Surgical mesh implant

Abstract

The present invention provides a surgical mesh implant comprising bacterial nanocellulose and methods of producing the surgical mesh implant.

Description

[0001] TITLE

[0002] SURGICAL MESH IMPLANT

[0003] CROSS REFERENCE TO RELATED APPLICATIONS The present application claims priority to U.S. Provisional Patent Application No. 63 / 605,724, filed December 4, 2023, the disclosure of which is incorporated by reference herein in its entirety.

[0004] STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

[0005] This invention was made with government support under Grant Nos. MCBPT T32, awarded by the National Institutes of Health, and AR065972, awarded by the National Institutes of Health. The government has certain rights in the invention.

[0006] BACKGROUND OF THE INVENTION

[0007] Hernia is defined as a pathological protrusion of tissue through a weakness, laxity or patency in a cavity wall or structure. Inguinal, femoral and ventral abdominal hernias are characterized by bowel protrusion through soft tissues and are detectable as a palpable pouch or bulge under the skin. Protruded tissues may be strangulated or incarcerated from hampered perfusion causing discomfort, pain, inability to perform daily activities, and even infection, ischemia, abscess formation, or necrosis, with severe cases necessitating emergency surgery.

[0008] Tension-free anatomical repair using mesh is regarded in many clinical settings as a superior option to direct suturing and tissue reanastomosis, particularly when reinforcing weak abdominal fascia in the setting of hernia surgery and repair. The surgical mesh for hernia was implemented to reduce recurrence rates, which remained high (30 - 40%) when suturing the defect site under tension. This method also was associated with increased morbidity as recurrent herniation was typically more severe, prone to infection, and often necessitated administration of steroids. Thus, the introduction of surgical mesh is an engineering solution for primary suturing of hernia repair. The gold standard mesh material for repair is a polypropylene-based microporous knitted thermoplastic fabric-like patch. However, polypropylene mesh often suffers complications upon implantation from poor biocompatibility, material shrinkage, and a propensity for scar tissue and bowel adhesion formation. Moreover, the implant must be able to withstand significant stresses associated with reinforcement of the abdominal wall and must possess anti-microbial activity to reduce the risk of postoperative infection. Mesh with pores too small allow for ingress of bacterial species but sterically blunt or hinder immune-mediated infiltration and surveillance of the implant. Mesh designs with submicron pores are particularly vulnerable and typically do not respond well to antibiotics once a viable biofilm is established. Clinical management in these cases often necessitates hospital readmission, re-operation, mesh explanation, and local debridement. Other approaches have developed hybrid or composite meshes with differential modification of polypropylene with various biopolymers, primarily through electrospinning. These mats tend to lack robust durability compared to nonhybrid counterparts. Durability is a key function considering that the implant must be able to withstand vigorous movement patterns and fluid flow upon implantation. Further, incorporating multiple techniques and materials raises processing complexity. Multiple layers may also increase the risk of mechanical imbalances, increase the mass of the implant, and raise the risk of foreign body reactions. Given these considerations, a simple, reproducible, versatile material and fabrication method is of interest for strategic improvement of mesh biomaterial capabilities, especially when interfacing with multiple in vivo tissues is desired.

[0009] Thus, there is a need in the art for an improved surgical mesh material with versatile physical and mechanical properties, while also meeting the rigorous requirements necessary for clinical use. The present invention meets this need.

[0010] SUMMARY OF THE INVENTION

[0011] In one aspect, the present invention relates to a method of preparing a bacterial nanocellulose surgical mesh, comprising the steps of: a) culturing one or more bacteria that produce cellulose; b) exposing the culture to an electric field; c) harvesting the resulting pellicle; and d) drying the pellicle. In some embodiments, step c) further comprises a step of washing and decellularizing the pellicle. In some embodiments, the step of washing and decellularizing the pellicle comprises washing the pellicle with an alkali solution. In some embodiments, the alkali solution is sodium hydroxide, lithium hydroxide, potassium hydroxide, or cesium hydroxide at a concentration of between 0.5 M and 2 M.

[0012] In some embodiments, the method of drying in step d) is accomplished by thermal evaporation or freeze-drying. In some embodiments, the method of thermal evaporation comprises oven-drying. In some embodiments, the method of freeze-drying comprises liquid nitrogen lyophilization.

[0013] In some embodiments, the bacterium of step a) is selected from a group consisting of Komagataeibacter , Acetobacter xylinum, Acetobacter, Achromobacter, Bacillus, Sarcina, Aerobacter, Agrobacterium, Escherichia, Azotobacter, Rhizobium, Enterobacter, Klebsiella, and Salmonella n some embodiments, the bacterium is Acetobacter xylinum.

[0014] In some embodiments, the electric field of step b) is a constant electric field. In some embodiments, the electric field is at a potential of between about 1 V and about 20 V. In some embodiments, the electric field is at a potential of 10 V.

[0015] In another aspect, the present invention relates to a surgical mesh implant comprising one or more layers of bacterial nanocellulose, wherein each layer has a parietal side and a visceral side, wherein the layers are stacked on top of each other such that the parietal side of one is in contact with the visceral side of the next layer, and wherein the surgical mesh implant comprises a plurality of pores from the parietal side to the visceral side of the surgical mesh implant.

[0016] In some embodiments, the surgical mesh implant has a Young’s modulus of between about 0.1 MPa to about 1.4 MPa. In some embodiments, the surgical mesh implant has a Young’s modulus of about 1.4 MPa.

[0017] In some embodiments, the surgical mesh implant has an ultimate tensile strength of between about 0.1 MPa to about 85 MPa. In some embodiments, the surgical mesh implant has an ultimate tensile strength of about 85 MPa.

[0018] In some embodiments, the surgical mesh implant has a pore size of between about 0.1 pm to about 1.5 pm. In some embodiments, the pore size on the parietal side of the implant is larger than the pore size on the visceral side.

[0019] In some embodiments, the surgical mesh implant has a thickness of between about 1 mm to about 5 mm.

[0020] In some embodiments, the surgical mesh implant is a circle. In some embodiments, the surgical mesh implant has a diameter of about 1 mm to about 100 mm.

[0021] In some embodiments, the surgical mesh implant degrades less than 10% when the implant is exposed to collagenase type II for 5 days.

[0022] In some embodiments, the surgical mesh implant is impregnated with an antibiotic.

[0023] BRIEF DESCRIPTION OF THE DRAWINGS

[0024] The following detailed description of embodiments of the invention will be better understood when read in conjunction with the appended drawings. It should be understood, however, that the invention is not limited to the precise arrangements and instrumentalities of the embodiments shown in the drawings.

[0025] Fig. 1 depicts a flow diagram of an exemplary method of the present invention.

[0026] Fig. 2 depicts a schematic representation of the method of the present invention.

[0027] Fig. 3 depicts a schematic representation of the electric stimulation process and resulting fiber alignment.

[0028] Fig. 4, comprising Figs. 4A through 4D, depicts representative bacterial nanocellulose culture media and pellicle washing. Fig. 4A depicts a representative image of bacterial cell culture media. Fig. 4B depicts a representative bacterial nanocellulose mesh sample after the culturing process. Fig. 4C depicts a representative bacterial nanocellulose mesh sample after the decellularization wash. Fig. 4D depicts a representative sample of the bacterial nanocellulose mesh after several weeks in storage.

[0029] Fig. 5, comprising Figs. 5A through 5E, depicts various images of representative bacterial nanocellulose pellicles. Fig. 5A depicts a representative bacterial nanocellulose mesh sample after liquid nitrogen freezing and lyophilization. Fig. 5B depicts a representative bacterial nanocellulose mesh sample after oven drying. Fig. 5C depicts a representative scanning electron microscopy image of liquid nitrogen freeze- dried bacterial nanocellulose. Fig. 5D depicts a representative scanning electron microscopy image of oven-dried bacterial nanocellulose. Fig. 5E depicts representative liquid nitrogen freeze-dried and oven-dried bacterial nanocellulose samples prior to gold sputtering and scanning electron microscopy analysis.

[0030] Fig. 6, comprising Figs. 6A through 6F, depicts representative scanning electron microscopy images of various bacterial nanocellulose pellicles and their properties. Fig. 6A and Fig. 6B depict representative bacterial nanocellulose pellicles under varied drying conditions. Fig. 6C and Fig. 6D depict representative scanning electron microscopy images of control, unstimulated bacterial nanocellulose treated with oven drying and liquid nitrogen freeze drying. Fig. 6E depicts representative results of hydroexpansivity test of bacterial nanocellulose. Fig. 6F depicts representative results of the quantification of pore sizes of oven-dried versus liquid-nitrogen freeze-dried samples.

[0031] Fig. 7, comprising Figs. 7A through 7F, depicts representative images of bacterial nanocellulose after electrical stimulation protocol. Fig. 7A depicts representative scanning electron microscopy images of bacterial nanocellulose under control conditions. Fig. 7B depicts representative scanning electron microscope images of bacterial nanocellulose under electric stimulation. Fig. 7C and Fig. 7D depict histograms of the representative results of fiber orientation quantification. Fig. 7E and Fig. 7F depict rose plots of the representative results of fiber orientation quantification.

[0032] Fig. 8 depicts representative results of a degradation assessment of bacterial nanocellulose after one month incubation.

[0033] Fig. 9, comprising Figs. 9A through 9B, depicts representative results of a degradation assessment. Fig. 9A depicts a representative graph of mass retention of bacterial nanocellulose samples exposed to cellulase and collagenase type II. Fig. 9B depicts a representative graph of mass retention of 20% gelatin methacryloyl hydrogen samples exposed to cellulase and collagenase.

[0034] Fig. 10, comprising Figs. 10A through 10D, depicts representative fluorescence microscopy images of human skeletal myoblasts culture. Fig. 10A and Fig.1 OB depict representative human skeletal myoblast culture in bacterial nanocellulose showing 3D multi-plane patterning. Fig. IOC and Fig. 10D depict representative human skeletal myoblast culture in 2D controls.

[0035] Fig. 11, comprising Figs. 11 A through 1 IF, depicts representative results of the evaluation of human skeletal myofiber alignment. Fig. 11 A depicts a representative fluorescence microscopy image of unstimulated bacterial nanocellulose. Fig. 11B depicts a representative fluorescence microscopy image of electrically stimulated bacterial nanocellulose. Fig. 11C and Fig. 1 ID depict representative histograms of human skeletal myofiber alignment in unstimulated versus electrically stimulated bacterial nanocellulose. Fig. 1 IE and Fig. 1 IF depict representative rose plots of human skeletal myofiber alignment in unstimulated versus electrically stimulated bacterial nanocellulose.

[0036] Fig. 12, comprising Fig. 12A through 12C, depicts representative fluorescence microscopy images of human skeletal myoblast staining in bacterial nanocellulose. Fig. 12A depicts representative human skeletal myoblast staining in liquid nitrogen unaligned freeze-dried bacterial nanocellulose. Fig. 12B depicts representative human skeletal myoblast staining in unaligned oven-dried bacterial nanocellulose. Fig. 12C depicts representative human skeletal myoblast staining in electrically stimulated bacterial nanocellulose.

[0037] Fig. 13, comprising Figs. 13A through 13B, depicts representative images of bacterial nanocellulose suturing. Fig. 13 A depicts representative images of bacterial nanocellulose suturing. Fig. 13B depicts representative images of bacterial nanocellulose mesh sutured to resected porcine hindlimb muscle.

[0038] Fig. 14, comprising Figs. 14A through 14F, depicts representative images of bacterial nanocellulose post-processing after pellicle harvest. Fig. 14A depicts a representative image of a washed and decellularized bacterial nanocellulose pellicle sample. Fig. 14B and Fig. 14C depict representative images of liquid nitrogen freeze- dried bacterial nanocellulose pellicle samples. Fig. 14D and Fig. 14E depict representative images of oven-dried bacterial nanocellulose pellicle samples. Fig. 14F depicts representative image of a sample cut into a disk for cell culture with human skeletal muscle myoblasts.

[0039] Fig. 15 depicts representative results of a water-retention capacity assessment of bacterial nanocellulose pellicle samples. Fig. 16, comprising Figs. 16A to 16H, depicts representative results of mechanical testing. Fig. 16A depicts a representative experimental set-up. Fig. 16B, Fig. 16C, and Fig. 16D depict representative stress versus strain graphs of oven-dried bacterial nanocellulose samples and commercial samples. Fig. 16E depicts representative images of porcine hindlimb muscle alone or sutured to bacterial nanocellulose for mechanical testing. Fig. 16F depicts representative results of strength testing for porcine hindlimb muscle alone or sutured to bacterial nanocellulose. Fig. 16G depicts a schematic representation of porcine hindlimb muscle sutured to bacterial nanocellulose. Fig. 16H depicts representative results of the ultimate strength test of a liquid nitrogen freeze-dried bacterial nanocellulose sample.

[0040] Fig. 17, comprising Figs. 17A through 17B, depicts representative images of suture strength testing of commercial mesh samples before (Fig. 17A) and after (Fig. 17B) testing.

[0041] Fig. 18, comprising Figs. 18A through 18C, depicts representative results of mesh degradation testing of bacterial nanocellulose samples versus commercial mesh samples. Fig. 18A depicts representative results of mesh degradation of bacterial nanocellulose and commercial mesh in media. Fig. 18B depicts representative results of mesh degradation of bacterial nanocellulose and commercial mesh in media with collagenase type II. Figure 3 depicts representative images of bacterial nanocellulose and commercial mesh degradation.

[0042] Fig. 19, comprising Figs 19A through 19D, depicts representative fluorescence microscopy images of human skeletal muscle myoblasts staining. Fig. 19A and Fig. 19B depict representative images of myoblasts cultured in unstimulated bacterial nanocellulose. Fig. 19C depicts a representative image of myoblasts cultured in a 2- dimensional traditional culture plate. Fig. 19D depicts a representative image of human myoblasts cultured in gelatin methacryloyl.

[0043] Fig. 20, comprising Figs. 20A and 20B depicts representative results comparing human skeletal muscle myofiber alignment in gelatin methacryloyl versus commercial mesh samples. Fig. 20A depicts representative results of myoblasts cultured in gelatin methacryloyl. Fig. 20B depicts representative results of myoblasts cultured in a commercial mesh sample. Fig. 21, comprising Figs. 21 A through 21D, depicts representative confocal imaging of human skeletal muscle myoblasts cultured in bacterial nanocellulose versus commercial samples. Fig. 21A depicts a representative image of myoblasts cultured in bacterial nanocellulose. Fig. 2 IB and Fig. 21C depict representative images of myoblasts cultured in commercial mesh samples. Fig. 2 ID depicts a representative image of myoblasts cultured in gelatin methacryloyl.

[0044] Fig. 22, comprising Figs. 22A through 22E, depicts representative images of bacterial nanocellulose hydrogel preparation for tissue culture. Fig. 22A and Fig. 22B depict representative images of a 6 mm punched-out portions from washed, decellularized and sterilized bacterial nanocellulose pellicle. Fig. 22C depicts a representative image of punched portions from liquid nitrogen freeze-dried bacterial nanocellulose pellicle. Fig. 22D depicts a representative image of punched portions from oven dried bacterial nanocellulose pellicle. Fig. 22E depicts a stylized representation of seeded bacterial nanocellulose mesh samples with human skeletal muscle myoblasts.

[0045] Fig. 23, comprising Figs. 23A through 23C, depicts representative images and orientation assessment results of electrically stimulated bacterial nanocellulose fibrils. Fig. 23A depicts a representative scanning electron microscopy image of aligned nanocellulose fibrils. Fig. 23B depicts a representative histogram showing the orientation of nanocellulose fibrils. Fig. 23C depicts a representative rose plot showing the orientation of nanocellulose fibrils.

