Immune Characterization and Phase I Clinical Testing of an Adipose Extracellular Matrix-Derived Biomaterial for Soft Tissue Reconstruction

IMMUNE CHARACTERIZATION AND PHASE I CLINICAL TESTING OF AN ADIPOSE EXTRACELLULAR MATRIX-DERIVED BIOMATERIAL FOR SOFT TISSUE RECONSTRUCTION

by Amy Elizabeth Anderson

A dissertation submitted to Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy

Baltimore, Maryland October 2017

ABSTRACT

Adipose tissue is used by surgeons for a variety of applications, including soft tissue reconstruction and wound healing. However, harvesting adipose tissue from patients presents challenges such as donor-site morbidity, outcome variability, and insufficient tissue volumes.

To address this clinical challenge, we aimed to create an "off-the-shelf" adipose material using mechanical and chemical processing techniques which remove lipids and living cells while processing the tissue into an injectable form preferred by patients and physicians.

A Phase I clinical study conducted in eight healthy volunteers assessed the safety and tolerability of Acellular Adipose Tissue (AAT) – an adipose extracellular matrix-derived biomaterial intended for the repair of soft tissue defects in humans. The trial evaluated subcutaneous injections of AAT administered in redundant tissues previously scheduled for removal in an elective surgical procedure (i.e. panniculectomy) between 1 and 18 weeks post-injection. AAT was well-tolerated in this first-in-human study and demonstrated satisfactory participant comfort and physician ease-of-use ratings. No patients experienced an adverse or unanticipated event related to their study participation.

Additionally, we evaluated immune profiles of AAT in both human and murine tissue samples to quantify the presence of T cells, B cells, dendritic cells, macrophages, M1- polarized (inflammatory) macrophages, and M2-polarized (wound-healing) macrophages.

Our clinical results indicate that the populations of immune cells migrating into AAT from the surrounding tissue match those of native adipose in terms of cell type and gene expression. For some patients at early excision times, AAT was associated with pro- regenerative immune responses, including M2 macrophage polarization and increased IL-4 expression. Addition in vivo studies investigated the local immunological microenvironment

ii created by AAT in both non-traumatic and injured environments, as the ability to modulate immune response in a targeted approach may be used to improve wound healing. These studies also aimed to characterize the effect of xenogeneic ECM vs. syngeneic ECM in a mouse model of volumetric muscle loss (VML).

Overall, our preclinical and clinical findings show that AAT is a promising new therapeutic tool for soft tissue reconstruction and is safe for subcutaneous use in humans.

Further studies, including a Phase II trial, are necessary to determine clinical efficacy.

Advisor: Jennifer H. Elisseeff, Ph.D.

Thesis Committee: Damon S. Cooney, M.D., Ph.D. (Reader)

Jeff W. M. Bulte, Ph.D. (Chair)

William Guang Wong, Ph.D.

Drew Pardoll, Ph.D.

Patrick Byrne, M.D.

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ACKNOWLEDGEMENTS

I would like to acknowledge the many individuals who helped make this work possible. The members of the Elisseeff lab who provided intellectual and technical support were invaluable both in my personal development and in the success of this work. Dr. Zayna Nahas and Dr.

Iwen Wu pioneered much of the early process development and preclinical studies that would become the foundation of my thesis and enable us to obtain approval to begin human testing. Jessica Yang also helped introduce me to the adipose ECM project and provided much of my initial training. I am deeply thankful for their contributions (and meticulous record keeping), without which this project would never have existed. I also would like to acknowledge the people who helped make the clinical trial possible, including our team of physicians (Drs. Damon Cooney and Patrick Byrne), our trial manager (Carisa Cooney), our regulatory and manufacturing consultant (Susan Wade), and clinical fellows (Rachael Payne and Jeff Aston).

My advisor, Dr. Jennifer Elisseeff, has been a constant source of inspiration. My appreciation of her incredible work ethic and breadth of knowledge has grown continually over the past six years, because it is hard to imagine anyone more ambitious or dedicated to growing her ecosystem. Her direction of this project and help thinking through the tough questions has been invaluable – when I am hung up on details, she can always help me see the big picture. While setting a high bar as an investigator, Dr. Elisseeff still manages to be a kind and flexible supervisor who prioritizes the career goals of her students. It is uncommon to find all of these qualities in a mentor, and I am eternally grateful for the opportunities she has given me.

I worked alongside many fantastic people during my years in the Elisseeff laboratory; however, one person’s contribution to my work stands well above all the others. Alexis

Parrillo was my close partner and a dear friend throughout these studies and shares in every success. Whether we were disemboweling mice, wading through buckets of human fat or simply thinking though yet another experiment together, Alexis never failed to brighten my day and always helped turn daunting tasks into new achievements.

Lastly, I am grateful for the support of my friends and family, without whom I would not be where (or who) I am today. I could not have asked for a better group of people to share the graduate school experience with than my CMM classmates. I would like to thank my parents, Lisa and Tim Anderson, and sister, Kelsey, for their unwavering faith in my decisions, even when my choices took me far from home. Finally, I want to thank Ashkon – my best friend and partner in everything (outside the lab) – for being my imperturbable rock and for following me across the country to be a part of this adventure with me.

TABLE OF CONTENTS

ABSTRACT ...... ii ACKNOWLEDGEMENTS ...... iv TABLE OF CONTENTS ...... vi LIST OF FIGURES ...... viii LIST OF TABLES ...... ix 1. INTRODUCTION ...... 1 2. BIOCHEMICAL CHARACTERIZATIONS OF ACELLULAR ADIPOSE TISSUE (AAT) – AN ADIPOSE EXTRACELLULAR MATRIX-DERIVED BIOMATERIAL ...... 9 2.1. Introduction ...... 9 2.2. Materials and Methods ...... 13 2.2.1. Adipose processing and AAT manufacturing ...... 13 2.2.2. Collagen content ...... 14 2.2.3. SDS-PAGE ...... 14 2.2.4. Lipid content ...... 15 2.2.5. Cell migration ...... 15 2.2.6. Residuals testing ...... 17 2.2.7. Statistical analyses ...... 18 2.3. Results ...... 18 2.3.1. Adipose processing and AAT manufacturing ...... 18 2.3.2. Collagen content ...... 19 2.3.3. SDS-PAGE ...... 20 2.3.4. Lipid content ...... 21 2.3.5. Cell migration ...... 22 2.3.6. Residuals testing ...... 23 2.4. Discussion ...... 24 3. IMMUNE CHARACTERIZATION OF AAT IN MURINE MODELS OF WOUND HEALING ...... 31 3.1. Introduction ...... 31 3.2. Materials and Methods ...... 33 3.2.1. Subcutaneous injection of AAT in mice ...... 33 3.2.2. Volumetric muscle loss (VML) surgery in mice ...... 33 3.2.3. Histology ...... 34 3.2.4. Flow cytometry ...... 34

3.2.5. Real time quantitative polymerase chain reaction (RT-qPCR) ...... 36 3.2.6. Statistical analyses ...... 37 3.3. Results ...... 37 3.3.1. Subcutaneous injections of AAT in mice ...... 37 3.3.2. Volumetric muscle loss (VML) surgery in mice ...... 41 3.4. Discussion ...... 44 4. PHASE 1 CLINICAL TESTING OF AAT – SAFETY, TOLERABILITY, AND EXPLORATORY FINDINGS ...... 47 4.1. Introduction ...... 47 4.2. Materials and Methods ...... 50 4.2.1. Subcutaneous injection of AAT in healthy human volunteers ...... 50 4.2.2. Follow-up visits ...... 51 4.2.3. AAT excision...... 51 4.2.4. Safety and tolerability ...... 52 4.2.5. Photoimaging ...... 52 4.2.6. Surveys ...... 53 4.2.7. Histopathology ...... 53 4.2.8. Immunostaining ...... 53 4.2.9. Flow cytometry ...... 54 4.2.10. Statistical analyses ...... 55 4.3. Results ...... 55 4.3.1. Enrollment, demographics, and dosage ...... 55 4.3.2. Adverse Events (AEs) ...... 56 4.3.3. Panel Reactive Antibody (PRA) testing ...... 58 4.3.4. Photoimaging ...... 59 4.3.5. Surveys ...... 60 4.3.6. Histopathology and immunostaining ...... 62 4.3.7. Flow cytometry ...... 65 4.4. Discussion ...... 70 5. CONCLUSIONS AND FUTURE WORK ...... 74 6. REFERENCES ...... 77 7. CURRICULUM VITAE ...... 80

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LIST OF FIGURES

Figure 2.1. Hydroxyproline content of irradiated and non-irradiated AAT ...... 20

Figure 2.2. SDS-PAGE analysis performed on samples digested for 5 hours and 24 hours ...... 21

Figure 2.3. Lipid content of human and porcine AAT batches ...... 22

Figure 2.4. Migration assay results for the clinical batch of human AAT ...... 23

Figure 2.5. Assays to test the human clinical AAT batch for residual process chemicals ...... 24

Figure 3.1. Hematoxylin and eosin staining of subcutaneous human and porcine AAT implants ...... 38

Figure 3.2. Subcutaneous flow cytometry results at 1 and 3 weeks ...... 39

Figure 3.3. Gene expression in subcutaneous AAT at 3 weeks ...... 40

Figure 3.4. Volumetric muscle loss flow cytometry results at 1 week ...... 42

Figure 4.1. Injection site photos at time of excision ...... 59

Figure 4.2. Participant and physician surveys ...... 61

Figure 4.3. Histology (H&E) by individual participant ...... 63

Figure 4.4. Cell migration over time and revascularization at 18 weeks ...... 64

Figure 4.5. Flow cytometry gating strategy for myeloid populations and macrophage polarization. . 66

Figure 4.6. Macrophage polarization in AAT implants and untreated distal fat ...... 68

Figure 4.7. Myeloid populations in AAT implants versus untreated distal fat ...... 69

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LIST OF TABLES

Table 2.1. AAT batches assessed with biochemical assays...... 19

Table 3.1. Mouse flow cytometry panel for subcutaneous injection studies ...... 35

Table 3.2. Mouse flow cytometry panel for volumetric muscle loss studies ...... 35

Table 3.3. Real time quantitative PCR Primer Sequences ...... 36

Table 4.1. Human flow cytometry panel for myeloid cells ...... 54

Table 4.2. Overview of Study Participants ...... 56

Table 4.3. AAT Exposure Duration and Dosage ...... 56

Table 4.4. Anticipated Adverse Events Related to Injection Sites ...... 57

Table 4.5. Panel Reactive Antibody Results ...... 58

1. INTRODUCTION

Soft tissue defects are relatively common and can occur due to trauma, congenital disease or co-morbid conditions (i.e., HIV/AIDS), or surgical inventions (i.e., lumpectomy). As tissue loss is often permanent, these defects can impact not only cosmesis but also normal physiological function, including lack of support for distal extremities and contracture leading to restricted range of motion [1]. The gold standard for soft tissue reconstruction is autologous adipose transfer (otherwise known as fat grafting), a procedure first developed by

Gustav Neuber over a century ago [2, 3]. However, although these procedures are still commonly used today and many advances have been introduced, autologous fat transfer techniques have many limitations. Adipose grafts behave unpredictably and outcomes can often vary significantly from patient to patient. Clinically, volume loss following autologous fat transfer has been reported to be between 40-60% and usually occurs within the first 4-6 months [4]. Re-grafting is often needed and implanted adipose tissue frequently leads to post- operative calcifications [5]. The unpredictability of these procedures can result in costly secondary surgeries and co-morbidities related to tissue harvest. Harvest procedures often lead to scarring at the donor site and are limited by the volume of autologous tissue available in each patient [6]. Transplanted adipocytes are often damaged or subjected to hypoxic

1 conditions that result in the release of intracellular lipids, a potent pro-inflammatory signal.

These signals combined with a lack of vascularization within the graft can lead to tissue necrosis and calcifications which impact the quality and durability of the reconstructed tissue

[7]. Deformities are often tolerated or overlooked since the complexity of these repair techniques is often greater than the anticipated benefits.

The potential for new soft tissue augmentation technologies cuts through numerous markets in plastic surgery, dermatology, oral and maxillofacial surgery and general traumatic and reconstructive surgery. The US market size for these applications is significant with the cosmetic market alone reaching as much as $13 billion annually. The tissue filler market for example is close to $2 billion and is expected to grow at a rate of 8.4 percent annually through 2020. Other reconstruction applications that have significant markets include: lumpectomy (100,000 procedures per year), craniofacial reconstruction (200,000 per year), and HIV lipodystrophy (350,000 per year). However, these market numbers likely underestimate the true clinical need due to current limitations in the supply of human tissue and manual processing methods that are currently limiting the scale up potential of tissue- derived biomaterials.

Decellularized extracellular matrix (ECM) products are a type of biomaterial that have recently gained popularity in the field of regenerative medicine, although they have been studied and used in various clinical applications since as early as 1995 [8]. Unlike synthetic biomaterials such as polymers, ECM materials are tissue-derived and are created using physical, enzymatic, and chemical approaches that remove the living cells from almost any

2 type of animal or human tissue [9]. These decellularization processes disrupt cellular membranes and denature key intracellular components such as DNA, but leave behind the non-cellular component which is present in all tissues and organs. This non-cellular component is mainly structural in nature and is known as the extracellular matrix (ECM).

Although all ECMs are composed of water, proteins, and polysaccharides, the physical characteristics and exact composition of any specific ECM depends on its tissue of origin.

ECMs have many physiological roles, including providing a physical scaffold and initiating cues required for tissue homeostasis and differentiation. ECM also directs function by binding to growth factors and interacting with receptors on the cell surface [10].

