JP2017520517A - Sustained release angiogenesis modulating compositions and methods for inducing and modulating angiogenesis - Google Patents

Abstract

The present disclosure relates to compositions comprising human placental extract and biodegradable microparticles, sustained release angiogenesis modulating compositions, and compositions and methods for releasing placental extract over a period of time to a target, and blood vessels Methods are provided for inducing and / or modulating neoplasia and for identifying modulators of angiogenesis. The present disclosure also provides a method of making a composition comprising a placental extract capable of inducing and / or modulating angiogenesis.

Description

CROSS REFERENCE TO RELATED APPLICATIONS This application is filed on May 8, 2014 in application number 61 / 990,256, “Compositions and methods for inducing and modulating angiogenesis, and methods for identifying angiogenesis modulators and Priority is claimed on the US provisional application entitled "Assay", which is incorporated herein by reference in its entirety.

DESCRIPTION OF GOVERNMENT SUPPORTED RESEARCH OR DEVELOPMENT This invention was invented with government support under grant number HL088207 awarded by the National Institutes of Health. The government has certain rights in this invention.

  Angiogenesis allows the formation of blood vessels in physiological and pathological conditions ranging from wound healing to cancer. Angiogenesis regulation depends on both location and stimulation, and each situation may involve a unique combination of regulatory molecules.

  The inability to revascularize engineered organs and inability to revascularize infarcted areas is a major challenge for successfully bringing regenerative medicine treatment to the clinic. The ability to regulate angiogenesis in a definitive manner ranges from defining normal and pathological vascular physiology to tissue / organ regeneration, wound healing, infarct tissue repair and cancer inhibition. It is believed to have a significant impact in clinical use. A variety of different approaches have been taken to initiate angiogenesis and drive larger angiogenesis, including direct cell seeding (alone and co-culture), the use of stem cells and human-derived modulator / growth factor combinations Yes. To date, these in vitro approaches that typically use non-human animal compounds have been genetically regulated in the case of methods that control their discrete protein composition, non-human origin, tumor origin, or gene expression. Due to the lack, it has hardly been brought to the clinic.

  Current methods for inducing angiogenesis in vitro are by simple combinations of human-derived modulators, use animal-derived stimulators, or rely entirely on the use of live animals for evaluation Yes. When using these current in vitro models consisting of simple combinations of human and / or animal-derived modulators to test for effective angiogenesis inhibitors, the screening process is limited. This is because they do not represent a series of broad human in vivo molecular interactions. Modulating only selected molecular pathways also limits attempts to prevascularize engineered organs. This is because regulating angiogenesis requires induction of many metabolic pathways. Also, at present, the most popular and successful approach uses Matrigel ™, a substance derived from Angel Breath-Home-Swan mouse sarcoma cells, which is inappropriate for human therapy. It is thought that there is.

  Thus, it is believed that improved human-based methods for inducing and regulating angiogenesis can spur both drug development and regenerative medicine.

  Briefly described, embodiments of the present disclosure provide human placenta extract compositions comprising human placenta extracts, sustained release angiogenesis modulating compositions, methods of inducing angiogenesis and anti-inflammatory compositions and methods. .

  The present disclosure provides a composition comprising a human placenta extract associated with the microparticle so that the human placenta extract is released from the biodegradable microparticle after exposure to the cell in vivo or in vitro. To do. The human placenta extract is obtained from a human placenta sample, blood and solids are substantially removed from the extract, and the extract contains placental proteins including cytokines and growth factors that were present in the placenta sample. Including.

  An embodiment of the method of inducing angiogenesis of the present disclosure comprises contacting a sustained release angiogenesis modulating composition with a cell, wherein the sustained release angiogenesis modulating composition is live in a cell in vitro or in vivo. The biodegradable microparticles are associated with the human placenta extract of the present disclosure such that the human placenta extract is released from the microparticles over a period of time after exposure to the degradable microparticles. In embodiments, the method further comprises combining a sustained release angiogenesis modulating composition with the biomaterial to induce angiogenesis in the biomaterial.

  The present disclosure also provides a method of reducing inflammation in a subject by administering a human placenta extract of the present disclosure or a sustained release / human placenta extract composition of the present disclosure to a subject in need of treatment for inflammation. .

  Other methods, compositions, features and advantages of the present disclosure will become apparent to those skilled in the art upon review of the following drawings and detailed description. All such additional compositions, methods, features and advantages are included in the present description and are intended to be within the scope of this disclosure.

  Further aspects of the present disclosure will be more readily appreciated upon review of the detailed description of various embodiments of the present disclosure described below when considered in conjunction with the accompanying drawings.

1A-1K show the formation of human placental extract (hPE) (also referred to herein as human placental matrix or hPM), characterization of hPE film, and angiogenic network formed on hPE film. FIG. FIG. 1A shows an embodiment of obtaining hPM by homogenization of placental ECM followed by urea solubilization and dialysis. 1B to 1C are SEM images showing the surface morphology of hPE, and FIGS. 1D to 1E are angiogenic networks when HUVECs are seeded on hPm thin film at 4 × 10 ⁴ cells / cm ² and cultured for 3 days. It is a SEM image of formation. FIG. 1F shows rhodamine phalloidin (shown as “red” -gray branched pathway) and DAPI (“blue” —bifurcated pathway), showing branched cellular filamentous pseudopods during angiogenic sprouting 1 day later on placental extract. (Shown as light gray dots inside). FIG. 1G shows rhodamine phalloidin (shown as “red” —gray branched pathway) and DAPI (“blue” —within branched pathway) showing a mature angiogenic network undergoing extensive cell streaking after 3 days. (Shown as light gray dots). FIG. 1H shows calcein (“green” —shown as gray branch) and DAPI (“blue” —light gray in branch) during early stage cell streaking and angiogenic network formation after 1 day on placental extract. HUVEC stained with (shown as dots). FIG. 1I shows DAPI (indicated by “blue” -grey dotted path) staining showing cell line formation of HUVEC after 3 days on placental extract. FIG. 1J shows HUVEC seeded in tissue culture plates at 4 × 10 ⁴ cells / cm ² and cultured in endothelial cell medium for 3 days. FIG. 1K shows an angiogenic network by HUVECs adhered at 4 × 10 ⁴ cells / cm ² on a placenta extract adhered to the surface of a tissue culture plate with 100 μL PE / cm ² and then cultured in endothelial cell medium for 3 days. Shows the formation of.

Figures 2A-2C show biochemical analysis of hPE and genetic analysis of HUVEC seeded on hPE. FIG. 2A is a bar graph showing cytokine analysis performed using a sandwich-based human angiogenic antibody array. Data were normalized on a scale ranging from negative control values (0%) to positive control values (100%) (data represent 3 biological replicates). 2B and 2C are bar graphs showing normalized spectral abundance factors (%) for immune-related (2B) and angiogenesis-related (2C) BM-related proteins, determined using LC-MS / MS. The value of the normalized spectral abundance coefficient of fibrinogen is shown as the sum of the FGA value and the FGG value, and laminin is shown as the sum of the values of LAMA2, LAMA4, LAMA5, LAMB1, LAMB2, LAMB3 and LAMC1. Figure 2D shows gene analysis was performed on 100 [mu] L PE / cm ² on a 80,000 HUVEC seeded 3 days at a density of cells / cm ^2. Some angiogenesis-related proteins not present in the lysate, including VEGFA, were upregulated by HUVEC when seeded on hPE. Data represent 4 biological replicates. P values are calculated using Student's t-test with repeated 2 ^ (-delta Ct) values for each gene in the control and treatment groups.

Figures 3A-3D show in vitro angiogenic networks formed on tissue culture flasks coated with hPE and Matrigel. FIG. 3A shows calcein stained HUVECs on hPE and Matrigel at various cell seeding densities after 1d (day), 3d and 5d. The angiogenic network maturation rate, defined as the time to maximum tubule number / mm ^2, was adjusted in hPE samples by varying the cell seeding density. By quantitative analysis, the 40,000 cell / cm ² angiogenic network took up to day 3 to reach its maximum capillary density (tubule / mm ² ), whereas the 80,000 cell / cm ² network had its maximum capillary It was found that the density was reached in 1 day (data not shown). In Matrigel samples, the angiogenic network was less clear after the first day. Scale bar, 200 microns. FIG. 3B does not form angiogenesis when WPMY-1 myofibroblasts are seeded on placental extract (FIG. 3B.i) but formed when seeded on Matrigel (FIG. 3B.ii). It shows that. FIG. 3C shows quantitative image analysis (masking) to determine angiogenic network parameters including average tubule length (mm), tubule density, number of bifurcations, number of meshes and tubule width. FIG. 3C is a series of bar graphs showing a comparison of angiogenic network parameters induced by hPE and Matrigel®, for samples seeded at 80,000 cells / cm ² by day 1, Matrigel The sample reached its maximum average tubule length, tubule density, branching point and number of meshes, indicating that apoptotic balls were formed by day 3, but hPM network parameters were more stable over 5 days of culture. .

Figures 4A-4C show screening of the anti-angiogenesis-inhibiting tumor suppressor protein thrombospondin-1 using an hPE-based angiogenesis screening assay. FIG. 4A shows HUVECs in which TSP-1 was added to the culture medium and seeded on hPE, Matrigel and control culture flasks (uncoated) for 1 day and then stained with calcein AM. Scale bar, 200 μm. The graph of FIG. 4B shows the average total capillary length [mm] and the average number of branch points that both decrease linearly with increasing TSP-1 concentration in hPE coated flasks. The graph in FIG. 4C shows a comparison of the normalized percent reduction in angiogenic network coverage. The culture plates coated with hPE had significantly higher sensitivity to TSP-1 concentrations than the culture plates coated with Matrigel, with R ² values of 0.97 and 0.36, respectively.

Figures 5A-5C show in vitro angiogenesis on 3D tissue constructs. FIG. 5A is a schematic diagram showing placenta-derived cells, scaffolds and cytokines to induce in vitro angiogenesis in living scaffolds immersed in hPE (human umbilical vein) after 3 days of seeding and culture. . FIG. 5B shows that the HUVEC seeded tissue scaffold without hPE soaking did not form an angiogenic network. FIG. 5C shows a series of representative images of a bioscaffold immersed in hPE, showing the appearance of both angiogenic budding and invagination mechanisms after 3 days in culture. Budding angiogenesis was most prevalent at HUVEC cell seeding density of 20,000 cells / cm ² (FIGS. 5Ci-5Ciii). At a seeding density of 40,000 cells / cm ² , the appearance of both budding and invagination angiogenesis was observed (FIGS. 5C.iii.-5C.iv.), but a density of 60,000 cells / cm ² . In (FIG. 5Cv-5C.vi.), Angiogenic tubules were formed by invagination.

Figures 6A-6B show diagrams of in vivo angiogenesis in living scaffolds incubated with hPE. FIG. 6A is a schematic showing a decellularized HUV scaffold implanted in a rat model between fascia and muscle layer after 2 hours incubation in PE, Matrigel or phosphate buffered saline (control). is there. FIG. 6B shows a series of images showing the scaffold removed for analysis after 5d implantation. Significantly more fibrotic capsule formation occurred in the control and the biological scaffold incubated with Matrigel compared to the scaffold incubated with hPE (FIGS. 6Bi-6B.iii.). Bright field images taken through the front of a translucent bioscaffold sheet show significantly improved capillary network formation with scaffolds incubated with Matrigel and hPE (Figures 6B.iv.-6B.vi.) compared to controls And shows that there is the most mature capillary bed in the hPE scaffold, which indicates the formation of a vascular structure with a connected blood flow from arterioles to capillaries to venules (FIG. 6B.vi). (Circled with a dotted line)). Scaffolds incubated with hPE by hematoxylin and eosin staining (FIGS. 6B.vii.-6B.vi.) had the most scaffold modeling compared to control and Matrigel scaffolds. It was revealed. The control scaffold (FIG. 6B.vi.) has little remodeling of the original fiber orientation and is from the ablumenal surface of the HUV bioscaffold (shown in italic “l”) to the scaffold. Cell migration (migration) was also minimal. Scaffolds incubated with Matrigel had slightly less cell migration from the ablumen surface of the HUV compared to the hPE scaffold (FIGS. 6B.vii.-6B.ix.). When compared to the control, the scaffold incubated with Matrigel also has a heterogeneous cell distribution, less scaffold modeling than the scaffold incubated with hPE, and the scaffold incubated with hPE exhibits a new collagen fiber orientation. And cell distribution was more uniform.

7A-7E show an embodiment for the formation of an angiogenic network on a human umbilical vein scaffold (HUV) cultured using dynamic cell culture conditions. As shown by the schematics in FIGS. 7A and 7B, the cells were seeded after incubating the tubular HUV scaffold in placental extract for 2 hours and constructed in a double perfusion bioreactor under standard cell culture conditions for 5 days. Was cultured. Cells remained in the lumen of the scaffold and did not migrate (FIG. 7C). Cell streaking, the first stage of tubule formation, was sporadic (FIGS. 7D and 7E).

FIG. 8 shows an original image (left) of a tube and cell network growing on the PE and a zoom (right) of that region. The zoomed image shows how each capillary (dotted line) was identified and measured. The small circle on the right image indicates a branch point identified by a small dot with a number on the image pointed to by an arrow. Each hexagonal arrangement of capillaries identifies a mesh (dotted circle).

9A-B are a series of images (FIG. 9A) and bar graphs (FIG. 9B) showing the angiogenic ability of PE. The panel in FIG. 9A shows the cell morphology of microvessel tubules formed by HUVEC seeded on PE and cultured for 1, 3 and 5 days compared to HUVEC seeded on tissue culture plates. The capillary network began to form on day 1 and reached a more mature configuration on day 3 (see lower right enlarged view). On the fifth day, the network regressed. The number of meshes decreased and there were some isolated cells (white dots). The graph in FIG. 9B shows the morphological and tissue distribution characteristics of tubule-like networks formed on PE after 1, 3 and 5 days in culture. Capillary length, number of BPs and number of meshes were compared between 1, 3 and 5 days. Statistical differences (indicated by asterisks) in all three parameters were found between day 1 and day 5 cells. Statistical analysis was performed using a two-sided t-test with unequal variance at p <0.05.

FIGS. 10A-9B show the cellular morphology of the microvascular network formed by HUVEC seeded on PE and cultured for 5 days compared to HUVEC seeded on tissue culture plates (control). The control is shown in the inset in the upper right corner of the image of FIG. 10A. From top left, inoculation of PE only once on day 1, twice on days 1 and 3, and 3 times on days 1, 3 and 4. By increasing the number of inoculations (from 1 to 3), it is possible to notice that the capillary network has evolved to a more mature and long-lasting configuration. In the histogram in FIG. 10B, the capillary length, BP, and number of meshes are compared between day 1 and day 5. Statistical differences (indicated by asterisks) in all three parameters were found between cells that received one inoculation (1 INOC) and cells that received three inoculations (3 INOC). It was. Statistical analysis was performed using a two-sided t-test with unequal variance at p <0.05.

FIG. 11 shows a series of images showing HUVECs cultured for 3 days with PE stored in a humidified 6% CO ₂ incubator at 37 ° C. from day 1 to day 20. The numbers on each image indicate the incubation time (days). PE maintained its ability to induce angiogenesis even after 15 days of incubation.

12A to 12B are diagrams showing characteristics of gelatin microparticles. In FIG. 12A, the top row shows an SEM image of empty particles approximately 40 μm in size. The particles have a regular shape and a smooth surface. The lower row of FIG. 12A shows an SEM image of PE-added particles. The change in surface morphology and increase in size is probably due to PE sorption. The graph in FIG. 12B shows a size distribution analysis of gelatin microparticles by optical microscopy.

FIG. 13 is a diagram showing a decomposition profile of non-crosslinked microparticles. In vitro cumulative percent degradation of uncrosslinked microparticles is shown as a function of their dry weight. The total percent cumulative degradation was 18.65% ± 0.09 after 20 days. The inset graph shows degradation after 6 hours of incubation (6.45% ± 0.12). Error bars represent the mean ± standard deviation at n = 3.

Figures 14A-14B show protein release kinetics of empty and added microparticles. FIG. 14A shows the in vitro cumulative percent release from empty (no PE) and added (PE) microparticles as a function of their dry and added weight, respectively. The total percent cumulative release was 4.65% ± 0.11 and 4.65% ± 0.07, respectively, after 22 days. Error bars represent the mean ± standard deviation at n = 3. FIG. 14B shows the in vitro difference in percent release between added microparticles and empty microparticles. The first peak shows the release of PE from the surface of the particle and the second peak is probably due to massive erosion.

FIG. 15 shows PE delivery using gelatin microparticles. The image shows the response of HUVEC seeded at a density of 20,000 cells / cm ² to PE-added microparticles after 5 days in culture (D5) compared to HUVEC seeded with empty microparticles (control). Arrows point to cells under different conditions, indicating differences in shape. Some budding was present after 5 days of culture with PE-added particles, but no network formation was observed. Larger spheres are microparticles.

16A-16G show the effect of continuous delivery of hPE to HUVEC. FIG. 16A shows a control with HUVEC cultured at 20,000 cells / cm ² in angiogenic medium. Figures 16B-16D show a single inoculation of hPM on day 1 (Figure 16B), two inoculations on days 1 and 3 (Figure 16C) and three inoculations on days 1, 3 and 5 ( FIG. 16D) shows the HUVEC. 16E-16G are bar graphs showing angiogenesis quantification performed on the images of FIGS. 16B-16D after staining. Results are presented as the mean plus or minus mean standard error between measurements on 3 samples per point for each condition.

17A to 17F are diagrams showing evaluation of microparticles. Figures 17A-17C show the three protocols evaluated: single (2 minutes) homogenization (Figure 17A), double (2 minutes / 1 minute) homogenization (Figure 17B) and double (2 minutes / 20 seconds). FIG. 17 shows an optical microscope image for homogenization (FIG. 17C). The histograms of FIGS. 17D-17F show the size distribution in microns for particles formed in each of the three protocols (each batch prepared with 3 samples per point).

18A-18F added hPM for each of the three protocols, single (FIGS. 18A, 18D), 2/1 duplex (FIGS. 18B, 18E) and 2/20 duplex (FIGS. 18C, 18F). It is a scanning electron microscope (SEM) image of PLGA microparticles. 18A-18C show the microparticle shape and FIGS. 18D-18F show the surface porosity.

19A and 19F show a table of addition efficiency (FIG. 19A) and an assessment of release rate from the particles for each protocol (P2: single, P3: 2/1 duplex and P4 2/20 duplex). It is a figure which shows (FIG. 19B). The curve represents the cumulative release rate up to 21 days for each protocol.

FIG. 20 is a diagram showing SDS page analysis of the supernatant of microparticles. The images are of the gel after staining showing qualitative protein content in the supernatant of BSA and hPM-added microparticles, and the relative initial content added during microparticle preparation.

FIG. 21 shows angiogenic network stability between pure hPM added to the alginate matrix (top row) and sustained release of hPM from PLGA microparticles embedded in the matrix (second row). FIG. 2 is a series of images showing endothelial cells stained with calcein AM after 7, 14, 21 and 28 days in culture to assess gender differences. The bottom row shows two controls, one empty PLGA microparticle embedded in an alginate matrix and the second no microparticle. HUVEC remained generally circular throughout all time points for both controls.

