Biological tissue and method of manufacture

Abstract

A treated adipose tissue having an extracellular matrix derived from human adipose tissue and adipose cells having a composition of at least 10% intact adipose cells is described. A method of treating adipose tissue and a media loading tool are also described.

Description

[0001] Cross - Reference to Related Applications

[0002] This application claims priority to U.S. Provisional Application No. 63 / 383,574, filed on November 14, 2022, and U.S. Provisional Application No. 63 / 383,578, filed on November 14, 2022, the entire contents of both of which are incorporated herein by reference. Background Art

[0003] The field of aesthetic or reconstructive surgery typically results from diseases, traumas, or injuries that damage local soft tissues. These defects can lead to loss of normal tissue function, emotional trauma, and pain (Coleman, S.R. 2016. Plastic and Reconstructive Surgery 118, 108S - 120S). Alternatively, aesthetic applications include small - volume (facial contouring) or large - volume (breast and buttocks) treatments using synthetic or natural fillers. Current clinical procedures often use autologous fat transfer or commercially available fillers to aid in repair (Coleman, S.R. 2016. Plastic and Reconstructive Surgery 118, 108S - 120S., Gir, P. et al. 2012. Plastic and Reconstructive Surgery 130, 249 - 258). Autologous fat transfer uses the patient's own adipose tissue, via liposuction, to fill voids while minimizing host immune rejection and promoting adipogenesis (Serra - Renom, J.M. et al. 2010. Plastic and Reconstructive Surgery 125, 12 - 18). Although advantageous, the use of heterogeneous, untreated fat can lead to donor - site morbidity due to liposuction (Patrick Jr, C.W. 2001. The Anatomical Record 263, 361 - 366.), poor survival of the remaining fat cells, which can lead to necrosis (Young, D.A. et al. 2012. Biomed Mater 7, 024104.), and the delivery of free lipid and tissue fragments can cause cyst formation and calcification (Kokai, L.E. et al. 2020. Plast Reconstr Surg Glob Open 8, e2574.). These complications can result in poor aesthetic outcomes, depending on the injection site, such as contour irregularities, tissue bunching, inflammation, or even more severe problems such as fat embolism, stroke, and / or blindness (Vasavada, A. et al. 2023. Autologous Fat Grafting For Facial Rejuvenation., StatPearls Publishing LLC).

[0004] Due to the increasing demand for cosmetic surgery, commercially available fillers have become increasingly popular, and recent research has provided insights into the current progress in this field (Choi, J.H. et al. 2010. Tissue Eng Part B Rev 16, 413-426). Synthetic and natural materials have been widely used as soft tissue fillers, but with limited success. Synthetic polymers, such as polyethylene glycol or polylactic acid, offer injectability and / or compositional control and are biologically inert, resulting in reduced host integration, 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, K.J. et al. 2004. Plastic and Reconstructive Surgery 113, 260-269., Patrick, C.W. et al. 1999. Tissue Eng 5, 139-151).

[0005] However, synthetic polymers lack the appropriate native tissue composition and mechanical properties. In addition, biodegradable synthetic polymers exhibit poor mechanical properties (Young, D.A. et al. 2012. Biomed Mater 7, 024104.), poor volume retention (Patrick Jr, C.W. 2001. The Anatomical Record 263, 361-366.), and the biodegradable compounds can accumulate, causing an immune response or even toxicity (Fu, K. et al. 2000. Pharmaceutical Research 17, 100-106., K. et al. 1996. Biomaterials 17, 457-461).

[0006] To better recapitulate native tissue, natural fillers containing extracellular matrix (ECM) have shown increased volume retention, host integration, reduced immune response, and adipogenesis over time for in vitro (Flynn, L. E. 2010. Biomaterials 31, 4715 - 4724.), in vivo (Young, D. A. et al. 2012. Biomed Mater 7, 024104., Han, T. T. Y. et al., 2015, Biomaterials 72, 125 - 137.), and clinical studies (Gold, M. H. et al. 2020. J Cosmet Dermatol 19, 1044 - 1056., Dige A. et al. 2019. Gastroenterology 156, 2208 - 2216e2201). Natural ECM retains the 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 adipose tissue with minimal manipulation that retains native ECM as an adipose tissue allograft (Banyard, D. A. et al. 2019. Regenerative Medicine and Plastic Surgery: Skin andSoft Tissue, Bone, Cartilage, Muscle, Tendon and Nerves, 71 - 89, SpringerInternational Publishing.).

[0007] However, the use of chemical, biological, and physical treatments in the fabrication of tissues that retain ECM can lead to damage to ECM proteins, structure, or potentially damage the sample. There is a need in the art for injectable processed tissues with minimal manipulation that retain native adipocyte structure and ECM. The embodiments described herein meet this need. SUMMARY OF THE INVENTION

[0008] In one embodiment, the processed adipose tissue has an extracellular matrix derived from human adipose tissue and adipocytes having at least 10% intact adipocytes. In one embodiment, at least 30% of the intact adipocytes. In one embodiment, the intact adipocytes are between 30% and 70%. In one embodiment, the intact adipocytes are between 30% and 55%. In one embodiment, the intact adipocytes are between 45% and 70%. In one embodiment, the tissue structural proteins include at least one of collagen, glycoprotein, and proteoglycan retained from the human adipose tissue. In one embodiment, the tissue comprises a decellularized tissue having a reduced amount 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 between 1 mm and 4.5 mm. In one embodiment, a method of processing adipose tissue comprises the steps of: freezing and thawing human adipose tissue, separating the human adipose tissue into pieces by mechanical grinding, continuously rinsing the human adipose tissue with a nonionic surfactant, a crystalloid solution, and sterile water, and dispensing the human adipose tissue into a package using a loading tool. In one embodiment, the loading tool comprises: a wide-mouth bottle having a proximal end opening, a distal end opening, and a chamber disposed between the proximal end opening and the distal end opening; a piston actuated by a lever and configured to slide distally and flush along the wall of the chamber; and a bracket configured to stabilize the piston under the lever and form a hinge 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 opening and the distal opening, the adapter having a first sealable port. In one embodiment, the piston has a second sealable port. In one embodiment, the method comprises debriding the human adipose tissue to remove foreign or abraded tissue. In one embodiment, the method comprises rinsing the human adipose tissue with a nonionic 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 comprises rinsing the human adipose tissue with a nonionic surfactant, a sterile crystalloid solution, and sterile water by dispensing the human adipose tissue into a sealable container and gently agitating the container. In one embodiment, the method comprises rinsing the tissue in a nonionic 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 comprising 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 using the loading tool to evenly distribute the human adipose tissue in a container. In one embodiment, the method includes packaging the container in a sterile pouch.

[0009] In one embodiment, the media loading tool includes a wide-mouth bottle having a proximal end opening, a distal end opening, and a chamber disposed between the proximal end opening and the distal end opening; a piston actuated by a lever and configured to slide distally and flush along the wall of the chamber; and a bracket 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 opening and the distal opening, the adapter having a first sealable port. In one embodiment, the piston has a second sealable port. BRIEF DESCRIPTION OF THE DRAWINGS

[0010] Referring to the following description and drawings, the foregoing and other objects and features will become apparent, the description and the drawings being included to provide an understanding of the present invention in accordance with various embodiments and constituting a part of this specification, wherein like reference numerals represent like elements, and wherein:

[0011] Figure 1A A flowchart depicting an embodiment of a method for producing an adipose tissue filler allograft according to one embodiment is shown, Figure 1B A flowchart depicting a method for producing an adipose tissue filler allograft according to one embodiment is shown, and Figure 1C A flowchart depicting a method for processing adipose tissue according to one embodiment is shown.

[0012] Figure 2A and Figure 2B A perspective side view of a loading tool assembly according to one embodiment is shown.

[0013] Figure 2C A cross-sectional view of a loading tool wide-mouth bottle according to one embodiment is shown.

[0014] Figure 2D and Figure 2E A perspective view of a first adapter according to one embodiment is shown, and Figure 2F A perspective view of a second adapter according to one embodiment is shown.

[0015] Figure 3ADepicts a visual inspection chart for color inspection according to an embodiment.

[0016] Figure 3B Depicts a visual inspection chart for excess free lipid inspection according to an embodiment.

[0017] Figure 4 Depicts a graph showing DNA quantification results according to an embodiment.

[0018] Figure 5 Describes a graph showing the average DNA content in all samples tested according to an embodiment.

[0019] Figure 6 Depicts a table showing the average component quantification in percentage according to an embodiment.

[0020] Figure 7A and Figure 7B Depicts the component amounts of water, lipid, and solids (including ECM) in grams according to an embodiment.

[0021] Figure 8A Depicts a graph showing the relative amount of ECM protein expression between an embodiment and native adipose tissue. Figure 8B Depicts a heat map showing the major ECM components expressed in an embodiment compared to native adipose tissue.

[0022] Figure 9 A- Figure 9 D depict four histological images from adipose tissue. Figure 9 A and Figure 9 B depict the staining from untreated tissue. Figure 9 C and Figure 9 D depict the staining according to an embodiment.

[0023] Figure 10 A- Figure 10 D depict four histological images from adipose tissue. Figure 10 A and Figure 10 B depict the staining from untreated tissue. Figure 10 C and Figure 10 D depict the staining from an embodiment.

[0024] Figures 11 - 14 Depicts four histological images from adipose tissue. Figure 11 and Figure 12 Depicts the staining of untreated tissue. Figure 13 and Figure 14 Depicts the staining from an embodiment.