ECM scaffolds are commonly used in tissue engineering to promote the healing or regrowth of damaged tissue. Initially, they provide a physical substrate upon which the cells can be seeded and localized to a specific area. They also provide key biochemical and physical cues for adhesion, migration, proliferation, and differentiation which help cells to form fully functional tissues or organs. Implanted ECM scaffolds will eventually be remodeled and be replaced with the seeded cells’ own secreted matrices.

ECM has many beneficial uses and new applications that are currently being explored in regenerative medicine. In the past 15 years, many ECM materials have been brought to market and been successfully used clinically. Alloderm® (LifeCell Corp., NJ), an ECM product prepared from cadaveric human skin, has been used for dentistry, burn therapy, plastic surgery, and hernia repair for over 13 years. The wide utility of acellular dermis is also evidenced by its use in abdominal hernia repair and skin grafting procedures; however,

3 the form of this substance (sheets ranging from 0.23-3.3mm in thickness) limits its use in soft tissue defect reconstruction. A micronized form of AlloDerm® does exist (Cymetra®MAT,

LifeCell Corp.); however, it does not have the longevity of the sheet form and is degraded by the host over time [11]. Many other decellularized allografts have also been successfully marketed and used clinically for reconstruction. In one case, a complex scaffold made of porcine small intestinal sub-mucosa ECM was used to treat a quadriceps defect in a 19-year- old marine three years post-injury, resulting in remarkable improvement after only 4 months

[12].

The complex protein and polysaccharide composition and unique physical structures of an

ECM cannot be mimicked using any synthetic biomaterial currently available, and thus these biomimetic scaffolds effectively modulate signal transduction and both directly and indirectly regulate cellular function similar to natural ECM [13]. The remarkable successes of ECM products already on the market and the potential benefits of ECM materials due to their biomimetic properties indicate that the possible clinical applications of these materials are vast. However, no adipose ECM product has yet been introduced into clinical use for correcting soft tissue defects.

Adipose tissue is where the body stores excess energy and regulates metabolic homeostasis by synthesizing and secreting various compounds. It is made up mostly of adipocytes, but also consists of blood cells, endothelial cells, adipose precursor cells, pericytes, and other cells in the stromal vascular fraction. Adipocytes can increase primarily in size but also in number in order to store excess energy produced from food [14]. Adipose is also a robust

4 source of mesenchymal stem cells, called adipose-derived stem cells (ASCs), which have the potential to differentiate into multiple lineages [15].

As per their role as energy suppliers for the body, adipose cell signaling can have a significant impact on overall health. It has been noted clinically that transplanting autologous fat can have a positive impact on surrounding tissues. This includes improvements in both scarring and aging skin. Perhaps more strikingly, autologous fat transfers have also improved radiation damage, damaged vocal cords, and chronic ulceration. These improvements may be related to undifferentiated cells, such as ASCs, in the adipose tissue [16]. These clinical observations indicate an important connection between adipose tissue and wound healing.

Adipose tissue is a practical raw material source for producing ECM biomaterials for several reasons. To start, adipose is relatively abundant and easy to harvest, whether from a deceased tissue donor or in a minimally invasive procedure for autologous use. An adult human can have a body fat composition of anywhere from less than 10% (a lean individual) to more than 50% (an obese individual) [17]. Additionally, adipose contains secreted factors that are beneficial for angiogenesis, anti-inflammation, and anti-apoptosis [18, 19].

Importantly, adipose tissue can also play a role in immune modulation. The metabolic processes carried out by adipose tissue, including adipocyte expansion and thermogenesis, can activate both the adaptive and innate immune system [20]. All of these qualities of adipose tissue make it an ideal candidate for a decellularized ECM biomaterial for use in wound healing and reconstruction applications.

An adipose-derived extracellular matrix presents a promising solution for repair of soft tissue defects. From the physical perspective adipose tissue (natural or processed) has unique and ideal mechanical properties for soft tissue reconstruction, characterized by a compliant and highly elastic nature. Compared to dermis, adipose ECM is more elastic, softer and have a molecular blueprint that offers an allogeneic scaffold, which can facilitate host tissue regeneration without eliciting immune rejection. The components of the adipose-derived scaffold are all naturally occurring adipose tissue proteins and proteoglycans that are highly conserved, even between different species. This means cells are not exposed to any foreign materials or synthetic polymers that can induce a foreign body response and subsequent fibrous capsule formation. This also enhances host tissue integration since matrix components can be easily degraded by cell-secreted enzymes as tissue remodeling takes place [21-23]. Mechanisms for matrix turnover are already established in host cells, avoiding any concerns over proper clearance of scaffold materials from the body. These factors contribute to the biocompatibility of ECM-based scaffolds and highlight their utility in regenerative medicine.

When addressing the challenges of wound healing and reconstruction, it is also important to understand the role of the immune system. Immune cells are important in every step of the wound healing process, from debridement to new tissue and scar formation. Neutrophils are the first responders arriving at the wound site approximately 24 hours after injury. Their main role is to debride the wound and decrease the likelihood of infection. Approximately

48-96 hours after injury, macrophages migrate in and promptly become the predominant cell population. They contribute to and conclude wound debridement and secrete cytokines and

6 growth factors that play a key role in cell recruitment and regulation during tissue repair, including both angiogenesis and new matrix deposition. On approximately the fifth day following injury, T lymphocytes migrate into the wound and regulate the proliferation phase of tissue repair [24].

As the immune system’s contribution to and regulation of tissue development and regeneration is becoming better understood [25], the field of tissue engineering is beginning to approach regeneration with the immune system in mind. Recent studies have identified the role of Type 2 helper T cells (TH2) in the biomaterial scaffold directed tissue repair [26] and examined the role of macrophages in the remodeling process after implantation of a surgical mesh [27]. Macrophages play a key role in wound healing; these cells are highly plastic and exist on a phenotypic spectrum ranging from “M1” macrophages (typically described as pro-inflammatory or classically activated) to “M2” macrophages (which are considered regulatory or homeostatic) [27]. Macrophage heterogeneity [28] and its implications for wound healing [29] can have a huge impact on the design of biomaterials to elicit pro-regenerative responses.

The work in this thesis describes the immunological characterization and Phase I clinical testing of an adipose extracellular matrix product, called Acellular Adipose Tissue (AAT).

This product was developed at Johns Hopkins University in the laboratory of Dr. Jennifer

Elisseeff and is intended to fill the clinical need for an "off-the-shelf" soft tissue repair technology for volume augmentation and soft tissue reconstruction. AAT provides a structure that mimics normal soft tissue and a matrix to promote cell migration and new fat tissue

7 formation. Extensive preclinical studies have characterized the physical properties and evaluated compatibility and efficacy in vivo [30, 31], including an evaluation of new adipose tissue development in athymic mice and biocompatibility in immune competent rodents. The in vivo behavior of the AAT was evaluated in multiple animal models including mouse, rat, and swine in comparison to the current clinical gold standard of autologous fat grafting.

Overall, these results highlight the biocompatibility of the AAT implants and their ability to provide soft tissue volume replacement. They also show an advantage autologous fat grafting which may cause calcification due to an inflammatory response to released intracellular lipids [3, 5].

2. BIOCHEMICAL CHARACTERIZATIONS OF ACELLULAR

ADIPOSE TISSUE (AAT) – AN ADIPOSE EXTRACELLULAR

MATRIX-DERIVED BIOMATERIAL

2.1. Introduction

Adipose tissue is commonly used by surgeons for a variety of applications, including those related to soft tissue reconstruction and wound healing [32, 33]. However, harvesting tissue from each patient presents a source of patient morbidity, outcome variability, and oftentimes inadequate volumes to address large defects. To address this clinical challenge, we aimed to create an "off-the-shelf" adipose material using mechanical and chemical processing techniques. Techniques were developed to remove lipids (a source of inflammation) and living cells while processing the tissue into an injectable form preferred by patients and physicians.

Acellular Adipose Tissue (AAT) was first developed in the Biomaterials and Tissue

Engineering Laboratory in the Translational Tissue Engineering Center (TTEC) at the Johns

Hopkins School of Medicine (JHM). It is now being further translated into clinical testing through a three way collaboration between Aegeria Soft Tissue (AST), Johns Hopkins

University (JHU), and the Armed Forces Institute of Regenerative Medicine (AFIRM) – II,

U.S. Army Medical Research and Materiel Command (USAMRMC). AST licensed the technology from JHU and developed the manufacturing and quality control systems required for GMP production and the clinical testing. Through this collaborative effort in translational medicine, the parties hope to apply the basic research from the laboratories of Jennifer H.

Elisseeff, Ph.D., to improve treatments for traumatic soft tissue injuries. AST’s adipose

ECM technology is expected to fill the significant clinical need for a simple and effective off-the-shelf soft tissue repair technology for small and large volume augmentation.

AAT is derived from cadaveric adipose tissue that is processed using mechanical and chemical methods to remove lipids and living cells. Adipose tissue is rinsed in peracetic acid, detergent, and Dulbecco's phosphate buffered saline (DPBS), then the resulting decellularized extracellular matrix is snap frozen in liquid nitrogen and mechanically processed using a knife mill. The tissue is then re-hydrated with saline and packaged into sterile syringes. The tissue processing is performed in Class A biosafety cabinets using aseptic processing techniques. For clinical use, the final packaged product is also gamma irradiated at 15-18 kGy and released in accordance with ISO 11137. AAT is composed primarily of structural proteins and extracellular matrix components derived from adipose tissue. As such, AAT provides a natural structure to mimic normal soft tissue in a sterile preparation and injectable format preferred by patients and physicians.

Early development of AAT (prior to the work outlined in this thesis) focused on testing various chemicals and processing methods. The goal was to balance mild, biocompatible

10 reagents and processing techniques with adequate removal of lipids. Delipidizing reagents were selected to minimize disruption of the tissue matrix and maintain biocompatibility, while manual mechanical processing methods were used to render the decellularized adipose tissue into an injectable product without altering the proteomic composition or biocompatibility of AAT. Initial process development research resulted in a lab-scale protocol that used mild reagents with manual mechanical processing. Later research focused on scale-up of manual techniques such that GMP processing could be performed for pilot clinical testing, including validation of different milling methods to replace manual processing.

After defining the first AAT processing method at lab-scale, the characteristics of the resulting AAT were monitored with respect to structural (DSC) and mechanical properties

(extrusion from syringe, rheology), morphology (SEM), and protein composition

(proteomics). As we moved towards scale-up, these product characteristics were continually evaluated to ensure that updated processing methods produced a consistent biomaterial which achieved our goal of an injectable, off-the-shelf soft tissue-replacement material with minimal lipids. The preclinical data provided in our original IND filing (approved on July

24th, 2015) confirm that:

(1) Lipids can be removed from adipose tissue without altering the basic extracellular

matrix organization.

(2) Ultrastructure of tissue is maintained during processing and after cutting into smaller

pieces.

(3) Rheological (mechanical) properties of the processed adipose are nearly identical to

lipoaspirate.

(4) Thermal properties of the adipose tissue shift significantly when lipids are removed

but further processing and sterilization produces minimal changes, thus suggesting

minimal changes to the AAT structure.

(5) Proteomic analyses of the extracellular matrix proteins reveal a complex mixture of

structural proteins typical of adipose-derived extracellular matrix tissue, particularly

collagen family proteins.

(6) AAT promotes migration of adipose-derived stem cells (ASCs) into the tissue matrix

in vitro.

Building on early development work done in the Elisseeff lab which characterized the physical properties of AAT, the studies outlined in this chapter sought to define the biochemical characteristics of AAT prior to Phase I clinical testing. Biochemical assays performed included an in vitro cell migration assay, residual chemicals testing, a lipid content assay, and a collagen content assay.

There were two main purposes for collecting this data: to get a better understanding of the biochemical composition and properties of the material, and to start building a database with the goal of understanding the batch–to–batch differences in AAT. This information will help define the expected variability from both tissue donors and from any changes in the manufacturing process, and will be critical for scaling up the manufacturing protocols for later stage clinical trials and eventual commercialization. The biochemical characterizations

12 also allowed us to study how the terminal sterilization process of gamma irradiation might change the properties of the final product.

2.2. Materials and Methods

2.2.1. Adipose processing and AAT manufacturing

To manufacture the product, adipose tissue is processed using chemical and mechanical techniques. Cadaveric human adipose tissue is acquired from a tissue bank or OPO. The bulk adipose tissue is cut into approximately 1-square inch blocks and stored frozen at less than -20°C. Frozen tissue is thawed and warmed to approximately 37°C for processing. The majority of the lipids/oils are removed from the tissue by processing using a mechanical press. The pressed tissue is washed in room temperature Dulbecco's modified Phosphate

Buffered Saline solution (DPBS) and collected in a mesh screen to concentrate. The tissue is then treated with peracetic acid for at least three hours at room temperature. After rinsing with DPBS/HEPES buffer, the mechanically pressed concentrate is stirred in an

EDTA/Triton-X detergent solution overnight. The tissue is rinsed repeatedly with DPBS buffer all visible trace of detergent or bubbles is removed, after which the tissue is rinsed an additional three times to remove any residual detergent and return the tissue to a neutral pH.

Finally, the pH of the rinsed product is re-confirmed to be between 6.0 - 7.0. After a final concentration in a mesh screen, the resulting decellularized extracellular matrix is flash frozen in liquid nitrogen and homogenized using a knife mill. The milled tissue is collected in a single container and adjusted to 91% moisture (approximately 9% human adipose extracellular matrix by weight) as measured with an infrared moisture analyzer. The re-

13 hydrated tissue is packaged into syringes. The tissue processing is performed in Class A manufacturing areas using aseptic processing techniques. Packaged product is gamma irradiated at 15-18 kGy and released in accordance with ISO 11137.