  Before describing the present disclosure in greater detail, it will be understood that the present disclosure is not limited to the particular embodiments described and, of course, may vary. It will also be understood that the terminology used herein is for the purpose of describing particular embodiments only and is not intended to be limiting.

  When indicating a range of values, each intervening value between the upper and lower limits of the range, up to one-tenth of the lower limit unit, unless the context clearly indicates otherwise, and the stated range It will be understood that any other value or intervening value is encompassed by the present disclosure. The upper and lower limits for these smaller ranges, apart from the limits specifically excluded in the stated ranges, can be included independently in the smaller ranges and are also included in the present disclosure. Where the stated range includes one or both of the limits, ranges excluding either or both of those included limits are also included in the disclosure.

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

  Any publications and patents cited herein, which are incorporated by reference, are hereby incorporated by reference to disclose and describe the methods and / or materials to which the cited publications relate. The citation of any publication is for disclosure prior to the filing date and should not be construed as an admission that the disclosure is not prior to such publication by prior disclosure. In addition, the publication dates listed may differ from the actual publication dates that need to be independently confirmed.

  As will become apparent to those skilled in the art upon reading this disclosure, each individual embodiment described and shown herein has distinct components and characteristics, which depart from the scope or spirit of this disclosure. Without, it can be easily separated from any of the features of some other embodiments, or can be combined with such features. Any described method can be performed in the order of events recited or in any other order which is logically possible.

  Embodiments of the present disclosure employ techniques such as medical, biochemistry, molecular biology, biology, pharmacology, etc. that are within the skill of the art unless otherwise indicated. Such techniques are explained fully in the literature.

  It should be noted that as used in the specification and the appended embodiments, the singular forms “a”, “an”, and “the” include the plural unless the context clearly dictates otherwise. Thus, for example, reference to “a cell” includes a plurality of cells. In this specification and in the embodiments that follow, reference will be made to a number of terms that are defined to have the following meanings unless a contrary intention is apparent.

  Prior to describing the various embodiments, the following definitions are provided, and the following definitions are used unless otherwise indicated.

(Definition)
In describing the disclosed subject matter, the following terms are used in accordance with the definitions set forth below.

  The terms “polypeptide” and “protein” as used herein refer to an amino acid polymer of three or more consecutive amino acids linked by peptide bonds. The term “polypeptide” includes proteins, protein fragments, protein analogs, oligopeptides and the like. The term “polypeptide” is a polypeptide as defined above, encoded by a nucleic acid, produced by recombinant techniques (isolated from a suitable source such as birds), or synthesized. Contemplate. The term “polypeptide” further contemplates a polypeptide as defined above comprising a chemically modified amino acid or an amino acid covalently or non-covalently linked to a labeled ligand.

  The terms “polynucleotide”, “oligonucleotide” and “nucleic acid sequence” are used interchangeably herein and refer to a coding sequence (in vitro or in vivo when placed under the control of appropriate regulatory or regulatory sequences). Polynucleotides or nucleic acid sequences that are transcribed and translated into polypeptides in), regulatory sequences (eg, translation initiation and termination codons, promoter sequences, ribosome binding sites, polyadenylation signals, transcription factor binding sites, transcription termination sequences, upstream and downstream Including, but not limited to, regulatory domains, enhancers, silencers, and regulatory sequences (DNA sequences to which transcription factors bind to alter the activity of the promoter of the gene to positive (induction) or negative (repression)). The terms described herein do not suggest a limitation on length or synthetic origin.

  The term “gene” as used herein refers to a nucleic acid sequence (both RNA or DNA) that encodes genetic information for the synthesis of whole RNA, whole protein, or any part of such whole RNA or whole protein. Including). A “gene” refers to a heritable unit that typically corresponds to a sequence of DNA that occupies a particular location on a chromosome, and includes genetic instructions for characteristics or traits in an organism. The term “gene product” refers to the RNA or protein encoded by a gene.

  The terms “treat”, “treat” and “treatment” are approaches for obtaining beneficial or desired clinical results. Specifically, beneficial or desired clinical outcomes include detectable or undetectable symptoms relief, disease severity reduction, disease stabilization (eg, no worsening), disease progression delay or slowdown, disease progression Including but not limited to substantial prevention of spread, improvement of disease status or temporary alleviation and mitigation (partial or complete). In addition, “treat”, “treat” and “treatment” may also be therapeutic in terms of partial or complete cure for the disease and / or adverse effects resulting from the disease. As used herein, the term “prophylactically treat” or “prophylactically treat” refers to completely, substantially or partially preventing a disease / condition or one or more of its symptoms in a host. Say. Similarly, “delaying the start of a condition” can also be included in “preventive treatment” and refers to the effect of increasing the time to the actual start of a condition in patients who are prone to the condition. Say.

  “Administering” means introducing a compound of the present disclosure into a subject. “Administration” can also refer to the act of providing (eg, by prescribing) a composition of the present disclosure to a subject.

  The terms “organism”, “subject” or “host” refer to humans, mammals (eg, cats, dogs, horses, mice, rats, pigs, hogs, cows and other cattle in need of treatment. , Refers to any organism in need of treatment, including birds (eg, chickens) and other living species, in particular, the term “host” includes humans, and as used herein, the term “human host”. Or “human subject” is used generically to refer to a human host, and in this disclosure the term “host” typically refers to a human host, so when used alone in this disclosure, “host The term "" refers to a human host, unless the context clearly indicates that it is intended to represent a non-human host. Or do not show symptoms of multiple overt, it inherited a risk of developing one or more of these conditions can be defined as hosts that have physiologically or other in the form.

  The term “expression”, as used herein, refers to the process that a constitutive gene undergoes to produce a polypeptide. This is a combination of transcription and translation. Expression generally refers to “expression” of a nucleic acid to produce a polypeptide, but refers to “expression” of a polypeptide, which indicates that the polypeptide is produced through expression of the corresponding nucleic acid. It is generally acceptable.

  “Angiogenesis” is a physiological process involving the growth of new blood vessels. Angiogenesis is an important part of biological processes such as growth and development, wound healing, embryogenesis and the like. Excessive angiogenesis can occur when diseased cells produce abnormal amounts of angiogenic growth factors that overwhelm the effects of natural angiogenesis inhibitors. An imbalance between the production of angiogenic growth factors and angiogenesis inhibitors can cause inappropriately regulated growth or inhibition of blood vessels. When new blood vessels grow excessively or insufficiently, they become angiogenesis-dependent diseases or angiogenesis-related diseases. Angiogenesis-related diseases can include, for example, but are not limited to, cancer, precancerous tissue, tumors, heart infarction and stroke. Excessive angiogenesis can include cancer, diabetic blindness, age-related macular degeneration, rheumatoid arthritis, psoriasis, and more than 70 other conditions. Insufficient angiogenesis can include coronary artery disease, stroke and delayed wound healing, and is also a factor in tissue engineering as discussed in more detail in this disclosure.

  As used herein, the term “modulate” and / or “modulator” directly or indirectly promotes / activates / induces / increases a specific function and / or trait in a cell / organism or This generally refers to interference / inhibition / reducing action. In some examples, a modulator may increase or decrease an activity or function relative to its natural state or compared to the average level of activity normally expected. Modulation can include causing overexpression or underexpression of the peptide (eg, by acting to upregulate or downregulate the expression of the peptide), or interact directly with the peptide of interest to activate Can be increased and / or decreased. Modulation also includes causing an increase or decrease in a specific biological activity or biological event, such as angiogenesis or a biological event associated with angiogenesis.

  As used herein, “upregulate” refers to the effect of increasing the expression and / or activity of a protein or other gene product. “Down-regulation” refers to reducing the expression and / or activity of a protein or other gene product.

  The term “isolated cell or population of cells” as used herein refers to an isolated cell or cells that have been excised from a tissue or grown in vitro by tissue culture techniques. The term “cell or population of cells” refers to the isolated cells described above or may refer to cells in vivo in animal or human tissue.

  The term “tissue” refers to a tissue that cooperatively performs a biological function and / or performs a biological purpose, such as forming all or part of an organ in an organism (eg, connective tissue, endothelial tissue). A generalized group of cells. A “tissue” generally includes a group of similar cells, or all cells of the same type, but a tissue may also include more than one type of cell when the group of cells serves a common purpose as a whole. .

  As used herein, the term “biocompatibility” refers to a living biological material and / or biological system (without any undue stress, toxicity or harmful effects on the biological material or system). For example, the ability to coexist with cells, cell components, living tissues, organs, etc.).

  The term “biological scaffold” refers to any biocompatible substrate (naturally occurring or synthetic) that has sufficient structural stability to support the growth of living biological material (eg, living cells). Say. In embodiments of the present disclosure, the biocompatible scaffold material is derived from a naturally occurring substrate such as, but not limited to, a decellularized human umbilical vein scaffold, eg, from a living organism, but further processed and That can be treated or produced from a natural source). In embodiments, the biological scaffold of the present disclosure has a three-dimensional structure (rather than a planar, two-dimensional structure) to support three-dimensional growth of viable cells.

  As used herein, the term “biodegradable” refers to dissolution, alteration, or other degradation over time in a natural environment (eg, in a living organism or living culture) that degrades its structural integrity. A substance that is lost and no longer exists in its original structural form. In an embodiment of the present disclosure, the biodegradable material dissolves / degrades over a period of time in the host organism.

  As used herein, the term “engineered” indicates that the engineered object is created and / or modified by a human. Although engineered objects may contain naturally derived materials, the objects themselves have been modified somewhat by human intervention and design.

  As used herein, the term “test compound” can include peptides, peptidomimetics, small molecules, nucleic acid sequences, or other compounds that can affect living cells or organisms. In some embodiments, a “test compound” can be a compound such as a chemical or peptide suspected of having a modulating effect on biological activity, function, or response to another compound. For example, in the present disclosure, a “test compound” may have a modulating effect on angiogenesis, such as increasing angiogenic activity, decreasing angiogenic activity, and / or modulating the effects of different angiogenic modulators. Can be a suspected compound.

  As used herein, the term “removed” or “substantially removed” indicates that an amount of a substance or compound has been separated from another composition, but the removed substance is fully detected. It is not necessary that all traces of the material removed so as to be impossible are absolutely present in the remaining composition. For example, if blood is “removed” or “substantially removed” from the composition, this indicates that a substantial portion of the blood in the composition has been removed, but some Blood or blood components may still be detected in trace amounts by precise screening (eg “substantially removed” requires that the composition does not contain 100% of the “removed” component Instead, the composition or substance is about 99% removed, about 95% removed, or about 90% removed, or the above exemplified components removed. Any percentage or range within the percentage range may be removed).

(Discussion)
Embodiments of the present disclosure include methods and compositions for inducing angiogenesis, as well as methods and compositions for modulating angiogenesis, and methods for making compositions for modulating angiogenesis. The disclosure also includes methods for identifying angiogenesis modulators and assays for identifying angiogenesis modulators. Embodiments of the present disclosure further include methods and compositions for delivering compositions for modulating angiogenesis. In embodiments, the present disclosure provides a placental extract capable of inducing and / or modulating angiogenesis in vitro and / or in vivo in tissue constructs and / or natural tissues, and tissue constructs and / or natural tissues Methods and compositions for delivering placental extract to cells therein. The disclosure also includes placental extracts that can be used in assays to identify compounds that modulate angiogenesis. Furthermore, the present disclosure includes a composition of a delivery vehicle supplemented with placental extract for the controlled release of the extract to an in vivo or in vitro cell population to induce angiogenesis.

  Angiogenesis is a complex process that depends on both location and stimulation, and in each case the ability to modulate these processes can involve complex combinations of regulatory molecules. The control of angiogenesis is further complicated by different formation mechanisms, two of which are best understood are intussusception and sprouting. Invagination is characterized by the insertion of stromal cell columns into the lumen of existing blood vessels, and budding is characterized by endothelial cell sprouting towards angiogenic stimulation in tissues that previously had no microvessels. Many molecules have been found to regulate angiogenesis, and more may be discovered in the future. This diversity of angiogenesis inducers has driven the continued search and development of angiogenesis regulators for use in the study of angiogenesis, drug screening, and treatment with regenerative medicine.

  Conventional models for studying angiogenesis rely entirely on the use of animal derived stimulants or the use of live animals for evaluation. In vivo animal studies provide a more accurate model for comparing the complexity of biomolecular pathways and mechanisms that occur during human angiogenesis. Standard in vivo angiogenesis models include the rabbit corneal neovascularization assay, the in vivo / in vitro chick chorioallantoic membrane assay and the rat mesenteric fenestration assay. If possible, in vitro angiogenesis models are selected to better control complex biological phenomena. However, this limits the study to a limited number of molecular species, such as VEGF. In this complex cascade, a more complex or multi-factor “mix” may be necessary to promote sufficient angiogenesis, and what can be obtained using a single molecule (or several) is itself May be limited.

For an in vitro angiogenesis model, the mouse-derived basement membrane matrix (BMM) or “Matrigel” assay is the preferred model, but this assay introduces a degree of in vivo complexity into the in vitro model, and the results are in vivo. This seems to be comparable to the result of. However, this assay is not suitable for clinical use because it is derived from Angel Breath-Home-Swan (EHS) mouse sarcoma cells and requires the sacrifice of a large number of animals ¹¹ . Several in vitro human-derived modulators have been used to model angiogenesis. Historically, they are based on single regulators (FGF, TGF-β, VEGF) and lack the diversity of cytokines and chemical gradients inherently found in vivo ¹² . In view of the interspecies differences ^13,14 associated with animal-derived models and the complexity of obtaining multiple protein formulations from human recombinant proteins, a robust human-derived approach (more complex of multiple proteins at near physiological ratios) Will have a significant impact on mechanistic studies, angiogenic drug screening, and the ability to enhance the clinical transformation of regenerative medicine therapy. Furthermore, the ability to modulate the angiogenesis process to represent different mechanisms and stages of formation provides an improved platform that characterizes key molecules and molecular pathways during angiogenesis. Let's go.

  The present disclosure provides methods for inducing and modulating angiogenesis in vitro and in vivo. In addition to inducing in vitro and in vivo angiogenesis, this model allows for the regulation of the rate of microvascular network maturation and the selective modeling of budding and invading angiogenesis. In vivo, it has been shown that human placenta extract (PE) significantly enhances capillary formation and eliminates fibrosis using a collagen-based bioscaffold with components added.

  The present disclosure describes such methods for inducing and modulating angiogenesis in vitro and in vivo using a complex set of tunable fully human biomolecules derived from human placenta. This approach uses controlled fractionation and separation techniques to obtain complexes of active human biomolecules isolated from human placenta. In addition to inducing and modulating in vitro angiogenesis and in vivo angiogenesis, the disclosed methods and compositions modulate the rate of microvascular network maturation and selectively model budding and invading angiogenesis. To make it possible. In embodiments, the methods and compounds of the present disclosure induce and modulate angiogenesis both in polymeric and ex vivo derived tissue scaffolds. These methods make it possible to regulate the rate of microvascular network maturation. In vivo, using a collagen-based bioscaffold with components added, it has been shown that the human placenta extract of the present disclosure significantly enhances capillary formation and eliminates fibrosis.

  Sustained delivery of growth factors that lead to angiogenesis is also a challenge faced by successful modulation of angiogenesis to promote angiogenesis with respect to tissue engineering approaches. The present disclosure also provides methods and compositions for controlled release of the disclosed compositions for modulating angiogenesis both in vitro and in vivo.

(Human placenta extract)
The present disclosure provides compositions and methods for inducing and / or modulating angiogenesis, including human placenta extract (PE or hPE). In embodiments of the present disclosure, PE obtains a sample from human placenta, removes blood from the placenta sample to produce a crude placenta extract (crude PE), and mixes the crude PE with urea or other protein solubilizers Solubilize proteins present in the extract, remove residual solids from the crude extract, and dialyze the urea-placenta extract mixture to remove substantial amounts of urea from the mixture to produce human PE It is produced by this. Human PE is a matrix-like compound, sometimes referred to herein as human placenta matrix (hPM).

  In embodiments, the process for making human PE is performed at a temperature between about −86 ° C. and about 5 ° C. In embodiments, the human placenta extract is made at a temperature of about 4 ° C. or less.

  In an embodiment, the process of removing blood from a placenta sample to produce a crude placenta extract comprises homogenizing a human placenta sample with a buffer, centrifuging the homogenized sample, and blood Discarding the supernatant containing. This process involves removing substantially all of the blood from the sample (eg, the sample is about 99% blood removed, about 95% blood removed, about 90% blood removed , Etc.) can be repeated multiple times (eg, a few times or more) until crude PE is produced. In certain embodiments, the buffer is a sodium chloride solution (NaCl).

  In embodiments, the protein in the crude placenta extract is solubilized by mixing the crude placenta extract with a protein solubilizer. In embodiments, the protein solubilizer does not permanently destroy the protein or permanently inactivate the protein (eg, solubilization reversibly denatures the protein to solubilize the protein. The protein should be able to refold, such as when the agent is removed), and can be any compound or mixture of compounds that can solubilize (eg, denature) the protein. In embodiments, the protein solubilizer can be, but is not limited to, urea, guanidine-HCl or other similar compounds. In an embodiment, the protein solubilizer is urea and the crude extract is mixed with the urea composition by homogenizing the crude extract with urea. In embodiments, urea is mixed with the crude extract for a period of between about 12 to about 36 hours. In an embodiment, urea is mixed with the crude extract for about 24 hours. In an embodiment, the urea solution is a urea buffer having a urea concentration of about 0.5M or higher. In embodiments, the urea is about 2M or more, about 4M urea or more, up to about 15M. In embodiments, the urea solution can have a concentration of about 0.5M to about 15M. In other embodiments, the protein solubilizer is guanidine-HCl having a concentration of about 0.5M to about 15M. In an embodiment, guanidine-HCL has a concentration of about 6M. The methods and compositions described below are described using urea as a solubilizer, but other suitable solubilizers can replace urea with, for example, but not limited to, those discussed above. It is understood.

  In an embodiment, after mixing with urea or other protein solubilizer, solids are removed from the solubilized protein-crude extract mixture (eg, urea-crude extract mixture). In embodiments, the solids are removed by centrifuging the PE mixture and discarding the pellets (including solids). This step can be repeated multiple times. After removing the solids, the PE mixture (eg, supernatant) is dialyzed to remove urea or other protein solubilizers from the placental extract. In an embodiment, the dialysis solution is TBS. In an embodiment, the dialysis solution is replaced after a period of time (eg, 1 hour, 2 hours, 3 hours, etc.), and dialysis is repeated several times (eg, 2, 3, 4 times, etc.) to substantially all urea. (Eg, placenta extract has about 99% urea removed, about 95% urea removed, etc.). In an embodiment, the PE may be centrifuged again to remove any remaining solids (such as polymerized proteins). In an embodiment, the residual PE is a transparent to pinkish viscous material. Further details regarding embodiments of the disclosed process for making placental extracts of the present disclosure can be found in the examples below.

  Thus, embodiments of the present disclosure also include PE made by the method of the present disclosure. In an embodiment, the present disclosure removes blood from a sample obtained from a human placenta sample to produce crude PE, and the crude placenta extract is converted to a protein solubilizer (eg, but not limited to urea, guanidine-HCl). Etc.) to solubilize the protein in the crude extract, separate the solid material from the solubilized protein-PE mixture, dialyze the PE mixture, and protein solubilizer (eg urea) from the mixture PE produced by removing human to produce human PE.