[0046] Fig. 24, comprising Figs, 24A through 24G, depicts representative scanning electron microscopy images of bacterial nanocellulose versus commercial mesh samples. Fig. 24A and Fig. 24D depict representative scanning electron microscopy images of liquid nitrogen freeze-dried bacterial nanocellulose. Figs. 24B and 24E depict representative scanning electron microscopy images of commercial Marigen™ samples. Fig. 24C and 24F depict representative scanning electron microscopy images of commercial Phoenix™ mesh samples. Fig 24G depicts a representative side-by-side image of bacterial nanocellulose mesh versus commercial mesh.

[0047] Fig. 25, comprising Figs. 25A through 25C, depicts representative results from myofiber maturation assessments in bacterial nanocellulose versus commercial meshes and gelatin methacryloyl. Fig. 25 A depicts a representative fusion index and nuclei counting for bacterial nanocellulose. Fig. 25B and Fig. 25C depict representative fusion indices and nuclei counting for bacterial nanocellulose, commercial Marigen™, commercial Phoenix™, and gelatin methacryloyl.

[0048] Fig. 26, comprising Figs. 26A through 26H, depicts representative results of antimicrobial activity assessments in bacterial nanocellulose versus commercial meshes. Fig. 26A depicts representative methicillin-resistant Staphylococcus aureus (MRSA) growth with various meshes treated with an antibiotic. Fig. 26B depicts representative MRSA growth with various meshes treated with a polyelectrolyte. Fig. 26C depicts representative Pseudomonas aeruginosa growth with various meshes treated with an antibiotic. Fig. 26D depicts representative Pseudomonas aeruginosa growth with various meshes treated with a polyelectrolyte. Fig. 26E depicts representative MRSA growth with various meshes treated with an antibiotic or polyelectrolyte after 1 day. Fig. 26F depicts representative MRSA growth with various meshes treated with an antibiotic or polyelectrolyte after 5 days. Fig. 26G depicts representative Pseudomonas aeruginosa growth with various meshes treated with an antibiotic or polyelectrolyte after 1 day. Fig. 26H depicts representative Pseudomonas aeruginosa growth with various meshes treated with an antibiotic or polyelectrolyte after 5 days.

[0049] Fig. 27, comprising Figs. 27A through 27C, depicts a representative finite element model of pillar deflection in response to boundary load. Fig. 27A and Fig. 27B depict representative models of pillar geometry. Fig. 27C depicts a representative model of the deflection of flexible pillars in response to a load.

[0050] Fig. 28, comprising Figs. 28A through 28F, depicts a representative hydrogel curing chamber for in vitro myoblast maturation platform. Fig. 28A and Fig. 28B depict a representative computer-assisted design of a hydrogel curing chamber. Fig. 28C depicts a representative stereolithography printed hydrogel curing chamber. Fig. 28D depicts a representative computer-assisted design of a hydrogel curing chamber. Fig. 28E and Fig. 28F depict representative stereolithography printed hydrogel curing chamber.

[0051] Fig. 29, comprising Figs. 29A through 29D, depicts representative models of pillar geometry. Fig. 29A, Fig. 29B and Fig. 29C depict simulated models. Fig. 29D depicts representative stereolithography prints of models. Fig. 30 depicts representative results of finite element comparison of maximum flexural capacity as a function of pillar material and geometry.

[0052] Fig. 31, comprising Figs. 31 A through 3 ID, depicts representative images of hydrogel curing under tension between flexible pillars. Fig. 31A depicts a representative image of von Mises Stress forces acting upon flexible pillars. Fig, 3 IB depicts a computer-assisted design of hydrogen curing. Fig. 32C depicts a representative image of hydrogels in standard 12-well plate. Fig. 3 ID depicts a simulated model of the volume displacement of the earbud structure.

[0053] Fig. 32, comprising Figs. 32A through Fig. 32E, depict representative images of earbud pillar geometries. Fig. 32A, Fig. 32B, and Fig. 32C depict representative computer-assisted designs of earbud pillar geometries. Fig. 32D and Fig. 32E depict representative images of custom pillar geometries printed by stereolithography .

[0054] Fig. 33, comprising Figs. 33A through 33C, depicts a representative computer-assisted design of well-plates for an electrical stimulation bioreactor. Fig. 33A depicts a representative computer-assisted design of custom well-plates. Fig. 33B depicts a representative computer-assisted design of custom well-platter stopper. Fig. 33C depicts a representative computer-assisted design of a custom well-plate showing its measurements.

[0055] Fig. 34, comprising Figs. 34A through 34D, depicts representative images of stereolithography printed electrical stimulation platform. Fig. 34A depicts a representative computer-assisted design of an electrical stimulation platform. Fig. 34B and Fig. 34C depict representative images of a stereolithography printed electrical stimulation platform. Fig. 34D depicts a schematic representation of an electrical stimulation process.

[0056] Fig. 35, comprising Figs. 35A though 35D, depicts representative models of bacterial nanocellulose mesh deformation. Fig. 35 A and Fig. 25B depict representative models of bacterial nanocellulose mesh deformation in response to physiologic intraabdominal pressure. Fig. 35D depicts a representative photographic demonstration of bacterial nanocellulose mesh deformation in response to manipulation and stress forces. DETAILED DESCRIPTION

[0057] It is to be understood that the figures and descriptions of the present invention have been simplified to illustrate elements that are relevant for a clear understanding of the present invention, while eliminating, for the purpose of clarity many other elements found in related systems and methods. Those of ordinary skill in the art may recognize that other elements and / or steps are desirable and / or required in implementing the present invention. However, because such elements and steps are well known in the art, and because they do not facilitate a better understanding of the present invention, a discussion of such elements and steps is not provided herein. The disclosure herein is directed to all such variations and modifications to such elements and methods known to those skilled in the art.

[0058] Definitions

[0059] Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which the invention pertains. Although any methods and materials similar or equivalent to those described herein can be used in the practice for testing of the present invention, exemplary materials and methods are described herein. In describing and claiming the present invention, the following terminology will be used.

[0060] It is also to be understood that the terminology used herein is for the purpose of describing particular embodiments only, and is not intended to be limiting.

[0061] The articles “a” and “an” are used herein to refer to one or to more than one (i.e., to at least one) of the grammatical object of the article. By way of example, “an element” means one element or more than one element.

[0062] “About” as used herein when referring to a measurable value such as an amount, a temporal duration, and the like, is meant to encompass variations of ±20%, ±10%, ±5%, ±1%, or ±0.1% from the specified value, as such variations are appropriate.

[0063] As used herein, “biocompatible” refers to any material, which when implanted in a mammal, does not provoke an adverse response in the mammal. A biocompatible material, when introduced into an individual, is not toxic or injurious to that individual, nor does it induce immunological rejection of the material in the mammal.

[0064] As used herein, a “culture” refers to the cultivation or growth of cells, for example, tissue cells, in or on a nutrient medium. As is well known to those of skill in the art of cell or tissue culture, a cell culture is generally begun by removing cells or tissue from a human or other animal, dissociating the cells by treating them with an enzyme, and spreading a suspension of the resulting cells out on a flat surface, such as the bottom of a Petri dish. There, the cells generally form a thin layer of cells called a “monolayer” by producing glycoprotein-like material that causes the cells to adhere to the plastic or glass of the Petri dish. A layer of culture medium, containing nutrients suitable for cell growth, is then placed on top of the monolayer, and the culture is incubated to promote the growth of the cells.

[0065] As used herein, “inoculum” refers to a small amount of material containing bacteria, viruses, or other microorganisms that is used to start a culture.

[0066] As used herein, “pellicle” refers to a thick floating biological membrane formed at the liquid-air interface of a cell culture.

[0067] As used herein, “scaffold” refers to a structure, comprising a biocompatible material that provides a surface suitable for adherence and proliferation of cells. A scaffold may further provide mechanical stability and support. A scaffold may be in a particular shape or form so as to influence or delimit a three-dimensional shape or form assumed by a population of proliferating cells. Such shapes or forms include, but are not limited to, films (e.g., a form with two dimensions substantially greater than the third dimension), ribbons, cords, sheets, flat discs, cylinders, spheres, 3-dimensional amorphous shapes, etc.

[0068] As used herein, “tissue engineering” refers to the process of generating a tissue ex vivo for use in tissue replacement or reconstruction. Tissue engineering is an example of “regenerative medicine,” which encompasses approaches to the repair or replacement of tissues and organs by incorporation of cells, gene, or other biological building blocks, along with bioengineered materials and technologies.

[0069] The terms “patient,” “subject,” “individual,” and the like are used interchangeably herein, and refer to any animal amenable to the systems, devices, and methods described herein. The patient, subject or individual may be a mammal, and in some instances, a human.

[0070] Ranges: throughout this disclosure, various aspects of the invention can be presented in a range format. It should be understood that the description in range format is merely for convenience and brevity and should not be construed as an inflexible limitation on the scope of the invention. Accordingly, the description of a range should be considered to have specifically disclosed all the possible subranges as well as individual numerical values within that range. For example, description of a range such as from 1 to 6 should be considered to have specifically disclosed subranges such as from 1 to 3, from 1 to 4, from 1 to 5, from 2 to 4, from 2 to 6, from 3 to 6 etc., as well as individual numbers within that range, for example, 1, 2, 2.7, 3, 4, 5, 5.3, and 6. This applies regardless of the breadth of the range.

[0071] Description

[0072] The present invention relates in part to a bacterial nanocellulose permanent surgical mesh implant and methods of making them. In surgical repair settings, mesh materials are applied to a defect site for tissue reinforcement and tension- free repair. Bacterial nanocellulose as a material for surgical meshes is of high interest because of its high purity and tunable mechanical and physical properties. Bacterial nanocellulose surgical meshes are biocompatible, flexible, strong, durable and cost- effective surgical meshes, and are a strong candidate to replace conventional surgical meshes for tension-free tissue repair.

[0073] The present invention is based in part on the unique properties of bacterial nanocellulose (BNC) when prepared by the methods described herein. BNC is of high purity and exhibits intrinsic and modifiable properties that are of high interest as a cost- effective biomaterial. As such, in one aspect, the present invention relates, in part, to a method of preparing a bacterial nanocellulose surgical mesh, comprising the steps of: a) culturing one or more bacteria that produce cellulose; b) exposing the culture to an electric field; c) harvesting the resulting pellicle; and d) drying the pellicle.

[0074] In another aspect, the present invention provides a bacterial nanocellulose surgical mesh implant formed using the methods of the present invention. Methods of making a BNC mesh

[0075] In one aspect, the present invention relates to methods of preparing a bacterial nanocellulose surgical mesh. In various embodiments, the method comprises the steps of: a) culturing one or more bacteria that produce cellulose; b) exposing the culture to an electric field; c) harvesting the resulting pellicle; and d) drying the pellicle.

[0076] In some embodiments, step a) comprises culturing one or more bacteria that produce cellulose. In some embodiments, the bacteria produce nanocellulose. In some embodiments, the bacteria are one or more bacteria of one or more genera selected from the group consisting of Komagataeibacter, Acetobacter xylinum, Acetobacter, Achromobacter, Bacillus, Sarcina, Aerobacter, Agrobacterium, Escherichia, Azotobacter, Rhizobium, Enterobacter, Klebsiella, and Salmonella. In some embodiments, the bacteria is Acetobacter xylinum.

[0077] In some embodiments, step a) of the method comprises culturing the selected bacteria for at least 1 day. In some embodiments, the method comprises culturing the selected bacteria for at least 2 days. In some embodiments, the method comprises culturing the selected bacteria for at least 3 days. In some embodiments, the method comprises culturing the selected bacteria for at least 4 days. In some embodiments, the method comprises culturing the selected bacteria for at least 5 days. In some embodiments, the method comprises culturing the selected bacteria for at least 6 days. In some embodiments, the method comprises culturing the selected bacteria for at least 1 week. In some embodiments, the method comprises culturing the selected bacteria for at least 2 weeks. In some embodiments, the method comprises culturing the selected bacteria for at least 3 weeks. In some embodiments, the method comprises culturing the selected bacteria for at least 1 month.

[0078] In some embodiments, step a) of the method comprises culturing the selected bacteria for between 1 and 40 days. In some embodiments, the method comprises culturing the selected bacteria for between 2 and 36 days. In some embodiments, the method comprises culturing the selected bacteria for between 4 and 32 days. In some embodiments, the method comprises culturing the selected bacteria for between 6 and 28 days. In some embodiments, the method comprises culturing the selected bacteria for between 8 and 24 days. In some embodiments, the method comprises culturing the selected bacteria for between 10 and 20 days. In some embodiments, the method comprises culturing the selected bacteria for between 12 and 16 days. In some embodiments, the method comprises culturing the selected bacteria for about 1 day, about 2 days, about 3 days, about 4 days, about 5 days, about 6 days, about 7 days, about 8 days, about 9 days, about 10 days, about 11 days, about 12 days, about 13 days, about 14 days, about 15 days, about 16 days, about 17 days, about 18 days, about 19 days, about 20 days, about 21 days, about 28 days, about 35 days, or about 40 days.

[0079] In some embodiments, bacterial culture media comprises any suitable bacterial culture medium. In some embodiments, the bacterial culture medium comprises one or more sugars. In some embodiments, the one or more sugars are one or more sugars selected from the group consisting of glucose, sucrose, dextrose, maltose, fructose, mannose, xylose, and trehalose.

[0080] In some embodiments, the bacterial culture medium comprises glucose. In some embodiments, the bacterial culture medium includes one or more nutrient supplements. In some embodiments, the one or more nutrient supplements are selected from the group consisting of yeast extract, peptone, tryptone, tryptone peptone, trypticase, and trypticase peptone. In some embodiments, the bacterial culture medium comprises yeast extract and peptone.

[0081] In some embodiments, the bacterial culture medium comprises one or more buffering agents. In some embodiments, the one or more buffering agents are selected from the group consisting of NalbPCh, Na2HPO4, KH2PO4, K2HPO4, NaKHPO- 4, ascorbic acid and citric acid. In some embodiments, the bacterial culture medium comprises NalbPC , and citric acid.

[0082] In some embodiments, the culture medium has a pH of between 1 and 9. In some embodiments, the culture medium has a pH of between about 2 and 8. In some embodiments, the culture medium has a pH of between about 3 and 7. In some embodiments, the culture medium has a pH of between about 4 and 6. In some embodiments, the culture medium has a pH of about 1, about 2, about 3, about 4, about 4.1, about 4.2, about 4.3, about 4.4, about 4.5 about 4.6, about 4.7, about 4.8, about 4.9, about 5, about 5.1, about 5.2, about 5.3, about 5.4, about 5.6, about 5.7, about 5.8, about 5.9, about 6, about 7, about 8, or about 9. In some embodiments, the culture medium has a pH of about 5.

[0083] In some embodiments, the solution is subjected to an autoclave cycle for sterilization.

[0084] In some embodiments, the method comprises tuning the pellicle size. Pellicle size and shape can be tuned by selecting suitable well plate types and sizes and by modifying cell culture time or through electric stimulation. In some embodiments, the pellicle is of any shape useful for implantation. In some embodiments, the pellicle is a circle, oval, square, rectangle, rounded square, rounded rectangle, or dogbone. In some embodiments, the pellicle is a circle. In some embodiments, the pellicle has a diameter between about 1 mm and about 100 mm. In some embodiments, the pellicle has a diameter between about 5 mm and about 50 mm. In some embodiments, the pellicle has a diameter between about 10 mm and about 25 mm. In some embodiments, the pellicle has a diameter of about 1 mm, about 2 mm, about 3 mm, about 4 mm, about 5 mm, about 6 mm, about 7 mm, about 8 mm, about 9 mm, about 10 mm, about 12 mm, about 14 mm, about 16 mm, about 18 mm, about 20 mm, about 22 mm, about 24 mm, about 26 mm, about 28 mm, about 30 mm, about 35 mm, about 40 mm, about 50 mm, about 55 mm, about 60 mm, about 65 mm, about 70 mm, about 75 mm, about 80 mm, about 85 mm, about 90 mm, about 95 mm, or about 100 mm.

