this heterogeneity is informative rather than problematic. Primary fibroblasts isolated from different donors, including those with IPF, exhibit considerable variability in traction force generation and spreading behavior, even when cultured on identical rigid substrates — a reflection of donor-specific differences in mechanosensitivity and cytoskeletal regulation[71,72]; the dECM platform of the present disclosure preserves that individuality while still driving a uniform pro-fibrotic behavior in cells. The ability of a single, pathologically relevant stiffness increase (~ 1.5 kPa — > 16 kPa) to elicit a consistent pro-fibrotic phenotype, despite inter-donor variation in morphology, highlights the utility of the aCNF -reinforced dECM platform for studying early mechanobiological events in lung fibrosis. Because the stiff matrix preserves native fibrillar architecture and viscoelastic relaxation, it offers a physiologically grounded alternative to purely synthetic substrates while delivering the mechanical cues necessary to activate key fibrogenic pathways. Fibrous lung-dECM hydrogels as a 3-D contraction assay and drug-screening platform

[00135] The interstitial space of the lung is a tension-bearing fibrous meshwork in which collagen I/III, elastin and matri cellular proteins are organised into sub-micrometre bundles. Mimicking this milieu in vitro is important because fibroblasts sense rigidity not just through bulk elastic modulus but through the ability of individual fibres to align, buckle and transmit force. Flat polyacrylamide or PEG gels — although tunable in stiffness — lack this hierarchical architecture and can mis-represent mechanobiology. Decellularized extracellular matrix (dECM) from porcine lung offers a pragmatic compromise between fidelity and availability: its collagen-to-elastin ratio, glycosaminoglycan content and nonlinear stress-strain response closely mirror those of human lung ECM, while yielding reproducible scaffolds at organ scale[73-75]. By covalently reinforcing porcine dECM with trace aldehyde-functionalized cellulose nanofibres (aCNFs) a family of hydrogels was obtained whose stiffness spans the healthy -to-fibrotic range (~1.5 to 16 kPa) without compromising the fibrillar topology, thereby creating a realistic 3D environment for cells.

[00136] A central profibrotic cue in this environment is transforming growth factor- (TGF-0). TGF-01 is secreted as an inactive complex composed of the cytokine, a latency-associated peptide (LAP) containing an RGD motif, and a latent-TGF-0 binding protein (LTBP) that tethers the complex to the ECM. When fibroblasts contract against the matrix, tension transmitted through av-containing integrins (avf> I . avf>3. avf>5. avf>6. av08) unfolds LAP, liberating active TGF-0, which then binds the type II receptor, recruits ALK5 (type I), and triggers SMAD2/3 phosphorylation and a myofibroblast gene program[76-79]. Because latent-TGF-0 activation is rate-limiting for fibrosis progression, blocking av integrins has emerged as a promising anti-fibrotic strategy. CWHM-12 is a high-affinity, broad-spectrum av-integrin antagonist that sterically occludes the RGD-binding cleft, preventing mechanical activation of latent TGF-0. In mouse models of lung, liver, kidney and skeletal-muscle fibrosis, systemic or local delivery of CWHM-12 at sub-micromolar plasma levels attenuates collagen deposition, lowers a-SMA expression and improves tissue mechanics without overt toxicity[80,81]. Therefore 1 pM was selected — an order of magnitude above reported in-cell ICAo values — to ensure near-complete target occupancy during the 3-D culture period. [00137] Ice-cold pre-gel suspensions were prepared for two formulations: soft native dECM (1.5 % w/v) and stiffRdECM 0.6 % (dECM 1.5 % w/v + 0.6 % w/v aCNF). Primary human lung fibroblasts from three independent donors were harvested at 70 - 80 % confluence and resuspended at 1 x 10⁶ cells mL ¹ directly into each pre-gel on ice. Pregel solutions were dispensed into sterile PDMS molds and incubated overnight (37 °C, 5 % CO2) to allow complete thermo-driven self-assembly and aldehy de-amine crosslinking (Figure 11 A). The next morning (defined as day 1), gel discs were released from the molds and transferred to 24-well plates containing 1 mL fibroblast growth medium, which was exchanged daily. On day 4, half of the gels in each stiffness group received 1 pM CWHM-12 (0.1 % DMSO carrier); the remaining gels received an equivalent volume of DMSO vehicle. Cultures were maintained until day 7. Macroscopic contraction was quantified on days 1, 4, and 7 by photographing each disc top-down on a calibrated stage and measuring the radius (R) in ImageJ. The contraction index was expressed as R/Ro, where Ro is the initial radius measured immediately after mold removal (Figure 11B). Radii for each donor were plotted individually (Figure 11C). For a-SMA analysis, gels were fixed on day 7 and Z-stacks spanning at least 100 pm in depth were acquired on a confocal macroscope and projected images were created as shown in Figure 12A.

[00138] HLF encapsulated within the soft native-dECM matrix induced a marked reduction in hydrogel diameter, reaching an average R/Ro of - 0.95 for donors 2 and 3 and 0.91 for donor 1 within the first 24 h and converging near 0.80 by day 7, consistent with the ability of cellular traction forces to overcome the low cohesive modulus of the network. In contrast, hydrogels reinforced with 0.6 % aCNF resisted compaction: donor 1 stabilized almost immediately at - 0.90 R/Ro, donors 2 and 3 began near 0.95 R/Ro and levelled at 0.93, illustrating that a three-fold increase in storage modulus imposes a mechanical threshold that any donor would interact and respond differently. Administration of the broad-spectrum av-integrin antagonist CWHM-12 (1 pM, day 4) did not further influence contraction of stiff hydrogels and was equally ineffective in soft matrices from donors 2 and 3; however, in donor 1 the drug curtailed additional compaction after day 4, terminating at 0.85 R/Ro instead of 0.80, showing that interindividual variability in the tension required to activate latent TGF- via av-integrins. These observations align with classical collagen-gel studies showing that bulk contraction is permitted only when the fiber network is sufficiently compliant and continuously connected, and that additional cross-linking or fiber densification suppresses global shortening even though local fiber sliding may persist[64, 82-84], They also accord with more recent work in engineered fibrous hydrogels where local buckling and strain-stiffening create donor-dependent limits to macroscopic compaction despite large differences in nominal G'[64, 82-85],

[00139] Collectively, the data demonstrate that the aCNF -reinforced lung-dECM platform closely recapitulates the stiffness-sensitive contractile phenotype of fibroblasts while exposing clinically relevant heterogeneity in baseline traction forces and drug responsiveness that is likely to underlie variable patient outcomes in anti-fibrotic therapy.

