High-density microchannels

Abstract

To provide improved scaffold biomaterials to substitute hydrogels, and methods for their production.SOLUTION: Provided herein are scaffold biomaterials including at least one bundle of microchannels, the bundle of microchannels having a plurality of decellularised microchannels isolated from plant or fungal tissue, the decellularised microchannels being arranged substantially parallel to each other within the bundle. Also provided are methods and uses of such scaffold biomaterials and bundles, as well as methods for the production of such scaffold biomaterials and bundles.SELECTED DRAWING: None

Description

[Technical Field]

[0001] The present invention relates generally to scaffold biomaterials. More particularly, the present invention relates to high-density decellularized microchannel bundles derived from plant tissue and their use as scaffold biomaterials. [Background technology]

[0002] The biomaterials industry, estimated to have a market value of US$90 billion, is driven by novel materials derived from natural sources, synthetic polymers, metals, and ceramics. These materials can form biocompatible, three-dimensional, highly porous scaffolds with nano- and micro-scale structures that promote the growth of living cells. For example, there is intense interest in novel biomaterials that support the infiltration and proliferation of living cells for potential applications in tissue engineering and regenerative medicine.

[0003] Biomaterial scaffolds have applications in multiple sectors, including dentistry and cosmetic surgery, clinical and medical therapy (e.g., regenerative medicine, wound healing, tissue engineering and repair, etc.), and research and development (including industrial and academic research in biomedical sciences).

[0004] Commercially available biomaterials often require complex and time-consuming production methods, which translates into high costs for end users even when they are not approved for human use. Furthermore, most commercially available biomaterials are derived from human / animal sources, posing the potential risk of rejection by the body and / or adverse immune responses and / or infectious disease transmission. Source materials may also have negative environmental impacts, potentially leading to concerns about unethical sourcing. Additionally, some commercially available biomaterials lose their shape after implantation, potentially reducing the success of tissue repair / replacement.

[0005] The field generally lacks viable commercially available biomaterial solutions to replace directional, channel-like tissue and organ replacements that promote tissue repair and / or regeneration. For example, in the field of spinal cord injuries (SCI), most academic research currently focuses on injectable hydrogels containing various factors intended to regulate and promote neuronal growth. However, hydrogels often lack directional and neuronal guidance capabilities.

[0006] Alternative, additional, and / or improved scaffold biomaterials and methods for producing the same are desirable. Summary of the Invention [Means for solving the problem]

[0007] Provided herein are materials (biomaterials) that can be used to mimic the directional channel-like microscopic structures of certain tissues in humans, plants, and animals. Examples of such directional tissues include the extracellular matrices (ECM) of the spinal cord, vascular channels, lymphatic tissue, nervous tissue, and many others. It is an object of the present invention to provide biocompatible materials with directional and / or channel-like structures, as well as methods for producing, and using the same.

[0008] In one embodiment, provided herein is a scaffold biomaterial comprising at least one bundle of microchannels, the bundle of microchannels comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle.

[0009] In another embodiment of the above scaffold biomaterial, the decellularized microchannels may comprise decellularized xylem and / or phloem channels.

[0010] In another embodiment of any of the above scaffold biomaterials, the decellularized xylem and / or phloem channels can be individually isolated from the plant or fungal tissue, can be grouped into one or more vascular bundles isolated from the plant or fungal tissue, or any combination thereof.

[0011] In another embodiment of any of the above scaffold biomaterials, the decellularized microchannels are substantially adhesive-free, eg, physically bound together by entanglement.

[0012] In yet another embodiment of any of the above scaffold biomaterials, multiple decellularized microchannels may be glued together in a bundle.

[0013] In yet another embodiment of any of the above scaffold biomaterials, multiple decellularized microchannels may be adhered together in a bundle by a biocompatible adhesive, optionally a biodegradable adhesive, optionally a low volume expansion adhesive.

[0014] In another embodiment of any of the above scaffold biomaterials, the biocompatible adhesive may comprise a PEG-based, polyurethane-based, gelatin-based, or fibrin-based adhesive.

[0015] In yet another embodiment of any of the above scaffold biomaterials, the biocompatible adhesive may comprise a fibrin-based adhesive.

[0016] In yet another embodiment of any of the above scaffold biomaterials, the decellularized microchannels may be cellulose-based, chitin-based, lignin-based, hemicellulose-based, or pectin-based, or any combination thereof.

[0017] In another embodiment of any of the above scaffold biomaterials, the density of decellularized microchannels within the bundles may be greater than the density of microchannels within plant or fungal tissue.

[0018] In yet another embodiment of any of the above scaffold biomaterials, the plant or fungal tissue is selected from the group consisting of apple thixosaur (Apple) tissue, fern (Pteridophyta) tissue, turnip (Brassica rapa) root tissue, ginkgo biloba branch tissue, horsetail (Equisetum aestivum) tissue, hermocallis hybrid leaf tissue, and the like. , kale (Brassica oleracea) stem tissue, conifer Douglas fir (Douglas fir) tissue, cactus fruit (pitaya) flesh tissue, Maculata Vinca tissue, water lotus (Nelumbo nucifera) tissue, tulip (Tulipa gesneriana) petal tissue, plantain (banana) tissue, broccoli (Brassica oleracea) A) Stem tissue, stem tissue of maple leaves (Acer psuedoplatanus), sugar beet (sugar beet) primary root tissue, leek (onion) tissue, orchid (Orchidaceae) tissue, turnip (Brassica rapa) stem tissue, leek (Allium ampeloprasum) ) tissue, maple (Acer spp.) tree branch tissue, celery (Apium graveolens) tissue, leek (onion) stem tissue, pine tissue, aloe vera tissue, watermelon (Citrullus lanatus var. lanatus) tissue, creeping jenny (Solanum spp.) tissue, cactus (cactae) tissue, Lychnis alpina tissue, rhubarb (Rheum rhabarbarum) tissue, pumpkin pulp (Cucurbita pepo) tissue, Dracaena (Asparagaceae) stem Tissue, Tradescantia reflexa (Tradescantia reflexa) stem tissue, Asparagus (Asparagus officinalis) stem tissue, Mushroom (fungal) tissue, Fenne Fennel tissue, rose (Rosa spp.) tissue, carrot (Daucus carota ota) tissue, or pear (pomoea) tissue, or by direct genome modification or genetically modified tissue produced by selective breeding, or any combination thereof.

[0019] In yet another embodiment of any of the above scaffold biomaterials, the plant or fungal tissue may comprise celery, asparagus, or both.

[0020] In another embodiment of any of the above scaffold biomaterials, the scaffold biomaterial may further comprise living cells, particularly non-native cells, on and / or within at least one of the decellularized microchannels.

[0021] In yet another embodiment of any of the above scaffold biomaterials, the living cells may be animal cells.

[0022] In yet another embodiment of any of the above scaffold biomaterials, the living cells may be mammalian cells.

[0023] In another embodiment of any of the above scaffold biomaterials, the living cells may be human cells.

[0024] In another embodiment of any of the above scaffold biomaterials, the decellularized microchannels may be separated from the plant or fungal material by liquid-based extraction.

[0025] In another embodiment of any of the above scaffold biomaterials, the separation may be by microchannel liquid-based extraction from the plant or fungal tissue.

[0026] In another embodiment of any of the above scaffold biomaterials, the liquid-based extraction may comprise at least one of an acid extraction, an acid and peroxide extraction, a salt extraction, or an alkaline extraction.

[0027] In another embodiment of any of the above scaffold biomaterials, the acid and peroxide extraction may comprise heating the plant or fungal tissue in an acid and peroxide solution.

[0028] In another embodiment of any of the above scaffold biomaterials, the acid and peroxide solution may comprise glacial acetic acid and 30% hydrogen peroxide in a ratio of 5:1 to 1:5, such as 3:1 to 1:3.

[0029] In another embodiment of any of the above scaffold biomaterials, the acid and peroxide solution may comprise a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v).

[0030] In other embodiments of any of the above scaffold biomaterials, the heating step may comprise heating for a period of up to 30 minutes or longer.

[0031] In another embodiment of any of the above scaffold biomaterials, the heating step may comprise heating for 30 minutes.

[0032] In another embodiment of any of the above scaffold biomaterials, the heating step may include boiling the acid and peroxide solution.

[0033] In another embodiment of any of the above scaffold biomaterials, the acid and peroxide solution can include a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v), and the solution is heated to boiling for 30 minutes.

[0034] In another embodiment of any of the above scaffold biomaterials, the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid and peroxide solution.

[0035] In another embodiment of any of the above scaffold biomaterials, the salt extraction may comprise heating the plant or fungal tissue in a salt solution.

[0036] In another embodiment of any of the above scaffold biomaterials, the saline solution may comprise LiCl or NaCl.

[0037] In another embodiment of any of the above scaffold biomaterials, the salt solution may comprise a salt concentration of about 0.5M to 10M, for example, 0.5M to 3M.

[0038] In another embodiment of any of the above scaffold biomaterials, the saline solution may comprise LiCl or NaCl at a salt concentration of about 3M.

[0039] In other embodiments of any of the above scaffold biomaterials, the heating step may comprise heating for a period of up to 30 minutes or longer.

[0040] In another embodiment of any of the above scaffold biomaterials, the heating step may comprise heating for 30 minutes.

[0041] In another embodiment of any of the above scaffold biomaterials, the heating step may include boiling the saline solution.

[0042] In another embodiment of any of the above scaffold biomaterials, the saline solution may include about 3 M LiCl, and the solution may be heated to boiling for 30 minutes.

[0043] In another embodiment of any of the above scaffold biomaterials, the saline solution may comprise about 3 M NaCl, and the solution may be heated to boiling for 15 minutes.

[0044] In another embodiment of any of the above scaffold biomaterials, the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the salt solution.

[0045] In another embodiment of any of the above scaffold biomaterials, the alkaline extraction may comprise heating the plant or fungal tissue in an alkaline solution.

[0046] In another embodiment of any of the above scaffold biomaterials, the alkaline solution may comprise sodium hydroxide (NaOH).

[0047] In another embodiment of any of the above scaffold biomaterials, the alkaline solution may comprise an alkaline concentration of about 0.5 to 10M, for example, 0.5M to 3M.

[0048] In another embodiment of any of the above scaffold biomaterials, the alkaline solution may comprise sodium hydroxide at an alkaline concentration of about 0.5M to 1M.

[0049] In another embodiment of any of the above scaffold biomaterials, the heating step may comprise heating for up to 30 minutes.

[0050] In another embodiment of any of the above scaffold biomaterials, the heating step may comprise heating for 30 minutes.

[0051] In another embodiment of any of the above scaffold biomaterials, the heating step may include boiling the alkaline solution.

[0052] In another embodiment of any of the above scaffold biomaterials, the alkaline solution can include about 0.5 M NaOH, and the solution is heated to boiling for 5 minutes.

[0053] In another embodiment of any of the above scaffold biomaterials, the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the alkaline solution.

[0054] In another embodiment of any of the above scaffold biomaterials, the acid extraction may comprise heating the plant or fungal tissue in an acid solution comprising an acid.

[0055] In another embodiment of any of the above scaffold biomaterials, the acid solution may comprise an acidic acid or hydrochloric acid.

[0056] In another embodiment of any of the above scaffold biomaterials, the acid solution may comprise 50% acetic acid.

[0057] In other embodiments of any of the above scaffold biomaterials, the heating step may comprise heating for a period of up to 30 or more minutes.

[0058] In another embodiment of any of the above scaffold biomaterials, the heating step may comprise heating for 30 minutes.

[0059] In another embodiment of any of the above scaffold biomaterials, the heating step may include boiling the acid solution.

[0060] In another embodiment of any of the above scaffold biomaterials, the acid solution can include 50% acetic acid, and the solution is heated to boiling for 30 minutes.

[0061] In another embodiment of any of the above scaffold biomaterials, the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid solution.

[0062] In another embodiment of any of the above scaffold biomaterials, the decellularization step may occur before the separation step.

[0063] In another embodiment, provided herein is the use of any of the scaffold biomaterials as described herein for supporting animal or plant cell growth, promoting tissue regeneration, promoting vascularization, treating and / or repairing spinal cord injury, repairing or reconstructing or replacing plant or animal tissue, filtering and / or separating solutions or materials, modifying or regulating growth of plants, material transport, fabrication to mimic desired shapes or objects, and / or for microfluidics, or any suitable application where biocompatible micrometer-scale channels may be useful.

[0064] In another embodiment of the above use, the scaffold biomaterial is used for the repair or reconstruction of plant or animal tissue. or replacement, and the plant or animal tissue may include damaged microvasculature, or plant tissue damaged by vascular disease (such as wilt), or damaged vasculature of wood infested by insects (such as emerald ash borers).

[0065] In another embodiment of any of the above uses, the scaffold biomaterial is for the repair or reconstruction of plant or animal tissue, which may comprise a damaged microvasculature, in which vascularization is impaired or impaired.

[0066] In yet another embodiment of any of the above uses, the scaffold biomaterial may be for the repair or reconstruction of animal tissue, wherein the animal is a human.

[0067] In yet another embodiment of any of the above uses, the scaffold biomaterial may be for plant modification or growth regulation, wherein the modification may be modification of the implantation process for accelerated growth.

[0068] In another embodiment of any of the above uses, the scaffold biomaterial may be for material transport, wherein the material transport is for drug delivery applications.

[0069] In yet another embodiment of any of the above uses, the scaffold biomaterial may be for heat transfer or heat exchange microfluidics.

[0070] In another embodiment, there is provided a method of supporting cell proliferation, comprising: providing any of the scaffold biomaterials as described herein; introducing one or more cells into the scaffold biomaterial; Provided herein is a method comprising:

[0071] In another embodiment, there is provided a method for promoting tissue regeneration, promoting angiogenesis, treating and / or repairing spinal cord injury, or repairing or reconstructing or replacing plant or animal tissue, comprising: providing any of the scaffold biomaterials as described herein; implanting the scaffold biomaterial at a site in need thereof; Provided herein is a method comprising:

[0072] In another embodiment, there is provided a method for the filtration and / or separation of a solution or material, comprising the steps of: providing any of the scaffold biomaterials as described herein; passing a solution or material through the scaffold biomaterial to filter and / or separate components from the solution based on size exclusion; Provided herein is a method comprising:

[0073] In another embodiment, there is provided a method for modifying or regulating the growth of a plant, comprising: providing any of the scaffold biomaterials as described herein; Implanting the scaffold biomaterial into a plant to provide accelerated growth. Provided herein is a method comprising:

[0074] In another embodiment, there is provided a method for material transport, comprising: providing any of the scaffold biomaterials as described herein; using the scaffold biomaterial to transport the material to the site where it is needed; Provided herein is a method comprising:

[0075] In another embodiment of the above method, the material may be a drug and the scaffold biomaterial may be used for drug delivery.

[0076] In another embodiment, there is provided a method for preparing a structure that mimics a desired shape or object, comprising the steps of: providing any of the scaffold biomaterials as described herein; shaping or positioning the scaffold biomaterial to mimic a desired shape or object; Optionally, attaching a scaffold biomaterial for structural reinforcement. Provided herein is a method comprising:

[0077] In another embodiment, there is provided a method of exchanging or transferring heat in a microfluidic process, comprising: providing any of the scaffold biomaterials as described herein; Provided herein is a method comprising using the microchannels of the scaffold biomaterial to carry one or more fluids, wherein the one or more fluids are in close proximity to allow for heat exchange or heat transfer.

[0078] In another embodiment, there is provided a method for isolating and decellularizing microchannels from plant or fungal tissue, comprising: isolating a microchannel from the plant or fungal tissue; decellularizing the microchannel; Optionally, sterilizing the microchannel. Provided herein is a method comprising:

[0079] In another embodiment of the above method, the step of isolating the microchannels may include mechanical separation of the microchannels or the vascular bundles containing the microchannels, or both, from the surrounding plant or fungal tissue.

[0080] In yet another embodiment of the above method, the separating step may be performed by gentle peeling or by cutting.

[0081] In another embodiment of any of the above method(s), the decellularized microchannels may be separated from the plant or fungal material by liquid-based extraction.

[0082] In another embodiment of any of the above method(s), the separation may be by microchannel liquid-based extraction from the plant or fungal tissue.

[0083] In another embodiment of any of the above method(s), the liquid-based extraction may include at least one of an acid extraction, an acid and peroxide extraction, a salt extraction, or an alkaline extraction.

[0084] In another embodiment of any of the above method(s), the acid and peroxide extraction may include heating the plant or fungal tissue in an acid and peroxide solution.

[0085] In another embodiment of any of the above method(s), the acid and peroxide solution may comprise glacial acetic acid and 30% hydrogen peroxide in a ratio of 5:1 to 1:5, for example, 3:1 to 1:3.

[0086] In another embodiment of any of the above method(s), the acid and peroxide solution may comprise a 1:1 glacial acetic acid:hydrogen peroxide solution (30%).

[0087] In another embodiment of any of the above method(s), the heating step comprises heating for a period of up to 30 minutes or longer, e.g., up to 45 minutes or up to 1 hour. It may include.

[0088] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0089] In another embodiment of any of the above method(s), the heating step may include boiling the acid and peroxide solution.

[0090] In another embodiment of any of the above method(s), the acid and peroxide solution can include a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v), and the solution is heated to boiling for 30 minutes.

[0091] In another embodiment of any of the above method(s), the liquid-based extraction may further include mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid and peroxide solution.

[0092] In another embodiment of any of the above method(s), the salt extraction may comprise heating the plant or fungal tissue in a salt solution.

[0093] In another embodiment of any of the above method(s), the saline solution may include LiCl or NaCl.

[0094] In another embodiment of any of the above method(s), the salt solution may comprise a salt concentration of about 0.5M to 10M, for example, 0.5M to 3M.

[0095] In another embodiment of any of the above method(s), the saline solution may include LiCl or NaCl at a salt concentration of about 3M.

[0096] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 minutes or longer, such as up to 45 minutes or up to 1 hour.

[0097] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0098] In another embodiment of any of the above method(s), the heating step may include boiling the salt solution.

[0099] In another embodiment of any of the above method(s), the salt solution can include about 3 M LiCl, and the solution can be heated to boiling for 30 minutes.

[0100] In another embodiment of any of the above method(s), the saline solution may include about 3 M NaCl, and the solution may be heated to boiling for 15 minutes.

[0101] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the salt solution.

[0102] In another embodiment of any of the above method(s), the alkaline extraction may comprise heating the plant or fungal tissue in an alkaline solution.

[0103] In another embodiment of any of the above method(s), the alkaline solution may include sodium hydroxide (NaOH).

[0104] In another embodiment of any of the above method(s), the alkaline solution may comprise an alkaline concentration of about 0.5 to 10 M, for example, 0.5 M to 3 M.

[0105] In another embodiment of any of the above method(s), the alkaline solution may include sodium hydroxide at an alkaline concentration of about 0.5M to 1M.

[0106] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 minutes or longer, such as up to 45 minutes or up to 1 hour.

[0107] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes or longer, for example, up to 45 minutes or up to 1 hour.

[0108] In another embodiment of any of the above method(s), the heating step may include boiling the alkaline solution.

[0109] In another embodiment of any of the above method(s), the alkaline solution can include about 0.5 M NaOH, and the solution is heated to boiling for 5 minutes.

[0110] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the alkaline solution.

[0111] In another embodiment of any of the above method(s), the acid extraction may comprise heating the plant or fungal tissue in an acid solution comprising an acid.

[0112] In another embodiment of any of the above method(s), the acid solution may comprise an acidic acid or hydrochloric acid.

[0113] In another embodiment of any of the above method(s), the acid solution may include 50% acetic acid.

[0114] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 or more.

[0115] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0116] In another embodiment of any of the above method(s), the heating step may include boiling the acid solution.

[0117] In another embodiment of any of the above method(s), the acid solution can include 50% acetic acid, and the solution is heated to boiling for 30 minutes.

[0118] In another embodiment of any of the above method(s), the liquid-based extraction comprises an acid The method may further include mechanically agitating, e.g., stirring, the plant or fungal tissue in the solution.

[0119] In another embodiment of any of the above method(s), the decellularization step may occur before the separation step.

[0120] In another embodiment, there is provided a method for preparing a scaffold biomaterial, comprising: isolating the microchannel from the plant or fungal tissue, or providing a microchannel that is already isolated from the plant or fungal tissue; decellularizing the microchannel; bundling the microchannels together so that the microchannels are positioned substantially parallel to one another; thereby providing a scaffold biomaterial comprising bundled microchannels.

[0121] In yet another embodiment of the above method, the step of isolating the microchannels may include mechanical separation of the microchannels or the vascular bundles containing the microchannels, or both, from the surrounding plant or fungal tissue.

[0122] In another embodiment of any of the above method(s), the separating step may be performed by gentle peeling or by cutting.

[0123] In another embodiment of any of the above method(s), the decellularized microchannels may be separated from the plant or fungal material by liquid-based extraction.

[0124] In another embodiment of any of the above method(s), the separation may be by microchannel liquid-based extraction from the plant or fungal tissue.

[0125] In another embodiment of any of the above method(s), the liquid-based extraction may include at least one of an acid extraction, an acid and peroxide extraction, a salt extraction, or an alkaline extraction.

[0126] In another embodiment of any of the above method(s), the acid and peroxide extraction may include heating the plant or fungal tissue in an acid and peroxide solution.

[0127] In another embodiment of any of the above method(s), the acid and peroxide solution may comprise glacial acetic acid and 30% hydrogen peroxide in a ratio of 5:1 to 1:5, for example, 3:1 to 1:3.

[0128] In another embodiment of any of the above method(s), the acid and peroxide solution may comprise a 1:1 glacial acetic acid:hydrogen peroxide solution (30%).

[0129] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 minutes or longer, such as up to 45 minutes or up to 1 hour.

[0130] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0131] In another embodiment of any of the above method(s), the heating step may include boiling the acid and peroxide solution.

[0132] In another embodiment of any of the above method(s), the acid and peroxide solution can include a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v), and the solution is heated to boiling for 30 minutes.

[0133] In another embodiment of any of the above method(s), the liquid-based extraction may further include mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid and peroxide solution.

[0134] In another embodiment of any of the above method(s), the salt extraction may comprise heating the plant or fungal tissue in a salt solution.

[0135] In another embodiment of any of the above method(s), the saline solution may include LiCl or NaCl.

[0136] In another embodiment of any of the above method(s), the salt solution may comprise a salt concentration of about 0.5M to 10M, for example, 0.5M to 3M.

[0137] In another embodiment of any of the above method(s), the saline solution may include LiCl or NaCl at a salt concentration of about 3M.

[0138] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 minutes or longer, such as up to 45 minutes or up to 1 hour.

[0139] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0140] In another embodiment of any of the above method(s), the heating step may include boiling the salt solution.

[0141] In another embodiment of any of the above method(s), the salt solution can include about 3 M LiCl, and the solution can be heated to boiling for 30 minutes.

[0142] In another embodiment of any of the above method(s), the saline solution may include about 3 M NaCl, and the solution may be heated to boiling for 15 minutes.

[0143] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the salt solution.

[0144] In another embodiment of any of the above method(s), the alkaline extraction may comprise heating the plant or fungal tissue in an alkaline solution.

[0145] In another embodiment of any of the above method(s), the alkaline solution may include sodium hydroxide (NaOH).

[0146] In another embodiment of any of the above method(s), the alkaline solution may comprise an alkaline concentration of about 0.5 to 10 M, for example, 0.5 M to 3 M.

[0147] In another embodiment of any of the above method(s), the alkaline solution may include sodium hydroxide at an alkaline concentration of about 0.5M to 1M.

[0148] In another embodiment of any of the above method(s), the heating step may include heating for up to 30 minutes or longer, such as up to 45 minutes or up to 1 hour.

