Influence of negative pressure would therapy and surface topography on soft-tissue responses around percutaneous devices

Percutaneous osseointegrated devices are being implemented and developed worldwide to improve the quality of life of amputee patients. Unfortunately, these devices disrupt the skin barrier and illicit wound healing responses which can increase the risk of infection. Unable to attach to, or integrate with, the device, the epidermis migrates alongside the device in an attempt to re-establish continuity. This phenomenon is known as epidermal downgrowth and leads to sinus tract formation, creating a portal for bacterial colonization and subsequent infection. In order to reduce the incidence of infection and improve the long-term success of percutaneous devices, there is a need to limit epidermal downgrowth by establishing a robust and stable skin-device seal. One strategy to limit epidermal downgrowth around percutaneous devices involves the application of negative pressure wound therapy (NPWT). However, it is unknown if epidermal downgrowth will remain limited once NPWT is discontinued. Additionally, the influence of NPWT on soft-tissue characteristics around percutaneous devices that exhibit limited downgrowth is unknown. Another strategy which has shown potential to improve soft-tissue outcomes around percutaneous devices involves the modification of device surface topography. Two commonly used topographies are porous and smooth. However, these topographies are unable to prevent epidermal downgrowth which can increase the risk of infection. The goal of this iv dissertation is to: 1) determine the ability of NPWT to prevent the progression of epidermal downgrowth after its discontinuation; 2) evaluate the ability of a microgrooved topography to limit epidermal downgrowth around percutaneous devices; 3) characterize soft-tissue responses, including vascularity and inflammation, to NPWT application which may play a role in limiting downgrowth. The purpose of this work is to contribute to the establishment of a robust skin-device seal with limited downgrowth for improved clinical adoption and effectiveness of percutaneous osseointegrated devices.

INFLUENCE OF NEGATIVE PRESSURE WOUND THERAPY AND SURFACE TOPOGRAPHY ON SOFT-TISSUE RESPONSES AROUND PERCUTANEOUS DEVICES by Divya Rajya Laxmi Pawar A dissertation submitted to the faculty of The University of Utah in partial fulfillment of the requirements for the degree of Doctor of Philosophy Department of Biomedical Engineering The University of Utah May 2019 Copyright © Divya Rajya Laxmi Pawar 2019 All Rights Reserved The University of Utah Graduate School STATEMENT OF DISSERTATION APPROVAL The dissertation of Divya Rajya Laxmi Pawar has been approved by the following supervisory committee members: Kent N. Bachus , Chair Jayant Agarwal , Member Robert D. Bowles , Member David W. Grainger , Member Vladimir Hlady , Member 12/14/18 Date Approved 12/14/18 Date Approved 12/14/18 Date Approved 12/14/18 Date Approved 12/14/18 Date Approved and by David W. Grainger , Chair/Dean of the Department/College/School of Biomedical Engineering and by David B. Kieda, Dean of The Graduate School. ABSTRACT Percutaneous osseointegrated devices are being implemented and developed worldwide to improve the quality of life of amputee patients. Unfortunately, these devices disrupt the skin barrier and illicit wound healing responses which can increase the risk of infection. Unable to attach to, or integrate with, the device, the epidermis migrates alongside the device in an attempt to reestablish continuity. This phenomenon is known as epidermal downgrowth and leads to sinus tract formation, creating a portal for bacterial colonization and subsequent infection. In order to reduce the incidence of infection and improve the long-term success of percutaneous devices, there is a need to limit epidermal downgrowth by establishing a robust and stable skin-device seal. One strategy to limit epidermal downgrowth around percutaneous devices involves the application of negative pressure wound therapy (NPWT). However, it is unknown if epidermal downgrowth will remain limited once NPWT is discontinued. Additionally, the influence of NPWT on soft-tissue characteristics around percutaneous devices that exhibit limited downgrowth is unknown. Another strategy which has shown potential to improve soft-tissue outcomes around percutaneous devices involves the modification of device surface topography. Two commonly used topographies are porous and smooth. However, these topographies are unable to prevent epidermal downgrowth which can increase the risk of infection. The goal of this dissertation is to: 1) determine the ability of NPWT to prevent the progression of epidermal downgrowth after its discontinuation; 2) evaluate the ability of a microgrooved topography to limit epidermal downgrowth around percutaneous devices; 3) characterize soft-tissue responses, including vascularity and inflammation, to NPWT application which may play a role in limiting downgrowth. The purpose of this work is to contribute to the establishment of a robust skindevice seal with limited downgrowth for improved clinical adoption and effectiveness of percutaneous osseointegrated devices. iv TABLE OF CONTENTS ABSTRACT .......................................................................................................... iii LIST OF FIGURES ..............................................................................................vii LIST OF ABBREVIATIONS .................................................................................. x ACKNOWLEDGEMENTS ....................................................................................xii Chapters 1 INTRODUCTION ............................................................................................. 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8 1.9 1.10 1.11 Percutaneous Osseointegrated Devices ......................................... 1 Current Status of Percutaneous Osseointegrated Devices ............. 2 Failure Modes of Percutaneous Devices ......................................... 4 The Structure of Skin ....................................................................... 6 Cutaneous Wound Healing.............................................................. 8 Foreign Body Response ................................................................ 17 Healing Around Percutaneous Devices ......................................... 19 History of Percutaneous Osseointegrated Device Development ... 21 Strategies to Improve Soft-Tissue Responses to Percutaneous Devices .......................................................................................... 24 Summary and Experimental Approach .......................................... 29 References .................................................................................... 32 2 PERI-PROSTHETIC TISSUE REACTION TO DISCONTINUATION OF NEGATIVE PRESSURE WOUND THERAPY AROUND POROUS TITANIUM PERCUTANEOUS DEVICES ........................................................................ 49 2.1 2.2 2.3 2.4 2.5 2.6 2.7 Abstract ......................................................................................... 49 Introduction.................................................................................... 50 Materials and Methods .................................................................. 52 Results .......................................................................................... 58 Discussion ..................................................................................... 60 Acknowledgements ....................................................................... 66 References .................................................................................... 67 3 EVALUATION OF SOFT-TISSUE RESPONSES AROUND LASER MICROOGROOVED TITANIUM PERCUTANEOUS DEVICES .................... 76 3.1 3.2 3.3 3.4 3.5 3.6 3.7 Abstract ......................................................................................... 76 Introduction.................................................................................... 77 Materials and Methods .................................................................. 80 Results .......................................................................................... 85 Discussion ..................................................................................... 87 Acknowledgements ....................................................................... 92 References .................................................................................... 93 4 INFLUENCE OF NEGATIVE PRESSURE WOUND THERAPY ON PERIPROSTHETIC TISSUE VASCULARIZATION AND INFLAMMATION AROUND POROUS TITANIUM PERCUTANEOUS DEVICES ................... 103 4.1 4.2 4.3 4.4 4.5 4.6 4.7 Abstract ....................................................................................... 103 Introduction.................................................................................. 104 Materials and Methods ................................................................ 108 Results ........................................................................................ 116 Discussion ................................................................................... 119 Acknowledgements ..................................................................... 