[00140] Consistent with earlier observations, encapsulated fibroblasts also exhibited a clear rise in a-SMA when they were cultured inside the stiff RdECM 0.6 % matrix, yet the magnitude of that rise differed markedly among the three donors, ranging from a modest 1.6-fold to a robust 3.4-fold increase over the soft dECM baseline (Figure 12). This heterogeneity echoes previous single-cell RNA-seq analyses showing that primary lung fibroblasts retain donor-specific epigenetic programs that modulate actomyosin tone and myofibroblast susceptibility even after extensive in-vitro passaging, and it underscores the need to preserve patient-level variability when assessing anti- fibrotic candidates[86,87]. Where the stiff network promotes a-SMA assembly is readily explained by mechanical activation of latent TGF- complexes anchored to the dECM fibrils: tension transmitted through avf> integrins unfolds the latency-associated peptide and liberates active TGF- 1, which in turn engages ALK5 and drives SMAD2/3- dependent transcription of ACTA2. In the present system, CWHM-12 — a nanomolar antagonist of av-containing integrins — attenuated that mechanochemical loop in a donorspecific manner, mirroring the contraction data in Figure 12 and revealing tight coupling between tissue-scale force generation and intracellular myofibroblast markers. Donor 1 illustrated a high-tension phenotype: soft gels compacted aggressively, stiff gels compacted little, and CWHM-12 reduced bulk contraction as well as a-SMA, yet the drug could not return a-SMA to baseline, consistent with residual integrin-independent activation of TGF- or with autocrine feed-forward signaling once a critical a-SMA threshold is crossed. Donors 2 and 3, by contrast, adopted a lower-tension set-point; their soft gels reached the same end-radius as donor 1, but contraction plateaued at an earlier time and CWHM-12 entirely normalized a-SMA (donor 2) or brought it close to baseline (donor 3), implying that in these donors av-integrin engagement was the dominant driver of myofibroblast commitment (Figure 13A and B). Importantly, stiff gels in all donors were largely refractory to additional mechanical or pharmacologic modulation — a-SMA remained elevated and uncompacted discs preserved their original diameter — reinforcing the concept that once matrix modulus rises above the tens-of-kilopascal range mechanical deformations fall below the threshold required to propagate sufficient strain along individual fibers to release latent TGF- . The three-dimensional renderings bolster this mechanistic view: cells in soft gels appeared densely packed because collective traction brought them into closer proximity, whereas in stiff gels the inter-cellular spacing remained wide, implying that mechanical decoupling limited both compaction and cell- to-cell paracrine reinforcement (Figure 13C). Together, these findings align with seminal collagen-matrix studies by Hinz and Wipff, who demonstrated that pharmacologic inhibition of avP3/p5 abolishes traction-mediated TGF-P activation[76], and with more recent work in strain-stiffening hyaluronan fibers by Baker et al., who showed that local buckling and realignment of fibers are prerequisites for force-dependent cytokine release[23]. The data of this study extend those observations to a lung-derived dECM platform, confirming that modest aldehyde-mediated stiffening suffices to reach a mechanical percolation limit where most fibroblast populations can no longer generate enough strain to mobilize latent TGF-P, yet a subset of high-contractile donors still pushes the system into a pathologic state — a clinically relevant scenario given that IPF progression often correlates with “super-activator” fibroblast clones. Finally, the muted effect of CWHM-12 in soft gels from donors 2 and 3 underscores the caveat that anti- integrin therapies may exhibit variable efficacy across patients and highlights the utility of the current fibrous, lung-mimetic hydrogel for stratifying such responses prior to clinical translation.