  The present disclosure thus provides an extract obtained from a human placenta (eg, from a human placenta sample) that substantially removes blood and solids and retains (some or all) the placental protein that was present in the placenta sample. Including human placenta extract. In embodiments, placental proteins include cytokines and growth factors.

  Analysis of the PE of the present disclosure reveals that PE contains many proteins, including many cytokines and growth factors. In an embodiment of the placental extract of the present disclosure, the extract comprises at least 20 different cytokines. In some embodiments, the extract comprises up to 40 different cytokines. Other embodiments include at least 50 cytokines. Some cytokines that may be present in the PEs of the present disclosure include those listed in the examples below. For example, some of the cytokines that may be present in the PE of the present disclosure include, but are not limited to, angiogenin, Acrp30Ag, IGFBP-1, NAP-2 and Fas / TNFGSF6, and RANTES and MIF.

  Cytokines and growth factors and other placental compounds present in the placental extracts of the present disclosure can induce angiogenesis in endothelial cell cultures, tissues, tissue constructs, engineered biological scaffolds, and the like. The placental extracts of the present disclosure can induce angiogenesis in vitro and in vivo. The placental extracts of the present disclosure can stimulate endothelial cell growth. In embodiments, the human PE of the present disclosure can modulate angiogenesis. Compared to other conventional compounds used to induce angiogenesis, such as BMM (compounds containing a single purified angiogenesis modulator (eg, purified VEGF-alpha or SDF-1) and purified fibrin) and the present disclosure The PE stimulates an increase in endothelial cell angiogenesis growth (eg tubules and network formation) and a decrease in myofibroblast angiogenesis growth (eg tubule formation) compared to BMM. The PE of the present disclosure also stimulates different growth and / or differentiation patterns of various cell lines (eg, stem cells, smooth muscle cells, etc.) compared to BMM, so that the growth / differentiation pattern of such cells is It can be distinguished from growth using BMM.

  The PE of the present disclosure can also upregulate various genes in endothelial cells compared to endothelial cells grown in the absence of PE. Some such genes include angiogenesis-related genes, extracellular matrix remodeling genes and vascular development genes. Some angiogenesis-related genes include, but are not limited to, ANGPTL4, CXCL3, human growth factor (HGF), ANGPT2, PGF, TYMP, VEGFA, HIF1A and FGF1. Some extracellular matrix remodeling genes that can be induced by the placental extracts of the present disclosure include, but are not limited to, MMP2, MMP9, COL4A3 and LAMA5. Vascular development genes include, but are not limited to, CDH2, HAND2, LECT1 and MDK.

(How to regulate angiogenesis)
The present disclosure also includes a method of inducing angiogenesis in a cell culture, comprising growing endothelial cells in the presence of the human placenta extract of the present disclosure. In an embodiment, a placental extract of the present disclosure obtained from a human placenta sample that has been processed to remove blood and solids, mixed with urea, and dialyzed to remove urea, comprising cytokines and growth factors Cell cultures are grown in the presence of placental extract containing placental protein. In an embodiment, the endothelial cell is a human endothelial cell. In yet another embodiment, the cell is a human umbilical vein endothelial cell (HUVEC). In an embodiment of the method of inducing angiogenesis in a cell culture, the cells are seeded at a density of at least about 40,000 cells / cm ² . In embodiments, the cells are seeded at a density of at least about 80,000 cells / cm ² . In embodiments, cell cultures can be grown on plates containing growth media and placenta extracts of the present disclosure.

The present disclosure relates to a method for inducing angiogenesis in a biomaterial in vivo comprising incubating the biomaterial in a composition comprising the human placenta extract of the present disclosure and implanting the biomaterial into a host. Including. In an embodiment, the biomaterial comprises naturally derived material and / or cells. In an embodiment, the biomaterial comprises an engineered bioscaffold comprising a human derived substrate material. In embodiments, the engineered bioscaffold includes a human umbilical vein scaffold. In embodiments, the human umbilical vein scaffold is decellularized. In some embodiments, the biomaterial is seeded with endothelial cells such as, but not limited to, human endothelial cells (eg, HUVEC). In an embodiment of the present disclosure, the biomaterial comprises an engineered scaffold material, including a human umbilical vein scaffold seeded with HUVEC. In some embodiments, the HUVECs are seeded on the biological scaffold at a cell density of at least about 40,000 cells / cm ² . In embodiments, HUVEC are seeded at a density of at least about 80,000 cells / cm ² . In embodiments, the biomaterial is incubated in the placenta extract for at least about 2 hours.

(Angiogenesis into biomaterials and engineered bioscaffolds)
The present disclosure also includes methods for angiogenesis in biomaterials including, but not limited to, engineered biomaterials, naturally derived biomaterials, and other biomaterials implanted in a host. Treating the biomaterial with the placental extract of the present disclosure, in addition to angiogenesis in the biomaterial, aids biotolerability, reduces inflammation, reduces rejection and scar formation, etc. It can also be used to pretreat biomaterials for in-vivo applications. Thus, the placenta extract of the present disclosure and the composition comprising the placenta extract of the present disclosure can be used to “add components” to any number of biomaterials to improve the outcome of such transplantation. .

  The present disclosure is specifically engineered, such as an implantable engineered biological scaffold comprising a human-derived substrate material incubated in a composition comprising a human placenta extract of the present disclosure. Also includes biomaterials. The biological scaffold of the present disclosure can be transplanted into a mammal such as a human. The bioscaffold of the present disclosure can include any biomaterial suitable for implantation into a host. Examples of bioscaffolds for use in the present disclosure include, but are not limited to, engineered bioscaffolds including tissues, matrix materials, any number of naturally occurring biomaterials, and the like. In embodiments, the bioscaffold is a 2D or 3D bioscaffold. In an embodiment, the biological scaffold comprises a human derived substrate material. In an embodiment, the biological scaffold comprises a decellularized human umbilical vein scaffold. In embodiments, cells such as, but not limited to, human cells, human endothelial cells (eg, human umbilical vein endothelial cells (HUVEC)), stem cells, other pluripotent cells, etc. are seeded in a living scaffold.

  As described in the Examples below, the biological scaffolds of the present disclosure incubated in the placental extracts of the present disclosure have more angiogenesis than the biological scaffolds incubated in the angiogenesis-inducing compound BMM or the control compound ( Induces, for example, angiogenesis) and less fibrosis. Biological scaffolds incubated in the placental extract of the present disclosure are also immunosuppressive and vascular to pro-inflammatory positive macrophages (eg, CD86 (M1)), in contrast to biological scaffolds incubated in BMM or controls. The proportion of pro-neoplastic positive macrophages (eg CD205 (M2)) was higher.

(Angiogenesis screening assay)
The placental extract of the present disclosure induces angiogenesis in cell culture and thus provides a good assay for identifying and screening for angiogenesis modulators. Thus, the present disclosure also includes methods and assays for identifying angiogenesis modulators.

  In an embodiment, the method comprises growing a culture of human endothelial cells in the presence of a compound comprising a human placenta extract of the present disclosure and contacting the human endothelial cell culture with a test compound. Since placental extract induces angiogenesis in cell culture, a test compound can be identified as an angiogenesis modulator if angiogenesis is less or more than expected. Thus, this method also determines the amount of angiogenesis in the culture and whether the amount of angiogenesis in the cell culture is greater than the amount of angiogenesis in the growth of the culture in the absence of the test compound. Or, if small, identifying the test compound as an angiogenesis modulator. In embodiments, an increase in the amount of angiogenesis compared to a culture grown in the absence of the test compound indicates that the test compound induces angiogenesis. A decrease in the amount of angiogenesis compared to cultures grown in the absence of the test compound indicates that the test compound inhibits angiogenesis. As described in the Examples below, screening for the compound thrombospondin-1 (TSP-1) according to the methods of the present disclosure identified this compound as an inhibitor of angiogenesis. The present disclosure also provides an assay for screening test compounds to identify modulators of angiogenesis, including a culture of endothelial cells grown in the presence of the human placenta extract of the present disclosure. The assays of the present disclosure can be used with the methods of the present disclosure to identify modulators of angiogenesis.

(Sustained delivery methods and compositions)
Compositions and methods for sustained release of PE of the present disclosure as the angiogenic effect of placenta extract may not persist for a long time in vivo or in biomaterials (eg, tissue, cell culture, bioscaffold, etc.) Are also provided in this disclosure.

  Embodiments of the present disclosure include a composition comprising human PE (hPE) of the present disclosure combined with biodegradable microparticles to provide a sustained release / human PE composition. PE-added biodegradable microparticles provide a sustained release angiogenesis modulating composition. Thus, in an embodiment, a sustained release angiogenesis modulating composition comprises human PE of the present disclosure and biodegradable microparticles, wherein the human PE is microscopic so that human PE is released from the microparticles. Combined with particles. In embodiments, human PE is obtained from a human placenta sample, blood and solids are substantially removed from the extract, and the extract contains placental proteins (cytokines and growth factors present in the placenta sample). Including). The human PE used in the sustained release angiogenesis modulating composition can be any embodiment of the human PE described above.

  In embodiments, human PE is released from the microparticles after being exposed to the cells in vivo or in vitro. In embodiments, when biodegradable microparticles are placed in a host in vivo or contacted with a biological scaffold seeded with cell culture or cells in vitro, the biodegradable microparticles begin to degrade and PE is degraded. Release to surrounding cells, tissues, etc. over time. In embodiments, the biodegradable microparticles release the first “burst” of PE and then slowly release the PE over a sustained period of time (eg, days, weeks, etc.). The release profile of the microparticles can be controlled by changing the size of the microparticles and the degree of crosslinking. In embodiments, the microparticles are cross-linked and the degree of cross-linking can be controlled to modify the particle's release parameters.

  In an embodiment, the biodegradable microparticles are made of gelatin. Further details regarding embodiments of gelatin biodegradable microparticles are described in Example 2 below. In an embodiment, the biodegradable microparticle comprises a mixture of microparticles of different sizes. It is thought that sustained release of PE can be obtained by including microparticles of various sizes. This is because particles of different sizes release PE at different rates.

  In an embodiment, the biodegradable microparticles are poly (lactic acid-co-glycolic acid) (PLGA) microparticles. In embodiments, hPE is added (eg encapsulated) in PLGA microparticles. In embodiments, PLGA microparticles loaded with hPE of the present disclosure have an average particle size of about 10 to about 1000 μm. In an embodiment, PLGA microparticles are made by an oil-in-water emulsion process that includes a double homogenization step. In an embodiment, PLGA microparticles are prepared by preparing a first emulsion by mixing a first aqueous solution (W1) comprising hPE of the present disclosure with a PLGA oily solution and homogenizing PLGA and W1 in a first homogenization step. Adding the first emulsion to a second aqueous solution (W2) containing a solvent (eg, but not limited to, for example, but not limited to, polyvinyl alcohol) in water to form a second emulsion (water-in-oil-in-water emulsion); Made by evaporating the solvent. In an embodiment, a second homogenization step is used to homogenize the second emulsion. In embodiments, the first homogenization is from about 1 to about 2 minutes. In some embodiments, no other homogenization step is used. In other embodiments, a second homogenization step is added. In embodiments, the second homogenization is from about 20 seconds to about 1 minute. Further details regarding embodiments of methods for preparing hPE-added PLGA microparticles are provided in Example 3 below. In embodiments, hPE-doped PLGA microparticles made using a single homogenization step have a size of about 100 to about 1000 μm in diameter. In embodiments where the single homogenization step is about 2 minutes, the resulting microparticles have a size in the range of about 50 to about 500 μm in diameter with an average diameter of about 260 to about 290 μm. In embodiments, hPE-added PLGA microparticles made using a second homogenization step of about 1 minute range in size from less than about 20 to about 100 μm in diameter and have an average diameter of about 36 to about 40 μm. In embodiments, hPE-added PLGA microparticles made using a second homogenization step of about 20 seconds range in size from less than 20 to about 200 μm and have an average diameter of about 85 to about 95 μm.

  The disclosure also includes a method of using the sustained release angiogenesis modulating composition described above for the sustained release of PE. In embodiments, the method uses sustained release compositions to provide sustained release of human PE in vivo or in vitro, such as, but not limited to, biomaterials (eg, cell cultures) in vivo or in vitro. Obtaining sustained release of human PE over time into tissues, tissue constructs, biological scaffolds, etc.). In embodiments, the methods of the present disclosure provide cells (eg, cells in culture, cells in vivo, cells in biomaterials or cells in contact with engineered biomaterials) into cells. The above sustained release vessel comprising biodegradable microparticles combined with the placental extract of the present disclosure such that human placenta extract is released from the microparticles into the biomaterial over a period of time after exposure to the microparticles Contacting with a neogenesis modulating composition. In embodiments, the cell is an endothelial cell such as, but not limited to, human umbilical vein endothelial cells (HUVEC).

  In embodiments, the sustained release angiogenesis modulating composition is associated with a biomaterial. In an embodiment, the biomaterial is a cell culture. In embodiments, the biomaterial is a tissue construct and / or tissue matrix that includes a cell culture. In an embodiment, the biomaterial is an alginate matrix comprising cells (eg, human cells, eg, HUVEC) embedded in the alginate matrix, and a sustained release angiogenesis modulating composition (eg, hPE-added biodegradable microdegradable). Particles) are also embedded in or in contact with the alginate matrix. In embodiments, a biomaterial associated with a sustained release angiogenesis modulating composition is implanted into a subject to induce angiogenesis into the biomaterial. In embodiments, the biomaterial is an engineered bioscaffold including, but not limited to, a human-derived substrate material, such as a decellularized human umbilical vein scaffold, seeded with human endothelial cells. is there. In embodiments, the subject is a mammal. In embodiments, the subject is a human. Other variations of the method of inducing angiogenesis using the sustained release angiogenesis modulating compositions of the present disclosure are possible and exemplary embodiments of the method are described in more detail in the examples below.

(Anti-inflammatory composition and method of use)
As described in the examples below, it was also found that the human PE of the present disclosure and the sustained release / human PE composition of the present disclosure have an anti-inflammatory effect on surrounding cells and tissues. Thus, the compositions of the present disclosure also include anti-inflammatory compositions comprising the human PE or sustained release / human PE compositions of the present disclosure. The methods of the present disclosure are methods (eg, reduce, alleviate, offset) of treating inflammation of a subject or subject tissue by exposing the subject or tissue to a human PE or sustained release human / PE composition of the present disclosure. And prevent).

  Further details regarding the tests and methods of the present disclosure are provided in the examples below. The specific examples below are to be construed as illustrative only and in no way limit the remaining disclosure. Based on the description herein, one of ordinary skill in the art would be able to utilize the present disclosure to its fullest extent, without further elaboration. All publications cited herein are hereby incorporated by reference in their entirety.

  The embodiments of the present disclosure, particularly any “preferred” embodiments, are merely possible examples of implementations set forth only for a clear understanding of the principles of the present disclosure. Many alterations and modifications may be made to the above-described embodiments of the present disclosure without substantially departing from the spirit and principles of the present disclosure. All such modifications and variations are intended to be included herein within the scope of this disclosure and protected by the following embodiments.

  The following examples are provided to provide those skilled in the art with a complete disclosure and description of how to perform the methods disclosed herein and how to use the compositions and compounds. Efforts have been made to ensure accuracy with respect to numbers (eg, amounts, temperature, etc.) but some errors and deviations are possible. Unless indicated otherwise, parts are parts by weight, temperature is in degrees Centigrade, and pressure is near atmospheric. Standard temperature and pressure are defined as 20 ° C. and 1 atmosphere.

  Note that ratios, concentrations, amounts and other numerical data may be expressed herein in a range format. Such a range format is used for simplicity and brevity, so whether each number and subrange is explicitly stated, as well as the number explicitly stated as a range limit. Thus, it should be understood that it should be construed flexibly in a manner that includes all individual numerical values or subranges within that range. For example, a concentration range of “about 0.1% to about 5%” includes not only explicitly stated concentrations of about 0.1 wt% to about 5 wt%, but also individual concentrations within the indicated range (eg, 1% 2%, 3% and 4%) and subranges (eg 0.5%, 1.1%, 2.2%, 3.3% and 4.4%) should also be construed is there. In certain embodiments, the term “about” may include traditional rounding with numeric significant digits.

  While embodiments of the present disclosure have been generally described so far, the following examples illustrate some further embodiments of the present disclosure. Embodiments of the present disclosure are described in the context of the following examples and corresponding text and figures, which are not intended to limit the embodiments of the present disclosure to this description. On the contrary, it is intended to cover all alternatives, modifications and equivalents included within the spirit and scope of the embodiments of the present disclosure.

Example 1
Induction and regulation of angiogenesis in ex vivo derived biological scaffolds using placenta-derived extracts This example describes a method of inducing angiogenesis using complex human placenta extracts (PE). PE is derived from human placenta and can induce angiogenesis in 2D and 3D in vitro models and in vivo within bioengineered tissue implants. This example also describes the positive screening of thrombospondin-1 as an anti-angiogenic protein using placenta extract with increased sensitivity compared to current in vitro models. In particular, this model makes it possible to adjust for the rate and type of angiogenesis (invagination vs. budding), and shows many advantages over conventional approaches and broad application in the fields of regenerative medicine and medicine.

Limiting mass transfer within tissues is a challenge in producing effective biomaterials. Even if this challenge can be temporarily overcome to improve cell migration within the human bioscaffold, the creation of effective vasculature will deliver nutrients over time to thick, densely packed materials Remains a fundamental goal. In adults, new blood vessels are created primarily by the physiological process of angiogenesis ⁴⁷ , which ultimately leads to the formation of nutrient-rich vascular networks. This example demonstrates that angiogenesis can be induced in human umbilical vein (HUV) vascular grafts and that angiogenesis provides a long-term nutrient delivery system.

  Successful angiogenesis in engineered organs and in vivo repair of infarcted tissue with angiogenesis regulators are major challenges in bringing regenerative medicine treatment to the clinic. A variety of different approaches have been taken to initiate angiogenesis and drive larger angiogenesis, including direct cell seeding (alone and co-culture), stem cells and human-derived modulator / growth factor combinations . To date, these in vitro approaches that typically use non-human animal compounds have hardly been brought to clinical practice.

  An important issue in this area is that the most popular and successful approach (Matrigel or basement membrane matrix) is derived from Angel Breath-Home-Swan mouse sarcoma cells and is therefore unsuitable for human therapy It is. Thus, approaches or mechanisms using human-based materials (those that actively promote angiogenesis in both in vitro and in vivo systems) are considered to have significant impact. This example provides human placenta extract (hPE) that can induce angiogenesis in 2D and 3D in vitro models and in vivo in engineered tissue implants. PE is a complex of active human biomolecules, and this example shows that in addition to inducing in vivo and in vitro angiogenesis in ex vivo derived human umbilical vein vascular grafts, this model Prove that you can adjust the stage. This example also demonstrates that PE enhances capillary formation and reduces fibrosis using a collagen-based bioscaffold with added ingredients.

Method Derivation of placenta extract. Full-term placenta was collected within 12 hours of delivery from UF Health Hands Hospital (Gainsville, FL). The umbilical cord and fetal membrane were removed and the placenta was dissected into a 2 cm cube and frozen. After 12 hours of progressive freezing to -86 ° C at a rate of -1 ° C / min, the placenta cubes were transferred to a cold room maintained at 4 ° C where the remaining procedures were completed. Once at 4 ° C., 100 grams of tissue was mixed with 150 mL of 3.4 M cold NaCl buffer (198.5 g NaCl, 12.5 ml 2 M Tris, 1.5 g EDTA and 0.25 g NEM in 1 L distilled water). . The NaCl buffer / tissue mix was homogenized to paste using a Tissuetek homogenizer at 3200 RPM, then centrifuged at 7000 RPM for 15 minutes and separated from the supernatant. This NaCl wash process was repeated two more times, with each supernatant being discarded and blood removed.