[0085] In some embodiments, the pellicle has a thickness of between about 0.1 mm and about 5 mm. In some embodiments, the pellicle has a thickness of between about 0.2 mm and about 4 mm. In some embodiments, the pellicle has a thickness of between about 0.3 mm and about 3 mm. In some embodiments, the pellicle has a thickness of between about 0.4 mm and about 2 mm. In some embodiments, the pellicle has a thickness of between about 0.5 mm and about 1 mm. In some embodiments, the pellicle has a thickness of about 0.1 mm, about 0.2 mm, about 0.3 mm, about 0.4 mm, about 0.5 mm, about 0.6 mm, about 0.7 mm, about 0.8 mm, about 0.9 mm, about 1 mm, about 1.2 mm, about 1.4 mm, about 1.6 mm, about 1.8 mm, about 2 mm, about 2.2 mm, about 2.4 mm, about 2.6 mm, about 2.8 mm, about 3 mm, about 3.2 mm, about 3.4 mm, about 3.6 mm, about 3.8 mm, about 4 mm, about 4.2 mm, about 4.4 mm, about 4.6 mm, about 4.8 mm, and about 5 mm. Unless stated otherwise, embodiments describing the pellicle of the present invention also describe the surgical mesh implant of the present invention, and vice versa.

[0086] In some embodiments, methods of the invention comprise step b), applying an electric field to the bacterial culture. In some embodiments, applying an electric field to the bacterial culture manipulates the bacterial nanocellulose fiber orientation during in vitro synthesis. In some embodiments, applying an electric field across the cell culture, aligns bacteria in a process known as electrophoresis. In some embodiments, aligning bacteria along the direction of the electric field results in continuously forming glucan chains, thereby producing a uniaxial surface patterning in the pellicles of bacterial nanocellulose. In some embodiments, modifying certain parameters, such as electric field strength, electric field direction, permittivity of the liquid media, surface potential between the bacterial cell and the liquid media, and the viscosity of the liquid media, the alignment may be modified to produce any suitable pattern. In some embodiments, the surface patterning of the bacterial nanocellulose mesh impacts the direction of myofiber development in human skeletal muscle tissue. As an example, human skeletal muscle tissue relies on linear organization for function and voluntary movement generation, therefore linear uniaxial surface patterning may be appropriate. However, the surface patterning of the bacterial nanocellulose pellicles may be modified to exhibit any suitable pattern for specific tissue engineering needs. The surface patterning of the bacterial nanocellulose mesh has further implications for the tensile strength and porosity of the surgical mesh.

[0087] In some embodiments, step b) comprises generating an electric field across the well plate containing the bacterial cell culture. In some embodiments, the method comprises establishing an electrical field across the culture vessel. In some embodiments, an electric field is generated by contacting the bacterial culture medium with two or more electrodes. For examples, each well of a standard 6-well plate can be fixed with a customized stopper, comprising foramen on opposite sides of each stopper, such that electrodes and leads can pass through, into the bacterial cell culture media at opposite ends of the well plate. Additionally, the number of electrodes and their positioning can be modified to produce different nanocellulose fiber alignments and hence different surface patterning of the surgical mesh. Once electrodes have been inserted into the cell culture media, a voltage may be applied to establish an electric field potential across the culture vessel. In some embodiments, the voltage is a constant DC voltage. In some embodiments, the voltage can be modified, such that the desired patterning is achieved. In various embodiments, the voltage used can be a constant DC voltage of 1 V, 2 V, 3 V, 4 V, 5 V, 6 V, 7 V, 8 V, 9 V, 10 V, 11 V, 12 V, 13 V, 14 V, 15 V, 16 V, 17 V, 18 V, 19 V or 20 V.

[0088] In some embodiments, the exposure of the bacterial cell culture to an electric field is performed when bacterial nanocellulose pellicles can be observed in the culture vessel.

[0089] In some embodiments, the bacterial cell culture is exposed to an electric field for between one hour and about 1 week. In some embodiments, the bacterial cell culture is exposed to an electric field for between about 12 hours and about 6 days. In some embodiments, the bacterial cell culture is exposed to an electric field for between about 1 day and about 5 days. In some embodiments, the bacterial cell culture is exposed to an electric field for between about 2 days and about 4 days. In some embodiments, the bacterial cell culture is exposed to an electric field for about 12 hours, about 14 hours, about 16 hours, about 18 hours, about 20 hours, about 22 hours, about 24 hours, about 1.5 days, about 2 days, about 2.5 days, about 3 days, about 3.5 days, about 4 days, about 4.5 days, about 5 days, about 5.5 days, about 6 days, about 6.5 days, or about 7 days.

[0090] In some embodiments, methods of the invention comprise a step c) harvesting the resulting bacterial nanocellulose pellicle. In some embodiments, step c) further comprises a step of washing the pellicle. In some embodiments, the pellicle is washed with an alkali solution. In some embodiments, the pellicle is washed with a solution of sodium hydroxide, potassium hydroxide, lithium hydroxide, or cesium hydroxide. In some embodiments, the pellicle is washed with a solution of sodium hydroxide. In some embodiments, the sodium hydroxide solution is at a concentration of about 0.1 M, about 0.2 M, about 0.3 M, about 0.4 M, about 0.5 M, about 0.6 M, about 0.7 M, about 0.8 M, about 0.9 M, about 1 M, about 1.1 M, about 1.2 M, about 1.3 M, about 1.4 M, about 1.5 M, about 1.6 M, about 1.7 M, about 1.8 M, about 1.9 M, about 2 M, about 2.5 M, about 3 M, about 3.5 M, about 4 M, about 4.5 M, about 5 M, about 6 M, about 7 M, about 8 M, about 9 M, or about 10 M. In some embodiments, the bacterial nanocellulose pellicles are then harvested from the well plates and washed with a 1 M solution of sodium hydroxide.

[0091] In some embodiments, step c) further comprises decellularizing the resulting bacterial nanocellulose pellicle. In some embodiments, the step of washing the pellicle with an alkali solution decellularizes the pellicle. In various embodiments, the bacterial nanocellulose pellicles are washed one or more times until the solution reaches a neutral pH. In some embodiments, the pellicles are washed with water, distilled water, Millipore® water, or deionized water.

[0092] In some embodiments, the method comprises step d) drying the pellicle. In some embodiments, the method of drying alters the pore size and / or mechanical properties (for example tensile strength and Young’s modulus) of the surgical mesh. Pore size and mechanical properties are dependent on liquid absorption capacity and water content in the wet state. Higher liquid absorbance of bacterial nanocellulose is associated with larger pore sizes and lower Young’s moduli. Further, the effect of porosity has important implications for tissue integration, risk of bowel adhesion and risk of biofdm development as a post-surgical complication. Differential porosity may assist with tissue integration and reduce the risk of bowel adhesion, while too small pore sizes increase the risk of microbial activity by hindering immune-mediated infdtration and surveillance of the implant. In some embodiments, pore size can be tuned by selecting a suitable drying method.

[0093] In some embodiments, the pore size is about 0.1 pm, 0.2 pm, 0.3 pm, 0.4 pm, 0.5 pm. 0.6 pm. 0.7 pm, 0.8 pm. 0.9 pm, 1.0 pm, 1,1 pm, 1.2 pm, 1.3 pm, 1.4 pm, or 1.5 pm, or any range between about 0.1 pm and about 1.5 pm.

[0094] In some embodiments, the bacterial nanocellulose pellicle is dried via stepwise solvent exchange (for example and without limitation, acetone, hexane, or ethanol can be used). In some embodiments, the bacterial nanocellulose pellicle is dried via critical point drying. In some embodiments, the bacterial nanocellulose pellicle is treated via oven drying. In some embodiments, the bacterial nanocellulose pellicle is dried via freeze drying. In some embodiments, the bacterial nanocellulose pellicle is freeze dried by liquid nitrogen lyophilization.

[0095] In various embodiments, the method further comprises sterilizing the bacterial nanocellulose pellicle. The bacterial nanocellulose pellicle may be sterilized by any suitable means known in the art, including, but not limited to, exposure to UV radiation, exposure to X-ray radiation, exposure to gamma radiation, and autoclaving.

[0096] Bacterial Nanocellulose Surgical Mesh

[0097] In another aspect, the present invention relates, in part, to a surgical mesh implant formed using one or more bacterial nanocellulose pellicles produced by any method of the present invention. In some embodiments, the surgical mesh implant comprises one or more pellicles of bacterial nanocellulose with a parietal side and a visceral side. The bacterial nanocellulose surgical mesh provides an alternative surgical mesh option for tension-free tissue defects.

[0098] In some embodiments, the surgical mesh is biocompatible. In some embodiments, the surgical mesh is non-resorbable. Biocompatibility of the bacterial nanocellulose surgical mesh derives from the high purity of nanocellulose produced via bacterial synthesis which occurs through oxidative fermentation under static culture conditions and forms a pure crystalline polymer free of immunogenic or pro- inflammatory phytochemicals, making bacterial nanocellulose an attractive option for use in biomedical applications.

[0099] In some embodiments, the surgical mesh comprises webs of micro-scale filaments in a stochastic assembly that lead to the formation of interconnected pores in a 3D matrix. In some embodiments, the micro-scale filaments are further comprised of nanofibrils linked together via either hydrogen bonds or van der Waal’s interactions. In some embodiments, each nanofibril has a length that extends up to 1 mm. In some embodiments, the nanofibrils are further comprised of a network of P-1, 4 glucan chains held together via weak inter- and intra-molecular forces. In some embodiments, each nanofibril has a diameter of 20 - 100 nm.

[0100] In some embodiments, the surgical mesh comprises bacterial nanocellulose pellicles with fibers that are randomly aligned. In some embodiments, the surgical mesh comprises bacterial nanocellulose pellicles with fibers aligned in a single direction. In some embodiments, the surgical mesh comprises bacterial nanocellulose pellicles with fibers aligned in two or more different directions. In some embodiments, the surgical mesh comprises bacterial nanocellulose pellicles with fibers that may be aligned in any suitable pattern.

[0101] In some embodiments, the surgical mesh is a pliable and spongy aerogel. In some embodiments the surgical mesh is translucent, thin and rigid. In some embodiments, the surgical mesh may have any suitable shape. In some embodiments, the surgical mesh is planar, such as in the form of a sheet. In other embodiments, the mesh can be shaped into a three-dimensional structure, such as a sphere. The surgical mesh can have any suitable thickness. In some embodiments, the surgical mesh has a thickness that is between about 0.1 mm and about 5 mm. In some embodiments, the bacterial nanocellulose pellicle has a thickness of between about 0.2 mm and about 4 mm. In some embodiments, the pellicle has a thickness of between about 0.3 mm and about 3 mm. In some embodiments, the pellicle has a thickness of between about 0.4 mm and about 2 mm. In some embodiments, the pellicle has a thickness of between about 0.5 mm and about 1 mm. In some embodiments, the pellicle has a thickness of about 0.1 mm, about 0.2 mm, about 0.3 mm, about 0.4 mm, about 0.5 mm, about 0.6 mm, about 0.7 mm, about 0.8 mm, about 0.9 mm, about 1 mm, about 1.2 mm, about 1.4 mm, about 1.6 mm, about 1.8 mm, about 2 mm, about 2.2 mm, about 2.4 mm, about 2.6 mm, about 2.8 mm, about 3 mm, about 3.2 mm, about 3.4 mm, about 3.6 mm, about 3.8 mm, about 4 mm, about 4.2 mm, about 4.4 mm, about 4.6 mm, about 4.8 mm, and about 5 mm.

[0102] In various embodiments, the surgical mesh may be of any suitable size or shape. In some embodiments, the surgical mesh is planar with a top-down shape of a circle, oval, square, rectangle, rounded square, rounded rectangle, or dogbone. In some embodiments, the surgical mesh is a circle. In some embodiments, the surgical mesh has a diameter between about 1 mm and about 100 mm. In some embodiments, the surgical mesh has a diameter between about 5 mm and about 50 mm. In some embodiments, the surgical mesh has a diameter between about 10 mm and about 25 mm. In some embodiments, the surgical mesh has a diameter of about 1 mm, about 2 mm, about 3 mm, about 4 mm, about 5 mm, about 6 mm, about 7 mm, about 8 mm, about 9 mm, about 10 mm, about 12 mm, about 14 mm, about 16 mm, about 18 mm, about 20 mm, about 22 mm, about 24 mm, about 26 mm, about 28 mm, about 30 mm, about 35 mm, about 40 mm, about 50 mm, about 55 mm, about 60 mm, about 65 mm, about 70 mm, about 75 mm, about 80 mm, about 85 mm, about 90 mm, about 95 mm, or about 100 mm.

[0103] In some embodiments, the surgical mesh has an ultimate tensile strength of at least about 0.1 MPa. In some embodiments, the ultimate tensile strength is at least about 0.6 MPa, at least about 1 MPa, at least about 2 MPa, at least about 3 MPa, at least about 4 MPa, at least about 5 MPa, at least about 6 MPa, at least about 7 MPa, at least about 8 MPa, at least about 9 MPa, at least about 10 MPa, at least about 12 MPa, at least about 14 MPa, at least about 16 MPa, at least about 18 MPa, at least about 20 MPa, at least about 25 MPa, at least about 30 MPa, at least about 35 MPa, at least about 40 MPa, at least about 45 MPa, at least about 50 MPa, at least about 55 MPa, at least about 60 MPa, at least about 65 MPa, at least about 70 MPa, at least about 75 MPa, at least about 80 MPa, or at least about 85 MPa, or in any range between about 0.1 MPa to about 85 MPa. In some embodiments, the surgical mesh has a Young’s modulus of at least about 0.1 MPa. In some embodiments, the surgical mesh has a Young’s modulus of at least about 0.2 MPa, at least about 0.3 MPa, at least about 0.4 MPa, at least about 0.5 MPa, at least about 0.6 MPa, at least about 0.7 MPa, at least about 0.8 MPa, at least about 0.9 MPa, at least about 1 MPa, at least about 1.1 MPa, at least about 1.2 MPa, at least about 1.3 MPa, at least about 1.4 MPa, or at least about 1.5 MPa. In some embodiments, the surgical mesh implant has a Young’s modulus of between about 0.1 MPa to about 1.5 MPa. In some embodiments, the surgical mesh implant has a Young’s modulus of between about 0.1 MPa to about 1.4 MPa. In some embodiments, the surgical mesh implant has a Young’s modulus of between about 0.1 MPa to about 1.3 MPa. In some embodiments, the surgical mesh implant has a Young’s modulus of about 1.4 MPa. In some embodiments, the bacterial nanocellulose surgical mesh can be sutured to native muscle tissue. In some embodiments, the surgical mesh does not experience degradation when implanted into a patient In some embodiments, the surgical mesh is fully non-resorbable. In some embodiments, the surgical mesh can be integrated into new tissue growth.

[0104] In some embodiments, the pore size can be 0.1 pm, 0.2 pm, 0.3 pm, 0.4 pm, 0.5 pm. 0.6 pm. 0.7 pm, 0.8 pm. 0.9 pm, 1.0 pm, 1,1 pm, 1.2 pm, 1.3 pm, 1.4 pm, or 1.5 pm, or any range between about 0.1 pm and about 1.5 pm. In some embodiments, the pore size on the parietal side of the membrane may be larger than the pore size on the visceral side. In some embodiments, the pore size on the visceral side of the surgical mesh may be larger than the pore size on the parietal side.