2.2.2. Collagen content

Irradiated and non-irradiated AAT from the demonstration batch (Lot TS-0240) were assayed for hydroxyproline content according to an SOP derived from the manufacturer's protocol

(Hydroxyproline Assay Kit, Sigma-Aldrich). A collagen control from bovine Achilles tendon (Sigma-Aldrich) was used to calculate the conversion factor between hydroxyproline and collagen. Briefly, AAT samples were lyophilized prior to the assay to eliminate any variation in water content resulting from syringe extrusion. Following acid-hydrolysis of all samples in 6 M HCl (120°C for 3 hours), samples were diluted, transferred to wells, vacuum dried, then incubated at 60°C in assay reagents prior to reading absorbance at 560 nm. To eliminate any effect from endogenous interfering compounds, a spiked sample control was included. All samples were assayed at four concentrations in triplicate to ensure absorbance measurements fell within the linear range of the assay's standard curve and to calculate the mean relative hydroxyproline and collagen content of each sample.

2.2.3. SDS-PAGE

AAT samples were digested using collagenase (Collagenase CLSPA, Worthington

Biochemical) in order to solubilize the proteinaceous matrix. Digestion mixtures were prepared by combining 100 mg of each AAT sample and either 10 units or 20 units of collagenase enzyme in a total volume of 1 ml DPBS buffer containing calcium and magnesium (Quality Biologic). Samples were incubated at 37°C for 5 hours or 24 hours.

Following incubation, samples were centrifuged to remove insoluble AAT fragments.

Protein samples were then boiled with Laemmli sample buffer (Bio-Rad) containing β- mercaptoethanol, loaded on Criterion 4-20% Tris-HCl gels (Bio-Rad) and run for 50 minutes at 200 volts on a Criterion Cell (Bio-Rad). Protein bands were visualized with a Coomassie blue reagent (SimplyBlue SafeStain, Life Technologies).

2.2.4. Lipid content

The total lipid content of biomaterials was quantified using a triglyceride colorimetric assay.

To extract lipids, adipose samples and AAT were minced into 1 mm or smaller pieces to disrupt physical barriers and release lipids. Organic extraction was conducted by the

Schwartz method for detection of triacylglycerol in cells and tissues. The triglyceride colorimetric assay uses enzymatic hydrolysis of the triglycerides by lipase to glycerol and free fatty acids. All enzymatic reactions were carried out using Infinity TG Reagent (Thermo

Fisher Scientific) according to the manufacturer's protocol. The glycerol released was subsequently measured by a coupled enzymatic reaction system producing a brilliant purple color. The absorbance was measured at 540nm. The concentration was determined using a glycerol standard curve and reported as a weight percentage of the product or a control sample (such as native human adipose tissue).

2.2.5. Cell migration

This assay measured the migration of human adipose-derived stem cells (ASCs) through a membrane in response to molecular signals. Serum-free media was used as the negative control and media with 10% fetal bovine serum (FBS) was used as the positive control. The ability of AAT to signal the ASCs to migrate was measured by counting the cells which pass

15 through the membrane. The result was expressed as a percentage of cell migration relative to the controls.

Human adipose-derived stem cells (ASCs) were thawed and grown in basic growth media supplemented with 1 ng/mL basic fibroblastic growth factor (bFGF), passaged and split once at approximately 80% confluency, with media changes every 2-3 days. Upon reaching 80% confluency a second time, cells were serum starved for 24 hours. After starvation, cells were trypsinized and a single cell suspension was created (300,000 cells/mL) in serum free media.

Samples were prepared by adding AAT to serum free media at a 1% (v/v) concentration, vortexing, incubating at room temperature for 15 minutes, and centrifuging to remove large chunks of AAT that might stick to the transwell membrane. Test sample(s), 10% FBS, 1%

PBS and serum free media (600 µl) are added in triplicate wells of a 24 well plate, a transwell insert is placed on top, and the plate is allowed to acclimate in an incubator. After acclimation, 100 μL of the cell suspension is added into the upper chamber of each transwell

(300,000 cells per well) and incubated at 37˚C for 6 hours. All cells remaining on the upper side of the membrane are removed using a cotton swab and any excess AAT is removed from the lower membrane by rinsing in DPBS. Transwells were fixed in methanol for 15 minutes at room temperature, stained with DAPI, and then imaged within 72 hours. Cell nuclei are counted in a 50x field of view. Cell migration is calculated relative to positive and negative control groups.

2.2.6. Residuals testing

A semi-quantitative method was developed using Quantofix EDTA test strips and validated using standard solutions of known EDTA concentration. A control batch of AAT was manufactured without EDTA to serve as a negative control during validation. To perform the test, a known concentration of EDTA is spiked into the control batch of AAT to ensure that none of the components of the AAT would interfere with the validity of the test strips.

Samples are centrifuged and supernatant is collected and applied to the test strips. Strips are read according to the manufacturer's instructions.

Residual Triton X-100 levels in AAT are quantified by reverse phase high-performance liquid chromatography (HPLC), with a control batch (manufactured without Triton X-100) serving as a negative control in these experiments. Triton X-100 standards are prepared in water and run along with the AAT samples. AAT and spiked control samples are prepared for HPLC by repeated centrifugation at >12,000 rpm to remove insoluble proteins and collect aqueous supernatant. Soluble proteins are then precipitated using methanol-chloroform extraction. Both the aqueous and organic layers are collected and combined. The protein pellet is washed with chloroform and the supernatant is also combined with the sample

(discarding the protein pellet). Samples are concentrated by freeze-drying in a lyophilizer until completely dry, then resuspended in a consistent volume of pure water for HPLC. An isocratic reverse phase separation is performed using an HC-C18(2) column and two mobile phases: HPLC-grade water and 100% acetonitrile. The peak corresponding to Triton X-100 is measured and quantified relative to standards.

2.2.7. Statistical analyses

Statistical analysis was performed using GraphPad Prism software. In grouped analyses with a single variable, significance was determined by one-way analysis of variance (ANOVA) using the Holm-Sidak correction for multiple comparisons where applicable (α = 0.05).

Significance in grouped analyses with two variables was calculated using two-way ANOVA with Tukey post-hoc testing (α = 0.05). P values less than 0.05 were considered statistically significant (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001). Plotted values represent the arithmetic or geometric mean (RT-qPCR data only) of the data set. Error bars represent +/- one standard deviation or geometric standard deviation (RT-qPCR only).

2.3. Results

2.3.1. Adipose processing and AAT manufacturing

For the biochemical assays described in this chapter, several lots of human adipose-derived

AAT (hAAT) and one lot of porcine adipose-derived AAT (pAAT) were assessed. Many biochemical assays were developed and tested on an AAT demonstration lot (TS-0240) produced from human donor tissue under GMP conditions. This batch was manufactured according to the proposed clinical standards in the facility which would also manufacture clinical supplies of AAT; analytical data obtained from this batch was used to determine product specifications and was submitted to the FDA to secure regulatory approval for Phase

I testing. After IND approval, a second batch of clinical-grade hAAT was manufactured in

October 2015 for Phase I clinical testing (TS-0268); this lot was assessed biochemically to gather batch information and demonstrate compliance with product specifications, including

18 cell migration, lipid content, levels of residual chemicals, and other qualitative assessments.

Additional non-clinical batches were manufactured in the laboratory (non-GMP facility) according to Good Laboratory Practices for later use in animal studies (hAAT: TS-0267, pAAT: PA1). Unlike the clinical lot, the two non-GMP batches used in animal testing were not terminally sterilized by gamma irradiation, though sterility was carefully maintained during tissue processing. A summary of the AAT batches described in this chapter is included in Table 2.1 below.

Table 2.1. AAT batches assessed with biochemical assays Species Batch/lot # Manufacturing Sterilization Description

Human TS-0240 GMP Gamma- GMP demonstration batch; utilized irradiated for assay development and defining

product specifications Human TS-0670 Lab None Lab-scale batch; used for animal studies

Human TS-0680 GMP Gamma- GMP clinical batch; tested and irradiated released for use in Phase 1 clinical

testing Pig PA1 Lab None Lab-scale batch; used for animal studies Pig PA2 Lab, with None Control batch manufactured without modified process EDTA and Triton X-100; used to develop assays for residual process chemicals

2.3.2. Collagen content

Irradiated and non-irradiated AAT from the demonstration batch (TS-0240) were assayed for hydroxyproline content, along with a pure collagen control. This assay showed that the irradiated hAAT is 87.0 ± 5.1% collagen by dry weight and non-irradiated hAAT is 89.0 ±

15.1% collagen by dry weight (Figure 1). Bovine collagen was used as the positive control to validate the assay and was measured to be 100.6 ± 9.1 % collagen by dry weight (Figure

2.1). These results are consistent with the measurements found in the literature and are expected due to the relatively high collagen content of most native ECMs.

Figure 2.1. Hydroxyproline content of irradiated and non-irradiated AAT. Collagen content of AAT samples was determined using a conversion factor between hydroxyproline and collagen calculated from the bovine collagen control. Error bars represent standard deviation from the mean.

2.3.3. SDS-PAGE

Irradiated and non-irradiated samples from the AAT demonstration batch (TS-0240) were digested with a purified bacterial collagenase and the resulting supernatants were analyzed by a gel electrophoresis method using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), a discontinuous polyacrylamide gel support medium containing sodium dodecyl sulfate (SDS) to denature the proteins. The conditions tested did not reveal visual differences in the distribution or abundance of proteins solubilized by collagenase in the irradiated and non-irradiated samples. Increasing incubation time over 5 hours did not result in greater sample digestion or change the resulting protein distribution or abundance (Figure 2.2).

Figure 2.2. SDS-PAGE analysis performed on samples digested for 5 hours and 24 hours. Each sample was treated with two different enzyme concentrations (10 U and 20 U per 100 mg sample of AAT). Control lanes contain only the collagenase enzyme, loaded with equal amount of enzyme as samples digested in 10 U of total enzyme. Blank lane(s) contain ultrapure water in place of digested sample.

2.3.4. Lipid content

Triglycerides present before and after adipose tissue processing were extracted and quantified to determine the fraction of total lipids removed during manufacturing (for the purposes of this assay, unprocessed adipose from the demonstration batch TS-0240 was used to calculate the initial glycerol concentration for all human batches, as unprocessed human adipose was not available from TS-0267 and TS-0268). Glycerol concentrations less than

0.41 umol/mL of TG reagent produced a linear standard curve which was used to calculate the total glycerol content of the sample (equivalent to triglyceride content). In both pig and human derived AAT batches, relative glycerol content was less than 2 μg/mg of sample, corresponding to > 97% lipid removal in all batches, whether lab-manufactured or GMP- manufactured (Figure 2.3).

Figure 2.3. Lipid content of human and porcine AAT batches. Glycerol content of a human lab batch (TS-0267), a pig lab batch (PA1), and two GMP human batches (TS-0240 and clinical batch TS-0268) were compared to the glycerol content of the corresponding native adipose to determine the percentage of lipid removal during manufacturing.

2.3.5. Cell migration

An in vitro cell migration assay was conducted on both irradiated and non-irradiated samples from the human clinical batch (TS-0268). In a pilot experiment (TS-0240), images taken of the transwell membrane showed that the non-irradiated AAT was more likely to stick to the membrane, potentially indicating a difference in mechanical properties. A second assay was performed with the samples centrifuged to remove large chunks that might stick to membrane, thus ensuring migration would only be triggered by interaction with soluble factors. In this second assay, the non-irradiated AAT resulted in somewhat less cell migration than the irradiated batch (38.94% and 51.87% respectively). However, this result was not statistically significant. Both AAT samples promoted significantly more migration than the negative and buffer controls (Figure 2.4). Relative cell migration was within the expected range of the product specification (20% - 70%).

Figure 2.4. Migration assay results for the clinical batch of human AAT. Microscope images show differential stickiness, with clumps of non-irradiated AAT stuck to the membrane (arrows), but none for irradiated AAT. Only significance relative to serum free condition is shown (one-way ANOVA).

2.3.6. Residuals testing

Residual chemical testing was performed on the clinical batch of human AAT and a negative control batch manufactured without the chemicals of interest. Both EDTA and Triton X-100 were found to be well within acceptable safety limits for the intended AAT dosage range in humans. Both test methods were validated using known standards and a spiked control to ensure that none of the components of the AAT would interfere with the validity of the test strips or HPLC method. The results of both validation measures are shown in Figure 2.5.

Quantofix EDTA test strips indicated that there was no detectable residual EDTA remaining in the final clinical product. Triton X-100 levels were, on average, 52.524 μg/ml of AAT after processing, corresponding to a 99.50% removal of Triton X-100.

Figure 2.5. Assays to test the human clinical AAT batch for residual process chemicals. Quantofix EDTA test strips were used to test for EDTA and HPLC was used to test for Triton X-100. A batch of porcine AAT was manufactured without the chemicals of interest to serve as a negative and spiked control in these experiments.

2.4. Discussion

AAT is similar to a number of tissue transplant products and is processed according to Good

Tissue Practices and Good Manufacturing Practices. Early development work (prior to this thesis) defined the AAT manufacturing process and identified mild chemical reagents which would minimize any in situ reaction to implanted material and preserve the structure of adipose ECM proteins. Mechanical processing methods used to render the decellularized adipose tissue into an injectable product were also validated, ensuring that the proteomic composition and biocompatibility of AAT remained consistent during initial process scale up. Preclinical animal studies also defined the behavior of AAT in vivo, such as its ability to recruit and differentiate adipose stem cells to form new adipose tissue and retain volume over time. These properties suggested a clinical indication for soft tissue replacement and informed many aspects of clinical development.

