Biological tissue and manufacturing method

Abstract

Processed adipose tissue is described, having an extracellular matrix derived from human adipose tissue and adipocytes with at least 10% intact adipocyte composition. Methods for processing adipose tissue and media loading tools are also described.

Description

[Technical Field]

[0001] (CROSS-REFERENCE TO RELATED APPLICATIONS) This application claims priority to U.S. Provisional Application No. 63 / 383,574, filed November 14, 2022, and U.S. Provisional Application No. 63 / 383,578, filed November 14, 2022, both of which are incorporated by reference herein in their entireties. [Background technology]

[0002] Cosmetic or reconstructive surgery is often the result of disease, trauma, or injury that damages local soft tissues. These defects can cause loss of normal tissue function, trauma, and pain (Coleman, SR 2016. Plastic and Reconstructive Surgery 118, 108S-120S). Alternatively, cosmetic applications include the treatment of small volumes (facial contouring) or large volumes (breasts and buttocks) using either synthetic or natural fillers. Current clinical procedures often use autologous fat grafting or commercially available fillers to assist in repair (Coleman, SR 2016. Plastic and Reconstructive Surgery 118, 108S-120S., Gir, P. et al. 2012. Plastic and Reconstructive Surgery 130, 249-258). Autologous fat grafting uses a patient's own adipose tissue via liposuction to fill voids while minimizing host immune rejection and promoting adipogenesis (Serra-Renom, JMet et al. 2010. Plastic and Reconstructive Surgery 125, 12-18). While advantageous, the use of heterogeneous, unprocessed fat can lead to donor-site morbidity from liposuction (Patrick Jr, CW 2001. The Anatomical Record 263, 361-366), poor viability of residual adipocytes, which can lead to necrosis (Young, DA et al. 2012. Biomed Mater 7, 024104), and delivery of free lipid and tissue debris can result in cyst formation and calcification (Kokai, LE et al. 2020. Plast Reconstr Surg Glob Open 8, e2574).Depending on the location of the injection, these complications can result in unaesthetic results such as contour irregularities, tissue buildup, inflammation, or even more serious problems such as fat embolism, stroke, and / or blindness (Vasavada, A. et al. 2023. Autologous Fat Grafting For Facial Rejuvenation., StatPearls Publishing LLC).

[0003] Over-the-counter fillers have become increasingly popular due to the increasing demand for cosmetic surgery, and recent research provides insight into current advances in this field (Choi, J. Het al. 2010. Tissue Eng Part B Rev 16, 413-426). Synthetic and natural materials have been used extensively as soft tissue fillers, but with limited success. Synthetic polymers such as polyethylene glycol or polylactic acid provide injectable materials and / or controlled compositions, are biologically inert, and cause reduced host uptake, granuloma formation (Cohen, G. et al. 2009. Journal of drugs in dermatology: JDD 8, 486-489), and / or immune rejection (Levenberg, S. et al. 2004. Curr Top Dev Biol 61, 113-134; Cronin, KJ et al. 2004. Plastic and Reconstructive Surgery 113, 260-269; Patrick, CW et al. 1999. Tissue Eng 5, 139-151).

[0004] However, synthetic polymers lack the appropriate natural tissue composition and mechanical properties. Furthermore, biodegradable synthetic polymers have poor mechanical properties (Young, DA et al. 2012. Biomed Mater 7, 024104), poor volume retention (Patrick Jr, CW 2001. The Anatomical Record 263, 361-366), and biodegradable compounds may accumulate, causing immune responses or even toxicity (Fu, K. et al. 2000. Pharmaceutical Research 17, 100-106, Maeder, K. et al. 1996. Biomaterials 17, 457-461).

[0005] To better mimic native tissue, natural fillers containing extracellular matrix (ECM) have demonstrated increased volume retention over time, host integration, reduced immune response, and adipogenesis in vitro (Flynn, LE 2010. Biomaterials 31, 4715-4724), in vivo (Young, DA et al. 2012. Biomed Mater 7, 024104, Han, TTY et al. 2015. Biomaterials 72, 125-137), and in clinical trials (Gold, MH et al. 2020. J Cosmet Dermatol 19, 1044-1056, Dige A. et al. 2019. Gastroenterology 156, 2208-2216 e2201). Native ECM retains structural and biological properties that enhance cell-cell and cell-matrix interactions for increased cell growth and tissue regeneration (Boudreau, N. et al. 2006. Cell 125, 429-431). These results have led to increased interest in minimally engineered adipose tissue that retains its native ECM for use as adipose tissue allografts (Banyard, DA et al. 2019. Regenerative Medicine and Plastic Surgery: Skin and Soft Tissue, Bone, Cartilage, Muscle, Tendon and Nerves, 71-89, Springer International Publishing).

[0006] However, the use of chemical, biological, and physical treatments in the production of ECM-retaining tissues can cause destruction of ECM proteins, structure, or potentially damage the sample. There is a need in the art for minimally manipulated, injectable, processed tissues that retain native adipocyte structure and ECM. The embodiments described herein fulfill this need. Summary of the Invention

[0007] In one embodiment, the processed adipose tissue comprises an extracellular matrix derived from human adipose tissue and adipocytes having at least 10% intact adipocyte composition. In one embodiment, the intact adipocyte composition is at least 30%. In one embodiment, the intact adipocyte composition is 30%-70%. In one embodiment, the intact adipocyte composition is 30%-55%. In one embodiment, the intact adipocyte composition is 45%-70%. In one embodiment, the tissue structural proteins comprise at least one of collagen, glycoproteins, and proteoglycans retained from human adipose tissue. In one embodiment, the tissue comprises decellularized tissue having reduced amounts of allogeneic components, cellular components, and free lipids. In one embodiment, the tissue is configured for implantation or injection into a patient. In one embodiment, the tissue is configured to pass through an opening of 1 mm to 4.5 mm. In one embodiment, a method for processing adipose tissue includes freezing and thawing human adipose tissue, separating the human adipose tissue into small pieces by mechanical comminution, sequentially rinsing the human adipose tissue with a non-ionic surfactant, a crystalloid solution, and sterile water, and dispensing the human adipose tissue into packages using a loading tool. In one embodiment, the loading tool comprises a jar having a proximal end opening, a distal end opening, and a chamber disposed therebetween; a piston configured to be actuated by a lever to slide distally and flush along the wall of the chamber; and a stand configured to stabilize the piston under the lever and form a hinged connection with the lever. In one embodiment, the proximal end opening and the distal end opening are each at least partially surrounded by a threaded surface. In one embodiment, the loading tool comprises an adapter configured to connect to at least one of the proximal and distal openings of the jar, the adapter having a first sealable port. In one embodiment, the piston has a second sealable port. In one embodiment, the method comprises debriding human adipose tissue to remove extraneous or damaged tissue.In one embodiment, the method comprises rinsing the human adipose tissue with a non-ionic surfactant, a sterile crystalloid solution, and sterile water by distributing the human adipose tissue into a centrifuge tube and centrifuging the human adipose tissue. In one embodiment, the method comprises rinsing the human adipose tissue with a non-ionic surfactant, a sterile crystalloid solution, and sterile water by distributing the human adipose tissue into a sealable container and gently shaking the container. In one embodiment, the method comprises rinsing the tissue in a non-ionic surfactant at least twice. In one embodiment, the method comprises rinsing the tissue in a crystalloid solution at least twice. In one embodiment, the method comprises rinsing the tissue in sterile water at least twice. In one embodiment, the method comprises drying the human adipose tissue in a centrifuge tube containing a drying sieve. In one embodiment, the method comprises decanting the tissue after each rinse and removing any decanted liquid and cellular components. In one embodiment, the method comprises using a loading tool to evenly distribute the human adipose tissue into the container. In one embodiment, the method includes packaging the container in a sterilization pouch.

[0008] In one embodiment, the medium loading tool includes a jar having a proximal end opening, a distal end opening, and a chamber disposed therebetween; a piston configured to be actuated by a lever to slide distally and flush along the wall of the chamber; and a stand configured to stabilize the piston under the lever and form a hinged connection with the lever. In one embodiment, the proximal end opening and the distal end opening are each at least partially surrounded by a threaded surface. In one embodiment, an adapter is configured to connect to at least one of the proximal end opening and the distal end opening, the adapter having a first sealable port. In one embodiment, the piston has a second sealable port. [Brief explanation of the drawings]

[0009] The foregoing and other objects and features will become apparent with reference to the following description and the accompanying drawings, which are included to provide an understanding of the invention according to its various embodiments and which form a part of this specification, and in which like numerals represent like elements.