[00141] Across the three donor-derived fibroblast populations encapsulated within soft and stiff dECM-based hydrogels, quantitative morphometric analysis revealed donor-specific yet interpretable trends in cell and nuclear shape (Figure 13). Projected cell area, a representative for cell spreading and matrix interaction, exhibited non-uniform responses to increased stiffness (Figure 13 A). In donor 1, a slight reduction in projected area was observed in stiff matrices compared to soft, indicating a subtle decrease in cellmatrix engagement. In contrast, donor 2 cells showed a measurable increase in area in stiff hydrogels, suggesting enhanced spreading or cytoskeletal tension, while donor 3 again followed a pattern of slight reduction, resembling donor 1. Interestingly, in stiff matrices treated with CWHM-12, cell area consistently trended toward the profile of cells in soft matrices, implying that inhibition of av integrin-mediated signalling partially reverted the stiffness-induced morphological shift. A similar pattern was observed for projected nuclear area, which often reflects nuclear deformability and cytoskeletal coupling (Figure 13B). In donor 1, the nucleus became slightly enlarged in stiff gels, while in donor 2 the increase was negligible. Donor 3 uniquely showed a decrease in nuclear area with increased stiffness; however, upon treatment with CWHM-f2, nuclear size increased toward soft-gel levels, mirroring the pattern observed for whole-cell area. This normalization in nuclear dimensions supports the hypothesis that integrin-mediated cytoskeletal forces modulate nuclear tension, which can be pharmacologically reversed even in a mechanically stiff context. Cell eccentricity, an indicator of elongation or anisotropy, was more variable across donors (Figure 13C). Donors 1 and 2 displayed a slight increase in eccentricity on stiff matrices compared to soft, potentially reflecting enhanced alignment or directional tension within a higher stiffness network. Donor 3 deviated from this pattern, with a small decrease in eccentricity in the stiff condition. These subtle changes suggest that not all fibroblasts respond to stiffness by adopting an elongated phenotype, particularly within a fibrous 3D context where fibre alignment and local matrix compliance also influence cell shape. For nuclear eccentricity, a more consistent pattern emerged (Figure 13D). In all donors, nuclear eccentricity decreased in stiff hydrogels relative to soft, reflecting a shift toward more rounded nuclear morphology. The observed reduction in nuclear eccentricity within stiff fibrous hydrogels is indicative of increased cytoskeletal tension, a relationship supported by prior studies. For instance, Keeling et al. demonstrated that actomyosin-generated cytoskeletal tension regulates nuclear shape and mechanics, with increased tension leading to more rounded nuclear morphologies in stiff environments[88]. Similar work highlighted that scaffold architecture and tensile deformation influence nuclear shape, mediated by the actin cytoskeleton. These findings confirm the role of cytoskeletal forces in modulating nuclear morphology in response to mechanical cues within fibrous matrices[89-91]. The fact that CWHM-12 treatment increased nuclear eccentricity in donor 3 again points to mechanical feedback mechanisms mediated through integrin-cytoskeleton-nucleus coupling that can be modulated pharmacologically. Taken together, these data show that donor identity strongly influences how fibroblasts adapt their morphology in response to mechanical stiffening, but integrin inhibition can partially reverse these adaptations. While classical models predict increased spreading and elongation with increasing stiffness, this was not universally observed here, likely due to the fibrous nature of the dECM and the presence of dynamic cell-matrix interactions that modulate force transmission in complex ways. Prior work has shown that matrix architecture and pericellular fibre mechanics often override bulk modulus in shaping cellular morphology [23, 70], The findings of the present study are consistent with this interpretation and reinforce the idea that patient-specific fibroblast responses to stiffness and to drug treatments cannot be reduced to a single morphological pattern, but rather must be analysed across multiple shape metrics and biological contexts. This platform provides the resolution needed to make such distinctions and could serve as a powerful tool for preclinical phenotyping of human fibroblast behavior in the context of fibrosis and its modulation.

Conclusion

[00142] In this study, a tunable fibrous hydrogel system was developed by reinforcing lung-derived decellularized extracellular matrix (dECM) with aldehyde- functionalized cellulose nanofibers (aCNFs), enabling precise modulation of matrix stiffness while preserving native ECM architecture. This mechanically adaptable platform successfully mimics the microenvironmental features of both healthy and fibrotic lung tissue, allowing for direct investigation of fibroblast behavior in physiologically relevant 2D and 3D contexts. The results demonstrate that increasing hydrogel stiffness promotes fibroblast activation, marked by enhanced a-smooth muscle actin (a-SMA) expression and nuclear localization of YAP, while altering cellular morphology in a donor-specific manner. It was shown that pharmacological inhibition of av-integrins using CWHM-12 elicits variable responses among donors, impacting both matrix contraction and fibrotic marker expression in a context-dependent fashion. These findings underscore the role of donor heterogeneity in shaping fibroblast mechanobiology and therapeutic responsiveness. By enabling quantification of cell morphology, contraction kinetics, and mechanosensitive signaling within a human lung-mimetic matrix, this platform offers a powerful tool for drug testing and fibrosis modeling. It supports not only mechanistic studies of fibroblast activation, but also the development of stratified screening approaches that account for patient variability — a step toward personalized anti-fibrotic therapies. Example 3

Surface Modification of CNCs with Aldehyde Groups

[00143] Aldehyde functional groups were introduced onto the surface of CNCs through oxidation using sodium periodate (NalC ) as an oxidising agent[42], [73], A 1.0 wt % CNC suspension was prepared, and NaIO4 was added at a weight ratio of 0.6: 1 (NaIO4: CNC). The properties of the CNC are as follows: width: 5-20nm; Length: 100- 250 nm, Surface Group: Hydroxyl, sulfonic group. The reaction mixture was stirred at room temperature under aluminum foil cover to protect the reaction from the light. After 2 hours of reaction, the reaction was quenched by adding 1 mL of ethylene glycol. The resulting suspension of aldehyde-modified CNCs was subjected to dialysis against deionized water (DI) with 14 water changes using a cellulose membrane (12 - 14 kDa cutoff). Subsequently, the a-CNC suspension was concentrated to the desired concentration exceeding 4.0 wt % through evaporation of excessive water.

[00144] CNF dispersion was prepared from CNFs; Cellulose Lab, New Brunswick, Canada, Product Number: CNF-Slurry-SMC, according to the method described in Example 1.

[00145] The incorporation of aldehyde-functionalized cellulose nanocrystals (aCNCs) at 0.5 % and 1.0 % w/v did not lead to a significant increase in the mechanical properties of the dECM-based hydrogels, as evidenced by the minimal changes observed in storage modulus (G'), loss modulus (G"), and compression modulus compared to native dECM. In contrast, equivalent concentrations of aldehyde- functionalized cellulose nanofibers (aCNFs) produced a marked, concentrationdependent enhancement in all measured parameters. This disparity reflects fundamental differences in morphology, aspect ratio, and network-forming potential between the two nanomaterials. Unlike CNFs, which are long, entangled fibrils with high aspect ratios capable of physically reinforcing the hydrogel matrix, CNCs are rigid, rod-like particles with short lengths that do not form percolating networks. As a result, CNCs are more readily dispersed and less likely to contribute to mechanical interlocking or network cohesion. Additionally, although both materials were functionalized with aldehyde groups to promote Schiff-base crosslinking with amine-rich dECM proteins, the route of functionalization may also play a role. CNFs in this study were sequentially treated — first carboxylated via TEMPO oxidation to increase surface reactivity and dispersion, and then oxidized with sodium periodate to introduce aldehyde groups. This dual-step modification may enhance crosslinking efficiency and retention within the ECM matrix. In contrast, CNCs were directly oxidized with sodium periodate, which may result in a lower effective density of aldehyde groups or a less optimal spatial configuration for crosslinking with the ECM. Together, these findings suggest that both the nanoscale morphology and the functionalization strategy influence the ability of cellulose derivatives to mechanically reinforce soft protein-based hydrogels. Contrary to CNCs, CNFs offer an effective platform for mechanical modulation in tissuemimetic systems requiring stable, tunable stiffness.