127 References .................................................................................. 127 5 SUMMARY, CONCLUSIONS AND FUTURE WORK .................................. 140 5.1 5.2 5.3 Summary and Conclusions.......................................................... 140 Challenges and Future Work ....................................................... 144 References .................................................................................. 155 APPENDIX: XPS ANALYSIS OF DEVICE SURFACE ...................................... 160 vi LIST OF FIGURES Figures 1.1 Schematic of the components of the OPRA (Osseointegrated Prostheses for the Rehabilitation of Amputees) system .............................................. 45 1.2 Schematic of epidermal downgrowth progression around percutaneous devices ..................................................................................................... 46 1.3 Application of negative pressure wound therapy for 4 weeks over percutaneous devices limited epidermal downgrowth in a guinea pig model ....................................................................................................... 47 1.4 Progression of epidermal downgrowth with increased device in situ time 48 2.1 (A) A photograph showing the porous coated subdermal and smooth device percutaneous post ........................................................................ 70 2.2 A representative photomacrograph showing the three regions used for qualitative analyses .................................................................................. 71 2.3 Qualitative differences in blood vessels were seen between the groups . 72 2.4 A representative set of photomicrographs showing (A) normal epidermal thickness and epithelial tissues at the 3-point junction for the (B) Untreated Group, (C) NPWT Group, and (D) Discontinued Group ........................... 73 2.5 A representative photomicrograph of the neutrophilic inflammatory crust (area within dash line) present externally at the 3-point junction (marked by *) in all the groups adjacent to the device and percutaneous post ...... 74 3.1 A set of photographs showing the three topographies used for this study: A) microgrooved, B) porous and C) smooth (machine finish) ................... 97 3.2 A representative set of photographic images of the skin-device exit sites at 4 weeks post-implantation: A) Microgrooved, B) Porous and C) Smooth Groups ..................................................................................................... 98 3.3 A representative set of H&E stained photomacrographs: A) Microgrooved, B) Porous and C) Smooth Groups ........................................................... 99 3.4 A representative H&E stained photomicrograph showing the skin-device interface of the microgrooved device (A) ................................................ 100 3.5 A representative H&E stained photomicrograph showing the skin-device interface of the porous device (A) .......................................................... 101 4.1 A) A magnified photograph of the gauze and semi-occlusive dressing with the NPWT delivery tube inserted onto the dressing and further secured with semi-occlusive dressings to obtain an airtight seal ......................... 132 4.2 A photograph of the animal dorsum indicating the tissue collection sites ............................................................................................................... 133 4.3 Top panel - A representative set of photographic images of skin-implant exit site at 4 weeks post-implantation ..................................................... 134 4.4 A set of representative photomicrographs of peri-prosthetic tissues showing the immunohistochemical staining for CD31 and counterstained with hematoxylin ..................................................................................... 135 4.5 A bar-chart showing the blood vessel densities quantified in 3 regions . 136 4.6 A set of photomicrographs of Untreated and NPWT treated peri-prosthetic tissues at the wound edge showing immunohistochemical staining for CD68 ...................................................................................................... 137 4.7 A bar-chart showing the CD68 positive cell densities quantified in 2 regions ................................................................................................... 138 4.8 A representative set of photomicrographs showing the soft tissue-device interfaces: (A) Untreated Group showing a sealed interface with the epidermis contacting the subdermal device, denoted by the white arrow ............................................................................................................... 139 A.1 High-resolution O 1s spectra obtained from XPS for all device components ............................................................................................................... 166 A.2 High-resolution Ti 2p spectra obtained from XPS for all device components............................................................................................ 167 A.3 High-resolution C 1s spectra obtained from XPS for all device components ............................................................................................................... 168 viii A.4 High-resolution N 1s spectra obtained from XPS for all device components ............................................................................................................... 169 A.5 High-resolution Al 2p spectra obtained from XPS for all device components............................................................................................ 170 A.6 High-resolution Si 2p spectra obtained from XPS for smooth and microgrooved devices ............................................................................ 171 ix LIST OF ABBREVIATIONS Abbreviations OPRA.................. Osseointegrated Prostheses for the Rehabilitation of Amputees ILP ..................................................................................... Integral Leg Prosthesis OPL..................................................................... Osseointegrated Prosthetic Limb POP ..................................................... Percutaneous Osseointegrated Prosthesis CD68............................................................................Cluster of Differentiation 68 CD31............................................................................Cluster of Differentiation 68 PDGF ..................................................................... Platelet Derived Growth Factor VEGF .............................................................. Vascular Endothelial Growth Factor FGF..................................................................................Fibroblast Growth Factor IL ............................................................................................................ Interleukin TGF-β ................................................................ Transforming Growth Factor beta EGF ................................................................................ Epidermal Growth Factor HGF .............................................................................. Hepatocyte Growth Factor TGF-α .............................................................. Transforming Growth Factor alpha MMPs............................................................................. Matrix Metalloproteinases IFN-γ ........................................................................................... Interferon gamma NPWT ............................................................. Negative Pressure Wound Therapy TNF-α........................................................................ Tumor Necrosis Factor alpha IAF ................................................................................... Institute Armand Fappier PMMA ...........................................................................Poly (Methyl) Methacrylate H&E ....................................................................................... Hematoxylin & Eosin XPS................................................................... X-ray Photoelectron Spectroscopy ANOVA ...................................................................................Analysis of Variance O, Ti, N, C, Al, Si............. Oxygen, Titanium, Nitrogen, Carbon, Aluminum, Silicon DE ............................................................................................... Double Epidermis SE ................................................................................................ Single Epidermis GT ............................................................................................. Granulation Tissue CD............................................................................................ Caesarean Derived IHC...................................................................................... Immunohistochemistry NBF................................................................................ Neutral Buffered Formalin DAB ..................................................................................... 3-3’ diaminobenzidine HA ................................................................................................... Hydroxyapatite α-SMA .......................................................................... alpha Smooth Muscle Actin CRP .......................................................................................... C Reactive Protein PCR ............................................................................Polymerase Chain Reaction MMA ........................................................................................ Methylmethacrylate xi ACKNOWLEDGEMENTS I would like to thank my advisor, Dr. Kent Bachus, for his continued support, guidance and mentorship throughout my graduate school tenure. I would also like to express my gratitude to my committee members, Drs. Jayant Agarwal, Robert Bowles, David Grainger and Vladimir Hlady, for challenging me, for providing guidance and for helping me progress in the right direction. I would like to take this opportunity to thank Dr. Sujee Jeyapalina for being an immense source of knowledge and for helping me work through scientific challenges. Additionally, I also thank the past and present members of the Orthopaedic Research Laboratory for their discussions and support. A special thanks to Saranne Mitchell for being a good friend and mentor. Gratitude to Troy D’Ambrosio for giving me the opportunity to be a part of the Lassonde New Venture Development Program. I would like to thank my family, Mahipati, Bharati, Devavrat and Nani, for being there every step of the way. Their tremendous support and encouragement guided me through the numerous joys and frustrations of this process. They cheered me on, picked me up and motivated me to push through to complete this journey. Thank you to all my friends, without whom life would not have been as exciting. I am abundantly grateful to my husband, Arad, for being an integral part of this journey. Your constant reassurance, support, encouragement, love and patience made each day less overwhelming. CHAPTER 1 INTRODUCTION 1.1 Percutaneous Osseointegrated Devices Percutaneous devices penetrate the body through the skin barrier and serve as a conduit between the host tissue and the external environment.1 Some percutaneous devices, such as fixator pins and glucose sensors, are designed to function in the body for a short period of time. In contrast, other types of percutaneous devices such as dental implants,2 bone-anchored hearing aids,3 peritoneal dialysis catheters4 and infusion pumps are designed to reside in the body and function for long durations of time. In the US, percutaneous osseointegrated prosthetic devices are being researched and developed to revolutionize the standard of care for amputee patients.5-7 An estimated 185,000 Americans undergo a limb amputation annually.8 This number is projected to grow with an increase in the aging population and progressing rates of vascular diseases.9 A study by Ziegler Graham et al. estimated that approximately 3.6 million people in the United States will experience limb loss by 2050.9 Currently, the standard of care for amputee patients is the socket type prosthetic which is designed to create a tight fit around the residual limb and is held in place through suction, belts or cuffs. However, socket 2 prostheses have numerous shortcomings which severely impact the quality of life of patients.10,11 Problems reported with socket use include pressure sores,10 skin necrosis,12 discomfort13,14 and loss of functional mobility.14,15 Due to these problems, percutaneous osseointegrated devices were developed as an alternative to the socket prosthetics. In general, percutaneous osseointegrated devices consist of an osseointegrated stem implanted into the residual bone, a subdermal barrier and a percutaneous post extending through the soft-tissue to provide a docking site for the external prosthetic. The direct integration of the device to the residual bone of the patients eliminates soft-tissue compression due to socket use and helps alleviate some of the socket-associated problems. Additionally, percutaneous osseointegrated devices have been shown to improve the performance of daily activities, including walking ability and sitting comfort,13,16 while offering a conduit for enhanced neural control allowing for better control of the limb prosthetic.17,18 While percutaneous devices show great promise in helping patients regain the ability to perform some daily activities, there are concerns regarding the quality of the soft-tissue-device interface, which could limit the longevity, functionality and widespread translation of these devices. 1.2 Current Status of Percutaneous Osseointegrated Devices Rickard Branemark was the first to use a percutaneous osseointegrated device for rehabilitating a transfemoral amputee patient in 1990.19 In the following years, the field of percutaneous osseointegrated devices saw a vast expansion 3 with numerous research groups around the world independently developing different implant systems.20-23 Based on clinical trials, the system designed by Branemark underwent various iterations to accommodate other amputation levels.24-26 This standard system is known as OPRA (Osseointegrated Prostheses for the Rehabilitation of Amputees, Integrum, Sweden) and consists of an externally threaded cylinder (fixture), implanted into the medullary canal of the residual bone, and a percutaneous component called abutment which connects to the fixture (Figure 1.1). The implant is made of medical grade titanium alloy Ti6Al4V.20 A nano-textured surface treatment is applied to the bone-implant interface and the percutaneous component is polished smooth. A second system, known as ILP (Integral Leg Prosthesis, Orthodynamics, Germany), was developed by Hans Grundei and further implemented by Horst Aschoff.27-29 The implant is made of cobalt-chrome-molybdenum alloy and has a macro-porous texture (Spongiosa-Metal) on the bone-implant interface and a smooth polished percutaneous component. Similar to the ILP design, another system was introduced in Australia by Munjed Al Muderis and in Holland by Van der Meent and is called OPL (Osseointegrated Prosthetic Limb, Permedica s.p.a., Italy).30-32 The OPL system consists of a Ti6Al4V stem with a plasma sprayed rough titanium coating, implanted in the bone. The percutaneous component contains a smooth polished, titanium niobium coating. All three systems are commercially available. Other implant systems which are currently undergoing clinical trials are the ITAP (Intraosseous Transcutaneous Amputation Prosthesis, Stanmore Implants Worldwide, United Kingdom),16 COMPRESS (Zimmer Biomet, USA)33 and the 4 POP (Percutaneous Osseointegrated Prosthesis, DJO Global, USA).34 1.3 Failure Modes of Percutaneous Devices Percutaneous devices protrude through the skin barrier which protects the body from the outer environment and thus elicit a variety of responses within the peri-prosthetic soft-tissues (tissues surrounding percutaneous devices). This may ultimately lead to device failure.35,36 Furthermore, the percutaneous device can act as a physical barrier and prevent the skin from re-establishing continuity, thereby increasing complications associated with wound non-closure.37 Andreas von Recum was one of the first to describe the following modes of failure associated with percutaneous devices (Figure 1.2): marsupialization (epidermal downgrowth), permigration, avulsion and infection.36 Since then, several groups have reported similar outcomes.5,38,39 Epidermal downgrowth is said to occur when the epidermis proliferates and migrates parallel to the device surface, leading to the formation of an empty space (sinus tract) between the migrating epidermis and device. Due to the lack of epidermal-device attachment, the sinus tract may get filled with inflammatory cells and cell debris and becomes a source for bacterial colonization and subsequent infection. Permigration was described to occur primarily around porous devices with a non-porous core. In this process, the epidermis starts migrating internally through the pores, while fibroblasts in the underlying dermis begin to establish an immature provisional matrix. The migrating epidermis slowly lyses through the immature connective tissue and reaches the non-porous core of the device. After reaching the core, the 5 epidermis begins to migrate apically along the length of the device and reaches the device bottom, at which point the device is effectively extruded. Avulsion occurs due to the presence of repeated mechanical injury to the device. Mechanical irritation leads to the build-up of interfacial stresses that can result in tissue destruction. Tissue injury may also result in inflammation which over time can serve as a source for infection. Finally, in the scenario that the device becomes infected, the peri-prosthetic tissue becomes infiltrated with neutrophils and other inflammatory cells. The presence of prolonged inflammation arrests the epidermal proliferation and migration, leading to a lack of epidermal-device contact. Consequently, tissue integration with the device is impaired, reducing device stability and ultimate device extrusion. Epidermal downgrowth and lack of soft-tissue-device integration are the primary mechanisms which lead to device exposure and increase the risk of percutaneous device infection. It must be noted that infections are regularly, but not always, related to the lack of soft-tissue-device integration.1 Other reasons for device-site infections include, but are not limited to, an introduction of microorganisms during the surgical procedure, an existing infection at the implantation site and inadequate daily cleaning of the