[0010] [Figure 1A] FIG. 1A shows a flow diagram illustrating an embodiment of a method for manufacturing an adipose tissue filler allograft according to one embodiment, FIG. 1B shows a flow diagram illustrating a method for manufacturing an adipose tissue filler allograft according to one embodiment, and FIG. 1C shows a flow diagram illustrating a method for processing adipose tissue according to one embodiment. [Figure 1B] FIG. 1A shows a flow diagram illustrating an embodiment of a method for manufacturing an adipose tissue filler allograft according to one embodiment, FIG. 1B shows a flow diagram illustrating a method for manufacturing an adipose tissue filler allograft according to one embodiment, and FIG. 1C shows a flow diagram illustrating a method for processing adipose tissue according to one embodiment. [Figure 1C] FIG. 1A shows a flow diagram illustrating an embodiment of a method for manufacturing an adipose tissue filler allograft according to one embodiment, FIG. 1B shows a flow diagram illustrating a method for manufacturing an adipose tissue filler allograft according to one embodiment, and FIG. 1C shows a flow diagram illustrating a method for processing adipose tissue according to one embodiment. [Figure 2A] 1 depicts an alternative side view of a loading tool assembly according to one embodiment. [Figure 2B] 1 depicts an alternative side view of a loading tool assembly according to one embodiment. [Figure 2C] FIG. 10 is a cross-sectional view of a loading tool jar according to one embodiment. [Figure 2D] 10 illustrates an alternative view of the first adapter according to one embodiment. [Figure 2E] 10 illustrates an alternative view of the first adapter according to one embodiment. [Figure 2F] 10 illustrates an alternative view of a second adapter according to one embodiment. [Figure 3A] 1 illustrates a visual inspection chart for color inspection according to one embodiment. [Figure 3B]1 shows a visual inspection chart for an excess free lipid test according to one embodiment. [Figure 4] 1 shows a graph illustrating DNA quantification results according to one embodiment. [Figure 5] 1 shows a graph depicting the average DNA content in all samples tested according to one embodiment. [Figure 6] 10 shows a table illustrating average component quantification as a percentage according to one embodiment. [Figure 7A] 1 shows the component amounts of water, lipids, and solids (including ECM) in grams, according to one embodiment. [Figure 7B] 1 shows the component amounts of water, lipids, and solids (including ECM) in grams, according to one embodiment. [Figure 8A] 1 shows a graph depicting the relative amounts of ECM protein expression between embodiments compared to native adipose tissue. [Figure 8B] 1 shows a heat map illustrating the major ECM components expressed in one embodiment compared to native adipose tissue. [Figure 9] 1 shows four histological images from adipose tissue: A and B show staining from untreated tissue; C and D show staining according to one embodiment. [Figure 10] Figure 1 shows four histological images from adipose tissue: A and B show staining from untreated tissue; C and D show staining from one embodiment. [Figure 11] Four histological images from adipose tissue are shown: Figures 11 and 12 show staining from untreated tissue, and Figures 13 and 14 show staining from one embodiment. [Figure 12] Four histological images from adipose tissue are shown: Figures 11 and 12 show staining from untreated tissue, and Figures 13 and 14 show staining from one embodiment. [Figure 13] Four histological images from adipose tissue are shown: Figures 11 and 12 show staining from untreated tissue, and Figures 13 and 14 show staining from one embodiment. [Figure 14]Four histological images from adipose tissue are shown: Figures 11 and 12 show staining from untreated tissue, and Figures 13 and 14 show staining from one embodiment. [Figure 15A] Figures 15A and 15B show images of histological staining of samples of the present invention at different time points. Figure 15A shows staining at 2 months. Figure 15B shows staining at 3 months. Figure 16 shows staining at more than 4 months. Figure 17 shows staining at 6 months. [Figure 15B] Figures 15A and 15B show images of histological staining of samples of the present invention at different time points. Figure 15A shows staining at 2 months. Figure 15B shows staining at 3 months. Figure 16 shows staining at more than 4 months. Figure 17 shows staining at 6 months. [Figure 16] Figures 15A and 15B show images of histological staining of samples of the present invention at different time points. Figure 15A shows staining at 2 months. Figure 15B shows staining at 3 months. Figure 16 shows staining at more than 4 months. Figure 17 shows staining at 6 months. [Figure 17] Figures 15A and 15B show images of histological staining of samples of the present invention at different time points. Figure 15A shows staining at 2 months. Figure 15B shows staining at 3 months. Figure 16 shows staining at more than 4 months. Figure 17 shows staining at 6 months. [Figure 18] Figures 18A and 18B show images of histological staining of samples of the present invention at different time points: Figure 18A shows staining at over 2 months, Figure 18B shows staining at over 3 months, and Figure 18C shows staining at 4.5 months. [Figure 19] Figure 19 shows images of histological staining of samples of the invention at different time points: Figure 19A shows staining at less than 1 month, Figure 19B shows staining at 1.5 months, and Figure 19C shows staining at 2.5 months. [Figure 20] 1 shows a graph illustrating the mechanical hardness over time for various samples of the present invention. [Figure 21] 1 shows the mechanical hardness over time of one sample of the present invention from one donor. [Figure 22] 1 shows the mechanical hardness over time of one sample of the present invention from a second donor. [Figure 23]Shown are images of staining from samples of human ASCs grown in control medium in the absence of test samples, stained with Oil Red O at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and (E) 2 weeks. [Figure 24] Shown are images of staining of human ASCs grown in differentiation medium in the absence of test samples with Oil Red O stained at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) 2 weeks. [Figure 25] Shown are images of staining of human ASCs grown in control medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) 2 weeks. [Figure 26] Shown are images of staining of human ASCs in control medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) 2 weeks. [Figure 27] Shown are images of staining of human ASCs in control medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) 2 weeks. [Figure 28] Shown are images of staining of human ASCs grown in differentiation medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 1 week, C) 2 weeks, and D) 2 weeks. [Figure 29] Shown are images of staining of human ASCs grown in differentiation medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 1 week, C) and D) 2 weeks, and E) and F) 2 weeks. [Figure 30] Shown are images of staining of human ASCs grown in differentiation medium in the presence of test samples with Oil Red O stained at A) 24 hours, B) 1 week, C), D) 2 weeks, E) and F) 2 weeks. [Figure 31] Shown are images of staining of human ASCs grown in control medium in the presence of a test sample identified as "Conventional" with Oil Red O staining at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) 2 weeks. [Figure 32]Shown are images of staining of human ASCs grown in differentiation medium in the presence of a test sample identified as "conventional" with Oil Red O staining at A) 24 hours, B) 1 week, C), D) 2 weeks, and E) and F) 2 weeks. [Figure 33] 1 shows a table showing the results of a human ASC-induced differentiation assay of various samples of the present invention and conventional samples. [Figure 34] 1 shows a graph illustrating absorbance values ​​for samples of a test embodiment of the present invention, conventional samples, and control samples. [Figure 35] Shown are images of test wells with a test sample identified as "Conventional" A) before test sample was added, B) 24 hours after test sample was added, C) 48 hours, D) 72 hours, and E) with 1% EV staining. [Figure 36] Shown are images of a test well with a test sample identified as "invention" A) before test sample was added, B) 24 hours after test sample was added, C) 48 hours, D) 72 hours after test sample was added, and E) with 1% EV staining. [Figure 37] Shown are images of control wells with primary Human Dermal Blood Endothelial Cells (HDBECs) identified as "Positive Control" and complete growth medium A) before the test sample was added, B) 24 hours after the test sample was added, C) 48 hours after the test sample was added, and D) 72 hours after the test sample was added. [Figure 38] Shown are images of control wells with primary HDBECs and basal medium identified as "Negative Control" A) before test sample was added, B) 24 hours after test sample was added, C) 48 hours after test sample was added, and D) 72 hours after test sample was added. [Figure 39] 1 shows a table showing results from an endothelial proliferation assay. [Figure 40] 1 shows a graph depicting results from an endothelial proliferation assay. [Figure 41]1 shows a table showing results from an endothelial proliferation assay averaged across samples of the invention, samples from conventional tissues, and positive and negative controls. [Figure 42] 1 shows a graph showing results from an endothelial proliferation assay averaged across samples of the invention, samples from conventional tissues, and positive and negative controls. [Figure 43] Image J analysis using the adiposoft plugin to calculate adipocyte area is shown. [Figure 44] Formula 1 is shown below. [Figure 45] Images are shown with five areas analyzed for each sample staining for comprehensive evaluation. [Figure 46] H&E staining of the first treated tissue is shown. [Figure 47] H&E staining of the second treated tissue is shown. [Figure 48] H&E staining of conventionally processed tissue is shown. [Figure 49] 1 is a table showing the adipocyte composition ranges of the first embodiment and the adipocyte composition ranges of the second embodiment. [Figure 50] Graph showing compositional analysis of adipose tissue showing lipid content (75-80%), water content (15-20%), and protein content (3-5%). [Figure 51] 1 is a table of the compositional analysis of the first allograft embodiment. [Figure 52] 1 is a graph showing quantification of a first tissue constituent for representative donor 1. [Figure 53] 10 is a table of the compositional analysis of a second allograft embodiment. [Figure 54] 1 is a table comparing adipose tissue composition excluding data for one representative donor 3. DETAILED DESCRIPTION OF THE INVENTION

[0011] It should be understood that the drawings and descriptions of the present invention have been simplified to show elements relevant to a clearer understanding of the invention, while excluding, for clarity, many other elements found in systems and methods of the art. Those skilled in the art may recognize that other elements and / or steps are desirable and / or necessary 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 invention, a discussion of such elements and steps is not provided herein. The disclosure herein covers all such variations and modifications to such elements and methods known to those skilled in the art.

[0012] Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. Although any methods and materials similar or equivalent to those described herein can be used in the practice or testing of the present invention, the preferred methods and materials are described.

[0013] As used herein, each of the following terms has the meaning associated with it in this section.

[0014] 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.

[0015] As used herein, when referring to a measurable value (e.g., amount, duration, etc.), "about" is meant to encompass variations of ±20%, ±10%, ±5%, ±1%, and ±0.1% from the particular value (where such variations are appropriate).

[0016] Ranges: Throughout this disclosure, various aspects of the invention may 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. Where appropriate, the description of a range should be considered to have all the possible subranges as well as individual numerical values ​​within that range specifically disclosed. For example, the description of a range such as 1 to 6 should be considered to have specifically disclosed subranges such as 1 to 3, 1 to 4, 1 to 5, 2 to 4, 2 to 6, 3 to 6, etc., as well as individual numbers within that range, e.g., 1, 2, 2.7, 3, 4, 5, 5.3, and 6. This applies regardless of the breadth of the range.

[0017] Referring now in detail to the drawings, wherein like reference numerals indicate like parts or elements throughout the several views, in various embodiments, minimally manipulated tissue filler allografts are presented herein.