[00146] While the present disclosure has been described with reference to examples, it is to be understood that the scope of the claims should not be limited by the embodiments set forth in the examples, but should be given the broadest interpretation consistent with the description as a whole.

[00147] All publications, patents and patent applications are herein incorporated by reference in their entirety to the same extent as if each individual publication, patent or patent application was specifically and individually indicated to be incorporated by reference in its entirety. Where a term in the present disclosure is found to be defined differently in a document incorporated herein by reference, the definition provided herein is to serve as the definition for the term.

FULL CITATIONS FOR DOCUMENTS REFERRED TO IN THE

DISCLOSURE

[1] P. Jain, H. Kathuria, and N. Dubey, “Advances in 3D bioprinting of tissues/organs for regenerative medicine and in-vitro models,” Biomaterials, vol. 287, p. 121639, Aug. 2022, doi: 10.1016/j. biomaterials.2022.121639.

[2] S. Chaudhary and E. Chakraborty, “Hydrogel based tissue engineering and its future applications in personalized disease modeling and regenerative therapy,” Beni-Suef University Journal of Basic and Applied Sciences , vol. 11, no. 1, p. 3, Jan. 2022, doi: 10.1186/s43088-021-00172-1.

[3] M. Mirbagheri, V. Adibnia, B. R. Hughes, S. D. Waldman, X. Banquy, and D. Kun Hwang, “Advanced cell culture platforms: a growing quest for emulating natural tissues,” Materials Horizons, vol. 6, no. 1, pp. 45-71, 2019, doi: 10.1039/C8MH00803E.

[4] X. Wan, Z. Liu, and L. Li, “Manipulation of Stem Cells Fates: The Master and Multifaceted Roles of Biophysical Cues of Biomaterials,” Advanced Functional Materials, vol. 31, no. 23, p. 2010626, 2021, doi: 10.1002/adfm.202010626.

[5] W. Hu, J. Shi, W. Lv, X. Jia, and K. Ariga, “Regulation of stem cell fate and function by using bioactive materials with nanoarchitectonics for regenerative medicine,” Science and Technology of Advanced Materials, vol. 23, no. 1, pp. 393-412, Dec. 2022, doi: 10.1080/14686996.2022.2082260.

[6] Z. Liu, X. Wan, Z. L. Wang, and L. Li, “Electroactive Biomaterials and Systems for Cell Fate Determination and Tissue Regeneration: Design and Applications,” Advanced Materials, vol. 33, no. 32, p. 2007429, 2021, doi: 10.1002/adma.202007429.

[7] Y. Liu et al., “Stiffness-mediated mesenchymal stem cell fate decision in 3D- bioprinted hydrogels,” Burns & Trauma, vol. 8, p. tkaa029, Jan. 2020, doi: 10.1093/bumst/tkaa029.

[8] R.-Z. Tang and X.-Q. Liu, “Biophysical cues of in vitro biomaterials-based artificial extracellular matrix guide cancer cell plasticity,” Materials Today Bio, vol. 19, p. 100607, Apr. 2023, doi: 10.1016/j.mtbio.2023.100607.

[9] W. Xie et al., “Static and Dynamic: Evolving Biomaterial Mechanical Properties to Control Cellular Mechanotransduction,” Advanced Science, vol. 10, no. 9, p. 2204594, 2023, doi: 10.1002/advs.202204594.

[10] X. Cun and L. Hosta-Rigau, “Topography: A Biophysical Approach to Direct the Fate of Mesenchymal Stem Cells in Tissue Engineering Applications,” Nanomaterials, vol. 10, no. 10, Art. no. 10, Oct. 2020, doi: 10.3390/nanol0102070.

[11] S. W. Wong, S. Lenzini, R. Giovanni, K. Knowles, and J.-W. Shin, “Matrix biophysical cues direct mesenchymal stromal cell functions in immunity,” Acta Biomaterialia, vol. 133, pp. 126-138, Oct. 2021, doi: 10.1016/j. actbio.2021.07.075.

[12] X. Cai, K.-C. Wang, and Z. Meng, “Mechanoregulation of YAP and TAZ in Cellular Homeostasis and Disease Progression,” Frontiers in Cell and Developmental Biology, vol. 9, 2021, Accessed: Oct. 22, 2022. [Online]. Available: https://www.frontiersin.org/articles/10.3389/fcell.2021.673599

[13] F. Liu et al., “Mechanosignaling through YAP and TAZ drives fibroblast activation and fibrosis,” American Journal of Physiology-Lung Cellular and Molecular Physiology, vol. 308, no. 4, pp. L344-L357, Feb. 2015, doi: 10.1152/ajplung.00300.2014.

[14] B. N. Narasimhan, M. S. Horrocks, and J. Malmstrom, “Hydrogels with Tunable Physical Cues and Their Emerging Roles in Studies of Cellular Mechanotransduction,” Advanced NanoBiomed Research, vol. 1, no. 10, p. 2100059, 2021, doi: 10.1002/anbr.202100059.