  The pellet is then homogenized in 100 mL of 4 M urea buffer (240 g urea, 6 g Tris base and 9 g NaCl in 1 L distilled water), stirred on a magnetic stir plate for 24 hours and then centrifuged at 14000 RPM for 20 minutes. (Sorvall RC6 + centrifuge, Thermo Scientific, NC, USA). The supernatant was removed and an 8000 MW dialysis tube (Spectrum Laboratories, Inc., CA, USA) in 1 L TBS (6 g Tris base and 9 g NaCl in 1 L distilled water) and 2.5 ml chloroform for sterilization. Dialyzed with The buffer was exchanged four more times with fresh TBS, each 2 hours apart. Finally, the contents of the dialysis tube are centrifuged at 3000 RPM for 15 minutes (Allegra X-12R centrifuge, Beckman Coulter, Inc., CA, USA) to remove polymerized protein and the supernatant (pink viscosity) The solution was collected and stored at -86 ° C until use.

  Biomolecular composition analysis. Relative cytokine levels were measured using RayBiotech, Inc. From a sandwich immunoassay array (human cytokine antibody array C series 1000, Inc, GA, USA). Chemoluminescence was detected using a Photo / Analyst Luminaryfx workstation (Fotodyne Incorporated, WI, USA) and signal intensity was measured using TotalLab 100 software (Nonlinear Dynamics, Ltd, UK). Relative abundance of basement membrane biomolecules was determined using MS Bioworks (Nano LC / MS / MS using a Waters NanoAcquisition HPEC (Waters, Milford, Mass.) System coupled with Orbitrap Velos Pro (ThermoFisher, Waltham, Mass.). Arbor, MI). Proteins were identified from the primary sequence database using the Mascot database search engine (Boston, Mass.).

RT-PCR analysis of cells from hPL-induced angiogenesis network. Relative angiogenic gene expression was determined using a 384 well RT ² human angiogenic RT ² Profiler PCR array (PAHS-024A, Qiagen, CA, USA). ECs were detached from the culture plate using Accutase (Innovative Cell Technologies, San Diego, Calif.) And stored immediately in 100 μl RNAlater. RNA was extracted using the RNeasy mini kit (Qiagen, CA, USA), and genomic DNA was digested using the RNase-free DNase kit (Qiagen, CA, USA). Purified RNA was reverse transcribed into cDNA using RT ² first strand kit (SA Biosciences, TX, USA) with incubation at 42 ° C for 15 minutes and then the reaction was stopped by incubation at 95 ° C for 5 minutes. . The cDNA was then mixed with RT ² SYBR green master mix (SA Biosciences, TX, USA) and loaded onto a 384 well human angiogenic PCR array. Using the Bio Rad CFX384 real-time system (Bio-Rad, CA, USA), the loaded array plate was subjected to a 10 minute denaturation cycle at 95 ° C and an annealing / extension cycle of 30 seconds at 40 ° C for 60 minutes Finally, a melting curve was obtained with a temperature increase from 60 ° C. to 95 ° C. at a rate of 0 ° C. per second. Data were analyzed using the ΔΔCt method and the RT ² Profiler PCR array data analysis template v4.0 software package (Quiagen, CA, USA).

  Human umbilical vein endothelial cell isolation and myofibroblast culture. Endothelial cells are detached from the vessel wall using a 1 mg / ml solution of bovine type I collagenase in phosphate buffered saline (Gibco, Invitrogen, NY, USA), thereby allowing human umbilical vein (UF Health Hands Hospital, (Collected from Gainsville, FL). In all experiments, the first derived human umbilical vein endothelial cells (HUVEC) were used between passages 1-3. For growth, cells were cultured using complete VasukuLife basal medium (VascuLife VEGF Medium Complete Kit, Lifeline, MD, USA). For angiogenesis experiments, VascuLife basal medium was mixed with 500 mL (VascueLife VEGF Medium Complete Kit, Lifeline with 25 ml glutamine, 0.5 ml hydrocortisone, 0.5 ml ascorbic acid, 10 ml FBS and 1.25 μl bFGF. , MD, USA), an endothelial cell medium was prepared. Human myofibroblasts (CRL 2854) were used between passages 5-10 (ATCC, Manassas, VA) and cultured with low glucose DMEM supplemented with 10% FBS.

Preparation of placental extract-derived angiogenesis assay. Unless otherwise stated, 32 μl of placental extract was thawed and pipetted into each well of a 96 well plate. The bottom of each well was uniformly coated with the extract using an orbital shaker at 30 RPM for 1 minute. The coated plate was then incubated for 30 minutes at 37 ° C. The HUVECs were then planted by pipetting directly at 20,000 cells / cm ² , 40,000 cells / cm ² or 80,000 cells / cm ² . Each concentration was investigated at multiple time points including Day 1, Day 3, and Day 5. Thrombospondin-1 was tested as an angiogenesis inhibitor using final concentrations of 0, 5, 10, 20, and 35 μg / μL diluted in endothelial cell medium.

Morphological characterization of angiogenic networks. Network formation was analyzed after staining with calcein AM (Invitrogen-Life Technologies, NY, USA) at a concentration of 2 μg / mL with endothelial cell culture medium. Stained cells were incubated for 30 minutes at 37 ° C., 5% CO ₂ in the dark, and then images were taken using a Zeiss Axiovert 200 inverted fluorescence microscope (Zeiss, Thornwood, NY). Images were analyzed and tubule length, tubule width, branch point and other network features were determined using ImageJ 1.45s (NIH, Bethesda, MD). The branch points were manually assigned as the positions of all nodes where the branches merge or the tubules emerge, and the capillary length was evaluated by determining the curve length from the branch point to the connection branch point. Capillary width measurements were taken in three different zones per capillary, two zones 10 μm each from the start and end points and one zone in the middle of the curve length. The percent coverage area was determined by processing the image with a “mean” measurement after using the “binary >> convert to mask” of the imageJ function. In the TSP-1 experiment, the final value was normalized to the sample without component addition and calculated as a percentage of the “1” value relative to the total count of pixel values and was obtained as “% cover area”.

  For scanning electron microscopy analysis of morphology, microvascular networks grown on glass slides were fixed in 2.5% glutaraldehyde, washed with PBS, fixed with 1% osmium tetroxide solution, 25% Progressively dehydrated with 50%, 75%, 85%, 95% and 3 × 100% ethanol solutions. The sample was then critical point dried, coated with gold / palladium and imaged using a Hitachi S-4000 FE-SEM.

Derivation of human umbilical vein scaffold and incubation of placenta extract. Placenta were collected from UF Health Hands Hospital Florida (Gainsville, FL) and HUVs were dissected using the ³² automated method described previously. The dissected HUV sample was decellularized in 1% SDS (Thermo Scientific, Rockford, IL) solution in solvent / tissue mass 20: 1 (w: v). Samples were decellularized on an orbital shaker plate at 100 rpm for 24 hours and then rinsed with PBS, followed by 37 ° C. in 70 U / mL DNase I solution (Sigma-Aldrich, St. Louis, MO) in PBS. Incubate overnight at Samples were terminally sterilized with 0.2% peracetic acid / 4% ethanol (Sigma-Aldrich, St. Louis, MO) solution for 2 hours and finally pH balanced with PBS (7.4). ). After decellularization, the scaffold was cut into 1.5 cm × 1.5 cm × 0.075 cm sheets, pre-frozen to −85 ° C., and then milllock tabletop at −85 ° C. under 10 mT vacuum for 24 hours. Lyophilized using a manifold lyophilizer (Kingston, NY). Immediately prior to cell seeding, the scaffolds were soaked in hPE, Matrigel or PBS (control) for 2 hours and seeded.

  Animal implant revascularization study. Male Sprague-Dawley rats (6 months old, 200 g) were purchased from Charles River Laboratories (Wilmington, Mass., USA) and all procedures were approved by the University of Florida IACUC (UF # 201207728). The terminally sterilized HUV scaffold was incubated in a biological hood for 2 hours in 5 mL hPE, MATRIGEL or PBS (control), respectively. The animals were anesthetized using isoflurane inhalation and subcutaneous pockets were created on the left and right side of the back by blunt preparation using scissors. One scaffold was inserted into each subcutaneous pocket and the skin was sutured using 4-0 sutures (Coviden, Mansfield, MA). Five days after transplantation, animals were euthanized and samples were removed for analysis.

  To analyze capillary network formation, immediately after removal from the animal, the fibrotic capsule was dissected with a scalpel and the HUV sample was placed on a glass slide. The entire image of the translucent scaffold sheet was taken using an Imager M2 optical microscope (Zeiss, Oberkochen, Germany) equipped with an Axiocam HRm digital camera (Zeiss, Oberkochen, Germany). To quantify cell migration and scaffold modeling, tissue samples were embedded in Neg-50 cryosection media and cut into 7 μm sections (Microm HM550 cryostat, Thermo Scientific, Waltham, Mass.), Standard Hematoxylin and eosin (H & E) staining (Richard-Alan Scientific, Kalamazoo, MI).

statistics. Results are reported as mean ± standard deviation. Linear regression was performed using SPSS (IBM, Somers, NY). Rt-PCR data was analyzed using RT ² Profiler PCR array data analysis software v3.2 (SABiosciences, Valencia, CA).

Angiogenesis induction in perfusion bioreactor cultures and HUV bioscaffolds. Cell-seeded tubular constructs were cultured for 5 days in a double perfusion bioreactor (FIG. 7) using a lumen flow rate of 4 mL / min at 60 pulses / min. Shear stress on the vessel wall was calculated using the Haagen-Poisseille equation under the assumption that the medium flow was uniform and laminar, the vessel was inelastic, cylindrical, and linear: ¹³⁴
(Where Q is the average volume flow rate and μ is the kinematic viscosity of water at 37 ° C. (equal to 0.000692 kg / (m ^* s)) ^134. Shear stress is 0 dynes / cm ² during each pulse From 0.02 dynes / cm ^{2 The} environment was maintained under standard cell culture conditions of 37 ° C. and 5% CO _2. Intrasystem pressures flowed both on the abluminal side and on the luminal side A negligible level (<2 mmHg) was maintained in the circuit so that there was no pressure gradient across the scaffold.The culture medium in the bioreactor was replenished every 2 days. A 10 cm long tubular scaffold was dissected into the annulus for analysis.

Results Derivation and Characterization of Human Placenta Extract After initial mechanical homogenization and centrifugation, the hPE derivation method utilized a urea step to linearize and solubilize the molecules. This was followed by dialysis separation to remove urea and refold the biomolecule to its original conformation (FIG. 1A). All steps in this derivation were performed in a cold room at 4 ° C. The final solution of PE was clear and highly viscous and consisted of biomolecules between 8 kD and 868 kD.

hPE could be used to soak biomaterials or could be a thin film for tissue culture assays. The reproducibility of hPE was assessed by analysis of standard deviations of total protein content in n 1/4 3 batches of hPE (each batch created with equal mass of tissue from 3 separate donors). , Have been shown to have similar reproducibility and protein content as Matrigel®. Scanning electron microscopy images show hPE surface morphology (FIGS. 1B-1C) and angiogenic network formation when HUVECs are seeded on hPE thin films at 4 × 10 ⁴ cells / cm ^{2 and} cultured for 3 days (FIG. 1D-1E).

  The angiogenic ability of human placenta extract (hPE) was first characterized by seeding primary human umbilical vein endothelial cells (HUVEC) on tissue culture plates (TCP) coated with hPE. Early stage cell streaking and budding were visible within 1 hour of cell seeding (data not shown) and the angiogenic network continued to mature until the end of the 3d experiment (FIGS. 1F-1G). The length of individual cell cords (multicellular) increased significantly from seeding day 1 (FIG. 1H) to day 3 (FIG. 1I). After 3 days in culture, the cells formed an extensive angiogenic network compared to the control sample (FIGS. 1J, 1K).

Biomolecular characterization of placental extract Of the 120 cytokines evaluated, 54 angiogenesis-related cytokines were detected in placental lysate (FIG. 2A). The most common angiogenesis-related chemokine was angiogenin, which is a potent stimulator of neovascularization ¹⁶ . There are also significant pro-angiogenic chemokines including, but not limited to, hepatocyte growth factor (HGF), fibroblast growth factor-4 (FGF4), leptin (LEP), ICAM-1, ICAM-2 and TIMP-2. was detected. LC-MS / MS confirms the presence of immune related proteins including annexins (ANXA1, ANXA2, ANXA4 and ANXA5), neutrophil defensins (DEFA1), interleukin enhancer binding factors (ILF2 and ILF3), IL27, ITBG1 and MRC1. Shown (FIG. 2B). Also angiogenesis-related basement membrane (BM) proteins including laminin (LAMA2, LAMA4, LAMA5, LAMB1, LAMB2, LAMB3 and LAMC1), fibronectin (FN1), heparin sulfate (HSPG2) and type 4 collagen (COL4A1, COL4A2 and COL4A3) Detected using LC-MS / MS (Figure 2C), each of which has been shown to play a key role in angiogenesis ^17-20 .

Endothelial cell gene expression within an angiogenic network induced by hPE Along with chemokine analysis, HUVEC gene analysis further supported the angiogenic nature of placental extracts. RT-PCR analysis showed that endothelial cells seeded 3d on hPE expressed a wide range of essential pro-angiogenic genes including hepatocyte growth factor, epidermal growth factor and placental growth factor (FIG. 2D). . Additional up-regulated genes include MMP2 and MMP9, which are proteolytic enzymes that help degrade the surrounding extracellular matrix to promote migration of endothelial cells and other cells associated with ECM remodeling. There is ²¹ . Type IV collagen was also upregulated, which is associated with the formation of basement membranes in the mature microvasculature ²² .

Modulation of capillary development and morphology in vitro Historically, in vitro assays have little or no control over the rate and stage of angiogenesis ⁹ . The present data show that in vitro hPE-based angiogenesis assays can be adjusted to control the maturation and morphology of angiogenic network formation by varying the initial cell seeding density. One day later, HUVEC seeded at a density of 40,000 cells / cm ² formed clearer tubules compared to seeding at a density of 80,000 cells / cm ² , but by day 5, Cells seeded at both densities had well defined tubules (FIG. 3A). These results show that by varying the cell seeding density, the maturation stage of network formation can be controlled when the culture is exposed to hPE. For example, quantitative image analysis of capillary network (average capillary length [mm], capillary density [# / mm ² ], branch point [# / mm ² ], mesh [# / mm ² ] and average capillary width [mm]) (FIG. 3C) results in a higher seeding density resulting in slower network maturation (maximum mean tubule length and time frame to reach number of meshes / mm ² ) and extended network formation time frame So it showed that more detailed analysis is possible. In comparison, lower cell densities result in faster maturation rates, which are more conductive to rapid screening approaches to test the effectiveness of anti-angiogenic agents for cancer treatment (for example) It is thought that there is.

  As a historical representative of the in vitro angiogenesis assay, the angiogenesis network induced by Matrigel was compared to the network induced by hPE (FIG. 3D). The morphology of the endothelial cell capillary network was first analyzed by exposing the cell culture to either Matrigel or hPE using calcein AM to determine viability and network structure. On day 1 after seeding, the Matrigel-coated plate showed that HUVEC formed a well-defined angiogenic tubule network, but after 3d the network structure collapsed into a spherical ball of apoptotic cells (Figure 3A, 3D). Some cell death was confirmed in the network induced by hPE, but no apoptotic ball formation was observed after an extended 5d period.

  During the late stages of angiogenesis, EC recruits smooth muscle cells (SMC) to stabilize blood vessels as the capillary network matures. Therefore, the effect of hPE and Matrigel on smooth muscle cell morphology was evaluated. Interestingly, in the absence of HUVEC, SMC seeded on Matrigel formed tubules after 1d (FIG. 3B.i), but on the hPE-coated plate, the typical “hills and valleys” Still formed (FIG. 3B.ii), indicating that there are significant differences in molecular signaling pathways between cell types. Since SMC is not known to form tubules in the early stages of microangiogenesis, these results may indicate that hPE-based angiogenesis is more accurate and corresponds to normal physiology.

Analysis of hPE-induced angiogenesis for drug screening applications In addition to its role in regenerative medicine, angiogenesis driven by human placenta extract (hPE) is used to screen angiogenesis-related drugs in vitro. Tested for ability. Matrigel was used as a control. Matrigel and hPE-based angiogenesis assays were screened against thrombospondin-1 (TSP-1), a glycoprotein with effective inhibitory activity against neovascularization. TSP-1 is a model drug for anti-angiogenic treatment of solid tumors ²³ . Control HUVECs seeded directly on tissue culture plates after 1d in culture were not affected by TSP-1 and the cells formed a typical cobblestone morphology. In contrast, HUVECs cultured on hPE-treated culture plates significantly reduced angiogenic network formation (FIG. 4A). The results show that total tubule length and branching point decrease linearly as a function of TSP-1 concentration (FIG. 4B). Importantly, these studies show that hPE-based assays are more sensitive to drug concentration compared to Matrigel-based assays. This is shown by a higher correlation between TSP-1 concentration and percent reduction in angiogenic network coverage, with R ² values of 0.97 and 0.36, respectively (FIG. 4C). This inhibition of network formation further confirms that actual angiogenesis is occurring rather than an unknown stress response. Furthermore, since hPE is derived from humans, this avoids inaccuracies based on species differences that can be brought about by screening using non-human systems ^13,14 .

In Vitro Angiogenic Network Formation within 3D Biological Scaffolds Successful angiogenesis to engineered organs is a major challenge in successfully developing regenerative therapy ²⁴ , ex vivo The ability of hPE to induce angiogenesis in the derived biological scaffold was analyzed. These studies show that hPE induced the formation of angiogenic networks in engineered (decellularized) human umbilical vein (HUV) biological scaffolds (FIG. 5A). Consistent with the assay in 2D culture plates, endothelial cells (EC) seeded on biological scaffolds developed into long elongated forms connected to multicellular cords forming a complex interconnected network.

In vivo angiogenesis occurs by a variety of mechanisms, most commonly budding or invagination. These data show that by varying cell density when incubated with hPE, budding versus invagination of angiogenesis can be regulated in vitro. At lower cell densities (2 × 10 ⁴ cells / cm ² ), network morphology on scaffolds incubated with hPE showed budding neovascularization (FIGS. 5C.i, 5C.ii), with intermediate densities (4 × At 10 ⁴ cells / cm ² ), the network morphology shows a combination of budding and invagination angiogenesis (FIGS. 5C.iii, 5C.iv), and at higher densities (6 × 10 ⁴ cells / cm ² ) The network morphology was more approximated by invaginated angiogenesis (FIGS. 5C.v, 5C.vi). Since budding generally occurs at an early stage of angiogenesis in low cell density regions where there are no capillaries, the correlation between cell density and a specific mechanism of angiogenesis is currently the case of in vivo capillaries and network formation. Support the understanding of In contrast, invagination occurs in areas of higher cell density where capillaries and endothelial cells already exist ^25,26 .