device after implantation.40 The most common complication reported with current clinical percutaneous osseointegrated prosthetic devices is superficial infection at the skin-device interface.41-43 A study in transhumeral amputee patients noted that 38% of osseointegrated arm prostheses developed superficial infections within 5 years of implantation.41 Another study in transfemoral amputees reported 58% infection 6 within 2 years.42 Although these superficial infections are typically treated using oral antibiotics, there are growing concerns regarding the frequent use and development of antibiotic resistance.44 If the initial antibiotic treatment is unsuccessful, then deep-bone infections can occur.41,42 These deep infections are hard to treat and generally require the removal of the entire implant.45 Other percutaneous osseointegrated devices such as dental implants also commonly face infections. Up to 56% of dental implants are afflicted with infections in the gingival soft-tissues surrounding the implants (peri-implant mucositis).46 Untreated, mucositis could lead to peri-implantitis, which is characterized by bone resorption and pocket formation around the implant and leads to implant failure.40 Regardless of the failure mode, the aforementioned complications result in a poor skin-device seal which can lead to a sinus tract formation between the skin and the device, and subsequent exposure of the underlying device. The sinus tract and exposed device can serve as a nidus for bacterial invasion and increase the risk factors for device infection.36 Therefore, the formation of a stable skin-device interface is important for long-term success.47 In order for percutaneous devices to be a viable standard of care, there is a need to limit epidermal downgrowth by establishing a stable tissue-percutaneous device interface. 1.4 The Structure of Skin Since a percutaneous device contacts the epidermal-dermal skin barrier, it is important to review and understand the cutaneous physiology of the host environment around the percutaneous devices. 7 The skin is a highly specialized and dynamic structure constituting of functionally distinct layers: epidermis, basement membrane (basal lamina), dermis and hypodermis. The epidermis is the outmost layer of the skin, composed mainly of keratinocytes (epidermal cells), which constitute approximately 90-95% of the cells in the epidermis. Within the epidermis, there are four distinct strata.48 The strata, from interior to exterior are: basale, spinosum, granulosum and corneum. The keratinocytes continually regenerate at the stratum basale and undergo progressive differentiation as they move toward the surface of the skin.49 Mitotically active keratinocytes from the stratum basale undergo differentiation into cells of the stratum spinosum, which then differentiate into keratinocytes of the stratum granulosum. The cells from the granulosum then transition to a terminally differentiated cornified cell type, which makes up the stratum corneum. In humans, it is estimated that keratinocytes take about 14 days to transition from the stratum basale to the stratum corneum.49 The stratum corneum functions as the primary skin barrier to prevent water loss and allows for the absorbance of environmental substances, as well as provides mechanical protection.49 The basement membrane, also referred to as the basal lamina, lies beneath the epidermis and is rich in glycogen, mucopolysaccharides and collagen IV. The basement membrane provides a dynamic link between the epidermis and the dermis. Additionally, the basement membrane functions to play a role in cell adhesion, differentiation, cell motility and in the transmission of extracellular signaling factors.50 The keratinocytes of the stratum basale are attached to the underlying basement membrane via adhesion structures known as 8 hemidesmosomes. Since the epidermis is avascular in nature, the proliferative keratinocytes in the epidermis receive oxygen and nutrients from the blood vessels in the underlying papillary dermis via the permeable basement membrane.49 The dermis consists of fibrous connective tissue which is made up of different dermal fibers (collagen and elastic fibers) and lies between the epidermis and the subcutaneous tissues. The primary cell type of the dermis is the fibroblast which plays a role in the synthesis and degradation of connective tissue matrix proteins.49 The dermis primarily functions to cushion the body from stress and strain, aiding in thermal regulation while providing elasticity and tensile strength to the skin. The dermis interacts with the epidermis during the repair and remodeling phase of wound healing.49 The dermis itself is approximately 15-40 times thicker than the epidermis and can be further divided into the papillary dermis and the reticular dermis.40,48 Beneath the dermis lies the hypodermis which is composed of adipose tissue and loosely woven collagen matrix. The hypodermis contains nerves, blood vessels, sweat glands, lymphatics and hair follicles. This layer of the skin insulates the body, supplies energy, cushions and protects the skin, and allows for skin mobility over the underlying structures.49 1.5 Cutaneous Wound Healing When the skin barrier is disrupted, the host tissue responds by undergoing wound healing. Wound healing is a complex and dynamic process involving a number of factors.49 The wound healing cascade is divided into four overlapping 9 and continuous phases: hemostasis, inflammation, re-epithelialization/granulation tissue formation and tissue remodeling.49,51 Tissue injury leads to disruption of blood vessels, blood extravasation and platelet release into the wound. In order to limit blood loss, hemostasis begins during which vasoconstriction, platelet aggregation and the coagulation cascade facilitate the formation of a blood clot. The hemostatic blood clot, composed of fibronectin and fibrin, functions as a provisional matrix for cell migration in the subsequent phases of wound repair. Platelets also secrete alpha-granules and growth factors that attract and activate neutrophils and later macrophages, fibroblasts and endothelial cells.51,52 In order to establish an immune barrier against invading micro-organisms, the inflammatory phase follows.52 The complement cascade is activated and neutrophils begin to arrive at the site of the cutaneous wound within 24-36 hours of injury.52 Neutrophils remove cellular debris, foreign particles, bacteria and function to primarily prevent infection. After 48-72 hours, monocytes appear at the wound site and differentiate into macrophages which aid in phagocytosis, killing of bacteria and removal of cellular debris. At the site of injury, macrophages encounter different stimuli and depending on the nature of the stimuli, macrophages become activated and acquire either an M1 or M2 phenotype.53 M1 macrophages are activated by interferon gamma (IFN-γ), lipopolysaccharide and tumor necrosis factor alpha (TNF-α).53 M1 macrophages are considered proinflammatory and are involved in antigen presentation and killing of intracellular pathogens. M2 macrophages are activated by IL-4 and IL-13, display an anti- 10 inflammatory expression profile and possess immunoregulatory and tissue remodeling characteristics.54 Immediately following tissue injury, macrophages exhibit primarily M1 characteristics through the production of TNF-α, IL-1 and IL6. As wound healing progresses and the remodeling phase of wound healing begins, macrophages transition to an M2 phenotype through the production of IL10.55,56 Recent studies have now challenged the traditional binary macrophage classification of M1 and M2.53,57 Evidence suggests that while the M1 and M2 phenotypes represent the polar ends of a continuum of phenotypes, various intermediate macrophage phenotypes exist in the spectrum. Furthermore, rather than distinct populations, macrophages can also develop mixed M1/M2 phenotypes. A recent study by Xue et al. identified nine different macrophage phenotypes in human macrophages based upon activation using chemical cues.58 Additionally, Xue et al. suggested that stimuli other than the traditional cues (IFNγ and IL-4), such as free fatty acids, may be involved in activating macrophages with mixed phenotypes.58 Overall, macrophages are versatile cells that display a variety of phenotypes, markers for which have not been well established and are part of ongoing research.55 The third stage of wound healing occurs 2-10 days post injury, and involves the coordinated action of two mechanisms: keratinocyte migration into the wound site, as well as fibroblast migration and extracellular matrix deposition in the wound bed.59,60 In this phase, keratinocytes play a major role in restoring the integrity of the injured epidermis through a process known as re-epithelialization.61 11 Keratinocytes are activated by growth factors and cytokines such as fibroblast growth factor (FGF), epidermal growth factor (EGF), hepatocyte growth factor (HGF) and transforming growth factor alpha (TGF-α).49,62 Additionally, the “free edge effect” caused by the absence of contact inhibition between neighboring cells at the damaged wound edge can also activate the keratinocytes. Upon activation, keratinocytes at the wound edge withdraw from terminal differentiation and undergo changes in morphology, gene expression and expression of cell surface receptors.63 In normal unwounded skin, keratinocytes attach