[15] M. Guvendiren, M. Perepelyuk, R. G. Wells, and J. A. Burdick, “Hydrogels with differential and patterned mechanics to study stiffness -mediated myofibroblastic differentiation of hepatic stellate cells,” Journal of the Mechanical Behavior of Biomedical Materials, vol. 38, pp. 198-208, Oct. 2014, doi: 10.1016/j.jmbbm.2013.11.008.

[16] T. A. Ulrich, E. M. de Juan Pardo, and S. Kumar, “The Mechanical Rigidity of the Extracellular Matrix Regulates the Structure, Motility, and Proliferation of Glioma Cells,” Cancer Research, vol. 69, no. 10, pp. 4167-4174, May 2009, doi: 10.1158/0008-5472. CAN-08-4859.

[17] J. Sapudom, L. Kalbitzer, X. Wu, S. Martin, K. Kroy, and T. Pompe, “Fibril bending stiffness of 3D collagen matrices instructs spreading and clustering of invasive and non-invasive breast cancer cells,” Biomaterials, vol. 193, pp. 47-57, Feb. 2019, doi: 10.1016/j.biomaterials.2018.12.010.

[18] Z. Ahmad et al., “Versatility of Hydrogels: From Synthetic Strategies, Classification, and Properties to Biomedical Applications,” Gels, vol. 8, no. 3, Art. no. 3, Mar. 2022, doi: 10.3390/gels8030167.

[19] R. D. Kasai et al., “A review on hydrogels classification and recent developments in biomedical applications,” International Journal of Polymeric Materials and Polymeric Biomaterials, vol. 72, no. 13, pp. 1059-1069, Sep. 2023, doi: 10.1080/00914037.2022.2075872.

[20] A. S. Hoffman, “Hydrogels for biomedical applications,” Advanced Drug Delivery Reviews, vol. 64, pp. 18-23, Dec. 2012, doi: 10.1016/j.addr.2012.09.010.

[21] E. Prince and E. Kumacheva, “Design and applications of man-made biomimetic fibrillar hydrogels,” Nat Rev Mater, vol. 4, no. 2, Art. no. 2, Feb. 2019, doi: 10.1038/s41578-018-0077-9.

[22] A. Avendano et al., “Integrated Biophysical Characterization of Fibrillar Collagen-Based Hydrogels,” ACS Biomater. Sci. Eng., vol. 6, no. 3, pp. 1408- 1417, Mar. 2020, doi: 10.1021/acsbiomaterials.9b01873.

[23] B. M. Baker et al., “Cell-mediated fibre recruitment drives extracellular matrix mechanosensing in engineered fibrillar microenvironments,” Nature Mater, vol. 14, no. 12, Art. no. 12, Dec. 2015, doi: 10.1038/nmat4444.

[24] M. S. Sakar, J. Eyckmans, R. Pieters, D. Eberli, B. J. Nelson, and C. S. Chen, “Cellular forces and matrix assembly coordinate fibrous tissue repair,” Nat Commun, vol. 7, no. 1, Art. no. 1, Mar. 2016, doi: 10.1038/ncommsl l036.

[25] B. Trappmann, B. M. Baker, W. J. Polacheck, C. K. Choi, J. A. Burdick, and C. S. Chen, “Matrix degradability controls multicellularity of 3D cell migration,” Nat Commun, vol. 8, no. 1, Art. no. 1, Aug. 2017, doi: 10.1038/s41467-017-00418-6.

[26] Z. Chen et al., “Printing Structurally Anisotropic Biocompatible Fibrillar Hydrogel for Guided Cell Alignment,” Gels, vol. 8, no. 11, Art. no. 11, Nov. 2022, doi: 10.3390/gels8110685.

[27] S. Qiao, T. Peijie, and J. Nan, “Crosslinking strategies of decellularized extracellular matrix in tissue regeneration,” Journal of Biomedical Materials Research Part A, vol. 112, no. 5, pp. 640-671, 2024, doi: 10.1002/jbm.a.37650. [28] J. Kort-Mascort et al., “Decellularized ECM hydrogels: prior use considerations, applications, and opportunities in tissue engineering and biofabrication,” Biomaterials Science, vol. 11, no. 2, pp. 400-431, 2023, doi: 10.1039/D2BM01273A.

[29] J. Yoon, H. Han, and J. Jang, “Nanomaterials-incorporated hydrogels for 3D bioprinting technology,” Nano Convergence, vol. 10, no. 1, p. 52, Nov. 2023, doi: 10.1186/s40580-023-00402-5.

[30] J. Kim, J. Jang, and D.-W. Cho, “Controlling Cancer Cell Behavior by Improving the Stiffness of Gastric Tissue-Decellularized ECM Bioink With Cellulose Nanoparticles,” Front. Bioeng. Biotechnol., vol. 9, Mar. 2021, doi: 10.3389/fbioe.2021.605819.

[31] D. Chimene, R. Kaunas, and A. K. Gaharwar, “Hydrogel Bioink Reinforcement for Additive Manufacturing: A Focused Review of Emerging Strategies,” Advanced Materials, vol. 32, no. 1, p. 1902026, 2020, doi: 10.1002/adma.201902026.

[32] M. A. Campea, M. J. Majcher, A. Lofts, and T. Hoare, “A Review of Design and Fabrication Methods for Nanoparticle Network Hydrogels for Biomedical, Environmental, and Industrial Applications,” Advanced Functional Materials, vol. 31, no. 33, p. 2102355, 2021, doi: 10.1002/adfm.202102355.

[33] S. M. Bakht, A. Pardo, M. Gomez-Florit, R. L. Reis, R. M. A. Domingues, and M. E. Gomes, “Engineering next-generation bioinks with nanoparticles: moving from reinforcement fillers to multifunctional nanoelements,” J. Mater. Chem. B, vol. 9, no. 25, pp. 5025-5038, Jun. 2021, doi: 10.1039/D1TB00717C.