[0149] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0150] In another embodiment of any of the above method(s), the heating step may include boiling the alkaline solution.

[0151] In another embodiment of any of the above method(s), the alkaline solution can include about 0.5 M NaOH, and the solution is heated to boiling for 5 minutes.

[0152] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the alkaline solution.

[0153] In another embodiment of any of the above method(s), the acid extraction may comprise heating the plant or fungal tissue in an acid solution comprising an acid.

[0154] In another embodiment of any of the above method(s), the acid solution may comprise an acidic acid or hydrochloric acid.

[0155] In another embodiment of any of the above method(s), the acid solution may include 50% acetic acid.

[0156] In another embodiment of any of the above method(s), the heating step may include heating for a period of up to 30 minutes or longer.

[0157] In another embodiment of any of the above method(s), the heating step may include heating for 30 minutes.

[0158] In another embodiment of any of the above method(s), the heating step may include boiling the acid solution.

[0159] In another embodiment of any of the above method(s), the acid solution can include 50% acetic acid, and the solution is heated to boiling for 30 minutes.

[0160] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid solution.

[0161] In another embodiment of any of the above method(s), the decellularization step may occur before the separation step.

[0162] In another embodiment of any of the above scaffold biomaterials, the decellularized microchannels are substantially adhesive-free, eg, physically bound together by entanglement.

[0163] In another embodiment of any of the above method(s), the bundling step may include gluing the microchannels together.

[0164] In yet another embodiment of any of the above method(s), the microchannels may be glued together by a biocompatible adhesive, optionally a biodegradable adhesive.

[0165] In yet another embodiment of any of the above method(s), the biocompatible adhesive may include a PEG-based, polyurethane-based, gelatin-based, or fibrin-based adhesive.

[0166] In another embodiment of any of the above method(s), the biocompatible adhesive may include a fibrin-based adhesive.

[0167] In yet another embodiment of any of the above method(s), the bundling step may include molding microchannels.

[0168] In yet another embodiment of any of the above method(s), the molding step may be performed using a mold in which the microchannels are filled.

[0169] In yet another embodiment of any of the above method(s), the molding may comprise biomedical grade silicone.

[0170] In another embodiment of any of the above method(s), the mold includes a passage for receiving the microchannel therein, the passage having a cross-sectional dimension given to the microchannel to be molded therein.

[0171] In yet another embodiment of any of the above method(s), the mold can be divided into two or more sections, which can be assembled around the microchannel for molding.

[0172] In another embodiment of any of the above method(s), adhesive can be added to the surface of the mold facing the microchannel, the microchannel can be filled into the mold, and the adhesive can be held in place while it cures.

[0173] In yet another embodiment of any of the above method(s), the mold may include a first section and a second section, and the molding step may include the steps of adding an adhesive to a surface of the first section facing the microchannel and to a surface of the second section facing the microchannel; placing the microchannel in contact with the adhesive in the first section and the second section; adding adhesive to the exposed surfaces of the microchannel in the first section, the microchannel in the second section, or both; placing the second section having the microchannel on the exposed surface of the microchannel in the first section opposite the first section, thereby assembling the mold; allowing the adhesive to substantially cure; and removing the mold.

[0174] In another embodiment of any of the above method(s), the method may further comprise cutting the bundled microchannels to a desired length.

[0175] In another embodiment of any of the above method(s), the method can further comprise cutting the bundled microchannels to a desired length, wherein the cutting step The tapping may be performed before the mold is removed.

[0176] In another embodiment of any of the above method(s), the method may further comprise the step of sterilizing the microchannel.

[0177] In yet another embodiment of any of the above method(s), the step of sterilizing may include exposing the microchannel to ethanol or a mixture of ethanol and water.

[0178] In yet another embodiment of any of the above method or methods, the microchannels may comprise xylem and / or phloem channels.

[0179] In yet another embodiment of any of the above method(s), the decellularization microchannels can be cellulose-based, chitin-based, lignin-based, lignocellulosic polymer-based, hemicellulose-based, or pectin-based, or any combination thereof.

[0180] In another embodiment of any of the above method(s), the density of decellularized microchannels within the bundle may be greater than the density of microchannels within the plant or fungal tissue.

[0181] In another embodiment of any of the above method or methods, the plant or fungal tissue is selected from the group consisting of apple thixopod (Apple) tissue, fern (Pteridophyta) tissue, turnip (Brassica rapa) root tissue, ginkgo biloba branch tissue, horsetail (Equisetum aestivum) tissue, daylily hybrid leaf tissue, kale (Brassica oleracea) stem tissue, conifer Douglas fir (Douglas fir) tissue, cactus fruit (Pitaya) flesh tissue, and Vinca maculata. tissue, aquatic lotus (Nelumbo nucifera) tissue, tulip (Tulipa gesneriana) petal tissue, plantain (Banana) tissue, broccoli (Brassica oleracea) stem tissue, maple leaf (Acer platanoides) stem tissue, beet (Beta vulgaris) primary root tissue, spring onion (Allium sativum) tissue, orchid (Orchidaceae) tissue, turnip (Brassica rapa) stem tissue, leek (Allium ampeloplastum) tissue, maple (Acer spp.) The microchannel-containing tissue may include tree branch tissue, celery (Apium graveolens) tissue, leek (Allium sativum) stem tissue, pine tissue, Aloe vera tissue, watermelon (Citrus lanatus var. lanatus) tissue, creeping jenny (Solanum kobainko) tissue, cactus tissue, Lychnis alpina tissue, rhubarb (Rheum labarbarum) tissue, pumpkin flesh (Cucurbita pepo) tissue, Dracaena (Asparagaceae) stem tissue, Tradescantia recutita (Tradescantia recutita) stem tissue, asparagus (Asparagus officinalis) stem tissue, mushroom (fungus) tissue, fennel (Fennel) tissue, rose (Rosa) tissue, carrot (Daucus carota) tissue, or pear (Pome fruit) tissue, or genetically modified tissue produced by direct genome modification or by selective breeding, or any combination thereof.

[0182] In yet another embodiment of any of the above method(s), the plant or fungal tissue may include celery, asparagus, or both.

[0183] In another embodiment of any of the above method(s), decellularizing the microchannel may include decellularizing by heat shock, treatment with a detergent, osmotic shock, lyophilization, physical lysis, electrical disruption, or enzymatic digestion, or any combination thereof, thereby removing cellular material and nucleic acids to provide a decellularized microchannel.

[0184] In another embodiment of any of the above method(s), the decellularization step may include treatment with sodium dodecyl sulfate (SDS).

[0185] In yet another embodiment of any of the above method(s), residual SDS may be removed by precipitating the salt residue containing SDS micelles from the microchannel using an aqueous divalent salt solution.

[0186] In another embodiment of any of the above method(s), treatment with dH2O, DI water, acetic acid, DMSO, or sonication, or any combination thereof, may be used to remove divalent salt solutions, salt residues, and / or SDS micelles.

[0187] In another embodiment of any of the above method(s), the divalent salt of the aqueous divalent salt solution may include MgCl or CaCl.

[0188] In another embodiment of any of the above method(s), the decellularization step can include treatment with an SDS solution of about 0.1% or about 1% SDS in water, and after decellularization, residual SDS can be removed by using an aqueous CaCl solution at a concentration of about 100 mM, followed by incubation in dH2O or DI water.

[0189] In yet another embodiment of any of the above method(s), the method may further comprise the step of introducing living plant or animal cells into the microchannels.

[0190] In another embodiment of any of the above method(s), the living cell may be a mammalian cell.

[0191] In another embodiment of any of the above method(s), the living cells may be human cells.

[0192] In another embodiment, provided herein is a decellularized microchannel produced by any of the method(s) described herein.

[0193] In another embodiment, provided herein is a scaffold biomaterial produced by any of the method(s) described herein.

[0194] In another embodiment of the above scaffold biomaterial, the scaffold biomaterial may be or may include a scaffold biomaterial as described herein.

[0195] In another embodiment, a mold: glue, one or more microchannels; one or more decellularizing agents; Scalpel or microtome, Sterile measuring devices, sterile saline, Tweezers and / or Instructions for carrying out any of the method(s) as described herein Provided herein are kits comprising any one, two, three, four or more of: [Brief explanation of the drawings]

[0196] These and other features will be better understood with regard to the following description and accompanying drawings. [Figure 1]Figure 1 shows (left): Isolated asparagus (AS) vascular bundle, approximately 3 cm long; (right): Microscopic cross section of an AS vascular bundle with the outline of the xylem channels clearly visible against the background. [Figure 2] Figure 2 shows (left): isolated celery (CL) vascular bundles, variable lengths 5-10 cm; (right): microscopic cross-sections of CL vascular bundles, with the outlines of xylem channels clearly visible against the ground tissue background. [Figure 3] Figure 3 (Left): Biomedical grade silicone mold (5 mm internal diameter, 1 cm length) for vascular bundle filling. The mold halves are pressed together. (Right): 2 mm thick section of the mold containing the adhered CL vascular bundle. [Figure 4] Figure 4 shows the AS channel bundle during the adhesive curing step. The mould halves are pressed together. [Figure 5] Figure 5 shows the CL channel bundle during the adhesive curing step. The mold halves are pressed together. [Figure 6] FIG. 6 shows (left): HDMC (AS) bundles stored in DPBS, (right): individual HDMC (AS) immediately after removal from the silicone mold. [Figure 7] FIG. 7 shows (left): HDMC(CL) bundles stored in DPBS, (right): individual HDMC(CL) immediately after removal from the silicone mold. [Figure 8] Figure 8 shows confocal laser scanning micrographs of HDMC (AS-based) samples after 1 week of GFP-3T3 cell culture followed by fixation in 4% paraformaldehyde. Visible cells (green - green fluorescent protein) in and around the xylem microchannels (blue - autofluorescence). (Left): Cross-section of a single CL vascular channel within an adhered bundle; cells are visible within the xylem channel. Note the cell density around the openings of the xylem and phloem channels. (Right): Cross-section (partial) of two vascular channels within an adhered bundle; cells were scattered throughout. [Figure 9]Figure 9 shows confocal laser scanning micrographs of HDMC (CL-based) samples after 1 week of GFP-3T3 cell culture followed by fixation in 4% paraformaldehyde. Visible cells (green - green fluorescent protein) are in and around the xylem microchannels (blue - autofluorescence). (Left): Cross-section of a single CL vascular channel within an adhered bundle; cells are visible within the xylem channel. Note that cell attachment to the fibrin glue deposits was localized to the right of the vascular channel. (Right): Cross-section of two vascular channels within an adhered bundle; cells were scattered throughout. [Figure 10] FIG. 10 shows a quarter AS section with visible vascular bundles pointed out by red arrows. [Figure 11] FIG. 11 shows an isolated AS vascular bundle in a weigh boat containing DPBS. [Figure 12] FIG. 1 shows an isolated CL vascular bundle in a weighing dish containing DPBS. [Figure 13] FIG. 10 shows AS HDMC flux during adhesive curing using a PEG-based adhesive. [Figure 14] Figure 1 shows AS HDMC in DPBS storage solution glued with a PEG-based adhesive. The VB was loosely bound and the HDMC was unable to maintain a cylindrical shape under the experimental conditions of the type of mold used. [Figure 15] Figure 1 shows CL HDMC bundles during adhesive curing, where volume expansion of the PEG-based Coseal adhesive caused separation of the silicone mold halves. [Figure 16] Figure 1 shows CL HDMC bundles adhered with PEG-based Coseal in DPBS storage solution, showing partial detachment of individual VBs. [Figure 17] FIG. 1 shows a side view of a bundle of AS strips bonded with a cyanoacrylate-based adhesive. [Figure 18] FIG. 1 shows the emitting surface of a glued (cyanoacrylate adhesive) AS strip bundle, with the individual channels faintly visible. [Figure 19]FIG. 1 shows AS channels in silicone mold halves bonded together during the curing period using biodegradable fibrin sealant (TISSEEL). [Figure 20] FIG. 10 shows the AS channel in the silicone mold during the glue curing step using fibrin glue. [Figure 21] FIG. 1 shows the appearance of the fibrin-adhered AS channel after removal from the silicone mold. [Figure 22] FIG. 1 shows a perspective view of an AS sample fixed with 4% paraformaldehyde after 1 week of GFP-3T3 cell culture (left panel) and a top view of the same sample (right panel). [Figure 23] CLSM image of an AS HDMC bundle sample fixed with 4% paraformaldehyde after 1 week of GFP-3T3 cell culture, showing cells (green) in and around the xylem microchannels (blue). [Figure 24] FIG. 1 shows the appearance of the AS channel in the food-grade silicone straw mold during the adhesive (fibrin) curing step. [Figure 25] FIG. 1 shows the appearance of adhered (fibrin) AS channel samples stored in sterile DPBS after cutting into 5 mm lengths. [Figure 26] FIG. 10 shows a side view of a CL channel bundle glued with cyanoacrylate adhesive. [Figure 27] Figure 1 shows the emitting surface of a CL channel bundle glued with cyanoacrylate adhesive, with individual channels faintly visible. [Figure 28] FIG. 10 shows CL channels in silicone mold halves bonded together using fibrin sealant during the curing period. [Figure 29] FIG. 10 shows the appearance of the CL channel in the silicone mold during the adhesive curing step using fibrin sealant. [Figure 30] FIG. 1 shows the appearance of fibrin-adhered CL channel bundles after removal from the silicone mold. [Figure 31]FIG. 1 shows a perspective view of a CL bundle sample fixed with 4% paraformaldehyde after 1 week of GFP-3T3 cell culture (left panel) and a top view of the same sample (right panel). [Figure 32] CLSM images of a CL bundle sample fixed with 4% paraformaldehyde after 1 week of GFP-3T3 cell culture, showing cells (green) within and around the xylem microchannels (blue). [Figure 33] FIG. 10 shows the appearance of the CL channel in the food-grade silicone mold during the adhesive (fibrin) curing step. [Figure 34] FIG. 10 shows the appearance of the glued CL channel after the top silicone mold half has been removed. [Figure 35] FIG. 1 shows the appearance of adhered CL channel samples that were cut into 5 mm lengths and then stored in sterile Dulbecco's Phosphate Buffered Saline (DPBS). [Figure 36] A) Confocal laser scanning fluorescence micrograph (Z-stack projection) of xylem channels and fibroblasts within HDMC / AS. The channel is visualized in red (autofluorescence; false color), and GFP-3T3 fibroblasts are visualized in green (green fluorescent protein; false color). B) A magnified view of the multichannel image of the red outline in panel A. C) A 3D volume rendering (Fiji / ImageJ; 3D viewer plugin) of the green fluorescent protein and xylem autofluorescence signals from the perspective of panel B. D) A rotational view of the 3D rendering showing the orientation of cells along the length of the channel. [Figure 37]A) Confocal laser scanning fluorescence micrograph (Z-stack projection) of xylem channels and fibroblasts in HDMC / CL. The channel is visualized in red (autofluorescence; false color), and GFP-3T3 fibroblasts are visualized in green (green fluorescent protein; false color). White arrows indicate cells outside the channel. Blue arrows indicate cells inside the channel. B) Zoom-in of one channel outlined in red in panel A. C) 3D volume rendering (Fiji / ImageJ; 3D viewer plugin) of the green fluorescent protein signal from the perspective of panel B. D) Rotational view of the 3D rendering showing cell orientation along the length of the channel. [Figure 38] Figure 1 shows histological staining of celery-derived HDMC bundles in the subcutaneous injection study of Example 8. Longitudinal section. Hematoxylin and eosin (A, B). [Figure 39] Figure 1 shows histological staining of HDMC bundles from celery in the subcutaneous injection study of Example 8. Transverse section. Hematoxylin and eosin (A, B). [Figure 40] Figure 1 shows histological staining of asparagus-derived HDMC bundles in the subcutaneous injection study of Example 8. Longitudinal section. Hematoxylin and eosin (A, B). [Figure 41] Figure 1 shows histological staining of asparagus-derived HDMC bundles in the subcutaneous injection study of Example 8. Transverse section. Hematoxylin and eosin (A, B). [Figure 42] FIG. 1 shows the percentage of bundles separated into individual VBs after treatment with various ratios of acetic acid:30% hydrogen peroxide and as a function of boiling time. [Figure 43] Decellularized celery placed on a clear plastic plate after boiling for 20 minutes in various concentrations of acid and peroxide solutions, showing the ratio of glacial acetic acid to 30% hydrogen peroxide (Figure 43A-3:1; Figure 43B-2:1; Figure 43C-1:1; Figure 43D-1:2; Figure 43E-1:3). [Figure 44]Figures 44A and 44B show asparagus strips treated with an acid and peroxide solution of 1:1 glacial acetic acid:30% hydrogen peroxide. Figure 44A shows asparagus strips freshly cut into slices in the acid / peroxide solution. Figure 44B shows the resulting extracted strips after removal from the acid / peroxide solution and washing with distilled water. Figure 44C shows a small portion of the strips that was placed in a subsequent water wash. [Figure 45] FIG. 1 shows the percentage of LiCl-treated flux separated into individual VBs as a function of salt concentration and boiling time. [Figure 46] FIG. 1 shows the percentage of NaCl-treated flux segregated into individual VBs as a function of salt concentration and boiling time. [Figure 47] FIG. 1 shows the percentage of NaOH-treated flux separated into individual VBs as a function of NaOH concentration and boiling time. [Figure 48] Figure 48A shows the size distribution of CL vascular bundles obtained from native tissue immersed in boiling LiCl saline for 15 minutes. Similar microstructural distributions were observed with other treatments. Figure 48A: Vascular bundles were stained with 0.1% calcofluor white, and images were thresholded and segmented for size analysis. Figure 48B: Histogram of vascular bundle diameter. The average size was 23.7 ± 1.5 μm. Values ​​are the mean and standard error of the mean. [Figure 49] Fluorescence microscopy images of CL vascular bundle (VB) ultrastructure preserved after boiling in 0.5 M LiCl for 10, 15, 20, and 30 minutes. Materials were stained with calcofluor white for 10 minutes, and images were converted to grayscale. [Figure 50] Fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 1 M LiCl for 10, 15, 20, and 30 minutes. Materials were stained with calcofluor white for 10 minutes, and images were converted to grayscale. [Figure 51]Fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 2 M LiCl for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 52] Fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 3 M LiCl for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 53] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 0.5 M NaCl for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 54] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 1 M NaCl for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 55] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 2 M NaCl for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 56] FIG. 1 shows fluorescence microscopy images of CL vascular bundle (VB) ultrastructure preserved after boiling in 3 M NaCl for 10, 15, 20, and 30 minutes. [Figure 57] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 0.5 M NaOH for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 58] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 1 M NaOH for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 59]Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 2 M NaOH for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 60] Fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in 3 M NaOH for 10, 15, 20, and 30 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 61] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in a 1:1 (v / v) ratio of peroxide and acetic acid for 10, 15, and 20 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 62] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in a 1:2 (v / v) ratio of peroxide and acetic acid for 10, 15, and 20 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 63] Figure 1 shows fluorescence microscopy images of CL vascular bundle (VB) microstructure preserved after boiling in a 2:1 (v / v) ratio of peroxide and acetic acid for 10, 15, and 20 minutes. The material was stained with calcofluor white for 10 minutes, and the image was converted to grayscale. [Figure 64] FIG. 1 shows HDMCs assembled from isolated CL vascular bundles boiled in water for 30 min (control condition). [Figure 65] FIG. 1 shows HDMCs assembled from NaOH-treated CL vascular bundles that were boiled for 30 minutes. [Figure 66] FIG. 1 shows CL macerated in dilute acetic acid and peroxide by boiling for 5 minutes as described in Example 10; and [Figure 67] FIG. 1 shows macerated celery with only the xylem and phloem fibers visible. DETAILED DESCRIPTION OF THE INVENTION

[0197] There has been a lack of materials (biomaterials) that can mimic the directional, channel-like microscopic structures of certain tissues in humans, plants, and animals. Examples of such directional tissues include the extracellular matrix (ECM) of the spinal cord, vascular channels, lymphatic tissue, nervous tissue, and many others. Biocompatible materials with directional and / or channel-like structures are desirable in the art.

[0198] Decellularized plant tissues can be used to provide scaffold biomaterials. Plant tissues may contain microchannels, such as xylem and phloem channels, which can provide some directionality to the scaffold biomaterial. However, in decellularized plant tissues, the number of functional structural elements (microchannels) may be limited by their natural density within the plant tissue. For example, material performance with respect to functional recovery after SCI may be limited by the number of microchannels available for neuronal growth in a given application. We have demonstrated that these channels can be efficiently integrated into the plant tissue. A method has now been developed for densifying to provide high density microchannel (HDMC) bundles, which can be used in a wide variety of applications, including the treatment and / or repair of spinal cord injury (SCI) and many others.

[0199] As described herein, in certain embodiments, microchannels present in the vascular bundles of plant tissues (i.e., xylem and phloem) can provide a guided scaffold for cell (e.g., neuron) growth and / or regeneration. Such decellularized plant structures can provide a more suitable surface for cell growth (in terms of attachment, biocompatibility, etc.) compared to certain artificial or synthetic substrates. It is shown herein that by extracting, isolating, and refilling these channels, these guided structural components can be densified and provide more pathways for cell growth. In certain embodiments, it is contemplated that densified bundles of microchannels can provide superior performance in certain applications compared to decellularized plant material without further modification resulting from increased microchannel densification. In certain instances where densification is beneficial, it is contemplated that unmodified decellularized material may be less effective as a scaffold per unit volume compared to densified vascular tissue.

[0200] In certain embodiments, the biomaterials described herein can be derived from cell wall architecture and / or vasculature found in the plant and fungal kingdoms to create 3D scaffolds that can promote and / or directionally guide cell infiltration, cell proliferation, vascularization, tissue repair, and / or tissue remodeling. As will be appreciated, biomaterials as described herein can be produced from any suitable part of a plant or fungal organism that has vasculature or microchannels (e.g., xylem and / or phloem, etc.), including, for example, seeds, roots, bark, leaves, stems, fruits, pulp, and cores. Biomaterials can include substances such as, for example, cellulose, chitin, lignin, lignocellulosic polymers, hemicellulose, pectin, and / or any other suitable biochemicals / biopolymers naturally found in these organisms.

[0201] In certain embodiments, unlike many commercially available biomaterials, plant / fungal-derived biomaterials such as those described herein may be substantially non-resorbable or poorly resorbable (i.e., they are not substantially degraded or absorbed by the body), although in certain embodiments, in embodiments in which adhesives are used, the adhesives may be biodegradable. The non-resorbable nature of these scaffolds may offer certain benefits. For example, in certain embodiments, the biomaterials described herein may be resistant to shape change and / or may retain their intended geometry over extended periods of time. In certain embodiments, because they have a minimal footprint compared to certain other products, they may be considered effectively invisible to the body and unlikely to elicit an immune response. As absorbable biomaterials degrade, their by-products often elicit harmful immune responses as well as induce oxidative stress and raise the pH in healing tissues, which can be avoided by using non-resorbable biomaterials.

[0202] It will be understood that unless otherwise stated, the meanings / definitions of the kingdoms Plantae and Fungi used herein are based on the Cavalier-Smith classification (1998).

[0203] Scaffold biomaterials Described herein are scaffold biomaterials, methods and uses thereof, and methods for producing same. It will be appreciated that the embodiments and examples are provided for illustrative purposes, directed to those skilled in the art, and are not meant to be limiting in any way.

[0204] In one embodiment, a scaffold biomaterial comprising at least one bundle of microchannels. Provided herein is a scaffold biomaterial, wherein a bundle of microchannels comprises a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle.