to the basement membrane by hemidesmosomes which provide mechanical stability. After wounding, dissolution of intercellular desmosomes (which maintain neighboring cell-cell contacts) and hemidesmosomes occurs and the keratinocytes start migrating laterally over the wound matrix.49,51 Keratinocytes also express a number of enzymes that facilitate the degradation of the underlying extracellular matrix in order to make a path for the migrating cells.64 One to two days after injury, the epidermal cells behind the actively migrating cells begin to proliferate.49,51 As re-epithelialization progresses, these proliferating cells produce laminin, collagen type IV and collagen type VII which will reconstitute the basement membrane.49 Two theories of wound reepithelialization currently exist and remain a subject of debate: leap frogging and tractor tread.63 The leap-frogging theory suggests that within the migrating edge, keratinocytes roll over the already adhered basal keratinocytes, and revert to a basal phenotype once in contact with the wound bed.65 The tractor tread theory suggests that the basal keratinocytes migrate over the wound bed and maintain 12 the desmosomal junctions to the neighboring cells, to pull the epidermis inward from the edge.63 The keratinocytes eventually revert to their normal phenotype and re-establish firm attachment to the basement membrane through hemidesmosomes and the underlying dermis. Regardless of the mechanism by which re-epithelialization occurs, once the keratinocytes from the opposite sides of the wound edges come in contact, re-epithelialization is considered to be complete, and keratinocytes begin to differentiate. During the re-epithelialization phase, the wound also undergoes fibroplasia and angiogenesis.49 Fibroplasia refers to the formation of granulation tissue and occurs approximately 4 days after injury.51 At this stage, the granulation tissue replaces the provisional matrix. Granulation tissue provides a substrate for keratinocyte migration in the later stages of wound repair that can help keratinocytes complete re-epithelialization.59 During granulation tissue formation, macrophages, fibroblasts and blood vessels accumulate in the wound bed. The macrophages release growth factors such as platelet derived growth factor (PDGF), vascular endothelial growth factor (VEGF), FGF and transforming growth factor beta (TGF-β) which stimulate the migration and proliferation of fibroblasts to the wound site.49 Once in the wound, the fibroblasts synthesize components of the new extracellular matrix such as collagen III, glycosaminoglycans and proteoglycans.49,52 The formation of new blood vessels, known as angiogenesis, occurs concurrently with granulation tissue formation. Capillaries from the surrounding wound edges develop and diffuse to the wound bed, establishing a robust 13 angiogenic network with high blood vessel density.52 Angiogenesis is a crucial process since blood vessels carry oxygen and nutrients to the wound in order to sustain cell metabolism and tissue repair. The inability of the tissue to form blood vessels during repair can lead to delayed or impaired wound healing as seen in diabetic patients.51 The endothelial cell, the chief cell of angiogenesis, is stimulated by hypoxia (lack of oxygen), FGF and VEGF.52 Macrophages, when stimulated by hypoxia,66,67 can release angiogenic growth factors such as VEGF and FGF.51,68 Once the wound bed is filled with the blood vessel rich granulation tissue, angiogenesis diminishes and a majority of the new blood vessels disintegrate due to apoptosis and the vascular density of the wound bed returns close to normal.51,69 In the fourth and final stage of wound healing, wound contraction and remodeling takes place. This phase occurs over a period of weeks, months or even years, but its initiation overlaps with the formation of granulation tissue during the proliferation phase.64 Wound remodeling is a coordinated event between fibroblasts, collagen and matrix metalloproteinases (MMPs). Fibroblasts first increase production of collagen and these increasing quantities of collagen subsequently signal the fibroblasts to decrease collagen synthesis.64 Extracellular matrix remodeling occurs as collagen production decreases and collagen breakdown increases. Proteolytic enzymes known as MMPs, produced by macrophages, keratinocytes and fibroblasts, play a key role in collagen degradation.51 A constituent of mature matrix, collagen type I replaces the premature collagen type III.49,70 In early wound healing, collagen in disorganized, but the new collagen matrix becomes oriented and cross-linked over time.64 14 Simultaneously, fibroblasts change their phenotype to myofibroblasts and the wound undergoes contraction and shrinks in size. The final result is a coordinated event between restoration of the epidermal barrier via re-epithelialization, and proliferation and remodeling of the underlying dermis to become a mature scar. Hemostasis, inflammation, re-epithelialization/granulation tissue formation and tissue remodeling are general processes that occur during wound repair in both humans and animals. Since a majority of the studies assessing wound healing around percutaneous devices are performed in animal models, it is important to discuss differences and similarities in wound healing characteristics between human and animal models. Small animals, such as the rats and guinea pigs, are frequently used in wound healing studies since they are readily available, relatively inexpensive, easy to handle and provide a suitable area of skin for wound healing studies.71,72 Despite these advantages, rodents differ anatomically and physiologically from humans. The epidermis of mice, rats and rabbits is much thinner than the human epidermis.73 One exception to this is the guinea pig which has a thicker epidermis than other rodents and a similar epidermal thickness as humans.74 The pig is considered an excellent model for wound healing due to similarities in anatomy, physiology and immunology with humans.74 For example, the epidermis of the pig is of a similar thickness (30-140 μm) to that of humans (50-120 μm).72 Due to the skin’s elasticity and its lack of attachment to the underlying subcutaneous connective tissue, rodents are referred to as “loose-skinned” animals. In contrast, humans and pigs are considered tight-skinned. Additionally, rodents possess a 15 panniculus carnosus muscle, which is absent in humans and pigs. The panniculus carnosus muscle contributes to the wound healing process in addition to myofibroblasts-based contraction. Additionally, the panniculus carnosus muscle is implicated in rapidly bringing the wound margins together. Due to the presence of this muscle, rodent models are considered limited because of the perception that rodent wounds heal primarily through contraction. Some studies suggest that 80% of wound closure in rodents occurs through contraction.75,76 On the other hand, other studies suggest that the majority of the contraction in rodent wounds occurs after re-epithelialization has completed.77,78 A recent study showed that in full thickness wounds in a murine model, re-epithelialization and contraction each contributed approximately 40-50% to the initial wound closure.78 This result indicates that re-epithelialization contributes to a significant portion of closure in rodent wounds. Furthermore, the study also suggested that splinting wounds to reduce contraction79 may alter normal healing processes and thus should be carefully considered.78 While Chen et al. showed that both re-epithelialization and contraction occur during wound healing in rodents, the percent of wound closure attributed to contraction due to panniculus carnosus and myofibroblasts is unknown. Regarding blood supply of the skin, the number, size and distribution of dermal blood vessels in humans is comparable to that of the pigs.74,80 Both humans and pigs have a sub-epidermal, dermal and lower vascular plexus system that allows for increased skin blood flow.73 On the other hand, rodents have a more rudimentary vasculature network which consists of one primary network above the 16 panniculus carnosus muscle and a second network below the adipose tissue.73 In rodents, hair may also influence blood flow as seen in a study in rats which reported that hair-baring skin exhibited decreased blood flow compared to skin with reduced hair.73 Quantitatively, there are no studies that compare blood vessel densities between rats and humans; however, one study in hairless guinea pigs reported that the dermal blood vessel densities were similar to that of humans.81 Many of the insights into immunological differences between animals and humans have come from research on murine models.82 However, significant differences between mice and humans exist in the development, activation and response to challenge, in both the innate and adaptive immunities.82 For example, human blood consists of 50-70% neutrophils and 30-50% lymphocytes, whereas mouse blood consists of 75-90% lymphocytes and 10-25% neutrophils.82 Studies have also suggested that human macrophages may be fundamentally different than rodent macrophages.83 Human monocytes and macrophages express the CD4 molecule which is absent on mouse macrophages.84 CD4 is an important molecule involved in the recognition of antigen plus MHC class II molecules during immune responses. The F4/80 antigen is highly expressed by murine macrophages and is commonly used as a marker for labelling murine macrophages; however, its human ortholog mucin-like hormone receptor, EMR1, is only expressed in eosinophils in humans.85 One of the primary differences between humans and mice macrophages are the effector molecules deployed to control infections.83 While it is well established that rodent macrophages produce nitric oxide (NO) in response to certain infectious agents, the production of NO by 17 human macrophages is less clear. NO is produced by macrophages in response to intracellular pathogens and exhibits anti-microbial and cytotoxic actions to protect the host from infection. In mouse macrophages, IFN-γ and lipopolysaccharide induce the expression of NO; however, the same inflammatory mediators do not induce NO expression in human macrophages.82,83 Overall, animal models serve as an initial step for assessing the wound healing response to a biomaterial or therapy, but the similarities and differences of the selected animal model compared to humans should be considered when analyzing the results. 