In vivo induction of angiogenesis An angiogenic response to component-added scaffolds (Matrigel and hPE) was evaluated 5 days after transplantation using a subcutaneous rat model (Figure 6A). Both the control and scaffolds incubated with Matrigel showed significant fibrosis around the scaffold, whereas the scaffold supplemented with hPE showed no discernable fibrosis around the implant (FIG. 6B. i-6B.iii.). Prevention of fibrosis in hPE samples is not limited to them, as detected by LC-MS / MS, but includes anti-inflammatory annexins (ANXA1, ANXA2, ANXA4 and ANXA5) ²⁷ , antimicrobial defensin peptides such as DEFA1 ²⁸ , As well as immune-related molecules including MRC1, which are known to bind to pathogens that may include viruses and bacteria.

  As shown by bright field microscopy, both the scaffolds incubated with Matrigel and hPE showed a significant increase in neovascularization compared to controls (FIGS. 6B.iv-6B.vi.). Although total angiogenesis appears to be comparable, hPE treatment shows the formation of a mature capillary bed, while Matrigel samples seemed to be less organized with no evidence of mature capillary bed formation . H & E stained sections migrated into and throughout the scaffolds in which cells were incubated in hPE, but in samples incubated with Matrigel, cell infiltration was reduced and cells in control samples were confined to the periphery of the scaffold (FIG. 6B.vii.-6B.ix.) The improvement in cell migration in both Matrigel and hPE incubated samples was the result of chemotaxis and growth factor signals adsorbed on the scaffold structure. Although both scaffolds supplemented with hPE and Matrigel improved cell migration relative to controls, cell remodeling between these sample groups showed variation. The dense cell area in the sample incubated with Matrigel shows the rest of the original HUV fiber, and its general structure appears to be almost completely remodeled with the scaffold incubated with hPE. Qualitatively more amorphous compared to the demonstrated fiber and cell structure (FIGS. 6B.vii.-6B.ix.).

Discussion Tissue regeneration, infarcted tissue and ischemic wound repair are three clinical areas where improved strategies for wound healing or organ replacement have significant clinical impact. The use of amniotic membranes and chorion in various applications has grown significantly over the past five years, and there is increasing evidence indicating that perinatal tissues have considerable clinical promise ^29-32 .

  The results herein describe in detail a novel approach to concentrate and deliver the physiological proportions of potent human-derived stimulants of angiogenesis and tissue remodeling. These data indicate that the onset of capillary formation (in vitro and in vivo), control of the EC phenotype during angiogenesis with the ability to modulate growth or maturation kinetics, and a significant reduction in in vivo tissue fibrosis Shows enhanced cellular activity towards

  The ability to regulate the in vitro maturation rate of capillary network formation and control the emergence of budding and invading angiogenic network forms provides a useful platform for further understanding of regulatory pathways during wound healing and organ regeneration Conceivable. Based on a comparison with Matrigel, the mechanism by which hPE stimulates cells appears to be fundamentally different. SMCs incubated with Matrigel initiated capillary-like formation, whereas SMCs exposed to hPE retain their typical hill and valley morphology, thus human-derived hPE is in more complex models A more representative model of physiological angiogenesis can be provided.

Some current models are based on human-derived (recombinant) modulators that rely on single or separate combinations of angiogenesis regulators ³³ . Separate combinations are useful to control variability and reduce the inherent complexity of miscellaneous approaches, but they limit the screening process and limit the ability of anti-angiogenic, tumor suppressor drugs. It does not represent a series of broad human in vivo molecular interactions that would be important when testing.

  The inherent complexity of hPE-based models can lead to advances in the pharmaceutical industry by providing a more effective screening approach for tumor suppressor drugs. Compared to current technology, exposing human EC to hPE has been shown to have increased sensitivity to angiogenesis-inhibiting drug concentration (TSP-1) and lower detection limits.

hPE-based models induce angiogenesis using a broad set of human-derived molecules in approximately physiological proportions. Modulating only selected molecular pathways would limit attempts to discover new anti-angiogenic drugs. This is because in vivo angiogenesis requires the induction of multiple metabolic pathways ^34,35 . Thus, even though a drug can regulate angiogenesis by interacting with any of these multiple pathways, it has little effect on inducing sufficient angiogenesis when the complexity of the local environment is lacking. It is thought not to. The results from in vivo analysis in this example provide further evidence that a complex PE affects a number of biochemical pathways, resulting in a wide range of effects. The data showed that hPE not only showed enhanced angiogenic properties, but also showed immune-reducing properties, as indicated by reduced fibrosis within the bioscaffold supplemented with hPE. . Complex Interactions in view by ^36, the molecular composition of hPE between angiogenesis and immunological molecule pathway for inducing capillary formation without significant immunological and inflammatory responses, clinical To provide an appropriate basis for the development of available technologies.

The positive results of the above studies are related not only to the presence of key growth factors (GF) and gene regulators in PE, but also to their presence in physiological proportions. It can be seen. For example, VEGF was upregulated in ECs induced by hPE, but the hPE solution did not contain detectable levels. This is in contrast to Matrigel (BMM), which contains an active concentration of VEGF in both standard and GFR (Growth Factor Reduced) versions. From the results herein showing more mature capillary bed formation, the presence of VEGF is only one of many contributing factors, and the presence of other regulatory factors plays a key role in blood vessel development. The possibility of performing is high. Similarly, when compared to human recombinant proteins used to initiate angiogenesis, these depend on a highly enriched (typically) single GF ^37-39 . Problems have been reported that the clinical use of single, highly reactive GF has had undesirable effects ^40-42 .

  The hPE angiogenesis model has been validated in vivo within 2D and 3D in vitro models as well as bioengineered tissue implants and can be readily adapted for various clinical or pharmaceutical applications. Together with its angiogenic and immunoreducing properties, it is derived from a physiologically healthy human vascular bed, making it unique among current angiogenesis models. The data presented here showed that hPE plays a critical role in several key clinical issues where the demand for a more successful alternative approach has become a clinical priority.

Reference for Example 1
1.Thurston, G., Murphy, TJ, Baluk, P., Lindsey, JR & McDonald, DM Angiogenesis in mice with chronic airway inflammation: strain-dependent differences. Am J Pathol 153, 1099-1112 (1998).
2. Cristofanilli, M., Charnsangavej, C. & Hortobagyi, GN Angiogenesis modulation in cancer research: novel clinical approaches. Nature reviews. Drug discovery 1, 415-426 (2002).
3. Lokmic, Z. & Mitchell, GM Engineering the microcirculation.Tissue Eng Part B Rev 14, 87-103 (2008).
4. Kurz, H., Burri, PH & Djonov, VG Angiogenesis and vascular remodeling by intussusception: from form to function.News in physiological sciences: an international journal of physiology produced jointly by the International Union of Physiological Sciences and the American Physiological Society 18, 65-70 (2003).
5. Burri, PH & Djonov, V. Intussusceptive angiogenesis--the alternative to capillary sprouting.Molecular aspects of medicine 23, S1-27 (2002).
6. Risau, W. Mechanisms of angiogenesis.Nature 386, 671-674 (1997).
7. Folkman, J. Angiogenesis in cancer, vascular, rheumatoid and other disease.Nature medicine 1, 27-31 (1995).
8. Jain, RK, Schlenger, K., Hockel, M. & Yuan, F. Quantitative angiogenesis assays: progress and problems.Nature medicine 3, 1203-1208 (1997).
9. Auerbach, R., Akhtar, N., Lewis, RL & Shinners, BL Angiogenesis assays: problems and pitfalls. Cancer metastasis reviews 19, 167-172 (2000).
10. Auerbach, R., Lewis, R., Shinners, B., Kubai, L. & Akhtar, N. Angiogenesis assays: a critical overview.Clinical chemistry 49, 32-40 (2003).
11.Kleinman, HK & Martin, GR Matrigel: basement membrane matrix with biological activity.Seminars in cancer biology 15, 378-386 (2005).
12. Cockerill, GW, Gamble, JR & Vadas, MA Angiogenesis: models and modulators.International review of cytology 159, 113-160 (1995).
13. Warren, MS et al. Comparative gene expression profiles of ABC transporters in brain microvessel endothelial cells and brain in five species including human.Pharmacological Research 59, 404-413 (2009).
14. Febbraio, M., Hajjar, DP & Silverstein, RL CD36: a class B scavenger receptor involved in angiogenesis, atherosclerosis, inflammation, and lipid metabolism.Journal of Clinical Investigation 108, 785-791 (2001).
15. Wang, Y. & Zhao, S. in Vascular Biology of the Placenta (San Rafael (CA); 2010).
16. Fett, JW et al. Isolation and characterization of angiogenin, an angiogenic protein from human carcinoma cells.Biochemistry 24, 5480-5486 (1985).
17. Xu, J. et al. Proteolytic exposure of a cryptic site within collagen type IV is required for angiogenesis and tumor growth in vivo.The Journal of cell biology 154, 1069-1080 (2001).
18. Kim, S., Bell, K., Mousa, SA & Varner, JA Regulation of Angiogenesis <i> in Vivo </ i> by Ligation of Integrin α5β1 with the Central Cell-Binding Domain of Fibronectin. The American journal of pathology 156, 1345-1362 (2000).
19. Iozzo, RV & San Antonio, JD Heparan sulfate proteoglycans: heavy hitters in the angiogenesis arena. Journal of Clinical Investigation 108, 349-355 (2001).
20. Patarroyo, M., Tryggvason, K. & Virtanen, I. in Seminars in cancer biology, Vol. 12 197-207 (Elsevier, 2002).
21. Rundhaug, JE Matrix metalloproteinases and angiogenesis. J Cell Mol Med 9, 267-285 (2005).
22. Kalluri, R. Basement membranes: structure, assembly and role in tumour angiogenesis.Nature reviews. Cancer 3, 422-433 (2003).
23. Lawler, J. Thrombospondin-1 as an inherent inhibitor of angiogenesis and tumor growth. J Cell Mol Med 6, 1-12 (2002).
24. Laschke, MW et al. Angiogenesis in tissue engineering: breathing life into constructed tissue substitutes.Tissue engineering 12, 2093-2104 (2006).
25. Adair, T. in Integrated systems physiology, from molecule to function to disease (Morgan & Claypool, 2011).
26. Djonov, V., Baum, O. & Burri, PH Vascular remodeling by intussusceptive angiogenesis. Cell and tissue research 314, 107-117 (2003).
27. Perretti, M. et al. Endogenous lipid-and peptide-derived anti-inflammatory pathways generated with glucocorticoid and aspirin treatment activate the lipoxin A4 receptor.Nature medicine 8, 1296-1302 (2002).
28. Paslakis, G. et al. The Putative Role of Human Peritoneal Adipocytes in the Fight against Bacteria: Synthesis of the Antimicrobial Active Peptide DEFA1-3. Nephron Experimental Nephrology 115, e96-e100 (2010).
29. Lee, S.-H. & Tseng, S. Amniotic membrane transplantation for persistent epithelial defects with ulceration. American journal of ophthalmology 123, 303-312 (1997).
30. Jin, CZ et al. Human amniotic membrane as a delivery matrix for articular cartilage repair.Tissue engineering 13, 693-702 (2007).
31. Bose, B. Burn wound dressing with human amniotic membrane. Annals of the Royal College of Surgeons of England 61, 444 (1979).
32. Daniel, J., Abe, K. & McFetridge, PS Development of the human umbilical vein scaffold for cardiovascular tissue engineering applications. ASAIO J 51, 252-261 (2005).
33. Vailhe, B., Vittet, D. & Feige, JJ In vitro models of vasculogenesis and angiogenesis. Laboratory investigation; a journal of technical methods and pathology 81, 439-452 (2001).
34. Pepper, M., Ferrara, N., Orci, L. & Montesano, R. Potent synergism between vascular endothelial growth factor and basic fibroblast growth factor in the induction of angiogenesis in vitro.Biochemical and biophysical research communications 189, 824- 831 (1992).
35. Sullivan, DC & Bicknell, R. New molecular pathways in angiogenesis.British journal of cancer 89, 228-231 (2003).
36. O'Byrne, KJ, Dalgleish, A., Browning, M., Steward, W. & Harris, A. The relationship between angiogenesis and the immune response in carcinogenesis and the progression of malignant disease. European journal of cancer 36, 151-169 (2000).
37. Montesano, R., Vassalli, J.-D., Baird, A., Guillemin, R. & Orci, L. Basic fibroblast growth factor induces angiogenesis in vitro.Proceedings of the National Academy of Sciences 83, 7297-7301 (1986).
38. Ferrara, N. & Alitalo, K. Clinical applications of angiogenic growth factors and their inhibitors.Nature medicine 5 (1999).
39. Zisch, AH, Lutolf, MP & Hubbell, JA Biopolymeric delivery matrices for angiogenic growth factors.Cardiovascular Pathology 12, 295-310 (2003).
40. Epstein, SE, Fuchs, S., Zhou, YF, Baffour, R. & Kornowski, R. Therapeutic interventions for enhancing collateral development by administration of growth factors: basic principles, early results and potential hazards.Cardiovascular Research 49, 532 -542 (2001).
41. Lee, RJ et al. VEGF gene delivery to myocardium deleterious effects of unregulated expression. Circulation 102, 898-901 (2000).
42.Hariawala, MD et al. VEGF improves myocardial blood flow but produces EDRF-mediated hypotension in porcine hearts. Journal of Surgical Research 63, 77-82 (1996).

(Example 2)
Introduction to controlled delivery of placental extracts for stimulation of angiogenesis Tissue engineering aims to build tissues and organs from scratch in vitro for transplantation into sick patients. However, this revolutionary alternative to transplantation involves the lack of the formation of appropriate vasculature to supply oxygen and nutrients to cells seeded in the graft. Thus, there is an urgent need for an effective method for inducing angiogenesis in tissue engineering constructs.

  To date, all methods that attempt to promote angiogenesis in engineered products have been unsuccessful as a result. The present disclosure provides a protocol for deriving pro-angiogenic extracts from human placenta that have been shown to induce and regulate early stages of angiogenesis, as described in the examples above.

  This example describes a 3D in vitro angiogenesis assay that promotes the formation of a capillary network and persists it over time. To this end, placental extracts were analyzed for their angiogenic potential and their biological activity over time. These studies have shown that frequent administration of the extract promotes the formation of a mature, long-lasting capillary network. Thus, biodegradable gelatin microparticles for the incorporation and controlled release of extracts were prepared and their degradation kinetics were studied. Finally, a 3D in vitro angiogenesis assay was developed. Placental extract-loaded microparticles were embedded in a type I collagen hydrogel scaffold seeded with HUVEC, demonstrating evidence of an early stage of microvascularization within the matrix.

Organ or tissue loss or failure is one of the most serious and expensive human health problems in modern society. Many approaches have been attempted to solve this problem, including surgical reconstruction, drug therapy, synthetic appliances, mechanical devices and implants. In the long term, these approaches usually fail. Because they do not completely replace the function of the lost organ or tissue ² , or in the case of transplantation, there is a great demand for organs and an insufficient number of donors.

One solution would be to develop an engineered organ or tissue. However, current tissue engineering approaches have limited success, in part due to nutrient barriers that arise across the thick biological scaffold. A major challenge for engineering thick (> 200 μm) tissue constructs is that cell growth and viability within 3D tissue constructs, as oxygen diffusion is typically limited to distances of 150-200 μm in tissue engineering scaffolds. Is to be impaired ^6,7 . Oxygen is a critical nutrient for cell survival, and without oxygen, cells in the scaffold die. Furthermore, poor transport conditions frequently result in the formation of a fibrotic capsule secreted by the cells. This capsule formation results in a lack of nutrient mass transfer within the scaffold, ultimately resulting in a low cell density. Thus, a method that facilitates delivery of oxygen and nutrients to cells seeded in a 3D tissue construct would be beneficial in engineering a tissue that can last for a long time. The placental extracts and methods of the present disclosure provide an approach to overcome oxygen and nutrient deficiencies, including inducing rapid development of nutrient-rich capillary systems within the scaffold. It is believed that performing the method to supply these essential nutrients not only to the edges of the construct but also to the center will help prevent the formation of fibrotic capsules.

  Angiogenesis is the center of tissue development and maintenance, and successful regulation promotes the controlled formation of established vascular networks in the transplanted graft. Several biological factors and molecular pathways are involved in the regulation of this complex process that is still partially unknown. In order to better understand the molecular mechanisms that initiate and control blood vessel growth, studies have determined in vitro or in vitro critical elements of angiogenesis under simple, well-defined and controlled conditions.

  The focus is on the development of an angiogenesis assay, a cell culture system that reproduces in vivo. A variety of different approaches have been used to promote in vitro angiogenesis, but to date they have hardly been transferred to clinical practice. The limitation of most angiogenesis models is that they are of animal origin (eg Matrigel base) or are completely dependent on the use of living animals (eg chick chorioallantoic membrane and rabbit corneal micropocket). Furthermore, they lack a variety of natural cytokines and chemical gradients in vivo. Consequently, the results of these assays are often disappointing due to the lack of long-lasting angiogenesis. Thus, a robust in vitro model of human origin would be useful for mechanistic studies and screening for angiogenic drugs for humans.

  In view of the fact that the placenta is a readily available tissue and is a rich source of vascular tissue and angiogenesis-related growth factors, in these studies angiogenesis is performed as described above and in Example 1. The placenta was used to derive a multiprotein human placenta extract or matrix (hPE or hPM), a viscous protein compound rich in promoting cytokines and angiogenesis-related growth factors. It has been shown that hPE can induce and regulate early stages of angiogenesis in vitro for a limited period of time. Furthermore, hPE has also been shown to significantly reduce fibrosis.

This example provides a method for sustained delivery of hPE such that growth factors within hPE remain functional long enough to allow the formation of a mature capillary network. For this purpose, we have developed the versatility of the microparticles, and the polypeptide efficiently encapsulated microparticles for their ability ²² to release their long-term continuous speed as a delivery system.

The microparticles are ²³ solid, nearly spherical particles with a large surface to volume ratio with a size in the range of 1-1000 μm. They are prepared from several different materials, both natural (eg starch, gum) and synthetic (eg polylactic acid and polyglycolic acid) using different methods such as hot melt extrusion, spray drying or solvent removal ²⁴ it can. The release rate of the microparticles can be adjusted by changing their size, and smaller particles dissolve faster than larger ones as the surface to volume ratio increases. For this reason, it is possible to adjust the delivery rate by combining particles of different sizes ²⁵ . Drug or protein release from microparticles usually occurs by simple matrix bioerosion. This process involves erosion of the particle surface, followed by bulk erosion and penetration of the release medium into the pores of the particle ^26,27 .

In this example, biodegradable gelatin microparticles were prepared and used for 3D in vitro angiogenesis assays. Angiogenesis assays, simple, clear and cell culture system for reproducing a determining factor of angiogenesis in vitro or in vivo under controlled conditions ^28. These can be two-dimensional or three-dimensional. In 3D in vitro assays, EC develops tubular structures both on the surface of the substrate and those that invade the ¹³ surrounding matrix, usually constructed with biogels.

  This example provides an embodiment of a 3D in vitro angiogenesis assay to promote the formation of a long-lasting vascular network within an implanted graft. In this embodiment, the assay is prepared by embedding human umbilical vein endothelial cells (HUVEC) with PE-added microparticles in a type I collagen matrix. The angiogenic response of HUVEC at different incubation times was then analyzed.