Though this product has been classified as a biologic by the FDA, AAT differs significantly from classic drugs or biologics. Traditional pharmacology and toxicology testing are not appropriate for this product because there is no identified pharmacological activity. The processed ECM proteins which make up the solid bulk of AAT may provide some local signaling which induces colonization of the material by the patient’s own adipocytes, but the mechanisms are unknown and likely complex. While AAT is a potentially bioactive material derived from natural tissues, it is primarily intended for soft tissue volume replacement and thus also resembles a medical device.

Lack of a well-defined pharmacological target or activity presents significant challenges in translating AAT to clinical use. The AAT drug substance had to be defined indirectly by characterizing multiple physical and biochemical properties rather than by any direct pharmacological assessment. Assays were developed to explore product composition (SDS-

PAGE protein separation, collagen content, lipid content), potency (cell migration), as well as levels of residual manufacturing chemicals. This testing was intended to monitor batch-to- batch variability during manufacturing and identify properties which might correlate to in vivo efficacy outcomes. These assays were also used to determine how different processing conditions or tissue sources altered the final product. For some of these analytical methods, irradiated and non-irradiated samples were tested in parallel to assess any effects of the sterilization process. Porcine and human batches were also compared, as both species of

AAT were used in early preclinical development and in more recent animal experiments (see

Chapter 3).

A colorimetric assay was employed to determine the collagen content of AAT.

Hydroxyproline (4-hydroxyproline) is a non-proteinogenic amino acid formed by the post- translational hydroxylation of proline, where it serves to stabilize the protein’s helical structure. Because hydroxyproline is largely restricted to collagen, the measurement of hydroxyproline levels can be used as an indicator of collagen content in tissue sample.

The absolute hydroxyproline content of AAT does not change significantly from irradiation

(as expected) and is only about 10% less than that of pure collagen. Irradiated AAT contains

110.2 µg of hydroxyproline per mg (dry weight), making it 87.0% collagen by weight.

Similarly, non-irradiated AAT contains 112.7 µg of hydroxyproline per mg (dry weight) and is therefore 89.0% collagen by dry weight. The conversion factor between hydroxyproline and collagen was determined by dividing the amounts of bovine collagen assayed by the amounts of hydroxyproline recovered. For this experiment, the calculated conversion factor was 7.9, which corresponds to a hydroxyproline content of 12.7% in collagen. This value very closely matches literature values which estimate the hydroxyproline content of collagen between 12.0% and 13.5% [34].

Although AAT is composed primarily of collagen, proteomic data has shown that AAT and other ECMs contain a wide variety of different proteins. To visual the protein “signature” of

AAT, bacterial collagenase was used to produce a water-soluble protein fraction for gel electrophoresis. Collagenases are endopeptidases that digest native collagen in the triple helix region. Unlike animal collagenases that split collagen in its native triple-helical conformation, bacterial collagenase is unique because it can degrade both water-insoluble

26 native collagens and water-soluble denatured ones. It can attack almost all collagen types, and is able to make multiple cleavages within triple helical regions [35]. After solubilization, protein separation by SDS-PAGE can be used to estimate relative molecular mass, to determine the relative abundance of major proteins in a sample, and to determine the distribution of proteins among fractions.

Digestion of AAT with collagenase solubilized a significant portion of solid material present in the sample. Though digestions were carried out for up to 24 hours, digestion for 5 hours was sufficient to achieve the maximum potential solubilization with this enzyme. SDS-PAGE analysis did not reveal visual differences in the distribution or abundance of proteins solubilized by collagenase in the irradiated and non-irradiated samples. These results provide a qualitative demonstration of AAT's proteomic diversity and suggest that the terminal sterilization step does not significantly alter the structure of AAT proteins or their susceptibility to degradation by collagenase.

In addition to the structural components of AAT, it was important to investigate potential contaminants that might reduce efficacy or otherwise alter the tissue response to AAT in vivo. Triglycerides account for the majority of the lipid content in native adipose tissue; therefore measurements of glycerol content correspond approximately to overall lipid content. Intracellular lipids are spilled from adipocytes when cell membranes are disrupted by chemical and physical processing; because these lipids possess inflammatory properties that may affect the host response following implantation [3, 5], the manufacturing process was designed to eliminate as much free lipid as possible. Since total amount of triglycerides

27 can vary significantly between adipose from different species or potentially even different human donors, it was important to discover if different AAT batches had similar amounts of triglyceride remaining after processing.

Lipid content was determined for several batches of human and porcine-derived AAT in order to compare the effectiveness and consistency of our lipid removal methods between different starting materials and manufacturing runs. The results demonstrate that different batches of AAT have very similar final lipid contents, regardless of tissue species or whether the product was manufactured in a GMP setting. This suggests that any differences observed in vivo are likely not related to variation in the lipid content of the materials. Since lipids present in the final product are a potential source of inflammation, lipid content may be directly correlated to key efficacy metrics such as volume persistence and immune responses.

In vivo studies have shown that cells infiltrate AAT following subcutaneous implantation, suggesting that there may be soluble factors present in the AAT that encourage migration.

The cell migration assay evaluates the effect of different concentrations of AAT on human

ASC migration using a transwell format. The results of these assays showed that both the irradiated and the non-irradiated samples of human AAT promote roughly equal cell migration of ASCs across a transwell membrane. Although non-irradiated AAT showed somewhat less migration than irradiated AAT, this could be due to the stickiness of the non- irradiated AAT interfering with cell counting rather than changes in biologic activity.

Though this assay provides a non-specific functional measurement of the soluble factors in

AAT, it has a number of limitations. There is the potential for significant variation between

28 different assays due to minor changes in culture conditions. Confluency of the ASCs prior to serum starvation appears to have a significant effect on cell migration, as evidenced by the large differences seen in migration in the control samples between different assays.

Therefore, migration results from different samples are best compared if they are obtained at the same time.

In response to a request from the FDA, two assays were developed to monitor the presence of residual process chemicals in the final AAT product. It was noted that Triton X-100 and

EDTA may have unintended activity when implanted, therefore the FDA believed these chemicals should be monitored in each GMP manufactured batch intended for clinical use.

Fortunately, the manufacturing process was designed to ensure removal unwanted residual chemicals, and the results of the residual chemical assays confirmed what had been assumed.

Greater than 99% of the Triton X-100 introduced during manufacturing is removed by rinsing, and the EDTA levels appear to fall under the lower limit of detection of the semi- quantitative test strips employed for the study. Care was taken to eliminate the possibility of interfering substances in the AAT samples; a control batch of AAT was manufactured without any Triton X-100 or EDTA to serve as negative control during assay validation.

Taken together, the results presented in this chapter represent a significant advancement in the clinical development of AAT to Phase I clinical testing. These assays were developed over many years to fulfill the FDA request for information on the product and were used during product release testing of the clinical trial GMP batch (TS-0268). Future clinical batches will also be tested by these methods to generate growing database of product information; this data will help define the process variability and inform future

29 manufacturing scale up. These biochemical characterizations ought to be performed whenever new batches are produced at lab-scale for animal use as well. Immunological results obtained from recent animal studies (presented in Chapter 3) have highlighted interesting differences between AATs in models of traumatic wound healing and non- traumatic subcutaneous injections. Comparing the biochemical properties of these AATs may shed light on new in vivo findings.

3. IMMUNE CHARACTERIZATION OF AAT IN MURINE

MODELS OF WOUND HEALING

3.1. Introduction

AAT is a being developed clinically for soft tissue augmentation in patients with stable tissue defects; however many studies have demonstrated the potential for ECM materials to also treat traumatic injuries [26, 36, 37]. AAT is extracellular matrix (ECM) derived from human adipose tissue that has been processed mechanically and with mild reagents to remove the lipid and cellular components, though the despite having no living cells, the final product is not an immune inert biomaterial. The processed ECM proteins which make up the solid bulk of AAT provide local signaling which induces colonization of the material by the patient’s own cells, but the mechanisms are unknown.

31 and examined the role of macrophages in the remodeling process after implantation of a surgical mesh [27]. Macrophages play a key role in wound healing; these cells are highly plastic and exist on a phenotypic spectrum ranging from “M1” macrophages (typically described as pro-inflammatory or classically activated) to “M2” macrophages (which are considered regulatory or homeostatic) [27]. Macrophage heterogeneity [28] and its implications for wound healing [29] can have a huge impact on the design of biomaterials to elicit pro-regenerative responses.

Dr. Elisseeff and Biomaterials and Tissue Engineering Lab at JHM recently published work in Science [26] investigating the immunological profile of various ECM-derived scaffolds by using flow cytometry to quantify the presence of T cells, B cells, vascular progenitor cells, dendritic cells, macrophages, M1-polarized (inflammatory) macrophages, and M2-polarized

(wound-healing) macrophages. These studies establish a critical role for the local immunological microenvironment in wound healing and suggest that the immune-modulating properties of ECM-based biomaterials such as AAT may be used in a targeted manner to facilitate wound healing.

Earlier preclinical in vivo testing of AAT evaluated compatibility and efficacy of volume retention in multiple models including mouse, rat, and swine [30, 38]. Because the AAT is derived from human adipose tissue, transplantation into other species represents a significant challenge to obtain meaningful results that are not complicated by immunological differences.

Human and animal research described therein has therefore aimed to investigate the immune composition of AAT and determine the resulting impact on wound healing and tissue regeneration. Adipose ECM from xenogeneic (human and pig) and syngeneic (C57BL/6 mice) sources were applied in mice to injured and non-injured tissues to isolate the effect of xenogeneic antigens, a significant limitation of all previous studies of ECM-mediated immune responses and wound healing.

3.2. Materials and Methods

3.2.1. Subcutaneous injection of AAT in mice

Animals were anesthetized using 4% isoflurane and maintained during the surgery using

2.5% isoflurane. The area was sterilized using 70% ethanol. Animals received two 0.20 cc injections of ECM into the subcutaneous space at superior and inferior positions on the dorsal side of the animal. Animals were then monitored until waking. At desired study end points (1 or 3 weeks), animals were sacrificed and samples were collected (implants and inguinal, axillary, and brachial lymph nodes). For flow cytometry and gene expression analysis, any skin was cut away from the implant. For histology, skin and implant were harvested together.

3.2.2. Volumetric muscle loss (VML) surgery in mice

Animals were anesthetized using 4% isoflurane and maintained during the surgery using

2.5% isoflurane. Hair was removed at the surgical site, and the area was sterilized with 70% ethanol. An incision was created from just above the knee to the hip. A 3 mm x 3 mm defect was created in the quadriceps muscle using surgical scissors. The defect was filled with 0.05 cc of either ECM material or sterile DPBS (Gibco) as a control. The incision was closed

33 using 3-5 sterilized wound clips (Roboz Surgical). Immediately following surgery, animals received carprofen (Rimadyl, Zoetis) subcutaneously at 5 mg/kg for pain. Animals were then monitored until waking. At the desired study end points (1, 3, or 6 weeks), animals were sacrificed and both quadriceps and all inguinal (local) lymph nodes, and axillary / brachial

(distal) lymph nodes were removed. All samples designated for gene expression analysis were immediately transferred into RNAlater (ThermoFisher), stored at 4˚C for 24 hours, and then moved to -80˚C if ribonucleic acid (RNA) isolations were not to be performed immediately. Quadriceps for flow cytometry analysis were processed immediately after removal. All animal procedures were performed in accordance with protocols approved by

Johns Hopkins Institutional Care and Use Committee (IACUC).

3.2.3. Histology

Samples were fixed in 10% formalin, serially dehydrated in graded ethanol solutions, and embedded in paraffin. Microtome sectioning was performed to obtain sections of 5 μm thickness. Slides were stained with hematoxylin and eosin (H&E).

3.2.4. Flow cytometry

Harvested tissue was finely diced in 1X DPBS on ice. Diced tissue was then digested in an enzyme solution consisting of 1.67 Wunsch U/ml Liberase TL (Sigma-Aldrich) and 0.2 mg/ml DNAse I (Roche) in serum-free RPMI 1640. Digested tissue was then filtered sequentially through 100 μm and 70 µm filters. Cells were then pelleted at 4°C at 300xg for

10 minutes. In some cases, the cell pellets were enriched for hematopoietic cells using

Lympholyte (Cedarlane) reagent. Remaining cells were then washed in DPBS (300xg for 5 minutes), then resuspended in a viability dye and stained for 30 minutes on ice, then washed,

34 then stained 45 minutes on ice with the flow cytometry antibody mixtures (see panel summary in Table 3.1 and Table 3.2 below). Stained cells were then washed and fixed using

Cytofix (BD Biosciences), then washed and stored in DPBS for up to 24 hours prior to data acquisition. Data was obtained using an LSRII flow cytometer (BD Biosciences) and analysis was conducted with FlowJo software.

Table 3.1. Mouse flow cytometry panel for subcutaneous injection studies Conjugate Antigen Phenotype Staining Dilution AF488 CD3 T cells 1:250 BV421 CD19 B cells 1:400 PE-Cy7 F4/80 Macrophages 1:250 APC-Cy7 CD11c Dendritic cells 1:200 AF700 CD86 M1 macrophages 1:200 APC CD206 M2 macrophages 1:400

Table 3.2. Mouse flow cytometry panel for volumetric muscle loss studies Conjugate Antigen Phenotype Staining Dilution PerCP/Cy5.5 CD11c Dendritic cells 1:100 PE-Cy5 CD3 T cells 1:200 PE-594 (CF) Siglec-F* Eosinophils 1:200 PE CD206 M2 macrophages 1:250 Pacific Blue Ly6g Neutrophils 1:400 PE-Cy7 F4/80 Macrophage 1:250 APC CD86 M1 macrophages 1:400 BV605 CD45 Hematopoetic cells 1:150 AF488 MHCII I-A/I-E Antigen presentation 1:200 AF700 CD11b Myeloid cell 1:400 BV510 Ly-6C Monocytes 1:300 Unless otherwise noted, all antibodies were obtained from Biolegend. Antibodies marked with (*) were obtained from BD Biosciences.