[0205] As will be appreciated, a scaffold biomaterial, in certain embodiments, can be a material having a three-dimensional structure (although in certain embodiments, it can be extremely thin in all of the x, y, and z dimensions), and the arrangement of decellularized channels provides directional porosity determined by the size, density, and orientation of the microchannels. In certain embodiments, a scaffold biomaterial can be suitable for providing a scaffold upon which living cells can infiltrate and / or grow and / or proliferate.

[0206] In certain embodiments, microchannels can generally comprise any suitable vascular or other channel-type structure derived from or isolated from plant or fungal tissue. In certain embodiments, microchannels can comprise, for example, xylem and / or phloem channels. In certain embodiments, microchannels can be decellularized such that cellular material and nucleic acids of the plant or fungal tissue are removed from the microchannels. In certain embodiments, microchannels can be individually isolated from plant or fungal tissue, grouped into one or more vascular bundles isolated from the plant or fungal tissue, or any combination thereof. In certain embodiments, microchannels can be cellulose-based, chitin-based, lignin-based, hemicellulose-based, lignocellulosic polymer-based, or pectin-based, or any combination thereof.

[0207] In certain embodiments, the microchannels may be about 50 to about 150 microns in diameter, or any subrange or value occurring therebetween, and have a continuous length determined based on the size of the biomaterial source or by cutting one or more microchannels to a desired length. In certain embodiments, a mix of microchannels of different diameters and / or lengths may be used, or the microchannels may be sorted / selected so that microchannels of similar diameters and / or lengths are used.

[0208] As will be understood, the cellular material and nucleic acid of the plant or fungal tissue may include intracellular contents, such as cellular organelles (e.g., chloroplasts, mitochondria), nuclei, cellular nucleic acids, and / or cellular proteins, which may be substantially removed, partially removed, or completely removed from the plant or fungal tissue, from the microchannel, and / or from the scaffold biomaterial. It will be recognized that trace amounts of such components may still be present in the decellularized plant or fungal tissue described herein. It will also be understood that references herein to decellularized microchannels will reflect that such cellular material found in the plant or fungal source of the microchannel has been substantially removed from the microchannel—although this does not exclude the possibility that the decellularized microchannels, in certain embodiments, may generally contain or comprise subsequently introduced or reintroduced cells, cellular material, and / or nucleic acids, e.g., animal or human cells, of any type.

[0209] Indeed, in certain embodiments, the decellularized microchannels and / or scaffold biomaterials as described herein may further comprise living cells on and / or within at least one of the decellularized microchannels. In certain embodiments, the living cells may be animal, mammalian, or human cells. In certain embodiments, the living cells may comprise plant and / or fungal cells.

[0210] In certain embodiments, plant and / or fungal cells can be added to the microchannels to further modify the channels, after which decellularization can be performed, e.g., prior to implantation. It is contemplated that decellularization processes may be performed. By way of example, in certain embodiments, the decellularization process may denature and / or remove native cell wall proteins from the scaffold, and selected and / or engineered moieties, such as polysaccharide proteins (e.g., polysaccharides conjugated to mammalian surface anchorages and ligand-binding domains), may be added to the walls of the decellularized scaffold. In certain embodiments, anchorage proteins may be added. In certain embodiments, cells that may be added / deposited onto the decellularized scaffold may include those that can produce useful or functional proteins or macromolecules of interest in the microchannel structures. In certain embodiments, proteins with mammalian surface anchorage points may be added to the scaffold, for example, to accelerate integration into surrounding tissue. In certain embodiments, cells may be genetically modified to produce such moieties or proteins, or the cells may be added to the scaffold for a period of time and then removed, washed off, or killed. Examples of proteins or moieties that may be added include mammalian growth factors for specific cell types (e.g., to promote neuronal growth).

[0211] In certain embodiments, a microchannel bundle may comprise any suitable grouping or cluster of decellularized microchannels. In the examples described herein below, the bundles were formed using adhesives. However, it will be understood that in certain embodiments, bundles may alternatively be formed using a variety of other adhesive-free techniques, including, but not limited to, techniques using sealants, mechanical packing or interlocking, or, for example, chemical crosslinking of adjacent isolated vasculature. In certain embodiments, bundles may be formed by suspending microchannels in a cellulose-based hydrogel or other such hydrogel. In certain embodiments, individual microchannels and / or vascular bundles may be combined together, for example, by threading around each other. In certain embodiments, it is contemplated that compaction / packing of isolated (possibly decellularized) vasculature into a larger covering and / or fixed vasculature (possibly decellularized) may be performed.

[0212] In certain embodiments of any of the above methods, the bundling step may include physically binding or physically cross-linking the microchannels and / or VBs without the use of adhesives. Such a process may be referred to as entanglement of the microchannels. The microchannels may be physically bound or otherwise encased such that separation of the microchannels and / or VBs requires the use of force. Different dimensions and lengths (Kuhn segments or persistence lengths) may be used to create different entanglement and alignment profiles. Entanglement may result from agitation using a stirring device such as a rod. Entanglement may occur during liquid-based extraction.

[0213] As described in detail herein, in certain embodiments, isolated decellularized plant or fungal vasculature (i.e., xylem, phloem, and / or vascular bundles thereof) can be bundled (and optionally glued) together to provide channel densification beyond that found in plant or fungal source materials. In certain embodiments, these bundles can be engineered to be biocompatible and may be suitable for further functionalization. In certain embodiments, it is contemplated that such high-density microchannel (HDMC) bundles can be used in a wide variety of applications, which may include, but are not limited to, use as extracellular matrices, cell scaffolds, and specialized structural supports.

[0214] As will be appreciated, in certain embodiments, the decellularization microchannels of a bundle may be arranged substantially parallel to one another within the bundle to provide a substantially common longitudinal direction or alignment of the channels of the bundle. Figures 4 and 5, described in more detail below, depict examples of bundles with decellularization microchannels arranged substantially parallel to one another (i.e., all microchannels in the left bundle of Figure 4 are arranged substantially parallel to one another, substantially Oriented in the same direction (left-right), all microchannels in the right bundle of Figure 4 are arranged substantially parallel to each other and oriented in substantially the same direction (front-back), and in the case where the decellularized microchannels are bundled by entanglement, substantially parallel may refer to a common direction or alignment of the bundle.

[0215] In certain embodiments, multiple decellularized microchannels of a bundle of scaffold biomaterials can be glued together within the bundle. In certain embodiments, multiple decellularized microchannels can be glued together within the bundle with a biocompatible adhesive, optionally a biodegradable adhesive. While it is typically desirable for the adhesive to be biodegradable so that it degrades over a period of time after implantation of the scaffold biomaterial, embodiments in which the adhesive is not biodegradable are also contemplated herein. In certain embodiments, the adhesive can be an adhesive that has good adhesion under moist conditions. In certain embodiments, the biocompatible adhesive can include a PEG-based, polyurethane-based, gelatin-based, or fibrin-based adhesive. In certain embodiments, the biocompatible adhesive can include a fibrin-based adhesive.

[0216] In the examples described below, fibrin glue has been shown to be particularly effective for bonding HDMC bundles and scaffold biomaterials. However, it will be understood that various other adhesives and / or sealants are also contemplated. For example, it is contemplated that in certain embodiments, fibrin glue may be substituted with other adhesive / sealant types, such as synthetic adhesives. Such adhesives may be advantageous in certain instances, for example, due to a less pronounced immune response after implantation.

[0217] PEG-based adhesives with additives can be used to allow for their biodegradation, but the inventors have found that the swelling of PEG-based adhesives makes them less desirable compared to fibrin adhesives. In general, in certain embodiments, a wide variety of suitable adhesive systems can be used in the fabrication of HDMC bundles, with the understanding that biodegradability of the adhesive, while typically desirable, is not required. In certain embodiments, it is contemplated that, for example, PEG-, polyurethane-, or gelatin-based adhesives or sealants can be used. In certain embodiments, the adhesive can be selected to be both biocompatible and degradable, for example, after implantation.

[0218] In certain embodiments, the adhesive may be a fibrin glue or a cyanoacrylate glue. In certain embodiments, the adhesive may include a fibrin sealant. In certain embodiments, the adhesive may include TISSEEL (available from Baxter), Evicel (available from Ethicon), or Vitagel (available from Stryker). In certain embodiments, the adhesive may include a PEG sealant. In certain embodiments, the adhesive may include Coseal (available from Baxter) or Duraseal (available from Covidien). Examples of adhesives are also described in Vyas et al., "Comparison of Hemostatic Agents Used in Vascular Surgery," Expert Opin Biol Ther., 2013, 13(12): 1663-1672, the entire text of which is incorporated herein by reference.

[0219] In embodiments where bonding, particularly molding, the HDMC bundle is desired, the adhesive may be selected to have low adhesion to the mold structure itself, low expansion properties as the adhesive sets / cures, or both, to prevent shattering of the microchannels. It is contemplated that in certain embodiments, a non-expansive adhesive or an adhesive with low expansion may be used. In certain embodiments, the adhesive may have a volume expansion of less than about 20%, less than about 19%, less than about 18%, less than about 17%, less than about 16%, less than about 15%, less than about 14%, less than about 13%, less than about 12%, less than about 11%, less than about 10%, less than about 9%, less than about 8%, less than about 7%, less than about 6%, less than about 5%, less than about 4%, less than about 3%, less than about 2%, less than about 1%, or substantially no volume expansion (i.e., 0%), or any range bounded by the upper end of any of these values ​​or any range between any two of these values, for example. An adhesive having a periphery can be selected.

[0220] In certain bonding embodiments, a shrinkable adhesive (or an adhesive that shrinks when dry) may be used. In such embodiments, the adhesive may typically be selected to provide little or minimal shrinkage to avoid damaging the bundle.

[0221] In certain embodiments, the adhesive may be biocompatible and / or bioinert. In certain embodiments, the adhesive may adhere well to lignocellulose, not adhere strongly to non-lignocellulosic materials, may be substantially non-immunogenic, may allow for sterile preparation, may not involve extensive preparation and / or processing steps prior to use, may set relatively quickly (e.g., within about 10 minutes), may have a low or no endotoxin load, may not penetrate lignocellulose so as not to block microchannels, may not degrade lignocellulose, may not alter its surface chemistry and / or affect or degrade its material properties, may not degrade or may degrade very slowly under storage conditions (e.g., 4°C, phosphate buffered saline), may not undergo autocatalytic chemical changes over time that significantly alter its properties, or any combination thereof.

[0222] In certain embodiments, an adhesive may be selected that is substantially biocompatible and substantially non-toxic. In certain embodiments, it may be desirable for the adhesive to be substantially bioinert and provoke little or no immune response. In certain embodiments, it may be desirable for the adhesive to be selected so that it is absorbable after cellular infiltration and / or implantation. In embodiments where molding is desired and a silicone-based mold is to be used, it may be desirable for the adhesive to be substantially non-sticky to silicone. In embodiments intended for therapeutic and / or in vivo applications, it may be desirable to use an adhesive approved by a regulatory agency, such as the FDA. Various surgical adhesives are commercially available, for example. A summary of surgical adhesives / sealants is provided in Yepremyan et al., Surgical Glue: A Brief Overview, Austin Journal of Biomedical Engineering, 2014, 1(4): 1020, and Annabi, et al., Elastic Sealants for Surgical Applications, European Journal of Pharmaceutics and Biopharmaceutics, 2015, http: / / dx.doi.org / 10.1016 / j.ejpb.2015.05.022. In certain embodiments, it may be desirable to select a synthetic adhesive over an adhesive of natural origin, as bioinertness is more prevalent among synthetic adhesives. While fibrin-based adhesives are shown to be particularly effective in the examples below, in embodiments where immune response susceptibility is particularly high, it may be desirable to use synthetic or other bioinert adhesives, for example, to avoid even a small immune response near the implantation site. Fibrin-based adhesives and synthetic adhesive materials are reviewed, for example, in Srinivasan et al., "Novel Synthetic Adhesive as an Effective Alternative to Fibrin-Based Adhesives," World Journal of Hepatology, 2017, 9(24): 1030-1039, the entire text of which is incorporated herein by reference.

[0223] Examples of adhesives of natural origin that may be considered include: Natural polymers (e.g., polysaccharides, polypeptides, etc.). Various types are known, including collagen / gelatin-based, chitosan-based, dextran-based, and alginate-based adhesives.

[0224] Gelatin-type adhesives are known for their biocompatibility and degradability, and an example of a gelatin-type adhesive approved for sale in Europe and with FDA fast track status is LifeSeal™, available from Life-Bond (www.life-bond.com / index.php / lifesealtm / ).

[0225] Fibrin-type adhesives, also known as fibrin sealants, are commonly used and are FDA-approved. There are many different types of fibrin adhesives, including, for example, adhesives based on bovine, porcine, and / or human thrombin.

[0226] Biomimetic adhesives are also contemplated. These types of adhesives are typically inspired by geckos, caddisflies, various worms, etc., and are known in the art, although most are still in the research stage.

[0227] Examples of synthetic adhesives that may be considered include: PEG-based adhesives: PEG-based adhesives are synthetic and generally considered to be biocompatible. Coseal (Baxter Inc.) is a widely available example of a PEG-based adhesive. While the inventors encountered issues with adhesive volume expansion and / or adhesion to silicone molds (if used) in initial testing with Coseal (not shown), it is believed that with appropriate adjustments, the effects of expansion and adhesion can be minimized. Polyurethane adhesives: Polyurethane adhesives are synthetic and generally considered biocompatible. These types are actively researched adhesives. Examples include VIVO (Adhesys Medical GmbH) (polyurethane-based) and / or TissuGlu (Cohera Medical Inc) (polyurethane-based, FDA-approved). Cyanoacrylates: Cyanoacrylates are considered synthetic but have toxic decomposition properties. Although these adhesives may be product-related and therefore somewhat limited in applicability, it is contemplated that such adhesives may still be useful where toxicity can be avoided, such as in topical use. Other adhesives: Examples of other synthetic adhesives that are contemplated include, for example, MeTro (Elastagen Ltd) - a recombinant tropoelastin-based and / or UV-curable adhesive.

[0228] In certain embodiments, for example, chemical cross-linking of microchannels with each other (simple cell cross-linking) can be used. Microtubules can be placed in an external enclosure, wrapping, or membrane by It is contemplated that by encasing the microchannels, gluing by sewing or tying the microchannels together, or any combination thereof, may be reduced or avoided entirely. In certain embodiments, decellularized microchannels may be interwoven or meshed together, which may, in certain embodiments, improve performance under tension. In certain embodiments, the channels may be flexible to allow such interwoven or meshed without significant damage. In certain embodiments, the interwoven may be performed during the molding step.

[0229] In certain embodiments, it is contemplated that the use of adhesives and / or sealants may be reduced or eliminated, for example, by molding processes that allow the individual pieces to fit tightly together.

[0230] In certain embodiments of the scaffold biomaterials described herein, the density of decellularized microchannels within the bundles may be higher than the density of microchannels within the plant or fungal tissue source. In certain embodiments, the density of decellularized microchannels may be higher than, for example, 1 cm 2 In certain embodiments, this may be based on a measurement of the number of channels per cm, e.g. 2 Based on the number of vascular bundles per cm 2 Based on the number of vasculature per cm 2 Density can be determined based on the number of individual xylem and / or phloem channels per 1 mm diameter. In certain embodiments, density can be measured in terms of the number of microchannels in a circular cross-sectional area of ​​a particular diameter. In certain preparations, samples have been measured at, for example, about 20 channels per 5 mm diameter circular cross-sectional area, although higher densities are also contemplated. .

[0231] In certain embodiments, it may be desirable to mimic axonal projections of the motor and / or sensory systems. In certain embodiments, it may be desirable to align the corticospinal tract, rubrospinal tract, raphe spinal tract, reticulospinal tract, propriospinal tract, spinothalamic tract, and / or spinal sensory axons. It is contemplated that HDMCs as described herein may be used in certain embodiments to mimic the 3D location of injured axons within a subject. In certain embodiments, HDMCs may comprise multiple subunits / channels rather than a single fixed graft. Furthermore, each SCI injury is usually unique and, in rare cases, affects the entire spinal cord. As such, the ability to specifically position HDMCs / channels in a 3D orientation that mimics an individual subject's specific injured axonal tract may be of particular interest in certain embodiments.

[0232] In certain embodiments, the plant or fungal tissue may generally include any suitable plant or fungal tissue or part containing at least some microchannels and / or vasculature (e.g., but not limited to, xylem and / or phloem channels) that may be isolated from surrounding plant or fungal tissue / structures.

[0233] In certain embodiments of the scaffold material(s) described above, the plant or fungal tissue is selected from the group consisting of apple thixosa (Apple) tissue, fern (Pteridophyta) tissue, turnip (Brassica rapa) root tissue, ginkgo biloba branch tissue, horsetail (Equisetum aestivum) tissue, daylily hybrid leaf tissue, kale (Brassica oleracea) stem tissue, conifer Douglas fir (Douglas fir) tissue, cactus fruit (Pitaya ) flesh tissue, Vinca maculata tissue, Aquatic lotus (Nelumbo nucifera) tissue, Tulip (Tulipa gesneriana) petal tissue, Plantain (Banana) tissue, Broccoli (Brassica oleracea) stem tissue, Maple leaf (Acer sycamore) stem tissue, Beet (Beta vulgaris) primary root tissue, Green onion (Allium sativum) tissue, Orchid (Orchidaceae) tissue, Turnip (Brassica rapa) stem tissue, leek (Allium ampeloplum) tissue, maple (Acer spp.) tree branch tissue, celery (Apium graveolens) tissue, leek (Allium sativum) stem tissue, pine tissue, aloe vera tissue, watermelon (Citrus lanatus var. lanatus) tissue, creeping jenny (Solanum kobainko) tissue, cactus tissue, Lychnis alpina tissue, rhubarb (Rheum labarbarum) tissue, pumpkin flesh (Cucurbita pepo) tissue, Dracaena (Asparagaceae) stem tissue, Tradescantia recutita (Tradescantia recutita) stem tissue, asparagus (Asparagus officinalis) stem tissue, mushroom (fungus) tissue, fennel (Foeniculum vulgare) tissue, rose (Rosa spp.) tissue, carrot (Daucus carota) tissue, or pear (Pome fruit) tissue. Further examples of plant and fungal tissues are described in Example 18 of WO 2017 / 136950, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," which is incorporated herein by reference in its entirety.

[0234] In certain embodiments, plant or fungal tissues can be genetically modified, either by direct genomic modification or by selective breeding, to create additional plant or fungal constructs that can be configured to physically mimic the tissue and / or functionally promote a target tissue effect. One of skill in the art, given the teachings herein, will be able to select a suitable scaffold biomaterial to suit a particular application.

[0235] In certain embodiments, the polymeric material may be characterized, for example, based on physical characteristics that can be measured and tailored to a particular application, such as size, structure (porous / tubular), hardness, strength, hardness, and / or ductility. Based on this, an appropriate tissue can be selected for a particular application.

[0236] Additionally, chemical properties, such as reactivity, etc., can be considered to select the material to suit a particular application. Coordination number, enthalpy of formation, heat of combustion, stability, toxicity and / or type of bonding may also be considered. Such characteristics (physical and chemical) may also affect the decellularization and and / or may be directly modified before or after functionalization.

[0237] In certain embodiments, microchannels may be sourced from the same tissue or part of a plant or fungus, or from different parts or tissues of a plant or fungus. In certain embodiments, microchannels may be sourced from the same individual plant or fungus, or from multiple plants or fungi of the same species. In certain embodiments, microchannels may be sourced from multiple plants or fungi of different species, such that a bundle contains vasculature from more than one species. In certain embodiments, microchannels may be selected to provide a particular mechanism. For example, in certain embodiments, microchannels having diameters falling within a particular size range may be selected for inclusion in a bundle to impart particular structural characteristics to the scaffold biomaterial.

[0238] In certain embodiments, the plant or fungal tissue may include celery, asparagus, or both.

[0239] In certain embodiments, the scaffold biomaterial can be a scaffold biomaterial configured to physically mimic a tissue of a subject and / or functionally promote a target tissue effect in a subject. Methods of using such scaffold biomaterials described herein can, in certain embodiments, include selecting a scaffold biomaterial as described herein in which the decellularized microchannels are configured to physically mimic a tissue of a subject and / or functionally promote a target tissue effect in a subject. One of skill in the art will be able to select a suitable scaffold biomaterial to suit a particular application, given the teachings herein.

[0240] Various methods can be used to generate scaffold biomaterials as described herein. By way of example, in certain embodiments of the above-described scaffold biomaterials, the decellularized microchannels can include microchannels that have been decellularized by heat shock, treatment with detergents (e.g., SDS, Triton X, EDA, alkyline treatment, acid, ionic surfactants, nonionic surfactants, and zwitterionic surfactants), osmotic shock, lyophilization, physical lysis (e.g., hydrostatic pressure, critical point decellularization, CO2 critical point decellularization), electrical disruption (e.g., non-thermal irreversible electroporation), or enzymatic digestion, or any combination thereof. In certain embodiments, biomaterials as described herein can be obtained from plants and / or fungi by utilizing a decellularization process that can involve any of several approaches (either individually or in combination), including, but not limited to, heat shock (e.g., rapid freeze-thawing), chemical treatment (e.g., detergents), osmotic shock (e.g., distilled water), freeze-drying, physical lysis (e.g., pressure treatment), electrical disruption, and / or enzymatic digestion.

[0241] In certain embodiments, microchannels will typically be isolated from the plant tissue first and then decellularized. In such a method, decellularization may be easier, as the microchannels are typically similar in size and can be decellularized at similar rates. However, it will be understood that in certain embodiments, the plant tissue may be decellularized first, and then the microchannels may be isolated from the decellularized plant tissue. In such an approach, the decellularization conditions may be adjusted accordingly to match the characteristics of the plant tissue being decellularized. (For example, in certain embodiments, larger samples of plant tissue may be subjected to longer or more invasive decellularization protocols compared to smaller samples of plant tissue that are more easily accessible to decellularization reagents.)

[0242] In certain embodiments, a decellularized microchannel may comprise a microchannel that has been decellularized by treatment with a detergent or surfactant, including, but not limited to, sodium dodecyl sulfate (SDS), Triton X, EDA, alkaline treatment, acid, ionic detergents, non-ionic detergents, and zwitterionic detergents.

[0243] In yet further embodiments, the decellularized microchannel may include a microchannel that has been decellularized by treatment with SDS. In yet another embodiment, residual SDS may be removed from the microchannel by washing with an aqueous divalent salt solution. The aqueous divalent salt solution may be used to precipitate / disrupt salt residues containing SDS micelles from the solution / scaffold, and dH2O, acetic acid, or dimethyl sulfoxide (DMSO) treatment or sonication may be used to remove salt residues or SDS micelles. In certain embodiments, the divalent salt of the aqueous divalent salt solution may include, for example, MgCl2 or CaCl2.

[0244] In another embodiment, the microchannels can be decellularized by treatment with a 0.01-10%, e.g., between about 0.1% and about 1%, SDS solution in a solvent, e.g., water, ethanol, or another suitable organic solvent, or, e.g., about 0.1% SDS or about 1% SDS, and residual SDS may be removed by using an aqueous CaCl solution at a concentration of about 100 mM, followed by incubation in dH2O. In certain embodiments, the SDS solution may be at a concentration greater than 0.1%, which may facilitate decellularization and may be accompanied by increased washing to remove residual SDS. In certain embodiments, the microchannels can be decellularized by treatment with an SDS solution of about 0.1% SDS in water, and residual SDS may be removed by using an aqueous CaCl solution at a concentration of about 100 mM, followed by incubation in dH2O.

[0245] Further examples of decellularization protocols that can be adapted to generate decellularized microchannels for scaffold biomaterials as described herein can be found in WO 2017 / 136950, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," the entire text of which is incorporated herein by reference.