[0018] One strategy for preserving the native extracellular matrix (ECM) in minimally manipulated adipose tissue is decellularization. Decellularization is a process that aims to improve biocompatibility or minimize immune responses by removing tissue-resident cells while leaving the ECM intact (Fu, RH et al. 2014. Cell Transplant 23, 621-630). Process steps can vary, but generally require continuous washing in chemical and biological agents, which can range from several days to over a week (Flynn, LE 2010. Biomaterials 31, 4715-4724., Aamodt, JMet al. 2016 Biomaterials 86, 68-82., Hinderer S. et al. 2016. Adv Drug Deliv Rev 97, 260-269). The use of chemical, biological, and physical treatments can cause destruction of ECM proteins (e.g., glycosaminoglycans) (Reing JE et al. 2010. Biomaterials 31, 8626-8633.), disruption of structure (Sesli, M. et al. 2018. Turk J Biol 42, 537-547.), or potentially damage the sample (Hayes, DJ et al. 2022. Biomaterials and Biosystems 7, 100053). In general, decellularization of adipose tissue requires precise control over a series of processing steps (approximately 2–5 days depending on the tissue and protocol) (Wang, L. et al. 2013. Acta Biomaterialia 9, 8921–8931., Mohiuddin, OA et al. 2019. 57–70 Springer International Publishing) to ensure all cellular components are removed while ensuring minimal damage to the residual ECM (Choi, J. Set al. 2011. Journal of Biomedical Materials Research Part A 97A, 292–299).The use of minimally manipulated adipose tissue requires fewer processing steps while still providing the natural adipose extracellular matrix (ECM) and tissue integrity for soft tissue fillers. Minimal manipulation of adipose tissue, or processing that does not alter the original relevant properties related to the tissue's utility for reconstruction, repair, or replacement (21 CFR 1271.3(f)(1)), provides a safe (Services, USDoHaH2020 ed Food and Drug Administration) and standardized approach to providing clinicians with off-the-shelf minimally manipulated adipose tissue products (MMAPs).

[0019] The present invention relates to minimally manipulated adipose tissue allografts that can be injected or surgically implanted into patients requiring soft tissue replacement. The adipose tissue allografts herein retain the native ECM and adipocytes of the donor tissue. Furthermore, the adipose tissue allografts also retain structural proteins such as collagen, glycoproteins, and proteoglycans. The retained cellular components are necessary to maintain the structural integrity and volume of the allograft. The adipose tissue allografts also contain greatly reduced amounts of cellular and allogeneic components (e.g., DNA and nucleic acids), which may induce an immune response that, if severe enough, may lead to immune rejection of the allograft. The allografts further retain components such as ECM, lipids, and water, the amounts of which can be varied across the manufacturing process embodiments described herein to produce various embodiments of the present invention. Adipose tissue filler allografts indeed exhibit higher levels of adipocytes, ECM structure and structural proteins, and greatly reduced amounts of DNA and nucleic acids upon histological staining (see Example 2).

[0020] The present disclosure further provides embodiments of methods for preparing minimally manipulated adipose tissue allografts capable of retaining at least a portion of intracellular components. The methods herein include several steps, discussed in detail below, for lysing the tissue of cellular and allogeneic components while avoiding denaturation and washout of structural proteins. The cellular and allogeneic components are removed by a freeze-thaw process, a mechanical grinding process, and rinsing with mild detergent, sterile saline, and sterile water. These processes weaken and / or disrupt the cellular membranes of cells in the tissue, thus helping to decellularize the tissue. Furthermore, rinsing also separates intracellular components from the rest of the tissue. These processes are also gentle enough to help retain necessary structural components of the tissue (e.g., extracellular matrix, adipocytes, and structural proteins). The methods described herein are minimally manipulated methods that fall within the scope of the 361 HCT / P guidelines.

[0021] 1A, a method 100 for processing adipose tissue generally includes freeze-thaw cycling (step 101), debridement and sectioning of human donor adipose tissue (step 102), size reduction (step 103), fluid handling (step 104), pre-packaging storage (step 105), quality control testing (step 106), and packaging (step 107). These processes are uniquely combined to affect the amount of tissue components, including adipocytes, free lipids, extracellular matrix, water, and oxygen, as well as to remove voids and homogenize the allograft.

[0022] The method begins by harvesting human cadaveric adipose tissue from a donor using any suitable method, which may include, but is not limited to, suction, incision, scraping, and other surgical methods known in the art. In other embodiments, the donor tissue may be harvested and stored at frozen temperatures.

[0023] 1B, in one embodiment, step 101 includes freezing the harvested donor tissue and thawing the harvested donor tissue for a period not exceeding 120 hours. This freeze-thaw cycle is the first step in decellularizing tissue, as cell membranes can be disrupted during freeze-thawing.

[0024] Step 102 involves debridement of the tissue to remove any extraneous or damaged tissue. The tissue is debridemented in a sterile field, and the amount of time the tissue is left open is consistently recorded. The adipose tissue is measured, weighed, and inspected for scars and extraneous tissue such as visible fascia, arteries, veins, or muscles. The tissue is debridemented using methods known in the art to remove all extraneous and damaged adipose tissue. Suitable methods may include, but are not limited to, the use of scalpels, forceps, scrapers, and other methods known in the art. Any extraneous and damaged tissue may be discarded.

[0025] Step 103 involves mechanically separating the tissue into small pieces. In some embodiments, separation can be achieved by cutting, grinding, crushing, or any other suitable method. In some embodiments, the donor tissue can be cut into small pieces. These pieces may be cubes of approximately 4 cm x 4 cm or less. Further separation can be achieved by feeding the adipose tissue cubes into a mechanical grinder. In some embodiments, the mechanical grinder can be a meat grinder or any other suitable grinder known in the art. Any excess connective tissue resulting from the tissue separation process can be discarded.

[0026] Mechanical comminution contributes to the decellularization process, and the shear force of the mill has the ability to tear the cell membranes of some cells. Mechanical comminution of adipose tissue also allows the product to be subjected to sufficient force to separate the tissue into fine particles. This provides the necessary injectability, allowing the product to pass through containers, syringes, or cannulas with diameters of 1.5 mm to 4.5 mm or greater. In one embodiment, the processed tissue is injectable through an opening with a diameter of 1.5 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 2 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 2.5 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 3 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 3.5 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 4 mm. In one embodiment, the processed tissue is injectable through an opening with a diameter of 4.5 mm. Furthermore, the mechanical comminution step also applies a force to the tissue that is gentle enough to keep at least 10% of the adipocytes in the tissue intact. Intact adipocytes, in one embodiment, maintain the structure of natural adipocytes. Intact adipocytes, in one embodiment, maintain the functions of natural adipocytes for cushioning and support. Intact adipocytes, in one embodiment, maintain the adipocyte tissue structure and extracellular matrix (ECM). Intact adipocytes, in one embodiment, maintain the adipocyte tissue structure, ECM, triglycerides, and connective tissue. In one embodiment, the tissue is treated with adipocytes having at least 10% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 15% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 20% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 25% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 30% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 40% intact adipocyte composition.In one embodiment, the tissue is treated with adipocytes having at least 50% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 55% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 60% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 65% intact adipocyte composition. In one embodiment, the tissue is treated with adipocytes having at least 70% intact adipocyte composition. Adipocytes help maintain the structural integrity and volume of the allograft, making it a better implant for soft tissue. Other methods known in the art for size reduction (e.g., cryo-milling) do not allow for the retention of adipocytes.

[0027] Step 104 involves liquid processing of the product. This step is broken down into several substeps, including successive rinses with non-ionic surfactant, sterile crystalloid solution, and sterile water. After each rinse, the tissue is decanted to separate it from any unwanted liquid or cellular components.

[0028] Step 104a involves separating the tissue from free lipids. In some embodiments, separation can be accomplished by distributing the pulverized adipose tissue into centrifuge tubes in equal portions. The tissue can then be centrifuged at a speed of 900-3655 RCF for an extended period of time. The free lipids can be separated from the adipose tissue by decanting the adipose tissue through a sieve or other methods known in the art. Any decanted liquid, oil, or cell pellets are discarded. The tissue can then be rinsed with sterile warm water. The tissue is returned in equal portions to the centrifuge tubes, and sterile warm water is added in an approximately 1:1 ratio of sterile warm water to tissue. The centrifuge tubes can be gently shaken to disperse the adipose tissue in the water. The centrifuge tubes can then be centrifuged at ambient temperature at 900-3655 RCF. The adipose tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell pellets are discarded.

[0029] Step 104b involves rinsing the adipose tissue with a non-ionic surfactant. This step further contributes to cell lysis, and the surfactant weakens cell membranes, helping to decellularize the tissue. In some embodiments, the surfactant product used can be a 0.01% Triton X-100 solution. In some embodiments, rinsing is performed by dispensing the adipose tissue into a centrifuge tube and adding a 0.01% Triton X-100 solution. The solution can be added at a 1:1 ratio to the adipose tissue. The centrifuge tube can be gently shaken for a period of time, such as 100, 200, 300, 400, 500, 600, 800, or 1,000 seconds or more. The centrifuge tube can then be centrifuged at a speed of 900-3655 RCF. In some embodiments, rinsing can be performed by dispensing the adipose tissue into a sealable container. A 0.01% Triton X-100 solution is then added at an approximately 1:1 ratio to the adipose tissue. The sealable container is then shaken several times during the rinse period (e.g., 1, 2, 3, 4, 5, 6, 8, 10 minutes, or more). The adipose tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell pellets discarded. The detergent rinse can be repeated as needed.

[0030] The adipose tissue is dried. In some embodiments, drying can be accomplished by centrifuging the tissue. Centrifugation can be performed at a speed of 900-3655 RCF for, for example, 1, 2, 3, 4, 5, 6, 8, or 10 minutes. The adipose tissue is decanted through a sieve, and any decanted liquid, oil, or cell pellet is discarded.

[0031] Step 104c includes rinsing the tissue in sterile crystalloid solution. The adipose tissue is rinsed with sterile crystalloid solution. In some embodiments, the sterile crystalloid solution can be 0.9% sterile saline. In some embodiments, rinsing may be performed by dispensing the adipose tissue into a centrifuge tube. 0.9% sterile saline is added to the tube at a solution-to-tissue ratio of approximately 1:1. The centrifuge tube is gently shaken. The centrifuge tube is then centrifuged at ambient temperature at a speed of 900-3655 RCF. In some embodiments, rinsing may be performed by dispensing the adipose tissue into a sealable container. 0.9% sterile saline is added to the adipose tissue at a solution-to-tissue ratio of approximately 1:1. The sealable container is gently shaken several times during the rinsing period. After rinsing, the tissue may then be decanted through a sieve, and any decanted liquid, oil, or cell pellets are discarded. The sterile saline rinse may be repeated multiple times.