3.2.5. Real time quantitative polymerase chain reaction (RT-qPCR)

Tissue was thawed and removed from RNAlater, rinsed in PBS and then homogenized in

TRIzol (Life Technologies) using fine scissors and RNase-free pestles. RNA was isolated by chloroform extraction and the aqueous layer containing RNA was transferred to a fresh tube containing an equal volume of 70% ethanol. The mixture was then applied to RNeasy Mini columns (Qiagen) and purified according to the manufacturer's instructions. RNA was eluted in RNase-free water and quantified using a Qubit 2.0 fluorometer (Invitrogen). RNA was treated to remove residual DNA using a cocktail containing DNase I, 10x DNase buffer and

RNaseOUT inhibitor according to reagent protocols (Life Technologies). Complementary deoxyribonucleic acid (cDNA) synthesis was conducted using Superscript Reverse

Transcriptase (RT) III enzyme as per manufacturer's instructions (Life Technologies). Real time quantitative polymerase chain reaction (qPCR) was conducted on Applied Biosystems

Real Time PCR machines using SYBR Green as a reporter. Primer sequences are included in

Table 3.3.

Table 3.3. Real time quantitative PCR Primer Sequences Gene Forward Primer (5' → 3') Reverse Primer (5' → 3') Arg1 ACAAGACAGGGCTCCTTTCAG TAAAGCCACTGCCGTGTTCA Retnlα CAGCTGATGGTCCCAGTGAAT AGTGGAGGGATAGTTAGCTGG Tnfα ATGGCCTCCCTCTCATCAGT TGGTTTGCTACGACGTGGG Il-4 GGTCACAGGAGAAGGGACGC AGCACCTTGGAAGCCCTACA Il-10 CAGGACTTTAAGGGTTACTTGGGT GCCTGGGGCATCACTTCTAC Il-1β TGCCACCTTTTGACAGTGATG AAGCTGGATGCTCTCATCAGG iNos CTTGGTGAAGGGACTGAGCTG GTTCTCCGTTCTCTTGCAGTTG Ifnγ CGGCACAGTCATTGAAAGCC TGTCACCATCCTTTTGCCAGT Gata3 CTCCTTGCTACTCAGGTGATCG AGGGAGAGAGGAATCCGAGT B2m CACTGAATTCACCCCCACTGA TCTCGATCCCAGTAGACGGT

3.2.6. Statistical analyses

3.3. Results

3.3.1. Subcutaneous injections of AAT in mice

To assess the in vivo response to the material, subcutaneous (SQ) injections of either hAAT or pAAT were made at superior and inferior positions on the dorsal side of the animal. The implants were harvested and analyzed at 1 or 3 weeks to assess short-term and longer term immune cell responses. Histological analysis showed minimal acute inflammatory response at both 1 and 3 weeks (Figure 3.1). Histological sections also show some significant differences in morphology between the hAAT and pAAT. The pAAT has a more textured appearance with larger pieces of material, while the hAAT looks smoother and more homogenous. Cell infiltration occurs from the surrounding tissues into the implant, indicating that the material promotes cell migration and corroborating the results of the in vivo cell migration assay. It is also interesting to note the brown fat pad adjacent to the implant in the

37 superior position (visible in the pAAT 3-week top section) which could potentially impact the cellular response to the material.

pAAT – 1 week hAAT – 1 week

Superior

Inferior

pAAT – 3 week hAAT – 3 week

Superior

Inferior

Figure 3.1. Hematoxylin and eosin staining of subcutaneous human and porcine AAT implants after 1 and 3 weeks. Implants labeled as “top” were injected at a superior position and implants labeled as “bottom” were injected at an inferior position.

The immune cell profile of the subcutaneous implants after 1 or 3 weeks in vivo were assessed using flow cytometry (Figure 3.2). Overall, the flow analysis showed that human and porcine AAT had similar immune cell profiles at each time point. A greater percentage of the cells migrating into the implant were CD206+ macrophages (M2 polarized) than

CD86+ (M1 polarized) macrophages, suggesting that the biomaterial skews macrophage polarization towards an M2 phenotype. When considering this result, it is important to understand that macrophage polarization is spectrum rather than a binary change, so macrophages could potentially be somewhere between an M1 and an M2 phenotype. It was also noted that the level of macrophage activation (F4/80hi relative to F4/80lo) was higher at 1 week than at 3 weeks. The percentage of CD3+ T cells in the implant is significantly higher at 3 weeks than at 1 week, indicating that the T cell response begins prior to 1 week and increases to a peak at some later time point.

h i P e r c e n t o f L iv e C e lls F 4 /8 0 M a c r o p h a g e A c tiv a tio n

8 0 *** 1 5 8

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/ *** * ** 6

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t h A A T - 3 w e e k s

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o N

5 ( p A A T - 3 w e e k s

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o C 0 0 0 h i lo F 4 /8 0 F 4 /8 0 C D 1 1 c + C D 3 + C D 1 9 +

h i P e r c e n t o f F 4 /8 0 + + C D 2 0 6 M a c r o p h a g e s C D 8 6 M a c r o p h a g e s M a c r o p h a g e P o p u la tio n

* *** 6 0 0 0 ** ** 2 0 0 0 1 0 0 * ***

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)

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D C C 1 0 0 2 0 1 0 0 5 0 0 0 0 h i lo h i lo F 4 /8 0 F 4 /8 0 F 4 /8 0 F 4 /8 0 C D 2 0 6 + C D 8 6 +

Figure 3.2. Subcutaneous flow cytometry results at 1 and 3 weeks. Implants were pooled for each animal to ensure adequate cell number for analysis. Statistical significance calculated using two-way ANOVA with TUKEY post-hoc testing for everything except F4/80hi macrophage activation for which a one-way ANOVA with TUKEY post-hoc testing was used. * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001.

Gene expression analysis performed on the SQ implants at 3 weeks post-injection showed that iNos, an M1 gene, and Arg1, an M2 gene, were both significantly increased in almost all of the implants. The increase in iNos gene expression in the pAAT far implant was not considered statistically significant. This information is interesting given the higher percentage of CD206+ M2 macrophages than CD86+ M1 macrophages in the implant observed in the flow cytometry data. Other genes including Il-4 were elevated in the implant relative to normal muscle, but these results were not statistically significant (Figure 3.3).

M 1 g e n e s

* 1 0 0 0 0 * * * * * * * * * U n in ju r e d m u s c le * * * * 1 0 0 0 * * * * P ig A A T , i n f e r io r * *

H u m a n A A T , i n f e r io r 1 0 0

Q P ig A A T , s u p e r io r R 1 0 H u m a n A A T , s u p e r io r

0 . 1 i N o s I L - 1  T n f 

M 2 g e n e s

* * * * * * * * * * 1 0 0 0 * * * * * * * * * * * * U n in ju r e d m u s c le * P ig A A T , i n f e r io r 1 0 0 H u m a n A A T , i n f e r io r

Q 1 0 P ig A A T , s u p e r io r R H u m a n A A T , s u p e r io r

0 . 1 I L - 4 A r g 1 I L - 1 0 R e t n l 

Figure 3.3. Gene expression in subcutaneous AAT at 3 weeks. Implants in the superior position and inferior positions were analyzed separately relative to gene expression in uninjured quadriceps muscle. Significance calculated using two-way ANOVA with TUKEY post-hoc testing. Asterisks with no line indicate significance compared to uninjured. * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001.

3.3.2. Volumetric muscle loss (VML) surgery in mice

A mouse volumetric muscle wound (VML) model was used to study the response to both pAAT and hAAT in a wound environment. In this experiment, a critical sized defect was created in the mouse quadriceps muscle and was filled with a biomaterial or saline as a control. To explore whether there was an effect related to xenogenic AAT in the mouse wound model, these experiments also included mouse-derived AAT produced from C57BL/6 mice as a syngeneic ECM control. Response to the biomaterials was assessed after 1 week.

Flow cytometry results of the 1 week VML study included a mouse AAT (mAAT) group to analyze the immune response to a genetically-matched ECM material (Figure 7). There were significantly more immune cells in quadriceps treated with hAAT and pAAT than those treated with mAAT or saline. Overall, the immune cell profile of wounds treated with mAAT more closely resembled those treated with saline than the xenogeneic AAT groups. The percentage of myeloid cells (CD45+CD11b+) was higher in both pAAT and hAAT. The xenogeneic AATs also recruited a much stronger eosinophil response (Siglec-F+MHCII-).

Interestingly, wounds treated with mAAT were the only group with a statistically significant increase in the proportion of T cells compared to uninjured quads. However, this there was no significant difference in the absolute number of T cells at the wound site between the different ECM treatments (data not shown). The data also indicates that pAAT and hAAT promote greater skewing of polarized macrophages to an M2-like phenotype than mAAT, though overall mAAT is still somewhat M2-polarizing. Saline treatment promotes a more

M1-like phenotype than any ECM treatment, as determined by the relative proportions of

CD206+CD86- and CD86+CD206- macrophages.

I m m u n e C e l l s ( C D 4 5 + % ) M y e l o i d C e l l s ( C D 1 1 b + % ) * * * * * * * * * * 1 0 0 * 9 0 * * * * * * * * * * * * *

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Figure 3.4. Volumetric muscle loss flow cytometry results at 1 week. Groups labeled as “uninj.” are uninjured quads from age-matched animals. Statistical significance calculated using a one-way ANOVA with Tukey post-HOC testing. Significance without lines indicate significance compared to uninjured. * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001.

RT-qPCR analysis of the wounded muscle 1 week after treatment showed that wounds treated with mouse syngeneic AAT were not significantly different than wounds treated with saline or uninjured muscle in any of the genes tested (Figure 3.5). Most of these M2 genes - including Il-4, Arg1, and Retnlα - were significantly increased in pAAT and hAAT treated wounds compared to saline treated wounds and uninjured muscle. Increases were also observed in M1 genes relative to saline treatment; though these increases were generally

42 similar between different ECMs. Most importantly, expression of Il-4 increased more than

100-fold in pAAT and hAAT relative to control groups, whereas mAAT also increased but was not significantly different than saline. Taken together, these results are consistent with flow cytometry analysis of macrophage polarization and indicate a difference in the profile of immune cells migrating to wounds treated with syngeneic ECM than those treated with xenogeneic ECMs.

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Figure 3.5. Gene expression in AAT-treated muscle defects at 1 week. Statistical significance is calculated relative to saline controls. * < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001.

3.4. Discussion

Preclinical studies showed promising results for the effective use of AAT to promote pro- regenerative immune responses with and without the presence of tissue trauma. In both VML and subcutaneous studies in mice, AAT demonstrated good tissue integration with cell migration into the implant. No significant inflammatory response was noted in these experiments. Results obtained from the subcutaneous implants indicate that macrophages begin migrating in from the surrounding tissue within 1 week. By three weeks, however, the macrophages are no longer as dominant and adaptive cells such as T cells have begun to appear. This mimics the immune response to a wound during which macrophages enter the wound first followed by lymphocytes [24]. RT-qPCR analysis showed an increase in gene expression for both iNos, the inducible form of nitric oxide synthase, and Arg1, Arginase 1.

The iNos gene generates nitric oxide and is an important enzyme in the macrophage inflammatory response [39]. Arg1 is an enzyme that metabolizes arginine and is highly expressed on M2 macrophages. Resident adipose tissue-associated macrophages typically have M2-like polarization and express Arg1 and are important for balancing inflammation in fat tissue and maintaining metabolism. Interestingly, Arg1 and iNos compete to metabolize arginine when they are co-expressed [40]. The co-expression of iNos and Arg1 likely indicates that the macrophages present within the AAT at 3 weeks are neither purely M1 nor

M2, though they may be skewed towards M2 at a population level.

Flow cytometry analysis of the AAT-treated quadriceps muscles at 1 week after critical injury showed significant infiltration of immune cells into the wound site. Overall the flow data suggest similarity between the xenogeneic AATs, including an increased percentage of

44 recruited immune cells (CD45+), eosinophils (CD45+ CD11b+ SiglecF+ MHCII-), and M2 macrophages (CD206+CD86-) relative to saline and uninjured controls. Although the differences were not significant, syngeneic mouse AAT was also somewhat more M2- polarizing than saline alone, and recruited greater percentages of immune cells. However, the macrophages present in mouse AAT tended to be more M1-polarized (CD86+) than in the other AATs. Gene expression analysis also showed that significantly more interleukin 4

(Il-4) was present in wounds treated with both pig and human AAT than those treated with saline or mouse AAT. This indicates that in some contexts, AAT promotes the migration of immune cells which trigger the release of this key pro-regenerative cytokine. This increased expression of Il-4 is not correlated with an increase in the proportion or absolute number of T cells in the wound, but may be due to the increased activity of myeloid cells orchestrating the upregulation of Il-4. Missing from these analyses are quantifications of different T cells subsets, particularly TH2 helper T cells, which are essential for creating a pro-regenerative microenvironment.

The combined results of our syngeneic versus xenogeneic AAT studies indicate that pro- regenerative immune responses to ECM are likely not driven by non-specific damage- associated molecular patterns (DAMPs) inherent to all ECMs [41]. It is possible that a response related to foreign antigens in both the pig and human AAT is significantly driving the observed increases in Il-4 expression, eosinophils, or M2 macrophage polarization.