[0246] In certain embodiments, the decellularized microchannels and / or scaffold biomaterials as described herein may further comprise living cells on and / or within at least one of the decellularized microchannels. In certain embodiments, the living cells may be animal cells, mammalian cells, or human cells. In certain embodiments, the cells may be cells introduced or seeded into and / or onto the scaffold biomaterial and / or decellularized microchannels, or cells that infiltrate into or onto the scaffold biomaterial and / or decellularized microchannels after implantation of the scaffold biomaterial and / or decellularized microchannels into, for example, a living animal or plant subject.

[0247] Methods and uses of scaffold biomaterials Scaffold biomaterials as described herein may include one or more high-density microchannel bundles. Many of the design considerations and contemplated uses of the scaffold biomaterials described herein may be related to those described for the scaffold biomaterials in International Publication No. WO 2017 / 136950, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," the entire text of which is incorporated herein by reference, although the biomaterials described herein may benefit from increased channel density and / or controllable channel arrangement, which may be substantially parallel to one another. Thus, the biomaterials described herein may be particularly advantageous for applications where, for example, channel directionality is desirable.

[0248] In certain embodiments, biomaterials as described herein may have applications, for example, in biomedical laboratory research and / or clinical regenerative medicine in human and / or veterinary applications. Such biomaterials may be useful as scaffolds that can be used as research tools for industrial / academic biomedical researchers in biomedical implants, in sensing devices and drug delivery vehicles, and / or in other suitable applications in which scaffolds may be used.

[0249] In certain embodiments, scaffold biomaterials as described herein can be used for the regeneration of blood vasculature. The wide range of available structures can allow for the artificial fabrication of vessel-like structures and / or can provide suitable conditions for angiogenesis (natural blood vessel formation).

[0250] In certain embodiments, scaffold biomaterials as described herein can be used as simple or complex tissues. For example, scaffolds can be used to replace / regenerate simple (skin, bone) or complex (spinal cord, muscle, nerves, blood vessels, etc.) tissues after accident, deformity, aesthetic, injury, or other damage to the tissue.

[0251] In certain embodiments, a method of providing a structural scaffold for supporting animal cell growth, promoting tissue regeneration, promoting vascularization, tissue replacement, or in cosmetic surgery in a subject in need thereof, comprising: providing a scaffold biomaterial according to any of the scaffold biomaterials described herein; Implanting the scaffold biomaterial into a subject. Provided herein is a method comprising:

[0252] In certain embodiments, the high-density microchannel (HDMC) approach described herein can provide a directional pathway for cell infiltration. "Directivity" can arise not only from the interior of the microchannel, but also from its outer surface and the inter-channel space. Figures 36 and 37 show evidence of cell growth along both the inside and outside of the microchannel within the HDMC. Thus, HDMCs may be of particular interest in tissue repair applications where cellular connections or ligation are severed. Directional biomaterials may be particularly desirable in such cases, for example. Because neurons in the spinal cord are highly organized in tracts, and their proper function relies on directional connections, it is contemplated that HDMCs may be of particular interest for spinal cord injury (SCI) treatment, for example. In certain embodiments, it is contemplated that HDMCs can provide an end-to-end support structure that can provide neuronal reconnection along appropriate tracts. Other structures that may be of particular interest for repair with HDMCs as described herein may include vasculature channels. In certain embodiments, it is contemplated that HDMC may act as joints and / or connectors for repairing particularly fine vasculature.

[0253] In certain embodiments, bidirectional microchannels provide nano- and / or micro-topographical cueing that can drive neuron and / or axon guidance. (Straley KS, Foo CWP, Heilshorn, incorporated herein by reference) SC (2010) Biomaterial design strategies for the treatment of spinal cord injuries. J Neurotrauma 27:1-19; Hoffman-Kim D, Mitchell J a, Bellamkonda RV (2010) Topography, cell response, and nerve regeneration. Annu Rev Biomed Eng 12:203-31; See Hoffman-Kim D, Mitchel J a, Bellamkonda RV (2010) Topography, cell response, and nerve regeneration. Annu Rev Biomed Eng 12:203-31.

[0254] In certain embodiments, HDMCs may allow individual channels to be aligned with the subject's own damaged spinal tracts and / or provide a better biomimetic of the lost architecture of the damaged tissue (see also Ige, OO, Umoru, LE, & Aribo, S. (2012). Natural Products: A Minefield of Biomaterials. ISRN Materials Science, 2012, 1-20. https: / / doi.org / 10.5402 / 2012 / 983062, incorporated herein by reference). Rostral axons may migrate through the scar tissue to bypass the lesion and reconnect with caudal axons. However, because the 3D coordinates of the lesion may be lost due to random axonal migration through the dense fibrin fibrils of the formed scar tissue, it is unlikely that the exact axon tract reconnected to the same rostral stump. The extent of recovery that can be achieved from axonal reconnection may be due to the natural plasticity of the central nervous system established through long-term physical therapy (Anderson, MA, O'Shea, TM, Burda, J., incorporated herein by reference). E., Ao, Y., Barlatey, SL, Bernstein, AM, ... Sofroniew, MV (2018). Required growth facilitators propel axon regeneration across complete spinal cord injury. Nature, 561(7723), 396-400. See also https: / / doi.org / 10.1038 / s41586-018-0467-6. The HDMC approach described herein may provide Thus, if the 3D coordinates of the entire damaged tissue are maintained, in certain embodiments, it is contemplated that the ability to accurately route axons rostral and caudal stump pathways may be re-established, which may increase the subject's motor recovery and reduce reliance on CNS plasticity during the patient's physical therapy.

[0255] In certain embodiments, a scaffold biomaterial may be implanted into the spinal cord and may promote repair or regeneration after spinal cord injury, may provide a structural implant for tissue replacement and / or tissue regeneration in a subject, may provide a structural implant for skin grafting and / or skin regeneration in a subject, may provide a structural implant for regeneration of blood vasculature in a target tissue or area or subject, may provide a bone replacement, bone filler or bone graft material, and / or may promote bone regeneration in a subject, may provide tissue replacement of skin, bone, muscle, nerves, vasculature or other damaged or malformed tissue in a subject, and / or may provide a structural implant for cosmetic surgery.

[0256] In certain embodiments, the scaffold biomaterial may be implanted in the spinal cord and may promote repair and / or regeneration following acute and / or chronic spinal cord injury in the central and / or peripheral nervous system.

[0257] In one embodiment, provided herein is the use of any of the scaffold biomaterials and / or microchannel bundles as described herein for supporting animal or plant cell growth, promoting tissue regeneration, promoting vascularization, treating and / or repairing spinal cord injury, repairing or reconstructing or replacing plant or animal tissue, filtering and / or separating solutions or materials, modifying or regulating growth of plants, material transport, fabrication to mimic desired shapes or objects, and / or microfluidics, or any suitable application where biocompatible micrometer-scale channels may be useful.

[0258] In certain embodiments, the scaffold biomaterial may be for the repair or reconstruction or replacement of plant or animal tissue, including damaged microvasculature, or plant tissue damaged by vascular disease (such as wilt disease), or damaged vasculature in trees infested by insects (such as emerald ash borers).

[0259] In certain embodiments, the scaffold biomaterial is a biomaterial for the repair or reconstruction of plant or animal tissue. The plant or animal tissue may contain a damaged microvasculature, and angiogenesis is impaired or impaired.

[0260] In another embodiment, the scaffold biomaterial may be for the repair or reconstruction of animal tissue, wherein the animal is a human.

[0261] In yet another embodiment, the scaffold biomaterial can be for plant modification or growth regulation, where the modification is modification of the implantation process for accelerated growth.

[0262] In yet another embodiment, the scaffold biomaterial can be for material transport, the material transport being for drug delivery applications.

[0263] In yet another embodiment, the scaffold biomaterial can be for heat transfer or heat exchange microfluidics.

[0264] In yet another embodiment, there is provided a method of supporting cell proliferation, comprising: Providing a scaffold biomaterial as described herein; introducing one or more cells into the scaffold biomaterial; Provided herein is a method comprising:

[0265] In yet another embodiment, there is provided a method for promoting tissue regeneration, promoting angiogenesis, treating and / or repairing spinal cord injury, or repairing or reconstructing or replacing plant or animal tissue, comprising: Providing a scaffold biomaterial as described herein; implanting the scaffold biomaterial at a site in need thereof; Provided herein is a method comprising:

[0266] In another embodiment, there is provided a method for the filtration and / or separation of a solution or material, comprising the steps of: Providing a scaffold biomaterial as described herein; passing a solution or material through the scaffold biomaterial to filter and / or separate components from the solution based on size exclusion; Provided herein is a method comprising:

[0267] In yet another embodiment, there is provided a method for modifying or regulating the growth of a plant, comprising: Providing a scaffold biomaterial as described herein; Implanting the scaffold biomaterial into a plant to provide accelerated growth. Provided herein is a method comprising:

[0268] In certain embodiments, transplantation can be performed surgically by identifying damaged or dysfunctional vasculature within the plant and replacing it with a scaffold biomaterial. In certain embodiments, such repair can prevent the loss of one or more distal portions of the plant. In certain embodiments, the scaffold biomaterial can be functionalized with one or more plant growth factors to promote or accelerate growth. In certain embodiments, the transplant can incorporate genetically modified cells obtained from the intended host or another host. Such cells can, for example, accelerate growth or confer resistance to certain plant diseases. In certain embodiments, plant R genes can be included; for example, the transplant can be used to incorporate cells with engineered R genes.

[0269] In another embodiment, there is provided a method for material transport, comprising: Providing a scaffold biomaterial as described herein; using the scaffold biomaterial to transport the material to the site where it is needed; Provided herein is a method comprising:

[0270] In another embodiment, the material can be or can include a drug, and the scaffold biomaterial can be used for drug delivery. In certain embodiments, the scaffold biomaterial can be loaded with a drug and administered to a subject at a designated site. In certain embodiments, the scaffold biomaterial can provide, for example, a controlled, slow release of a drug over time, thus avoiding subjecting the subject to multiple injections.

[0271] In certain embodiments, it is contemplated that scaffold biomaterials as described herein can be used for drug delivery to provide localization in drug delivery schemes. Vehicles such as alginate beads have been investigated as a means of encapsulating drugs and releasing them over time in a controlled manner. However, this type of delivery system can have problems delivering drugs to the designated location where they are needed, as the encapsulating material can be washed away. In certain embodiments, scaffold biomaterials as described herein can be designed to anchor the drug encapsulation system and remain in place so that the drug can be delivered to the intended target. Removal of the scaffold biomaterial can be used to stop drug delivery as needed. In certain embodiments, scaffold biomaterials as described herein can be used to anchor a drug encapsulation tool / vehicle, or can directly act as a drug encapsulation tool / vehicle, or both.

[0272] In another embodiment, there is provided a method for preparing a structure that mimics a desired shape or object, comprising the steps of: Providing a scaffold biomaterial as described herein; shaping or positioning the scaffold biomaterial to mimic a desired shape or object; Optionally, attaching a scaffold biomaterial for structural reinforcement. Provided herein is a method comprising:

[0273] In yet another embodiment, there is provided a method of exchanging or transferring heat in a microfluidics process, comprising: Providing a scaffold biomaterial as described herein; using microchannels to carry one or more fluids, the one or more fluids being in close proximity to allow for heat exchange or heat transfer; Provided herein is a method comprising:

[0274] In certain embodiments, one or more fluids may be conveyed in different microchannels by counter-flow or co-flow or a combination thereof.

[0275] In certain embodiments, the HDMC bundles and / or scaffold biomaterials as described herein are useful in a variety of applications, which may include, but are not limited to, any one or more of the following: treatment of spinal cord injury; repair / reconstruction of human, plant, or animal tissue (e.g., repair / reconstruction of damaged microvasculature, especially in cases where vascularization is not possible or has been impaired in any way, and / or in repairing plant tissue damaged by vascular disease, e.g., wilt disease, and / or to replace damaged vasculature in trees infested by insects such as emerald ash borers); solution / material filtration and / or separation; plant modification / growth regulation (e.g., modifying transplant processes for accelerated growth); material transport (e.g., drug delivery applications); fabricated to mimic a desired shape or object; and / or microfluidics (e.g., heat transfer / exchange); or any other suitable application in which biocompatible micrometer-scale channels may be useful. It is contemplated that the invention may be used in various ways.

[0276] Methods for isolating and decellularizing microchannels and preparing scaffold biomaterials:

[0277] Methods for isolating and decellularizing microchannels and for preparing scaffold biomaterials are described in detail herein, as well as experimental examples of such methods in the Examples section below.

[0278] Further examples of decellularization protocols that can be adapted to generate decellularized microchannels for scaffold biomaterials as described herein can be found in WO 2017 / 136950, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," the entire text of which is incorporated herein by reference.

[0279] In one embodiment, there is provided a method for isolating and decellularizing microchannels from plant or fungal tissue, comprising the steps of: isolating a microchannel from the plant or fungal tissue; decellularizing the microchannel; Optionally, sterilizing the microchannel. Provided herein is a method comprising:

[0280] It will be appreciated that in certain embodiments, the microchannels are typically separated from the plant or fungal tissue prior to decellularization, but it is also contemplated that in certain embodiments, the order may be switched such that decellularization may be performed before the microchannels are separated from the plant or fungal tissue.

[0281] In certain embodiments, decellularization may be performed prior to separating the microchannels from the plant or fungal tissue, hi certain embodiments, such approaches may benefit from performing a perfusion method to facilitate penetration of the decellularizing agent into all relevant areas of the plant or fungal tissue.

[0282] In certain embodiments, perfusion methods can be applied to decellularize plant or fungal tissue before channels are isolated, and perfusion can use substantially the same solutions and reagents as those already described herein for performing decellularization. A pathway for the decellularization solution to enter the plant or fungal tissue vasculature can be provided (e.g., using a needle tip). It is contemplated that such an approach may be particularly interesting for certain tissues, such as celery, which can easily wick fluids through its vasculature. In certain embodiments, such an approach can be used to primarily or solely decellularize the microchannel structure of plant or fungal tissue. For example, in certain embodiments, perfusion can be used for targeted decellularization (relatively rapid) or complete decellularization (longer time), for example, based on the use of multiple fluid injection sites.

[0283] In another embodiment of the above method, the step of isolating the microchannels may comprise mechanically separating the microchannels or the vascular bundles containing the microchannels, or both, from the surrounding plant or fungal tissue. In yet another embodiment, the step of separating may be performed by gentle peeling or by cutting.

[0284] In certain embodiments, the separation of one or more structures, e.g., microchannels, from plant or fungal tissue allows the separation of one or more structures of interest from the surrounding plant or fungal tissue. The separation step may involve one or more manual steps that may be performed to extract or separate the structure from the plant or fungal tissue. Such manual steps may involve cutting, slicing, peeling, and / or other physical separation techniques. As will be appreciated, for large-scale operations, such manual steps may be burdensome. As described herein, the inventors have therefore developed extraction techniques that may be less burdensome and / or easily amendable, for example, for scale-up. Thus, in certain embodiments, the separating step may involve a liquid-based extraction to isolate one or more structures from the plant or fungal tissue.

[0285] As will be understood, liquid-based extraction can include any suitable process for treating and extracting one or more structures from plant or fungal tissue (which can be either native plant or fungal tissue, or decellularized plant or fungal tissue, or a combination thereof) using a liquid extraction solution. In some cases, the plant or fungal tissue can be processed prior to liquid-based extraction, such as by dividing it into small pieces or strips with a suitable device, for example, a scalpel or mandoline.

[0286] Liquid-based extraction will be understood by those skilled in the art as a process for chemically treating and extracting vascular bundles (VBs) and / or microchannels from native or decellularized plant or fungal tissues using a liquid solution. Liquid-based extraction can be understood as the maceration of native or decellularized plant or fungal tissue to obtain intact substructures, such as microchannels. Liquid-based extraction can result in the isolation of a single microchannel or a bundle of microchannels. When native plant or fungal tissues are used, the liquid-based extraction conditions can result in partial or complete decellularization of the native tissue. In such cases, the tissue can be further treated using any of the decellularization methods discussed herein.

[0287] In the embodiments described herein, the liquid solution can be an acid solution, an acid and peroxide solution, a salt solution, or an alkaline solution for acid extraction, acid and peroxide extraction, salt extraction, or alkaline extraction, respectively. In some cases, two or more treatments or solutions are used simultaneously or sequentially.

[0288] One or more solutions may be combined with the plant or fungal tissue and heated to a desired temperature, e.g., boiling. In some cases, the solution is heated or boiled for a period of time, e.g., 0.1 to 30 minutes. Periods longer than 30 minutes, e.g., up to 1 hour or longer, are also contemplated. Any amount of time within the range or any value therebetween (which may be rounded to the nearest 0.1), or any subrange spanning any two of these values, is contemplated.

[0289] In another embodiment of any of the above method(s), acid extraction can include mixing the plant or fungal tissue in an acid solution containing an acid. The mixture of plant or fungal tissue and acid solution is then heated. As will be appreciated, the acid solution can generally include any suitable acid capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable acids include, but are not limited to, acetic acid, boric acid, carbonic acid, hydrochloric acid, citric acid, hydrofluoric acid, nitric acid, oxalic acid, phosphoric acid, sulfuric acid, boron trifluoride, oxalic acid, malonic acid, succinic acid, malic acid, and the like. Other categories of acids are contemplated, such as carboxylic acids, including, but not limited to, linear saturated dicarboxylic acids, branched dicarboxylic acids, unsaturated dicarboxylic acids, substituted dicarboxylic acids, and aromatic dicarboxylic acids. The acid may be dissolved / mixed in a suitable solvent to form an acid solution. Typically, the solvent may include water, but other solvents or combinations of solvents may also be used, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, ethanol, ethanol, ethanol, ammonium hydroxide ... Also contemplated are acetone, dimethylformamide, dimethylsulfoxide, and amphiphilic solvents or colloids. By way of example, in certain embodiments, the acid solution may comprise an aqueous solution of 50% acetic acid.

[0290] In another embodiment of any of the above method(s), acid and peroxide extraction can include mixing the plant or fungal tissue in an acid and peroxide solution containing an acid and peroxide. The mixture of plant or fungal tissue and acid and peroxide solution is then heated. As will be appreciated, the acid / peroxide solution can generally include any suitable acid and peroxide capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable acids include, but are not limited to, acetic acid, boric acid, carbonic acid, hydrochloric acid, citric acid, hydrofluoric acid, nitric acid, oxalic acid, phosphoric acid, sulfuric acid, boron trifluoride, oxalic acid, malonic acid, succinic acid, malic acid, etc. Other categories of acids are contemplated, including, but not limited to, linear saturated dicarboxylic acids, branched dicarboxylic acids, unsaturated dicarboxylic acids, substituted dicarboxylic acids, and aromatic dicarboxylic acids. Examples of suitable peroxides include, but are not limited to, hydrogen peroxide, lithium peroxide, barium peroxide, dibenzoyl peroxide, benzoyl peroxide, and methyl ethyl ketone peroxide. Peroxides can form peroxyacids when combined with acids such as peracetic acid. Acid and peroxide solutions can include glacial acetic acid and 30% hydrogen peroxide in ratios of 5:1 to 1:5, e.g., 3:1 (containing 13.05 M acetic acid and 2.45 M hydrogen peroxide) to 1:3 (containing 4.35 M acetic acid and 7.35 M hydrogen peroxide). The acid and peroxide can be dissolved / mixed in a suitable solvent to form an acid and peroxide solution. Typically, the solvent may comprise water, although other solvents or solvent combinations are contemplated, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethylsulfoxide, and amphiphilic solvents or colloids. By way of example, in certain embodiments, the acid / peroxide solution may comprise an aqueous solution of 1:1 glacial acetic acid:30% hydrogen peroxide (v / v).

[0291] In another embodiment of any of the above method(s), salt extraction can include mixing the plant or fungal tissue in a salt solution. The mixture of plant or fungal tissue and salt solution is then heated. As will be appreciated, the salt solution can generally include any suitable salt capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable salts can be monovalent, e.g., LiCl and NaCl, divalent, e.g., MgSO4 and CaCl2, trivalent, e.g., AlCl3, etc. Suitable cations include, but are not limited to, lithium, sodium, potassium, magnesium, calcium, iron, copper, zinc, aluminum, and ammonium. Suitable anions include, but are not limited to, chloride, bromide, acetate, carbonate, citrate, fluoride, nitrate, phosphate, sulfate, iodide, and borate. Pharmaceutical salts, such as ibuprofenate, are also contemplated. The selection of a suitable salt can also depend on desired properties, such as ionic activity, screening, coordination size, Debye length, or the desired ionic strength. The salt solution can have a suitable salt concentration, for example, about 0.5 M to 10 M, or any value therebetween (which may be rounded to the nearest 0.1), or any subrange spanning any two of these values. The salt can be dissolved / mixed in a suitable solvent to form the salt solution. Typically, the solvent can include water, but other solvents or solvent combinations, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethyl sulfoxide, and amphiphilic solvents or colloids, are also contemplated. By way of example, in certain embodiments, the salt solution may comprise an aqueous solution of NaCl or LiCl having a salt concentration of about 0.5M to 3M.

[0292] In another embodiment of any of the above method(s), alkaline extraction can include mixing the plant or fungal tissue in an alkaline solution. In some cases, the mixture of plant or fungal tissue and alkaline solution is heated. As will be appreciated, the alkaline solution can generally include any suitable base capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable bases include, but are not limited to, sodium hydroxide (NaOH), potassium hydroxide (KOH), carbonic acid, nitric acid, phosphoric acid, sulfuric acid, ammonia, calcium hydroxide, magnesium hydroxide, lithium hydroxide, zinc hydroxide, sodium carbonate, sodium bicarbonate, butyllithium, sodium azide, sodium amide, sodium hydride, sodium borohydride, and lithium diisopropylamine. The alkaline solution can have a suitable alkaline concentration, for example, about 0.5 to 10 M, or any value therebetween (which may be rounded to the nearest 0.1), or any subrange spanning any two of these values. The base can be dissolved / mixed in a suitable solvent to form the alkaline solution. Typically, the solvent can include water, but other solvents or solvent combinations, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethyl sulfoxide, and amphiphilic solvents or colloids, are also contemplated. By way of example, in certain embodiments, the alkaline solution can include an aqueous solution of NaOH having a base concentration of about 0.5 M to 1 M.

[0293] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the alkaline solution. The solution may be stirred using a stir bar or other suitable method, e.g., a mechanical or magnetic stirrer.

[0294] Methods for liquid-based extraction can be tailored to produce VB and / or microchannels of desired morphology and / or ultimate yield strength. Methods can also be tailored to isolate VB and / or microchannels from desired plants or fungi. For example, longer heating or boiling times can result in softened microchannels and / or VB (see Tables 1-8). Softened VB and / or microchannels may be desirable in applications where easily moldable microchannels are desired. Tailoring the mechanical properties of isolated microchannels and / or VB can lead to directing cell growth and differentiation and controlling the mechanical properties of the resulting engineered tissue. Hardened VB and / or microchannels may be desirable for applications where strength is desired, such as structures for cell growth. Higher concentrations of acid, base, salt (e.g., greater than 10 M) and acid / peroxide (e.g., greater than 5:1 to 1:5) and shorter boiling times are contemplated.

[0295] In yet another embodiment, there is provided a method for preparing a scaffold biomaterial, comprising the steps of: isolating the microchannel from the plant or fungal tissue; decellularizing the microchannel; bundling the microchannels together so that the microchannels are positioned substantially parallel to one another; Thereby, provided herein is a method comprising the steps of providing a scaffold biomaterial comprising bundled microchannels.

[0296] It will be appreciated that in certain embodiments, the microchannels are typically separated from the plant or fungal tissue prior to decellularization, however, it is contemplated that in certain embodiments, the order may be switched such that decellularization may be performed before the microchannels are separated from the plant or fungal tissue.