[0105] In some embodiments, the surgical mesh provides a scaffold for the ingrowth of new tissue to reinforce the defective muscle. The surgical mesh can support proliferation, migration, and differentiation of cells. In some embodiments, these cells may be human skeletal muscle myoblasts.

[0106] The surgical mesh of the present invention provides a surgical mesh alternative for surgical repair. In some embodiments, the invention can be used as a tension-free repair option for ventral abdominal, inguinal, and femoral herniation. In some embodiments, the surgical mesh can be used as a tension-free tissue repair option for other tissue defects, including but not limited to compartment syndrome, ischemia, volumetric muscle loss, pelvic floor laxity, organ prolapse, and other abdominal, pelvic floor, and soft tissue defects including hernia, fistula, and incisional defects, myocardial defects, and valvular defects.

[0107] It will be understood by those of skill in the art that numerous and various modifications can be made without departing from the spirit of the present disclosure. Therefore, it should be clearly understood that the forms disclosed herein are illustrative only and are not meant to limit the scope of the present disclosure.

[0108] EXPERIMENTAL EXAMPLES

[0109] The invention is further described in detail by reference to the following experimental examples. These examples are provided for purposes of illustration only, and are not intended to be limiting unless otherwise specified. Thus, the invention should in no way be construed as being limited to the following examples, but rather, should be construed to encompass any and all variations which become evident as a result of the teaching provided herein.

[0110] Without further description, it is believed that one of ordinary skill in the art can, using the preceding description and the following illustrative examples, make and utilize the present invention and practice the claimed methods. The following working examples therefore are not to be construed as limiting in any way the remainder of the disclosure. Example 1 : Engineering a Bacterial Nanocellulose Surgical Mesh

[0111] Cellulose is one of the most abundant polymers on Earth consisting of glucose monomers (CeHioOs) arranged via -1,4-glycosidic linkages into long fibrillar chains. These chains interact via intra- and inter-molecular hydrogen bonding, owing to an abundance of hydroxyl groups, condensing into nanofibers. The repeating unit within the polymer is the disaccharide cellobiose, forming the basis for a wide array of Saturday and elastic materials. (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Andree, V., et al. Mater. Res. Express 2021, 8, 025402; Ul-Islam, M., et al. Cellulose 2013, 20, 253-263) Bacterial nanocellulose (BNC) is an emerging biomaterial, with its chemical composition lending hydrophilicity, biodegradability, and chemical modification capacity. (Ul-Islam, M., et al. Cellulose 2013, 20, 253-263; Kuzmenko, V., et al. Mater. Sci. Eng. C 2013, 33, 4599- 4607; Lin, N., et al. Eur. Polym. J. 2014, 59, 302-325; K lemm, D., et al. Mater. Today 2018, 21, 720-748; Barja, F. J. Biomed. Res. 2021, 35, 310-317.) In culture produced as a cellulosic, gelatinous pellicle at the liquid-air interface of carbohydrate rich medium containing synonymous Komagataeibacter or Gluconacetobacter xylinum or Acetobacter xylinum (A. xylinum) - as well as genera such a Acetobacter, Achromobacter, Bacillus, Sarcina, Aerobacter, Agrobacterium, Escherichia, Azotobacter, Rhizobium, Enterobacter, Klebsiella, Salmonella, and others. (K lemm, D., et al. Mater. Today 2018, 21, 720-748; Rangaswamy, B. E., et al. Int. J. Polym. Sci. 2015, 2015, 280784; Ahmed, J., et al. Biotechnol. Adv. 2020, 41, 107549) During aerobic fermentation, A. xylinum and other synthetic species extrude cellulose protofibrils through their cell wall into the extracellular space. (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649) Secreted chains interact to form 3D, randomly oriented, webbed, crystalline networks of nanofibril ribbons 20-100 nm in diameter and up to several micrometers in length. (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Abol-Fotouh, D„ et al. Sci. Rep. 2020, 10, 3491)

[0112] Through continuous synthesis and deposition, a porous matrix of pure cellulose is formed. (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P , et al. Microb. Biotechnol. 2019, 12, 633-649) Though cellulose produced by bacteria is chemically identical to plant cellulose, BNC is produced as a pure polymer and does not contain phytochemicals or immunogenic, pro-inflammatory contaminants (such as lignin, pectin, and hemi-cellulose). (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Keegstra, K. Plant Physiol. 2010, 154, 483-486; Guerriero, G., et al. J. Integr. Plant Biol. 2010, 52, 161-175; Cheng, Y.-W., et al. ACS Biomater. Sci. Eng. 2020, 6, 3046-3054; Delmer, D. P. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999, 50, 245-276) Thus, no additional purification is necessary after BNC synthesis, highlighting one of several notable advantages. In addition, the unique porous structure allows for additional mechanical strength, a high Young’s modulus, and additional water retention capacity compared to plant cellulose. (Lin, N., et al. Eur. Polym. J. 2014, 59, 302-325; Barja, F. J. Biomed. Res. 2021, 35, 310-317; Delmer, D. P. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999, 50, 245-276)

[0113] Due to its durability, elasticity, and other physiologically relevant mechanical properties, BNC is a promising biomaterial for use in culture of human skeletal muscle myoblasts (HSMM) and other skeletal muscle associated tissues. Further, BNC shows promise as biomaterial for medical implants, as it has previously received significant attention as an absorptive bandage or dressing in wound healing due to its hydrophilicity, liquid retention capacity, and nontoxicity. (Barja, F. J. Biomed. Res. 2021, 35, 310-317; Ahmed, J., et al. Biotechnol. Adv. 2020, 41, 107549; Zwimer, J., et al. Sci. Rep. 2021, 11, 2127; Pang, M., et al. Eur. Polym. J. 2020, 122, 109365; Picheth, G. F., et al. Int. J. Biol. Macromol. 2017, 104, 97- 106) The crystalline chemical structure of BNC provides durability and resistance to degradation in the presence of human immune- mediated and tissue remodeling enzymes, thus conferring long-term stability in vivo and providing opportunities for surgical fixation and permanent implantation. (Pang, M., et al. Eur. Polym. J. 2020, 122, 109365)

[0114] BNC also possesses unique physical and mechanical properties as a pure polymer, which can be tuned during and post synthesis of the BNC pellicle fibers in vitro. (Andree, V., et al. Mater. Res. Express 2021, 8, 025402; Zwirner, J., et al. Sci. Rep. 2021, 11, 2127; Pang, M., et al. Eur. Polym. J. 2020, 122, 109365; Singh, G., et al. Biomed. Mater. Bristol Engl. 2021, 16, 062004) P-1,4 glucan chains are extruded from the pole of BNC producing bacterial rods forming fine ribbons that undergo bundling to form microfibrils. Under these conditions, the random movement of bacteria produce an isotropic array of glucan chains forming a nanonetwork from extruded nanofibers. (McNamara, J. T., et al. Annu. Rev. Biochem. 2015, 84, 895-921; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Rangaswamy, B. E , et al. Int. J. Polym. Sci. 2015, 2015, 280784; Wang, L., et al. Carbohydr. Polym. 2020, 249, 116829) The BNC pellicle is ultimately produced at the liquid-air interface where bacterial P-1,4 glucan chain extrusion occurs under static culture conditions at 26-28 °C. (Wang, L., et al. Carbohydr. Polym. 2020, 249, 116829; Bodea, I. M., et al. Polymers 2021, 13)

[0115] Drying methods are known to produce varied results with respect to structural, mechanophysical properties, and surface character, including porosity and surface hornification. (Ul-Islam, M., et al. Cellulose 2013, 20, 253-263; K lemm, D., et al. Mater. Today 2018, 21, 720-748; Bodea, I. M., et al. Polymers 2021, 13) Previous reports have achieved a spongy aerogel consistency through step-wise solvent exchange, critical point drying, and freeze drying, which produce no significant changes in pellicle thickness, and maintain a pliant dry aerogel with little brittleness. Reduced porosity and significant thickness reduction can be achieved through oven drying, producing compact, translucent sheets which are more fragile, yet still pliable. (Ul-Islam, M., et al. Cellulose 2013, 20, 253-263; Bodea, I. M., et al. Polymers 2021, 13) Further, numerous studies cite the importance of hydrogel patterning to properly direct myofiber development in human and murine skeletal muscle tissue engineering models. Bacterial nanocellulose fiber orientation in the hydrogel matrix can be manipulated under electric field stimulation, producing highly aligned fibrous substrate for myriad applications. (Cheng, Y.-W., et al. ACS Biomater. Sci. Eng. 2020, 6, 3046-3054; Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Reid, G., et al. Bioeng. Basel Switz. 2020, 7)

[0116] A bottom-up approach of biomaterial nano-synthesis of high purity from a carbohydrate-rich microbial ecosystem under electrical stimulation was pursued in order to produce a tailored product for applications in biomedicine.

[0117] The materials and methods employed in these experiments are now described. Bacterial Nanocellulose Electrical Stimulation for Tissue Engineering Applications

[0118] Bacterial nanocellulose fiber orientation can be manipulated through an applied electric field to culture inoculum during in vitro synthesis. Here, we designed a custom bioreactor to deliver electric stimulation to BNC pellicles during late-stage incubation and synthesis, thus delivering uniaxial surface patterning to each pellicle through BNC fiber alignment.

[0119] Assuming a bacterial cell acts like a charged particle with a dipole moment in an infinite ionic liquid under a non-uniform electric field, the cell will be driven towards the region of maximal gradient of the applied field. The translational di electrophoretic force EEon the charged particle is given by: where IJ.Cdescribes the induced dipole moment of the bacterial cell. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 31, 1041-1047; Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235- 244)

[0120] If we assume electrostatic interactions between particles in E of diameter d, the dipole-dipole interparticle attraction is proportional to ~ E2and produces particle pairs aligned in the direction of E. Assuming the starting spherical particles are placed in parallel with field lines from E, then the time t required to rotate particle pairs in parallel with E is given by: t ~ 102p • (E0E2).

[0121] In this equation £0is a constant, the permittivity of free space (8.8542 x 1012F / m).

[0122] For the geometries relevant to this study, we assume the establishment of a uniform electric field E. Establishment of a uniform electric field in an ionic liquid allows for mobile ions to propagate parallel to the electric field lines closest to the centroid of the channel or well in a coronal cross-section. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484) This occurs through dual charge accumulation processes. Ions closest to the well or channel wall accumulate due to electrostatics; ambient thermal processes are insufficient for diffusion into the bulk, thus forming a statically bound Stern layer. The electrostatic force dissipates medially within this layer, however, allowing for progressively uninhibited charge flow of an ionic fluid along a perpendicular axis from the well or channel wall. Electro-osmotic (EO) flow describes the net drag forces exerted as mobile ions propagate across the bulk, given by: where the velocity vE0of the ionic fluid is dependent on iE0the mobility of the fluid and the magnitude of the applied electric field E under ideal EO flow. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 31, 1041-1047; Hanaor, D„ et al. J. Eur. Ceram. Soc. 2012, 32, 235-244) To produce fiber alignment via electrical stimulation we estimated the electric field effect on fiber production given the viscosity and flow of an ionic fluid (liquid media). A charged layer will form around electrically charged bacterial cells placed in a uniform electric field as a surface charge is screened by a diffuse layer of oppositely charged ions. An electric field exerts opposing forces on the diffusion layer and cell surface, though the opposing diffusion layer force is ultimately applied to the cell surface through viscous drag. A resultant Coulomb force proportional to the net charge accumulation of surface adsorbed species will drive cellular propagation toward the region of highest opposite potential in a process known as electrophoresis. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 31, 1041-1047; Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235-244; Pohl, H. A., et al. J. Electrochem. Soc. 1960, 107 (5), 390)

[0123] Through electrophoresis, a charged cell (modeled as a sphere) will thus be driven by the Coulombic force as a function of the net charge of the bacterial cell at a velocity vcgiven by: describing a dependence on the mobility of the bacterial cell [band the magnitude of velocity proportional to that of the applied field. (Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 31, 1041-1047; Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235- 244) In electrohydrodynamic processes, given a sphere of diameter d moving with velocity v in a medium of viscosity 77, the viscous drag force FDis governed by the equation:

[0124] FD= 3n ■ 77 ■ d ■ vEQ, and where:

[0125] Assuming constant velocity and steady movement of the charged cell. (Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235-244) Mobility in EO flow is further described by: feo as a function of the permittivity of the fluid (liquid media), ( the zeta potential, describing the electrokinetic surface potential between the ionic bulk fluid and the well or channel wall, and 77 the viscosity. (Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 1, 1041-1047; Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235-244) Given the sample equation for mobility as a function of permittivity of the liquid media, surface potential between the bacterial cell and the liquid media, and the viscosity, and the strength of the applied electric field, the net velocity of the bacterial cell vNETin an applied field is given by: dependent on the sum of the EO and electrokinetic mobilities, respectively. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Pohl, H. A., et al. J. Electrochem. Soc. 1960, 107 (5), 390)

[0126] Thus, from these relationships and geometries we consider the applied electric field uniform and disregard the effect of the dielectrophoretic force (FE). We further deduce a medium of low viscosity and a high electric field strength will produce rapid alignment, assuming bacterial cells in the bulk are charged, free-moving spherical particles, and that they produce glucan chains continuously. (Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Sano, M. B , et al. Ann. Biomed. Eng. 2010, 38, 2475- 2484; Hanaor, D., et al. J. Eur. Ceram. Soc. 2011, 31, 1041-1047; Hanaor, D., et al. J. Eur. Ceram. Soc. 2012, 32, 235-244; Aguilar-Agon, K. W., et al. J. Cell. Physiol. 2019, 234, 23547-23558) From these physics we selected a potential of 10 V (approximately 0.2 V / cm) for continuous stimulation of bacterial culture at the air-media interface for the last three days of culture time.

[0127] The surface patterning of BNC mesh reported herein has important implications for tissue engineering, especially with respect to highly linearized tissue morphologies and ultrastructures exemplified in skeletal muscle, otherwise known as voluntary muscle. (Cheng, Y.-W., et al. ACS Biomater. Sci. Eng. 2020, 6, 3046-3054; Reid, G., et al. Bioeng. Basel Switz. 2020, 7; Chen, S., et al. Biomaterials 2015, 73, 23- 31; An, Y., et al. Chin. Med. J. (Engl.) 2014, 127, 4130-4139) Human skeletal muscle relies on linear organization for function and voluntary movement generation. (Reid, G., et al. Bioeng. Basel Switz. 2020, 7; Feinberg, A. W., et al. Biomaterials 2012, 33, 5732- 5741; Tamargo, M. A., et al. ACS Biomater. Sci. Eng. 2021, 7, 5215-5229) Human skeletal muscle tissue possesses natural regeneration capacity in response to minor stresses and injuries. Through mechanisms under active investigation, a signaling cascade activates muscle stem cells (satellite cells) located within their niche, under the basal lamina along myofibers. Severe muscle defects resulting in incomplete tissue regeneration in vivo from injury, denervation, ablation, infection debridement, or aggressive debulking surgeries can all cause functional deficits in patients and impact quality of life.

[0128] In this work, the influence of BNC in vitro surface patterning and postprocessing is studied in the context of human skeletal muscle tissue engineering. Applications in healthcare are envisaged where novel reconstructive surgery approaches using next-generation, personalized-medicine-based biomaterials and implants are warranted. This applies, in particular, to surgical mesh materials for hernia repair and other soft tissue reconstructions, where surgical teams are still limited by inadequacies in available surgical mesh materials, either due to biocompatibility, long-term durability, cost, or a combination thereof.