However, there are likely factors other than species contributing to the differences observed between AAT treatments. Biochemical characterizations of the mouse AAT may reveal a key difference that helps to explain the loss of the pro-regenerative phenotype. More rapid

45 resorption of the mouse AAT compared to the xenogeneic materials may also be preventing the immune system from mounting a full M2/Th2 response. Additional studies will be needed to confirm the mechanism or mechanisms driving these phenotypes. Future work will investigate an allogeneic ECM material in the wound environment in order to better understand the implications of these results for the translation of AAT to clinical applications.

4. PHASE 1 CLINICAL TESTING OF AAT – SAFETY,

TOLERABILITY, AND EXPLORATORY FINDINGS

4.1. Introduction

Soft tissue loss may occur from trauma (i.e. burns, battlefield injury, and accidents), cancer

(i.e. lumpectomy or other resections), surgery, or simply from the aging processes. The loss of soft tissue is frequently permanent and causes disfiguring depression of skin where tissue is lost, which cannot only impact cosmesis but also function. In recent years, a substantial number of patients have acquired severe soft tissue deficits as a result of battlefield injuries.

Functional deficits which result from soft tissue loss include: lack of soft tissue support for distal extremities, such as amputated limbs, and restrictive scar contracture.

Many types of tissue are used for volume correction by reconstructive surgeons; however, adipose tissue grafting continues to be the gold standard procedure to repair soft tissue defects [2, 3]. Adipose tissue is employed to repair soft tissue defects because of its highly elastic physical properties and bulking capabilities. Nevertheless, adipose tissue transfer techniques have a number of limitations [42-44]. Current surgical interventions to replace

47 soft tissue primarily involve the harvest of autologous adipose tissue, and as such yield highly variable results and donor site morbidity. During the fat grafting procedure, adipocytes are ripped from their native environment and reinjected at another site of the body. Adipocytes are fragile by nature (due to the oversized lipid droplet they contain) and are thus easily ruptured by the negative pressure applied during liposuction. Once injected in their new site, adipocytes in the center of grafts no longer have access to a blood supply; these metabolically active cells undergo necrosis. Surgeons have developed numerous protocols to improve viability of the grafted cells by injecting into small channels rather than a bolus, using more involved preparations of lipoaspirate such as centrifugation or washing, and adding stromal vascular cells to aid in graft survival [45]. However, the unpredictable rates of resorption and lack of standardization of techniques remain a problem. Patients with acquired or congenital soft tissue deformities often live with their defects for years while undergoing multiple reconstructive procedures which can fail or provide unsatisfactory results, leaving patients both functionally and psychologically impaired.

For these reasons, soft tissue reconstruction remains a challenge in the healthcare industry.

Currently, large volume defects are treated with synthetic implant devices. For smaller tissue defects, synthetic and biological materials have been employed to provide temporary reconstruction. There are a number of soft tissue fillers currently approved for marketing in the US for correction of age-induced wrinkles, generally for augmenting small volumes of soft tissue. There are no products on the market today that fully resemble the AAT product; however there are some synthetic or biological products with similar indications. AAT has some characteristics of a number of current products from the perspectives of composition

48 and indication, such as acellular dermis (Alloderm™ and Cymetra™), that have a strong history of clinical use as tissue transplant products [46, 47]. From the composition perspective, AAT is a tissue-derived product similar to tissue products derived from bone, skin, amnion, small intestine, and pericardium.

AAT is expected to fill a significant clinical need for a simple and effective “off-the-shelf” soft tissue repair technology for small and large volume augmentation. The product could potentially be used for reconstruction of body defects, such as breast reconstruction, facial reconstruction, and cosmetic applications. The AST technology was developed and is designed to provide a natural structure to mimic normal soft tissue and promote new fat tissue growth in a sterile preparation that has been shown to promote adipose deposition in animal models. This novel and proprietary “off-the-shelf” solution can be prepared in multiple forms (injectable and bulk sheets) for different applications.

The proposed indication for AAT as a soft tissue acellular matrix is for replacing or supplementing missing, inadequate, or damaged soft tissue. As an acellular matrix with immune-modulating and other bioactive properties, AAT serves as a physical scaffold and provides biological signals to allow the recipient’s cells to repopulate the implanted material.

AST and the Elisseeff laboratory intend to develop this product for correction of soft tissue loss from trauma (i.e. burns, battlefield injury, and accidents) and cancer (i.e. lumpectomy or other resections). It is hoped that as clinical trials advance, safety will be proven for larger implant volumes as well as stable correction of larger deformities. These applications represent a significant unmet need in the healthcare industry.

Preclinical data suggest that the AAT biomaterial is safe for use in small and large animals; however, it is unknown if AAT is safe or efficacious in humans. A first-in-human, prospective Phase I trial was therefore conducted in healthy volunteers to assess the safety and tolerability of AAT intended for the repair of soft tissue defects in humans. The primary objective of this study was to assess the safety of AAT injections in healthy human subjects.

Secondary objectives included assessments of the biocompatibility of explanted implants using histological methods and the tolerability of AAT injections as reported in patient and physical satisfaction surveys. On an exploratory basis, the immune cell populations and cell migration into the implant were also characterized.

Together, these analyses will permit a comprehensive evaluation of the safety profile of this biomaterial for subcutaneous use. By conducting this proof-of-concept study in healthy human volunteers we will clinically evaluate an off-the-shelf soft tissue reconstructive material (AAT) that, if successful, will improve the quality of life outcomes and limit the need for invasive procedures for individuals with severe soft tissue defects. If our results are promising, acellular adipose tissue (AAT) may be a viable alternative to autologous tissue transplantation for correction of certain soft tissue deformities.

4.2. Materials and Methods

4.2.1. Subcutaneous injection of AAT in healthy human volunteers

An open-label pilot study was conducted in healthy volunteers undergoing elective surgery for the removal of redundant tissue (n=8). All participants were treated with the study

50 intervention: AAT injected subcutaneously in areas excised as a part of their previously planned elective surgical procedure (abdominoplasty or panniculectomy). A total volume of up to 4 mL of AAT was injected per participant, with 1 and 2 mL volumes for individual injection sites. Injections were made under local anesthesia and sterile conditions using a blunt needle and following standard injection procedures. All injections were performed in an outpatient setting with approval from the Johns Hopkins Institutional Review Board.

4.2.2. Follow-up visits

Participants completed a 1-week post-injection follow-up visit to assess all injection sites for

AEs/SAEs/UP/Es. Participants returned (depending upon whether or not they have implants) for follow-up visits at Weeks 2, and 4, and at the time of their final excision surgery. Non- excision follow-up assessments consisted of a physician assessment of the injected area, photographic documentation of the injection sites, review of concomitant medications, and recording of any unanticipated or serious adverse events potentially associated with the injection procedure. Findings were recorded in the study CRFs and any AEs/SAEs were reported to the medical monitor.

4.2.3. AAT excision

At the end of their assigned study time point, participants had all AAT implants removed simultaneously during their elective surgery after 1 week (±2d, n=2), 2 weeks (±2d, n=2), 4 weeks (±2d, n=2), 6 weeks (±2d, n=1) or 18 weeks (±2d, n=1) in situ. The predetermined redundant tissue containing the implants was excised by the surgeon and collected by the study team. The tissue was placed in a sterile container and transported to the lab in

51 compliance with relevant institutional safety requirements. Upon arrival in the lab, implants were labeled and excised to include a thin circumferential layer of native tissue and prepared for histopathological analysis, including but not limited to hematoxylin and eosin staining and flow cytometry (fluorescence-activated cell sorting, or FACS). Untreated adipose samples (>10 cm away from injection sites) were also collect from each participant.

4.2.4. Safety and tolerability

The primary outcome of safety was determined by the incidence and rate of adverse events.

Secondary outcomes were histopathological analysis of implants performed at the excision date between 1 – 18 weeks post-injection, as well as assessment of tolerability through participant-reported comfort and physician-reported ease-of-use with the intervention. Panel reactive antibody (PRA) testing was also conducted to evaluate any systemic immune reaction producing HLA (human leukocyte antigen)-reactive antibodies against the AAT implants at 4 and 12 weeks post-injection compared to a baseline PRA assessment.

4.2.5. Photoimaging

Standardized 2D photography was performed in order to document AAT injection locations and volume maintenance over time. Implant locations were documented using reliable physiological landmarks (i.e., the belly button for the abdomen) with the patient positioned such that the redundant tissue is supported. Landmarks were documented to ensure reproducibility for the patient’s return visits.

4.2.6. Surveys

Implants were evaluated by participants and physicians for fullness at the injection site and softness, smoothness, and naturalness of the implants at the Injection and Excision Study

Visits. Participants also rated comfort during and after the injection while physicians rated ease of use and overall appearance.

4.2.7. Histopathology

The section of pannus incorporating all AAT implants was excised from each subject during their abdominoplasty or panniculectomy procedure and histopathology analyses were performed on a single implant from each participant. Samples were fixed in 10% formalin, serially dehydrated in graded ethanol solutions, cleared in xylene, and embedded in paraffin.

Samples were sectioned at 5 µm thickness and stained with hematoxylin and eosin (H&E) staining protocol or Masson’s trichrome. For each participant, a trained pathologist scored sections from the AAT implant and an area of distal adipose as an untreated control site.

4.2.8. Immunostaining

Multispectral immunohistochemistry performed by sequential rounds of immunostaining and antigen retrieval using Opal reactive fluorophores and methodology (Perkin Elmer). Briefly, antigen retrieval was performed using heat-induced epitope retrieval with sodium citrate buffer. Deparaffinized and rehydrated slides were boiled in a microwave for 15 min in 10 mM sodium citrate, 0.05% Tween-20, pH 6. Sections were incubated in 3% hydrogen peroxide for 15 min, blocked (4% normal goat serum/1% BSA or 10% BSA in 0.05%

Tween-20 in TBS) for 30 minutes at room temperature (RT), and incubated with primary antibody for 30 minutes at RT. Secondary detection was performed using an Opal Multiplex

IHC Detection Kit (Perkin Elmer). Sections were incubated in the appropriate HRP-polymer conjugate for secondary antibody labeling (10 min, RT), then in Opal substrate working solution (10 min, RT). Sections were then re-microwaved and the protocol was repeated for each primary antibody. After sequential immunostaining, sections were incubated in Opal

DAPI working solution (5 min, RT), mounted in DAKO fluorescent mounting media and coverslipped. Slides were stored at 4°C until imaging.

4.2.9. Flow cytometry

Specimens were finely diced in 1X DPBS on ice and then digested in an enzyme solution consisting of 1.67 Wunsch U/ml Liberase TL (Sigma-Aldrich) and 0.2 mg/ml DNAse I

(Roche) in serum-free RPMI 1640. Digested tissue was then filtered sequentially through 100

μm and 70 µm cell strainers. The cell suspension was pelleted at 4°C at 300xg for 10 minutes and washed twice in DPBS (300xg for 5 minutes), and cells were counted. Cells were stained with LIVE/DEAD Aqua viability dye (Thermo Fisher) for 30 minutes on ice, washed, then stained for 45 minutes on ice with the flow cytometry antibody mixtures (Biolegend, see

Table 4.1). Stained cells were fixed using Cytofix reagent (BD Biosciences) and stored in

DPBS for up to 24 hours prior to data acquisition. Data was obtained using an LSRII flow cytometer (BD Biosciences) and analysis was conducted using FlowJo software.

Table 4.1. Human flow cytometry panel for myeloid cells Conjugate Antigen Phenotype Staining Dilution

BV605 CD45 Hematopoietic cells 1:20

AF700 CD11b Myeloid cell 1:200

AF488 CD11c Dendritic cells 1:20

APC/Cy7 CD68 Macrophage 1:20

PerCP/Cy5.5 CD14 Monocytes 1:20

APC CD15 Granulocyte 1:20

BV421 CD80 M1 macrophages 1:20

PE CD163 M2 macrophages 1:20

PE/Cy7 HLA-DR, DP, DQ Antigen presentation 1:20

4.2.10. Statistical analyses

Significance in grouped analyses with two variables was calculated using two-way ANOVA with Tukey post-hoc testing (α = 0.05). P values less than 0.05 were considered statistically significant (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001). Plotted values represent the arithmetic mean of the data set. Error bars represent +/- one standard deviation.

4.3. Results

4.3.1. Enrollment, demographics, and dosage

All screened patients (n=8) met inclusion criteria and were enrolled in the study. All study participants were non-Hispanic Caucasians and underwent abdominoplasty or panniculectomy procedures (Table 4.2). Study participants were injected with up to 4 mL of

AAT subcutaneously, and all implants were excised by 18-weeks post-injection. A summary of AAT dosage and excision time points in providing in Table 4.3.

Table 4.2. Overview of Study Participants Study Age Sex BMI Date of Amount Injected Excision Date Off Reason Off ID Injection Date Study Study Completed 01 64 M 71 4/25/16 Two 1 mL injections 5/24/16 7/19/16 study Withdrew from 02 30 F 70 8/19/16 Two 1 mL injections 8/31/16 11/29/16 study* Completed 03 52 F 26 8/26/16 One 2 mL injection 8/31/16 11/16/16 study Completed 04 27 F 26 9/14/16 Two 1 mL injections 9/30/16 11/21/16 study Completed 05 54 F 56 9/21/16 Two 1 mL injections 10/17/16 12/1/16 study Completed 06 51 F 27 9/21/16 Two 1 mL injection 9/27/16 12/14/16 study Completed 07 44 F 24 10/13/16 One 2 mL injection 2/17/17 4/6/17 study Two 1 mL injections Completed 08 62 F 38 10/14/16 & one 2 mL injection 11/22/16 1/3/17 study *Participant 02 withdrew from the study prior to completing the 6-week post-excision visit and 12-week post- injection PRA. This participant withdrew because she lived several hours away from our institution and no longer desired to come to follow-up visits.