[0032] Step 104d involves rinsing the tissue in sterile water. The adipose tissue is rinsed with sterile water. In some embodiments, the sterile water may be warm. In some embodiments, rinsing is performed by dispensing the adipose tissue into a centrifuge tube. Sterile water is added to the centrifuge tube at a solution-to-tissue ratio of approximately 1:1. The centrifuge tube is gently shaken for approximately 2, 5, 10, 15, or 20 seconds. The tube is then centrifuged at ambient temperature at a speed of 900-3655 RCF. In some embodiments, rinsing may be performed by dispensing the adipose tissue into a sealable container. Sterile water is added to the sealable container at a solution-to-tissue ratio of approximately 1:1. The sealable container is gently shaken several times during the rinsing period. After rinsing, the tissue may then be decanted through a sieve, and any decanted liquid, oil, or cell pellets are discarded. The sterile water rinse may be repeated multiple times.

[0033] Step 104e involves removing excess water from the adipose tissue. In some embodiments, the excess water is removed by loading a dry sieve into the centrifuge tube. A wipe is cut around the perimeter of the dry sieve and placed on top of the sieve. The adipose tissue can be distributed in equal amounts into the tubes, and the tubes can be centrifuged at ambient temperature. The tubes can be centrifuged at speeds of 900-3655 RCF. Excess water can be drained through the bottom opening of the centrifuge tube. The drained, unwanted liquid can be discarded. Step 104e can be repeated for multiple centrifugation cycles (e.g., 2, 3, 4, 5, or more).

[0034] Step 105 involves pre-packaging the resulting adipose tissue allograft. Pre-packaging involves combining the tissue filler products and gently mixing until a uniform consistency is achieved. The adipose tissue product may be prepared for packaging by evenly distributing the product into centrifuge tubes. The tubes may be centrifuged at a speed of approximately 900-3655 RCF for a predetermined amount of time (e.g., 200, 300, 400, 500, 600, 800, or 1,000 seconds).

[0035] 1C , a method 150 for processing adipose tissue includes steps 152 of freezing and thawing human adipose tissue, step 154 ​​of separating the human adipose tissue into small pieces by mechanical comminution, step 156 of sequentially rinsing the human adipose tissue with a non-ionic surfactant, a sterile crystalloid solution, and sterile water, and step 158 of dispensing the human adipose tissue into packages using a loading tool. In one embodiment, the method includes debridement of the human adipose tissue to remove extraneous or damaged tissue. In one embodiment, the method includes rinsing the human adipose tissue with a non-ionic surfactant, a sterile crystalloid solution, and sterile water by dispensing the human adipose tissue into a centrifuge tube and centrifuging the human adipose tissue. In one embodiment, the method includes rinsing the human adipose tissue with a non-ionic surfactant, a sterile crystalloid solution, and sterile water by dispensing the human adipose tissue into a sealable container and gently shaking the container. In one embodiment, the method includes rinsing the tissue multiple times in a non-ionic surfactant. In one embodiment, the method includes rinsing the tissue multiple times in sterile crystalloid solution. In one embodiment, the method includes rinsing the tissue multiple times in sterile water. In one embodiment, the method includes drying the human adipose tissue in a centrifuge tube containing a drying sieve. In one embodiment, the method includes decanting the tissue after each rinse and removing any decanted liquid and cellular components. In one embodiment, the method includes evenly distributing the human adipose tissue into a container using a loading tool. In one embodiment, the method includes packing the container into a sterile pouch.

[0036] In some embodiments, tissue products can be dispensed via a loading tool, such as that shown in Figures 2A-2F, according to one embodiment. The loading tool can be used in conjunction with adipose tissue manufacturing processes to enable centrifugation, atmospheric control, moisture content, and composition control, physical handling, and dispensing of adipose allografts and similar tissues and media. The tool can be made to be sterilized via autoclave and does not require external power. The loading tool can retain a moldable and flowable media during centrifugation, seal the media during atmospheric transfer, remove unwanted voids and gas pockets from the media, and dispense the media to fit a variety of different containers. In one embodiment, the loading tool 200 comprises a centrifuge-compatible jar 250 or container with sealable openings at both ends, such as a proximal opening 252 and a distal opening 254. The sealable openings can include, for example, a threaded exterior for mating with a twist cap to seal the opening. The openings can be configured to allow efficient decantation during liquid processing, then one-way gas exchange during atmospheric control, and then void-free distribution during dispensing. A first adapter 270 (also referred to as a container adapter), such as that shown in FIGS. 2D and 2E, can mate with the distal opening 254 on the jar 250 via a threaded surface and provide a sealable access port 276 with a proximal opening 272 and a distal opening 274 to facilitate the distribution of medium from the jar 250 into the container. A connecting structure 278, such as a threaded opening for a shoulder screw, may be included to assemble the separable parts of the first adapter 270. A second adapter 280 (also referred to as a nitrogen adapter), such as that shown in FIG. 2F, can mate with the distal opening 254 on the jar 250 via a threaded surface and provide a sealable access port 286 with a proximal opening 282 and a distal opening 284 for fluid or gas communication with the jar 250, for example, to remove gas or liquid from the medium in the jar 250. This sealable access port 286 may also be connectable to a valved conduit to create a sealed fluid communication.

[0037] The piston 202, stand 204, locking bar 206, jar 250, and nitrogen adapter 280 together facilitate creating an airtight container around the culture medium. Ports on the nitrogen adapter and piston 275 allow for one-way gas exchange to remove undesirable gases, such as oxygen, from the culture medium. The ports may have controllable valves, for example, to control fluid and gas communication. The lever 210 and container adapter 270 cooperate with other components to distribute the culture medium in a void-free and hypoxic manner. The lever 210 can be attached to the stand 204 with a hinge connection 212. A locking bar 206 can be added to prevent movement of the lever 210.

[0038] The loading tool can be completely disassembled and sterilized. The materials are compatible with various sterilization methods, such as steam sterilization and ethylene oxide. The loading tool allows for atmospheric control through sealed atmosphere exchange for product packaging. The tissue is contained in a sealed space with ports to facilitate vacuum removal and flow-through replacement of the naturally occurring atmosphere, and optional refilling with an inert gas. Atmospheric exchange allows for storage using a variety of inert gases. The loading tool gently pushes the tissue directly into the container in a controlled manner with minimal atmospheric exposure, creating a filled container with minimal voids. The loading tool can be adapted to various tip configurations for use with the desired container. The loading tool jar is also centrifuge-compatible in both shape, strength, and mass. The aforementioned centrifuge-compatible jar has sealable openings on both the top and bottom for selective removal of high- or low-density tissue, solution, and / or debris. This, along with the proximal and distal openings of the jar, facilitates product rinsing and allows the same container to be used for product filling / packaging. The loading tool allows for water content control through suspension centrifugation. A jar and dry sieve assembly for floating tissue, with or without the addition of a permeable membrane that suspends the tissue above the empty space during centrifugation to remove excess water and / or other solutions present in the tissue. The tool and its components are reusable, as the components can be autoclaved.

[0039] The use of a loading tool for the packaging process allows for efficient dispensing of allograft material within the primary package, which can improve the allograft's shelf stability, cosmetic performance, and allows for controlled dispensing at the point of use. The loading tool is prepared for use by loading the centrifuge jar containing the adipose tissue onto the adapter. The loading tool's piston is slightly depressed to engage the O-ring, and the locking bar is inserted and latched. A nitrogen adapter is attached to the base of the jar, connecting vacuum to the piston.

[0040] The loading tool is then used to purge oxygen from the tissue. The vacuum pump is adjusted (e.g., to about -5, -10, -15, -20 Hg, or higher) and the vacuum line valve is opened. The nitrogen line valve is opened sufficiently to reduce the vacuum (e.g., to about -5, -10, -15 Hg for 5, 10, or 15 seconds). The nitrogen line valve is turned off and opened a second time to reduce the vacuum a second time. The nitrogen line valve is turned off, then opened a third time and left on. The vacuum pump is turned off while leaving the vacuum line open and bringing the pressure to 0 Hg with nitrogen. The nitrogen line valve is turned off.

[0041] The loading tool is then prepared for packaging. After the pressure has normalized, the locking bar is removed and the piston is lowered to contact the adipose tissue. The vacuum line valve is closed and the nitrogen adapter is removed. The container is then attached to the container adapter.

[0042] The tissue can now be packaged. The loading tool 200 is used in combination with a container adapter 270 to package the tissue into a final primary container. As mentioned herein, a syringe 290 is referenced according to some embodiments, but more generally, some type of container, such as a syringe, cannula, or other type of container, can be adapted for use.

[0043] Step 106 involves performing several quality control tests to ensure the adipose tissue allograft is up to standard.

[0044] In some embodiments, the tissue is visually inspected for color. If the tissue is off-white in color and / or has various shades / tones of yellow according to the chart shown in Figure 3A, the product passes inspection. If the tissue is not off-white in color or various shades / tones of yellow according to the chart shown in Figure 3A, the product fails inspection. If the tissue fails the color inspection test, reprocessing must be performed.

[0045] Reprocessing involves evenly distributing the adipose tissue into centrifuge tubes and immersing the adipose tissue in sterile 0.9% saline to achieve a saline-to-adipose tissue ratio of approximately 1:1. The centrifuge tubes are gently shaken for approximately 2, 5, 10, 15, or 20 seconds to disperse the adipose tissue in the saline. The centrifuge tubes are then centrifuged at 900-3655 RCF for 5 minutes. The tissue may then be decanted through a sieve, and any cell pellets, liquid, or oil may be discarded. This process is repeated, with the adipose tissue gently mixed in the centrifuge tube and returned to the centrifuge. The tissue is then centrifuged at 900-3655 RCF for 3 minutes. The tissue may then be decanted through a sieve, and any undesirable liquid or cellular components may be discarded. Visual color inspection is repeated. Reprocessing may be repeated as many times as necessary until the product is acceptable. A final visual color inspection is recorded.