[0129] Evaluation of Bacterial Nanocellulose Post-Processing for Physical and Mechanical Tunability Culture of BNC in prepared media incubated at 26 °C produced pellicles over the course of three weeks (Fig. 2). After washing and decellularization, pellicles treated via oven drying produced thin, translucent phenotypes, while those processed via liquid nitrogen freeze drying produced spongy aerogels (Figs. 4, 6).

[0130] The image processing tool Imaged was utilized to assess pore size in postprocessed BNC pellicles under two protocols as described herein. The software tool Imaged was used for image analysis (Fig. 6).

[0131] The distribution of pore sizes in the liquid nitrogen freeze-dried BNC pellicles possessed, on average, significantly larger pores than oven-dried BNC pellicles, differing by more than 50% (p < 0.05). Upon physical examination, oven-dried BNC pellicles were more translucent in character and had reduced thickness compared to liquid nitrogen freeze-dried counterpart samples (Figs. 5, 6).

[0132] The microarchitecture of electrically stimulated BNC pellicles consisted of highly aligned fibers compared to control BNC pellicles grown under static culture conditions free of electric field implementation. Greater than 80% uniaxial fiber alignment within ± 20 ° of axis midline is reported under 10 V stimulation conditions for approximately three days in vitro (Fig. 7).

[0133] Gross-scale analysis of surface character or thickness did not reveal differences between the stimulated and unstimulated BNC pellicles. No differences in gross physical character or mechanical properties were observed between the stimulated and unstimulated BNC.

[0134] Degradation Evaluation

[0135] Hydrogels collected, dry blotted, and weighed at each time point during the degradation study were compared to initial conditions. Initial weights (dry and wet) determined prior to enzyme or mock culture treatment were used as the baseline with little variation over the study period (Fig. 8).

[0136] Bacterial nanocellulose and gelatin methacryloyl (GelMA) hydrogels incubated for one month under mock cell culture conditions were analyzed over the study period. Neither oven-dried nor liquid nitrogen freeze-dried BNC exhibited physical signs of degradation. Comparatively, GelMA gross morphology revealed shrinkage after one month in mock in vitro culture, however, hydrogel mass loss over the study period was not significant but may be attributable to increased water mass (Fig. 8). Mechanical testing was pursued to further evaluate hydrogel integrity after the study period, as previously described, with no significant changes in BNC mechanical strength after the degradation challenge.

[0137] Under mock culture conditions BNC treated with high concentrations of protease

[0138] (collagenase type II) and cellulase, BNC was resistant to protease degradation at 50 U / mL and 5 U / mL but rapidly degraded in the presence of cellulase at both concentrations (1 mg / mL and 10 mg / mL). Comparatively, 20% GelMA hydrogels were used as a control and underwent degradation under both enzymatic conditions with nearly a 50% reduction in sample mass under the highest concentrations of cellulase and collagenase type II (10 mg / mL and 50 U / mL, respectively) over a period of 12 h at 37 °C (Fig- 9).

[0139] Bacterial nanocellulose exhibited robust resistance to degradation after 12 h in DI water as well as both concentrations of collagenase type II, which is present in humans as matrix metalloproteinase-8. Conversely, GelMA exhibited evidence of degradation over 12 h under all experimental conditions, apart from the lowest concentration of cellulase (1 mg / mL) and the DI water control setting (Fig. 9).

[0140] Evaluation of Human Skeletal Muscle Myoblasts in Fabricated Bacterial Nanocellulose Hydrogels

[0141] In an evaluation of human skeletal muscle myoblasts (HSMM) in fabricated bacterial nanocellulose and GelMA hydrogels, HSMM 2.5D culture was pursued in all BNC and GelMA hydrogels for one month. Staining revealed robust proliferation covering full hydrogel surface area, physiologic development, and sustained maintenance of myofibers in BNC hydrogels. Both BNC and GelMA hydrogels had high levels of HSMM proliferation at early (two weeks) timepoints. Myoblasts grown in 2.5D culture in BNC hydrogels were more likely to exhibit mature morphological markers of myofiber development including fusion and multinucleation compared to GelMA samples after one month. Myoblasts grown in GelMA over one month did not exhibit physiologic morphology when analyzed via staining for signs of myofiber development (Figs. 10-12).

[0142] When HSMM 2.5D culture in BNC was compared to standard 2D culture, BNC constructs provided enhanced substrate suitability for myofiber alignment compared to a standard well plate coated with Matrigel (Fig. 10). Confocal imaging of HSMM 2.5D culture in BNC versus GelMA revealed robust proliferation and highly aligned myofiber morphology in the stimulated BNC experimental group. The majority of myofibers cultured in BNC hydrogels were aligned within ± 20 ° from the central axis (Figs. 10-12).

[0143] After one month, hydrogels were then collected for mechanical testing to further investigate physical findings. Upon physical examination, GelMA hydrogels were comparatively difficult to handle and prone to breakage compared to BNC hydrogels which retained their character and physical integrity over the study period. The ultimate strength of liquid nitrogen freeze-dried and oven-dried BNC samples was found to be 0.52 ± 0.14 and 27 ± 5.9 MPa, respectively while GelMA’s ultimate strength was several orders of magnitude lower at 3.3 ± 1.1 kPa after one month.

[0144] The Young’s moduli of the hydrogels were determined from the initial slope of the stress versus strain curve after performing a uniaxial tension test. As anticipated, there was batch-to-batch variability in the production of BNC mesh products and respective mechanical properties.

[0145] Despite batch-to-batch variations in development, each product met the capacity for support of HSMM and physical durability for the proposed healthcare indications. Bacterial nanocellulose constructs effectively maintained physiologic myofibers compared to other conditions and materials tested, suggesting BNC may be used as an implant material for skeletal muscle associated soft tissue repair. The implementation of surgical meshes in soft tissue fixation and repair is instrumental in myriad reconstructive surgeries. Surgical mesh materials are often essential in duroplasty, hernioplasty, post-mastectomy breast reconstruction, pelvic floor prolapse repair, and other surgical procedures. In particular, hernioplasty is a common procedure performed in the setting of symptomatic soft-tissue herniation, a condition affecting more than 20 million people worldwide and with a lifetime risk of hernia of nearly one-third in men. The mainstay of hernia repair is tension-free fixation using surgical mesh. While this method is employed mainly to reduce the rate of hernia recurrence, nextgeneration approaches are necessary in order to improve upon current material products and solutions. Namely, mesh products must meet strict criteria including thresholds for tensile strength, biocompatibility, as well as prevention of infection, adhesions, and hernia recurrence.

[0146] Meshes derived from biological materials typically have the best biocompatibility scores. Decellularized biomaterials such as acellular dermal matrix are defined by chemical non-toxicity and a paucity of immunogenic epitopes. Implantation of these materials blunts the severity of any foreign body reaction marked by inflammation, edema, thrombosis, calcification, fibrosis, seroma formation, bowel adhesions, granuloma formation, and abscesses or fistulas. While performability in anti-inflammatory and cellular integration categories is excellent, many biologically derived materials lack durability, however, and are degraded or resorbed over time, highlighting limitations where permanent repair solutions are required. In addition, materials like acellular dermal matrix used for surgical repair and tissue reconstruction are derived from cadavers and are thus highly unsustainable and costly. Therefore, an ideal surgical mesh material with versatile physical and mechanical properties, while also meeting the rigorous requirements necessary for clinical use is of notable interest.

[0147] The methods used are described herein.

[0148] Bacterial Nanocellulose Culture

[0149] Acetobacter xylimis (ATCC® 3767™) were purchased through ATCC. SkBM™-2 Skeletal Muscle Myoblasts Basal Medium and HSMM (CC-2580) were purchased from Lonza Bioscience. Gelatin from porcine skin (type A), methacrylic anhydride, and dopamine hydrochloride were supplied by Sigma-Aldrich. Sodium hydroxide (NaOH) pellets and dimethyl sulfoxide (DMSO) were purchased from Thermo Fisher Scientific. Glucose, yeast extract, bacto-Peptone, NaFLPCL, citric acid, and photoinitiator 2-hydroxy-4'-(2-hydroxyethoxy)-2-methylpropiophenone (Irgacure 2959) were provided by Sigma- Aldrich. Dulbecco’s phosphate-buffered saline (DPBS) was supplied by Thermo Fisher Scientific. Biopsy Punch, 6 mm (273692) was supplied by KRUUSE. Immunohistochemistry IgG anti-dystrophin antibody (Ab 15277) was purchased through Abeam. Goat anti-rabbit Alexa-Fluor™ 568 (1 :400; cat. no. 81-6114), phalloidin Alexa-Fluor™ 488 (1 :1000; Invitrogen, cat. no, A12379), and DAPI (Img / mL) were purchased through Invitrogen.

[0150] The bacterial strain ATCC® 3767™1was selected for in vitro biosynthesis and production of BNC pellicles for this study. Here, BNC was cultured over the course of several weeks in prepared media. Bacterial culture media was prepared by mixing glucose (20 g L '), yeast extract (5 g L1), bacto-Peptone (5 g L1), NaH2PO4 (2.7 g L1), Citric Acid (1.5 g L1). The solution was pH adjusted to 5.0 and subjected to autoclave cycle for 45 min at 121 °C. (Lin, N., et al. Eur. Polym. J. 2014, 59, 302-325; Barja, F. J. Biomed. Res. 2021, 35, 310-317)

[0151] Bacterial culture plates underwent multiple selection rounds to consistently produce robust BNC pellicles. Briefly, inoculum from productive wells during the first growth cycle of BNC was aspirated from liquid inferior to parental pellicles and seeded onto new plates with freshly prepared, warmed media at a ratio of 1 :50. This selected inoculum stock was maintained in vitro for the duration of the study and the inoculation procedure was repeated for each new growth cycle. The size of the pellicles could be tuned through well plate selection procedures, well plate size, as well as culture time prior to harvesting.

[0152] An apparatus was designed using AutoCAD Fusion 360 for electrical stimulation of BNC meshes in vitro using a standard power supply. A custom-designed well plate lid was 3D printed using FormLabs Form3 printers and a biocompatible Surgical Guide resin. The lid was then retrofitted to a standard well plate allowing for copper wire electrode incorporation into the tissue culture space for establishment of voltage potential across the well plate.

[0153] After establishment of BNC pellicles for two weeks in vitro in designed well plates, an electrical field was introduced for the remaining culture time. A potential of 10 V under direct current was applied across each well plate and BNC pellicles were stimulated for a total of three days in vitro to under a constant electric field in aqueous conditions. After three weeks in culture at 26 °C, the BNC pellicles were harvested and washed with 1 M NaOH in a cleaning and decellularization step. Multiple wash out steps were pursued with deionized (DI) water until the solution reached neutral pH for future work involving biological studies and materials.

[0154] Bacterial nanocellulose pellicles were then placed and stored in DI water at 4 °C in preparation for further differential processing. Washed and sterilized BNC underwent post-processing for tuning mechanical and physical properties. Hydrated meshes underwent oven drying at 60 °C for 12 h on aluminum sheets. The remaining meshes were treated and processed via liquid nitrogen lyophilization. Meshes were submerged in liquid nitrogen and rapidly frozen until completely frozen stiff, with care to avoid cracking. Meshes were then transported to a Labconco Free-Zone 2.5 L -50 °C Benchtop Freeze Dryer and were lyophilized overnight. Oven-treated and freeze-dried meshes were then re-sterilized with UV radiation for 2 min prior to cell culture. Remaining meshes were autoclaved and sealed for biomedical applications.

[0155] Scanning Electron Microscopy

[0156] Analyses of the surface morphology were attained using an ultra-high- resolution field emission gun scanning electron microscopy (SEM) instrument with an acceleration voltage of 10.0 kV (NOVA 200 Nano SEM; FEI, Hillsboro, OR, USA). To reduce perturbations in surface readouts from charge accumulation, BNC mesh samples were sputtercoated with a thin layer of gold-palladium in a nitrogen atmosphere (Agar Sputter Coater, PlanoGmbH).

[0157] Electric field stimulated BNC pellicles were decellularized, washed, and freeze dried using the liquid nitrogen protocol, then prepped for SEM imaging as described. SEM images of stimulated BNC samples were analyzed using ImageJ and a MATLAB script to determine fiber orientation order parameter. (Bolivar-Monsalve, E. J., et al. Bioprinting 2021, 21, e00125) The same image analysis technique was utilized for myofiber alignment in stimulated BNC samples versus unstimulated BNC samples seeded with human skeletal muscle myoblasts.

[0158] Gelatin Methacryloyl Synthesis Dialyzed and freeze-dried gelatin methacryloyl was prepared as previously described.35Photocrosslinking of GelMA was pursued using the commercially available photoinitiator 2-hydroxy-l-[4-(2-hydroxyethoxy)phenyl]-2methyl-l-propanone (Irgacure 2959) under aqueous conditions. Freeze-dried GelMA was dissolved in MiliQ DI water. The solution was first heated to 50 °C for 10 min with subsequent addition of the photoinitator Irgacure 2959. The warm aqueous 20% GelMA solution was placed in preformed molds designed and printed using AutoCAD Fusion 360 and FormLabs Form3 3D printers for photocrosslinking with 0.5 wt% Irgacure 2959 under UV light (35 mW / cm2) with an OmniCure® Series 2000 Spot UV Curing System (Excelitas Technologies) for 2 min.

[0159] Degradation Studies

[0160] Both BNC and GelMA hydrogels were subjected to one month in mock culture in vitro conditions to assess for degradation. Hydrogels were placed in SkBM™-2 Skeletal Muscle Myoblasts Basal Medium prepared from SkBM™-2 Skeletal Muscle Myoblasts Basal Medium Bullet Kit (Lonza Bioscience, CC-3246) and maintained at 37 °C for one month.

[0161] Further, BNC mesh and GelMA samples were treated separately with two biological enzymes, collagenase type II (MMP-8) and cellulase, under incubation at 37 °C. Collagenase II was prepared at concentrations of 50U / mL and 5 U / mL in sterile- fdtered 1% Dulbecco’s Phosphate Buffered Saline (Sigma- Aldrich). Cellulase was prepared at concentrations of 1 mg / mL and 10 mg / mL in sterile-filtered Dulbecco’s Phosphate Buffered Saline (Sigma-Aldrich). GelMA hydrogels were also treated under the same conditions. All samples were assessed for a total of 12 h in vitro under enzymatic treatment.

[0162] Cell Culture Studies

[0163] Stimulated BNC hydrogels were compared to unstimulated control BNC hydrogels, 20% GelMA hydrogels, and traditional 2D culture. Hydrogel specimens were cut into 6 mm disks using a KRUUSE Biopsy Punch, 6 mm (273692). Disk were then sterilized with ethanol and allowed to dry; sterilized hydrogels were placed in 24-well plate in preparation for cell seeding. Each well was seeded with 200k human skeletal muscle myoblasts (Lonza Bioscience, CC-2580). HSMM stock was first thawed and underwent multiple passages before cell seeding experiments between passage 4 and passage 7. Wells containing 2D culture were first coated with Matrigel prior to cell seeding. Human skeletal muscle myoblasts were cultured using SkBM™-2 Skeletal Muscle Myoblasts Basal Medium (Lonza Bioscience, CC-3246) over three weeks in vitro. Media was replaced every other day.