Experimental Method Endothelial cell isolation and culture. Pre-isolated from human umbilical vein and described by Jaffe et al. (Culture of Human Endothelial Cells Derived from Universal Veins. 52, 2745-2756; 1973, incorporated herein by reference for isolation and culture of HUVEC). The human umbilical vein endothelial cells (HUVEC) cultured in advance as described above were detached from the flask using acutase (Fisher Scientific) and centrifuged at 1000 rpm for 5 minutes. The cell pellet is resuspended in medium (25 ml glutamine, 0.5 ml hydrocortisone, 0.5 ml ascorbic acid, 10 ml FBS, 1.25 μL VEGF and 1.25 μL bFGF in 500 ml VascuLife basal medium). HUVECs were counted using a hemocytometer.

Analysis of hPM angiogenesis as a function of time. To evaluate how hPE incubation time affects HUVEC angiogenic response, cells were seeded in tissue culture plates coated with hPE and incubated with hPE for 1, 3 or 5 days. Briefly, a placental matrix was prepared using the isolation methods described in the above examples. The vial containing the extract was thawed and pipetted into a well of a 96 well plate (100 μL per cm ² ). By Pipet cell solution directly at 20,000 cells / cm ^2, the HUVEC were prepared for planting. Angiogenic medium was added to each sample prepared on the plate (200 μL per cm ² ) and the plate was placed in a humidified 5% CO ₂ incubator at 37 ° C.

To determine how hPE retains biological activity over time, hPE was stored for various times including 20, 15, 9, 7, 5, 3 and 1 days. (In an incubator with 5% CO ₂ and 37 ° C.). Tissue culture 96-well plates were then coated with hPE membrane from each time point and seeded with HUVEC as described above. Cells were cultured for 3 days to qualitatively characterize the angiogenic network.

  Analysis of hPE angiogenesis as a function of inoculation frequency. The response of HUVEC to the number of hPE inoculations was also analyzed. Initially, cells were seeded on day 1 on hPE membranes, and hPE was mixed with culture medium for the addition of hPE components after day 1. Cells were cultured for 5 days, some groups of samples received only one hPE inoculation on day 1 (seeding day), others inoculated twice on day 1 and day 3, The other group received 3 inoculations on day 1, day 3 and day 4.

Preparation of gelatin microparticles. Gelatin microparticles were prepared using Method ³³ described by Tabata et al. (Incorporated herein by reference for the preparation of gelatin microparticles). All reagents used were obtained from Fisher Scientific. Briefly, a 10% wt aqueous solution of type B gelatin was prepared by adding 1 g gelatin to 9 mL deionized water. The temperature was raised to 45 ° C. and the solution was added dropwise to 375 ml of warm (45 ° C.) olive oil through a syringe and 21-G needle under constant stirring at 400 rpm. After 10 minutes, the emulsion temperature was lowered to 15 ° C. and stirring was maintained for 30 minutes to induce gelation. 100 mL of cold acetone was added and the emulsion was stirred for 1 hour. Fine particles were removed by vacuum filtration, washed with acetone and dried. Once dried, they were placed in an aqueous solution containing 0.1% wt Tween 80 and 0.5% wt glutaraldehyde. The solution was continuously stirred for 15 hours at 125 rpm and 4 ° C. to facilitate cross-linking of the microparticles. Cross-linked microparticles were collected by vacuum filtration, washed with deionized water, and stirred in 100 mL of 10 mM glycine aqueous solution to block any unreacted glutaraldehyde. After 1 hour, the microparticles were collected again by filtration, washed with deionized water, and lyophilized. 100 μL of pure hPM per mg of microparticles was added to the cross-linked lyophilized gelatin microparticles by incubating. The mixture was vortexed at maximum speed and incubated overnight at 4 ° C. to cause adsorption.

  In vitro release kinetics from hPE-added microparticles. In order to assess the degradation kinetics of gelatin microparticles, 10 mg (dry weight) of empty uncrosslinked microparticles were incubated in 1 ml phosphate buffered saline (PBS) at pH 7.4, 37 ° C., Protein release kinetics were compared with release from hPE-added and un-added crosslinked microparticles using a protein assay kit. Briefly, the supernatant of each specimen was collected periodically (after 1, 2, 3 and 6 hours and then daily) and replaced with fresh PBS. While measuring absorbance at 562 nm using a plate reader (Bio Teck Sinergy 2 plate reader, Bio Teck Instrument, Inc., Winooski, Vermont; USA), the Micro BCA assay kit (Thermo Scientific, Mass., Mass. USA) was used. The protein in the supernatant was quantified. The protein concentration was compared to a freshly prepared standard of bovine serum albumin at concentrations ranging from 200 to 0.5 μg / mL. Final values are obtained as total released protein from the start of the experiment and normalized as a function of the empty uncrosslinked microparticle dry weight or the estimated wet weight before the start of the experiment. Absorbance values are normalized to that of phosphate buffered saline (pH 7.4). The linear range of the assay was 2-40 μg / mL and the detection limit was 0.1 μg / mL. Each experiment was performed with 3 samples per point.

Preparation of 3D in vitro angiogenesis assay. A non-planar 3D in vitro angiogenesis assay was prepared using a type I collagen matrix embedded with HUVEC and PE-added microparticles. A collagen hydrogel matrix was prepared by mixing 8 mL cold Vitrogen collagen, 1 mL sterile PBS and 1.166 mL 0.1 M NaOH solution. The color transition of the solution from red to purple showed a change in pH. The pH of the solution was checked with pH test paper and adjusted to 7.4 by adding a few drops of 0.1M NaOH or 0.1M HCl solution. Prior to gelation, 3 mL of HUVEC in solution was added at a concentration of 40,000 cells / mL and 2 mg of added microparticles prepared as previously described ¹⁷ were added. Thereafter, the collagen / cell / microparticle solution was mixed by pipetting until homogeneous, and 500 μL was pipetted into each well of a 48-well plate. To induce collagen gelation, the culture plate was placed in a 37 ° C. oven for 30 minutes. The final thickness of the collagen hydrogel was 6.6 mm. Finally, 100 μL of angiogenic medium per cm ² was added to each well (containing the hydrogel with seeded cells, gelled and embedded microparticles), and the plate was placed in a humidified 5% CO ₂ incubator at 37 ° C. .
Analysis method

  Fluorescent staining with calcein AM. Calcein AM staining was performed using a survival killing assay (Invitrogen-Life Technologies, NY, USA). Briefly, calcein AM was pipetted directly into the medium present in the culture wells at a final concentration of 2 μg / ml. Stained cells were incubated at 37 ° C. for 30 minutes, and these were then observed using an inverted fluorescence microscope (Zeiss Axiovert 200 inverted fluorescence microscope).

Qualitative and quantitative network formation analysis. Analysis of cellular responses to experimental conditions in an in vitro model of angiogenesis was performed in previous studies (each incorporated herein by reference) ^34-37 in several semi-qualitative and quantitative methods. ing. Both morphological (average tubule length) and topological (number of branch points and number of meshes) parameters were considered. Because they allow for the characterization of the spatial organization of ECs in capillary-like networks.

  After cell staining, the field of view selected for each sample was taken. The resulting images were analyzed using ImageJ 1.45s (Wayne, Rasband-National Institutes of Health, USA).

Bifurcation points (BP), which are nodes where the branches meet or from which the tubules emerge, were identified and counted. A mesh of ³⁸ cell networks identified by an avascular region surrounded by blood vessels arranged in a hexagon was also manually counted. Finally, the length of the narrow tube was evaluated by drawing a straight line (white dotted line in FIG. 8 of the zoom image) along each thin tube, and the measurement of the straight line was automatically calculated by software.

  Scanning electron microscopy (SEM). The microparticle surface structure was investigated by scanning electron microscopy (S-4000 FE-SEM, Hitachi High Technologies, TX, USA). The lyophilized sample was mounted on an aluminum sample stage using double-sided graphite tape and coated with gold and palladium using a scatter (deskV, Denton Vacuum). Sample images were then taken with a Hitachi S-4000 FE-SEM (TX, USA).

Statistical analysis. The experiment was performed using three samples per point. All charted data are presented as mean ± standard deviation. Data analysis was performed using SPSS (IBM, Somers, NY). The significance test was calculated using a two-sided Student's t-test with unequal variance and no correspondence. The significance level was set at p <0.05.
result

  PE angiogenic ability. The data collected shows that the extent of HUVEC's angiogenic response to PE is strongly influenced by the incubation time and the number of inoculations given PE.

  For incubation time, HUVECs incubated with PE formed an angiogenic-like network whose morphology varied as a function of incubation time (FIGS. 9A-B). On the first day after sowing, HUVEC did not yet form a clear network (FIG. 9A, day 1). The mesh was numerous (102.67 ± 3.52) and small, and short tubules were observed (average tubule length: 78.85 ± 3.24 μm). The number of branch points (BP) was large (105 ± 9.85), but it was difficult to identify in the presence of several cell clusters. On the third day of culture (the third day in FIG. 9A), a well-formed network appeared. This is the presence of fewer BPs (71 ± 6.25), longer tubules (average length: 136.43 ± 15.23 μm) and wider and fewer numerous meshes (61.33 ± 8.02) Also confirmed by. On day 5 of culture (FIG. 9A, day 5), the network was still visible, but began to decompose. The tubules were longer (170.56 ± 16.51 μm) but fewer and thinner. Meshes were difficult to identify and the number decreased (36.33 ± 6.02). For BP, the number decreased slightly (63 ± 4). The degradation of the network can be attributed to the fact that the cells have already used up the angiogenic molecules contained within the PE. No tubule formation was observed in the control plate (upper right corner of each image), thus indicating that the change in cell morphology was not due to stress or other external factors.

  A semi-quantitative analysis of the spatial composition of HUVEC is shown in the histogram of FIG. 9B. Bars indicate average tubule length, number of BPs and number of meshes. It can be noted that a statistical difference was found between day 1 and day 5 considering all three parameters.

  In this study, capillary-like network formation was affected by the number of PE inoculations (FIGS. 10A-B). The average tubule length increased from cells that received a single inoculation to those received three times, but both the number of BP and mesh decreased (FIGS. 10A and 10BB). There was no clear network formation after a single inoculation of PE. Although the mesh was numerous (76.67 ± 19.74), it did not spread as confirmed by the average tubule length (129.71 ± 12.88 μm) (FIG. 10A, first panel). Furthermore, there were several BPs (110.78 ± 15.40) and cell clusters, thus indicating that the cells were not well connected. Cells that received two inoculations formed a network, but some clusters were still present (FIG. 10A, second panel). This may suggest that the network is not fully mature. However, while the average tubule length increased (139.91 ± 8.93 μm), the number of meshes and branch points decreased (52.67 ± 19.74 and 84.33 ± 11.86, respectively), thus the network Have probably changed towards a clearer structure. Cells that received 3 inoculations had a wider but fewer mesh (43.89 ± 6.52) and a reduced number of BPs (80.67 ± 11.92) (FIG. 10A, 3 Eye panel). The tubule length increased (153.89 ± 8.54 μm), suggesting that the tubules joined together to improve the network structure. Also, in this case, no tubule formation was observed in the control plate (upper right corner of each panel).

  The bar graph in FIG. 10B shows a semi-quantitative analysis of the spatial organization of HUVEC. Bars showing average tubule length, number of BPs and number of meshes are divided into three groups according to the number of PE inoculations received (1, 2 or 3). Data show an increase in mean tubule length from cells that received one inoculation (129.71 ± 12.88 μm) to cells that received three times (153.89 ± 8.54 μm), a more mature network of cells Suggested that they are organized. This finding indicates that both the number of meshes (from 76.67 ± 19.74 to 43.89 ± 6.52) and the number of BPs (from 110.78 ± 15.40 to 80.67 ± 11.92). This is also supported by the observed decrease. From the statistical analysis performed, differences were found in all three parameters between cells that received one inoculation and those that received three.

With respect to the retained biological activity of the extract, the extract maintains its angiogenic capacity until storage for 15 days in a humidified 5% CO ₂ incubator at 37 ° C. (FIG. 11). No significant difference was observed between days 1 and 7 of storage. In view of all these features of PE, PE was considered suitable for constant and sustained release over time.

Characterization of gelatin microparticles. Size distribution analysis of gelatin microparticles of some preparations indicated that these size ranges were between 20.86 ± 3.53 and 124.083 ± 13.01 μm. Nearly 80% of these had diameters in the range of 20-80 μm (FIG. 12B). SEM inspection of the microparticles shown in FIG. 12A showed a difference in surface morphology between the empty microparticles and the added microparticles. Empty microparticles (upper row) showed a smooth surface and regular shape. After the addition (bottom row), the particles were larger and had an irregular surface. These two appearances mean (i) PE attachment (adsorption) to the surface and (ii) PE penetration (absorption) ²⁷ inside the particles.

  FIG. 13 shows the degradation profile of non-crosslinked microparticles. This experiment was performed only with empty microparticles to evaluate the degradation kinetics of gelatin in PBS. An initial burst was observed. After 6 hours, the cumulative percent release was 6.45% ± 0.12 (FIG. 13, inset). The burst was followed by a slower release. After 20 days, the total cumulative release was 18.65% ± 0.09 (FIG. 13, main figure).

  FIG. 14A shows in vitro degradation of empty microparticles (black, no PE) and in vitro release of microparticles with added PE (grey, PE). In both cases, a small initial burst was observed. After 48 hours, the cumulative percent release from the added microparticles was 1.03% ± 0.08, but for empty microparticles it was 0.76% ± 0.05, almost constant in both cases Release continued. The cumulative percent release from added and empty microparticles after 22 days was approximately the same, as highlighted in the graph, were 4.65% ± 0.07 and 4.65% ± 0.11, respectively. On day 23, the release from empty microparticles was slightly greater than that from additive particles (5.18 ± 0.05 and 4.92 ± 0.09, respectively). The experiment was terminated based on evidence that most of the PE was released.

  FIG. 14B shows the percent release difference between the added and empty microparticles. Two peaks can be observed. The first peak is from hour 1 to day 5, and the second is from day 5 to day 22. The first peak indicates that PE bound to the particle surface was first released in a shorter period of time. The second peak, possibly due to erosion of a large amount of particles, lasts longer and the total cumulative release is larger. This indicates that PE is gradually released from the pores or channels formed in the microparticles by the release medium.

Despite the presence of an initial “burst”, the release kinetics shown from the microparticles obtained by filtration are close to the zero order profile, which means that the release is nearly constant. We argue that this may be due to the coexistence of different sized particles in the same sample. Release rate is affected by particle size, more precisely, smaller microparticles have a larger release as a result of an increase in surface area to volume ratio. This phenomenon has already been used to modulate drug or protein release to achieve a controlled near zero order release profile ²⁵ .

The total cumulative release from both empty and added microparticles is somewhat lower than the values reported in the ^39,40 literature, usually over 20%. This can be attributed to a high degree of crosslinking.

3D in vitro angiogenesis assay. After determining the optimal cell density and PE volume, an angiogenesis assay was prepared as described in the method above. After 3 days of culture, there was no sign of tubule formation and no difference was seen between HUVEC seeded with PE-added microparticles and control (results not shown). After 5 days in culture, cell morphology changed and budding was observed, but no capillary-like network was formed yet (FIG. 15). Budding is an early stage of angiogenesis, thus the presence of budding after 5 days in culture demonstrates an angiogenic response from HUVECs ⁴¹ . Since no network was formed 3 days after seeding, the amount of PE released by the particles may have been insufficient to promote angiogenesis within the observed time frame. However, budding formation indicates that PE has been released and an angiogenic response has begun. The release rate of the microparticles can be optimized to promote tubule formation within a shorter time frame.

DISCUSSION AND CONCLUSION The findings of this example demonstrate that the angiogenic response of HUVEC is influenced not only by incubation time but also by the number of PE inoculations received. In particular, in early experiments, the capillary network began to form 1 day after sowing and became well defined after 3 days and began to degrade after 5 days. It has also been shown that HUVECs form more mature capillary networks when they receive more than one PE inoculation. Furthermore, in this example, the network did not degrade after 5 days, as in the experiment discussed above. Finally, the extract maintains its angiogenic capacity until the 15th day of storage. In view of all these features of PE, PE can be considered suitable for constant and sustained release over time. Thus, with the goal of improving its ability to induce and regulate angiogenesis, drug delivery methods for multiprotein mixtures have been developed using biodegradable gelatin microparticles to further conserve biological activity. , Controlled and prolonged the delivery of hPE.

  For this purpose, biodegradable gelatin microparticles were prepared as a delivery system for PE. The absorption and adsorption of hPE on gelatin particles was evaluated by SEM inspection and analysis of in vitro release kinetics. SEM analysis shows an increase in size and a change in morphology between the additive and empty particles, which is a result of hPE adhesion (adsorption) to the surface and / or penetration (absorption) into the particles. is there. The release kinetics shows two peaks, the first peak is believed to be the result of the release of protein bound on the surface of the microparticles, which is smaller than the second peak and occurs in a shorter period of time. It was. The second peak was thought to be due to massive erosion. This is because it had a higher total cumulative release for a longer period of time. Despite the presence of an initial “burst”, the release kinetics were close to the zero order profile after the first day. The analysis revealed that the particles are suitable for use as a mediator for the sustained release of the extract in a 3D in vitro angiogenesis assay.

  These studies also show that when incorporated into 3D type I collagen matrix, hPE-added gelatin microparticles induced cells to long morphology and early stages of tubule formation with some cell sprouting after 5 days in culture ( FIG. 15, right). Since budding is an early stage of angiogenesis, the presence of budding implies an angiogenic response from HUVEC. The observed total cumulative release from both empty and loaded microparticles was approximately 5% after 23 days, as reported in the literature when a single protein is released from gelatin microparticles. Significantly lower than the 20-30% value. Despite the low cumulative release, the concentration of hPE released by the particles was still high enough to produce an angiogenic response within the collagen hydrogel. Thus, collagen hydrogels embedded with hPE-added gelatin microparticles can be used in in vitro angiogenesis assays, ranging from screening anti-angiogenic cancer drugs to mechanistic studies of tubule and angiogenesis network formation. There is a possibility.

  Overall, this example demonstrated that biodegradable gelatin microparticles can be used as a mediator for sustained release of multiprotein hPE in a 3D angiogenesis assay. Cell sprouting was observed after 5 days in culture using hPE-added gelatin microparticles, but no interconnected capillary network was formed. Increasing the amount of hPM released by the particles or extending the culture time can promote the formation of a more interconnected capillary network.