1.6 Foreign Body Response In its initial stages, the wound healing process around an implanted device is similar to the normal wound healing process. In addition, a host response to the foreign body also takes place. The sequence of reactions that occur following the implantation of a material/device is: blood-material interactions, provisional matrix formation, acute inflammation, chronic inflammation, granulation tissue formation, foreign body reaction, and fibrosis/fibrous capsule formation.86 The size, shape, chemical and physical properties of the device/biomaterial influence the intensity and duration of the host response. All implants and devices contain materials that are recognized by the blood as foreign. Immediately upon implantation, bloodmaterial interactions occur and proteins adsorb on and around the device, leading to the formation of a thrombus or blood-clot.87 This blood protein deposition on the surface of the device is described as provisional matrix formation and consists of 18 fibrin, activated platelets, inflammatory cells and endothelial cells. During the same time, an acute inflammatory phase also occurs and is marked by the presence of fluid (edema) and emigration of leukocytes (predominantly neutrophils). Similar to the normal wound healing process, neutrophils, monocytes, macrophages and fibroblasts migrate to the injury site. However, at this point, the normal wound healing cascade deviates and chronic inflammation begins due to the presence of persistent inflammatory stimuli.88 The chemical and physical properties of the implant, the continued presence of the implant and motion at the tissue-implant interface can all lead to chronic inflammation. This phase is characterized by the presence of macrophages, monocytes and lymphocytes. The chronic inflammatory phase is usually of a short duration, lasting no longer than 2 weeks, but persistence of lymphocytes and monocytes at extended time points may indicate an infection. However, in the cases of biocompatible materials or reduced inflammatory stimuli, chronic inflammation resolves leaving monocytes and macrophages to interact at the interface of the implanted device. After the resolution of acute and chronic responses, granulation tissue formation occurs characterized by the proliferation of blood vessels and fibroblasts. Eventually, the granulation tissue is replaced by dense connective tissue, known as fibrosis or fibrous encapsulation.88 The foreign body reaction at the device interface is controlled by macrophages, which may fuse together to form foreign body giant cells. The end stage of the foreign body response towards the device involves walling off the device by a fibrous capsule to isolate the implant and prevent it from interacting with the surrounding tissue environment.89 19 1.7 Healing Around Percutaneous Devices The normal wound healing cascade described in section 1.5 and the foreign body response discussed in section 1.6 are complicated by the presence of the percutaneous post that permanently protrudes through the skin barrier. Thus, wound healing around the percutaneous post is complex and yet to be fully described. Epidermal downgrowth, a major failure modality of percutaneous devices (described in section 1.3), is attributed to the re-epithelialization phase of the wound healing cascade.35,36,90 Re-epithelialization involves the proliferation and migration of keratinocytes over the underlying granulation tissue in order to cover the wound defect.51 In open wounds without devices, once the migrating keratinocytes from the opposing wound edges meet, re-epithelialization phase is said to be completed and the migratory keratinocytes revert to the differentiating phenotype. However, due to the presence of the percutaneous post, the surrounding wound edges are unable to meet. In a frustrated attempt to close the defect, the epidermis migrates adjacent to the granulation tissue, alongside the device surface, leading to epidermal downgrowth. Factors that influence wound healing and epidermal downgrowth around percutaneous devices include vascularity, inflammation and mechanical forces. Studies involving dental implants suggest that a reduced blood supply may lead to negative healing outcomes in gingival tissues surrounding the dental implants, which may ultimately lead to tissue downgrowth.91,92 Inflammation caused by bacterial colonization at the skindevice interface may also negatively impact wound healing around percutaneous 20 devices and cause epidermal downgrowth.2 The rate of downgrowth may be exacerbated due to shear/tensile forces at the interface between the skin and the device, which may continuously break any attachment the epidermis makes with the device.36 Interfacial forces around percutaneous devices may be increased in subdermal only devices compared to bone-anchored devices. Bone anchoring percutaneous devices can help limit interfacial forces between the skin and the device by stabilizing the device to the bone.93,94 Another factor that may influence motion around devices is the thickness of the surrounding peri-prosthetic softtissues. Studies have suggested that thinning the soft-tissue minimizes the relative movement between the surrounding skin and the percutaneous post.26,42 Thinning of the surrounding tissue is a currently utilized surgical technique for the implantation of commercially available percutaneous osseointegrated implants.24 A third factor which has been shown to influence interfacial forces between the tissue and the device is the device surface topography. Inclusion of a textured topography such as porous coating allows for better integration of the surrounding soft-tissue leading to a reduction of tissue movement compared to smooth device surfaces.5 Therefore, there is a need to investigate strategies that can allow for a stable and firm attachment of the epidermis to the device. These strategies should also help modulate inflammation, as well as achieve vascularization in the underlying tissue to maintain the epidermal-device contact over time. 21 1.8 History of Percutaneous Osseointegrated Device Development Percutaneous osseointegrated devices consist of a soft-tissue-device interface and a bone-device interface. The creation of a stable skin-device seal has been the common objective of studies, with some using unanchored or subdermal only devices and others using bone-anchored devices to assess softtissue responses to devices. The following section summarizes the pertinent literature regarding both bone-anchored and unanchored devices with a focus on soft-tissue responses. In the early 1970s, Winter investigated the skin-implant interface around both porous and solid percutaneous implants (not bone-anchored) in a porcine model up to 10 weeks.95 Implants made of polyethylene, porous polytetrafluoroethylene film and Millipore of varying pore sizes were tested. The author observed that the epidermis had migrated along the surface of solid materials, deep into the underlying tissues. Winter called this phenomenon epidermal downgrowth and concluded that it would subsequently result in the creation of a sinus tract, leading to infection.95 Around the same time, Hall et al. also investigated tissue responses to several porous and solid materials, in the dorsum of varying animal models (not bone-anchored), up to 12 months.90,96 Some implant materials tested were: nylon velour, Dacron velour and vitreous carbon. The authors suggested that if the epidermis is unable to establish a “good contact” with the device surface, then epidermal downgrowth will occur.90 Furthermore, Hall also postulated that when a break in the epidermal tissue occurs due to device implantation, wound healing leads to the formation of a granulation bed around the 22 device. The epidermal cells migrate upon this granulation bed, along the device surface, in an attempt to meet the cells from the opposite side which leads to epidermal downgrowth.90 Thus, Hall et al. were one of the first to provide an explanation for the mechanism of epidermal downgrowth. In the late 1970s, Mooney et al. implanted stainless steel intramedullary devices (bone-anchored) with an unpolished carbon collar as an interface between the skin and the underlying device, in three above-elbow amputee patients.97 However, all the devices were removed after 6 months post-implantation, due