[0164] Fluorescence Microscopy

[0165] Hydrogels from cell culture plates were harvested after three weeks and fixed for 20-30 min using a 4% paraformaldehyde solution. Fixed hydrogels and well plates were then permeabilized in 0.1% Triton X-100 (Invitrogen) for 30 min and blocked in 2% bovine serum albumin solution blocking buffer for 1-2 h. Cell staining for F-actin, nuclei, and dystrophin, was performed according to the manufacturer’s protocol. F-actin and nuclei were stained with phalloidin Alexa-Fluor™ 488 (1 :1000; Invitrogen, cat. no, A12379) and DAPI (Img / mL). Dystrophin was stained with Abeam IgG anti-dystrophin antibody (Ab 15277). Secondary staining was performed with goat anti-rabbit Alexa- Fluor™ 568 (1 :400; Invitrogen, cat. no. 81-6114). Fixed and stained samples were imaged using a Zeiss Observer fluorescence microscope.

[0166] Mechanical Testing

[0167] Hydrogel stiffness was determined by applying a uniaxial tension test to cut 10 mm x 2 mm specimens. Tests were administered using an Instron 5943 Single Column Universal Testing system (Illinois Tool Works, Inc.) at room temperature. Hydrogels were clamped at each end in the dynamic mechanical analysis instrument using two steel clamps and subjected to a uniaxial crosshead displacement rate of 4 mm / min. Testing was performed until hydrogel rupture occurred for each specimen.

[0168] Confocal Imaging

[0169] Human skeletal muscle myoblasts were seeded in electric field stimulated and patterned BNC hydrogels, unstimulated BNC hydrogels, as well as 20% GelMA hydrogels with a seeding density of 100k cells / mL. Human skeletal muscle myoblasts were maintained in culture with SkBM™-2 media exchange every other day. Hydrogels from cell culture plates were harvested after one month. Hydrogels were fixed, permeabilized, and blocked using the aforementioned protocol, and were then stained with F-actin and nuclei were stained with phalloidin Alexa-Fluor™ 488 (1 : 1000; Invitrogen, cat. no, A12379) and DAPI (Img / mL). Hydrogels were imaged using a STELLARIS 5 confocal microscope (Leica Microsystems) and assessed.

[0170] Statistical Analysis

[0171] Statistical differences between samples were calculated using a Student’s two-tailed t-test or two-way ANOVA for independent readings or multiple comparisons, respectively.

[0172] Example 2: Bacterial Nanocellulose Competitive Analysis against Clinical Matrices Here, several processing parameters were investigated and optimized to ensure robust, reliable BNC synthesis including inoculum density, incubation temperature, carbohydrate source and concentration, pH, electric field stimulation and charge flow, as well as systematic bacterial colony selection for enhanced pellicle synthesis. Further, after in vitro treatment and optimization, BNC constructs were subjected to different post-processing techniques for mechanophysical property tuning, either via liquid nitrogen lyophilization or oven drying at overnight at 60 °C. Bacterial nanocellulose has attracted considerable interest in healthcare settings, due to its advantageous properties (mechanical, structural, and physical) and polymeric purity, making it better suited than plant cellulose for biomedical applications. (Abol-Fotouh, D., et al. Sci. Rep. 2020, 10, 3491; Ahmed, J., et al. Biotechnol. Adv. 2020, 41, 107549; Andree, V., et al. Mater. Res. Express 2021, 8, 025402; Roberts, E., L. et al. ACS Mater. Au 2023) It was noted that BNC properties can be tuned and optimized for various applications in healthcare. (Pang, M., et al. Eur. Polym. J. 2020, 122, 109365; Andree, V., et al. Mater. Res. Express 2021, 8, 025402; Wang, L., et al. Carbohydr. Polym. 2020, 249, 116829; Bodea, I. M., et al. Polymers 2021, 13 (13)) BNC was explored as a robust, pliable biomaterial with promising characteristics for biomedical applications as a non-resorbable surgical mesh. In surgical repair settings, mesh materials are applied to a defect site for tissue reinforcement and tension-free repair (Fig. 13). (Bayion, K., et al. Membranes 2017, 7; Alkhoury, F., et al. Surg. Laparosc. Endosc. Percutan. Tech. 2011, 21, 82-85; Deeken, C. R., et al. Surg. Endosc. 2012, 26, 566-575; Afonso, J. S., et al. Int. Urogynecology J. 2008, 19, 375- 380; Li, X., et al. J. Meeh. Behav. Biomed. Mater. 2014, 37, 48-55) In abdominal, femoral, or inguinal hernia repair, mesh fixation and implantation assist in establishing a barrier between protruded viscera and the abdominal wall, cavity, or inguinal canal. (Alkhoury, F., et al. Surg. Laparosc. Endosc. Percutan. Tech. 2011, 21, 82-85; Smart, N. J., et al. The Surgeon 2012, 10, 159-171; Eric, S., et al. Abdominal Wall Treatment Devices, 2010) Barrier re-establishment, in turn, reduces the risk of recurrent herniation (recurrence rate is reduced from approximately 30-40% to 5-10% with mesh implantation compared to direct suturing at the defect site). (Alkhoury, F., et al. Surg. Laparosc. Endosc. Percutan. Tech. 2011, 21, 82-85; Deeken, C. R., et al. Surg. Endosc. 2012, 26, 566-575; Saha, T., et al. OpenNano 2022, 7, 100046) Upon fixation with fibrin glue, sutures, or staples, the mesh must facilitate and undergo tissue integration with minimal foreign body response elicitation. (Deeken, C. R., et al. Surg. Endosc. 2012, 26, 566-575; Smart, N. J., et al. The Surgeon 2012, 10, 159-171; Koscielny, A., et al. Langenbecks Arch. Surg. 2018, 403 (2), 255-263; Anderson, J. M., et al. Semin. Immunol. 2008, 20, 86-100)

[0173] Balancing mesh durability as a permanent, non-resorbable implant with the delicate and demanding requirements for optimized biocompatibility has historically posed challenges. (Alkhoury, F., et al. Surg. Laparosc. Endosc. Percutan. Tech. 2011, 21, 82-85; Deeken, C. R., et al. Surg. Endosc. 2012, 26, 566-575; Li, X., et al. J. Meeh. Behav. Biomed. Mater. 2014, 37, 48-55; Smart, N. J., et al. The Surgeon 2012, 10, 159— 171; Saha, T , et al. OpenNano 2022, 7, 100046; Anderson, J. M., et al. Semin. Immunol. 2008, 20, 86-100) Further, gold standard meshes suffer from structural constraints and changes upon implantation, in addition to limited tissue integration capacity and ongoing risk of implant-associated nosocomial infections, particularly with multi-drug resistant Staphylococcus Aureus (MRSA). (Saha, T., et al. OpenNano 2022, 7, 100046; He, L., et al. Front. Surg. 2022, 9; Narkhede, R., et al. Indian J. Surg. 2015, 77, 322-326; Wilson, R. B., et al. J. Gastrointest. Surg. Off. J. Soc. Surg. Aliment. Tract 2022, 26, 950-964)

[0174] The ideal mesh should minimize the foreign body response and infection risk, promote physiologic tissue ingrowth, facilitate ingress and egress of native products, cells, and soluble factors, protect against defect recurrence, as well as maintain flexibility, elasticity, and appropriate stiffness. Further the ideal mesh should be non-carcinogenic, non-immunogenic, sterilizable, and guard against abscess, serosa, fistula, or bowel adhesion formation in the acute and chronic setting post-implantation. (Smart, N. J., et al. The Surgeon 2012, 10, 159-171; Saha, T., et al. OpenNano 2022, 7, 100046; Ghanashyam, A., et al. 2022) While the clinical gold standard clinical mesh products reliably meet some of the aforementioned qualities and characteristics, mesh material improvements are still needed for integration into clinical practice.

[0175] The shape and structure of mesh pores impact proper integration of the implant since foreign body response severity scales with surface area of the implant. To reduce scar tissue and bridging fibrosis, pores must be sufficiently large to prevent granuloma fusion around neighboring mesh fibers and pores, while simultaneously allowing for easy integration of macrophages, fibroblasts, extracellular matrix fibers, and other immunocompetent surveillance cells. (Li, X., et al. J. Meeh. Behav. Biomed. Mater. 2014, 37, 48-55; Saha, T., et al. OpenNano 2022, 7, 100046) In addition, based on literature, abdominal wall repair represents a unique and notable challenge as implanted mesh substrates are often fixed in physical contact with different body cavities, thus a tailored development solution is warranted to promote healthy tissue integration and defect resolution while the implant interfaces with different tissue types (z.e., parietal versus visceral) on each side of the mesh. (Alkhoury, F., et al. Surg. Laparosc. Endosc. Percutan. Tech. 2011, 21, 82-85; Deeken, C. R., et al. Surg. Endosc. 2012, 26, 566-575; Koscielny, A., et al. Langenbecks Arch. Surg. 2018, 403 (2), 255-263)

[0176] Here two-dimensional (2D) cell culture with human skeletal muscle myoblasts (HSMM) with optimized BNC mesh was compared with commercially available counterparts, Marigen™ and Phoenix™, FDA-approved for wound healing indications, as a demonstration of biocompatibility and promise for applications in healthcare. (Barja, F. J. Biomed. Res. 2021, 35, 310-317; Ahmed, J., et al. Biotechnol. Adv. 2020, 41, 107549; Pang, M., et al. Eur. Polym. J. 2020, 122, 109365; Dorweiler, B., et al. Ges. Gefasschirurgie 2018, 23 (Suppl 2), 46-55)

[0177] Marigen™ and Phoenix™ are both commercially available wound matrices indicated for the support of soft tissue and healing. Marigen™ is derived from fish skin, approximating extracellular matrix or a natural biopolymer hydrogel, while Phoenix™ is a synthetic commercial polymer.

[0178] Enhanced Durability of Bacterial Nanocellulose versus Commercial Mesh Materials Different processing of BNC produces markedly varied outputs in mechanophysical parameters, as reported previously. (Baija, F. J. Biomed. Res. 2021, 35, 310-317; Andree, V., et al. Mater. Res. Express 2021, 8, 025402; Bodea, I. M., et al. Polymers 2021, 13 (13); Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38 (8), 2475-2484; Ul-Islam, M., et al. Cellulose 2013, 20, 253-263) Differences in physical character as well as enhanced absorption capacity of aerogel-like liquid nitrogen freeze-dried BNC compared to oven-dried BNC are shown. (Figs. 14, 15).

[0179] When compared to commercial matrix samples, BNC pore sizes were significantly smaller in the dry state, but liquid nitrogen freeze-dried BNC, in particular, showed higher propensities for swelling capacity and liquid absorption (Fig. 15).

[0180] Young’s moduli were tested in the dry state for each material. Marigen™ commercial matrix produced the highest values (2.1 SD + 0.24 MPa). By comparison, gelatin methacryloyl (GelMA) control hydrogels had the weakest values (0.133 SD ± 0.006 kPa) by several orders of magnitude. It was found that the Young’s modulus of liquid nitrogen freeze-dried bacterial nanocellulose (1.4 SD ± 0.012 MPa) fell between Marigen™ and Phoenix™ (0.15 SD ± 0.012 MPa).

[0181] Each experimental sample was readily manipulated and loaded into an Instron Mechanical Testing System for facile evaluation; in comparison, GelMA samples were mechanically weak and posed problems for the operator during loading and clamping. Bacterial nanocellulose and commercial sample mechanical testing revealed Young’s moduli in a similar regime (Fig. 16) to GelMA control hydrogels.

[0182] Suture strength testing was performed for BNC mesh samples secured to resected porcine hindlimb tissue. Biceps femoris tissue was resected from previously frozen porcine hindlimb under meticulous surgical dissection. Biceps femoris tissue samples were then cut into 10 mm x 2 mm sections for suturing and mechanical evaluation. The inferior boundary of each porcine tissue sample was secured to the superior boundary of sample mesh with five interrupted stiches using Ethicon 2-0 sutures and loaded onto an Instron Mechanical Testing System via glass slides for suture strength testing. Uniaxial crosshead displacement was pursued at a rate of 4 mm / min and proceeded until complete specimen rupture.

[0183] Bacterial nanocellulose mesh samples were compared to commercial Phoenix™ samples in the evaluation of suture strength. All BNC samples sutured to porcine muscle remained intact and the majority of ruptures occurred within the tissue at the suture line. All samples demonstrated elastic and plastic deformation prior to rupture. In contrast to BNC sutured samples, the majority of Phoenix™ sutured samples ruptured within the commercial mesh matrix rather than in the skeletal muscle tissue during testing. Sutured samples were also compared to native porcine muscle controls with no significant difference in the ultimate strength of native muscle versus muscle sutured to BNC constructs (Figs. 16, 17).

[0184] When subjected to 37 °C incubation and 37 °C incubation with extracellular matrix protease for five days Marigen™ rapidly degraded, with wet mass loss over the study period under both conditions. Neither Phoenix™ nor BNC showed signs of degradation in the enzyme-free incubation; however, mass loss occurred in Phoenix™ samples after five days of incubation in collagenase type II extracellular matrix remodeling enzyme conditions.

[0185] Conversely, BNC did not show appreciable signs of degradation under either condition, demonstrating robust resistance to enzyme treatment, even at supraphysiologic concentrations. The GelMA hydrogels demonstrated some resistance in the mock in vitro trial but degraded completely by day three when exposed to collagenase type II (Fig. 18).

[0186] In Vitro Maturation of Human Skeletal Muscle Myoblasts Test

[0187] The materials and methods are now described. Acetobacter xylinus (ATCC® 3767™) were purchased through ATCC.

[0188] SkBM™-2 Skeletal Muscle Myoblasts Basal Medium and HSMM (CC-2580) were purchased from Lonza Bioscience. Gelatin from porcine skin (type A), methacrylic anhydride, and dopamine hydrochloride were supplied by Sigma-Aldrich. Sodium hydroxide (NaOH) pellets and dimethyl sulfoxide (DMSO) were purchased from Thermo Fisher Scientific. Glucose, yeast extract, bacto-Peptone, NaFLPCU, citric acid, and photoinitiator 2-hydroxy-4'-(2-hydroxyethoxy)-2-methylpropiophenone (Irgacure 2959) were provided by Sigma- Aldrich. Dulbecco’s phosphate-buffered saline (DPBS) was supplied by Thermo Fisher Scientific. Biopsy Punch, 6 mm (273692) was supplied by KRUUSE. Immunohistochemistry IgG anti-dystrophin antibody (Ab 15277) was purchased through Abeam. Goat anti-rabbit Alexa-Fluor™ 568 (1 :400; cat. no. 81-6114), phalloidin Alexa-Fluor™ 488 (1 :1000; Invitrogen, cat. no, A12379), and DAPI (Img / mL) were purchased through Invitrogen.

[0189] The BNC-producing strain Acetobacter xylinus (ATCC® 3767™) was selected herein for the initial in vitro biosynthesis of the desired mesh product. The BNC pellicles were synthesized and treated as previously described.1,2’20Briefly, Acetobacter xylinus (A. xylinus), now Gluconacetobacter xylinus or Komagataeibacter xylinus, was cultured for approximately three weeks in prepared carbohydrate-rich media containing glucose (20 g L1), yeast extract (5 g L1), bacto-Peptone (5 g L1), NaFhPCh (2.7 g L1), and citric acid (1.5 g L1). After mixing the media was pH adjusted to 5.0 and autoclaved for 45 min at 121 °C. (Barja, F. J. Biomed. Res. 2021, 35, 310-317; Lin, N., et al. Eur. Polym. J. 2014, 59, 302-325)

[0190] After three weeks in culture all pellicles were harvested, then washed and decellularized with 1 M NaOH. The aqueous solution was then titrated back to neutral pH using multiple rinse steps with MilliQ H2O. (Barja, F. J. Biomed. Res. 2021, 35, 310— 317) Pellicles were then sterilized and stored at 4 °C until further processing. Mechanophysical property tuning was achieved through post-harvest processing. (Barja, F. J. Biomed. Res. 2021, 35, 310-317; Andree, V., et al. Mater. Res. Express 2021, 8, 025402) Pellicles were treated via oven drying at 60 °C for 12 h on aluminum sheets or liquid nitrogen lyophilization using a Labconco Free-Zone 2.5 L -50 °C Benchtop Freeze Dryer with treatment overnight to produce thin membranes or spongy aerogel-like membranes, respectively, for cell culture.