[34] Y. Bao, J. He, K. Song, J. Guo, X. Zhou, and S. Liu, “Functionalization and Antibacterial Applications of Cellulose-Based Composite Hydrogels,” Polymers, vol. 14, no. 4, Art. no. 4, Jan. 2022, doi: 10.3390/polyml4040769.

[35] S. M. F. Kabir, P. P. Sikdar, B. Haque, M. A. R. Bhuiyan, A. Ah, and M. N. Islam, “Cellulose-based hydrogel materials: chemistry, properties and their prospective applications,” Prog Biomater, vol. 7, no. 3, pp. 153-174, Sep. 2018, doi: 10.1007/s40204-018-0095-0.

[36] K. J. De France, T. Hoare, and E. D. Cranston, “Review of Hydrogels and Aerogels Containing Nanocellulose,” Chem. Mater., vol. 29, no. 11, pp. 4609- 4631, Jun. 2017, doi: 10.1021/acs.chemmater.7b00531.

[37] A. K. Rana, E. Frollini, and V. K. Thakur, “Cellulose nanocrystals: Pretreatments, preparation strategies, and surface functionalization,” International Journal of Biological Macromolecules, vol. 182, pp. 1554-1581, Jul. 2021, doi: 10.1016/j.ijbiomac.2021.05.119.

[38] F. Rol, M. N. Belgacem, A. Gandini, and J. Bras, “Recent advances in surface- modified cellulose nanofibrils,” Progress in Polymer Science, vol. 88, pp. 241- 264, Jan. 2019, doi: 10.1016/j.progpolymsci.2018.09.002.

[39] M. Dabaghi et al., “A Robust Protocol for Decellularized Human Lung Bioink Generation Amenable to 2D and 3D Lung Cell Culture,” Cells, vol. 10, no. 6, p. 1538, Jun. 2021, doi: 10.3390/cellsl0061538.

[40] A. Isogai and L. Bergstrom, “Preparation of cellulose nanofibers using green and sustainable chemistry,” Current Opinion in Green and Sustainable Chemistry, vol. 12, pp. 15-21, Aug. 2018, doi: 10.1016/j.cogsc.2018.04.008.

[41] S. P. Bangar et al., “Surface modifications of cellulose nanocrystals: Processes, properties, and applications,” Food Hydrocolloids , vol. 130, p. 107689, Sep. 2022, doi: 10.1016/j.foodhyd.2022.107689. [42] F. Torabizadeh, M. Fadaie, E. Mirzaei, S. Sadeghi, and G.-R. Nejabat, “Tailoring structural properties, mechanical behavior and cellular performance of collagen hydrogel through incorporation of cellulose nanofibrils and cellulose nanocrystals: A comparative study,” International Journal of Biological Macromolecules, vol. 219, pp. 438-451, Oct. 2022, doi: 10.1016/j.ijbiomac.2022.08.006.

[43] K. Sahajpal et al., “Dynamic protein and polypeptide hydrogels based on Schiff base co-assembly for biomedicine,” J. Mater. Chem. B, vol. 10, no. 17, pp. 3173— 3198, May 2022, doi: 10.1039/D2TB00077F.

[44] S. Zhang, D. Huang, H. Lin, Y. Xiao, and X. Zhang, “Cellulose Nanocrystal Reinforced Collagen-Based Nanocomposite Hydrogel with Self-Healing and Stress-Relaxation Properties for Cell Delivery,” Biomacromolecules, vol. 21, no. 6, pp. 2400-2408, Jun. 2020, doi: 10.1021/acs.biomac.0c00345.

[45] R. H. J. de Hilster et al. , “Human lung extracellular matrix hydrogels resemble the stiffness and viscoelasticity of native lung tissue,” American Journal of Physiology-Lung Cellular and Molecular Physiology, vol. 318, no. 4, pp. L698- L704, Apr. 2020, doi: 10.1152/ajplung.00451.2019.

[46] A. J. Booth et al., “Acellular Normal and Fibrotic Human Lung Matrices as a Culture System for In Vitro Investigation,” Am J Respir Crit Care Med, vol. 186, no. 9, pp. 866-876, Nov. 2012, doi: 10.1164/rccm.201204-07540C.

[47] F. Liu et al., “Distal vessel stiffening is an early and pivotal mechanobiological regulator of vascular remodeling and pulmonary hypertension,” JCI Insight, vol. 1, no. 8, Jun. 2016, doi: 10.1172/jci.insight.86987.

[48] K. S. Saleh et al., “Engineering Hybrid-Hydrogels Comprised of Healthy or Diseased Decellularized Extracellular Matrix to Study Pulmonary Fibrosis,” Cel. Mol. Bioeng, vol. 15, no. 5, pp. 505-519, Oct. 2022, doi: 10.1007/sl2195-022- 00726-y.

[49] S. O. Sarrigiannidis, J. M. Rey, O. Dobre, C. Gonzalez-Garcia, M. J. Dalby, and M. Salmeron- Sanchez, “A tough act to follow: collagen hydrogel modifications to improve mechanical and growth factor loading capabilities,” Materials Today Bio, vol. 10, p. 100098, Mar. 2021, doi: 10.1016/j.mtbio.202L 100098.

[50] R. Parenteau-Bareil, R. Gauvin, and F. Berthod, “Collagen-Based Biomaterials for Tissue Engineering Applications,” Materials, vol. 3, no. 3, Art. no. 3, Mar. 2010, doi: 10.3390/ma3031863.

[51] F. Chua, J. Gauldie, and G. J. Laurent, “Pulmonary Fibrosis,” Am J Respir Cell Mol Biol, vol. 33, no. 1, pp. 9-13, Jul. 2005, doi: 10.1165/rcmb.2005-0062TR.