[0046] In some embodiments, a syringe passage test is performed. A syringe is filled to the brim with product (approximately 14 cc). The syringe is connected to a single outlet cannula. The product is squeezed through the cannula. If the entire contents of the syringe pass through the cannula with finger pressure, the product passes the syringe passage test. If no tissue can be squeezed out with finger pressure on the plunger and / or the cannula becomes clogged, the product fails the syringe passage test. If the product fails the syringe passage test, it is reworked and the test is repeated.

[0047] Reprocessing involves squeezing the tissue through a 3-4 mm sieve using a scraper into a bowl. Collect only the tissue that passes through the sieve. Gently mix the tissue until a uniform consistency is achieved. The adipose tissue is then equally distributed among centrifuge tubes and immersed in sterile water at a sterile water to adipose tissue ratio of approximately 1:1. Gently shake the centrifuge tubes for approximately 2, 5, 10, 15, 20 seconds, or more to disperse the adipose tissue in the sterile water. The tubes are then centrifuged at 900-3655 RCF (e.g., 200, 300, 400, 500, 600, 800, 1,000 seconds, or more). Decant the adipose tissue through the sieve and discard any cell pellet, liquid, or oil. Gently mix the adipose tissue in the centrifuge tube, return it to the centrifuge, and centrifuge at 900-3655 RCF (e.g., for 60, 90, 120, 150, 180 seconds, or longer). Decant the tissue again through the sieve and discard any cell pellet, liquid, or oil. Repeat the syringe passage test; reprocessing may be repeated as many times as necessary until the product is acceptable. Record the final syringe passage test.

[0048] In some embodiments, the product is visually inspected for excess free lipids according to the chart shown in Figure 3B. If the free lipids do not form an oily crust on or around the tissue according to the chart, the tissue is acceptable. If the free lipids form an oily crust on or around the tissue according to the chart, the tissue is unacceptable and therefore fails the test. If the product fails, it is reprocessed and the test is repeated.

[0049] Reprocessing involves evenly distributing the adipose tissue into centrifuge tubes and immersing the tissue in sterile water to achieve an approximately 1:1 ratio of sterile water to adipose tissue. The tubes are gently shaken (e.g., for 5, 10, 15, 20, 25 seconds, or more) to disperse the adipose tissue in the water. The tubes are then centrifuged at 900-3655 RCF for 5 minutes. The tissue is decanted through a sieve and any cell pellet, liquid, or oil is discarded. The adipose tissue is gently mixed in the centrifuge tube, returned to the centrifuge, and centrifuged at 900-3655 RCF. The tissue is again decanted and any cell pellet, liquid, or oil is discarded. The free lipid test is repeated, and reprocessing may be repeated as many times as necessary until the product is acceptable. Final test results are recorded.

[0050] In some embodiments, an absorption measurement test is performed. Absorbent paper is placed on a clean, flat, nonporous surface. A pea-sized sample of the product is placed on the absorbent paper. The tissue is allowed to sit for 15 minutes. Using a ruler, the distance migrated is measured from the point where the tissue contacts the paper to the furthest edge of the lipid trail. If the migrated lipid trail is 5 mm or less on the absorbent paper, the product is acceptable. If the migrated lipid trail is more than 5 mm on the absorbent paper, the product is unacceptable. If the product fails the absorption measurement test, reprocessing is performed and the test is repeated. Reprocessing involves evenly distributing the adipose tissue into centrifuge tubes and immersing the tissue in sterile water to achieve a sterile water to adipose tissue ratio of approximately 1:1. The tubes are gently shaken for approximately 2, 5, 10, 15, or 20 seconds to disperse the adipose tissue in the water. The tubes are then centrifuged at 900-3655 RCF for 5 minutes. The tissue is decanted through a sieve and any cell pellet, liquid, or oil is discarded. Gently mix the adipose tissue in the centrifuge tube, return it to the centrifuge, and centrifuge for 3 minutes at 900-3655 RCF. Decant the tissue again and discard any cell pellet, liquid, or oil. Visual inspection and absorption measurement tests may be repeated as necessary until the product is acceptable. Record the final test results.

[0051] Step 107 involves packaging the product. Packaging involves using a loading tool to load the product into the containers. The loading tool is fully set up and the pistons on the loading tool are used to load the tissue product into the containers, backfilling each container. The containers are then capped. A torque wrench may be used to tighten each cap. The loading step is repeated until all product is packaged into containers. All containers may be inspected for compliance with product specifications, cosmetic quality, air pockets, and completeness of fill.

[0052] In some embodiments, each container is packaged within a pouch. The pouch may be a foil or foil chevron pouch. Each container may be packaged with the cap facing the chevron and away from the opening of the pouch. Each pouch may be vacuum sealed. Each seal may be inspected to ensure it is uniform across the width of the pouch, wrinkle-free, and complete. A unique allograft ID label number may be affixed to each pouch.

[0053] In some embodiments, each pouch may be packaged inside a pouch having a larger width and length. The inner pouch may be vacuum sealed. A single sealed pouch with a syringe inside may be placed inside the larger pouch. In some embodiments, the larger pouch may be a polyester or Tyvek chevron-shaped pouch. The smaller pouch may be placed inside the larger pouch so that the chevrons are aligned. Each larger pouch may be heat-sealed. Each seal may be inspected to ensure it is uniform, wrinkle-free, and complete across the width of the pouch. [Example]

[0054] Experimental Example The present invention will now be described with reference to the following examples, which are provided for illustrative purposes only and the present invention should in no way be construed as being limited to these examples, but rather as encompassing any and all variations that become evident as a result of the teachings provided herein.

[0055] Without further description, it is believed that one skilled in the art can, using the preceding description and the following illustrative examples, make and use the present invention and practice the claimed methods. The following working examples therefore specifically illustrate preferred embodiments of the present invention, and are not to be construed as limiting in any way the remainder of the disclosure.

[0056] Experimental Example 1 Soft tissue defects can cause loss of normal tissue function, trauma, and pain. Cosmetic or reconstructive surgery is needed, but current procedures have variability in volume retention, vascularization induction, and complete wound healing. Minimally engineered adipocyte products (MMAPs) offer a non-surgical approach to cosmetic or reconstructive surgery that may overcome these issues. The study conducted here evaluated the safety, biocompatibility, and volume retention of the present invention (minimally engineered adipocyte filler allografts, hereafter referred to as "allografts"). Results showed that the allografts were biocompatible with human systems, as observed in vitro by suppressed T-cell proliferation and viability, and no significant immune responses or safety risks were observed in vivo when the allografts were implanted in athymic mice for 3 months. Furthermore, in vivo allograft implants demonstrated complete wound closure, and all mice remained in good health throughout the study. Allograft implants retained their matrix and exhibited increased vascularization and local adipose tissue organization, as observed by histopathology. Finally, allograft volume retention was observed in vivo, with high-volume implants (200 μl) maintaining visible implant volume for up to three months. Our findings indicate that allografts are versatile MMAPs that synergistically integrate with surrounding tissues, creating a potential off-the-shelf product for use in cosmetic or reconstructive surgery.

[0057] This study presents the novel human MMAP adipose tissue allograft as an excellent potential volume filler for cosmetic or trauma-based soft tissue replacement. The allograft's natural yellow color and ease of handling for implantation (including its bulk and fluidity) allow for a variety of applications and delivery methods (e.g., surgical implantation or injection). No adverse immune responses were reported in either in vitro or in vivo studies. Mice implanted with either low or high implant volumes experienced no health issues and remained in good health for the entire 3-month duration of the study. High-volume implants maintained better volume retention and were more firm upon removal, but showed signs of tissue encapsulation and potential preliminary signs of fat necrosis, while low-volume implants were resorbed much more rapidly into native tissue. Finally, implants were remodeled, and histopathologists observed evidence of enhanced collagen formation, increased vascularization, and altered local adipose organization. Our findings indicate that the allograft is a versatile human MMAP that interacts synergistically with surrounding tissue, creating a potential off-the-shelf product for surgical use.

[0058] Experimental Example 2 This study evaluated the compositional characteristics of adipose allografts, a connective tissue composed of clusters of cells (adipocytes) surrounding a thick extracellular matrix (ECM) (Mariman, EC, et al., 2010. Cellular and molecular life sciences, 67(8), 1277-129.), including, among others, preadipocytes, fibroblasts, vascular endothelial cells, and macrophages (FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use, July 2020). Adipose tissue provides cushioning and support, particularly for the skin, stores energy in the form of lipids, and insulates the body, among other things (FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use, July 2020), and provides expression and secretion of factors and proteins with important endocrine functions, such as adiponectin, TNFα, IL-6, and other cytokines (Kershaw, EE et al., 2004. The Journal of Clinical Endocrinology & Metabolism, 89(6), 2548-2556). The adipose allograft of the present invention retains the unique natural properties of adipose tissue, namely, adipocyte structure and extracellular matrix.

[0059] The materials and methods for this experiment are described here. Fat allograft samples were analyzed by multiple methods to characterize their composition. Fat allograft samples were subjected to DNA quantification using a DNA assay kit. Quantifying the DNA content of the samples confirms the effectiveness of reducing DNA content after the processing method used to generate the fat allograft. The DNA assay consists of extracting and isolating DNA from the samples using a series of buffers and then purifying it using a spin column. The isolated DNA is then quantified using a spectrophotometer.

[0060] The adipose allograft sample, the previous product iteration adipose allograft, and the unprocessed adipose sample (also referred to as raw) were subjected to compositional analysis to determine the major components of the sample, specifically water, lipids, and solids (including ECM), which were quantifiable on a weight-to-weight (gram) percentage basis, and the results were compared.

[0061] Adipose allograft samples and raw samples were evaluated to determine their structural protein content (including ECM) and protein retention in the allograft compared to native tissue by proteomics performed by a third-party vendor. Proteomics studies help identify proteins in the samples and confirm their presence by mass spectrometry.

[0062] The adipose allograft and live samples were processed for histology, embedded in paraffin, sectioned, stained, and imaged by a third-party vendor. Images of the tissue samples showing microscopic structure were generated to compare the adipose allograft samples with unprocessed samples and to confirm whether structural components were retained in the allograft compared to the native tissue. Samples were stained with Masson's trichrome and hematoxylin and eosin (H&E). Masson's trichrome allows visualization of connective tissue (including the ECM) in the sample. H&E allows visualization of tissue structure and DNA in the nuclei of adipocytes.