[0191] The BNC pellicles were produced and underwent post-processing both in vitro through electrical stimulation, and after harvesting. Prior to harvesting, selected pellicles were subjected to a 10 V potential under direct current for three days in a custom-designed in vitro stimulation apparatus, while control pellicles received no stimulation. (Wang, L., et al. Carbohydr. Polym. 2020, 249, 116829; Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38 (8), 2475-2484)

[0192] Gelatin Methacryloyl Synthesis

[0193] Gelatin methacryloyl was prepared as previously described. (Van Den Bulcke, A. I., et al. Biomacromolecules 2000, 7, 31-38) Briefly, dialyzed and freeze- dried GelMA was dissolved at a concentration of 20% in warmed MiliQ H2O at 50 °C for 10 min. The commercially available photoinitiator 2-hydroxy-l-[4-(2- hy droxy ethoxy )phenyl]-2-methyl-l -propanone (Irgacure 2959) was then added to the aqueous solution. Photocrosslinking of GelMA hydrogels was pursued in pre-formed molds with 0.5 wt% Irgacure 2959 under UV light with an Omni Cure® Series 2000 Spot UV Curing System standard filter (Excelitas Technologies) for 2 min.

[0194] Mechanical Testing

[0195] Hydrogel specimens were prepared in 10 mm x 2 mm sections for tensile testing and mechanical evaluation. A uniaxial tension test was administered for each dry specimen and each sutured specimen using an Instron 5943 Single Column Universal Testing system (Illinois Tool Works, Inc.) at room temperature. Samples were secured using adhesive tape and loaded into the testing system via metallic pneumatic clamps. Testing was performed for each sample at a uniaxial crosshead displacement rate of 4 mm / min and proceeded until complete hydrogel rupture occurred.

[0196] Degradation Studies

[0197] Hydrogels were maintained under mock in vitro conditions at 37 °C and subjected to enzymatic degradation conditions over the study period. Degradation experiments were undertaken using the extracellular matrix remodeling protease, collagenase type II (MMP-8) and a SkBM™-2 Skeletal Muscle Myoblasts Basal Medium control condition. Hydrogels were assessed over the course of 5 days for signs of degradation. Collagenase II was prepared at concentrations of 50 U / mL in SkBM™-2 Skeletal Muscle Myoblasts Basal Medium.

[0198] Cell Culture Studies

[0199] Hydrogel specimens were cut into disks using a KRUUSE Biopsy Punch, 6 mm (273692) and sterilized prior to cell seeding and culture. Prepared BNC and 20% GelMA hydrogels were compared to commercial (Marigen™ and Phoenix™) specimens and unbleached, unmodified plant-based cellulose for 2.5D culture as well as 2D culture controls.

[0200] Electrically stimulated as well as control (no stimulation) BNC samples were selected for in vitro studies. Human skeletal muscle myoblasts (CC-2580) were purchased from Lonza Bioscience, thawed, and underwent several passages at 60% confluence prior to experimental investigation.

[0201] Hydrogel specimens were loaded into cell culture wells and seeded with 200k human skeletal muscle myoblasts. Cell studies were conducted between passages 4 and 7. Control wells containing 2D culture were first coated with Matrigel prior to cell seeding. Human skeletal muscle myoblasts were seeded at a concentration of 200k cells / ml and cultured using SkBM™-2 Skeletal Muscle Myoblasts Basal Medium. A time course study tracked proliferation from approximately two weeks to over one month in vitro. SkBM1M-2 Skeletal Muscle Myoblasts Basal Medium was replaced every other day.

[0202] Staining and Immunohistochemistry

[0203] For immunofluorescence staining hydrogel samples were fixed in 4% paraformaldehyde for 30 min and permeabilized with 0.1% Triton X-100 for an additional 30 m in phosphate-buffered saline (PBS) at room temperature. Samples were blocked for 2 h in 2% bovine serum albumin (BSA) in PBS with washes performed between each step. After incubation in BSA immunohistochemistry was performed with Abeam IgG anti -dystrophin antibody (Abl 5277) followed by was performed with phalloidin Alexa-Fluor™ 488 (1 :1000; Invitrogen, cat. no, A12379) and DAPI (1 mg / mL). Hydrogel samples were washed between each staining step and blot dried before transfer onto slides with ProLong™ Gold Antifade Mountant (Thermo Fischer, cat. no. P10144) and secured with a coverslip.

[0204] Fluorescence Microscopy

[0205] Hydrogels and 2D controls from cell culture plates were isolated after approximately two weeks or after one month and imaging studies were performed. Fixed and stained samples were imaged using a Zeiss Observer fluorescence microscope.

[0206] Confocal Imaging

[0207] Stained hydrogel samples were imaged at room temperature using a STELLARIS 5 confocal microscope (Leica Microsystems) and assessed for physiologic morphology and ultrastructure. Image acquisition was performed using Leica LAS X Life Science Microscope Software Platform.

[0208] Scanning Electron Microscopy

[0209] Dry fdm surface analyses of BNC morphological features were obtained using an ultrahigh-resolution field emission gun scanning electron microscopy (SEM) instrument with an acceleration voltage of 10.0 kV (NOVA 200 Nano SEM; FEI, Hillsboro, OR, USA). The BNC dry film was fixed to the surface of an aluminum SEM holder using carbon adhesive tape. To reduce signal perturbations in morphological readouts, BNC and commercial samples were sputter coated prior to imaging using a thin layer of gold-palladium in a nitrogen atmosphere (Agar Sputter Coater, PlanoGmbH).

[0210] Fusion Index

[0211] Confocal images were analyzed using established Fiji (ImageJ) ViaFuse-V and ViaFuse-F macros and compared to manual counts to assess for myotube fusion as a marker of cell viability and differentiation. (Aguilar-Agon, K. W., et al. J. Cell. Physiol. 2019, 234, 23547-23558; Chen, S., et al. Biomaterials 2015, 73, 23-31; Heher, P., et al. Acta Biomater. 2015, 24, 251-265; Hinkle, E. R., et al. Skelet. Muscle 2021, 11 , 28) For manual comparison, confocal images were first unstacked prior to analysis. The DAPI channel images were converted to binary, and a threshold was set to isolate nuclei. Prior to counting, individual nuclei were then further isolated using a 50 pm2to 250 pm2boundary condition. Small holes in the nuclei were filled and a watershed function was implemented to de-cluster and to isolate adjacent nuclei. Next, particle analysis was operationalized over the processed binary images. A mask was used on myotube images for morphologic ultrastructure delineation. For calculation of the fusion index, DAPI and myotube staining images were merged. Finally, the total number of nuclei outside of positively stained cells and the total number of nuclei in mononucleated myotubes were recorded and implemented into the fusion index calculation as a ratio.

[0212] Anti-Microbial Assay

[0213] Hydrogels were treated with 2% anti-microbial solution in bacterial culture broth and incubated with multi-drug resistant Staphylococcus aureus (MRSA) or Pseudomonas aeruginosa (P. aeruginosa) at an initial OD of 0.06 across all conditions. Liquid-nitrogen-dried and oven-dried BNC were compared to Phoenix™ and Marigen™ commercial samples as well as a GelMAG anti-microbial hydrogel positive control. Negative control wells contained bacteria with no hydrogel. To demonstrate drug loading, anti -microbial solutions were prepared using antibiotics (1% cephalexin and 1% ciprofloxacin) for broad spectrum activity or 2% polyelectrolyte pDDA, which is often used in detergents.

[0214] To load either antibiotic agent, dry hydrogels were incubated in 2% antimicrobial solution overnight at 4 °C to allow for maximum hydroexpansion and passive absorbance of anti-microbial solution into the hydrogel matrix. Hydrogel samples were also compared to hydrogels prepared with polyelectrolyte solution loaded into the hydrogel matrix during curing with visible light (450-520 nm). Prior to inoculation with MRSA or P. aeruginosa, hydrogel samples were UV sterilized for 2 min on each side at an intensity of 35 mW / cm2using an OmniCure® Series 2000 Spot UV Curing System (Excelitas Technologies). After hydrogels were treated with OD 0.06 bacterial suspension, they were incubated at 35 °C. At certain timepoints, suspended media was collected to assessed for bacterial survival through optical density measured in a microplate reader (BioTek Synergy). Suspended media was also diluted and plated on respective agar plates (tryptic soy broth for MRSA and lysogeny broth for P. aeruginosa) overnight at 35 °C. The next day CFU were counted.

[0215] Bacterial Nanocellulose Boosts in Vitro Maturation of Human Skeletal Muscle Myoblasts Human skeletal muscle myoblasts cultured in aligned, electrically stimulated BNC hydrogels demonstrated enhancement myofiber alignment and markers of viability and maturity including fusion indices compared to all samples evaluated (Figs, 19-21, 25).

[0216] Both electrically stimulated and unstimulated BNC samples used for HSMM tissue culture evaluation (Fig. 22). Electrically stimulated BNC revealed aligned nanocellulose microfibrils, designed as a micropatteming platform to support HSMM (Figs. 21, 23). Though post-processing influenced BNC mechanophysical properties, all BNC samples supported high levels of HSMM proliferation in vitro. Stimulated versus unstimulated BNC were similar in gross-scale physical character. Gross-scale comparison of BNC to commercial samples also revealed physical similarities (Fig. 24).

[0217] Fluorescence staining and microscopy revealed physiologic-like morphology and ultrastructure in BNC samples over a four-week period in vitro. All BNC and GelMA hydrogels evaluated had high levels of HSMM proliferation at early (two week) timepoints.

[0218] Unbleached, unmodified plant-based cellulose constructs had the lowest sustained proliferation and maintenance of HSMM compared to bacterial nanocellulose and commercial mesh constructs, likely owing to differences in substrate purity and mechanophysical character (Figs. 19, 21)

[0219] Fusion Index

[0220] While all BNC and GelMA samples had similar indices of myotube fusion and viability (fusion index) at over four weeks in vitro, BNC nuclei count as a function of area (approximately 350 + 60 nuclei / mm2) at the end of the study period was significantly higher than other samples, including commercial samples Phoenix™ and Marigen™ (104 ± 23 and 70 ± 20 nuclei / mm2, respectively) and GelMA (93 + 15 nuclei / mm2) hydrogels (Fig. 25).

[0221] Anti-Microbial Activity

[0222] Antibiotic and antimicrobial polyelectrolyte loading and bactericidal activity metrics in BNC mesh samples were compared to FDA-approved matrices (Marigen™ and Phoenix™). All samples loaded with 2% antimicrobial poly electrolyte or 2% broad spectrum antibiotic exerted bacteriostatic activity. A measure of colony forming units (CFU) at day five revealed equal antimicrobial activity between BNC and commercial samples. The average CFU counts at day five for liquid nitrogen freeze-dried BNC and oven-dried BNC were 88 ± 16, and 34 ± 13, respectively, for antibiotic treated samples. By comparison, the average CFU counts at day five for Phoenix™ and Marigen™ were 72 ± 23, and 45 ± 18, respectively, treated under the same conditions. Control samples at day five with no drug loading revealed significant bacterial growth over five days. The average control CFU counts (555 ± 34) were significantly larger than both BNC drug-loaded sample types (p<0.0001). The BNC antimicrobial drug loading was also improved compared to positive control glycidyl methacrylate-modified gelatin (GelMAG) cured with poly-(diallyldimethylammonium chloride) (pDDA) antimicrobial hydrogel (Fig. 26).

[0223] Statistical differences between samples were calculated using a Student’s two-tailed t test or two-way ANOVA for independent readings or multiple comparisons, respectively.

[0224] The BNC swelling capacity, in particular liquid nitrogen freeze-dried BNC, outperformed commercial wound care and tissue reconstruction matrices, highlighting its suitability for in vitro cellular integration through diffusion into the swollen matrix. (Barja, F. J. Biomed. Res. 2021, 35, 310-317; Pang, M., et al. Eur. Polym. J. 2020, 122, 109365; Dorweiler, B., et al. Ges. Gefasschirurgie 2018, 23 (Suppl 2), 46-55; Boso, D., et al. Mater. Basel Switz. 2020, 13, 2483) Interestingly, HSMM culture in BNC dramatically augmented myotube and formation and physiologic myofiber hypertrophy, as well as nuclei density compared to traditional GelMA hydrogels and other commercially available mesh products.

[0225] In light of clinical design considerations for surgical mesh, BNC samples underwent suture strength testing, evaluating the suturable performability of BNC in concert with soft tissues. In this case, BNC samples were found to be easily suturable and did not break, tear, or rupture when sutured to porcine skeletal muscle, even during mechanical testing. Rupture events that occurred during mechanical testing of BNC and porcine skeletal muscle constructs were exclusively confined to the skeletal muscle tissue itself, suggesting a comparatively higher degree of mechanical strength within the BNC mesh. These findings contrasted with the evaluation of the Phoenix™ commercial matrix, which experienced rupture prior to the porcine skeletal muscle upon mechanical testing and evaluation. While mechanical mismatch is a concern for any implant, several indications exist for a more durable repair solution with respect to the implanted mesh, especially in the setting of prior soft tissue laxity.

[0226] Further, BNC outperformed commercial matrices, as well as GelMA hydrogels, in long-term durability studies, showing no signs of degradation even in the presence of supraphysiologic enzyme concentrations over several days, while demonstrating comparable mechanical properties, including Young’s modulus, to commercially available samples. Thus, these findings position BNC as an attractive candidate for biomedical implant applications and future in vivo tests and development where permanent, non-resorbable materials are indicated to repair clinically intractable soft-tissue defects or laxities.

[0227] Example 3: A Computational Study for Skeletal Muscle Tissue Engineering

[0228] A functional limitation of skeletal muscle tissue engineering lies in the immature capacity to generate faithful, physiologic, biomimetic scaffolds and constructs. (Aguilar-Agon, K. W., et al. J. Cell. Physiol. 2019, 234, 23547-23558; An, Y., et al. Chin. Med. J. (Engl.)

[0229] 2014, 127, 4130-4139; Boso, D., et al. Mater. Basel Switz. 2020, 13, 2483; Chen, S., et al. Biomaterials 2015, 73, 23-31; Egusa, H., et al. Tissue Eng. Part A 2013, 19, 770-782; Grasman, J. M., et al. Acta Biomater. 2015, 25, 2-15; Heher, P., et al. Acta Biomater.