[0297] In certain embodiments of the above methods, isolating the microchannels may include mechanically separating the microchannels or the vascular bundles containing the microchannels, or both, from the surrounding plant or fungal tissue. In certain embodiments, the separating step may be performed by gentle peeling or by cutting.

[0298] In certain embodiments, separating one or more structures, e.g., microchannels, from plant or fungal tissue may include one or more manual steps that may be performed to extract or separate one or more structures of interest from the surrounding plant or fungal tissue. Such manual steps may involve cutting, slicing, peeling, and / or other physical separation techniques. As will be appreciated, for large-scale operations, such manual steps may be burdensome. As described herein, the inventors have therefore developed extraction techniques that may be less burdensome and / or easily amendable, for example, for scale-up. Thus, in certain embodiments, the separating step may include a liquid-based extraction to isolate one or more structures from the plant or fungal tissue.

[0299] As will be understood, liquid-based extraction can include any suitable process for treating and extracting one or more structures from plant or fungal tissue (which can be either native plant or fungal tissue, or decellularized plant or fungal tissue, or a combination thereof) using a liquid extraction solution. In some cases, the plant or fungal tissue can be processed prior to liquid-based extraction, such as by dividing it into small pieces or strips with a suitable device, for example, a scalpel or mandoline.

[0300] Liquid-based extraction will be understood by those skilled in the art as a process for chemically treating and extracting vascular bundles (VBs) and / or microchannels from native or decellularized plant or fungal tissues using a liquid solution. Liquid-based extraction can be understood as the maceration of native or decellularized plant or fungal tissue to obtain intact substructures, such as microchannels. Liquid-based extraction can result in the isolation of a single microchannel or a bundle of microchannels. When native plant or fungal tissues are used, the liquid-based extraction conditions can result in partial or complete decellularization of the native tissue. In such cases, the tissue can be further treated using any of the decellularization methods discussed herein.

[0301] In the embodiments described herein, the liquid solution can be an acid solution, an acid and peroxide solution, a salt solution, or an alkaline solution for acid extraction, acid and peroxide extraction, salt extraction, or alkaline extraction, respectively. In some cases, two or more treatments or solutions are used simultaneously or sequentially.

[0302] One or more solutions may be combined with the plant or fungal tissue and heated to a desired temperature, e.g., boiling. In some cases, the solution is heated or boiled for a period of time, e.g., 0.1 to 30 minutes. Periods longer than 30 minutes, e.g., up to 1 hour or longer, are also contemplated. Any amount of time within the range or any value therebetween (which may be rounded to the nearest 0.1), or any subrange spanning any two of these values, is contemplated.

[0303] In another embodiment of any of the above method(s), acid extraction may include mixing the plant or fungal tissue in an acid solution comprising an acid. The mixture of plant or fungal tissue and acid solution is then heated. As will be appreciated, the acid solution may generally comprise any suitable acid capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable acids include: Examples of suitable acids include, but are not limited to, acetic acid, boric acid, carbonic acid, hydrochloric acid, citric acid, hydrofluoric acid, nitric acid, oxalic acid, phosphoric acid, sulfuric acid, boron trifluoride, malonic acid, succinic acid, malic acid, etc. Other categories of acids are contemplated, including, but not limited to, linear saturated dicarboxylic acids, branched dicarboxylic acids, unsaturated dicarboxylic acids, substituted dicarboxylic acids, and aromatic dicarboxylic acids. The acid may be dissolved / mixed in a suitable solvent to form an acid solution. Typically, the solvent may include water, but other solvents or solvent combinations, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethyl sulfoxide, and amphiphilic solvents or colloids, are also contemplated. By way of example, in certain embodiments, the acid solution may include an aqueous solution of 50% acetic acid.

[0304] In another embodiment of any of the above method(s), acid and peroxide extraction can include mixing the plant or fungal tissue in an acid and peroxide solution containing an acid and peroxide. The mixture of plant or fungal tissue and acid and peroxide solution is then heated. As will be appreciated, the acid / peroxide solution can generally include any suitable acid and peroxide capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable acids include, but are not limited to, acetic acid, boric acid, carbonic acid, hydrochloric acid, citric acid, hydrofluoric acid, nitric acid, oxalic acid, phosphoric acid, sulfuric acid, boron trifluoride, oxalic acid, malonic acid, succinic acid, malic acid, etc. Other categories of acids are contemplated, including, but not limited to, linear saturated dicarboxylic acids, branched dicarboxylic acids, unsaturated dicarboxylic acids, substituted dicarboxylic acids, and aromatic dicarboxylic acids. Examples of suitable peroxides include, but are not limited to, hydrogen peroxide, lithium peroxide, barium peroxide, dibenzoyl peroxide, benzoyl peroxide, and methyl ethyl ketone peroxide. Peroxides can form peroxyacids when combined with acids such as peracetic acid. Acid and peroxide solutions can include glacial acetic acid and 30% hydrogen peroxide in ratios of 5:1 to 1:5, e.g., 3:1 (containing 13.05 M acetic acid and 2.45 M hydrogen peroxide) to 1:3 (containing 4.35 M acetic acid and 7.35 M hydrogen peroxide). The acid and peroxide can be dissolved / mixed in a suitable solvent to form an acid and peroxide solution. Typically, the solvent may comprise water, although other solvents or solvent combinations are contemplated, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethylsulfoxide, and amphiphilic solvents or colloids. By way of example, in certain embodiments, the acid / peroxide solution may comprise an aqueous solution of 1:1 glacial acetic acid:30% hydrogen peroxide (v / v).

[0305] In another embodiment of any of the above method(s), salt extraction can include mixing the plant or fungal tissue in a salt solution. The mixture of plant or fungal tissue and salt solution is then heated. As will be appreciated, the salt solution can generally include any suitable salt capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant / fungal tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable salts can be monovalent, e.g., LiCl and NaCl, divalent, e.g., MgSO4 and CaCl2, trivalent, e.g., AlCl3, etc. Suitable cations include, but are not limited to, lithium, sodium, potassium, magnesium, calcium, iron, copper, zinc, aluminum, and ammonium. Suitable anions include, but are not limited to, chloride, bromide, acetate, carbonate, citrate, fluoride, nitrate, phosphate, sulfate, iodide, and borate. Pharmaceutical salts, such as ibuprofenate, are also contemplated. The selection of a suitable salt may also depend on the desired properties, such as ionic activity, screening, coordination size, Debye length, or desired ionic strength. The salt solution may have a suitable salt concentration, for example, about 0.5 M to 10 M, or any value therebetween (which may be rounded to the nearest 0.1), or any value between these values. The range may include any subrange spanning any two of the above. The salt may be dissolved / mixed in a suitable solvent to form a salt solution. Typically, the solvent may include water, but other solvents or solvent combinations are also contemplated, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethyl sulfoxide, and amphiphilic solvents or colloids. By way of example, in certain embodiments, the salt solution may include an aqueous solution of NaCl or LiCl having a salt concentration of about 0.5M to 3M.

[0306] In another embodiment of any of the above method(s), alkaline extraction can include mixing the plant or fungal tissue in an alkaline solution. In some cases, the mixture of plant or fungal tissue and alkaline solution is heated. As will be appreciated, the alkaline solution can generally include any suitable base capable of osmotic shock of the plant / fungal tissue, decomposition of the outer plant tissue structure, disruption of hydrogen bonds within the plant / fungal tissue, and / or disruption of the plant / fungal polymer crystalline structure to extract tissue components, such as VBs and / or microchannels. Examples of suitable bases include, but are not limited to, sodium hydroxide (NaOH), potassium hydroxide (KOH), carbonic acid, nitric acid, phosphoric acid, sulfuric acid, ammonia, calcium hydroxide, magnesium hydroxide, lithium hydroxide, zinc hydroxide, sodium carbonate, sodium bicarbonate, butyllithium, sodium azide, sodium amide, sodium hydride, sodium borohydride, and lithium diisopropylamine. The alkaline solution can have a suitable alkaline concentration, for example, about 0.5 to 10 M, or any value therebetween (which may be rounded to the nearest 0.1), or any subrange spanning any two of these values. The base can be dissolved / mixed in a suitable solvent to form the alkaline solution. Typically, the solvent can include water, but other solvents or solvent combinations, such as, but not limited to, propanol, ethanol, methanol, ammonia, acetic acid, acetone, dimethylformamide, dimethyl sulfoxide, and amphiphilic solvents or colloids, are also contemplated. By way of example, in certain embodiments, the alkaline solution can include an aqueous solution of NaOH having a base concentration of about 0.5 M to 1 M.

[0307] In another embodiment of any of the above method(s), the liquid-based extraction may further comprise mechanically agitating, e.g., stirring, the plant or fungal tissue in the alkaline solution. The solution may be stirred using a stir bar or other suitable method, e.g., a mechanical or magnetic stirrer.

[0308] Methods for liquid-based extraction can be tailored to produce VB and / or microchannels of desired morphology and / or ultimate yield strength. Methods can also be tailored to isolate VB and / or microchannels from desired plants or fungi. For example, longer heating or boiling times can result in softened microchannels and / or VB (see Tables 1-8). Softened VB and / or microchannels may be desirable in applications where easily moldable microchannels are desired. Tailoring the mechanical properties of isolated microchannels and / or VB can lead to directing cell growth and differentiation and controlling the mechanical properties of the resulting engineered tissue. Hardened VB and / or microchannels may be desirable for applications where strength is desired, such as structures for cell growth. Higher concentrations of acid, base, salt (e.g., greater than 10 M) and acid / peroxide (e.g., greater than 5:1 to 1:5) and shorter boiling times are contemplated.

[0309] In certain embodiments, the liquid-based extraction may result in substantial sterilization (concurrent sterilization). In other cases, the isolated microchannel and / or VB is sterilized after using any of the sterilization methods described herein.

[0310] In certain embodiments, HDMC bundles can be formed from multiple vascular bundles (including xylem and phloem channels) obtained from a plant. and / or may be further processed to isolate phloem channels, which can then be used to form HDMC bundles that may have even higher densities. As will be appreciated, higher densities can be achieved by removing more of the non-vascular plant tissue surrounding the microchannels. Non-vascular tissue may generally refer to parenchyma and thick-walled tissue. In certain embodiments, it is contemplated that channel density can also be increased, for example, by selecting thinner-walled or smaller microchannels, by cross-sectional area (which may be plant species dependent and may be selected for the desired application, e.g., so as not to be smaller than the cells to be accommodated), and / or by reducing the adhesive used in the inter-channel spaces, for example, by modifications to the molding / adhesion steps (usually macroscale, mechanical considerations).

[0311] In certain embodiments of any of the above methods, the bundling step can include gluing the microchannels together. In certain embodiments, the microchannels can be glued together with a biocompatible adhesive, optionally a biodegradable adhesive. In certain embodiments, the biocompatible adhesive can include a PEG-based, polyurethane-based, gelatin-based, or fibrin-based adhesive. In certain embodiments, the biocompatible adhesive can include a fibrin-based adhesive.

[0312] In certain embodiments of any of the above methods, the bundling step may include molding the microchannel. In certain embodiments, the molding step may be performed using a mold into which the microchannel is filled. In certain embodiments, the mold may include biomedical grade silicone. In certain embodiments, the mold may include polypropylene plastic or Teflon or another non-stick material or coating. In certain embodiments, the mold may include a non-stick material to which the adhesive does not or only weakly adheres, may be biocompatible, may be disposable, may be autoclavable, may be moldable or somewhat flexible, or any combination thereof. In certain embodiments, the mold may include a passage for receiving the microchannel therein, the passage having a cross-sectional dimension that imparts to the microchannel to be molded therein. In certain embodiments, the mold may be divided into two or more sections, which can be assembled around the microchannel for molding. In certain embodiments, adhesive may be added to the surface of the mold facing the microchannel, and the microchannel is filled into the mold and held therein while the adhesive cures. In certain embodiments, the mold includes a first section and a second section, and the molding step includes the steps of adding an adhesive to a surface of the first section facing the microchannel and to a surface of the second section facing the microchannel; placing the microchannel in contact with the adhesive in the first section and the second section (the microchannel may be divided between the first and second sections); adding adhesive to the exposed surfaces of the microchannel in the first section, the microchannel in the second section, or both; placing the second section having an associated microchannel on the exposed surface of the microchannel associated with the first section opposite the first section, thereby assembling the mold; allowing the adhesive to substantially harden; and removing the mold.

[0313] In certain embodiments of any of the above methods, the bundling step may include physically binding or physically cross-linking the microchannels and / or VBs without the use of adhesives. Such a process may be referred to as entanglement of the microchannels. The microchannels may be physically bound or otherwise entangled such that separation of the microchannels and / or VBs requires the use of force. Different entanglement and alignment profiles may be created by the use of different dimensions and different lengths (Kuhn segments or persistence lengths). Entanglement may also be achieved by agitation using an agitating device such as a rod. Entanglement can occur during liquid-based extraction.

[0314] In certain embodiments of any of the above methods, the method may further include cutting the bundled microchannels to a desired length, hi certain embodiments, cutting the bundled microchannels to a desired length may be performed before the mold is removed.

[0315] In yet another embodiment of any of the above methods, the method may further include the step of sterilizing the microchannel. In another embodiment, the step of sterilizing may include exposing the microchannel to ethanol or a mixture of ethanol and water.

[0316] In certain embodiments of any of the above methods, the microchannels may comprise xylem and / or phloem channels.

[0317] In another embodiment of any of the above methods, the decellularization microchannels can be cellulose-based, chitin-based, lignin-based, hemicellulose-based, or pectin-based, or any combination thereof.

[0318] In yet another embodiment of any of the above method(s), the density of decellularized microchannels within the bundle may be greater than the density of microchannels within the plant or fungal tissue.

[0319] In yet another embodiment of any of the above method or methods, the plant or fungal tissue is selected from the group consisting of apple thixopod (Apple) tissue, fern (Pteridophyta) tissue, turnip (Brassica rapa) root tissue, ginkgo biloba branch tissue, horsetail (Equisetum aestivum) tissue, daylily hybrid leaf tissue, kale (Brassica oleracea) stem tissue, conifer Douglas fir (Douglas fir) tissue, cactus fruit (Pitaya) flesh tissue, and kale (Brassica oleracea) stem tissue. Vinca tissue, water lotus (Nelumbo nucifera) tissue, tulip (Tulipa gesneriana) petal tissue, plantain (Banana) tissue, broccoli (Brassica oleracea) stem tissue, maple leaf (Acer platanoides) stem tissue, beet (Beta vulgaris) primary root tissue, spring onion (Allium sativum) tissue, orchid (Orchidaceae) tissue, turnip (Brassica rapa) stem tissue, leek (Allium ampeloplastum) tissue, maple (Acer spp.) ) tree branch tissue, celery (Apium graveolens) tissue, leek (Allium sativum) stem tissue, pine tissue, aloe vera tissue, watermelon (Citrullus lanatus var. lanatus) tissue, creeping jenny (Solanum kobanko) tissue, cactus tissue, Lychnis alpina tissue, rhubarb (Rheum labarum) tissue, pumpkin pulp (Cucurbita pepo) tissue, Dracaena (Asparagaceae) stem tissue, Tradescantia recutita (L.) The microchannel-containing tissue may include microchannel-containing tissue derived from tradescantia (Tradescantia recutita) stem tissue, asparagus (Asparagus officinalis) stem tissue, mushroom (fungal) tissue, fennel (Fennel) tissue, rose (Rosa) tissue, carrot (Daucus carota) tissue, or pear (Pome fruit) tissue, or genetically modified tissue produced by direct genome modification or by selective breeding, or any combination thereof. In another embodiment, the plant or fungal tissue may include celery, asparagus, or both. Further examples of plant and fungal tissues are described in Example 18 of International Publication No. WO 2017 / 136950, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," the entire contents of which are incorporated herein by reference.

[0320] Examples of decellularization protocols that can be adapted to generate decellularized microchannels for scaffold biomaterials as described herein are described in International Publication No. 2017 / 111444, entitled "Decellularized Cell Wall Structures from Plants and Fungus and Use Thereof as Scaffold Materials," which is incorporated herein by reference in its entirety. It can be found in brochure no. 36950.

[0321] Various methods can be used for decellularization. By way of example, in certain embodiments, decellularization can include decellularization by heat shock, treatment with detergents (e.g., SDS, Triton X, EDA, alkaline treatment, acid, ionic detergents, non-ionic detergents, and zwitterionic detergents), osmotic shock, lyophilization, physical lysis (e.g., hydrostatic pressure), electrical disruption (e.g., non-thermal irreversible electroporation), or enzymatic digestion, or any combination thereof. In certain embodiments, the decellularization process can include any of several approaches (either individually or in combination), including, but not limited to, heat shock (e.g., rapid freeze-thawing), chemical treatment (e.g., detergents), osmotic shock (e.g., distilled water), lyophilization, physical lysis (e.g., pressure treatment), electrical disruption, and / or enzymatic digestion.

[0322] In certain embodiments, decellularization may involve treatment with a detergent or surfactant, including, but not limited to, sodium dodecyl sulfate (SDS), Triton X, EDA, alkaline treatment, acid, ionic detergents, non-ionic detergents, and zwitterionic detergents.

[0323] In yet further embodiments, the decellularized microchannel may include a microchannel that has been decellularized by treatment with SDS. In yet another embodiment, residual SDS may be removed from the microchannel by washing with an aqueous divalent salt solution. The aqueous divalent salt solution may be used to precipitate / disrupt salt residues containing SDS micelles from the solution / scaffold; dH2O, acetic acid, or dimethyl sulfoxide (DMSO) treatment or sonication may also be used to remove salt residues or SDS micelles. In certain embodiments, the divalent salt of the aqueous divalent salt solution may include, for example, MgCl2 or CaCl2.

[0324] In another embodiment, the microchannels can be decellularized by treatment with a 0.01-10%, e.g., between about 0.1% and about 1%, SDS solution, or e.g., about 0.1% SDS or about 1% SDS, in a solvent, e.g., water, ethanol, or another suitable organic solvent, and residual SDS may be removed by using an aqueous CaCl solution at a concentration of about 100 mM, followed by incubation in dH2O. In certain embodiments, the SDS solution may be at a concentration greater than 0.1%, which may facilitate decellularization and may be accompanied by increased washing to remove residual SDS. In certain embodiments, the microchannels can be decellularized by treatment with an SDS solution of about 0.1% SDS in water, and residual SDS may be removed by using an aqueous CaCl solution at a concentration of about 100 mM, followed by incubation in dH2O.

[0325] In another embodiment, any of the above method(s) may further comprise introducing living plant or animal cells into the microchannels. In another embodiment, any of the above method(s) may further comprise culturing living plant or animal cells on and / or within the scaffold biomaterial. In one embodiment, the living cells may comprise mammalian cells, e.g., human cells.

[0326] In certain embodiments, the microchannels and / or scaffold biomaterials may be functionalized with specific growth factors selected for a particular cell type of interest, which may in certain embodiments include, for example, providing growth factors attached to the scaffold.

[0327] In certain embodiments, particularly for spinal cord injury applications, the invention provides a method for promoting repair and / or recovery. It is contemplated that patient-derived neural progenitor cells may be added to a scaffold as described herein to achieve this.

[0328] In another embodiment, provided herein is a decellularized microchannel produced by any of the above method(s).

[0329] In yet another embodiment, provided herein is a scaffold biomaterial produced by any of the above method(s).

[0330] In another embodiment, Type, glue, one or more microchannels; one or more decellularizing agents; Scalpel or microtome, Sterile measuring devices, sterile saline, Tweezers and / or Instructions for carrying out any of the methods as described herein Provided herein are kits comprising any one, two, three, four or more of:

[0331] In the examples described below, AS and CL plant tissue sources are identified as offering particularly desirable characteristics. AS is well characterized and works extremely well, while CL offers advantages over AS, such as ease of vascular bundle extraction and a higher number of xylem channels per vascular bundle. However, it will be understood that a wide variety of other plant source tissues can be used. In the examples below, custom-designed molds made of biomedical-grade silicone were used to package / densify and adhere the vascular bundles. A variety of other mold materials are also contemplated, and it will be understood that, in general, any suitable material that does not substantially adhere to adhesives / sealants and / or HDMC bundles, which may be suitably biocompatible or otherwise compatible with the HDMC bundles being produced, can be used. [Example] [Example]

[0332] Preparation of high density microchannels (HDMC) and their biocompatibility analysis Described herein are methods for isolating and decellularizing individual vascular bundles or vascular tissues from any suitable plant, with the goal of avoiding or minimizing the introduction of processing residues or contaminants. The bundling / densification of these vascular channels or tissues with various adhesives and sealants (including naturally occurring and / or synthetic adhesives) has been investigated. A technique for creating custom-designed molds that provide adhesion of vascular bundles (or generally any plant microchannel structure) in a 3D arrangement without substantially causing damage or introducing impurities has been described.

[0333] In this study, we exploited transport structures present in plant vascular bundles (i.e., xylem and phloem channels) to prepare high-density microchannel (HDMC) bundles for various applications. In plants, xylem and phloem channels primarily transport water and sugars, respectively, and are physically and compositionally distinct from surrounding tissues in the plant. They can be visualized as long, continuous, tube-shaped structures, the walls of which typically possess a relatively higher lignin content compared to other structures in the plant. Lignin can impart structural rigidity to these structures, and as described herein, rigidity is achieved by isolating these channels from surrounding non-vascular ground tissues. This may aid in the isolation (mechanical) of the

[0334] As will be appreciated, the chemical composition and structural layout of vascular bundles can vary and may be highly dependent on the particular plant species. In these studies, asparagus-sourced (AS) and celery-sourced (CL) source materials have been identified as providing favorable structural characteristics for application of the decellularized vascular bundles, for example, as a guided support matrix for the growth and regeneration of certain cell types. As will be appreciated, the choice of plant source can be tailored for the particular intended application of the resulting HDMC.

[0335] In these studies, asparagus (family: Asparagaceae) and celery (family: Apiaceae) were selected as source plant materials. These plants were found to have certain favorable characteristics for HDMC fabrication, including: vascular bundles that can be isolated from surrounding ground tissue (i.e., non-vascular tissue) due to differences in structural carbohydrate composition between the vascular and ground tissues; xylem channel diameters large enough to accommodate a variety of cell types and structures (including, for example, neuronal cell bodies); and a relatively high density of xylem and phloem channels within the vascular bundles.

[0336] Figure 1 shows an isolated AS vascular bundle, approximately 3 cm long (left panel), and a microscopic cross-section of the AS vascular bundle (right panel), with the outline of the xylem channels clearly visible against the background. Figure 2 shows an isolated CL vascular bundle, 5-10 cm visible length (left panel), and a microscopic cross-section of the CL vascular bundle (right panel), with the outline of the xylem channels clearly visible against the background tissue.

[0337] As detailed herein, isolated vascular structures, e.g., plant vascular bundles, can be glued together to produce bundles of controllable (or arbitrary) diameter based on the materials selected and used, with the controllable (or arbitrary) length typically limited only at the upper end by the length of the isolated individual structures used to prepare the HDMC (unless sequential longitudinal fusion or linking of two or more structures is performed to obtain longer lengths). In certain embodiments, HDMCs can be bundled together or cut and generally recombined into any desired shape and / or length. Typically, HDMCs are cut to a desired length suitable for the desired application.

[0338] Lignocellulose-containing plant vasculature is considered unique due to its biocompatibility, its structural integrity, and its orientation. As described herein, these mechanisms are imparted to HDMC, and it is contemplated herein that these mechanisms may be useful for several different applications, including, but not limited to, extracellular matrix repair and replacement, even where function may be highly dependent on its orientation (e.g., in the extracellular matrix of the spinal cord). In embodiments in which an adhesive is used to prepare HDMC, it is generally contemplated that the adhesive may be biocompatible (optionally, but preferably) and biodegradable, e.g., to leave the primarily vascular plant structure in the desired orientation and location within the host or physical support.