to the development of chronic infection. The authors suggested that some possible reasons for infection included the presence of mechanical stresses leading to irritation at the skin-device interface and poor tissue vascularization around the device.97 Moreover, Squier et al. investigated the influence of surface porosity on softtissue attachment and epidermal downgrowth in the back skin of pigs (not boneanchored).98 The results indicated that the presence of vascularized soft-tissue adjacent to the device may serve as a barrier against downgrowth.98 A review by Grosse-Siestrup et al. recommended parameters for percutaneous device design, based on investigations of naturally occurring percutaneous analogs such as horn, fingernail, hoof, tooth and antlers.99 The term three-phase junction was introduced, which is defined as the point where the air, the device and the tissue meet. The authors noted that the three-phase junction, referred to as the 3-point junction throughout the rest of this document, is important in determining the fate of the percutaneous device. The review proposed adding a subcutaneous component to 23 the percutaneous device, positioned next to muscles or bones, which can allow for tissue ingrowth to stabilize the device and also reduce the impact of mechanical forces and torques at the 3-point junction. Additionally, they suggested that the skin penetrating component should be cylindrical with a circular cross-section to minimize sharp corners that may cause stress peaks in the skin. Regarding the surgical procedure, a two-stage surgical procedure was recommended over a onestage procedure. In a two-stage approach, the subdermal component is first implanted followed by a period of recovery to allow for tissue integration into the implant. In the second stage, the percutaneous component is implanted and attached to the subdermal component. In contrast, in a one-stage procedure, the entire device is implanted in a single surgery.99 Starting in the late 1980s and going into the 21st century, John A. Jansen’s group conducted several studies to evaluate parameters that may influence percutaneous devices and are as follows: different animal models (guinea pig94, rabbit47,94 and goat100), placement location of the implants (cranium, tibia47 and the dorsum94,101) and different subcutaneous flange materials (titanium mesh and Dacron velour.)101-103 Results from the initial studies indicated that bone-anchored devices may be efficient in mitigating mechanical stresses at the skin-implant interface, resulting in limited downgrowth, compared to unanchored devices.47,94 However, as the studies progressed, the authors noted that in lieu of bone anchoring, incorporation of an appropriate subcutaneous flange may also help limit epidermal downgrowth.100,101 A subcutaneous flange consisting of a titanium fiber mesh elicited reduced inflammation compared to a Dacron velour flange.102,103 24 In the late 1990s and the following decades, research in the field of percutaneous devices was focused on investigating strategies to improve the periprosthetic soft-tissue responses to percutaneous devices. These studies will be discussed in the following section, according to the strategies they evaluated. 1.9 Strategies to Improve Soft-Tissue Responses to Percutaneous Devices Numerous strategies have been attempted to improve peri-prosthetic tissue responses with a goal to promote epidermal and dermal integration and reduce epidermal downgrowth. These strategies include surgical approaches (one-stage surgery and two-stage surgery),7,104 biological approaches (protein-based surface coatings,105-107 antimicrobial coatings, mesenchymal stem cell therapy108 and negative pressure wound therapy104,109) and engineering approaches (application of different materials,110,111 device design and surface topography112). While the aforementioned strategies have shown varying degrees of success, advances in wound healing therapies such as negative pressure wound therapy and device surface topography have shown potential and will be discussed below. 1.9.1 Negative pressure wound therapy Negative pressure wound therapy (NPWT) is a commonly used clinical treatment modality, used to achieve healing in acute and chronic wounds113,114 such as pressure ulcers, diabetic ulcers, skin grafts, abdominal wounds, traumatic wounds and burn wounds.115,116 Application of NPWT generates mechanical 25 forces at the wound edges which induce wound shrinkage. NPWT is discontinued once the desired goals have been achieved, for example when the wound has been optimized for surgical closure, the wound has achieved closure or when the wound fails to show signs of improvement such as no change in size.116,117 Although the duration of treatment varies, NPWT is rarely continued for more than 3 months.116 The primary mechanisms of action of NPWT include promotion of angiogenesis,66,118 granulation tissue formation,119 cellular proliferation and differentiation.119 Additionally, the mechanical stress induced due to NPWT application promotes wound healing by stimulating cell proliferation and differentiation.120 It is suggested that NPWT increases angiogenesis by promoting the expression of pro-angiogenic factors, VEGF and FGF.66,121,122 In addition, it is also postulated that NPWT regulates inflammation,116,123 although there is ambiguity regarding the specific role of NPWT and whether it increases124,125 or decreases126,127 inflammation in the wound bed. Overall, studies show that NPWT promotes healing and wound closure in clinical wounds.123,128 Recently, Mitchell et al. investigated the application of NPWT over percutaneous devices.104 NPWT was applied continuously for 4 weeks around porous coated devices in a guinea pig model (not bone-anchored). Results indicated that NPWT treated percutaneous devices exhibited significantly reduced epidermal downgrowth compared to percutaneous devices which did not receive therapy (Figure 1.3). However, the application of continuous NPWT around percutaneous devices is clinically impractical as a long-term treatment option. Furthermore, unlike clinical wounds, the presence of the percutaneous post makes 26 it difficult to determine if the wound has stabilized and if NPWT can be discontinued. Thus, there is a need to investigate if epidermal downgrowth around percutaneous devices will continue to be limited once NPWT has been discontinued. While NPWT was shown to limit epidermal downgrowth around percutaneous devices, the soft-tissue responses facilitated by NPWT, which may lead to limited downgrowth, are unknown. As indicated in section 1.7, it is suggested that the lack of vascularization and inflammation may negatively influence wound healing around percutaneous devices and lead to epidermal downgrowth. Thus, it is possible that NPWT may stimulate vascularization and modulate inflammation in peri-prosthetic soft-tissues, leading to limited downgrowth. Therefore, the soft-tissue responses around percutaneous devices after NPWT application need to be investigated. 1.9.2 Surface topography In order to limit soft-tissue complications, the percutaneous device surface should facilitate the establishment of a robust epidermal-device seal and should promote soft-tissue integration. To achieve this goal, numerous studies have investigated the use of porous surfaces. Using a rabbit model, Isackson et al. determined that porous coated titanium percutaneous devices (not boneanchored) had a significantly decreased risk of infection, due to enhanced softtissue integration, compared to smooth devices.129,130 Similarly, a study in an ovine model (bone-anchored) showed that porous coated percutaneous devices were 27 able to prevent infections up to 9 months, compared to smooth percutaneous devices.7 It was postulated that porous coated devices may prevent infection due to their ability to significantly limit the rate of epidermal downgrowth compared to the smooth devices.5,7 However, in a 2-year follow-up, the porous coated devices exhibited progressive epidermal downgrowth, with approximately 17% samples showing infection.131 Jeyapalina and colleagues also investigated the soft-tissue morphology around porous coated titanium percutaneous devices, in a 3-month porcine dorsum model (not bone-anchored).132 The soft-tissue response exhibited a migratory epidermis at the interface and an underlying region of granulation tissue. The authors noted that the peri-prosthetic tissues reflected characteristics of continuous wound healing, which may suggest a mechanism for epidermal downgrowth. Overall, while the porous coated devices limit the rate of epidermal downgrowth and offer initial protection against infection when compared to smooth devices, they are unable to completely halt downgrowth (Figure 1.4). This may ultimately threaten the long-term survival and functionality of these percutaneous devices. In order to determine an optimal surface to obtain epidermal attachment, Pendegrass et al. performed an in vitro study to evaluate the effect of different surface topographies on the proliferation and attachment of keratinocytes.133 Machine-finished, smooth-polished, sand-blasted and hydrofluoric acid-etched, titanium alloy discs were tested. Results showed that smooth-polished surfaces increased the attachment and proliferation keratinocytes. It should be noted that this study did not evaluate the response of keratinocytes to porous surfaces. 