[0025] Figure 15-Figure 17 Depicts histological staining images of the samples of the present invention at different time points. Figure 15A Shows the staining at 2 months. Figure 15B Shows the staining at 3 months. Figure 16 Shows the staining at greater than 4 months. Figure 17 Shows the staining at 6 months.

[0026] Figure 18 Depicts histological staining images of the samples of the present invention at different time points. Figure 18 A shows the staining at greater than 2 months. Figure 18 B shows the staining at greater than 3 months. Figure 18 C shows the staining at 4.5 months.

[0027] Figure 19 Depicts histological staining images of the samples of the present invention at different time points. Figure 19 A shows the staining at less than 1 month. Figure 19 B shows the staining at 1.5 months. Figure 19 C shows the staining at 2.5 months.

[0028] Figure 20 Depicts a graph showing the mechanical hardness of various samples of the present invention over time.

[0029] Figure 21 Shows the mechanical hardness of a sample of the present invention from one donor over time.

[0030] Figure 22 Shows the mechanical hardness of a sample of the present invention from a second donor over time.

[0031] Figure 23 Depicts images of the staining of human ASC samples grown in control medium in the absence of test samples at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and (E) at 2 weeks with Oil Red O staining.

[0032] Figure 24 Depicts images of the staining of human ASC grown in differentiation medium in the absence of test samples at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) at 2 weeks with Oil Red O staining.

[0033] Figure 25 Depicts images of the staining of human ASC grown in control medium in the presence of test samples at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) at 2 weeks with Oil Red O staining.

[0034] Figure 26Depicts images of Oil Red O-stained human ASCs in control medium in the presence of test samples at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) at 2 weeks.

[0035] Figure 27 Depicts images of Oil Red O-stained human ASCs in control medium in the presence of test samples at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) at 2 weeks.

[0036] Figure 28 Depicts images of Oil Red O-stained human ASCs growing in differentiation medium in the presence of test samples at A) 24 hours, B) 1 week, C) 2 weeks, and D) at 2 weeks.

[0037] Figure 29 Depicts images of Oil Red O-stained human ASCs growing in differentiation medium in the presence of test samples at A) 24 hours, B) 1 week, C), D) 2 weeks, and E) and F) at 2 weeks.

[0038] Figure 30 Depicts images of Oil Red O-stained human ASCs growing in differentiation medium in the presence of test samples at A) 24 hours, B) 1 week, C), D) 2 weeks, and E) and F) at 2 weeks.

[0039] Figure 31 Depicts images of Oil Red O-stained human ASCs growing in control medium in the presence of test samples identified as "conventional" at A) 24 hours, B) 48 hours, C) 1 week, D) 2 weeks, and E) at 2 weeks.

[0040] Figure 32 Depicts images of Oil Red O-stained human ASCs growing in differentiation medium in the presence of test samples identified as "conventional" at A) 24 hours, B) 1 week, C), D) 2 weeks, and E) and F) at 2 weeks.

[0041] Figure 33 Depicts a table presenting the results of the human ASC-induced differentiation assay for various samples of the present invention and conventional samples.

[0042] Figure 34 Depicts a graph showing the absorbance values of samples of the test embodiments of the present invention, conventional samples, and control samples.

[0043] Figure 35 Depicts images of test wells with test samples identified as "conventional" at A) before adding the test sample, B) 24 hours after adding the test sample, C) 48 hours, D) 72 hours, and E) in the case of 1% EV stain.

[0044] Figure 36 Depicts images of test wells with test samples identified as "the present invention" at A) before adding the test sample, B) 24 hours after adding the test sample, C) 48 hours, D) 72 hours, and E) in the presence of 1% EV stain.

[0045] Figure 37 Depicts images of control wells with primary human dermal microvascular endothelial cells (HDBECs) identified as "positive control" and complete growth medium at A) before adding the test sample, B) 24 hours after adding the test sample, C) 48 hours, and D) 72 hours.

[0046] Figure 38 Depicts images of control wells with primary HDBECs identified as "negative control" and basal medium at A) before adding the test sample, B) 24 hours after adding the test sample, C) 48 hours, and D) 72 hours.

[0047] Figure 39 Describes a table presenting the results of the endothelial proliferation assay.

[0048] Figure 40 Depicts a graph presenting the results of the endothelial proliferation assay.

[0049] Figure 41 Depicts a table presenting the results of the endothelial proliferation assay averaged across samples of the present invention, samples from conventional tissues, and positive and negative controls.

[0050] Figure 42 Depicts a graph presenting the results of the endothelial proliferation assay averaged across samples of the present invention, samples from conventional tissues, and positive and negative controls.

[0051] Figure 43 Shows ImageJ analysis for calculating adipocyte area using the adiposoft plugin.

[0052] Figure 44 Shows Equation 1.

[0053] Figure 45 Shows an image with five regions stained and analyzed for each sample for comprehensive evaluation.

[0054] Figure 46 Shows H&E staining of the first treated tissue.

[0055] Figure 47 Shows H&E staining of the second treated tissue.

[0056] Figure 48Shows a routinely processed tissue stained with H&E.

[0057] Figure 49 Is a table of the adipocyte composition ranges of the first embodiment and the adipocyte composition ranges of the second embodiment.

[0058] Figure 50 Is a graph showing the compositional analysis of adipose tissue, which shows a lipid content (75%-80%), a water content (15%-20%), and a protein content (3%-5%).

[0059] Figure 51 Is a table of the compositional analysis of the first allograft embodiment.

[0060] Figure 52 Is a graph showing the quantification of the first tissue components of representative donor 1.

[0061] Figure 53 Is a table of the compositional analysis of the second allograft embodiment.

[0062] Figure 54 Is a table of the results of the comparison of adipose tissue composition excluding the data of 1 representative donor 3.

[0063] DETAILED DESCRIPTION It should be understood that the drawings and description of the present invention have been simplified to illustrate elements relevant to a clearer understanding of the present invention, while eliminating many other elements that exist in the systems and methods in the art for the sake of clarity. Those of ordinary skill in the art can recognize that other elements and / or steps are desirable and / or required when implementing the present invention. However, since such elements and steps are well known in the art and since they do not facilitate a better understanding of the present invention, no discussion of such elements and steps is provided herein. The present disclosure relates to all such changes and modifications of such elements and methods known to those skilled in the art.

[0064] 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.

[0065] As used herein, each of the following terms has a meaning related to the meaning in this section.

[0066] The article "a" is used herein to refer to one or more than one (i.e., at least one) grammatical object of the article. For example, "an element" means one element or more than one element.

[0067] As used herein, when referring to measurable values such as amounts, lengths of time, etc., "about" is intended to encompass variations of ±20%, ±10%, ±5%, ±1%, and ±0.1% relative to the specified value, as such variations are appropriate.

[0068] Ranges: Throughout this disclosure, various aspects of the invention may be presented in 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 immutable limitation on the scope of the invention. Where appropriate, the description of a range should be considered to have specifically disclosed all the possible sub-ranges as well as the individual numerical values within that range. For example, a description of a range such as 1 to 6 should be considered to have specifically disclosed sub-ranges such as 1 to 3, 1 to 4, 1 to 5, 2 to 4, 2 to 6, 3 to 6, etc., as well as the individual numerical values within that range, such as 1, 2, 2.7, 3, 4, 5, 5.3, and 6. This applies regardless of the width of the range.

[0069] Reference is now made in detail to the drawings, in which like reference numerals designate identical parts or elements throughout the several views, and in various embodiments, minimally manipulated tissue filler allografts are provided herein.

[0070] A strategy for preserving the native extracellular matrix (ECM) in minimally manipulated adipose tissue is decellularization. Decellularization is a process aimed at improving biocompatibility or minimizing the immune response by removing tissue-resident cells while keeping the ECM intact (Fu, R.H. et al. 2014. Cell Transplant 23, 621-630). The method steps can vary, but generally involve successive washes in chemical and biological reagents, which can range from a few days to more than a week (Flynn, L.E. 2010. Biomaterials 31, 4715-4724., Aamodt, J.M. et 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 lead to the destruction of ECM proteins (e.g., glycosaminoglycans) (Reing J.E. et al. 2010. Biomaterials 31, 8626-8633.), structure (Sesli, M. et al. 2018. Turk JBiol 42, 537-547.), or may damage the sample (Hayes, D.J. et al. 2022. Biomaterials and Biosystems 7, 100053). Generally, decellularization of adipose tissue requires precise control of a series of processing steps (depending on the tissue and protocol, approximately 2-5 days) (Wang, L. et al. 2013. Acta Biomaterialia 9, 8921-8931., Mohiuddin, O.A. et al. 2019. 57-70 Springer International Publishing) to ensure minimal damage to the residual ECM while ensuring the removal of all cellular components (Choi, J.S. et 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 native adipose extracellular matrix (ECM) and tissue integrity for soft tissue fillers. Minimally manipulating adipose tissue or treating it without altering the original relevant characteristics related to the tissue's utility for reconstruction, repair, or replacement (21 CFR 1271.3(f)(1)) provides a safe (Services, U.S.D.o.H.a.H. 2020 ed U.S. Food and Drug Administration (Food and Drug Administration)) and standardized method, thus providing clinicians with an off-the-shelf, minimally manipulated adipose tissue product (MMAP).