[0063] Next, we discuss experimental results. During the development of adipose allografts, it was essential that the process include a series of steps to improve the reduction of cellular and allogeneic components through decellularization while preserving structural components such as the extracellular matrix (ECM). Decellularization is the process of removing any allogeneic or xenogeneic cellular antigens from tissue that may induce an immune response (Bruyneel AAN, et al., Artif Organs. 2017). Insufficient reduction of cellular and allogeneic components can trigger the innate immune system, which is the part of the immune system that is nonspecific and attacks any substance considered foreign. This can lead to inflammation, which, if severe enough, can result in graft rejection (Kasravi M, et al., Biomater Res. 2023). Crapo et al. stated that decellularization techniques cannot remove 100% of cellular material (Crapo PM, et al., Biomaterials. 2011). Therefore, the development of adipose allografts has focused on reducing the cellular and allogeneic components in the final product compared to native tissue.

[0064] Figure 4 shows the average DNA concentration (both raw and fat allograft samples) across three donors. The results indicate lower DNA concentrations in fat allograft samples compared to raw. Overall, fat allograft samples contained less DNA than their untreated counterparts, as supported by Figure 5. Figure 4 also shows sample variability. One challenge in obtaining a meaningful count of DNA quantity per sample is donor variability. Each donor has unique characteristics that can lead to discrepancies in the data, as seen in Figure 6.

[0065] During the development of adipose allografts, it was useful to understand their compositional characteristics to accurately describe the product and compare it to native tissue or other grafts. For comparison purposes, a compositional analysis was developed to determine the major components of adipose tissue-based samples, specifically water, lipid, and solid (including ECM) components. The average sample component amounts obtained for each form of adipose tissue-based sample, i.e., the first and second sample embodiments and raw, are shown in Figures 6, 7A, and 7B. Variability was observed between samples of the same type, which may be due to donor and sample variability, as each donor is unique in composition and sample homogenization is not always achieved. The results appear to indicate a relationship between solid components and water content. This may be due to the hydrophilic nature of proteins in the ECM, which favors adhesion (Yang, L. et al., RSC Advances, 2017). On average, the proportion of solid components (including ECM) increased in the samples of Example 1 compared to Example 2. In future studies, it is recommended to increase the number of samples analyzed to increase the reliability of the results.

[0066] During the development of adipose allograft processing, nonionic surfactants were used as mild detergents to reduce cellular and allogeneic components while avoiding denaturation and washout of structural proteins. Proteomic studies were used to confirm the presence of structural proteins in adipose allografts compared to native adipose tissue. As shown in Figures 8A and 8B, the following proteins were retained in four different allograft samples compared to native tissue from the same donor: adipose allografts retain the major adipose collagens (COL4, COL6) as well as the major fibrillar collagens (COL1, COL3) and minor fibrillar collagen (COL5). These proteins are involved in the formation of the extracellular matrix (ECM) and are some of the most abundant proteins in adipose tissue (Mori S, et al. Int J Biol Sci. 2014, Ricard-Blum S. Cold Spring Harb Perspect Biol. 2011). Adipose allografts also retain glycoproteins, including key adipose ECM proteins fibronectin and laminin-α, as well as emilin 1 and 2, adiponectin (associated with adipocyte proliferation), and fibrillin 1 (contributing to adipogenesis) (Ricard-Blum S. Cold Spring Harbor Perspective Biol. 2011). Furthermore, adipose allografts retain various proteoglycans. An important component of proteoglycans is glycosaminoglycan (GAG), the major ECM protein in adipose tissue, which is also involved in basement membrane development, water retention, and growth factor transport across the ECM (Pessentheiner AR, et al. 2020).

[0067] During the development of adipose allografts, the grafts are prepared through minimally manipulative processing methods that preserve the adipose tissue architecture in the graft, as in native tissue. Histological images of the tissue samples, displaying the microscopic structure, were collected and compared with images of native adipose tissue and the adipose allograft to confirm that the adipose tissue architecture was preserved. To observe the microscopic structural composition of the tissue, tissue samples were prepared using two distinct stains: Masson's Trichrome and H&E. Masson's Trichrome is a useful stain to use because it targets connective tissues such as collagen, keratin, fibrin, and other structural components (Zhou X, et al. Bio Protoc. 2017). Masson's Trichome stain is a three-color stain that identifies connective tissue with blue / green, cell nuclei with red / purple, and cytoplasm with pink. H&E is a useful stain to use because it targets cell structures and nuclei (Zhou X, et al. Bio Protoc. 2017). H&E is a double stain that identifies cell nuclei stained purple / blue by hematoxylin and cytoplasm stained red / pink by eosin.

[0068] Histological images of the tissue samples are shown in Figures 9-14 and demonstrate that the adipose allografts retained structural components, such as the adipocyte structure (represented by honeycomb-like structures) and connective tissue network (including ECM), comprising the adipose tissue, compared to native tissue. This observation is consistent across the various samples. The histological images shown in Figures 9-14 also demonstrate an increased amount of connective tissue (including ECM) staining relative to the individual adipocyte structures in the adipose allograft samples compared to native tissue. This observation is consistent across the various samples.

[0069] Experimental Example 3 This study evaluated the real-time aging performed on adipose tissue-based research allografts. Real-time aging studies provide data to determine the shelf life of a product and the effects of aging on the material. During the study, the product was placed on a shelf and exposed to environmental changes, such as fluctuations in temperature and humidity, simulating real-world conditions that a product may experience during its life cycle. This study simulated real-time aging conditions to collect data on the aging of adipose tissue-based grafts for research purposes only.

[0070] Materials and methods are described herein. Human adipose tissue was designated for study with appropriate donor consent and permission. Donors were tested and found to be negative for the following infectious diseases: antibodies to human immunodeficiency virus (types 1 and 2), nucleic acid test (NAT) for HIV-1, hepatitis B surface antigen, hepatitis B core antigen antibodies, antibodies to hepatitis C virus, and syphilis. Pretreatment cultures of the collected tissue were obtained and found to be negative for the following pathogenic highly virulent microorganisms: Clostridium; fungi (yeasts, molds); and Streptococcus pyogenes. The adipose tissue was collected in a manner that excluded contamination and / or cross-contamination.

[0071] Transportation, storage, processing, and packaging of the adipose tissue was performed and documented according to the methods disclosed herein. The adipose tissue was processed in a manner that minimized contamination and / or cross-contamination. The grafts were subjected to gamma irradiation at a minimum dose of 15 kGy at a third-party contractor according to their procedures. Radiation dosimetry records were obtained. Following irradiation, the grafts were transported to a location designated for the study at ambient temperature and in stable environmental conditions. Prior to evaluation, the grafts were stored at ambient temperature. Evaluation time points were approximately several months apart.

[0072] At least one graft unit was subjected to bioburden testing prior to irradiation. Additional bioburden testing was performed on at least one graft after irradiation to confirm the absence of microbial contamination. A minimum of one graft unit was evaluated at selected time points. Each graft tissue was assigned to maximize the number of evaluations. All evaluations were voluntary, dependent on graft availability. Quality control testing was performed in-house. Grafts were prepared for histology imaging to assess the structural characteristics of the product on a microscopic scale and potential changes over time. Mechanical testing was performed. In vitro cell viability assays were performed in-house to evaluate whether test samples could affect cell viability of either NIH / 3T3 or NCTC clone 929 (L cells, L-929, a derivative of strain L) mouse fibroblast cell lines.

[0073] The results of the experiments are described here. Bioburden testing was performed on adipose allograft study samples from four different donor lots prior to irradiation. Bioburden testing was also performed on adipose allograft study samples from two different donor lots after irradiation.

[0074] Quality control testing, histology, mechanical assays, and in vitro viability assays were performed throughout the study to evaluate the product's structural integrity, free lipid content, cell viability, and overall shelf life. Pre-irradiation bioburden testing indicated that no more than 14 CFU of contamination was present in the samples. Post-irradiation bioburden testing confirmed the absence of microbial contamination after irradiation, with no growth results. A total of 20 samples were evaluated for quality control testing of tissue size at various time points, from 0 to nearly 6 months. All samples passed evaluation at various time points. Results suggest that the product's tissue size remained consistent over time, up to nearly 6 months.

[0075] A total of 22 samples were evaluated for color quality control at various time points, from 0 to approximately 6 months. All samples passed the evaluation at various time points. The results suggest that the color of the product is acceptable over a period of up to approximately 6 months.

[0076] A total of 22 samples were evaluated for quality control testing of free lipid content at various time points, from 0 to nearly 6 months. All samples evaluated at 0 to 4 months passed the evaluation. Future studies may include evaluating triglyceride and glycerol content over time to quantify potential changes in free lipid content and trends.

[0077] Adipocytes and connective tissue structures were observable in histological images from samples stained with H&E and Masson's trichrome across all time points. Histological examination was performed on the samples. This histological data suggests that the adipose allograft samples retained their native adipose structure over time. Figures 15-17 show images of H&E-stained adipose allograft samples from a single donor at 2, 3, 4, and 6 months. Figure 18 shows images of adipose allograft samples stained from another donor at 2, 3, and 4.5 months. Figure 19 shows images of adipose allograft samples stained from another donor at 1, 1.5, and 2.5 months. Adipocytes and connective tissue structures were observable in most histological images from samples stained with Oil Red O across all time points. Histological images showing poor sample structure were observed to be prepared in the same order, suggesting poor sample preparation techniques from the histology laboratory.

[0078] The results of the mechanical assay revealed that a slight trend toward a decrease in mechanical stiffness of the adipose allograft samples appeared to be observed. The mechanical stiffness of the two separate donor samples, along with the untreated samples, was measured over time. However, there is no conclusive statistical evidence supporting this trend because the sample sizes per time point and donor were not large enough to perform statistical analysis.