[0339] In the study described in this example, HDMC was prepared from AS and CL and its biocompatibility was assayed.

[0340] HDMC preparation method: Vascular bundles were first isolated from the AS or CL stalks with a scalpel blade or by gentle peeling, respectively. A dissecting microscope was used to facilitate visualization, but was not necessary for a trained eye. The vascular bundles in both the AS and CL were visually distinguishable from the surrounding ground tissue. If necessary, the stalks were placed at 4°C to allow the vascular bundles to absorb colored water. Contrast could be further enhanced by leaving the specimens in colored water overnight. In either case, excess ground tissue surrounding the isolated vascular bundles could be removed with a scalpel blade. With this method, successful isolation of intact individual vascular bundles was achieved.

[0341] The isolated vascular bundles may then be cut to specific lengths (e.g., with a scalpel or microtome blade). The thickness of the vascular bundles can also be recorded with a digital caliper, taking care not to crush the channels between the caliper arms.

[0342] The vascular bundles were then decellularized, the entire contents of which are incorporated herein by reference. The decellularized bundles were decellularized in a 9-day process utilizing sodium dodecyl sulfate, based on the decellularization protocol detailed in International Publication No. 2017 / 136950, entitled "Cell Wall Structures from Plants and Fungus and Their Use as Scaffold Materials." The decellularized bundles were then sterilized by immersion in 70% ethanol and stored in sterile DPBS at 4°C until use.

[0343] Isolated vascular bundles were bundled together to provide a high-density microchannel (HDMC). In this study, the step of bundling the isolated vascular bundles was accomplished using a custom mold and adhesive. The custom mold was made of biomedical-grade silicone, which was found not to adhere to the fibrin glue used to glue the vascular bundles together in these studies. Prior to the development of the silicone mold, molds made from drinking straws were also used successfully. Drinking straw material is food-grade, and these straws generally possess a small inner diameter comparable to the diameter of a rat spinal cord and therefore may be suitable for certain applications. However, plastic and silicone straws were less effective as molds than biomedical-grade silicone molds because the adhesive adhered to the plastic straws and the silicone straws were more difficult to manipulate due to their flexibility.

[0344] Figure 3 shows the custom-made biomedical-grade silicone mold (5 mm inner diameter, 1 cm length) used for vascular bundle filling in this study (left panel). The mold halves were pressed together, and the right panel shows a 2 mm thick slice of the mold containing the adhered CL vascular bundle.

[0345] The bundling process was carried out under a laminar flow hood to minimize the chance of contaminants becoming lodged in the glued bundles. All handling was performed with autoclaved forceps. First, the fibrin glue kit and its dual syringe application system were prepared according to the manufacturer's instructions (TISSEEL kit; Baxter Healthcare, CA). (Catalog No. 1503152; Document No. 0716820). Individual channels (vascular bundles) to be glued together were then laid out in a sterile staging area. The channels were kept moist to avoid drying and collapse of the inner xylem and phloem. Finally, the silicone mold halves were lined up in a secondary sterile staging area and filled with vascular bundles using tweezers. Packing density varied depending on the inner diameter of the silicone mold and the average thickness of the isolated plant vascular bundles selected for use.

[0346] FIG. 4 shows the AS channel bundle during the adhesive curing step, with the mold halves pressed together, and FIG. 5 shows the CL channel bundle during the adhesive curing step, with the mold halves pressed together.

[0347] Once the mold halves were filled, the channels were removed but not separated. Two to three drops of adhesive were quickly deposited into the interior channel of each mold half, and the filled channel was immediately placed back into the mold. Two to three drops of adhesive were then deposited on top of the packaged channel. The mold halves were then pressed together by manipulation with tweezers. The mold was then left undisturbed for approximately 15 minutes to allow the adhesive to cure. The amount of adhesive to successfully bind the channels together was chosen appropriately, as too little adhesive could result in the channels breaking apart during the mold removal step, and too much adhesive could result in a "globe" of excessive adhesive adhering to the outside of the bundle. The location of the initial adhesive deposit was also found to be helpful in providing uniform adhesion / adhesion throughout the bundle.

[0348] In these studies, adhesives were carefully selected. It was desired that the adhesive be biocompatible, biodegradable, capable of adhering to wet lignocellulosic materials (but not mold materials such as silicone), nontoxic for internal use, and preferably fast-setting. Cyanoacrylate adhesives were also tested but were less effective than fibrin glues, as molds proved difficult to remove from the cyanoacrylate adhesive. Also, while physical adhesion was successful, it was noted that, according to literature reports on surgical adhesives and sealants, certain types of cyanoacrylate adhesives may produce toxic degradation products under certain conditions for internal use.

[0349] After the adhesive has set, excess channel length may be cut away before the bundle is removed from its mold. In these studies, this was best achieved using a microtome blade (very sharp). At this stage, precise lengths of bonded bundles can generally be produced without damaging the bundle by cutting directly through the mold. The mold may provide support compared to the exposed bonded bundle, which may shatter somewhat under certain cutting pressures.

[0350] The mold halves were gently pulled apart with tweezers to reveal the final product: a high-density microchannel (HDMC) containing an adhered plant vascular bundle. The HDMC bundles can be sterilized at this stage, for example, by immersion in 70% ethanol for 30 minutes followed by storage in sterile DPBS at 4°C. Figure 6 shows an HDMC (AS) bundle stored in DPBS (left panel) and an individual HDMC (AS) immediately after removal from the silicone mold (right panel). Figure 7 shows an HDMC (CL) bundle stored in DPBS (left panel) and an individual HDMC (CL) immediately after removal from the silicone mold (right panel).

[0351] Biocompatibility analysis of HDMC: The biocompatibility of HDMC was first evaluated by a one-week cell culture using a 3T3 fibroblast cell line expressing green fluorescent protein (GFP-3T3). Briefly, sections of HDMC were sliced ​​and cells were seeded on the cross-sectional surface of the sections. After one week under standard cell culture conditions (37°C, 5% CO), the sections were fixed with 4% paraformaldehyde solution for microscopy. Figures 8 (AS-based HDMC) and 9 (CL-based HDMC) demonstrate the presence of cells on similar HDMC within the microchannels. Together, the results support that the HDMC bundles were biocompatible.

[0352] Figure 8 shows confocal laser scanning micrographs of HDMC (AS-based) samples after 1 week of GFP-3T3 cell culture followed by fixation in 4% paraformaldehyde. Cells (green—green fluorescent protein) are visible in and around the xylem microchannels (blue—autofluorescence). The left panel shows a cross-section of a single CL vascular channel within the glued bundle, with cells visible within the xylem channel. Note the cell density around the openings of the xylem and phloem channels. The right panel shows a cross-section (partial) of two vascular channels within the glued bundle, with cells scattered throughout.

[0353] Figure 9 shows confocal laser scanning micrographs of HDMC (CL-based) samples after 1 week of GFP-3T3 cell culture followed by fixation in 4% paraformaldehyde. Cells (green - green fluorescent protein) are visible in the xylem microchannels (blue - autofluorescence) and in the xylem microchannels (blue - autofluorescence). The left panel shows a cross-section of a single CL vascular channel within the adhered bundle, with cells visible within the xylem channel. Note that cell attachment to the fibrin glue deposit was localized to the right of the vascular channel. The right panel shows a cross-section of two vascular channels within the adhered bundle, with cells scattered throughout.

[0354] Figures 8 (AS-based HDMC) and 9 (CL-based HDMC) demonstrate the presence of cells on similar HDMC within the microchannels. Taken together, the results support that the HDMC bundles were biocompatible. [Example]

[0355] Examples of Standard Operating Procedures (SOPs) developed for the extraction and isolation of plant or fungal (i.e., AS) vascular bundles (VB) An example of a standard operating procedure (SOP) developed for the extraction and isolation of plant or fungal vascular bundles and / or microchannels is described below. This SOP was used for the extraction and isolation of vascular bundles from AS sources.

[0356] Safety Statement Maintain awareness of the position of the scalpel blade and the side of the microtome blade to guard against accidental cuts. Consult all appropriate MSDSs. Wear appropriate personal protective equipment, including vinyl gloves, a lab coat, and goggles.

[0357] solution Dulbecco's phosphate-buffered saline DPBS (Hyclone, Catalog No. 350-000-CL) 500ml bottle Store in the dark at 4°C. Do not use if the expiration date has passed.

[0358] procedure Asparagus selection Asparagus bunches were selected from the produce section of a grocery store. Bunches consisting of large diameter (>1.0 cm) asparagus sprouts were preferably selected. After selection and purchase, the asparagus was immediately transported to a refrigerated storage environment (4°C). Asparagus bunches that are not processed within an hour of storage can be placed in shallow water (while maintaining refrigeration) to prevent the sprouts from drying out.

[0359] Asparagus bunches were preferably processed within 24 hours of purchase.

[0360] Generation of AS vascular bundles [Note: If other plant or fungal tissue has previously been treated, all tools and cutting mats can be disinfected with 70% ethanol and Accel TB solution before treatment.]

[0361] Wearing fresh gloves, the produce tag identifying the asparagus grower and PLU number was removed from the bunch and recorded. The asparagus sprouts were removed from their packaging rubber bands and rinsed under tap water. The sprouts were placed in a beaker of shallow water next to a cutting mat.

[0362] A large weighing boat was placed next to the cutting mat and a small amount (approximately 25 ml) of DPBS was poured into the boat, which was used to hold the isolated AS vascular bundles as they were generated.

[0363] A scalpel (Feather number 10) and microtome blade were prepared for use.

[0364] The buds were treated one by one. The following procedure was applied to each individual bud: The sprouts were removed from the shallow beaker of water and placed on a cutting mat. A microtome blade was used to cut off the bottom 2 cm (approximately) of the sprout. If the new end of the sprout still appeared discolored, dry, or otherwise damaged, an additional 1 cm segment was cut until the end consisted of fresh, undamaged tissue. A 3 cm piece (referred to as the "stump") measured from the freshly cut bottom end of the bud was cut using a microtome blade. This stump represented the "bottom portion" of the bud. The 3-cm stump was cut in half lengthwise using a microtome blade. The resulting halves were examined for visible vascular bundles near the cut surface (see, eg, Figure 10). Half of each stump was brought to a mandoline blade (adjusted for approximately 1 mm thick slices) and sliced ​​into approximately 1 mm thick slices until no more slices were produced, which were dropped directly into a weigh boat containing DPBS. The weighing boat containing the thin slices was placed on a cutting board. Slices were expected for vascular bundles that extended across the entire sheet. Vascular bundles extending over at least ¾ of the slice length were selected for isolation in the next step. Vascular bundles were individually excised from the slices using a scalpel blade or a microtome blade. Cutting with a scalpel blade was preferably performed straight and evenly (do not lift the blade, as this may create serrated edges on the side of the isolated VB). Care was taken to remove as much of the surrounding ground tissue as possible when extracting the VB. The isolated VB cuttings were not modified with any cutting tool as this may create uneven edges or serrated edges. The isolated vascular bundle was immediately visually inspected. Any of the following observations resulted in the isolated vascular bundle being discarded: · Kinks or breaks in the vascular bundles. Any VB with continuous vasculature less than 1.5 cm in length. Isolated vascular bundles passing visual inspection were immediately deposited into weigh boats (containing DBPS) using clean forceps (see Figure 11). All slices were processed one at a time to isolate the VB as described above. After all slices were processed, another 3 cm stump was cut from the asparagus sprout using a microtome blade, this stump representing the "middle" portion of the sprout. A 3 cm piece was cut and treated exactly like the first 3 cm stump to create more isolated vascular bundles. The extracted VBs may be sorted according to their origin in the asparagus sprout (bottom, middle, or top stump). The final 3 cm stump was cut and treated exactly like the first and second 3 cm stumps to create more isolated vascular bundles.

[0365] Asparagus sprouts were treated as described above until the target number of VBs was produced.

[0366] If any single asparagus sprout exhibited any of the following symptoms, processing was discontinued and the entire AS batch was discarded: Mold ·corruption ·Insect parasitism

[0367] If the sprouts showed any of the following symptoms, they were discarded and not processed (however, processing of the remaining sprouts could still proceed and the batch was not discarded): Dry cracks or cracks Excessively curved or otherwise misshapen buds.

[0368] Note: If the process was interrupted for more than 0.5 hours, the sprouts currently being processed were discarded and the remaining sprouts (in shallow water) were returned to a refrigerated environment (4°C). All remaining sprouts were preferably processed within 24 hours of their purchase.

[0369] Preparation of AS vascular bundles for decellularization Using forceps, the isolated AS vascular bundles in the weigh boat were placed into a 50 ml Falcon tube. The vascular bundles were counted and the total number per tube was indicated on the tube.

[0370] Note: A 50 ml centrifuge tube (Nunc) preferably contained no more than 60 AS vascular bundles. If a weigh boat contained more than 60 vascular bundles, multiple 50 ml tubes were used.

[0371] Each tube was labeled with the following information: Date in year-month-day format. ·Operator's three-letter identifying initials. Product codes beginning with the letters "AS". The number of AS vascular bundles contained in the tube is written on the tube cap.

[0372] If VB decellularization was desired, the tubes were immediately processed for decellularization according to the SOP set out in Example 4 below.

[0373] Disposal of plant tissue waste and cleaning of tools Waste asparagus and celery tissue was collected in disposable bags. The waste material was sorted as compostable material.

[0374] The cutting mat was rinsed with tap water. Dry plant tissue was gently removed. The mat was thoroughly disinfected with 70% ethanol solution and Accel TB solution.

[0375] Chef's knives, tweezers, scalpel blades and microtome blades were cleaned in the same manner as the cutting mats described above.

[0376] Metal tools were gently wiped dry using Kimwipes.

[0377] The DPBS solution in the weigh boat was collected in an appropriate liquid waste jug. [Example]

[0378] Examples of standard operating procedures (SOPs) developed for the extraction and isolation of vascular bundles (VB) from plants or fungi (i.e., CL) An example of a standard operating procedure (SOP) developed for the extraction and isolation of plant or fungal vascular bundles and / or microchannels is described below. This SOP was used for the extraction and isolation of vascular bundles from CL sources.

[0379] Safety Statement Be aware of the position of the scalpel blade and the side of the microtome blade to prevent accidental cuts. Refer to all appropriate MSDS. Wear appropriate personal protective equipment, including vinyl gloves, a lab coat, and goggles.

[0380] solution Dulbecco's phosphate-buffered saline DPBS (Hyclone, Catalog No. 350-000-CL) 500ml bottle Store in the dark at 4°C. Do not use if the expiration date has passed.

[0381] procedure Celery Selection Celery plants were selected from the produce section of a grocery store. The plants contained no visibly damaged stems, preferably firm stems. After selection and purchase, the celery plants were immediately transported to a refrigerated storage environment (4C). Celery plants were preferably processed within 24 hours of purchase.

[0382] Generation of CL vascular bundles [Note: If you have previously treated other plant or fungal tissue, disinfect all tools and cutting mats with 70% ethanol and Accel TB solution before proceeding.]

[0383] Wearing new gloves, the produce tag identifying the producer of the celery and the PLU number (if present) was removed from the celery and recorded in the logbook. The celery was rinsed with tap water.

[0384] I placed the celery next to the cutting mat. I used a small knife (Feather #10) and a chef's knife. A large weighing dish was placed next to the cutting mat and a small amount (approximately 50 ml) of DPBS was poured into the dish. This dish was used to hold the isolated AS vascular bundles as they were generated. One celery stalk was manually removed from the stalk and placed on the cutting mat. The following steps apply to each individual celery stalk: Using a chef's knife, I cut off the bottom 4-5cm of the stem, this part of the stem is usually thicker than the rest of the stem and often has dirt particles hiding on the surface. If there were leaves at the top of the stem, I used a chef's knife to cut off the top until the leafy part was gone. A 1- to 2-cm section was carefully removed manually from the top of the stem, taking care not to sever the vascular bundles that run near the outer surface of the stem. The exposed vascular bundles were peeled away and the entire length of the stalk was pulled. Peeling was done slowly to avoid damaging the vascular bundles while they were still embedded in the surrounding celery tissue. The exposed vascular bundles could be peeled away as a group or individually. The detached vascular bundles were placed on a cutting board, and loosely attached excess ground tissue was manually removed with gloved hands. The vascular bundle was cut into 6 cm long sections using a scalpel. The isolated vascular bundle was immediately visually inspected and discarded if any of the following findings were present: · Kinks or breaks in the vascular bundles. - Excessive spiraling, where vascular bundles tend to naturally curl up into strong loops. Freshly cut vascular bundles that passed visual inspection were placed in a weighing dish containing Dulbecco's phosphate buffered saline (DPBS) (see Figure 12).

[0385] All celery stalks except the innermost light-colored stalks of the plant were processed as described above. Unused inner stalks were discarded.

[0386] If even one celery stalk showed any of the following symptoms, processing was stopped and the entire CL batch was discarded: Mold ·corruption ·Insect contamination

[0387] If any of the following symptoms were observed in the shoots, the stems were discarded and not processed (however, processing of the remaining shoots could still continue and the batch was not discarded): Splitting or cracking due to drying Broken or split stems due to damage in the grocery store or during transportation

[0388] NOTE: If the process was interrupted for more than 0.5 h, the stems currently being processed were discarded and the remaining stems were returned to a refrigerated environment (4 °C). All remaining stems were preferably processed within 24 h of purchase.

[0389] Preparation for decellularization of CL vascular bundles Using forceps, the isolated CL vascular bundles in the weighing dish were placed into a 50 ml Falcon tube. The vascular bundles were counted and the total number per tube was labeled on the tube.

[0390] NOTE: The 50 ml Falcon tube contained no more than 50 CL vascular bundles. If the weighing dish contained more than 50 vascular bundles, multiple 50 ml Falcon tubes were used.

[0391] Each tube was labeled with the following information: Date in YYYY-MM-DD format. The operator's three-letter initials. · Generated code beginning with the letters "CL". The number of CL vascular bundles contained inside the tube was written on the tube cap.

[0392] The tubes were immediately processed for decellularization according to the SOP detailed in Example 4 below.

[0393] Disposal of plant tissue waste and cleaning tools Discarded asparagus and celery tissue was collected in disposable bags. Waste was separated as compostable material.

[0394] The cutting mat was rinsed with tap water. Dried plant tissue was carefully removed. The mat was thoroughly disinfected with 70% ethanol solution and Accel TB solution.

[0395] Chef's knives, tweezers, scalpel blades, and microtome blades were cleaned in the same manner as the cutting mat. Metal tools were gently wiped dry using Kimwipes. The DPBS solution in the weighing dish was collected in an appropriate liquid waste container. [Example]

[0396] Example of a standard operating procedure (SOP) developed for decellularization of vascular bundles (VB) from plants or fungi (i.e., AS and CL) using sodium dodecyl sulfate (SDS). An example of a standard operating procedure (SOP) developed for the decellularization of plant- or fungal-derived vascular bundles and / or microchannels using sodium dodecyl sulfate (SDS) is described below. This SOP was used to decellularize vascular bundles obtained from AS and CL sources. While the following SOP focuses primarily on plant tissue, it is believed that fungal tissue can be treated similarly. In certain embodiments, it is believed that some adjustments to the time in solution and / or concentration of SDS and / or CaCl can be made, for example, for fungal tissue.

[0397] solution 0.1% SDS solution SDS powder (Fisher, Catalog No. BP166-500) 0.5g Sterile water (Baxter, Cat. No. JF7624) final volume 500 ml Prepared in autoclaved bottles (Fisher, Cat. No. FB-800-500) Store at 20-25°C. Use within 7 days. 0.1M CaCl2 solution 5.549g CaCl2 powder (Acros Organics, catalog number 3496150000) Sterile water (Baxter, Cat. No. JF7624) final volume 500 ml Prepared in autoclaved bottles (Fisher, Cat. No. FB-800-500) Store at 20-25°C. Use within 7 days. Dulbecco's phosphate-buffered saline DPBS (Hyclone, Catalog No. 350-000-CL) 500ml bottle Store in the dark at 4°C. Do not use anything that has passed its expiration date. Sterile ethanol solution HistoPrep 95% RA (Fisher, Catalog No. HC13001GL) 368.4 ml Deionized water (Millipore) final volume 500 ml Autoclave under a biosafety cabinet (BSC). Prepared in a 100% ethanol bottle (Fisher, Cat. No. FB-800-500). Store at 20-25°C. Use only under BSC. Stable indefinitely.

[0398] procedure Note 1: The source and identity of all solutions used in the following procedure was recorded in the operator's log. Note 2: It is preferred to use the following procedure immediately after completing the SOPs described in Examples 2 and / or 3 above.

[0399] Decellularization with sodium dodecyl sulfate (SDS) (Days 1-4): Day 1: Freshly cut and isolated vascular bundles (VBs) were transferred from the DPBS storage solution into 50 ml centrifuge tubes (Nunc). To ensure proper mixing of the VBs in the SDS solution, no more than 60 VBs were placed in each tube. Each centrifuge tube was filled with 0.1% SDS solution (up to the 50ml fill line). The tube caps were then tightened firmly into place. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. The start time was recorded.

[0400] Day 2: After 20–24 h of exposure of VB to SDS solution on an orbital shaker, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the SDS solution was decanted from each tube into a lab sink. The tube was filled with unused 0.1% SDS solution and the cap was re-tightened. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0401] Day 3: Twenty to 24 hours after the end of the SDS solution change on day 2, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the SDS solution was decanted from each tube into a lab sink. The tube was filled with unused 0.1% SDS solution and the cap was re-tightened. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0402] Day 4: Twenty to 24 hours after the end of the SDS solution change on day 3, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the SDS solution was decanted from each tube into a lab sink. The tube was filled with unused 0.1% SDS solution and the cap was re-tightened. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0403] Removal of sodium dodecyl sulfate (SDS) by CaCl salt wash (days 5-6) Day 5: Twenty to 24 hours after the end of the SDS solution change on day 4, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the SDS solution was decanted from each tube into a lab sink. · The tube was filled with DI water up to the 50ml fill line. The water was decanted from each tube into a laboratory sink. The DI water rinse described in the previous two steps was repeated two more times. After decanting the third DI water rinse, each tube was filled to the 50ml fill line with 0.1M CaCl2 solution and re-capped. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0404] Day 6: Twenty to 24 hours after the end of the change from SDS to CaCl2 solution on day 5, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the CaCl2 solution was decanted from each tube into a laboratory sink. Each tube was filled to the 50 ml fill line with unused 0.1 M CaCl2 solution and recapped. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0405] Removal of CaCl2 salt by water washing (days 7-8): Day 7: Twenty to 24 hours after the end of the CaCl2 solution change on day 6, the shaker was stopped and the tubes were removed from the shaker. The caps of each tube were removed and the SDS solution was decanted from each tube into a lab sink. · The tube was filled with DI water up to the 50ml fill line. The water was decanted from each tube into a laboratory sink. The DI water rinse described in the previous two steps was repeated two more times. After decanting the DI water from the third rinse, each tube was filled to the 50ml fill line with water and re-capped. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0406] Day 8: On day 7, 20-24 hours after the end of the change from CaCl2 to aqueous solution, the shaker was stopped and the tubes were removed from the shaker. The cap of each tube was removed and the water was decanted from each tube into a laboratory sink. · The tube was filled with unused water up to the 50ml fill line and the cap was re-tightened. The tube was fixed to the plate of an orbital shaker (Corning LSE) using tape. The orbital shaker was set to rotate at 120 RPM with no time limit. The shaker was observed for at least 1 minute to ensure the tube was secured and was not in danger of coming loose. Time was recorded.