[0071] The present invention relates to a minimally manipulated adipose tissue allograft that can be injected or surgically implanted into a patient in need of soft tissue augmentation. The adipose tissue allograft herein retains the native ECM and adipocytes of the donor tissue. In addition, the adipose tissue allograft also retains 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 allograft also contains highly reduced amounts of cellular and allogeneic components such as DNA and nucleic acids that may induce an immune response which, if severe enough, may lead to immune rejection of the allograft. The allograft further retains components such as ECM, lipids, and water, the amounts of which can be varied by embodiments of the manufacturing methods described herein to produce various embodiments of the present invention. The adipose tissue filler allograft does exhibit a higher level of adipocytes as well as ECM structures and structural proteins and a highly reduced amount of DNA and nucleic acids in histological staining (see Example 2).

[0072] The present disclosure further provides an embodiment of a method for preparing a minimally manipulated adipose tissue allograft that retains at least a portion of the intracellular components. The method herein includes several steps discussed in detail below to lyse the tissue having cellular and allogeneic components while avoiding denaturation and elution of the structural proteins. The cellular and allogeneic components are removed by a freeze-thaw process, a mechanical grinding process, and rinsing with mild detergents, sterile saline, and sterile water. These processes help to weaken and / or disrupt the cell membranes of the cells in the tissue and thus decellularize the tissue. In addition, the rinsing also separates the intracellular components from the rest of the tissue. These processes are also gentle enough to help retain the necessary structural components of the tissue such as the extracellular matrix, adipocytes, and structural proteins. The method described herein is a minimally manipulated method within the 361HCT / P guidelines.

[0073] Now referring Figure 1A , a method for processing adipose tissue 100 generally includes the following steps: freeze-thaw cycle (step 101), human donor adipose tissue debridement and slicing (step 102), size reduction (step 103), liquid handling (step 104), pre-packaging storage (step 105), quality control inspection (step 106), and packaging (step 107). These processes are combined in a unique way to affect the amounts of tissue components including adipocytes, free lipids, extracellular matrix, water, and oxygen and to remove voids and homogenize the allograft.

[0074] The method begins with harvesting human cadaveric adipose tissue from a donor using any suitable method. Suitable methods can include, but are not limited to, aspiration, dissection, scraping, and other surgical methods known in the art. In other embodiments, donor tissue can be harvested and stored at cryogenic temperatures.

[0075] Now referring Figure 1B , in one embodiment, step 101 includes freezing the harvested donor tissue and allowing the harvested donor tissue to thaw for no more than a 120-hour period. This freeze-thaw cycle is the first step in decellularizing the tissue because cell membranes are disrupted during freeze-thaw.

[0076] Step 102 includes debriding the tissue to remove any foreign or abraded tissue. The tissue is debrided onto a sterile area, and the amount of time the tissue is open is consistently recorded. The adipose tissue is measured, weighed, and inspected for abraded and foreign tissue such as visible fascia, arteries, veins, or muscle. The tissue is debrided using methods known in the art to remove all foreign tissue and abraded adipose tissue. Suitable methods can include, but are not limited to, using a scalpel, forceps, scraper, and other methods known in the art. Any foreign and abraded tissue can be discarded.

[0077] Step 103 includes mechanically separating the tissue into small pieces. In some embodiments, the separation can be performed by cutting, grinding, crushing, or any suitable method. In some embodiments, the donor tissue can be sliced. These slices can be cubes approximately 4 cm × 4 cm or smaller. Further separation can be performed 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 obtained from the tissue separation process can be discarded.

[0078] Mechanical grinding aids in the decellularization process of cells, and the shearing force of the grinder can tear the cell membranes of some cells. Mechanical grinding of adipose tissue also subjects the product to sufficient force to separate the tissue into fine particles. This provides the product with the necessary injectability to allow it to pass through containers, syringes, or cannulas with a diameter of 1.5 mm - 4.5 mm or larger. In one embodiment, the treated tissue can be injected through an opening with a diameter of 1.5 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 2 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 2.5 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 3 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 3.5 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 4 mm. In one embodiment, the treated tissue can be injected through an opening with a diameter of 4.5 mm. Additionally, the mechanical grinding step also subjects the tissue to a force gentle enough to allow at least 10% of the adipocytes in the tissue to remain intact. In one embodiment, the intact adipocytes maintain the natural adipocyte structure. In one embodiment, the intact adipocytes maintain the natural adipocyte function to obtain cushioning and support. In one embodiment, the intact adipocytes maintain the tissue structure and extracellular matrix (ECM) of the adipocytes. In one embodiment, the intact adipocytes maintain the tissue structure, ECM, triglycerides, and connective tissue of the adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 10% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 15% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 20% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 25% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 30% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 40% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 50% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 55% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 60% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 65% intact adipocytes. In one embodiment, the tissue is treated with adipocytes composed of at least 70% intact adipocytes. Adipocytes help maintain the structural integrity and volume of allografts, making them better soft tissue implants. Other methods known in the art for size reduction, such as cryomilling, do not allow the retention of adipocytes.

[0079] Step 104 includes liquid handling of the product. This step is broken down into several sub-steps, which include successive rinsing with a non-ionic surfactant, sterile crystalloid fluid, and sterile water. After each rinse, the tissue is decanted to separate it from unwanted liquid or cellular components.

[0080] Step 104a includes separating the tissue from free lipids. In some embodiments, the separation can be performed by dispensing the ground adipose tissue in aliquots into centrifuge tubes. The tissue can then be centrifuged at a speed of 900 - 3655 RCF for a certain period of time. Decanting the adipose tissue through a sieve or other methods known in the art can separate the free lipids from the adipose tissue. Any decanted liquid, oil, or cell clumps are discarded. The tissue can then be rinsed with warm sterile water. The tissue is dispensed back into centrifuge tubes in aliquots, and warm sterile water is added at a ratio of approximately 1:1 warm sterile water to tissue. The centrifuge tubes can be gently agitated to disperse the adipose tissue in the water. The tubes are then centrifuged at 900 - 3655 RCF at ambient temperature. The adipose tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell clumps are discarded.

[0081] Step 104b includes rinsing the adipose tissue with a non-ionic surfactant. This step further aids in cell lysis, and the surfactant helps to weaken the cell membrane to decellularize the tissue. In some embodiments, the surfactant product used can be a 0.01% Triton X-100 solution. In some embodiments, the rinse is performed by dispensing the adipose tissue into centrifuge tubes and adding it to the 0.01% Triton X-100 solution. The solution can be added at a ratio of 1:1 solution to adipose tissue. The centrifuge tubes can be gently agitated for a certain period of time, such as 100, 200, 300, 400, 500, 600, 800, or 1,000 seconds or longer. The centrifuge tubes can then be centrifuged at a speed of 900 - 3655 RCF. In some embodiments, the rinse can be performed by dispensing the adipose tissue in a sealable container. Then, the 0.01% Triton X-100 solution is added at a ratio of approximately 1:1 solution to adipose tissue. The sealable container is then agitated several times during the rinse time (e.g., 1, 2, 3, 4, 5, 6, 8, 10 minutes or longer). The adipose tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell clumps are discarded. The surfactant rinse can be repeated as needed.

[0082] Dry the adipose tissue. In some embodiments, the tissue can be dried by centrifugation. The 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 clumps are discarded.

[0083] Step 104c includes rinsing the tissue in a sterile crystalloid fluid. The adipose tissue is rinsed with a sterile crystalloid fluid. In some embodiments, the sterile crystalloid fluid can be a 0.9% sterile saline solution. In some embodiments, the rinsing can be performed by dispensing the adipose tissue into a centrifuge tube. A 0.9% sterile saline solution is added to the tube at a ratio of approximately 1:1 solution to tissue. The centrifuge tube is gently agitated. Then, the centrifuge tube is centrifuged at a speed of 900 - 3655 RCF at ambient temperature. In some embodiments, the rinsing can be performed by dispensing the adipose tissue in a sealable container. A 0.9% sterile saline solution is added to the adipose tissue at a ratio of approximately 1:1 solution to tissue. The sealable container is gently agitated several times during the rinsing period. After rinsing, the tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell clumps are discarded. The sterile saline rinse can be repeated multiple times.

[0084] Step 104d includes rinsing the tissue in sterile water. The adipose tissue is rinsed in sterile water. In some embodiments, the sterile water can be warm. In some embodiments, the rinsing is performed by dispensing the adipose tissue into a centrifuge tube. Sterile water is added to the centrifuge tube at a ratio of approximately 1:1 solution to tissue. The centrifuge tube is gently agitated for approximately 2, 5, 10, 15, or 20 seconds. Then, the tube is centrifuged at a speed of 900 - 3655 RCF at ambient temperature. In some embodiments, the rinsing can be performed by dispensing the adipose into a sealable container. Sterile water is added to the sealable container at a ratio of approximately 1:1 solution to tissue. The sealable container is gently agitated several times during the rinsing period. After rinsing, the tissue can then be decanted through a sieve, and any decanted liquid, oil, or cell clumps are discarded. The sterile water rinse can be repeated multiple times.

[0085] Step 104e includes removing excess water from the adipose tissue. In some embodiments, excess water is removed by loading a drying sieve into a centrifuge tube. A wipe is cut around the circumference of the drying sieve and placed on top of the sieve. The adipose tissue can be equally dispensed into the tube, and the tube can be centrifuged at ambient temperature. The tube can be centrifuged at a speed of 900 - 3655 RCF. Excess water can be drained through the bottom opening of the centrifuge tube. Any unwanted liquid that is drained can be discarded. Step 104e can be repeated for multiple centrifugation cycles (e.g., 2, 3, 4, 5, or more).