[0079] A total of eight samples were evaluated in an in vitro cell viability assay at various time points, from 0 to approximately 6 months. An in vitro cell viability assay was performed on the various samples. All samples passed the assay, suggesting that fibroblast survival or growth was not adversely affected by the test samples up to approximately 4 months. Four months was chosen as the maximum time point because at least three samples were assayed at this time point, compared to a single sample at 6 months.

[0080] Experimental Example 4 This study evaluated the differentiation of adipose-derived stem cells (ASCs) into adipocytes and determined whether the presence of a test sample of tissue product altered the ASC differentiation characteristics over control or other test samples.

[0081] Adipose-derived stem cells (ASCs) have attracted increasing interest for therapeutic applications in regenerative medicine due to their multiple lineage attributes: ASCs are poorly immunogenic, have the capacity to self-renew, can migrate to injury sites, act via autocrine and paracrine pathways, and are multipotent (they can differentiate into adipocyte, osteoblast, and chondrocyte lineages).

[0082] Materials and methods are described here. ASCs were thawed from liquid nitrogen and prepared for seeding into T150 culture flasks. After the T150 flasks reached 90% confluency, 6,579 cells / cm were plated. 2The inserts were subcultured into individual wells of a 24-well plate. Designated cells were fed with control stem medium without the test sample, control stem medium supplemented with the test sample, differentiation medium without the test sample, differentiation medium supplemented with the test sample, conventional medium containing the control stem medium, and conventional medium containing the differentiation medium. The stem medium was replaced every 2–3 days until the cells in the wells designated as differentiation wells reached 85–90% confluency. After reaching confluency, the control stem medium was replaced with differentiation medium at the same feeding frequency. The day after seeding, inserts containing the test sample were added. Photographs were taken 24 hours, 48 ​​hours, 1 week, and 2 weeks after addition of the inserts. After 2 weeks, the cells were examined to determine whether differentiation had occurred. The cells were treated with fixative and then stained with Oil Red O to observe lipid formation. The Oil Red O was then eluted and read using a spectrophotometer.

[0083] The results are as follows. After two weeks of culture, the cells were stained with Oil Red O and evaluated for lipid formation. Figures 23-32 show the results after two weeks. Figure 23: Cells in control medium with no test sample added. Figure 24: Differentiation wells (also control wells) with no test sample added. Figure 25: Cells with control medium and Donor 1 test sample. Figure 26: Cells with control medium and Donor 2 test sample. Figure 27: Cells with control medium and Donor 3 test sample. Figure 31: Cells with control medium and conventional test sample. Figure 28: Cells with differentiation medium and Donor 1 test sample. Figure 29: Cells with differentiation medium and Donor 2 test sample. Figure 30: Cells with differentiation medium and Donor 3 test sample. Figure 32: Cells with differentiation medium and conventional sample. Figures 33 and 34 show the average absorbance, expressed as optical density (OD) at 510 nm, of test samples of embodiments of the present invention versus conventional products using a spectrophotometric assay. The level of OD correlated with the amount of Oil Red O staining in ASCs differentiated into adipocytes. All ASC groups grown in differentiation medium were found to have substantial differentiation into adipocytes, as indicated by lipid formation identified by Oil Red O staining. No significant differences in ASC differentiation into adipocytes were observed between ASCs grown in the presence of test sample adipose allografts containing differentiation medium (mean OD 1.5 ± 0.63 SD) and cells grown in differentiation medium without test sample (mean OD 0.87 ± 0.24 SD). No significant differences in ASC differentiation into adipocytes were observed between ASCs grown in the presence of conventional test sample with differentiation (mean OD 1.04 ± 0.09 SD) and cells grown in differentiation medium without test sample (mean OD 0.87 ± 0.24 SD). The results also suggest that ASC differentiation into adipocytes may be aided by the presence of test sample adipose allografts and conventional products, with slightly better differentiation observed in the presence of adipose allografts.

[0084] Experimental Example 5 In this experiment, the transwell paracrine effect of test samples on endothelial cell proliferation was determined. Primary human dermal blood endothelial cells (HDBECs) are a subpopulation of human dermal endothelial cells. They are isolated from the dermis of adult skin from a single donor. Cells are analyzed for CD31 positivity and podoplanin negativity by flow cytometry analysis. Blood endothelial cells have important functions in physiological processes such as vascular tone, capillary permeability, blood clotting, fibrolysis, and angiogenesis. Primary HDBECs were chosen for this assay due to their use in tissue engineering models (Pappalardo A et al., 2023. Sci Adv).

[0085] Materials and methods are described here. To determine the effect on cell proliferation, 7,895 cells / cm 2 Primary HDBECs were seeded into 24-well plates and grown in complete growth medium in the presence of test samples. After 3 days of culture, cells were stained with crystal violet (CV) and assessed for changes in total cell number. Positive and negative control wells were set up using complete growth medium versus basal medium, respectively, to observe cell proliferation of primary HDBECs in the absence of test samples. Cell proliferation was predicted from wells with primary HDBECs grown in complete growth medium, thus establishing the assay positive control. Cell proliferation was not predicted from wells with primary HDBECs grown in basal medium, thus establishing the assay negative control. After 3 days of culture, cells were stained with CV and assessed for changes in total cell number.

[0086] (A) Test well with test sample identified as "conventional" before test sample was added, then (B) 24 hours, (C) 48 hours, (D) 72 hours, and (E) 1% CV staining after the test sample was introduced into the insert. All three test wells for the conventional sample were similar in appearance, and a representative image from one well is shown in Figure 35.

[0087] Test wells with test samples identified as embodiments of the invention (A) before the test sample was added, then (B) 24 hours, (C) 48 hours, (D) 72 hours, and (E) after introducing the insert with the test sample at 1% CV staining. All three test wells of one embodiment of the invention were similar in appearance, and a representative image from one well is shown in Figure 36.

[0088] A) A control well with primary HDBECs and complete growth medium, identified as a "positive control," at 1% CV staining before the test sample was introduced into the test well, and then at B) 24 hours, C) 48 hours, and D) 72 hours after the insert with the test sample was introduced into the test well. All three control wells were similar in appearance, and a representative image from one well is shown in Figure 37.

[0089] A) Control well with primary HDBECs and basal medium, identified as a "negative control," before the test sample was introduced into the test well, and then B) 24 hours, C) 48 hours, and D) 72 hours after the insert with the test sample was introduced into the test well at 1% CV staining. All three control wells were similar in appearance, and a representative image from one well is shown in Figure 38.

[0090] The results are described below with reference to Figures 39-42. The data shown in the tables represent absorbance values ​​read from a microplate reader (CLARIOstar). After 3 days of culture, cells were stained with CV for changes in cell number. Optical density (OD) levels indicate the amount of CV dye eluted from fixed cells (absorbance -590 nm). No significant differences in total cell number were observed between primary HDBECs grown in the presence of test sample adipose allografts (mean OD 4.96 ± 0.71 SD) and primary HDBECs grown in complete medium without test sample (mean OD 4.92 ± 0.61 SD). These results suggest that the presence of test sample adipose allografts does not cause a negative effect on the cell proliferation of primary HDBECs in vitro. No significant differences in cell proliferation were observed between the conventional test sample and the adipose allograft test sample. The test samples (both adipose allograft and conventional) did not have a negative effect on the cell proliferation of primary HDBECs in vitro. No significant differences in cell proliferation were observed between the conventional and adipose allograft test samples. A significant difference in cell proliferation of primary HDBECs was observed, as expected, in the positive control (mean OD 4.92 ± 0.16 SD) compared to the negative control (mean OD 0.90 ± 0.61 SD). Primary HDBEC proliferation was observed in the positive control. Primary HDBEC proliferation was not observed in the negative control.

[0091] Experimental Example 6 The purpose of this experiment is to quantify the percent adipocyte composition for the treated tissue embodiments described herein.

[0092] Human cells, tissues, and cell- and tissue-based products (HCT / Ps) are defined in Title 21 of the Code of Federal Regulations (CFR) Part 1271 as products containing or consisting of human cells or tissues intended for implantation, transplantation, infusion, or transfer into a human recipient (FDA 2020). HCT / Ps are regulated only under Section 361 of the PHS Act if all stated criteria are met. One of the criteria for HCT / P regulation is that tissue processing must maintain minimal manipulation.

[0093] Minimal manipulation is defined in 21 CFR 1271.10(a) as processing that does not alter the original relevant properties of tissue related to the tissue's utility for reconstruction, repair, or replacement. Adipose tissue is composed primarily of adipocytes and surrounding connective tissue. FDA classifies the original relevant properties as primarily providing cushioning and support to the body. Therefore, to evaluate whether the manufacture of an adipose-derived HCT / P meets the criteria for minimal manipulation, consideration should be given to whether the processing alters the HCT / P's utility to provide cushioning and structural support (FDA 2020).

[0094] For example, the production of adipose-derived HCT / Ps in which cellular components have been completely removed from the surrounding connective tissue (decellularized) is considered more common than those minimally engineered by the FDA. This is because the decellularization process alters their ability to provide cushioning and support. To demonstrate that tissues processed according to embodiments described herein maintain the criteria for minimal manipulation, this report demonstrates how adipocyte structure and connective tissue content are preserved.

[0095] One of the most distinguishing characteristics of processed tissue is the retention of structurally intact adipocytes. To support the argument for maintaining minimal manipulation, quantification of adipocyte composition was performed for each product. In adipose tissue, hematoxylin and eosin (H&E) staining is a useful tool for distinguishing the surrounding connective tissue from adipocytes. Quantitative assessment of the structure of processed tissue was achieved using the adiposoft plugin for the Fiji adaptation of Image J analysis software (Figure 43). The adiposoft plugin is designed to identify individual adipocytes and calculate the inner area of ​​each, at which point the software generates a report. The percent adipocyte composition for each sample can be calculated by taking the ratio of adipocyte area to total sample area (Equation 1, Figure 44).

[0096] Adipocyte Structure Quantification: The primary goal of this experiment was to quantify the percent adipocyte composition content in the adipose allograft samples. This was accomplished by histological staining and Image J analysis.