[0407] Sterilization (9th day): Day 9: Twenty to 24 hours after the end of the water change on day 8, the shaker was stopped and the tubes were removed from the shaker. The time was recorded. The caps of each tube were removed and the water was decanted from each tube into a laboratory sink. Retightened the tube cap. The tube was sprayed with 70% ethanol solution and brought into a biosafety cabinet (BSC). Each tube was filled with sterile ethanol up to the 30 ml fill line and the tube cap was re-tightened. The tubes were placed upright in a tube rack so that all of the decellularized VB was immersed in the ethanol solution. The tube was allowed to stand for 15 minutes. After 15 minutes, the tube was inverted and allowed to stand for another 15 minutes. After 15 minutes, the tube was placed upright again and the cap was removed. The ethanol solution from each tube was removed with a 10 ml serological pipette and collected in a glass waste bottle in the laboratory sink. Each tube was filled to the 30 ml fill line with sterile DPBS using a 10 ml serological pipette, followed by gentle swirling. DPBS was aspirated from each tube using a sterile Pasteur pipette. Repeat the previous two steps two more times for a total of three sterile DPBS rinses. After the third rinse volume of DPBS was aspirated from each tube, each tube was filled with 30 ml of sterile DPBS and the tube caps were screwed firmly back into place. Each tube was labeled with the following additional information: production code, sterilization date, liquid solution content, and sterilization status. The tubes were kept in the dark at 4°C. [Example]

[0408] Fabrication of high density microchannel (HDMC) bundles from lignocellulosic materials (i.e., AS and CL) using fibrin glue Described below is an example of a standard operating procedure (SOP) developed for the fabrication of high-density microchannel (HDMC) bundles from lignocellulosic materials (i.e., AS and CL) using fibrin glue, which was used to bundle and glue vascular bundles obtained from AS and CL sources.

[0409] Safety Statement Always protect against needlestick injuries when handling hypodermic needles. Fibrin glue and fibrin glue derived products (including protein vials) were treated as biohazard level 2 materials. Consult all appropriate MSDSs. Appropriate personal protective equipment, including vinyl gloves, lab coat, and goggles, was worn and all work was performed under a laminar flow hood.

[0410] Solutions and Reagents Dulbecco's phosphate-buffered saline DPBS (Hyclone, Catalog No. 350-000-CL) 500ml bottle Store in the dark at 4°C. Do not use if the expiration date has passed.

[0411] TISSEEL Fibrin Sealant Kit (Baxter Healthcare, catalog number 1503152) Aprotinin (synthetic) solution 3000KIU / ml 1ml Sealer Protein Concentrate (Human) Reconstituted with Aprotinin Solution to 96-125mg / ml 1 ml of calcium chloride solution at 25 micromoles / ml Thrombin (human) reconstituted with calcium chloride solution to 500 IU / ml Ingredients stored at 2-25°C. Prepare the solution with a dual syringe system according to the manufacturer's instructions (see procedure below). .

[0412] Sterile ethanol solution HistoPrep 95% RA (Fisher, Catalog No. HC13001GL) 368.4 ml l Deionized water (Millipore) final volume 500 ml Prepared in autoclaved bottles (Fisher, Cat. No. FB-800-500) under a biosafety cabinet (BSC). Store at 20-25°C. Use only under BSC. Stable indefinitely.

[0413] Procedure - Preparing the Adhesive Preparation of Tisseel and Duploject NOTE: TISSEEL preparation follows manufacturer's instructions (Baxter Document ID 0716820). The procedures below are equivalent restates of these procedures to suit local laboratory conditions.

[0414] All four vials in the TISSEEL kit fit into the wells of a FIBRINOTHERM (Baxter) device. The device warms the contents of the vial to 37°C and maintains this temperature. A signal light appears when warming is complete. If the vial is refrigerated, warming may take up to 5 minutes.

[0415] After pre-heating was complete, all four vials were transferred to a biosafety cabinet (BSC).

[0416] The plastic flip-off caps on the aprotinin and sealer protein vials were removed without touching the rubber vial stoppers, which were disinfected with Kimwipes containing Accel TB disinfectant. The rubber stoppers were allowed to dry.

[0417] The aprotinin solution was transferred into a sealer protein vial using the blue graduated syringe and needle tip provided in the kit.

[0418] The sealer protein vial was transferred to the stirring well of the FIBRINOTHERM device (the vial already contained a small magnetic stir bar). The contents of the vial were stirred for 10 minutes. Mixing was complete when the vial solution was clear and free of air bubbles and particles. DUPLOJECT Dual The vial remained in the 37°C warming well until the syringe system was assembled. The flip-off plastic caps on the calcium chloride and thrombin vials were removed without touching the rubber vial stoppers, which were disinfected with Kimwipes containing Accel TB disinfectant solution. The stoppers were allowed to dry.

[0419] The calcium chloride solution was transferred into the thrombin vial using the black graduated syringe and needle tip provided in the kit.

[0420] The thrombin vial was gently swirled until clear and then transferred to a FIBRINOTHERM device for rewarming to 37°C.

[0421] The vials containing the two solutions (originally the sealer protein and thrombin vials) were transferred from the FIBRINOTHERM device to the BSC.

[0422] The rubber stoppers of both vials were wiped with a Kimwipe containing Accel TB and allowed to dry.

[0423] Using the second blue graduated syringe and needle tip from the kit, the solution from the sealer protein vial was drawn into the syringe.

[0424] The needle tip was removed from the syringe (discarded in a sharps container) and the syringe was clipped onto the red dual syringe-holding DUPLOJECT assembly.

[0425] Using the second black graduated syringe and needle tip from the kit, the solution from the thrombin vial was drawn into the syringe.

[0426] Remove the needle tip from the syringe (discard it in a sharps container) and place the syringe in the red DUPLOJECT assembly. Clipped onto the umbri.

[0427] The plastic connector was attached to the clipped end of the syringe.

[0428] An application cannula was attached to the end of the interface.

[0429] The completed DUPLOJECT assembly remains stable and horizontal until fibrin glue is needed. The tube was placed upright against a surface (such as a tube rack) and the fibrin glue was preferably used within 4 hours.

[0430] Vascular bundle (VB) connections Note: All materials described in this subsection were autoclaved for sterilization and removed from the autoclave only once within the BSC.

[0431] A tube containing the correct type (AS or CL) and quantity of decellularized and sterilized vascular bundles (20 VB per HDMC bundle) was selected from storage and brought into the BSC along with the following: Microtome blades Custom-made silicone molds (one mold per HDMC bundle) ·tweezers 24-well plates (sterilized and packaged; at least two plates were used when producing HDMC from one type of VB)

[0432] The solid contents of the VB tube were transferred to the inner surface of the lid of a 24-well plate using sterile tweezers, and 5 ml of sterile DPBS was added to prevent the VB from drying out.

[0433] Using sterile forceps, up to two silicone molds were removed from the autoclave pouch and placed inside the inner surface of a second 24-well plate lid with the internal channels open and facing upward (toward the operator). No more than four mold halves (two HDMCs total) were manipulated within a single plate lid.

[0434] Using a pair of sterile forceps, 10 VBs were packed into each channel of the mold half.

[0435] The packed VB was removed in bundles from each mold half, and the bundles removed from each half were placed immediately in front of the mold.

[0436] Using the DUPLOJECT system, three drops were placed along the length of the internal channel of each mold half. Fibrin glue was deposited.

[0437] Immediately after the adhesive was deposited, the bundled VB bundle was repositioned in the mold channel, and gentle pressure was applied with the tips of tweezers to pack the VB tightly into the adhesive-filled mold channel.

[0438] The fibrin glue was allowed to harden for 10 minutes, during which time photographs of the mold were taken.

[0439] After 10 minutes, replace the applicator tip of the DUPLOJECT system with a new, unclogged tip. and exchanged it for

[0440] Three drops of fibrin glue were deposited along the length of each packed VB bundle.

[0441] Immediately after the adhesive was deposited, the mold halves (each containing 10 packed VBs) were pressed together using tweezers. The mold was oriented so that the halves remained attached without the need for tweezers contact.

[0442] The fibrin glue was allowed to harden for 10 minutes, during which time photographs of the mold were taken.

[0443] After 10 min, the edge of the VB protruding from the silicone mold was cut off with a microtome blade and discarded.

[0444] The mold halves were gently separated using tweezers. The HDMC bundle product could be released by gently prying open one of the mold halves. A photograph of the released HDMC was taken.

[0445] NOTE: HDMC bundles may be sterilized at this stage by immersion in 30 mL of 70% ethanol solution for 30 min, followed by rinsing with 20 mL of DPBS 3x.

[0446] The HDMC bundle product was placed in 50 ml centrifuge tubes (Nunc) containing 10 ml sterile DPBS (one tube per HDMC).

[0447] The HDMC tubes were stored in the dark at 4°C. [Example]

[0448] Test examples of various adhesives An example of fabricating bonded high density microchannel (HDMC) bundles and testing various types of adhesives in a molding process is described below.

[0449] AS adhesion with PEG-based synthetic adhesive method: Decellularized AS vascular bundles (VB) produced by the SOP (decellularization) of Example 4 were used. Molding: Two halves of a biomedical grade silicone (Dow Corning Silastic MDX4-4210) mold were filled with VB: 10 VB per mold half; 20 VB per HDMC. Adhesive: Baxter's Coseal adhesive was used. Two drops were deposited inside each mold half, after which VB was packed into the channels. After five minutes, two more drops of adhesive were deposited on the packed VB in each mold half before the mold halves were pressed together (Figure 13). A 10-minute cure time was allowed before attempting to separate the mold halves. Aseptic assembly was performed in a BSC.

[0450] result: Coseal adhesive produced a cohesive gel immediately after deposition onto the mold halves. However, the expansion volume was significant, estimated at approximately 400% by volume. Due to the mold used and the specific setup of this experiment, which was not designed to accommodate the expansion, the adhesive expansion was detrimental, causing the mold halves to separate, and therefore the HDMC cylindrical shape was not maintained. Because Coseal adhesive adheres more strongly to the silicone mold than fibrin adhesive, it was also found that the HDMC bundles split in half if the adhesive was not carefully separated from the mold halves. The HDMC bundles resulting from this experiment were loosely bound and did not have a cylindrical shape when removed from the mold (see Figure 14). Although difficulties were encountered with the PEG-based adhesive in this experimental system, it is believed that these can be adapted to accommodate the swelling and tackiness of the PEG-based adhesive, for example, by using a larger mold and / or molding as described above, by using a lower microchannel density to allow for swelling, and / or by using a mold of a different material, for example, that has less adhesion to the PEG-based adhesive.

[0451] CL adhesion with PEG-based synthetic adhesive method: Decellularized CL vascular bundles (VB) were prepared according to the SOP (decellularization) of Example 4. Molding: Two halves of a biomedical grade silicone (Dow Corning Silastic MDX4-4210) mold were filled with VB: 10 VB per mold half; 20 VB per HDMC. Adhesive: Baxter's PEG-based Coseal adhesive was used. Two drops were deposited inside the mold halves, after which VB was packed into the channels. After 5 minutes, two more drops of adhesive were deposited on the VB packed into each mold half before the mold halves were pressed together (Figure 15). A 10-minute cure time was allowed before attempting to separate the mold halves. Aseptic assembly was performed in a BSC.

[0452] result: Coseal adhesive produced a cohesive gel immediately after deposition onto the mold halves. However, the expansion volume was significant, estimated at approximately 400% by volume. Due to the mold used and the specific setup of this experiment, which was not designed to accommodate expansion, adhesive expansion was detrimental, causing the mold halves to separate, and therefore the HDMC cylindrical shape was not maintained. Because Coseal adhesive adheres more strongly to the silicone mold than fibrin adhesive, it was also found that if the adhesive was not carefully separated from the mold halves, each half would remain adhered to its silicone mold half, causing the HDMC bundle to split into two. The HDMC bundles obtained from this experiment were loosely bound and did not have a cylindrical shape when removed from the mold (see Figure 16, where partial detachment of individual VBs is observed). While difficulties were encountered with the PEG-based adhesive in this experimental system, it is believed that adaptations can be made to accommodate the expansion and adhesive properties of the PEG-based adhesive by, for example, using a larger mold and / or the aforementioned molding, by using a lower microchannel density to allow for expansion, and / or by using a mold of a different material, e.g., one that adheres less well to the PEG-based adhesive.

[0453] AS bonding with cyanoacrylate adhesive (Krazy adhesive) method: Decellularized AS vascular bundles (channels) were prepared according to the SOP (decellularization) of Example 4 as described above. Molding: Two halves of a plastic straw (inner diameter 5 mm) were stuffed with a channel. Adhesion: Cyanoacrylate adhesive (Krazy glue) was used. The mold halves were pressed together. Before being poured into the packed channel, droplets were deposited. The method was not sterile and was assembled manually.

[0454] result: Bundles were successfully assembled. Under these conditions, the maximum packing density was approximately 34 channels in a 5 mm diameter, which compares favorably with the 5–9 channels observed in native AS pieces of similar diameter (see Figures 17–18).

[0455] AS adhesion using biodegradable fibrin sealant (TISSEEL) method: Decellularized AS vascular bundles (channels) were prepared according to the SOP (decellularization) in Example 4 above. Molding: Two halves of a custom-made silicone (Sylgard 184) mold (inner diameter 4 mm) were inserted into the The silicone material did not adhere to the fibrin sealant, and the bonded AS channel was easily removed from the mold. The molds were autoclaved in autoclave pouches before use. Adhesion: The adhesive was fibrin sealant. A drop was applied to the packed channel using the Baxter DUPLOJECT system for TISSEEL application before the mold halves were pressed together. Aseptic assembly was performed in a laminar flow hood using autoclaved forceps. Assembly was completed inside the lid of a sterile 24-well plate.

[0456] result: Adhesive bundles were successfully assembled. The maximum packing density under the conditions used was approximately 20 channels in a 4 mm diameter. This was a slightly reduced density relative to the cyanoacrylate adhesive. Figure 19 shows the AS channels in the connected silicone mold halves during the curing period.

[0457] The same experiment was repeated with minor modifications. The channel modification was reduced to 18 channels per bundle, and the average diameter of each individual channel was slightly increased. Figure 20 shows the appearance of the AS channels in the silicone mold during the adhesive curing step. Figure 21 shows the appearance of the fibrin-glued AS channels after removal from the silicone mold. Sterilization was performed by immersing the glued bundle samples in 70% ethanol for 30 minutes, followed by three washes with sterile DBPS. The samples were stored in sterile DPBS at 4°C before in vitro testing.

[0458] In vitro biocompatibility testing was then performed. GFP-3T3 fibroblasts were cultured on the adhered bundle samples for one week and then fixed with 4% paraformaldehyde. Cells initially deposited at the exposed channel openings. Confocal laser-scanning microscopy (CLSM) of the fixed samples was performed to determine the presence or absence of cells. Figure 22 (left panel) shows an oblique view of an AS sample after culturing GFP-3T3 cells for one week and fixing with 4% paraformaldehyde. The right panel shows a top view of the same sample.

[0459] Figure 23 shows CLSM images of AS HDMC bundles fixed with 4% paraformaldehyde after 1 week of GFP-3T3 cell culture, revealing cells (green) within and around the xylem microchannels (blue).

[0460] Results and Discussion: Multiple adherent HDMC bundles were successfully generated. After one week of culture, cells were visible both within the channel and on the surrounding material (adhesive, outside the channel). The presence of cellulose enabled visualization of the cells, and the autofluorescence of the xylem microchannels allowed visualization of the cellulose scaffold of AS.

[0461] These experiments were also repeated with some modifications, with mold halves formed from food-grade silicone drinking straws. The mold halves were autoclaved in autoclave pouches prior to use, as opposed to Sylgard 184 silicone, which is not food-grade. It was noted that manipulating the small mold halves with autoclaved tweezers was more difficult than, for example, manipulating the larger, custom-made Sylgard 184 mold halves. The same sterilization procedures as above were followed. was used, and the resulting HDMC bundles were intended for use in transplantation studies in rats (see Example 8 below).

[0462] We successfully generated multiple adhered AS bundles. Seven samples, each approximately 5 mm long and 5 mm in diameter, were generated. During storage at 4°C, some samples failed to adhere or separated, likely due to insufficient fibrin glue in some bundles. Figure 24 shows the appearance of the AS channel in the food-grade silicone mold during the glue curing step. Figure 25 shows the appearance of an adhered AS channel sample stored in sterile DPBS after cutting into 5 mm lengths.

[0463] Some samples were lost during storage, so the procedure was repeated but with a little more glue to keep the channels together within the silicone straw mold.

[0464] Multiple adherent AS bundles were successfully prepared. Four samples, each 5 mm long and 5 mm in diameter, were generated, and no loss of samples was observed during storage. The results of subcutaneous implantation are shown in Example 8 below.

[0465] CL adhesion using cyanoacrylate adhesive (strong instant adhesive / "Krazy adhesive") method: A decellularized CL vascular bundle (channel) was prepared according to the SOP (decellularization) of Example 4. Molding: The channel was packed into two halves of a plastic straw (inner diameter 5 mm). Gluing: Cyanoacrylate glue ("Krazy glue") was used. The mould halves were pressed together. A droplet was deposited in the packed channel before being compressed. Non-sterile manual assembly was performed.

[0466] result: The assembly of glued bundles was successfully achieved. The maximum packing density under these conditions was approximately 24 channels in a 5 mm diameter. Figure 26 shows the side view of a CL channel bundle glued with cyanoacrylate adhesive. Figure 27 shows the emitting surface of a CL channel bundle glued with cyanoacrylate adhesive, with individual channels faintly visible.

[0467] CL adhesion using biodegradable fibrin sealant (TISSEEL) method: A decellularized CL vascular bundle (channel) was prepared according to the SOP (decellularization) of Example 4. Molding: Two channels were custom-made in silicone (Sylgard 184) molds (inner diameter 4 mm). The silicone material did not adhere to the fibrin sealant, and the glued CL channel was easily removed from the mold. The molds were autoclaved in autoclave pouches before use. Adhesion: Biodegradable fibrin sealant was used as the adhesive. A drop was applied to the packed channel using the Baxter DUPLOJECT system for TISSEEL application before the mold halves were pressed together. Aseptic assembly was performed in a laminar flow hood using autoclaved forceps. Assembly was completed inside the lid of a sterile 24-well plate.

[0468] result: Bonded bundles were successfully assembled. The maximum packing density under the conditions tested was approximately 20 channels in a 4 mm diameter. Figure 28 shows the CL channels in the silicone mold halves connected together during the curing period.

[0469] This experiment was repeated to generate more bundle samples for further testing. The same fabrication procedure was used, but the channel packing density was modified to reduce to 18 channels per bundle to account for the variation (increase) in the average size of the CL channels. Sterilization was performed by immersing the bonded bundle samples in 70% ethanol for 30 minutes, followed by washing three times with sterile DBPS. The samples were stored in sterile DPBS at 4°C before in vitro testing.

[0470] In vitro biocompatibility testing was performed as follows: GPF-3T3 fibroblasts were cultured on the adhered bundle samples for 1 week and then fixed with 4% paraformaldehyde. The cells were first deposited on the exposed channel openings. Confocal laser scanning microscopy (CLSM) of the fixed samples was used to determine the presence or absence of cells.

[0471] result: We successfully generated multiple adherent CL bundles. After one week of culture, cells were visible both within the channel and on the surrounding material (adhesive, external to the channel). The presence of GFP protein allowed visualization of the cells, and the autofluorescence of the xylem microchannels allowed visualization of the cellulose scaffolding of the CLs.

[0472] Figure 29 shows the appearance of the CL channel in the silicone mold during the adhesive curing step. Figure 30 shows the appearance of the fibrin-adhered CL channel after removal from the silicone mold. Figure 31 shows an oblique view (left panel) of a CL bundle sample fixed with 4% paraformaldehyde after one week of GFP-3T3 cell culture, and a top view (right panel) of the same sample. Figure 32 shows a CLSM image of a CL sample fixed with 4% paraformaldehyde after one week of GFP-3T3 cell culture. Cells (green) can be seen inside and around the xylem microchannel (blue).

[0473] This experiment was repeated, but modified to use mold halves formed from food-grade silicone drinking straws to provide specimens for subcutaneous implantation in rats (see Example 8). Mold halves were placed in pouches and autoclaved prior to use. Manipulating the small mold halves with autoclaved forceps was more difficult than manipulating the larger custom-made Sylgard 184 mold halves. Using the same sterilization procedures as above, samples were intended for subcutaneous implantation in rats (see Example 8).

[0474] Results: We successfully generated multiple adherent CL bundle samples. Nine samples, each 5 mm long and 5 mm in diameter, were generated. During storage at 4°C, three of the nine samples ultimately failed to adhere or separated, likely due to insufficient fibrin adhesion of these three bundles.

[0475] Figure 33 shows the appearance of the CL channels in the food-grade silicone mold during the adhesive curing step. Figure 34 shows the appearance of the bonded CL channel bundle after removal from the top silicone mold half. Figure 35 shows the appearance of the bonded CL channel sample (stored in sterile DPBS) after cutting into 5 mm lengths. [Example]

[0476] Guidance properties and directionality of HDMC bundles To further investigate the ability of the HDMC bundles described herein to induce cell directionality, As described herein, a scaffold biomaterial capable of providing directionality and / or guidance for cell growth / invasion / proliferation is desired. While many scaffolds in the field are unable to provide directionality, the present inventors have developed a scaffold biomaterial comprising a high-density microchannel bundle as described herein, in which decellularized microchannels are arranged substantially parallel to one another within the bundle, that can provide directionality to cells growing on and / or within the bundle.

[0477] A description of one such study follows:

[0478] method: (HDMC Fabrication): HDMCs were fabricated using decellularized vascular bundles (VBs) of AS and CL raw materials, generally following the SOPs in Examples 3, 4, and 5. Briefly, isolated AS and CL VBs were decellularized as described herein, then molded and attached to HDMC / AS and HDMC / CL, respectively, using custom-made silicone (Dow, Sylgard 184) molds (Baxter Healthcare, TISSEEL Fibrin Sealant, Cat. No. 1503152). The VB density was 18 AS or CL VBs per HDMC. HDMCs were sterilized by immersion in 70% ethanol solution for 30 minutes, followed by three washes with sterile Dulbecco's phosphate-buffered saline (DPBS; Hyclone, Cat. No. 350-000-CL). They were washed and stored in DPBS at 4°C.

[0479] (Cell culture): GFP-3T3 fibroblasts (producing green fluorescent protein) were cultured with HDMC / AS and HDMC / CL for 1 week (medium was changed twice at regular intervals) in 10% fetal bovine serum (FBS; Hyclone) and 1% penicillin-streptomycin. The HDs were approximately 5 mm long and 5 mm in diameter and cultured in Dulbecco's Modified Eagle's Medium (DMEM; Hyclone) supplemented with phospholipase C (P / S). Sections of MC / AS and HDMC / CL were deposited into wells of a 24-well plate, and approximately 50,000 GFP-3T3 cells were deposited (in small droplets of medium) directly onto the open channel surface of the HDMC. Four hours after initial cell deposition in the droplets (to allow cells to invade and attach to the HDMC), 1 ml of medium was added to the wells containing the HDMCs. Two medium changes were performed before the end of the one-week culture period. Samples were incubated under standard cell culture conditions in a humidified 37°C, 5% CO2 incubator.