[0086] Step 105 includes pre-packaging the resulting adipose tissue allograft. The pre-packaging includes combining the tissue filler product and gently mixing until a uniform consistency is obtained. The adipose tissue product can be prepared for packaging by equally distributing the product into centrifuge tubes. The tubes can 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).

[0087] Now referring Figure 1C , a method 150 for processing adipose tissue includes the following steps: freezing and thawing human adipose tissue 152, separating the human adipose tissue into pieces 154 by mechanical grinding, continuously rinsing the human adipose tissue 156 with a non-ionic surfactant, sterile crystalloid solution, and sterile water, and dispensing the human adipose tissue into a package 158 using a loading tool. In one embodiment, the method includes the step of debriding the human adipose tissue to remove foreign or abraded tissue. In one embodiment, the method includes the step of rinsing the human adipose tissue with a non-ionic surfactant, sterile crystalloid solution, and sterile water by dispensing the human adipose tissue into centrifuge tubes and centrifuging the adipose tissue. In one embodiment, the method includes the step of rinsing the human adipose tissue with a non-ionic surfactant, sterile crystalloid solution, and sterile water by dispensing the human adipose tissue into a sealable container and gently agitating the container. In one embodiment, the method includes the step of rinsing the tissue multiple times in the non-ionic surfactant. In one embodiment, the method includes the step of rinsing the tissue multiple times in the sterile crystalloid solution. In one embodiment, the method includes the step of rinsing the tissue multiple times in the sterile water. In one embodiment, the method includes the step of drying the human adipose tissue in a centrifuge tube including a drying sieve. In one embodiment, the method includes the step of decanting the tissue and removing any decanted liquid and cellular components after each rinse. In one embodiment, the method includes the step of evenly dispensing the human adipose tissue in a container using a loading tool. In one embodiment, the method includes the step of packaging the container in a sterile pouch.

[0088] In some embodiments, according to one embodiment, the tissue product can be dispensed via a loading tool, such as Figures 2A - 2FAs shown. The loading tool can be used in combination with adipose tissue manufacturing methods to enable centrifugation, atmosphere control, water content, composition control, physical processing, and dispensing of adipose allografts and similar tissues and media. The tool can be sterilized via an autoclave and does not require external power. The loading tool can hold a formable and flowable medium during centrifugation, enclose the medium during atmosphere transfer, remove unnecessary voids and air traps from the medium, and dispense the medium to fit various different containers. In one embodiment, the loading tool 200 includes a centrifuge-compatible wide-mouth bottle 250 or container having 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 enable effective decantation during liquid handling, then one-way gas exchange during atmosphere control, and then void-free dispensing during dispensing. A first adapter 270, such as the adapter shown in Figure 2D and Figure 2E (alternatively referred to as a container adapter) can mate with the distal opening 254 on the wide-mouth bottle 250 via a threaded surface and provide a sealable inlet 276 having a proximal opening 272 and a distal opening 274 to facilitate dispensing the medium from the wide-mouth bottle 250 into a container. A connection structure 278, such as a threaded opening for a shoulder screw, can be included for assembling separable components of the first adapter 270. A second adapter 280, such as the adapter shown in Figure 2F (alternatively referred to as a nitrogen adapter) can mate with the distal opening 254 on the wide-mouth bottle 250 via a threaded surface and provide a sealable inlet 286 having a proximal opening 282 and a distal opening 284 for fluid or gas communication with the wide-mouth bottle 250, which can be used, for example, to remove gas or liquid from the medium in the wide-mouth bottle 250. This sealable inlet 286 can also be connected to a valved conduit to create a sealed fluid connection.

[0089] The piston 202, bracket 204, locking lever 206, wide-mouth bottle 250, and nitrogen adapter 280 together facilitate creating an airtight container around the medium. Ports 275 on the nitrogen adapter and piston enable one-way gas exchange to remove undesired gases, such as oxygen, from the medium. The port can have, for example, a controllable valve for controlling fluid and gas communication. The lever 210 and container adapter 270 work together with the other components to dispense the medium in a void-free and low-oxygen manner. The lever 210 can be attached to the bracket 204 at a hinged connection 212. A locking lever 206 can be added to prevent movement of the lever 210.

[0090] The loading tool can be completely disassembled and sterilized. The material is compatible with various sterilization methods such as steam sterilization, ethylene oxide, etc. The loading tool enables atmosphere control for product packaging through sealed atmosphere exchange. Tissue is contained in a sealed space with ports to facilitate removal of the naturally occurring atmosphere by vacuum and replacement in a flow-through manner, backfilling with an inert gas as needed. The atmosphere exchange enables preservation using various inert gases. The loading tool gently extrudes the tissue directly into a container with minimal atmosphere exposure in a controlled manner to produce a filled container with minimal voids. The loading tool can be adapted to various tip configurations for use with desired containers. The loading tool jars are also compatible with centrifugation in terms of shape, strength, and quality. The jars compatible with centrifugation as indicated previously have sealable openings at the top and bottom for selectively removing high-density or low-density tissue, solutions, and / or debris. This, along with the proximal and distal openings of the jar, facilitates product flushing and allows the same container to be used for product filling / packaging. The loading tool enables control of the water content through suspension centrifugation. The jar and drying sieve assembly suspend the tissue with or without a permeable membrane that suspends the tissue over an empty space during centrifugation to remove excess water and / or other solutions present in the tissue. Since the assembly can be autoclaved, the tool and its components are reusable.

[0091] Using the loading tool for the packaging process allows allografts to be effectively dispensed in the primary packaging, which can improve shelf stability, the cosmetic properties of the allograft, and enable controlled dispensing at the point of use. Prepare the loading tool for use by loading a centrifuge jar containing adipose tissue onto the adapter. Slightly depress the piston of the loading tool to engage the O-ring and insert and latch the locking rod. Attach the nitrogen adapter to the base of the jar and attach the vacuum to the piston.

[0092] Then, use the loading tool to purge oxygen from the tissue. Adjust the vacuum pump (e.g., approximately -5Hg, -10Hg, -15Hg, -20Hg or higher) and open the vacuum line valve. Open the nitrogen line valve enough to reduce the vacuum (e.g., to approximately -5Hg, -10Hg, -15Hg for 5, 10, or 15 seconds). Close the nitrogen line valve and open it a second time to reduce the vacuum a second time. Close the nitrogen line valve and then open it a third time and keep it open. Close the vacuum pump while keeping the vacuum line open and allow nitrogen to bring the pressure to 0Hg. Close the nitrogen line valve.

[0093] Then prepare the loading tool for packaging. Remove the locking rod after pressure normalization and lower the piston to contact the adipose tissue. Close the vacuum line valve and remove the nitrogen adapter. Attach a container adapter to a container.

[0094] The tissue can now be packaged. The loading tool 200 is used in combination with the container adapter 270 to package the tissue into the final primary container. As indicated herein, although the syringe 290 is mentioned according to several embodiments, more generally, several types of containers may be suitable for use, such as syringes, cannulas, or other types of containers.

[0095] Step 106 includes performing several quality control checks to ensure that the adipose tissue allograft meets the standards.

[0096] In some embodiments, the color of the tissue is visually inspected. According to Figure 3A the chart depicted in Figure 3A if the tissue color is off-white and / or has various shades / tints of yellow, the product passes the inspection. According to

[0097] If the tissue fails the color inspection test, rework must be performed. Rework includes equally distributing the adipose tissue into centrifuge tubes and submerging the adipose tissue in sterile 0.9% saline to achieve a roughly 1:1 saline-to-adipose tissue ratio. The centrifuge tubes are gently oscillated for approximately 2, 5, 10, 15, or 20 seconds to disperse the adipose tissue into the saline. Then, the centrifuge tubes are centrifuged at 900 - 3655 RCF for 5 minutes. Then, the tissue is decanted through a sieve, and any cell clumps, liquid, or oil can be discarded. This process is repeated, and the adipose tissue is gently mixed inside the centrifuge tube and placed back in the centrifuge. The tissue is centrifuged at 900 - 3655 RCF for 3 minutes. The tissue is decanted through a sieve, and any unwanted liquid or cellular components can be discarded. The visual color inspection is repeated. Rework can be repeated as many times as needed until the product is acceptable. Record the final visual color inspection.

[0098] In some embodiments, an injectability test is performed. A filled syringe is loaded with the product (roughly 14 cc). The syringe is connected to a single outlet cannula. The product is extruded through the cannula. If the entire contents of the syringe can be extruded through the cannula using the force of a finger, the product passes the injectability test. If the force of a finger on the plunger cannot extrude the tissue and / or the cannula becomes blocked, the product fails the injectability test. If the product fails the injectability test, rework is performed, and the test is repeated.

[0099] Rework includes squeezing the tissue through a 3 - 4 mm sieve into a bowl using a spatula. Only collect the tissue that passes through the sieve. Gently combine and mix the tissue together until a uniform consistency is obtained. Then, equally distribute the adipose tissue into centrifuge tubes and immerse it in sterile water to achieve a roughly 1:1 sterile water to adipose tissue ratio. Gently oscillate the centrifuge tubes for approximately 2, 5, 10, 15, 20 seconds or longer to disperse the adipose tissue into the sterile water. Then, centrifuge the tubes at 900 - 3655 RCF (e.g., for 200, 300, 400, 500, 600, 800, 1,000 seconds or longer). Decant the adipose tissue through the sieve and discard any cell clumps, liquid, or oil. Gently mix the adipose tissue in the centrifuge tube, place it back in the centrifuge, and centrifuge at 900 - 3655 RCF (e.g., for 60, 90, 120, 150, 180 seconds or longer). Decant the tissue through the sieve again and discard any cell clumps, liquid, or oil. Repeat the injectability test and rework can be repeated as needed multiple times until the product is acceptable. Record the final injectability test.