Reference for Example 2
1.http: //www.donatelifeny.org/about-donation/data/.
2. Kim, B.-S. & DJ, M. Development of biocompatible synthetic extracellular matrices for tissue engineering.Treds in Biothecnology 16, 224-230 (1998).
3. Bach, FH Xenotransplantation: problems and prospects. Annual review of medicine 49, 301-310 (1998).
4. Dvir, T., Timko, BP, Kohane, DS & Langer, R. Nanotechnological strategies for engineering complex tissues.Nature nanotechnology 6, 13-22 (2011).
5. Langer, R. & Vacanti, J. Tissue Engineering. Science 260, 920-926 (1993).
6. Martin, Y. & Vermette, P. Bioreactors for tissue mass culture: Design, characterization, and recent advances.Biomaterials 26, 7481-7503 (2005).
7. Muschler, GF, Nakamoto, C. & Griffith, LG Engineering Principles of Clinical Cell-Based Tissue Engineering.The Journal of Bone and Joint Surgery 1541-1558 (2004).
8. Laschke, MW et al. Angiogenesis in tissue engineering: breathing life into constructed tissue substitutes.Tissue engineering 12, 2093-104 (2006).
9. Soker, S., Machado, M. & Atala, a Systems for therapeutic angiogenesis in tissue engineering.World journal of urology 18, 10-8 (2000).
10. Adair, TH & Montani, J.-P. Angiogenesis. Colloquium Series on Integrated Systems Physiology: From Molecule to Function 2, 1-84 (2010).
11. Stein, I., Neeman, M., Shweiki, D., Itin, A. & Keshet, E. Stabilization of vascular endothelial growth factor mRNA by hypoxia and hypoglycemia and coregulation with other ischemia-induced genes. Molecular and Cellular Biology 15, 5363-8 (1995).
12. Pardali, E., Goumans, M.-J. & Ten Dijke, P. Signaling by members of the TGF-β family in vascular morphogenesis and disease,,. Trends in Cell Biology 20, 556-567 (2010).
13. Vailhe, B., Vittet, D. & Feige, J. In Vitro Models of Vasculogenesis and Angiogenesis. 81, 439-452 (2001).
14. Largo, RA, Ramakrishnan, VM & Ehrbar, M. Angiogenesis and Vascularity for Tissue Engineering Applications. (2010).
15. Carmeliet, P. Angiogenesis in health and disease.Nature medicine 9, 653-60 (2003).
16. Achen, MG & Stacker, SA The vascular endothelial growth factor family; proteins which which the development of the vasculature.International Journal of Experimental Pathology 79, 255-265 (1998).
17. Carmeliet, P. Mechanisms of angiogenesis and arteriogenesis.Nature medicine 6, 389-95 (2000).
18. Formiga, FR et al. Angiogenic therapy for cardiac repair based on protein delivery systems.Heart failure reviews 17, 449-73 (2012).
19. Iruela-Arispe, M. & Dvorak, H. Angiogenesis: A dynamic balance of stimulators and inhibitors.
Thrombosis and Haemostasis 78, 67-677 (1997).
20. Lee, K., Silva, EA & Mooney, DJ Growth factor delivery-based tissue engineering: general approaches and a review of recent developments. 153-170 (2011) .doi: 10.1098 / rsif.2010.0223
21. Moore, MC Modulation of nutrient deficiences occuring in engineered ex vivo tissue scaffolds. (2013).
22. Vasir, JK, Tambwekar, K. & Garg, S. Bioadhesive microspheres as a controlled drug delivery system. International Journal of Pharmaceutics 255, 13-32 (2003).
23. Chaturvedi, G. Microspheres technology and its application. (2009).
24. Determan, AS, Trewyn, BG, Lin, VS-Y., Nilsen-Hamilton, M. & Narasimhan, B. Encapsulation, stabilization, and release of BSA-FITC from polyanhydride microspheres. Journal of controlled release: official journal of the Controlled Release Society 100, 97-109 (2004).
25. Narayani, R. & Panduranga Rao, K. Gelatin microsphere cocktails of different sizes for controlled release of anticancer drugs. International journal of pharmaceutics 143, 255-258 (1996).
26. Tamizharasi, S., Rathi, J. & Rathi, V. Formulation and Evaluation of Pentoxifylline-loaded poli-e- caprolattone microspheres. Ind J of Pharm Sci 70, 333-5 (2008).
27. Cohen, S., Yoshioka, T., Lucarelli, M., Hwang, LH & Langer, R. Controlled Delivery Systems for Proteins Based on Poly (Lactic / Glycolic Acid) Microspheres. Pharmaceutical Research 8, 713-720 (1991 ).
28. Vernon, RB & Sage, EH A Novel, Quantitative Model for Study of Endothelial Cell Migration and Sprout Formation within Three-Dimensional Collagen Matrices. 133, 118-133 (1999).
29. Staton, C. a, Reed, MWR & Brown, NJ A critical analysis of current in vitro and in vivo angiogenesis assays.International journal of experimental pathology 90, 195-221 (2009).
30. Montesano, R. & Pepper, MS Three dimensional in vitro assay of endothelial cell invasion and capillary tube morphogenesis.Vascular Morphogenesis: In vivo, in vitro, in mente 79-110 (1998).
31. Auerbach, R., Lewis, R., Shinners, B., Kubai, L. & Akhtar, N. Angiogenesis assays: a critical overview.Clinical chemistry 49, 32-40 (2003).
32. Jaffe, EA, Nachman, RL, Becker, CG & Miinick, CR Culture of Human Endothelial Cells Derived from Umbilical Veins. 52, 2745-2756 (1973).

(Example 3)
Induction of placental extract release from PLGA microparticles to modulate angiogenesis Single administration of the human placenta extract of the present disclosure can be performed at an early stage of in vitro and in vivo angiogenesis, as described above in Example 1. Has been shown to induce and regulate. Since multiple doses of hPM over time have been shown to promote further capillary network development, controlled delivery further stabilizes network formation for an extended period of time, as described in Example 2 above. He made a hypothesis. In this example, hPE was encapsulated in poly (lactic acid-co-glycolic acid) (PLGA) microparticles to extend the release period. The microparticle preparation was optimized for hPE addition, characterized morphological characteristics (size, encapsulation efficiency, porosity) and profiled protein release.

  In order to overcome the problems identified above for inducing angiogenesis for transplanted tissues or tissue constructs, the disclosed method can use human placenta to induce capillary network formation and perform the above-described implementations. A human placenta extract called human placenta matrix (hPM) was derived in this example containing angiogenesis and immunoregulatory proteins as described in the examples. Endothelial cells (HUVEC) seeded on hPM in vitro have been shown above to form an angiogenic network with upregulation of angiogenic genes, and in vivo, the matrix is a biological scaffold with added components It was shown to inhibit tissue fibrosis while inducing angiogenesis in the body. Furthermore, endothelial cells that received multiple hPM inoculations at regular time points (culture days 1, 3 and 5) received only a single inoculation whose network began to degrade after 5 days. Formed a more stable and long-lasting angiogenic network compared to the cultured cells. Thus, approaches for controlled delivery of matrices over time have been investigated to allow for longer lasting and more stable capillary network formation.

The complex and heterogeneous nature of hPM can complicate the controlled release mechanism. For example, different charge and chain properties of proteins can have an impact on loading efficiency due to interaction with each other and with the materials used for controlled release. Both natural and synthetic microparticulate materials have been investigated for their ability to encapsulate proteins. Natural materials such as collagen, chitosan and alginate provide biocompatible and non-invasive encapsulation techniques, which have a degradation rate between about 7 and 10 days, which is somewhat sustained. Allows release, but may limit use for longer term sustained protein release ^{11, 12, 13} . Chemical or photochemical crosslinking can slow the degradation of these microparticles ^9,14 .

PLA copolymers are used for protein encapsulation in biomedical applications and are FDA approved biocompatible synthetic materials ^{15, 16, 17} . These polymers can provide long-lasting controlled release of proteins and can be used to create composites and multilayer microparticles ^15,18 . Of these materials, PLGA or poly (lactic-co-glycolic acid) has been used for the controlled release of certain growth factors (eg BMP, VEGF, bFGF) ^{19, 20, 21} .

Studies are evaluating single protein PLGA encapsulation, some are investigating the simultaneous encapsulation of multiple proteins ^22,23 , but studies evaluating the encapsulation of complex heterogeneous mixtures of proteins are Absent. This example describes the composition and encapsulation technique for hPM using PLGA microparticles and evaluates the effect of controlled release of this mixture on 3D cultures of endothelial cells.

  In view of the heterogeneous composition of hPM, in this embodiment described in this example, the PLGA synthesis protocol was optimized to be suitable for multiprotein hPM release for use in cell culture. Consideration was made including optimization of microparticle size (to be suitable for regenerative medicine), high loading and encapsulation efficiency, low initial burst and controlled release profile. Furthermore, it was confirmed that hPM was released from the microparticles at a concentration suitable for inducing angiogenesis using endothelial cells. After microparticle synthesis and addition, an analysis was performed to assess whether the encapsulation process was selective for hPM protein. The effect of controlled hPM release on the induction of angiogenesis was evaluated using a 3D culture system using hPM-added microparticles embedded in an alginate-based hydrogel seeded with HUVEC. Cell behavior was assessed at specific time points during 28 days and compared to cells that received controlled component addition of hPM from PLGA microparticles over the entire period of culture. The response of cells that received two direct inoculations was evaluated.

Method Effect of multiple rapid dose hPM inoculations. Using an orbital shaker for 1 minute at 30 rpm and uniformly coating the wells of a 96-well tissue culture plate with a pipette, in vitro using HUVEC, the hPM component addition profile during the formation of an angiogenic network The effect analysis was determined. The plate was then incubated at 37 ° C. for 30 minutes to warm hPM. HUVEC was suspended in angiogenic medium and pipetted onto hPM and then placed in a humidified 6% CO ₂ incubator at 37 ° C. For controls, HUVEC were cultured in angiogenic medium at 20,000 cells / cm ² . Three different profiles of hPM inoculation were compared. 1) hPM was inoculated and added only on day 1 (day of sowing), 2) hPM was inoculated on days 1 and 3, and 3) hPM was inoculated on days 1, 3 and 5. Cells were cultured for 7 days and the medium was replaced on days 3 and 5. As a control, cells were seeded directly at the bottom of the plate at the same density and the medium was replaced every 2 days. Cells were stained on day 7, imaged using a fluorescence microscope, and angiogenesis was quantified by analyzing the images as described in the “Angiogenesis Quantification” section.

  PLGA microparticle preparation. PLGA (poly (DL-lactide-co-glycolide)) microparticles were prepared using a water-in-oil-in-water emulsion (Protocol 1) (Duret, Cupertino, CA). One aqueous solution (W1) was prepared using the hPM protein mixture as described in Example 1, and 2 g of polyvinyl alcohol (wt 30,000-70,000, 87-90% hydrolysis, Sigma-Aldrich, St. The other aqueous solution (W2) was prepared by dissolving Louis, MO) in 100 mL DI water. Oil solution (O) was obtained by dissolving 90 mg of PLGA in 3 mL of chloroform until the solution appeared clear. W1 was added to O and homogenized at 20,000 rpm for 1 minute. Using a micropipette, the obtained primary emulsion was added dropwise to W2 while stirring at 300 rpm. When all the primary emulsion was added, the resulting secondary emulsion was lightly covered with aluminum foil and allowed to stir overnight (300 rpm) in a fume hood to evaporate the solvent. Next, the secondary emulsion was centrifuged at 1000 rpm for 10 minutes, the supernatant was removed, and the microparticles were washed twice more with DI water. The hardened microparticles were suspended in DI water, lyophilized for 48 hours and then stored at 4 ° C. until needed. For control, W1 was made by substituting bovine serum albumin (BSA) for hPM (Sigma-Aldrich, St. Louis, MO) to create single protein loaded PLGA microparticles.

  Due to the heterogeneous nature of hPM mixtures, encapsulation is a difficult element of controlled release from PLGA microparticles. Therefore, the impact of protocol modifications on size and release was analyzed. First, the effect of longer homogenization in the primary stage was evaluated (protocol 2: 2 minutes instead of 1 minute). The effects of additional homogenization steps introduced after formation of the secondary water-in-oil-in-water emulsion and before solvent removal were then investigated (Protocol 3: 1 min further homogenization; Protocol 4: 20 sec further homogenization). ). After each modification, the morphological characteristics and associated release rates of the microparticles were evaluated and compared before moving on to the optimization process.

  PLGA microparticle morphological characterization. The morphological characteristics of PLGA microparticles were evaluated using microscopy. The average size of the microparticles was evaluated using an inverted optical Leica microscope equipped with a color digital camera (Leica DM IL LED, Leica Microsystems Inc., IL, USA). The captured images were analyzed using the free software ImageJ 1.45s (Wayne, Rasband-National Institutes of Health, USA-http: //imagej.nih.gov/ij/). The diameter of each particle was determined by automatic tracing using Leica LAS software (Wetzlar, Germany) after manually tracing the particles.

  Scanning electron microscopy (SEM) (S-4000 FE-SEM, Hitachi High Technologies, TX, USA) was used to characterize the surface morphology and porosity of the microparticles. For shape and surface analysis, lyophilized samples were mounted on an aluminum sample stage using double-sided graphite tape and coated with gold and palladium using a scatter (deskV, Denton Vacuum). The coated samples were then examined at an accelerating voltage of 2 kV, photographed at low magnification to evaluate average dimensions and shape, and porosity and morphological feature changes at high magnification.

  Addition efficiency. Direct measurement of encapsulated protein after microparticle dissolution was used to determine the efficiency of hPM addition in PLGA microparticles. This is because Ravi. Development and characterization of polymeric microspheres for controlled release protein loaded drug delivery system. 70, (2008) and Igartua, M .; , Stability of BSA encapsulated into PLGA microspheres using PAGE and capillary electrophoresis. International Journal of Pharmaceuticals 169, 45-54 (1998), both of which are incorporated herein by reference for hydrolysis techniques, were carried out by adaptation of the hydrolysis techniques described in International Journal of Pharmaceuticals 169, 45-54 (1998). Briefly, 15 mg of lyophilized microspheres was digested with 5 mL of 0.1 M NaOH containing 5% w / v SDS to give a clear solution on a shaker for 15 h at room temperature. Hydrolyzed until The resulting clear solution was then neutralized to pH 7 by adding 1M HCl and centrifuged at 5000 rpm for 10 minutes. The protein concentration in the supernatant was analyzed using a UV-visible spectrophotometer at 562 nm and the Pierce BCA standard protein assay with 3 samples per point. Encapsulation efficiency was expressed as the ratio of actual protein content to theoretical protein content.

  Characterization of in vitro hPM release. To characterize the release of hPM from PLGA microparticles, 10 mg of added microparticles was suspended in 1 mL of phosphate buffered saline (PBS). Tubes were incubated in a shaking incubator and three samples were evaluated for each protocol. Every 2 days, the tube was centrifuged at 5000 rpm for 5 minutes, the supernatant was collected, and an equal volume of fresh PBS was added to the tube. The protein concentration was quantified by measuring the absorbance at 562 nm with a spectrophotometer. Comparison of the results from the standard Pierce BSA assay and the micro-protein BSA assay was performed to give a more reliable assessment of the released protein at each time point tested. Measurements were collected using 3 samples per point. The protein concentration was compared to freshly prepared standards ranging from 200 to 0.5 μg / mL. The average absorbance of standard PBS was subtracted from all measurements. The amount of protein in each sample was summed with the amount at each previous time point to construct a cumulative release curve.

  Since hPM is a complex protein mixture that includes growth factors, extracellular matrix proteins and biomolecules characterized by various charges and properties, these studies have been used to determine whether encapsulation occurs selectively. Protein release from hPM-added microparticles was analyzed. SDS-PAGE analysis was performed on the supernatant collected after overnight stirring. Contains supernatant from hPM-added and BSA-added microparticles, and hPM and BSA in PVA used at the same concentration as added to the microparticles (50 μL hPM diluted with 10 mg BSA and 5 mL DI water in 50 mL DI water) Four solutions were analyzed. A protein standard was used to evaluate the molecular weight of the detected protein. SDS-page was performed using a BIO-RAD electrophoresis system (Hercules, CA). Following electrophoresis, polyacrylamide gels were stained with Coomassie blue for 1 hour under continuous shaking. The protein band was then enhanced by destaining the gel in a 4: 1: 5 methanol: acetic acid: DI aqueous solution. The final gel was washed with DI water and imaged.

Alginate-hydrogel based 3D culture. An angiogenesis assay using added microparticles was created to evaluate its application for controlled hPM release. Human umbilical vein endothelial cells (HUVEC) were seeded and cultured in 3D alginate gel (1.5% alginate) in which hPM-added microparticles were embedded. Sample groups include cells suspended in alginate containing pure hPM, cells suspended in alginate containing embedded hPM-added microparticles, cells suspended in alginate without microparticles, and empty microparticles Cells suspended in alginate containing. For all sample groups, HUVEC was suspended in the medium and gently suspended in the alginate matrix before polymerization by pipetting 0.054M calcium chloride solution. The alginate-cell suspension was left to allow polymerization for 15 minutes, then the gel was washed with PBS, the culture medium was added, and the plates were incubated at 37 ° C. in a humidified 6% CO ₂ incubator.

In order to obtain a comparison result between the condition in which hPM was added directly to the matrix and the condition in which hPM was released from the microparticles, the concentration of hPM mixed in each gel was released after 21 days from the embedded microparticles. The protein concentration was equal (78 μg protein per cm ² ). The effect of sustained controlled hPM release from microparticles was determined by morphological characterization of the cellular network in the gel stained with calcein AM (Life Technologies, Grand Island, NY) and rapid dose inoculation of hPM and 7,14 , 21 and 28 days.

  Angiogenesis quantification. Quantification of the angiogenic output was performed using ImageJ 1.45s (NIH, Bethesda, MD). Images obtained using an inverted fluorescence microscope at a magnification of 5x were obtained using the following parameters: number of meshes formed by HUVEC over time, number of branch points and tubule-like structure as described in Example 1 Were processed to evaluate and quantify the length of

Statistical analysis. Experiments on both microparticles and 3D cultures were performed using 3 samples per point. All graphs and tabulated data are expressed as mean ± standard error of the mean. Analysis was performed using Excel (Microsoft Office) and Minitab 15 (Minitab, State College, PA). Significance was calculated using the ANOVA test, where more than 2 conditions were evaluated and specific differences were evaluated using a post hoc test. When comparing only two conditions, an unequal variance, unpaired two-tailed Student's t-test was used. The significance level was set at ^* p <0.05.

Results Effect of multiple rapid dose hPM inoculations. The formation of HUVECs in the angiogenic network was affected by the number of hPM component additions received over time (FIGS. 16A-16D). As shown in FIG. 16E, tubule length increased as a function of the number of inoculations. Although there was no significant difference in the number of meshes and branch points (FIGS. 16F-16G), an increasing and more mature trend of the mesh was observed as a function of the number of inoculations. Since mature angiogenesis is characterized by long tubules and a reduced number of branch points and meshes, the results confirm that continuous administration of hPL helps the endothelial cells to organize a stable network.

  Initial PLGA microparticle preparation: Protocol 1. The PLGA microparticles obtained from protocol 1 had a mean distribution of 447 ± 32 μm and a normal distribution with a size ranging from 100 to 1000 μm. Three batches of microparticles were prepared under the same conditions and a repeatable distribution could be demonstrated. The addition efficiency of hPM in PLGA microparticles was 64 ± 4% (data not shown).

  In vitro release analysis showed a low initial burst corresponding to 10% of the total amount of encapsulated protein (21.38 ± 0.83 μg / mL), but 51.23 ± of the initially encapsulated protein. 0.19% (134.56 ± 0.50 μg / mL) was released from PLGA microparticles after 21 days (data not shown). The release rate profile appeared to be linear and constant throughout the 21-day period evaluated, but the amount of hPM released from the microparticles at the end of the evaluation period was (as described above) the angiogenic response. Concentrations were significantly lower compared to the use of rapid dose infusion of hPM (500 μg / mL) to induce

  Control BSA loaded PLGA microparticles had an average size of 239.60 ± 9.45 μm and a normal distribution of sizes from 50 to 400 μm, with 70% of the microparticles ranging between 150 and 350 μm. The addition efficiency of BSA-added PLGA microparticles was 76 ± 3% of the total amount of protein added initially. In vitro release analysis showed a higher initial burst compared to hPM-loaded microparticles at 20% (51.52 ± 1.76 μg / mL) of the total amount of encapsulated BSA, but initially encapsulated 66.7 ± 9.55% (166.74 ± 23.8 μg / mL) of the released protein was released from PLGA microparticles after 21 days (data not shown). The release rate profile appeared to be linear and constant throughout the time period evaluated. The difference in results observed between hPM-added PLGA microparticles and BSA-added PLGA microparticles suggested that release kinetics could be improved through optimization of the preparation and addition protocol.