[0230] 2015, 24, 251-265; Hicks, M. R., et al. Nat. Cell Biol. 2018, 20 (1), 46-57; Khodabukus, A., et al. Biomaterials 2019, 198, 259-269; Nakayama, K. H., et al. Adv. Healthc. Mater. 2019, 8, el801168; Rao, L., et al. Nat. Commun. 2018, 9, 126) To overcome this limitation over recent decades, combinatorial, multifaceted approaches have been pursued using myriad cell types, biomaterials, and stimulation methods. (Aguilar-Agon, K. W , et al. J. Cell. Physiol. 2019, 234, 23547-23558; Chen, S., et al. Biomaterials 2015, 73, 23-31; Grasman, J. M., et al. Acta Biomater. 2015, 25, 2-15; Heher, P., et al. Acta Biomater. 2015, 24, 251-265; Hicks, M. R., et al. Nat. Cell Biol. 2018, 20 (1), 46-57; Khodabukus, A., et al. Biomaterials 2019, 198, 259-269; Nakayama, K. H., et al. Adv. Healthc. Mater. 2019, 8, el 801168; Rao, L., et al. Nat. Commun. 2018, 9, 126; Xi, H., et al. Cell Stem Cell 2020, 27, 158-176. elO) One such mechanism is the custom bioreactor, designed to support and dictate physiologic skeletal muscle functional maturation through structural engineering controls, appropriate chemical signaling, electromechanical stimulation, mechanotransduction cues, or a combination thereof. (An, Y., et al. Chin. Med. J. (Engl.) 2014, 127, 4130-4139; Heher, P„ et al. Acta Biomater. 2015, 24, 251- 265; Khodabukus, A., et al. Biomaterials 2019, 198, 259-269; Ronaldson-Bouchard, K., et al. Nature 2018, 556, 239-243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817; Yang, G. H, et al. Theranostics 2021, 11, 48-63)

[0231] Myoblasts cultured in soft extracellular matrix hydrogels like fibrin and collagen, and subjected to static or dynamic strain regimens, triggers the development of myotubes into concomitant alignment along the axis of strain within the construct. A singular strain axis also encourages endogenous phenotypic patterning and robust sarcomere organization in series. A number of strain and dynamic loading regimes have been explored to elucidate the role of mechanotransduction and relevant parameters in myogenic programming. (An, Y , et al. Chin. Med. J. (Engl.) 2014, 127, 4130-4139; Egusa, H., et al. Tissue Eng. Part A 2013, 19, 770-782; Heher, P., et al. Acta Biomater. 2015, 24, 251-265; Ronaldson- Bouchard, K., et al. Nature 2018, 556, 239-243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817; Dhahri, W., et al. Nat. Biomed. Eng. 2018, 2, 351-352) Rational designs have featured fibrin and other natural hydrogels for mechanical tissue training as they approximate the mechanical properties of native skeletal muscle tissue extracellular matrix. (Chen, S., et al., Biomaterials 2015, 73, 23-31; Heher, P., et al. Acta Biomater. 2015, 24, 251-265; Yang, G. H, et al. Theranostics 2021, 11, 48-63; Choi, Y - J., et al. Biomaterials 2019, 206, 160-169; Heid, S., et al. Acta Biomater. 2020, 113, 1- 22)

[0232] Within the context of this translational work, myriad next generation approaches were leveraged to maximize efficiency through computational design and validation as well as synergistic methodical testing and simulation within the scope of material science, fabrication, and biomaterial use (Figs. 27, 28). (Chen, S., et al., Biomaterials 2015, 73, 23-31; Ronaldson-Bouchard, K., et al. Nature 2018, 556, 239- 243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817) Specifically, to circumvent a trial-and-error approach, a robust systematic method for materials testing was pursued using high-fidelity computational models of the desired 3D biomechanical and electromagnetic stimulation system, modeled using the Computer Assisted Design (CAD) platform Autodesk Fusion360, and tested using the powerful finite element COMSOL Multiphysics package. (Ronaldson-Bouchard, K., et al. Nature 2018, 556, 239-243; Applebaum, M., et al. Cell Differ. 2015, 56, 77-98; Schoen, I., et al. Nano Lett. 2010, 10, 1823-1830) This fully comprehensive bioengineering approach was employed to optimize the timeframe and biomaterials usage in the study, as a novel platform for skeletal muscle tissue engineering (Figs. 29, 30).

[0233] Models of the system of interest were designed using Autodesk Fusion360 and imported in COMSOL Multiphysics for simulation testing based on a comprehensive list of pertinent material properties including Poisson’s ratio (v), Young’s modulus (E), and material density (p). Stress loads on the system were tested after setting the appropriate boundary conditions and employing the relevant physics (Fig. 27). These methods systematically assessed deformation of the desired geometries in response to physiologically relevant stress forces; and identical initial conditions were tested across an array of candidate materials and compared further using two different candidate geometries.

[0234] Based on flexibility inherent to the earbud geometric system and previously investigations of hydrogel suspension capacity, this geometry was first selected over the simple pillars for prototype printing and implementation (Figs. 31, 32). (Ronaldson-Bouchard, K., et al. Nature 2018, 556, 239-243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817) A custom-designed chamber for hydrogel synthesis and curing was built and printed based on this geometry (Fig. 28). Finally, based on standard 6-well plate dimensions, a tunable electrical stimulation bioreactor system was built using CAD, implementing custom-designed stoppers and tubing for local electric field establishment within each well of the plate, according to desired stimulation conditions (Figs. 33, 34).

[0235] From the custom bioreactor design, an electrodynamically controlled assembly and biofabrication of BNC under aerobic conditions is described to produce unilateral micropattemed mesh for tissue engineering and biomedical applications. A custom-built electrical stimulation system was designed to apply electric field stimulation to each well in the system, effecting precise control over bacterial cellulose production in uniaxially aligned networks (Figs. 33, 34). This engineering facilitates enhanced continuous BNC production, since A. xylinum is an obligate anaerobe and the electrolysis of water under an electric field produces oxygen in the culture media. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484; Jacek, P., et al. Microb. Biotechnol. 2019, 12, 633-649; Barja, F. J., Biomed. Res. 2021, 35, 310-317)

[0236] With respect to surgical mesh design, modifications to existing materials pose limitations due to additional manufacturing complexity, restricted permutability or tunability, as well as mechanophysical inadequacies associated with the starting material itself. (Bayion, K., et al. Membranes 2017, 7, 47; Ghanashyam, A., et al. 2022; Van Den Bulcke, A. I., et al. Biomacromolecules 2000, 1, 31-38; Li, X., et al. J. Meeh. Behav. Biomed. Mater. 2014, 37, 48-55; Saha, T., et al. OpenNano 2022, 7, 100046) A bottom-up approach is, therefore, desirable to address precision design requirements in microfabrication and surface patterning while optimizing for material durability and functionality in the appropriate clinical setting (Figs. 33-35). BNC was selected here due to its high chemical purity, physical durability, biocompatibility, hydroexpansivity, and long-term stability. (Grasman, J. M., et al. Acta Biomater. 2015, 25, 2-15; Nakayama, K. H., et al. Adv. Healthc. Mater. 2019, 8, el801168; Barja, F. J., Biomed. Res. 2021, 35, 310-317; Ghanashyam, A., et al. 2022; Lin, N., et al. Eur. Polym. J. 2014, 59, 302-325; Pang, M., et al. Eur. Polym. J. 2020, 122, 109365)

[0237] Theoretical Background Mechanosensing is central for physiologic cellular regulation and maintenance. Within the realm of skeletal muscle tissue engineering, sensitivity to physical parameters and mechanical stimuli is paramount as muscle tissue represents a highly dynamic biological system. Informed, local mechanosensing of deleterious injury or strain is implicated in satellite cell activation, regeneration capacities, and subsequent myofiber repair or hypertrophy.

[0238] To provide an appropriate mechano-stimulus and encourage linear myocyte patterning, a uniaxial hydrogel fixation model between flexible pillars was pursued. (Nakayama, K. H., et al. Adv. Healthc. Mater. 2019, 8, el 801168; Ronaldson- Bouchard, K., et al. Nature 2018, 556, 239-243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817; Tamargo, M. A., et al. ACS Biomater. Sci. Eng. 2021, 7, 5215-5229) Here dual pillar arrays modeled and fabricated using CAD software and 3D printing (Figs. 27-31) are described. The spring stiffness of the pillar geometries was assumed to be dependent on modeling geometries and material properties (including absolute dimensions, and Young’s moduli). For small-scale deformations (i.e., small deflections, 5, compared to the total possible deformation), an ideal spring model is employed according to Hooke’s Law:

[0239] F = k8, where k describes the spring constant of the material of Young’s modulus, E, given by:

[0240] . 3nEd4k = -

[0241] 64L3

[0242] Further, in the computational model, the base was fixed (i.e., rigid substrate), and maximum lateral displacement was evaluated when an inward deflection force was applied to the superomedial pillar face (i.e., a boundary load on a soft pillar). Here, the surface von Mises stress is computed maximally at the fixed boundary between the base and the deformable pillar (Figs. 27, 29-31). The maximum stress omaxis given by: umax ° d s'

[0243] Simulation Rationale and Mechanism For hydrogel curing a CAD model of a chamber was developed to accommodate pillar structures at the appropriate specifications (Figs. 28, 29). Dual pins were fixed onto the pillars to allow for circumferential hydrogel deposition and curing with pillars the flexed position, and resultant free hydrogel fixed under tension: with pillars removed from the chamber and allowed to resume their initial extended position.

[0244] The pairs of pillars were placed at the bottom of the curing chamber in the horizontal position to allow for hydrogel compaction around the vertical pillars during curing.

[0245] In order to determine optimum flexion as a function of material, a systematic computational modeling approach was pursued. Based on physiologically relevant tension parameters, a boundary load at the superomedial face of the pillar of 400 nN was applied to all material interfaces (Figs. 27, 29-31). Pillars were synthesized to allow for either static tension based upon hydrogel curing conditions or dynamic mechanical loading during auxotonic tissue contraction (Figs. 28, 29). Finite element analysis (FEA) was performed to compare pillar flexion using traditional (polydimethylsiloxane) PDMS versus 3D-pritable resins.

[0246] While PDMS has desirable ease of manipulation, cost-effectiveness, and biocompatibility, it also requires additional processing steps including mold design and curing protocols to produce desired geometries. Polydimethylsiloxane mechanical properties are also dependent on several factors including curing time, curing temperature, and current agent concentration, thus different Young’s moduli for PDMS were built into the model for evaluation. (Ronaldson-Bouchard, K., et al. Nature 2018, 556, 239-243; Ronaldson-Bouchard, K., et al. Nat. Protoc. 2019, 14, 2781-2817; Tamargo, M. A., et al. ACS Biomater. Sci. Eng. 2021, 7, 5215-5229) Polydimethylsiloxane also has non-selective absorption tendencies of hydrophobic compounds, including many small molecule drugs, thus 3D-printable resins were selected for comparison to save on fabrication time for the system. (Tamargo, M. A., et al. ACS Biomater. Sci. Eng. 2021, 7, 5215-5229)

[0247] The platform was designed to accommodate 24 tissues in separate wells suspended between flexible pillar geometries (Fig. 32). This was achieved using standard 24-well plate dimensions published from MatTek Corporation to design pillar arrays for tissue culture of human skeletal muscle myoblasts. Pillars were printed using FormLabs 3D printers and biocompatible resins (Figs. 28, 29, 32).

[0248] Fabrication of the Bacterial Nanocellulose Electrical Stimulation Platform

[0249] An electrical stimulation apparatus designed to simulate electrophoresis kinetics within each well of a standard 6-well plate design was developed. Many protocols often require expensive equipment and platinum materials for electrical stimulation, creating a barrier for access to these technologies and stimulation platforms. This barrier was overcome by developing a user-friendly protocol and basic copper wires; establishing a sacrificial system in which BNC pellicles were stimulated and terminally harvested upon completion of the three-day stimulation regimen. To ensure continuation of the study, prior to terminal stimulation, each well was aspirated, and the aspirate was used to inoculate the next culture plate. Rather than integrating custom Arduino software and complex hardware ecosystems, standard gel electrophoresis systems were repurposed in combination with widely available tissue culture plates to meet experimental needs (Figs. 33, 34).

[0250] Standard tissue culture plates were simply modified and retrofitted using a 3D printing approach. Re-usable, loosely fitting stoppers were designed to allow for adequate aeration of the bacterial inoculate and printed accordingly in reference to MatTek 6-well plate standard dimensions and geometric specifications. Each stopper was designed with oppositional foramen for built-in electrode incorporation (Fig. 34). The voltage and charge injection across each reactor in the electrical stimulator culture system were extensively evaluated, and stimulation conditions were selected based on previous literature. (Sano, M. B., et al. Ann. Biomed. Eng. 2010, 38, 2475-2484)

[0251] An electrical stimulator generated direct current transmitted to each well of the bioreactor system placed in an incubator with temperature and partial pressure settings appropriate for bacterial culture. Electrodes were placed in oppositional orientations in each well via retrofitted stopper inlets spanning the diameter of each well (Fig. 34).

[0252] Further investigation is warranted with other electrode materials including carbon graphite, titanium, platinum, stainless steel, and titanium-nitride coated titanium. To save costs associated with this technology, however, stimulation was pursued with readily available copper wires.

[0253] The disclosures of each and every patent, patent application, and publication cited herein are hereby each incorporated herein by reference in their entirety. While this invention has been disclosed with reference to specific embodiments, it is apparent that other embodiments and variations of this invention may be devised by others skilled in the art without departing from the true spirit and scope of the invention. The appended claims are intended to be construed to include all such embodiments and equivalent variations.

Claims

CLAIMSWhat is claimed is:

1. A method of producing a surgical mesh implant comprising the steps of: a) culturing a bacterium that produces cellulose; b) exposing the culture to an electric field; c) harvesting the resulting pellicle; d) drying the pellicle.

2. The method of claim 1, wherein step c) further comprises a step of washing and decellularizing the pellicle.

3. The method of claim 2, wherein the step of washing and decellularizing the pellicle comprises washing the pellicle with an alkali solution.

4. The method of claim 3, wherein the alkali solution is a sodium hydroxide, lithium hydroxide, potassium hydroxide, or cesium hydroxide solution at a concentration of between 0.5 M and 2 M.

5. The method of claim 1, wherein the method of drying in step d) is accomplished by thermal evaporation or freeze-drying.

6. The method of claim 5, wherein the method of thermal evaporation comprises oven drying.

7. The method of claim 5, wherein the method of freeze-drying comprises liquid nitrogen lyophilization.

8. The method of claim 1, wherein the bacterium of step a) is selected from a group consisting of Komagataeibacter, Acetobacter xylinum, Acetobacter, Achromobacter, Bacillus, Sarcina, Aerobacter, Agrobacterium, Escherichia, Azotobacter, Rhizobium, Enterobacter, Klebsiella, and Salmonella.

9. The method of claim 8, wherein the bacterium sAcetobacter xylinum.

10. The method of claim 1, wherein the electric field of step b) is a constant electric field.

11. The method of claim 10, wherein the electric field is at a potential of between about 1 V and about 20 V.

12. A surgical mesh implant comprising one or more layers of bacterial nanocellulose, wherein each layer has a parietal side and a visceral side, wherein the layers are stacked on top of each other such that the parietal side of one is in contact with the visceral side of the next layer, and wherein the surgical mesh implant comprises a plurality of pores from the parietal side to the visceral side of the surgical mesh implant.

13. The surgical mesh implant of claim 12, wherein the surgical mesh implant has a Young’s modulus of between 0.1 MPa to 1.5 MPa.

14. The surgical mesh implant of claim 12, wherein the surgical mesh implant has an ultimate tensile strength of between 0.1 MPa to 85 MPa.

15. The surgical mesh implant of claim 12, wherein the surgical mesh implant has a pore size of between 0.1 pm to 1.5 pm.

16. The surgical mesh implant of claim 12, wherein the pore size on the parietal side of the implant is larger than the pore size on the visceral side.

17. The surgical mesh implant of claim 12, wherein the implant has a thickness of between 1 mm to 5 mm.

18. The surgical mesh implant of claim 12, wherein the implant is a circle having a diameter of between 1 mm to 100 mm.

19. The surgical mesh implant of claim 12, wherein the implant degrades less than 10% when the implant is exposed to collagenase type II for 5 days.

20. The surgical mesh implant of claim 12, wherein the implant is impregnated with an antibiotic.

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