[0100] In some embodiments, visually inspect the product for excess free lipid according to the Figure 3B chart depicted in. According to the chart, if the free lipid does not form an oily regular substance at or around the tissue, the tissue is acceptable. According to the chart, if the free lipid forms an oily contour at or around the tissue, the tissue is unacceptable and thus fails the test. If the product fails, rework is performed and the test is repeated.

[0101] Rework includes equally distributing the adipose tissue into centrifuge tubes and immersing the tissue in sterile water to achieve a roughly 1:1 sterile water to adipose tissue ratio. Gently oscillate the tubes (e.g., for 5, 10, 15, 20, 25 seconds or longer) to disperse the adipose tissue into the water. Then, centrifuge the tubes at 900 - 3655 RCF for 5 minutes. Decant the tissue through the sieve and discard any cell clumps, liquid, or oil. Gently mix the adipose tissue inside the centrifuge tube, place it back in the centrifuge, and centrifuge at 900 - 3655 RCF. Decant the tissue again and discard any cell clumps, liquid, or oil. Repeat the free lipid test and rework can be repeated as needed multiple times until the product is acceptable. Record the final inspection result.

[0102] In some embodiments, absorption measurement tests are conducted. Place the absorbent paper on a clean, flat, and non-porous surface. Place a pea-sized product sample on the absorbent paper. Allow the tissue to settle for 15 minutes. Using a ruler, measure the travel distance from the point where the tissue contacts the paper to the farthest edge of the lipid trace. If the lip trace traveling on the absorbent paper is less than or equal to 5 mm, the product is acceptable. If the lipid trace traveling on the absorbent paper is greater than 5 mm, the product is unacceptable. If the product fails the absorption measurement test, it is reprocessed and the test is repeated. Reprocessing includes equally distributing the adipose tissue into centrifuge tubes and submerging the tissue in sterile water to achieve a roughly 1:1 sterile water to adipose tissue ratio. Gently agitate the tube for approximately 2, 5, 10, 15, or 20 seconds to disperse the adipose tissue into the water. Then, centrifuge the tube at 900 - 3655 RCF for 5 minutes. Decant the tissue through a sieve and discard any cell clumps, liquid, or oil. Gently mix the adipose tissue inside the centrifuge tube, place it back in the centrifuge, and centrifuge at 900 - 3655 RCF for 3 minutes. Decant the tissue again and discard any cell clumps, liquid, or oil. Repeat the visual inspection and absorption measurement tests, and repeat as needed until the product is acceptable. Record the final test results.

[0103] Step 107 includes packaging the product. Packaging includes loading the product into a container using a loading tool. The loading tool is fully set, and the tissue product is loaded into the container using a piston on the loading tool to backfill each container. Seal the container. A torque wrench can be used to tighten each lid. Repeat the loading step until all the product is packaged into containers. All containers can be inspected for product specifications, cosmetic quality, air traps, and fill completeness.

[0104] In some embodiments, each container is packaged inside a pouch. The pouch can be a foil or foil herringbone pouch. Each container can be packaged such that the lid faces the herringbone and away from the opening of the pouch. Each pouch can be vacuum-sealed. Each seal can be inspected to ensure it is uniform, free of wrinkles, and complete across the width of the pouch. A unique allograft ID tag number can be applied to each pouch.

[0105] In some embodiments, each pouch can be packaged inside a pouch with a larger width and length. The inner pouch can be vacuum-sealed. A single sealed pouch with a syringe inside can be placed inside the larger pouch. In some embodiments, the larger pouch can be a polyester or Tyvek herringbone pouch. The smaller pouch can be placed inside the larger pouch such that the herringbones are aligned. Each larger pouch can be heat-sealed. Each seal can be inspected to ensure it is uniform, free of wrinkles, and complete across the width of the pouch.

[0106] Experimental Example

[0107] The present invention will now be described with reference to the following embodiments. These embodiments are provided for illustrative purposes only, and the present invention should in no way be construed as limited to these embodiments, but rather should be construed to cover any and all modifications that become apparent as a result of the teachings provided herein.

[0108] Without further description, it is believed that one of ordinary skill in the art can make and utilize the present invention and practice the claimed methods using the foregoing description and the following illustrative embodiments. Accordingly, the following working examples specifically point out the preferred embodiments of the present invention and will not be construed as limiting the remainder of the disclosure in any way.

[0109] Experimental Example 1

[0110] Soft tissue defects can result in loss of normal tissue function, emotional trauma, and pain. Cosmetic and reconstructive surgery is needed; however, current surgeries have problems such as varying degrees of success in volume retention, angiogenesis induction, and complete wound healing. Minimally manipulated adipose products (MMAPs) provide a non-surgical method in cosmetic or reconstructive surgery that can overcome these problems. In the study conducted herein, the safety, biocompatibility, and volume retention of the present invention were evaluated: a minimally manipulated adipose filler allograft, hereinafter referred to as "allograft". The results demonstrated that, as observed in vitro by inhibited T cell proliferation and viability and in vivo, where the allograft was implanted in athymic mice for 3 months and no significant immune response or safety risk was observed, the allograft is biocompatible with the human system. Additionally, the in vivo allograft implants demonstrated complete wound closure, and all mice remained in good health throughout the duration of the study. As observed by histopathology, the allograft implants retained the matrix and increased angiogenesis and local adipogenesis were observed. Finally, volume retention of the allograft was observed in vivo in the case of high-volume implants (200 μl) maintaining a visible implant volume for up to 3 months. Our findings suggest that the allograft is a versatile MMAP that synergistically integrates with the surrounding tissue, thus creating a potential off-the-shelf product for use in cosmetic or reconstructive surgery.

[0111] This study presents a novel human MMAP adipose tissue allograft as an excellent volume filler candidate for aesthetic or trauma-based soft tissue augmentation. The natural yellowish response and ease of implant handling of the allograft, including its bulk volume and fluidity, permit various applications and delivery methods, such as surgical implantation or injection. No adverse immune responses were reported in either in vitro or in vivo studies. None of the mice implanted with low or high implant volumes suffered any health problems and remained in good health throughout the study duration, i.e., 3 months. High-volume implants maintained better volume retention and were firmer when removed, but showed signs of tissue encapsulation and potential initial signs of fat necrosis, while low-volume implants were absorbed into native tissue much more rapidly. Finally, the implants were remodelled, and the histopathologist observed evidence of enhanced collagen formation, increased angiogenesis, and local adipose tissue remodelling changes. Our findings suggest that the allograft is a versatile human MMAP that interacts synergistically with the surrounding tissue, thus creating a potential off-the-shelf product for surgical use.

[0112] Experimental Example 2

[0113] This study evaluated the compositional characterization of adipose allografts. Adipose tissue is a connective tissue composed of clusters of cells (adipocytes) surrounded by a thick extracellular matrix (ECM) (Mariman, E.C., et al., 2010. Cellular and molecular life sciences, 67(8), 1277-129.) and other cells including 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). In addition to other tissues, adipose tissue provides cushioning and support for the skin, stores energy in the form of lipids, and insulates the body, among other functions (FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use, July 2020), such as the expression and secretion of factors and proteins with important endocrine functions, such as adiponectin, TNFα, IL-6, and other cytokines (Kershaw, E.E. et al., 2004. The Journal of Clinical Endocrinology & Metabolism, 89(6), 2548-2556). The adipose allograft invention preserves the intrinsic natural characteristics of adipose tissue, namely the adipocyte structure and extracellular matrix.

[0114] The materials and methods of this experiment are now described. Adipose allograft samples were analyzed by multiple methods to characterize their composition. DNA quantification of adipose allograft samples was performed using a DNA assay kit. Quantifying the DNA content of the samples supported the efficacy of the processing method used to generate the adipose allografts in reducing the DNA content. The DNA assay consisted of extracting and separating DNA from the samples using a series of buffers and then purifying the DNA using a spin column. The separated DNA was then quantified using a spectrophotometer.

[0115] Perform a compositional analysis on adipose allograft samples (i.e., previous product iterations of adipose allografts) and untreated adipose samples (also referred to as native). This compositional analysis determines the main components of the samples, specifically water, lipids, and solids (including ECM), and this composition can be quantified based on weight / weight (in grams) percentages. Compare the results.

[0116] Evaluate adipose allograft samples and native samples by proteomics from a third-party vendor to identify the structural protein content (including ECM) and protein retention in the allografts compared to native tissue.. Proteomics studies are used to identify proteins in the samples and confirm their presence by mass spectrometry.

[0117] Process adipose allograft samples and native samples by a third-party vendor to obtain histology, embed in paraffin, cut, stain, and image. Generate images of the tissue samples presenting the microstructure of the tissue samples to compare the adipose allograft samples with the untreated samples and to support whether the structural components are retained in the allograft compared to native tissue. Stain the samples with Masson’s Trichrome and hematoxylin and eosin (H&E). Masson’s Trichrome visualizes the connective tissue (including ECM) in the samples. H&E visualizes the tissue structure and DNA in the nuclei of adipocytes.