[52] M. Dabaghi, M. Barreiro Carpio, N. Saraei, J. M. Moran-Mirabal, M. R. Kolb, and J. A. Hirota, “A Roadmap for Developing and Engineering in vitro Pulmonary Fibrosis Models,” Biophysics Reviews, vol. 4, no. 2, p. 021302, Jun. 2023, doi: 10.1063/5.0134177.

[53] N. C. Henderson, F. Rieder, and T. A. Wynn, “Fibrosis: from mechanisms to medicines,” Nature, vol. 587, no. 7835, Art. no. 7835, Nov. 2020, doi: 10.1038/s41586-020-2938-9.

[54] B. Hinz, “Mechanical Aspects of Lung Fibrosis,” Proc Am Thorac Soc, vol. 9, no. 3, pp. 137-147, Jul. 2012, doi: 10.1513/pats.201202-017AW.

[55] B. Hinz et al., “Recent Developments in Myofibroblast Biology: Paradigms for Connective Tissue Remodeling,” The American Journal of Pathology, vol. 180, no. 4, pp. 1340-1355, Apr. 2012, doi: 10.1016/j.ajpath.2012.02.004. [56] F. S. Younesi, A. E. Miller, T. H. Barker, F. M. V. Rossi, and B. Hinz, “Fibroblast and myofibroblast activation in normal tissue repair and fibrosis,” Nat Rev Mol Cell Biol, pp. 1-22, Apr. 2024, doi: 10.1038/s41580-024-00716-0.

[57] E. Brauer etal., “Collagen Fibrils Mechanically Contribute to Tissue Contraction in an In Vitro Wound Healing Scenario,” Advanced Science, vol. 6, no. 9, p. 1801780, 2019, doi: 10.1002/advs.201801780.

[58] S. Masterton and M. Aheame, “Influence of poly dimethylsiloxane substrate stiffness on comeal epithelial cells,” Royal Society Open Science, vol. 6, no. 12, p. 191796, Dec. 2019, doi: 10.1098/rsos.l91796.

[59] J. He, B. Fang, S. Shan, and Q. Li, “Mechanical stiffness promotes skin fibrosis through Piezo 1 -mediated arginine and proline metabolism,” Cell Death Discov., vol. 9, no. 1, pp. 1-9, Sep. 2023, doi: 10.1038/s41420-023-01656-y.

[60] G. Bartalena, R. Grieder, R. I. Sharma, T. Zambelli, R. Muff, and J. G. Snedeker, “A novel method for assessing adherent single-cell stiffness in tension: design and testing of a substrate-based live cell functional imaging device,” Biomed Microdevices, vol. 13, no. 2, pp. 291-301, Apr. 2011, doi: 10.1007/sl0544-010- 9493-3.

[61] J. L. Balestrini, S. Chaudhry, V. Sarrazy, A. Koehler, and B. Hinz, “The mechanical memory of lung myofibroblasts,” Integr. Biol., vol. 4, no. 4, p. 410, 2012, doi: 10.1039/c2ib00149g.

[62] K. E. C. Blokland et al., “Substrate stiffness engineered to replicate disease conditions influence senescence and fibrotic responses in primary lung fibroblasts,” Front. Pharmacol., vol. 13, p. 989169, Nov. 2022, doi: 10.3389/fphar.2022.989169.

[63] S. Asano et al., “Matrix stiffness regulates migration of human lung fibroblasts,” Physiol Rep, vol. 5, no. 9, p. e!3281, May 2017, doi: 10.14814/phy2.13281.

[64] L. G. Major et al., “Volume Adaptation Controls Stem Cell Mechanotransduction,” ACS Appl. Mater. Interfaces, vol. 11, no. 49, pp. 45520- 45530, Dec. 2019, doi: 10.1021/acsami.9bl9770.

[65] C. Kim et al., “Stem Cell Mechanosensation on Gelatin Methacryloyl (GelMA) Stiffness Gradient Hydrogels,” Ann Biomed Eng, vol. 48, no. 2, pp. 893-902, Feb.

2020, doi: 10.1007/s 10439-019-02428-5.

[66] D. L. Matera, A. T. Lee, H. L. Hiraki, and B. M. Baker, “The Role of Rho GTPases During Fibroblast Spreading, Migration, and Myofibroblast Differentiation in 3D Synthetic Fibrous Matrices,” Cel. Mol. Bioeng, vol. 14, no. 5, pp. 381-396, Oct.

2021, doi: 10.1007/sl2195-021-00698-5.

[67] C. D. Davidson et al., “Myofibroblast activation in synthetic fibrous matrices composed of dextran vinyl sulfone, "Acta Biomat er ialia, vol. 105, pp. 78-86, Mar. 2020, doi: 10.1016/j.actbio.2020.01.009.

[68] B. C. Heng et al., “An overview of signaling pathways regulating YAP/TAZ activity,” Cell. Mol. Life Sci., vol. 78, no. 2, pp. 497-512, Jan. 2021, doi: 10.1007/s00018-020-03579-8.

[69] S. Piccolo, S. Dupont, and M. Cordenonsi, “The Biology of YAP/TAZ: Hippo Signaling and Beyond,” Physiological Reviews, vol. 94, no. 4, pp. 1287-1312, Oct. 2014, doi: 10.1152/physrev.00005.2014.

[70] G. Brusatin, T. Panciera, A. Gandin, A. Citron, and S. Piccolo, “Biomaterials and engineered microenvironments to control YAP/T AZ-dependent cell behaviour,” Nature Mater, vol. 17, no. 12, pp. 1063-1075, Dec. 2018, doi: 10.1038/s41563- 018-0180-8. [71] A. Das, R. S. Fischer, D. Pan, and C. M. Waterman, “YAP Nuclear Localization in the Absence of Cell-Cell Contact Is Mediated by a Filamentous Actindependent, Myosin II- and Phospho-YAP-independent Pathway during Extracellular Matrix Mechanosensing,” Journal of Biological Chemistry, vol. 291, no. 12, pp. 6096-6110, Mar. 2016, doi: 10.1074/jbc.M115.708313.