Table 4.3. AAT Exposure Duration and Dosage Study Date of Date of Duration of Excision Time Cumulative AAT ID Injection Excision Exposure Point Dose 01 4/25/16 5/24/16 4 weeks, 1 day 4 weeks 2 mL 02 8/19/16 8/31/16 1 week, 5 days 2 weeks 2 mL 03 8/26/16 8/31/16 5 days 1 week 2 mL 04 9/14/16 9/30/16 2 weeks, 2 days 2 weeks 2 mL 05 9/21/16 10/17/16 3 weeks, 5 days 4 weeks 2 mL 06 9/21/16 9/27/16 6 days 1 week 2 mL 07 10/13/16 2/17/17 18 weeks, 1 day 18 weeks 2 mL 08 10/14/16 11/22/16 5 weeks, 5 days 6 weeks 4 mL

4.3.2. Adverse Events (AEs)

All eight subjects had at least one anticipated adverse event (AE) that was considered related to a study implant; however, these anticipated AEs were mild and localized to AAT injection sites (Table 4.4). Anticipated AEs experienced by patients included pain/tenderness (n=2 participants), erythema (n=4), bruising (n=4), hyperpigmentation (n=1), and textural change

(n=3). No subject experienced abrasion, edema, blistering, blanching, crusting, oozing, purpura, scabbing, ulcer, or scarring at their injection site(s), all of which are other common

AEs associated with injections.

Table 4.4. Anticipated Adverse Events Related to Injection Sites Excision Time 1 week 2 weeks 4 weeks 6 wk 18 wk Total Study ID 03 06 02 04 01 05 08 07 Pain/Tenderness Inj 1wk 1wk 3/8 (37.5%) Erythema Ex Ex 1wk 1wk 4/8 Ex (50%) Bruising Ex 1wk 2wk 1wk 4/8 (50%) Hyperpigmentation Ex 1/8 (12.5%) Textural change* 1wk 1wk 1wk 3/8 2wk 4wk 2wk (37.5%) Ex 4wk Other (define) Ex Ex Ex 3/8 Implant Implants Implant (37.5%) indurated indurated raised *Textural change was described as implant being “raised”, “indurated”, or “palpable” **Study Visit Key: Inj= Injection visit, 1wk= 1-week post-injection visit, 2wk= 2-week post-injection visit, 4wk= 4-week post-injection visit, Ex= Excision visit

In addition to the injection-associated AEs related listed above, three study participants (#01,

#02 and #05) were prescribed oral antibiotics postoperatively for cellulitis surrounding the surgical incision. Study participant #05 was also prescribed iron supplementation given the concern for anemia due to large drain output postoperatively. These were anticipated events given the procedure. One subject experienced a serious adverse event (SAE) which was not related to the study or study drug. Subject #01 underwent treatment for a severe infection following the panniculectomy; the medical monitor deemed that this SAE was a result of the surgical procedure only and related to the patient’s poor overall health and other co- morbidities. There were no other SAEs, serious adverse events, or subjects who discontinued

57 the study due to an AE. Laboratory results, physical examinations, and vital signs were unremarkable throughout the study.

4.3.3. Panel Reactive Antibody (PRA) testing

Baseline panel reactive antibody (PRA) results obtained prior to AAT injection were compared to results obtained 4 and 12 weeks post-injection for each study participant (Table

4.5). For 7 of 8 subjects, circulating HLA antibodies did not increase at either post-injection time point. One subject (#03) had a sub-clinical increase in HLA antibodies at 12 weeks post- injection (11 weeks post-excision). This minor change is likely not related to the study intervention, as it occurred nearly 11 weeks after removal of the implant. Therefore, we conclude none of the subjects had a systemic immunologic response which produced HLA antibodies to the AAT.

Table 4.5. Panel Reactive Antibody Results

BASELINE TESTING POST-INJECTION TESTING Excision Time Increase in IgG HLA antibodies at: PT # point IgG HLA Positive 4 weeks 12 weeks 03 1 week Yes No No Yes; lowest antibody detection level 06 1 week No No CPRA Low = 44 02 2 weeks Participant withdrew 04 2 weeks Yes No No 05 4 weeks No No No 01 4 weeks No No No 08 6 weeks No No No 07 18 weeks Yes No No

4.3.4. Photoimaging

Photos of the injection site on the day of excision are shown below for each individual

(Figure 4.1). Most anticipated AEs occurred during the first two weeks after injection, and most AEs were resolved by later time points. Mild erythema was the most common AE present at the time of excision (#06, 02, and 08). Some implants were raised or palpable under the skin (#03, 04, 05). One subject with an early excision time point still had mild bruising (#06). Participant 08 was still experiencing mild erythema, hyperpigmentation, and textural change at their 6 week excision time point; however, these changes were not obvious except on close inspection. The participant with the longest exposure (18 weeks) reported no

AEs at the time of excision (#07).

Figure 4.1. Injection site photos at time of excision. *For Participant #03, the injection site had to be photographed post-excision.

4.3.5. Surveys

Implants were evaluated by participants and physicians for fullness at the injection site and softness, smoothness, and naturalness of the implants at the Injection and Excision Study

Visits. Participants also rated comfort during and after the injection while physicians rated ease of use and overall appearance. All but one participant reported being very satisfied with their comfort during and after the injection. Participant appearance and fullness ratings varied, with most of the patients reporting and a high degree of satisfaction with fullness and that the implants felt very soft, smooth, and natural. Although appearance ratings were positive overall, some participants perceived their implants to become less soft and smooth over time.

Physicians were very satisfied with the overall appearance injected sites both at injection and at later time points. During some injections, the physician reported temporary needle clogging; however, this can be addressed by adding a homogenization step to the injection procedure in future studies. Physician ratings of injected site fullness, softness, naturalness, and smoothness ranged from neutral to positive and were essentially the same at both study visits (Figure 4.2).

Overall, both participants and physicians reported satisfaction with the study procedure and appearance of the implants. Participants tended to have higher levels of satisfaction than physicians. Appearance data will be more meaningful in the next phase of clinical testing when AAT implants will be used to fill defects rather than augment normal tissue. Deeper injections in the subcutaneous space will likely result in more natural volume correction.

Figure 4.2. Participant and physician surveys. (A) Participant comfort and appearance survey results obtained at injection and excision visits. (B) Physician ease of use and appearance survey results obtained at injection and excision visits.

4.3.6. Histopathology and immunostaining

Overall, histopathological assessment revealed minimal inflammatory responses with cell migration into the material from surrounding tissue in all participants (Figure 4.3). Though accurate volume measurements could not be taken on the skin surface with calipers during this study, histological analysis reveals that a significant amount of the original implant volume is retained in all participants. A trained pathologist assessed sections of untreated adipose tissue and AAT implant from each subject, reporting infiltration of polymorphonuclear cells (PMNs), lymphocytes, and histiocytes. Most implants had a thin band of fibrosis surrounding the implant, and subcutaneous fat necrosis was minimal or absent in all cases.

Cellular migration from the host tissue into the implant is apparent at host-implant boundary

(Figure 4.4A) and appears to increase with time in situ. By 18 weeks, there are significantly more cells present at the implant periphery and implant center than at earlier time points.

Additional multispectral immunohistochemistry analyses performed also reveal the formation of new blood vessels at the edge of the implant (CD31+ vascular endothelial cells) and infiltration of perivascular/adipose stem cells (CD34+) both around and within the implant at

18 weeks post-injection (Figure 4.4B).

Figure 4.3. Histology (H&E) by individual participant (AAT implant is outlined).

Figure 4.4. Cell migration over time and revascularization at 18 weeks. (A) Cellular migration into the acellular matrix from surrounding tissue. (B) Immunohistochemistry of DAPI (nuclei), CD31 (endothelial cells), and CD34 (perivascular and adipose stem cells) at 18 weeks post-injection. H = host, I = implant.

Rare multinucleate giant cells were seen in approximately half the cases and were usually associated with small pieces of polarizable material. The polarizable particles noted by the pathologist in this clinical lot of AAT were also previously noted in multiple animal studies submitted with our IND. Recently we were able to identify the material as cotton fragments derived from sterile gauze used to clean manufacturing equipment. Small threads of cotton became introduced during manufacturing and were milled into fragments along with the adipose matrix proteins. Though the cellular response to these particles appears mild, some fragments are seen encapsulated within multinucleated giant cells, indicated some immunogenic potential. For future clinical studies, manufacturing protocols will be altered to eliminate these particles.

4.3.7. Flow cytometry

A panel of immune surface markers was used to identify different populations of myeloid cells present inside and around the AAT implants, as well as in untreated distal adipose tissue, in 6 of 8 study participants (Figure 4.5). In 4 subjects, absolute cell counts were also taken during tissue processing.

Macrophage polarization was the main outcome assessed in these experiments, where CD80 and CD163 represented classically activated (M1) and alternately activate (M2) macrophages respectively. Macrophages were identified globally by surface expression of phenotypic markers (CD45+CD15-CD11bhiCD11c+CD14+MHCII+/-CD68+). Monocytes were defined similarly to macrophages (CD45+CD15-CD11bhiCD11c+CD14+MHCII+/-CD68-), although due to their extravasated location, it may be more accurate to classify these cells as immature

65 macrophages (CD68-) rather than monocytes. These two populations (CD68+ macrophages and CD68- immature macrophages/monocytes) are similarly polarized, therefore for simplicity they were combined for analyses of polarization in these studies.

Figure 4.5. Flow cytometry gating strategy for myeloid populations and macrophage polarization.

In the six subjects tested, the majority of macrophages present in both AAT and control fat were MHCII+, alternately activated M2 macrophages (CD163+CD80-). Subjects with later excision time points tended to have larger fractions of double-positive macrophages than those with earlier excisions; similarly, early excision time points appear to be associated with a larger proportion of unpolarized macrophages. Four subjects had increased percentages of

M1 (CD80+CD163-) macrophages at their injected site; however M1 macrophages were by far the least abundant macrophage phenotype in all AAT implants tested (Figure 4.6A). Four of six subjects show an apparent increase in the CD163 median fluorescence intensity (MFI) of macrophages in AAT implants relative to control adipose (Figure 4.6B). Increases in

CD80 MFI can also be seen in four cases (Figure 4.6C). Overall, the macrophage populations near and within AAT implants were fairly similar to subject-matched distal adipose. Obvious relative increases in M2 polarization are apparent in most cases; while somewhat less obvious are increases in M1 polarization (Figure 4.6D), as these increases are mostly accounted for by expansion of the double-positive macrophage population. MHC- macrophage populations were also analyzed, though these cells were much less abundant and much less polarized than MHCII+ macrophages (data not shown).

Dendritic cells expressed most of the same myeloid markers as macrophages and were defined as CD45+CD15-CD11bloCD11c+MHCII+/-, while granulocytes expressed

CD45+CD11bhiCD15+. A population of apparently non-myeloid MHCII-positive cells was also apparent in these tissue samples, as these cells were negative for all myeloid markers and possessing a lymphoid-like FSC x SSC scatter appearance (CD45+CD15-CD11c-

MHCII+).

Cell count data was obtained for four subjects (Figure 4.7). For most myeloid populations, including monocytes, macrophages, dendritic cells, non-myeloid MHCII+ cells, there were not statistically significant differences in cell counts between AAT and control tissue. As a percentage of CD45+ immune cells, subjects at later time points tended to have relatively fewer PMNs and relatively more macrophages within their implants. In the implants excised at 1 and 2 weeks (participant 006 and 004), the absolute number of M2 macrophages was triple that in control fat (p < 0.0001).

Figure 4.6. Macrophage polarization by percentage and median fluorescence intensity (MFI) in AAT implants and untreated distal fat. (A) Percentages of unpolarized (CD163-CD80-), M2 polarized (CD163+CD80-), M1 polarized (CD80+CD163-), and double-positive (CD163+CD80+) macrophages. (B) M2 polarization assessed by CD163 MFI of all macrophages. (C) M1 polarization assessed by CD80 MFI of all macrophages. (D) Macrophage polarization by participant (representative plots).

Figure 4.7. Myeloid populations in AAT implants versus untreated distal fat for four study participants. Quantification of polymorphonuclear cells (PMNs), macrophages (Mø), monocytes (mono), dendritic cells (DCs), and other MHCII+ populations are represented as a percentage of CD45+ cells and as absolute cell counts. Absolute cell counts are also shown for unpolarized, M2 (CD163+CD80-), M1 (CD80+ CD163-), and double-positive macrophages.

4.4. Discussion

Overall, AAT demonstrated satisfactory safety results in Phase I testing. Safety was evaluated with in human subjects respect to the duration and amount of AAT exposure, adverse events (AEs), including serious adverse events (SAEs) and other significant AEs, physical examination results, laboratory abnormalities, and vital signs. Biocompatibility and tolerability were assessed from histopathology results, panel reactive antibody (PRA) testing, and patient comfort / physician ease-of-use surveys.

None of the participants experienced SAEs or unanticipated AEs related to the study, or exited the study due to AEs. All AEs noted were expected and mild, including redness, bruising, textural changes, hyperpigmentation and tenderness at the injection site. Many other adverse events commonly associated with injections were not observed in any participant throughout the study (i.e.: scarring, ulceration, scabbing, purpura, oozing, crusting, blanching, blistering, edema or abrasions). The amount of material injected did not appear to correlate with any increase in negative inflammatory responses or increased severity of AEs, suggesting that the doses and exposures tested were within safe ranges for these individuals.

All participants were healthy and well-appearing at study visits and physical examinations were unremarkable, with one exception (unrelated SAE for participant 001). One participant was hospitalized for an infection following the panniculectomy procedure, but the study medical monitor ruled that this SAE was likely a result of the patient’s poor overall health and unrelated to AAT injection. Otherwise, the study physician(s) did not note any

70 unexplained changes in laboratory values during the study that were likely related to the study intervention.