[0118] Now describe the results of the experiment. During adipose allograft development, it is imperative that the process includes a series of steps to improve the reduction of cellular and allogeneic components by a decellularization method while leaving behind structural components such as the extracellular matrix (ECM). Decellularization is the process of removing any heterologous or xenogeneic cellular antigens from tissue that would induce an immune response (Bruyneel AAN, et al., Artif Organs. 2017). Inadequate reduction of cellular and allogeneic components may trigger the innate immune system, which is part of the nonspecific immune system and attacks any substance perceived as foreign. This can lead to inflammation and, if severe enough, to graft rejection (Kasravi M, et al., Biomater Res. 2023). Crapo et al. stated that decellularization techniques cannot remove 100% of the cellular material (Crapo PM, et al., Biomaterials. 2011). Thus, adipose allograft development focuses on the reduction of cellular and allogeneic components in the final product compared to native tissue.

[0119] Figure 4Shows the average DNA concentration in three donors, namely the original sample and the adipose allograft sample. The results show that the DNA concentration in the adipose allograft is lower compared to the original. Overall, as Figure 5 supported, the adipose allograft sample contains less DNA than its original counterpart. Figure 4 Also depicts sample variability. One challenge in obtaining a primary quantity of DNA per sample is donor variability. As Figure 6 shown, each donor has unique characteristics that can lead to inconsistent data.

[0120] During the development of adipose allografts, it is helpful to understand the compositional characterization of adipose allografts to accurately describe the product and compare it with native tissue or other grafts. For comparison purposes, compositional analysis is performed to determine the major components of adipose tissue-based samples, specifically the water, lipid, and solid (including ECM) components. Figure 6 、 Figure 7A and Figure 7B Present the average amount of sample components produced in each corresponding form of adipose tissue-based samples (i.e., the first sample embodiment and the second sample embodiment and the original). Variability between samples of the same type was observed, which may be attributed to donor and sample variability as the composition of each donor is unique and sample homogenization may not always be achieved. The results seem to indicate a relationship between the solid component and the water content. This may be attributed to the fact that the proteins in the ECM are hydrophilic, thus facilitating attachment (Yang, L. et al., RSC Advances, 2017). On average, the percentage of solid components (including ECM) in the sample of embodiment 1 increased compared to embodiment 2. In future studies, it is recommended to increase the number of samples analyzed to improve the confidence of the results.

[0121] During the development of adipose allograft processing, non-ionic surfactants are used as mild detergents to reduce cellular and allogeneic components while avoiding the denaturation and elution of structural proteins. Proteomic studies are used to confirm the presence of structural proteins in adipose allografts compared to native adipose tissue. As Figure 8A and Figure 8BAs shown above, compared to native tissue from the same donor, the following proteins are retained in four different allograft samples. Adipose allografts retain major adipose collagens (COL4, COL6), as well as major fibrillar collagens (COL1, COL3) and minor fibrillar collagen (COL5). These proteins are responsible for ECM formation 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, the key adipose ECM proteins fibronectin and laminin-α, as well as elastin microfibril interfacer-located protein 1, elastin microfibril interfacer-located protein 2, adiponectin (associated with adipocyte proliferation), and fibrillin-1 (contributing to adipocyte development) (Ricard-Blum S. Cold Spring Harb Perspect Biol. 2011). Additionally, adipose allografts retain various proteoglycans. The key component in proteoglycans is glycosaminoglycan (GAG), which is the major ECM protein in fat; it is also responsible for basement membrane development, water retention, and growth factor transport across the ECM (Pessentheiner AR, et al. 2020).

[0122] During the development of adipose allografts, the grafts are prepared by minimally invasive processing methods that maintain the structure of the adipose tissue in the graft as it is in native tissue. Histological images of tissue samples presenting the microstructure of the tissue samples are collected to confirm the retention of adipose tissue structure by comparing images of native adipose tissue with images of adipose allografts. Tissue samples are prepared with two different stains, namely Masson's trichrome and H&E, to observe the microscopic structural composition of the tissue. Masson's trichrome is a beneficial 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 trichrome stain is a three-color stain that identifies connective tissue in blue / green, nuclei in red / purple, and cytoplasm in pink. H&E is a beneficial stain to use because it targets cell structures and nuclei (Zhou X, et al. BioProtoc. 2017). H&E is a dual stain that identifies nuclei in purple / blue stained with hematoxylin and cytoplasm in red / pink stained with eosin.

[0123] Figures 9 - 14The histological images of the tissue samples depicted above show that the adipose allograft retains the structural components, such as the adipocyte structure (represented by the honeycomb structure) and the connective tissue network including adipose tissue (including ECM), compared to the native tissue. This observation was consistent across the samples. Figures 9 - 14 The histological images depicted in also show an increased amount of staining of the connective tissue (including ECM) compared to the individual adipocyte structures in the adipose allograft samples compared to the native tissue. This observation was consistent across the samples.

[0124] Experimental Example 3

[0125] This study evaluated the aging performed on adipose tissue-based research allografts in real time. The real-time aging study provided data to determine the shelf life of the product and the effect of aging on the material. During the study, the product was on the shelf and exposed to environmental changes such as temperature and humidity fluctuations, which simulated the real-world environment the product might experience during its life cycle. In this study, the conditions for simulating real-time aging were used to collect data on the aging of adipose tissue-based grafts solely for research purposes.

[0126] Materials and methods are now described. Donor adipose tissue was designated for research with appropriate donor consent and permission. The donor was tested and found negative for the following infectious diseases: antibodies against human immunodeficiency virus (types 1 and 2), nucleic acid testing (NAT) for HIV-1 in cells, hepatitis B surface antigen, hepatitis B core antigen antibody, antibodies against hepatitis C virus, and syphilis. Pretreatment cultures of the recovered tissue were obtained and found negative for the following pathogenic, highly virulent microorganisms: Clostridium spp., fungi (yeast, mold); and Streptococcus pyogenes. The adipose tissue was recovered in a manner to exclude contamination and / or cross-contamination.

[0127] The adipose tissue transportation, storage, handling, and packaging were performed and recorded according to the methods disclosed herein. The adipose tissue was handled in a manner to minimize contamination and / or cross-contamination. The grafts were gamma irradiated at a minimum dose of 15 kGy at a third-party contractor according to the graft protocol. Records of the radiation dosimetry were obtained. After irradiation, the grafts were transported to a location with stable environmental conditions designated for research at ambient temperature. The grafts were stored at ambient temperature before evaluation. The evaluation time points were approximately several months apart.

[0128] Perform a bioburden test on at least 1 graft unit before irradiation. After irradiation, perform an additional bioburden test on at least 1 graft to confirm no microbial contamination. Evaluate at least 1 graft unit at selected time points. Assign the tissue in each graft in a manner that maximizes the number of evaluations. All evaluations are optional as they depend on graft availability. Perform quality control checks internally. Prepare grafts for histological imaging to evaluate the structural characteristics of the product and potential changes over time at the microscopic scale. Conduct mechanical tests. Perform an in vitro cell viability assay internally to evaluate whether the test sample may affect cell viability, i.e., the NIH / 3T3 or NCTC clone 929 (L cells, L-929, derivative of strain L) mouse fibroblast cell line.

[0129] Now describe the experimental results. Perform a bioburden test on adipose allograft research samples from 4 different donor batches before irradiation. Also perform a bioburden test on adipose allograft research samples from 2 different donor batches after irradiation.

[0130] Perform quality control checks, histology, mechanical assays, and in vitro viability assays throughout the study to evaluate the structural integrity, free lipid content, cell viability, and overall shelf life of the product. The bioburden test before irradiation indicated that the contaminants present in the samples were no greater than 14 CFU. The bioburden test after irradiation supported no microbial contamination after irradiation and no growth results. Evaluate the tissue dimensions of a total of 20 samples at different time points from 0 to almost 6 months. All samples passed this evaluation at different time points. The results indicated that the product tissue dimensions remained consistent for up to almost 6 months.

[0131] Evaluate the color of a total of 22 samples at different time points from 0 to almost 6 months. All samples passed this evaluation at different time points. The results indicated that the product color was acceptable for up to almost 6 months.

[0132] Evaluate the free lipid content of a total of 22 samples at different time points from 0 to almost 6 months. All samples evaluated from 0 to 4 months passed this evaluation. Future studies could include evaluating triglyceride and glycerol content over time to quantify the free lipid content and potential changes in trends.

[0133] At all time points, adipocytes and connective tissue structures were observable in histological images of samples stained with H&E and Masson's trichrome. Perform histological analysis on the samples. This histological data indicated that adipose allograft samples retained their native fat structure over time. Figure 15- Figure 17Images of fat allograft samples from a single donor stained with H&E at time points of 2, 3, 4, and 6 months are shown. Figure 18 Images of stained fat allograft samples from another donor at 2, 3, and 4.5 months are shown. Figure 19 Images of stained fat allograft samples from another donor at 1, 1.5, and 2.5 months are shown. At all time points, adipocytes and connective tissue structures were observable in most histological images of samples stained with Oil Red O. Histological images showing poor sample structure were prepared in the same order, indicating poor sample preparation techniques from the histological laboratory.

[0134] Mechanical measurement results revealed that there appeared to be a slight trend of decrease in the mechanical hardness of fat allograft samples. Mechanical hardness over time for two different donor samples and an untreated sample. However, there was no conclusive statistical evidence to support the decreasing trend because the sample sizes for each time point and donor were not large enough to perform a statistical analysis.

[0135] A total of 8 samples were evaluated for in vitro cell viability at different time points from 0 to almost 6 months. In vitro cell viability assays were performed with various samples. All samples passed the assay, thus indicating that the survival or growth of fibroblasts was not negatively affected by the test samples for up to almost 4 months. Four months was chosen as the maximum time point because at least 3 samples were assayed at this time point compared to a single sample at 6 months.