[72] A. Totaro et al., “YAP/TAZ link cell mechanics to Notch signalling to control epidermal stem cell fate,” Nat Commun, vol. 8, no. 1, p. 15206, May 2017, doi: 10.1038/ncomms 15206.

[73] B. Sun, Q. Hou, Z. Liu, and Y. Ni, “Sodium periodate oxidation of cellulose nanocrystal and its application as a paper wet strength additive,” Cellulose, vol. 22, no. 2, pp. 1135-1146, Apr. 2015, doi: 10.1007/sl0570-015-0575-5.

Claims

Claims:

1. A composite hydrogel comprising a decellularized extracellular matrix (dECM) and surface-modified cellulose nanofibers (CNFs).

2. The composite hydrogel of claim 1, wherein the surface-modified CNFs comprise one or more functional groups on a surface of the CNFs, and wherein the groups are selected from aldehyde, methacrylate, carboxylate, carboxylic acid, vinyl, thiol, alkyne, azide, and hydrazide.

3. The composite hydrogel of claim 1 or 2, wherein the surface-modified CNFs comprise methacrylate functional groups on the surface of the CNFs.

4. The composite hydrogel of claim 1 or 2, wherein the surface-modified CNFs comprise aldehyde functional groups on the surface of the CNFs.

5. The composite hydrogel of any one of claims 1 to 4, wherein the surface- modified CNFs are present in the hydrogel in an amount of about 0.1 % w/v to about 2 % w/v.

6. The composite hydrogel of any one of claims 1 to 5, wherein the dECM is a digested dECM.

7. The composite hydrogel of any one of claims 1 to 6, wherein the dECM comprises ECM from a mammalian tissue.

8. The composite hydrogel of any one of claims 1 to 7, wherein the dECM comprises ECM from one or more of lung, liver, brain skin, kidney, and heart.

9. The composite hydrogel of any one of claims 1 to 8, wherein the dECM is present in the hydrogel in an amount of about 1 % w/v to about 3 % w/v.

10. The composite hydrogel of any one of claims 1 to 8, wherein the weight ratio of the dECM to the surface-modified CNFs is about 30:1 to about 1:2.

11. The composite hydrogel of any one of claims 1 to 10, wherein the composite hydrogel further comprises one or more additives.

12. The composite hydrogel of claim 11, wherein the one or more additives are selected from cytokines, cells, cell culture media, and bioactive growth factors.

13. The composite hydrogel of any one of claims 2 to 12, wherein the functional groups on the surface of the CNFs are further functionalized and/or conjugated to bioactive molecules.

14. The composite hydrogel of any one of claims 1 to 13, wherein the composite hydrogel is a fibrous hydrogel.

15. The composite hydrogel of any one of claims 1 to 14, wherein the composite hydrogel possesses improved mechanical properties compared to mechanical properties of a same hydrogel but without the surface-modified CNFs.

16. The composite hydrogel of claim 15, wherein the improved mechanical properties are selected from one or more of increased storage modulus, loss modulus, and compression modulus.

17. The composite hydrogel of any one of claims 1 to 16, wherein the composite hydrogel is reinforced while the physical structures of the hydrogels remain substantially unchanged.

18. The composite hydrogel of claim 17, wherein the physical structures are selected from one or more of fiber dimension and porosity, redistribution of internal stress and viscoelastic properties.

19. The composite hydrogel of any one of claims 1 to 18, wherein the composite hydrogel has compression moduli of greater than or equal to 7kPa.

20. The composite hydrogel of claim 19, wherein the composite hydrogel has compression moduli of about 7kPa to 25kPa.

21. The composite hydrogel of any one of claims 1 to 20, wherein the composite hydrogel has a reduced hydrolytic degradation compared to a hydrolytic degradation of a same hydrogel but without the surface-modified CNFs.

22. The composite hydrogel of any one of claims 1 to 21, wherein the composite hydrogel has similar macromolecule permeability compared to a macromolecule permeability of a same hydrogel but without the surface-modified CNFs.

23. The composite hydrogel of any one of claims 1 to 22 for use in tissue modelling, tissue engineering, as bioinks for bioprinting, analytical applications, and drug screening and delivery.

24. The composite hydrogel of for use of claim 23, wherein the tissue modelling is 2D or 3D cell culture and the analytical applications are selected from microfluidic platforms and gel contraction assays.

25. The composite hydrogel of any one of claims 1 to 22 for use in modelling healthy and diseased tissues.

26. The composite hydrogel of any one of claims 1 to 22 for use in forming a 3D model for fibroblast-to-myofibroblast transition.

27. The composite hydrogel of any one of claims 1 to 22 for use in modelling pulmonary fibrosis.

28. A method of preparing a composite gel of any one of claims 1 to 22 comprising combining a decellularized extracellular matrix (dECM) and a surface-modified cellulose nanofibers (CNFs).

29. The method of claim 28, wherein the dECM is in the form of a liquid solution.

30. The method of claim 29, wherein the dECM liquid solution comprises a buffer.

31. The method of claim 30, wherein the buffer is phosphate-buffered saline (PBS).

32. The method of any one of claims 29 to 31, wherein the pH of the dECM liquid solution is from about 7.2 to about 7.4.

33. The method of any one of claims 28 to 32, wherein the surface-modified CNFs are in the form of an aqueous suspension.

34. The method of claim 33, wherein the aqueous suspension of the surface- modified CNFs comprises a buffer.

35. The method of claim 34, wherein the buffer is Hank's Balanced Salt Solution (HBSS).