Secondary outcomes of tolerability and biocompatibility were assessed by patient and physician surveys, PRA testing, and histopathology. Satisfaction and tolerability survey ratings were positive overall, though appearance scores will be much more meaningful in future studies where AAT is used to fill a defect rather than augment normal tissue. As noted by the blinded pathologist, cellular and tissue responses to the implant, and within the implant, were generally mild, typically including infiltrate of immune cells with some collagen remodeling at the implant periphery. Rare multinucleate giant cells were seen in approximately half the cases, sometimes associated with small fragments of polarizable material (also noted in some previous studies and recently identified as cotton thread contaminant from the surgical gauze used in manufacturing). Overall, histological and PRA assessments revealed minimal inflammatory responses in all subjects. The implants also show good volume retention at up to 18 weeks post-injection, the latest time-point studied, and none of the implants showed any indication of encapsulation, cyst formation, or tissue necrosis.

One of the distinct advantages of the adipose ECM compared to fat grafting is that the high metabolic burden of transplanting adipocytes is bypassed. Instead, the adipose ECM is an acellular graft that relies on host cells to migrate into the scaffold and repopulate it to form new tissue. This also ensures that the rate of vascularization is coupled with the rate of new adipose tissue formation to generate sustainable adipose tissue with high cell viability. Cell

71 migration into the AAT implant coupled with regenerative remodeling is required for new tissue formation. At the latest time point of 18 weeks, some of the migrating cells were identified as vascular endothelial cells and perivascular/adipose stem cells, suggesting the potential for new tissue development as cells repopulate the matrix. Additional immunohistochemical analyses on other samples are ongoing, so it is possible that regenerative processes are occurring at earlier time points as well.

This study was designed to provide initial safety data for AAT to support the treatment of patients with soft tissue defects. Since subcutaneous injections of AAT were administered into redundant tissues scheduled for surgical removal, we had the opportunity to recover treated tissues and evaluate cellular responses at a number of points between 1 and 18 weeks.

Though this wide distribution of time points severely limited the power of statistical analyses, having this range allowed us to visualize changes in cell migration and implant morphology over a relatively long period, encompassing both early and late immune system responses. Excised implants assessed using histopathological methods revealed cell migration into the matrix but not any significant inflammation, both key components of long- term efficacy.

On an exploratory basis, the specific immune cell populations which localize with the implant were also characterized by flow cytometry. In preclinical studies, trauma and xenogeneic ECMs enhanced macrophage polarization and triggered significant eosinophil recruitment. In this non-traumatic environment, allogeneic human adipose ECM did not elicit as strong an immune response. Immune cell populations associated with AAT were, on the

72 whole, similar to the resident immune cells in normal adipose tissue. The most striking differences were observed at the level of macrophage polarization, where at some early time points (2 weeks post-injection or less) the absolute numbers of M2 macrophages were three times higher than in surrounding adipose. Though M1 macrophages were also presence and also sometimes also increased, these cells were several orders of magnitude less abundant than M2 or double-positive macrophages. A major limitation in statistical interpretation of these studies comes from the low number of AAT samples in each participant (usually n=1 for each outcome) as well as the heterogeneity of the subjects themselves. However, these results support findings from animal studies which identified favorable immune properties of

AAT.

Taken together, we believe these findings establish the safety of AAT for subcutaneous injection in humans and support continuation of our clinical development program into Phase

II testing to treat patients with soft tissue defects. Importantly, the histological analysis also shows that the product promotes cell migration and so has the potential to promote new tissue formation and permanent correction of soft tissue defects. Flow cytometry results also validate regenerative immunology principles previous only studied in animal models. These findings will have significant impact for field and future clinical development of this product.

5. CONCLUSIONS AND FUTURE WORK

This work describes the development of an adipose-tissue derived ECM material for soft tissue reconstruction, including many of the material characterizations and preclinical studies that lead to the first in human study. These preclinical results and others were critical in obtaining FDA approval for initial clinical testing. Consequently, the safety results obtained from this first clinical study will be leveraged to advance AAT to Phase II clinical testing to confirm safety and determine efficacy in patients. Our most recent animal studies have sought to identify the mechanisms of ECM-mediated immunomodulation and will be critical to help define clinical indications for AAT and inform future research.

In these preclinical studies, AAT demonstrated volume retention, significant tissue integration, and minimal inflammation. Subcutaneous implants attracted large proportions of macrophages around 1 week, followed later by the clearing of the macrophages and increased migration of T cells. In a mouse wound environment, pig and human AAT elicited a strong

M2-macrophage response while syngeneic mouse AAT elicited a more neutrally-polarized macrophage response that was similar to saline treated wounds. However, CD3+ T cell response was not significantly impacted by ECM tissue source, which may suggest a combination of factors (both species-specific and non-species specific properties of ECMs) is

74 responsible for the immune response to these materials. The ability of AAT to modulate immune response and induce a favorable pro-regenerative environment could potentially be harnessed to improve wound healing and reduce scarring after injury. This data indicates that

AAT could be a good substitute for autologous fat transfer in the treatment of soft tissue defects.

In Phase I clinical trial studies, AAT proved safe, biocompatible, and well-tolerated by all outcomes measured. No serious adverse events were reported, and all anticipated adverse events were mild and localized to the injection site. Both physicians and participants reported overall satisfaction with the comfort/ease of use and appearance of the injected area.

Importantly, implant volume was retained until the latest measured time point of 18 weeks, and the material integrated into the surrounding tissue rather than becoming isolated by fibrosis. Histological analysis of the implants also showed significant cell migration into the implant from surrounding tissue which generally increased as time went on. Although immune cells were observed within the implant, the lack of a systemic immune reaction suggests that any immune-modulation is occurring locally. All of this data indicates that

AAT is safe for use in humans and has the potential to support cell infiltration and new tissue formation. Additional histological analyses will be conducted to identify specific types of immune cells present in the AAT, particularly T cells and eosinophils, as these cells have demonstrated roles in orchestrating macrophage polarization and wound healing. It will be critical to determine whether the cellular response to AAT is comprised of pro-inflammatory immune cells, pro-regenerative immune cells, and/or stem cells to fully understand the functional impact of adipose ECM-associated immune modulation.

Future studies will determine the immune response to allogeneic ECM in a wound model and determine mechanisms contributing to the ECM-associated immune microenvironment.

Biochemical characterizations must be performed for each new manufactured lot to develop an understanding of batch-to-batch variability and help determine which properties of the material are correlated with successful clinical outcomes. These factors and others will be considered for design and validation of a scaled-up manufacturing process for future clinical trials involving significantly more participants. Together, this work will enable Phase II clinical testing, which will be conducted to test the safety and efficacy of AAT in filling small soft tissue defects in human patients.

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7. CURRICULUM VITAE

EDUCATION Johns Hopkins School of Medicine // Baltimore, MD 2011 – 2017 Ph.D., Cellular and Molecular Medicine (CMM)

University of California-Davis // Davis, CA 2007 – 2011 B.S., Biochemistry and Molecular Biology, Magna Cum Laude (GPA: 3.84/4.00)

RESEARCH EXPERIENCE Doctoral Research // Johns Hopkins School of Medicine 2012 – 2017 PI: Jennifer Elisseeff, Ph.D., Department of Biomedical Engineering . Clinical development of an adipose extracellular matrix (ECM)-derived biomaterial for soft tissue reconstruction and creation of pro-regenerative immune responses . Characterized local immune responses to ECM biomaterials in preclinical animal wound models and identified mechanisms of action driving macrophage polarization and eosinophil recruitment . Spearheaded Phase I clinical research including: human safety and tolerability, evaluation of immune responses, development of quality control assays, and execution of all SOPs required for compliance . Developed protocols to determine batch to batch variability and cell-based potency assays . Evaluated adipose stem cells (ASCs) as a starting material for engineering metabolically active brown adipose tissue to counter obesity and associated metabolic disorders . Investigated an injectable biomaterial for localized delivery and sustained release of small molecules to promote brown fat differentiation and improve metabolic homeostasis . Genetically engineered stem cells to secrete therapeutic proteins with inducible synthetic gene switch . Supervised junior scientists and worked within a large multi-disciplinary team, including: clinicians, trial managers, consultants, government representatives; maintained relationships with CROs and tissue banks

Graduate Intern, Biopharmaceutical Development // MedImmune 2014 – 2015 Drug Delivery and Device Development Group, Gaithersburg, MD . Worked on independent projects, proposed and investigated novel methods of antibody delivery to tumors

. Developed a sustained release hydrogel and validated assays for measuring sustained release in vitro . Collaborated with an industry partner to test antibody loading in novel DNA-based nanoparticles

Research Rotations // Johns Hopkins School of Medicine 2012 PI: Saul Sharkis, Ph.D., Department of Oncology . Isolated and transplanted a unique stem cell population and evaluated its capacity for pancreatic homing and beta cell regeneration following intravenous administration

PI: William Matsui, M.D., Ph.D., Department of Oncology . Screened small molecule drugs to target cancer stem cells in multiple myeloma, employed in vitro cancer models and flow cytometry to track colony formation, viability, and stem cell metabolism

Undergraduate Research // University of California-Davis 2008 – 2011 PI: Wolf-Dietrich Heyer, Ph.D., Department of Microbiology . Investigated protein interactions in DNA repair pathways (S. cerevisiae) . Produced yeast stains expressing recombinant tagged proteins (plasmid editing, PCR, transformations), identified protein interactions using affinity purification and Western blots

PUBLICATIONS (*co-first author) 1. Anderson AE, Parrillo AJ, Wu I, Payne R, Aston J, Cooney C, Cooney D, Byrne P, Elisseeff JH. “A Phase I open-label study evaluating the safety of acellular adipose tissue (AAT), a novel soft tissue reconstruction solution, in healthy volunteers.” (In preparation). 2. Anderson AE, Parrillo AJ, Wu I, Elisseeff JH. “Regenerative immune-modulation of extracellular matrices is enhanced in xenogeneic implants.” (In preparation). 3. Yang JP*, Anderson AE*, McCartney A, Ory X, Ma G, Pappalardo E, Bader J, Elisseeff JH. “Metabolically active 3-dimensional brown adipose tissue engineered from white adipose-derived stem cells.” Tissue Engineering Part A. 2017. 23(7-8): 253-262.

PODIUM PRESENTATIONS 1. Anderson AE, Parrillo AJ, Payne R, Aston J, Cooney C, Cooney D, Byrne P, Elisseeff JH. A Phase I open-label study evaluating the safety of acellular adipose tissue (AAT), a novel soft tissue reconstruction solution, in healthy volunteers. Military Health Systems Research Symposium. Kissimmee, FL, 2017.

2. Anderson AE, Parrillo AJ, Payne R, Aston J, Cooney C, Cooney D, Byrne P, Elisseeff JH. A Phase I open-label study evaluating the safety of acellular adipose tissue (AAT), a novel soft tissue reconstruction solution, in healthy volunteers. International Federation for Adipose Therapeutics and Science. San Diego, CA, 2016. 3. Anderson AE, Deans TL, Elisseeff JH. Engineering tissues with controllable genetic circuits. Tissue Engineering and Regenerative Medicine International Society Annual Meeting. Washington, D.C, 2014. 4. Anderson AE, Deans TL, Elisseeff JH. Engineering tissues with controllable genetic circuits. Maryland Stem Cell Research Fund Symposium. Silver Spring, MD, 2014.

POSTER PRESENTATIONS 1. Yang JP, Anderson AE, Elisseeff JH. Local induction of browning in white adipose tissue using a PLGA delivery system. Tissue Engineering and Regenerative Medicine International Society Annual Meeting. Washington, D.C, 2014. 2. Anderson AE, Deans TL, Elisseeff JH. Turning stem cells into therapeutic implants: tissue engineering meets synthetic biology. Cellular and Molecular Medicine Annual Retreat. Baltimore, MD, September 2013. 3. Anderson AE, Schwartz EK, Zhang XP, Heyer WD. Investigation of Mus81-Rad54 interactions and their roles in homologous recombination and DNA repair. University of California-Davis Undergraduate Research Conference. Davis, CA, May 2010.

LEADERSHIP AND COMMUNITY SERVICE

President // Hopkins Biotech Network (HBN) 2014 – 2017 . Led executive team of >15 professionals (Ph.D., M.B.A., M.P.H.) and established new strategic collaborations . Co-founded mentoring program that matches aspiring entrepreneurs with mentors in the HBN network . Oversaw organization-wide activities: networking, fundraising, program management, recruitment, event planning, volunteer coordination, and website development (www.hopkinsbio.org)

Director of Events and Marketing // Hopkins Biotech Network (HBN) 2013 – 2014 . Coordinated a team for effective marketing, planning and execution of all HBN events

TEACHING EXPERIENCE Pollard Scholar Teaching Assistant, Cellular & Molecular Medicine Program 2013 . Selective teaching position awarded for achieving highest grade in first year graduate courses

. Prepared lectures and presented review material to incoming first-year Ph.D. students

AWARDS & HONORS . Nominated for Forbes “30 under 30” List in Science by Johns Hopkins University, 2018 . Young Investigator Award, Honorary Mention, Military Health System Research Symposium, 2017 . Three-Minute Thesis Competition, 3rd place winner, CMM Program Retreat, 2016 . Pollard Scholarship, Johns Hopkins University, 2012 . Citation for Outstanding Performance in Research, UC Davis, 2011 . Integrated Studies Honors Program, UC Davis, 2007-2011 . Regent’s Scholarship, UC Davis, 2007-2011