[0136] Experimental Example 4

[0137] This study evaluated the differentiation of adipose-derived stem cells (ASCs) into adipocytes and determined whether the presence of test samples of tissue products, via controls or other test samples, altered the ASC differentiation properties.

[0138] Adipose-derived stem cells (ASCs) have drawn attention in the therapeutic applications of regenerative medicine due to the multiple properties of this lineage. ASCs have low immunogenicity and the ability to self-renew, can migrate to damaged sites, act through autocrine and paracrine pathways, and are pluripotent (able to differentiate into adipocyte, osteoblast, and chondrocyte lineages).

[0139] Materials and methods are now described. ASCs were thawed from liquid nitrogen and prepared for seeding in T150 culture flasks. After the T150 flasks reached 90% confluence, 6,579 cells / cm 2Subculture into each well of a 24-well plate. Designated cells receive control dry medium without the test sample, control dry medium supplemented with the test sample, differentiation medium without the test sample, differentiation medium supplemented with the test sample, regular control dry medium, and regular differentiation medium. Replace the dry medium every 2 - 3 days until the cells in the wells designated as differentiation wells reach 85% - 90% confluence. After reaching confluence, replace the control dry medium with the differentiation medium at the same feeding frequency. Add inserts with the test sample one day after inoculation. Take pictures 24 hours, 48 hours, 1 week, and 2 weeks after adding the inserts. Two weeks later, check the cells for differentiation. Treat the cells with a fixative and then stain with Oil Red O to observe lipid formation. Then, elute the Oil Red O and read the absorbance using a spectrophotometer.

[0140] The results are now described. After culturing for two weeks, the cells were stained with Oil Red O and lipid formation was evaluated. Figures 23 - 32 The results after 2 weeks are shown. Figure 23 Cells in the control medium without the addition of the test sample. Figure 24 Differentiation wells (also control wells) without the addition of the test sample. Figure 25 Cells in the case of control medium and donor 1 test sample. Figure 26 Cells in the case of control medium and donor 2 test sample. Figure 27 Cells in the case of control medium and donor 3 test sample. Figure 31 Cells in the case of control medium and regular test sample. Figure 28 Cells in the case of differentiation medium and donor 1 test sample. Figure 29 Cells in the case of differentiation medium and donor 2 test sample. Figure 30 Cells in the case of differentiation medium and donor 3 test sample. Figure 32 Cells in the case of differentiation medium and regular sample. Figure 33 and Figure 34Presents the average absorbance of a test sample of an embodiment of the present invention using spectrophotometry relative to the optical density (OD) at 510 nm of a conventional one. The OD level is related to the amount of Oil Red O staining in ASCs differentiated into adipocytes. It was found that all ASC groups grown in the differentiation medium were substantially differentiated into adipocytes as shown by lipid formation identified by Oil Red O staining. No significant difference in the differentiation of ASCs into adipocytes was observed between ASCs grown in the differentiation medium in the presence of the test sample adipose allograft (average OD 1.5 ± 0.63 SD) and cells grown in the differentiation medium without the test sample (average OD 0.87 ± 0.24 SD). No significant difference in the differentiation of ASCs into adipocytes was observed between ASCs grown in the presence of the test sample in conventional differentiation (average OD 1.04 ± 0.09 SD) and cells grown in the differentiation medium without the test sample (average OD 0.87 ± 0.24 SD). The results also showed that the differentiation of ASCs into adipocytes can be supported by the presence of the test sample adipose allograft and conventional ones, and slightly better differentiation was observed in the presence of the adipose allograft.

[0141] Experimental Example 5

[0142] In this experiment, the trans-well paracrine effect of the test sample on cell proliferation of endothelial cells was determined. Primary human dermal microvascular endothelial cells (HDBECs) are a subset of human dermal endothelial cells. They were isolated from the dermis of adult skin of a single donor. By flow cytometry analysis, the cells were analyzed to be CD31 positive and podoplanin negative. Vascular endothelial cells play a key role in physiological processes such as vasotension, capillary permeability, blood coagulation, fibrinolysis, and angiogenesis. Since primary HDBECs are used in tissue engineering models, primary HDBECs were selected for this trial (Pappalardo A et al., 2023. Sci Adv).

[0143] Now the materials and methods are described. To determine the effect on cell proliferation, 7,895 cells / cm 2Primary HDBECs were seeded in 24-well plates and the plates were grown in complete growth medium and in the presence of the test sample. After 3 days of culture, the cells were stained with crystal violet (CV) and the change in total cell number was evaluated. Positive control wells and negative control wells were established using complete growth medium relative to basal medium, in the absence of the test sample, to observe cell proliferation of primary HDBECs. Cell proliferation was expected in wells with primary HDBECs grown in complete growth medium, which established the assay positive control. No cell proliferation was expected in wells with primary HDBECs grown in basal medium, thus establishing the assay negative control. After 3 days of culture, the cells were stained with CV and the change in total cell number was evaluated.

[0144] (A) Before addition of the test sample, then at (B) 24 hours, (C) 48 hours, (D) 72 hours, and (E) after introduction of the insert with the test sample at 1% CV staining, test wells with the test sample identified as "conventional". The appearance of all three test wells with the conventional sample was similar, and a representative image of one well is presented in Figure 35 above.

[0145] (A) Before addition of the test sample, then at (B) 24 hours, (C) 48 hours, (D) 72 hours, and (E) after introduction of the insert with the test sample at 1% CV staining, test wells with the test sample identified as an embodiment of the present invention. The appearance of all three test wells of one embodiment of the present invention was similar, and a representative image of one well is presented in Figure 36 herein.

[0146] A) Before introduction of the test sample into the test well, then at B) 24 hours, C) 48 hours, D) after introduction of the insert with the test sample at 1% CV staining at 72 hours, control wells with primary HDBECs and complete growth medium identified as "positive control". The appearance of all three control wells was similar, and a representative image of one well is presented in Figure 37 above.

[0147] A) Before introduction of the test sample into the test well, then at B) 24 hours, C) 48 hours, and D) after introduction of the insert with the test sample at 1% CV staining at 72 hours, control wells with primary HDBECs and basal medium identified as "negative control". The appearance of all three control wells was similar, and a representative image of one well is presented in Figure 38 herein.

[0148] Now refer to Figures 39 - 42, the results are now described. The data presented in the table show the absorbance values read from a microplate reader (CLARIOstar). After 3 days of culture, the cells were stained with CV to observe changes in cell number. The optical density (OD) level shows the amount of CV dye eluted from the fixed cells (absorbance - 590 nm). No significant difference in total cell number was observed between primary HDBECs grown in the presence of the test sample adipose allograft (mean OD 4.96 ± 0.71 SD) and primary HDBECs grown in complete medium without the test sample (mean OD 4.92 ± 0.61 SD). These results indicate that the presence of the test sample adipose allograft does not have a negative impact on the cell proliferation of primary HDBECs in vitro. No significant difference in cell proliferation was observed between the conventional test sample and the adipose allograft test sample. The test samples, namely both the adipose allograft and the conventional one, do not have a negative impact on the cell proliferation of primary HDBECs in vitro. No significant difference in cell proliferation was observed between the conventional test sample and the adipose allograft test sample. As expected, a significant difference in cell proliferation of primary HDBECs was observed in the positive control (mean OD 4.92 ± 0.61 SD) compared to the negative control (mean OD 0.90 ± 0.16 SD). Cell proliferation of primary HDBECs was observed in the positive control. No cell proliferation of primary HDBECs was observed in the negative control.

[0149] Experimental Example 6

[0150] The purpose of this experiment was to quantify the percentage of adipocyte composition of the embodiments of the processed tissue described herein.

[0151] Human cells, tissues, and cell- and tissue-based products (HCT / P) are defined in Title 21 of the Code of Federal Regulations (CFR), Part 1271, as products that contain or consist of human cells or tissues and are intended for implantation, transplantation, infusion, or transfer into a human recipient (FDA 2020). HCT / P are regulated only under Part 361 of the PHS Act if all of the criteria set forth are met. One of the HCT / P regulatory criteria is that tissue processing must maintain minimal manipulation.

[0152] Minimal manipulation is defined in 21 CFR 1271.10(a) as a process that does not alter the original relevant characteristics of the tissue related to the tissue's utility for reconstruction, repair, or replacement. Since adipose tissue is mainly composed of adipocytes and surrounding connective tissue, the FDA classifies the original relevant characteristics as mainly providing cushioning and support for the body. Therefore, to evaluate whether the manufacturing of adipose-derived HCT / P meets the minimal manipulation standard, it should be considered whether the process alters the utility of the HCT / P to provide cushioning and structural support (FDA 2020).

[0153] For example, the manufacturing of adipose-derived HCT / P in which the cellular components are completely removed from the surrounding connective tissue (decellularization) will be considered by the FDA to be more than minimal manipulation. This is because the decellularization process alters its ability to provide cushioning and support. To show that the processed tissue according to the embodiments described herein adheres to the minimal manipulation standard, this report describes how to retain the adipocyte structure and connective tissue contents.

[0154] One of the most distinguishable features of the processed tissue is the retention of structurally intact adipocytes. The adipocyte composition of each product is quantified to support the argument for maintaining minimal manipulation. In adipose tissue, hematoxylin and eosin (H&E) staining is a useful tool for distinguishing the surrounding connective tissue from adipocytes. The quantitative evaluation of the processed tissue embodiments is completed using the Fiji-adapted adiposoft plugin for Image J analysis software( Figure 43 ). The adiposoft plugin is designed to identify individual adipocytes and calculate the area inside each adipocyte; at this time, the software generates a report. By obtaining the ratio of the adipocyte area to the total sample area, the percentage of adipocyte composition of each sample can be calculated (Equation 1, Figure 44 ).