  PLGA microparticle size optimization and morphology characterization. Size analysis from microparticles created using Protocol 2 homogenized primary emulsion for 2 minutes (1 minute more than Protocol 1) showed an average diameter of 276.78 ± 13.41 μm (FIGS. 17A, 17D). ). The microparticles created using Protocol 3 with a 1 minute homogenization step added to the secondary emulsion preparation step had an average size of 37.79 ± 1.30 μm (FIGS. 17B, 17E). However, the microparticles created using protocol 4 in which a 20 second homogenization step was added to the secondary emulsion had an average size of 91.85 ± 2.92 μm (FIGS. 17C, 17F). .

  The results from SEM analysis showed no significant difference in surface porosity between the different protocols. The surface of the microparticles obtained with the different modifications described appeared to be round shapes with all homogeneous surfaces (FIGS. 18A-18F). 18A-18C show the microparticle shape, and FIGS. 18D-F show the microparticle surface porosity of the particles prepared according to protocols 2, 3 and 4, respectively.

  Addition efficiency. Encapsulation efficiency did not appear to be linearly correlated with the size of the microparticles (FIG. 19A). Encapsulation efficiency increased to 73 ± 4% for protocol 2 and decreased to 63 ± 3% when the size was significantly reduced in protocol 3. The highest encapsulation efficiency was obtained using protocol 4 and was 79 ± 9% of the encapsulated protein with the initial total protein addition.

  In vitro release of hPM from optimized hPM-added microparticles. The release rate of hPM from microparticles created using an optimized protocol is a direct result of the difference in size and loading efficiency, and the release profile trends obtained by modifying the homogenization step are similar It seemed to do. The initial burst did not change significantly with modification of the duration of the primary emulsion homogenization step, from protocol 14.14 ± 0.31% (21.38 ± 0.83 μg protein / mL) in protocol 1 (data not shown). 2 to 6.42 ± 2.07% (18.88 ± 6.11 μg protein / mL). A significant increase in burst release was observed when secondary emulsion homogenization was introduced. Compared to 27.72 ± 3.73% for protocol 4 (106.33 ± 14.33 μg / mL), 23.75 ± 0.40% (60. 82 ± 1.03 μg / mL) was released during the first day (FIG. 19B).

  Cumulative release after 21 days correlated with microparticle size, with smaller microparticles allowing more hPM release in the assayed period. After 21 days of release, the microparticles from protocol 2 released 64.09 ± 1.70% (188.43 ± 5 μg / mL) of the total protein encapsulated, while MP from protocol 3 was Releases 98.62 ± 1.40% (252.48 ± 3.6 μg / mL) and MP from protocol 4 is 87.60 ± 1.12% (335.98 ± 4.72 μg / mL). Released (FIG. 19B). Further analysis to compare the P3 and P4 release profiles over an extended period of time showed that after 30 days the microparticles from protocol 3 were completely degraded and no longer released hPM, but the microparticles from protocol 4 , Showed that up to 93.12 ± 0.2% (357 ± 4.09 μg / mL) of the total protein encapsulated was released.

  Results from the SDS page were limited by the low concentration of protein in the supernatants collected and analyzed. However, the analysis showed uniform encapsulation of hPM in PLGA microparticles without any evidence of selective encapsulation for a particular protein. For both BSA-added and hPM-added microparticles, analysis of the supernatant collected after the microparticles solidified showed similar protein content with lower concentrations (FIG. 20).

Alginate-hydrogel based 3D culture. Results from the angiogenesis assay showed that controlled release of hPM can improve the stability of capillary-like structures in the endothelial cell seeding matrix. FIG. 21 shows that after 7 days of culture, only a few budding and tubule structures were observed in HUVECs cultured in alginate hydrogels embedded with hPM-added microparticles (and no maturation network was observed). Angiogenesis in a matrix containing hPM is relatively more mature in the same time frame, with an average tubule length of 207.90 ± 15.31 μm, 1.27 ± 0.84 mesh / mm ² , and 9.60. ± 0.70 branch point / mm ² . However, after 14 days in culture, the alginate gel embedded with hPM-added MP began to organize primary tubules with an average length of 137.41 ± 9.77 μm and 6.66 ± 0.47 branch points / mm ² , The matrix containing pure hPM showed degradation of the angiogenic network, no mesh was observed and only isolated tubules with an average length of 226.58 ± 25.64 μm were observed. At 21 days, the MP-embedded gel showed immature mesh formation, an average tubule length of 204.07 ± 12.75 μm and an average of 11 ± 1.25 branch points / mm ² . Overall, after 21 days in culture, the formed angiogenic network remained stable with only minor changes in the number of branch points / mm ² (FIG. 21). After 28 days in culture, the average tubule length is 168.88 ± 10.31 μm, the average number of branch points / mm ² is 12.93 ± 1.61, and the average mesh number is 3.17 per mm ^2. It was ± 0.93.

Discussion Proteins encapsulated with PLGA microparticles are drug delivery systems for controlled release of growth factors, and the commonly used fabrication technique is the water-in-oil-in-water method ^19,20,21 . In this study, a complex human derived multiprotein mixture (placental matrix or hPM) containing over 2600 angiogenic and antifibrotic proteins was encapsulated in PLGA microparticles. The results show that hPM loaded PLGA microparticles can be used to sustain and control the release of multiprotein fusions including pro-angiogenic and fibrous proteins over time. When used with endothelial cells during in vitro culture, hPM-added PLGA microparticles allow the formation of a stable angiogenic network.

  These studies revealed that hPM had an effect on PLGA addition and microparticle morphology due to the complex multiprotein composition. Compared to single protein (BSA) encapsulation, hPM-added microparticles showed a significantly larger average size than that with BSA. This may be the result of complex interactions between proteins in hPM and PLGA polymers with a wide variety of different molecular charges, pH and sizes compared to BSA-doped microparticles. SDS-PAGE analysis revealed that a wide distribution could be observed in the supernatant with the same composition as diluted hPM, regardless of the low concentration of protein released from hPM-added PLGA microparticles. These results suggested that the encapsulation of the protein mixture was homogeneous and not selective for a particular protein and that the protein composition of hPM was not affected during microparticle preparation.

It has previously been shown that both the speed and duration of the homogenization step affect the microparticle size ^{17 27} . This example revealed a correlation between the size of hPM-added PLGA microparticles and the in vitro release rate. Although optimization of the homogenization of the primary emulsion stage created microparticles with the desired controlled release characteristics, the efficiency of addition of hPM in these particles was somewhat lower compared to other production protocols evaluated. Optimization of the secondary emulsion stage homogenization produced microparticles with increased encapsulation of the protein solution (up to 78%) in the desired size range. Interestingly, the encapsulation efficiency did not correlate with the size of the microparticles, but instead was influenced by the length of homogenization of the secondary emulsion, which was a correlation between the heterogeneous protein mixture and the polymer. It may be the result of the formation of fragmented incomplete particles due to action.

Using the above optimized PLGA microparticle synthesis protocol (which allows higher hPM encapsulation efficiency, optimized microparticle size and linear release over 30 days compared to non-optimized protocol) Added microparticles were incorporated into an alginate-based angiogenesis assay to assess the endothelial cell (HUVEC) response to controlled hPM release over time. The 3D angiogenesis assay embeds hPM-added microparticles in ^{30, 31} alginate gels selected as the basis of the assay due to cost-effective processability and overall good extracellular material properties It was created by After 21 days in culture, the total amount of protein delivered from the microparticles (78 μg / cm ² ) is lower than the concentration delivered in previous 2D angiogenesis studies using rapid dose infusion (263 μg / cm ² ) , HUVEC formed neovascularized tubules in the gel in the region close to the embedded hPM loaded PLGA microparticles. Compared to the rapid dosing component addition of hPM directly mixed with an alginate matrix, the use of hPM-added microparticles allowed for prolonged and sustained angiogenic network formation over 28 days. Cells formed angiogenic networks earlier in alginate gels mixed directly with hPM, but these networks degraded faster compared to gels with hPM-added microparticles. After 28 days, hPM release from the microparticles was almost zero order, but the formation of a new angiogenic mesh could still be confirmed, indicating that the controlled release of hPM from the PLGA microparticles formed the angiogenic network. It was suggested that the stability of was improved.

  Overall, this example describes the optimization of a PLGA microparticle synthesis protocol for use in delivering complex multiprotein mixtures with low initial bursts and high encapsulation efficiency. The method developed here was then used to develop a new angiogenesis assay that allowed for the formation of a sustained angiogenic network within the alginate matrix. Importantly, the developed technology has applications in the delivery of any complex serum and protein mixture, as well as applications in a wide range of tissue engineering and regenerative medical systems. These techniques can be applied to different materials (eg, collagen, fibrin, laminin) by embedding serum-supplemented PLGA microparticles to develop improved in vitro angiogenesis assays that persist in the cell seeding matrix. A better understanding of the role played by sex serum delivery can be gained.

Conclusion This example describes an embodiment of the method of hPM encapsulation, and angiogenic structure formation was observed using an alginate-based angiogenesis assay. A sustained angiogenic response over an extended period of 21 days was observed in the 3D hydrogel culture system. This confirmed the effectiveness of a controlled hPM release approach to guide the formation and maintenance of capillary networks.

References for Example 3
1. Laschke, MW et al. Angiogenesis in tissue engineering: breathing life into constructed tissue substitutes.Tissue Eng 12, 2093-2104 (2006).
2. Carmeliet, P. & Jain, RK Molecular mechanisms and clinical applications of angiogenesis.Nature 473, (2011).
3. Adair, TH Angiogenesis. (Morgan & Claypool Publishers, 2010). At <http://books.google.com/books?id=ykn66NeaPakC>
4. Chu, H. & Wang, Y. Therapeutic angiogenesis: controlled delivery of angiogenic factors.Ther Deliv 3, (2012).
5. Xie, J. et al. Induction of angiogenesis by controlled delivery of vascular endothelial growth factor using nanoparticles.Cardiovasc Ther 31, e12-18 (2013).
6. Chung, J. & Shum-Tim, D. Neovascularization in Tissue Engineering.Cells 1, 1246-1260 (2012).
7.Hughes, CS, Postovit, LM & Lajoie, GA Matrigel: a complex protein mixture required for optimal growth of cell culture.Proteomics 10, 1886-1890 (2010).
8. Moore, MC Modulation of nutrient deficiences occuring in engineered ex vivo tissue scaffolds. (2013).
9. Pan, S.-C., Wu, L.-W., Chen, C.-L., Shieh, S.-J. & Chiu, H.-Y. Angiogenin expression in burn blister fluid: implications for its role in burn wound neovascularization. Wound Repair Regen 20, 731-739 (2012).
10. Shi, H., Han, C., Mao, Z., Ma, L. & Gao, C. Enhanced angiogenesis in porous collagen-chitosan scaffolds loaded with angiogenin.Tissue Eng Part A 14, 1775-1785 (2008) .
11. Chan, OCM, So, K.-F. & Chan, BP Fabrication of nano-fibrous collagen microspheres for protein delivery and effects of photochemical coupling on release kinetics.Journal of Controlled Release 129, 135-143 (2008).
12. Jay, SM & Saltzman, WM Controlled delivery of VEGF via modulation of alginate microparticle ionic crosslinking. Journal of Controlled Release 134, 26-34 (2009).
13. Jay, SM, Shepherd, BR, Bertram, JP, Pober, JS & Saltzman, WM Engineering of multifunctional gels integrating highly efficient growth factor delivery with endothelial cell transplantation.FASEB J 22, 2949-2956 (2008).
14. Chan, BP et al. Photochemical cross-linking for collagen-based scaffolds: a study on optical properties, mechanical properties, stability, and hematocompatibility.Tissue Eng 13, (2007).
15. Cohen, S., Yoshioka, T., Lucarelli, M., Hwang, L. & Langer, R. Controlled Delivery Systems for Proteins Based on Poly (Lactic / Glycolic Acid) Microspheres. Pharm Res 8, 713-720 ( (1991).
16. Lassalle, V. & Ferreira, ML PLA nano- and microparticles for drug delivery: an overview of the methods of preparation. Macromol Biosci 7, 767-783 (2007).
17. Lee, WL et al. Fabrication and drug release study of double-layered microparticles of various sizes.J Pharm Sci 101, 2787-2797 (2012).
18. Lee, WL, Shi, W.-X., Low, ZY, Li, S. & Loo, SCJ Modeling of drug release from biodegradable triple-layered microparticles.J Biomed Mater Res A 100, 3353-3362 (2012) .
19. Simon-Yarza, T. et al. PEGylated-PLGA microparticles containing VEGF for long term drug delivery.International Journal of Pharmaceutics 440, 13-18 (2013).
20. Kirby GTS, White LJ, Rahman CV, Cox HC, Qutachi O, Rose FRAJ, Hutmacher DW, Shakesheff KM, Woodruff MA. PLGA-Based Microparticles for the Sustained Release of BMP-2. Polymers (2011). Doi: 3 (1): 571-586.
21. Wang, Y. et al. Degradable PLGA scaffolds with basic fibroblast growth factor: experimental studies in myocardial revascularization.Tex Heart Inst J 36, (2009).
22. Acharya, AP, Lewis, JS & Keselowsky, BG Combinatorial co-encapsulation of hydrophobic molecules in poly (lactide-co-glycolide) microparticles. Biomaterials 34, 3422-3430 (2013).
23. Roman, BS et al. Co-encapsulation of an antigen and CpG oligonucleotides into PLGA microparticles by TROMS technology.European Journal of Pharmaceutics and Biopharmaceutics 70, 98-108 (2008).
24. Ravi, S. et al. Development and characterization of polymeric microspheres for controlled release protein loaded drug delivery system. 70, (2008).
25. Igartua, M. et al. Stability of BSA encapsulated into PLGA microspheres using PAGE and capillary electrophoresis. International Journal of Pharmaceutics 169, 45-54 (1998).
26. Bouyer, E., Mekhloufi, G., Rosilio, V., Grossiord, J.-L. & Agnely, F. Proteins, polysaccharides, and their complexes used as stabilizers for emulsions: alternatives to synthetic surfactants in the pharmaceutical field ? Int J Pharm 436, 359-378 (2012).
27. Giteau, A., Venier-Julienne, MC, Aubert-Pouessel, A. & Benoit, JP How to achieve sustained and complete protein release from PLGA-based microparticles? International Journal of Pharmaceutics 350, 14-26 (2008).
28. Klose, D., Siepmann, F., Elkharraz, K. & Siepmann, J. PLGA-based drug delivery systems: Importance of the type of drug and device geometry.International Journal of Pharmaceutics 354, 95-103 (2008) .
29. Acharya, G. et al. A study of drug release from homogeneous PLGA microstructures. Journal of Controlled Release 146, 201-206 (2010).
30. Rowley, JA, Madlambayan, G. & Mooney, DJ Alginate hydrogels as synthetic extracellular matrix materials.Biomaterials 20, 45-53 (1999).
31. Gandhi, JK, Opara, EC & Brey, EM Alginate-based strategies for therapeutic vascularization. Ther Deliv 4, 327-341 (2013).
32. Zisch, AH, Lutolf, MP & Hubbell, JA Biopolymeric delivery matrices for angiogenic growth factors.Cardiovasc Pathol 12, (2003).
33. Goodwin, AM In vitro assays of angiogenesis for assessment of angiogenic and anti-angiogenic agents. Microvascular Research 74, 172-183 (2007).

Claims (20)

A human placenta extract obtained from a human placenta sample, wherein blood and solids are substantially removed from said extract, said extract comprising a placental protein comprising cytokines and growth factors; Human placental extract, wherein the placental protein was present in the placental sample;
The human placenta extract, wherein the human placenta extract is released from the microparticles over a period of time after being exposed to the cells in vivo or in vitro. Combined with the composition. The placenta extract is
Removing blood from a sample obtained from the human placenta sample to produce a crude placenta extract;
Mixing the crude placenta extract with a protein solubilizer to solubilize the protein in the crude extract;
Separating solid material from the solubilized protein-placenta extract mixture;
2. The composition of claim 1, wherein the composition is made by dialysis against a solubilized protein-placenta extract mixture to remove the protein solubilizer from the mixture to produce a human placenta extract.   2. The composition of claim 1, wherein the placenta extract comprises at least 20 different cytokines.   The composition of claim 2, wherein the protein solubilizer is urea.   The composition of claim 1, wherein the placental extract has the ability to modulate angiogenesis.   The composition of claim 1, wherein the biodegradable microparticles comprise poly (DL-lactide-co-glycolide) (PLGA) microparticles.   The PLGA microparticles are those to which the human placenta extract (hPE) is added, and the hPE-added PLGA microparticles are those produced using an oil-in-water process involving double homogenization. The composition according to claim 7.   The composition of claim 1, wherein the biodegradable microparticles comprise gelatin microparticles.   The composition of claim 1, wherein the biodegradable microparticles comprise microparticles of different sizes. A method of inducing angiogenesis comprising contacting a cell with a sustained release angiogenesis modulating composition, wherein the sustained release angiogenesis modulating composition comprises:
The human placenta extract combined with the microparticles such that the human placenta extract is released from the microparticles over a period of time after exposure of the microparticles to the cells in vivo or in vitro with biodegradable microparticles Wherein the placenta extract comprises an extract obtained from a human placenta sample, blood and solids have been substantially removed from the extract, and the extract contains cytokines and growth factors. A placental protein comprising, wherein the placental protein comprises a human placenta extract that was present in the placental sample.   The method of claim 10, wherein the cell is an endothelial cell. The placenta extract is
Removing blood from a sample obtained from the human placenta sample to produce a crude placenta extract;
The crude placenta extract is mixed with a protein solubilizer to solubilize the protein in the crude extract,
Separating solid material from the solubilized protein-placenta extract mixture;
11. The method of claim 10, wherein the method is made by dialysis against a solubilized protein-placenta extract mixture to remove the protein solubilizer from the mixture to produce a human placenta extract.   The method according to claim 12, wherein the protein solubilizer is urea. 11. The method of claim 10, further comprising combining the sustained release angiogenesis modulating composition with a biomaterial.   15. The method of claim 14, further comprising inducing angiogenesis on the biomaterial in vivo by implanting the biomaterial into a subject.   15. The method of claim 14, wherein the biomaterial is an engineered bioscaffold comprising a human derived substrate material.   15. The method of claim 14, wherein the biological scaffold comprises a decellularized human umbilical vein scaffold seeded with human endothelial cells.   The method of claim 17, wherein the human endothelial cells are human umbilical vein endothelial cells (HUVEC).   The method of claim 15, wherein the subject is a human.   A method of reducing inflammation in a subject comprising administering to the subject a composition comprising a human placenta extract obtained from a human placenta sample, wherein blood and solids are substantially removed from said extract. Wherein the extract comprises a placental protein comprising cytokines and growth factors, and the placental protein was present in the placental sample.