[0480] (Fixation): At the end of the 1-week cell culture period, the medium was removed from the HDMC wells. The wells were washed three times with DPBS and then filled with 1 ml / well of 4% paraformaldehyde (PFA) fixative solution for 10 min, protected from light. At the end of the 10-min fixation period, the PFA solution was removed, and the wells were washed three times with DPBS, then filled with 1 ml / well of DPBS and stored at 4°C, protected from light. (Microscopy): HDMC samples were removed from the DPBS-containing wells of a 24-well plate and placed in a 5 mm plastic cell culture dish. The open channel edge (the edge where cells were deposited) was examined with a laser scanning confocal microscope. The surface was scanned using a 10x objective for areas where cells attached within and around the xylem channel opening. Both single optical sections and vertically stacked sections ("Z-stacks"; 63 sections spaced 2.8 microns apart) were captured at the region of interest. Optical sections were visualized using Fiji / ImageJ. 3D visualization of Z-stacks is a standard package in Fiji Rendered with a 3D viewer plugin.

[0481] result: The xylem channel structures of both HDMC / AS and HDMC / CL were directly visualized by autofluorescence (with a 405 nm excitation laser), and the presence of cells was indicated by a green fluorescent signal due to the presence of GFP in the cells (with a 488 nm excitation laser). Cells were visualized in and around the xylem channels of both HDMC / AS (Figures 36A and 36B) and HDMC / CL (Figures 37A and 37B). 3D reconstruction of optical sections revealed cell infiltration in and around the xylem channels (Figures 36C and 37C) and toward the interior of HDMC (Figures 36D and 37D).

[0482] Consideration: This cell culture study of the biocompatibility of HDMC / AS and HDMC / CL demonstrated that fibroblasts could attach and survive on the HDMC structures, confirming that HDMC is a biocompatible material. Fibroblasts were visualized both within and around the xylem microchannels, suggesting that the channel space of the AS and CL raw materials is suitable for cell infiltration and proliferation. Notably, 3D rendering of optical sections showed evidence of cell clusters growing deep into the HDMC material via microchannel-mediated guidance. This supports the idea that HDMC materials can serve as a substrate for directed cell growth both inside and outside of lignocellulosic microchannels. Therefore, HDMC can provide an engineered scaffold that promotes cell reconnection, for example, between previously severed links.

[0483] Figure 36A) shows a confocal laser scanning fluorescence micrograph (Z-stack projection) of xylem channels and fibroblasts within HDMC / AS. The channels are visualized in red (autofluorescence; false color), and GFP-3T3 fibroblasts are visualized in green (green fluorescent protein; false color). Figure 36B) shows a multichannel magnification of the red outline in panel A. Figure 36C) shows a 3D volume rendering (Fiji / ImageJ; 3D viewer plugin) of the green fluorescent protein and xylem autofluorescence signals from the perspective of panel B. D) Rotated view of the 3D rendering shows the orientation of the cells along the length of the channel.

[0484] Figure 37A) shows a confocal laser scanning fluorescence micrograph (Z-stack projection) of the xylem channel and fibroblasts in HDMC / CL. The channel is visualized in red (autofluorescence; false color), and GFP-3T3 fibroblasts are visualized in green (green fluorescent protein; false color). White arrows indicate the presence of cells outside the channel. Blue arrows indicate the presence of cells inside the channel. Figure 37B) shows a magnified view of a single channel outlined in red in panel A. Figure 37C) shows a 3D volume rendering (Fiji / ImageJ; 3D viewer plugin) of the green fluorescent protein signal from the perspective of panel B. Figure 3 7D) shows a rotated view of the 3D rendering, illustrating the orientation of the cells along the length of the channel.

[0485] Therefore, this study demonstrates that scaffold biomaterials containing HDMC bundles with decellularized microchannels from either AS or CL plant tissue sources can be used to induce cell guidance. These results demonstrate that cells were positioned both inside and outside the xylem channels of the HDMC bundles and were directional (i.e., guided by the directionality of the channels). [Example]

[0486] In vivo biocompatibility of scaffold biomaterials and HDMC bundles To further investigate the biocompatibility of the scaffold biomaterial and HDMC bundles as described herein, in vivo studies were performed.

[0487] method: Surgical procedure: Before surgical implantation, rats were subcutaneously injected with 0.9% saline and 0.05 mg / kg buprenorphine. Rats were anesthetized using isoflurane. The eyes were The right side of the rat was shaved from the hip to the shoulder. The skin was cleaned and sterilized with sterile solution. Three 1-2 cm incisions were made parallel to the anterior spine (near the lower, middle, and upper back). The incisions were made through the epidermis, dermis, and subcutaneous fat layer to the underlying muscle. HDMC bundle grafts (CL and AS) were implanted into each incision (one graft per incision). The incisions were sutured, and transdermal bupivacaine 2% was applied to the suture site. In addition, buprenorphine was administered subcutaneously 4-6 hours after the first injection. Specimens were collected at 4 and 8 weeks.

[0488] Histology: Collected samples were fixed in 10% formalin and placed in 70% ethanol before paraffin embedding. Serial sections were cut and stained with either hematoxylin and eosin (H&E) or Masson's trichrome.

[0489] Results and Discussion: Figures 38 and 39 show histological staining (hematoxylin and eosin; H&E) of longitudinal and transverse sections of HDMC / CL (celery-derived) material retrieved from the implantation site, respectively. In each case, panel B is an enlargement of the area in panel A. The CL-based microchannels (seen in Figure 39A) remain bundled together and are clearly visible.

[0490] Figures 40 and 41 show histological staining (H&E) of longitudinal and transverse sections of HDMC / AS (asparagus-derived) material retrieved from the implantation site, respectively. Although the AS-based microchannels (seen in Figure 40A) remained bundled, under the conditions tested, the boundaries between the microchannels were less distinct than those of HDMC / CL (Figure 39A).

[0491] A rough direct comparison of the histological staining of HDMC / CL (Figures 38 and 39) and HDMC / AS (Figures 40 and 41) suggests that the HDMC / AS material may be somewhat more biocompatible under the conditions tested. [Example]

[0492] Methods and procedures for liquid-based extraction of intact vascular bundle microchannels from native or decellularized vascular plants for HDMC generation Studies were conducted to determine alternative methods for isolating VB microchannels from native or decellularized vascular plants for HDMC production. Various different chemical treatments were used, including osmotic shock, temperature change, mechanical agitation, and maceration in acid and base solutions. The optimal solutions, concentrations, and times are determined depending on the goal of the procedure. The guiding principle for these steps is to break down the plant tissue into its underlying intact structures, which can be used alone or in combination with other plant materials.

[0493] method: Preparation of buffers and solutions 1. For NaCl, prepare 50 mL of 0 M, 0.5 M, 1 M, 2 M, and 3 M solutions. 2. For LiCl, prepare 50 mL of 0 M, 0.5 M, 1 M, 2 M, and 3 M solutions. 3. For acid / peroxide, prepare 50 mL solutions of glacial acetic acid to 30% hydrogen peroxide in ratios of 3:1, 2:1, 1:1, 1:2, and 3:1. 4. For base, prepare 50 mL of 0M, 0.5M, 1M, 2M, and 3M solutions of NaOH.

[0494] Preparation of plant tissues 5. Prepare strips of tissue by slicing the skin or removing the ends. For fruits such as apples or pears, peel and slice into equal pieces. For celery, asparagus, etc., cut off the white end and top. Cut into 6.3cm (or appropriate size) pieces and then slice using a mandoline slicer. 7. For plants such as asparagus and celery, leave the section containing the vascular bundles found in the xylem and phloem. For fruits such as apples and pears, use all of the inner flesh away from the core. 8. At this point, the native tissue can be treated with the conditions described below. Alternatively, the native tissue can be first decellularized according to the SDS-based methods of our previous work and patents before adding the treatments described below. 9. The remaining strip is placed in a beaker filled with the solution desired for isolating the microstructures.

[0495] Extraction procedure Materials needed: a. 50 mL tubes for various solutions and / or concentrations b. 200 mL beaker (use the exact same beaker throughout the procedure) c. Hot plate d. Thermometer e. Four 6-well plate lids (or any transparent rectangular plate lids for imaging) f. One 1 L beaker filled with distilled water (it is recommended to have another beaker nearby to hold 200 mL of distilled water to rapidly stop the reaction). g. Forceps h. Timer

[0496] Procedure: This procedure can be applied to any vascular plant tissue. Results are shown here using celery and asparagus. 1. Obtain 5 x 10CL strips from the first batch of each beaker. Take a photo of the strips on the table. Count the number of VBs in the strips (total number). 2. Prepare beakers with solutions for each treatment (e.g., 1 M NaCl = 5 beakers filled with 50 mL of 1 M NaCl each) - allow to come to room temperature or prepare solutions in advance. 3. Place the four beakers on a hot plate (for salt treatments, the hot plate may be kept on a bench, but for acid and base treatments, the entire procedure must be carried out in a fume hood). 4. Place 10 CL strips into each beaker and increase the temperature of the hotplate to 100 °C. 5. Once bubbles are visible, start a 30 minute (or interval) timer. Remove each beaker after 10, 15, 20, and 30 minutes, respectively. 6. Once the time is up, remove the beaker from the hot plate and add 200 mL of room temperature water (Caution: Do not add cold water to a hot beaker as the glass may shatter - add slowly) 7. Using a thermometer, carefully draw figure eights between the strands 30 times to gently separate any loose strands. (Note: Do not shear or separate strips that do not come loose easily.) 8. Pour off the solution - being careful not to lose any loose bundles, place the strips / removed bundles on the lid of the clear plate. Take a photo and note the number of bundles extracted (note if the strip separated easily or if the VB was clearly removed; note the % bundle extracted).

[0497] result: The effect of temperature and boiling time on the rate of liquid-based extraction of bundles from plant material is shown. The results below show that the parenchyma between the xylem and phloem bundles decomposes, allowing the sections to be cut into smaller pieces, and that the xylem and phloem bundles can be extracted over a prolonged period of treatment.

[0498] The experiments disclosed herein demonstrate that various liquid-based chemical treatments, such as salt (NaCl, LiCl), maceration (with acetic acid and peroxide), and base treatment (NaOH), can alter the mechanical and structural properties of plant materials and extract various parts of the plant, such as the vascular bundles. Conditions such as concentration and boiling time were evaluated for their ability to extract these materials and the effect of these solutions on the structural and mechanical properties of the materials.

[0499] Maceration is commonly used in the pulp and paper and food industries to soften and digest plant materials such as wood, bark, leaves, and fruit. In the food industry, fruits are often immersed in liquid to absorb the flavor of the liquid into the food, an example being sugar-coated strawberries to allow them to release their juice and soften. However, in both cases, these industrial processes cause ultrastructural degradation of the plant tissue, which is valuable for in vitro and in vivo tissue engineering applications.

[0500] Acid and Peroxide Treatment Results: Acid and peroxide can be used to both soften plant material and extract various parts of the plant to preserve microstructure. This was done with asparagus and celery, and the results are shown below. Figure 42 shows the results of extracting celery strips with acid and peroxide and boiling them to extract the vascular bundles contained in each strip. Here, boiling for 30 minutes with a 1:1 ratio of acid to 30% hydrogen peroxide results in the release of the greatest number of vascular bundles (Figures 43 and 44). It can be seen that ratios of 1:1, 1:2, and 1:3 were able to release bundles of vascular bundles from the decellularized strips. Similar results were obtained with salt and alkali treatments (discussed further below). Figure 44 shows asparagus strips treated with a 1:1 acid and peroxide solution of glacial acetic acid and 30% hydrogen peroxide. The approach described herein can be applied to vascular tissues of other plants to achieve the same extraction results. Figure 44 shows asparagus strips treated with an acid and peroxide solution of 1:1 glacial acetic acid:30% hydrogen peroxide.

[0501] Note that maceration at room temperature did not produce bundles, nor did boiling without a maceration solution. A combination of heating / boiling and maceration solution produces bundles. Furthermore, the same treatment can be applied to decellularized bundles with similar results, but with different boiling times and concentrations. Figure 43 shows previously decellularized celery subjected to the same conditions as the native strips of Figure 42. It can be noted that individual bundles could be extracted.

[0502] Salt Treatment Results: Salt treatment was also shown to be useful for extracting plant bundles. As with acid and peroxide, a range of salt concentrations and boiling times were tested to determine whether alternative solutions were more or less effective at extracting various plant parts and preserving their fine structure. Figures 45 and 46 show that longer boiling times were generally successful in extracting vascular bundles regardless of salt concentration, although the more concentrated solutions (3M and 1M) were shown to extract the greatest number of vascular bundles. The 2M solution was also shown to release a large number of bundles after 15 minutes, but this decreased after 20 and 30 minutes.

[0503] Results of alkaline treatment: In addition to acid treatment, various concentrations of NaOH base were used to test the removal of the bundles from the strips. The results showed that alkaline treatment significantly improved the removal of the bundles from the strips. It was clear that the strips were good candidates for separation. In this particular case, shorter boiling times (less than 15 minutes) at 0.5M appeared to result in the isolation of the greatest amount of vascular bundles (Figure 47). Higher concentrations, particularly 2M and 3M, did not allow the strips to be separated into individual vascular bundles. Optimal bundle extraction, yielding the greatest amount of individually isolated vascular bundles attached to or extracted cleanly from the surrounding parenchyma, appears to be around the 0.5-1M range, with boiling times between 10 and 20 minutes.

[0504] Vascular bundle microstructure: Vascular bundles were stained with 0.1% calcofluor white for 10 minutes. Cross-sections of HDMC were performed using a microtome blade in the PDMS mold. Images were acquired at 2.5x and 10x magnifications with an SZX16 stereomicroscope (1600x1200, ISO 100). Image processing was completed in Fiji (ImageJ). Raw images were converted to binary and then thresholded using adaptive thresholding techniques. Images were processed using noise reduction, speckle removal, and image segmentation functions. Images were then segmented for particle size analysis. SEM images of the microchannels revealed interconnected pores between individual channels. Because the pore size distribution is much smaller at the submicron scale, their structure is not the focus of this analysis.

[0505] Figure 48 shows the size distribution of CL vascular bundles obtained from native tissue immersed in boiling LiCl saline for 15 minutes. Similar microstructural distributions were observed with other treatments. In Figure 48A, vascular bundles were stained with 0.1% calcofluor white, and the image was thresholded and segmented for size analysis. Figure 48B shows a histogram of vascular bundle diameters. The average size was 23.7 ± 1.5 μm. Values ​​are the mean and standard error of the mean.

[0506] The images shown in Figures 49-63 summarize the morphology under several acid and peroxide (Figures 61-63), salt (Figures 49-56), and alkali (Figures 57-60) treatments. Tables 1-4 summarize the main observations and quantitative results. Tables 5-8 summarize the mechanical properties of the vascular bundles after treatment.

[0507] [Table 1]

[0508] [Table 2]

[0509] [Table 3]

[0510] [Table 4]

[0511] [Table 5]

[0512] [Table 6]

[0513] [Table 7]

[0514] [Table 8]

[0515] VB extracted using the procedure described in this example can be used to bundle HDMCs, as described herein. Figures 64 and 65 show HDMCs assembled from isolated CL vascular bundles boiled in water for 30 minutes (control condition) and in 1 M NaOH for 30 minutes. The NaOH bundles dehydrated rapidly and adhered quickly. In this example, cyanoacrylate adhesive was used to bundle the HDMCs. [Example]

[0516] Alternative methods and procedures for liquid-based extraction of intact vascular bundle microchannels from native or decellularized vascular plants for HDMC generation procedure: 1. I cut the celery into 2.5cm (1 inch) pieces and then sliced ​​them using a mandolin kitchen slicer. 2. Approximately 50-60 pieces were filled to the top with maceration solution (see Treatment Solutions 1-7 below). The sections were placed in a 1 L beaker until they were completely submerged. 3. The solution was brought to a boil (95-100°C) and mixed thoroughly every 5 minutes. 4. After boiling for 10 minutes, the strands began to separate from the rest of the strip. 5. After maceration was complete, the samples were removed from the maceration solution and washed several times with distilled water until the acetic acid and peroxide were removed.

[0517] After extended boiling, more bundles were observed to separate from the rest of the section, but the bundles were observed to soften due to the extended boiling time. Images of CL macerated in dilute acetic acid and peroxide by boiling for 15 minutes are shown in Figure 66.

[0518] Additionally, various solutions were tested for their potential as maceration products. These included: Treatment Solution 1: 1:1 glacial acetic acid and 30% hydrogen peroxide Treatment Solution 2: 1:1 glacial acetic acid and peroxide (as above), diluted to 50% with water. Treatment Solution 3: Glacial Acetic Acid Treatment solution 4: 50% diluted acetic acid Treatment solution 5: 95% ethanol Treatment Solution 6: 3 molar sodium chloride (NaCl) Treatment Solution 7: 4 molar hydrochloric acid (HCl)

[0519] Of these treatments, treatment solutions 1, 2, 4, 6, and 7 were able to remove the bundles, but the concentrated hydrochloric acid was too harsh as it broke down the bundles and shredded them into individual pieces beyond the point of use, and was therefore diluted two-fold for subsequent testing.

[0520] It was also found that celery and asparagus did not require slicing or other prior preparation of the samples before adding them to the maceration solution. For these two, considerable mixing was performed to remove the bundles from the pieces once the samples were removed from the maceration solution. The celery and asparagus were soft enough that the surrounding tissue could be squeezed out of the bundles (much easier with celery than with asparagus). The long fibers were removed from the solution and placed in a new beaker containing distilled water. The bundles were then washed by vigorously mixing and manually crushing the sample, while avoiding breaking the bundles. Each time the samples were mixed and washed, more and more surrounding tissue was removed (Figure 67).

[0521] As will be appreciated, a decellularization step (such as SDS-based decellularization as described herein) can be performed before or after the liquid extraction / maceration step, if desired.

[0522] One or more exemplary embodiments have been described by way of example, and those skilled in the art will appreciate that certain changes and modifications can be made without departing from the scope of the invention as defined in the claims.

Claims

1. A method for supporting cell growth, comprising the steps of: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and introducing one or more cells into the scaffold biomaterial;

2. A method for isolating and decellularizing microchannels from plant or fungal tissue, comprising: isolating a microchannel from the plant or fungal tissue; and decellularizing the microchannel; optionally, sterilizing the microchannel; The method.

3. The method of claim 2, wherein the step of isolating the microchannels comprises mechanical separation of the microchannels or the vascular bundles containing the microchannels, or both, from surrounding plant or fungal tissue, the separating step being carried out by gentle peeling or cutting, and / or wherein the acid and peroxide extraction comprises heating the plant or fungal tissue in an acid and peroxide solution.

4. The method of claim 2, wherein the step of isolating the microchannels comprises mechanical separation of the microchannels or the vascular bundles containing the microchannels, or both, from surrounding plant or fungal tissue, and wherein the step of isolating is carried out by liquid-based extraction of the microchannels from the plant or fungal tissue, and / or the liquid-based extraction comprises at least one of acid extraction, acid and peroxide extraction, salt extraction, or alkaline extraction, and wherein the acid and peroxide solution comprises glacial acetic acid and 30% hydrogen peroxide in a ratio of 3:1 to 1:3, and the acid and peroxide solution comprises a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v).

5. A method according to any one of claims 2 to 4, wherein the heating step comprises heating for up to 30 minutes or comprises heating for 30 minutes, wherein the heating step comprises boiling the acid and peroxide solution and / or wherein the acid and peroxide solution comprises a 1:1 glacial acetic acid:hydrogen peroxide solution (30% v / v) and the solution is heated to boiling for 30 minutes, and wherein the liquid-based extraction further comprises the step of mechanically agitating, e.g. stirring, the plant or fungal tissue in the acid and peroxide solution.

6. The method of claim 4, wherein the salt extraction comprises heating the plant or fungal tissue in a salt solution, and / or wherein the salt solution comprises LiCl or NaCl, the salt solution has a salt concentration of about 0.5M to 3M, the salt solution comprises LiCl or NaCl at a salt concentration of about 3M, and the heating comprises heating for up to 30 minutes or heating for 30 minutes.

7. The method of claim 6, wherein the heating step includes boiling the salt solution, and / or wherein the salt solution comprises about 3 M LiCl and the solution is heated to boiling for 30 minutes, or wherein the salt solution comprises about 3 M NaCl and the solution is heated to boiling for 15 minutes, and wherein the liquid-based extraction further includes mechanically agitating, e.g., stirring, the plant or fungal tissue in the salt solution.

8. The method of claim 4, wherein the alkaline extraction comprises heating the plant or fungal tissue in an alkaline solution, and / or wherein the alkaline solution has an alkaline concentration of about 0.5 to 3 M, the alkaline solution comprises sodium hydroxide (NaOH), the alkaline solution comprises sodium hydroxide at an alkaline concentration of about 0.5 M to 1 M, and the heating comprises heating for up to 30 minutes or heating for 30 minutes.

9. The method of claim 8, wherein the heating step comprises boiling the alkaline solution and / or the alkaline solution comprises about 0.5 M NaOH and the solution is heated to boiling for 5 minutes, preferably the liquid-based extraction further comprising a step of mechanically agitating, e.g. stirring, the plant or fungal tissue in the alkaline solution.

10. The method of claim 4, wherein the acid extraction comprises heating the plant or fungal tissue in an acid solution, and / or wherein the acid solution comprises acetic acid or hydrochloric acid, wherein the acid solution comprises 50% acetic acid, and wherein the heating comprises heating for up to 30 minutes or heating for 30 minutes, and wherein the heating comprises boiling the acid solution, wherein the acid solution comprises 50% acetic acid and the acid solution is heated to boiling for 30 minutes, and wherein the liquid-based extraction further comprises mechanically agitating, e.g., stirring, the plant or fungal tissue in the acid solution, and wherein the decellularizing step occurs before the separating step.

11. A method according to any one of claims 2 to 10, further comprising the step of introducing living plant or animal cells into the microchannel.

12. The method of claim 11, wherein the living cells are mammalian cells or human cells.

13. A decellularized microchannel produced by a method according to any one of claims 2 to 12.

14. Use of a scaffold biomaterial comprising at least one bundle of microchannels for promoting tissue regeneration, promoting angiogenesis, treating and / or repairing spinal cord injuries, or repairing or reconstructing or replacing plant or animal tissue, wherein the bundle of microchannels comprises a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by adhesion, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by fixing the microchannels.

15. A method for filtering and / or separating a solution or material, comprising the steps of: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and passing a solution or material through the scaffold biomaterial to filter and / or separate components from the solution based on size exclusion;

16. A method for modifying or regulating the growth of a plant, comprising the steps of: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and implanting the scaffold biomaterial into the plant to provide accelerated growth;

17. A method for material transport, comprising the steps of: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and using the scaffold biomaterial to deliver a material, such as a drug, to a site in need thereof; 18. A method for preparing a structure that mimics a desired shape or object, comprising: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and shaping or positioning the scaffold biomaterial to mimic a desired shape or object; optionally, adhering the scaffold biomaterial for structural reinforcement; The method.

19. A method for exchanging or transferring heat in a microfluidics process, comprising the steps of: providing a scaffold biomaterial comprising at least one microchannel bundle, the microchannel bundle comprising a plurality of decellularized microchannels isolated from plant or fungal tissue, the decellularized microchannels being arranged substantially parallel to one another within the bundle, and the plurality of decellularized microchannels being bundled together by gluing, by using a sealant, by mechanical filling, by mechanical entanglement, by crosslinking, by suspending the microchannels in a hydrogel, by threading the microchannels around one another, or by compressing / filling the microchannels, or by immobilizing the microchannels; and using the microchannels of said scaffold biomaterial to carry one or more fluids, said one or more fluids being in close proximity to allow for heat exchange or heat transfer;

20. A kit comprising one or more of the following: Type; glue; one or more microchannels; one or more decellularizing agents; Scalpel or microtome; Sterile measuring devices; Sterile saline; tweezers; and / or Instructions for carrying out the method according to any one of claims 1 to 10 and 15 to 19;

Images