[0155] Adipocyte structure quantification: The main purpose of this experiment is to quantify the percentage of adipocyte composition content in adipose allograft samples. This is accomplished by histological staining and Image J analysis.

[0156] Terminology :

[0157] HCT / P (Human cells and tissue products)

[0158] H&E (Hematoxylin and eosin)

[0159] Materials (including but not limited to) :

[0160] Formalin

[0161] 5mL conical tube

[0162] Nitrile gloves

[0163] Clamp

[0164] Procedures :

[0165] (1) Five tissues were fabricated from five tissue donors to account for donor variability.

[0166] (2) After fabrication, samples were taken from each product iteration, fixed in formalin in 5 mL conical tubes, and submitted for H&E histology staining.

[0167] (3) For each histological slide, five images of each sample were captured to obtain a general characterization ( Figure 45 ).

[0168] (4) For each sample stain, all five images were analyzed via Image J software to measure adipocyte area and total sample area. The average percentage of adipocyte composition was calculated by Formula 1.

[0169] Results

[0170] Figures 46 - 48 Depicted are H&E stained images of the first processed tissue and the second processed tissue ([[]] Figure 46 and Figure 47 ) processed according to the embodiments described herein, as well as a conventional processed tissue ([[]] Figure 48 ). For each sample stain, five sections were analyzed via Image J software and the average percentage of adipocyte composition was calculated.

[0171] 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 was evaluated by sampling from five independent donors. The adipocyte percentage range for the first tissue embodiment was calculated to be roughly between 45% - 70%, while the range for the second tissue embodiment was determined to be between 30% - 55%. The difference in adipocyte content can be attributed to the variation in the number of wash steps between the two processing procedures.

[0172] Discussion

[0173] Comparison of H&E staining showed that both the first and second tissue products contained a mixture of extracellular matrix (ECM) components (pink) and intact adipocytes (pink circles enclosing white spaces), while the conventional tissue showed only elements of ECM in its product. This unique feature distinguished the first and second tissue products from the conventional third product and confirmed the instrumental advantage in providing cushioning and support (the core feature of fat). In addition, the report provided evidence that the first / second tissue complied with the minimal manipulation requirements of HCT / P by preserving adipocyte structure and connective tissue content. There was no available processed tissue showing retention of both intact adipocytes and connective tissue content. Thus, both the first allograft product and the second allograft product incorporated unique structural features that had not been previously exploited.

[0174] FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use (July 2020).

[0175] Rasband, W.S., ImageJ, U.S. National Institutes of Health, Bethesda, Maryland, USA.

[0176] Experimental Example 7

[0177] The purpose of this experiment was to evaluate the adipose tissue composition of the processed tissue products according to the embodiments described herein. Tissue composition will be evaluated by measuring water content, lipid content, and protein content.

[0178] 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 major player in metabolic regulation (Luo 2016). Connective tissue is mainly composed of different types of extracellular matrix (ECM) proteins (including collagen, glycoproteins, and proteoglycans responsible for maintaining structural integrity and other roles). ECM also controls various physiological processes, including cell attachment, differentiation, survival, and development. In addition, evidence suggests that ECM networks are required for tissue remodeling and repair (Ruiz-Ojeda 2019).

[0179] Generally, the three most abundant components of adipose tissue are lipids, ECM proteins, and water. Over the years, many studies have been conducted to measure the compositional quantification in adipose tissue. McKee et al. performed a meta-analysis review of various papers, thus reporting proteomic estimates of the absolute and relative amounts of ECM proteins in adipose tissue (McKee 2019). Additionally, 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 typically consists of 75%-80% lipids, 15%-20% water, and 3%-5% protein (McKee 2019). The authors' findings on adipose component quantification will be used as a benchmark for the expected percentages of all three components in adipose allograft samples.

[0180] Results :

[0181] For this experiment, three processed tissues according to one allograft implementation and three processed tissues according to a second allograft implementation were made from three tissue donors; and each donor was divided and processed to make both a first product and a second product. Table 1( Figure 51 ) depicts the percentage of water, lipid, and protein content of the first allograft implementation samples tested. The bottom row of Table 1 shows the percentage ranges for each component: 19%-47% water content, 47%-80% lipid content, and 2%-7% protein content. Figure 52 Illustrates the compositional percentages of the first product made from tissue donor 1. Table 2( Figure 53 ) shows the compositional quantification of the second product samples. The percentages of the second product samples are as follows: 15%-36% water content, 60%-84% lipid content, and 2%-4% protein content.

[0182] Discussion of Results

[0183] The results showed that the similar compositional quantification of the first tissue sample and the second tissue sample was similar to the published literature on adipose tissue, with some minor differences. Compared to the second product samples, the first product samples showed slightly higher water content but lower lipid content. Additionally, the first product samples showed a higher range of protein content.

[0184] Differences in the compositional analysis of Donor 3 were noted. Since the calculated percentage composition for this donor did not fall within the range of Donors 1 and 2, the tests were repeated for Donor 3 to ensure the accuracy of the results. Based on the results of the second readings, it was confirmed that the results were accurate. The data confirmed that Donor 3 had a greater amount of connective tissue compared to the other two donors, which could also be the reason for the increased water content in the sample. Additionally, Donor 3 showed a higher protein content compared to Donors 1 and 2. Based on this information, the compositional range was reported as a single average to account for donor variability.

[0185] FDA Guidance, Regulatory Considerations for Human Cells, Tissues, and Cellular and Tissue-Based Products: Minimal Manipulation and Homologous Use (July 2020).

[0186] 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.

[0187] 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.

[0188] Ruiz-Ojeda FJ, Méndez-Gutiérrez A, Aguilera CM, Plaza-Díaz 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。

[0189] The disclosures of each and every patent, patent application, and publication cited herein are hereby incorporated by reference in their entirety. While the invention has been disclosed with reference to specific embodiments, it will be apparent to other skilled in the art that other embodiments and variations of the invention may be designed without departing from the true spirit and scope of the invention.

Claims

1. A processed adipose tissue, the processed adipose tissue comprising: An extracellular matrix derived from human adipose tissue; and A plurality of adipocytes having a composition of intact adipocytes of at least 10%.

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

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

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

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

6. The processed adipose tissue according to claim 1, the processed adipose tissue further comprising: A plurality of structural proteins retained from the human adipose tissue, the plurality of structural proteins including at least one of collagen, glycoprotein, and proteoglycan.

7. The processed adipose tissue according to claim 1, the processed adipose tissue further comprising: A decellularized tissue having a reduced amount of allogeneic components, cellular components, and free lipids.

8. The processed adipose tissue according to claim 1, the processed adipose tissue being configured for implantation or injection into a patient.

9. The processed adipose tissue according to claim 1, the processed adipose tissue being configured to pass through an opening between 1 mm and 4.5 mm.

10. A method of processing adipose tissue, the method comprising: Freezing and thawing human adipose tissue; Separating the human adipose tissue into pieces by mechanical grinding; Successively rinsing the human adipose tissue with a nonionic surfactant, a crystalline liquid, and sterile water; And Dispensing the human adipose tissue into a package using a loading tool.

11. The method according to claim 10, wherein the loading tool comprises: A wide-mouth bottle having a proximal end opening, a distal end opening, and a chamber disposed between the proximal end opening and the distal end opening; A piston actuated by a lever and configured to slide distally and flush along the wall of the chamber; And A bracket configured to stabilize the piston under the lever and form a hinged connection with the lever.

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

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

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

15. The method according to claim 10, the method further comprising: Debriding the human adipose tissue to remove foreign or abraded tissue.

16. The method according to claim 10, the method further comprising: The human adipose tissue is rinsed with a non-ionic surfactant, a sterile saline solution, and sterile water by dispensing the human adipose tissue into a centrifuge tube and centrifuging the human adipose tissue.

17. The method according to claim 10, the method further comprising: The human adipose tissue is rinsed with a non-ionic surfactant, a sterile saline solution, and sterile water by dispensing the human adipose tissue into a sealable container and gently agitating the container.

18. The method according to claim 10, the method further comprising: Rinsing the tissue in a non-ionic surfactant at least twice.

19. The method according to claim 10, the method further comprising: Rinsing the tissue in a crystalline saline solution at least twice.

20. The method according to claim 10, the method further comprising: Rinsing the tissue in sterile water at least twice.

21. The method according to claim 10, the method further comprising: Drying the human adipose tissue in a centrifuge tube including a drying sieve.

22. The method according to claim 10, the method further comprising: Decanting the tissue after each rinse and removing any decanted liquid and cellular components.

23. The method according to claim 10, the method further comprising: Using the loading tool to uniformly dispense the human adipose tissue in a container.

24. The method according to claim 19, the method further comprising: Packaging the container in a sterile pouch.

25. A medium loading tool, the medium loading tool comprising: A wide-mouth bottle having a proximal end opening, a distal end opening, and a chamber disposed between the proximal end opening and the distal end opening; A piston actuated by a lever and configured to slide distally and flush along the wall of the chamber; And A bracket configured to stabilize the piston under the lever and form a hinged connection with the lever.

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

27. The medium loading tool according to claim 25, the medium loading tool further comprising: An adapter configured to connect to the distal opening, the adapter having a first sealable port.

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

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