[0097] Terminology: HCT / P (Human Cell and Tissue Product) H&E (hematoxylin and eosin) Materials (including but not limited to): formalin 5mL conical tube Nitrile gloves forceps procedure: (1) Five tissues were prepared from five tissue donors to account for donor variability. (2) After manufacturing, samples were taken from each product replicate, fixed in formalin in 5 mL conical tubes, and subjected to H&E histological staining. (3) For each histology slide, five images were captured for each sample to obtain universal characterization (Figure 45). (4) For each sample staining, all five images were analyzed by Image J software to measure the adipocyte area and total sample area. The average adipocyte composition percentage was calculated by Equation 1.

[0098] result: 46-48 show H&E staining images of the first and second processed tissues, respectively, processed according to embodiments described herein (FIGS. 46 and 47), and conventionally processed tissue (FIG. 48). For each sample stain, five sections were analyzed using Image J software, and the average percent adipocyte composition was calculated.

[0099] Table 1 (Figure 49) shows the average results for the first tissue sample, and Table 2 shows the average results for the second tissue sample. Each product evaluation was sampled from five separate donors. The adipocyte percentage range for the first tissue embodiment was calculated to be approximately 45-70%, while the range for the second tissue embodiment was determined to be 30-55%. The difference in adipocyte content may be due to variations in the number of washing steps between the two processing procedures.

[0100] Consideration Comparison of H&E staining reveals that both the first and second tissue products contain a mixture of extracellular matrix (ECM) components (pink) and intact adipocytes (pink circles surrounding white spaces), whereas the conventional tissue exhibits only ECM elements within their products. This unique feature distinguishes the first and second tissue products from the conventional third product and demonstrates their useful advantage in providing the core properties of fat: cushioning and support. Furthermore, this report provides evidence of the first / second tissue's compliance with HCT / P minimum handling requirements through preservation of adipocyte structure and connective tissue content. No processed tissue is available that demonstrates the retention of both intact adipocytes and connective tissue content. Thus, both the first and second allograft products incorporate unique structural features not previously exploited.

[0101] FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use(July 2020). Rasband, WS, ImageJ, US National Institutes of Health, Bethesda, Maryland, USA.

[0102] Experimental Example 7 The purpose of this experiment was to evaluate the adipose tissue composition of tissue products processed according to embodiments described herein. The tissue composition was assessed by measuring the water content, lipid content, and protein content.

[0103] Adipose tissue is composed of connective tissue and various cell populations, including adipocytes, preadipocytes, endothelial cells, blood cells, fibroblasts, pericytes, and macrophages, and is increasingly recognized as a key player in metabolic regulation (Luo 2016). Connective tissue is primarily composed of different types of extracellular matrix (ECM) proteins, including collagens, glycoproteins, and proteoglycans, which are particularly involved in maintaining structural integrity. The ECM also regulates various physiological processes, including cell attachment, differentiation, survival, and development. Furthermore, evidence suggests that the ECM network is necessary for tissue remodeling and repair (Ruiz-Ojeda 2019).

[0104] Generally, the three most abundant components of adipose tissue are lipids, ECM proteins, and water. Over the years, numerous studies have been conducted to measure component quantification in adipose tissue. McKee et al. conducted a meta-analysis review of various papers reporting proteomic estimates of the absolute and relative abundance of ECM proteins in adipose tissue (McKee 2019). In addition, the authors analyzed empirical data to estimate the amounts of other tissue components, including water and lipid content. These amounts were combined with the relative amounts of ECM protein content to determine tissue composition (Figure 50). According to McKee et al., adipose tissue is typically composed of 75–80% lipid, 15–20% water, and 3–5% protein (McKee 2019). The authors' findings of fat component quantification are used as a benchmark for the expected proportions of all three components in adipose allograft samples.

[0105] result: For this experiment, three processed tissues according to one allograft embodiment and three processed tissues according to a second allograft embodiment were obtained from three tissue donors. Each donor was aliquoted and processed to create both a first product and a second product. Table 1 (Figure 51) shows the water, lipid, and protein content percentages for the first allograft embodiment samples tested. The bottom row of Table 1 shows the percentage range for each component: 19-47% water content, 47-80% lipid content, and 2-7% protein content. Figure 52 shows the percent composition for the first product produced from tissue donor 1. Table 2 (Figure 53) shows the component quantification for the second product samples. The percentages for the second product samples were as follows: 15-36% water content, 60-84% lipid content, and 2-4% protein content.

[0106] Discussion of results The results show that the first and second tissue samples have the same component quantification, which is similar to the published literature on adipose tissue, but with some slight differences.Compared to the second product sample, the first product sample shows slightly higher water content, but lower lipid content.In addition, the first product sample shows a higher protein content range.

[0107] Discrepancies were noted in the compositional analysis of Donor 3. Because the calculated composition percentages for this donor did not match the ranges for Donors 1 and 2, the test was repeated for Donor 3 to ensure accuracy of the results, which were confirmed to be accurate based on the results of the second reading. The data demonstrate that Donor 3 had a greater amount of connective tissue compared to the other two donors, which may also be responsible for the increased water content in the sample. Additionally, Donor 3 exhibits a higher protein content compared to Donors 1 and 2. Based on this information, the compositional range was reported as a single average value to account for donor variability.

[0108] FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use(July 2020).

[0109] McKee TJ, Perlman G, Morris M, Komarova SV.Extracellular matrix composition of connective tissues: a systematic review and meta-analysis.Sci Rep.2019 Jul 22;9(1):10542.doi:10.1038 / s41598-019-46896-0.PMID:31332239;PMCID:PMC6646303.

[0110] Luo L,Liu M.Adipose tissue in control of metabolism.J Endocrinol.2016 Dec;231(3):R77-R99.doi:10.1530 / JOE-16-0211.PMID:27935822;PMCID:PMC7928204.

[0111] Ruiz-Ojeda FJ, Mendez-Gutierrez A, Aguilera CM, Plaza-Diaz J.Extracellular Matrix Remodeling of Adipose Tissue in Obesity and Metabolic Diseases.Int J Mol Sci.2019 Oct 2;20(19):4888.doi:10.3390 / ijms20194888.PMID:31581657;PMCID:PMC6801592.

[0112] The disclosures of any and all patents, patent applications, and publications cited herein are incorporated herein by reference in their entireties. While the present invention has been disclosed with reference to specific embodiments, it will be apparent that other embodiments and variations of the invention may be devised by those skilled in the art without departing from the true spirit and scope of the invention.

Claims

1. 1. Processed adipose tissue, comprising: an extracellular matrix derived from human adipose tissue; and a plurality of adipocytes having at least 10% intact adipocyte composition.

2. 2. The processed adipose tissue of claim 1, wherein the intact adipocyte composition is at least 30%.

3. 2. The processed adipose tissue of claim 1, wherein the intact adipocyte composition is between 30% and 70%.

4. 2. The processed adipose tissue of claim 1, wherein the intact adipocyte composition is between 30% and 55%.

5. 2. The processed adipose tissue of claim 1, wherein the intact adipocyte composition is between 45% and 70%.

6. The processed adipose tissue of claim 1 , further comprising a plurality of structural proteins including at least one of collagen, glycoproteins, and proteoglycans retained from the human adipose tissue.

7. The processed adipose tissue of claim 1 , further comprising decellularized tissue having reduced amounts of allogeneic components, cellular components and free lipids.

8. 10. The processed adipose tissue of claim 1 configured for implantation or injection into a patient.

9. 10. The processed adipose tissue of claim 1, configured to pass through an opening of between 1 mm and 4.5 mm.

10. 1. A method for treating adipose tissue, comprising: Freezing and thawing human adipose tissue; separating the human adipose tissue into small pieces by mechanical comminution; rinsing the human adipose tissue successively with a non-ionic surfactant, a crystalloid solution, and sterile water; and dispensing said human adipose tissue into a package using a loading tool.

11. The loading tool comprises: a jar having a proximal end opening, a distal end opening, and a chamber disposed therebetween; a piston configured to be actuated by a lever to slide distally and flush along the wall of the chamber; a stand configured to stabilize the piston below the lever and form a hinged connection with the lever.

12. The method of claim 11 , wherein the proximal end opening and the distal end opening are each at least partially surrounded by a threaded surface.

13. The method of claim 11 , wherein the loading tool further comprises an adapter configured to connect to at least one of the proximal and distal openings, the adapter having a first sealable port.

14. The method of claim 13 , wherein the piston has a second sealable port.

15. 11. The method of claim 10, further comprising debridement of the human adipose tissue to remove extraneous or damaged tissue.

16. The method of claim 10, further comprising rinsing the human adipose tissue with a non-ionic surfactant, sterile saline, and sterile water by distributing the human adipose tissue into a centrifuge tube and centrifuging the human adipose tissue.

17. The method of claim 10, further comprising rinsing the human adipose tissue with a non-ionic surfactant, sterile saline, and sterile water by distributing the human adipose tissue into a sealable container and gently shaking the container.

18. 11. The method of claim 10, further comprising rinsing the tissue at least twice in a non-ionic detergent.

19. 11. The method of claim 10, further comprising rinsing the tissue in crystalloid saline at least twice.

20. 11. The method of claim 10, further comprising rinsing the tissue in sterile water at least twice.

21. 11. The method of claim 10, further comprising drying the human adipose tissue in a centrifuge tube containing a drying sieve.

22. 11. The method of claim 10, further comprising decanting the tissue after each rinse and removing any decanted liquid and cellular components.

23. The method of claim 10, further comprising using the loading tool to evenly distribute the human adipose tissue into a container.

24. 20. The method of claim 19, further comprising packaging the container in a sterilization pouch.

25. 1. A media loading tool comprising: a jar having a proximal end opening, a distal end opening, and a chamber disposed therebetween; a piston configured to be actuated by a lever to slide distally and flush along the wall of the chamber; a stand configured to stabilize the piston under the lever and form a hinge connection with the lever.

26. 26. The medium loading tool of claim 25, wherein the proximal end opening and the distal end opening are each at least partially surrounded by a threaded surface.

27. 26. The medium loading tool of claim 25, further comprising an adapter configured to connect to a distal opening having the first sealable port.

28. 26. The medium loading tool of claim 25, wherein the piston has a second sealable port.

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