28 Furthermore, the results of this in vitro study have yet to be validated using in vivo models. Regardless, enhanced proliferation of epidermal cells on smooth surfaces may be one of the reasons why the majority of commercially available titanium implants contain a smooth surface at the soft-tissue-implant interface.40 A second reason may be that the smooth implant surfaces are relatively easy to clean. Finally, it has also been theorized that smooth surfaces allow the epidermis to undergo downgrowth towards the underlying muscle or bone, to presumably create a biological seal with the muscle or bone.28 Even if such an attachment does occur, no animal or clinical study has demonstrated this theorized epidermal-bone attachment. A third topography which has been widely researched is the grooved surface topography. In vitro and in vivo studies demonstrated that grooved surfaces oriented epidermal cells along the long axis of the grooves and reduced epidermal downgrowth compared to smooth surfaces.134,135 Furthermore, grooves aligned circumferentially around the implant promoted perpendicular tissue orientation to the implant surface and exhibited less downgrowth than grooves aligned parallel to the long axis of the implants.39,136 In 2002, Chehroudi et al. observed that although grooved topographies significantly limited epidermal downgrowth compared to smooth devices at 24 weeks, the grooved topographies still exhibited some amount of downgrowth.137 One reason may be due to a lack of texturing within the grooves. Studies on dental implants have shown that inclusion of micro-texturing or nano-sized features within the grooves can enhance the strength of cellular adhesion which may limit epidermal downgrowth over longer 29 time periods.138,139 Recently, a method was developed to produce microchannels, known as Laser-Lok® technology, with micro- and nano-structures, on titanium surfaces.140-142 Numerous animal143-146 and clinical studies147-149 have concluded that the application of microchannels (8 µm width, 4 µm depth) over a 2 mm wide band on the collar portion of the dental implant induces orientation and attachment of the epidermis and the connective tissue, thereby preventing epidermal downgrowth. While the Laser-Lok® technology has shown great success in limiting gingival tissue downgrowth around dental implants, it is yet to be used in percutaneous devices for classical skin applications. The incorporation of microtextured surfaces to percutaneous devices, implanted in dermal tissue, may limit epidermal downgrowth. 1.10 Summary and Experimental Approach In summary, percutaneous devices disrupt the skin barrier and are susceptible to infection. The risk of infection is increased due to the formation of a sinus tract created as a result of epidermal downgrowth. The initiation and progression of epidermal downgrowth is the product of wound healing responses to the continual presence of the percutaneous device. Additionally, epidermal downgrowth may also be attributed to reduced vascularity and inflammation in the peri-prosthetic tissues. In order to improve the long-term functionality of percutaneous devices, there is a need to limit epidermal downgrowth by establishing a robust and stable soft-tissue-device seal. The central hypothesis governing this work was that strategies which modulate peri-prosthetic tissue 30 morphology and promote device tissue integration will limit epidermal downgrowth around percutaneous devices. Two potential strategies to limit epidermal downgrowth are the application of NPWT and the modification of device surface topography. The goal of this work was to investigate a) the effects of discontinuation of NPWT on epidermal downgrowth, b) the use of the Laser-Lok® (microgrooved) topography to limit epidermal downgrowth and c) the influence of NPWT on peri-prosthetic soft-tissue characteristics, leading to limited downgrowth. Chapter 2 is comprised of a peer-reviewed publication. The goal of this study was to investigate the influence of discontinuing NPWT treatment on epidermal downgrowth. A recent study showed that, in a guinea pig model, application of NPWT for 4 weeks (NPWT Group) to porous coated titanium percutaneous devices significantly limited epidermal downgrowth compared to untreated controls (Untreated Group).104 However, the application of NPWT as a long-term treatment option is clinically impractical. Thus, there was a need to assess whether the application of a short course of NPWT could have long-lasting consequences on limiting epidermal downgrowth after the treatment had been discontinued. In order to test this, historically successful104,109 porous coated titanium devices were implanted in the dorsum of guinea pigs using established aseptic surgical procedures.104,109 Post-surgery, animals received 4 weeks of NPWT treatment followed by 4 weeks of no treatment (Discontinued Group). Results showed that the Discontinued Group at 8 weeks, had significantly more downgrowth than the NPWT Group. On the other hand, no significant difference in downgrowth was found between the Discontinued Group and the Untreated Group. 31 The results suggested that NPWT can only limit epidermal downgrowth while being applied. In Chapter 3, the aim was to investigate the influence of the microgrooved topography to limit downgrowth around percutaneous devices. Previous studies have shown that application of circumferential microgrooves to dental implants effectively halts gingival tissue downgrowth.144,147 However, it is unknown whether the microgrooved topography will limit epidermal downgrowth around percutaneous devices implanted in normal skin. To test this, percutaneous devices were designed with either a microgrooved, porous or smooth topography on the post. The porous and smooth topographies served as controls. The devices were implanted into three groups of guinea pigs, for 4 weeks. The results indicated no significant difference in epidermal downgrowth, with all three groups showing measurable amounts of downgrowth. This may suggest that varying topographical features may not be effective in limiting epidermal downgrowth. In Chapter 4, the goal was to evaluate the influence of NPWT on soft-tissue characteristics around percutaneous devices. Previous studies in open wounds have shown that NPWT increases angiogenesis and modulates inflammation, leading to good wound healing outcomes.66,116 Studies in peri-prosthetic tissues without NPWT have indicated that a lack of vascularization and presence of inflammation may lead to epidermal downgrowth.2,91 However, the influence of NPWT on peri-prosthetic tissue characteristics that lead to limited downgrowth is unknown. To elucidate the effects of NPWT on wound healing characteristics in peri-prosthetic tissues, percutaneous devices were implanted in two groups of rats: 32 one group received NPWT for 4 weeks and the other served as an untreated control. The results indicated that the NPWT Group showed half as much downgrowth as the Untreated Group, although no significant difference in downgrowth was found. The NPWT Group showed a two-fold increase in blood vessel densities compared to the Untreated Group. Lastly, the NPWT Group showed a trend towards increased CD68 positive cell densities compared to the Untreated Group. Overall, these findings suggest that NPWT may help improve wound healing outcomes around percutaneous devices by modulating vascularity and inflammation. 1.11 References 1.Peramo A, Marcelo CL. Bioengineering the skin-implant interface: the use of regenerative therapies in implanted devices. 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Development of a soft tissue seal around bone-anchored transcutaneous amputation prostheses. Biomaterials 2006;27(23):4183-91. 39.Brunette DM, Chehroudi B. The effects of the surface topography of micromachined titanium substrata on cell behavior in vitro and in vivo. J Biomech Eng 1999;121(1):49-57. 40.Abdallah MN, Badran Z, Ciobanu O, Hamdan N, Tamimi F. Strategies for optimizing the soft tissue seal around osseointegrated implants. Adv Healthc Mater 2017;6(20). 41.Tsikandylakis G, Berlin O, Branemark R. Implant survival, adverse events, and bone remodeling of osseointegrated percutaneous implants for transhumeral amputees. Clin Orthop Relat Res 2014;472(10):2947-56. 42.Branemark R, Berlin O, Hagberg K, Bergh P, Gunterberg B, Rydevik B. A novel osseointegrated percutaneous prosthetic system for the treatment of patients with transfemoral amputation: a prospective study of 51 patients. Bone Joint J 2014;96-B(1):106-13. 36 43.Al Muderis M, Khemka A, Lord SJ, Van de Meent H, Frolke JP. 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Development of a new percutaneous access device for implantation in soft tissues. J Biomed Mater Res 1991;25(12):1535-45. 102.Paquay YC, de Ruijter JE, van der Waerden JP, Jansen JA. Tissue reaction to Dacron velour and titanium fibre mesh used for anchorage of percutaneous devices. Biomaterials 1996;17(12):1251-6. 103.Jansen JA, Walboomers XF. A new titanium fiber mesh-cuffed peritoneal dialysis catheter: an experimental animal study. J Mater Sci Mater Med 2001;12(10-12):1033-7. 104.Mitchell SJ, Jeyapalina S, Nichols FR, Agarwal J, Bachus KN. Negative pressure wound therapy limits downgrowth in percutaneous devices. Wound Repair Regen 2016;24(1):35-44. 105.Chimutengwende-Gordon M, Pendegrass C, Blunn G. Enhancing the soft tissue seal around intraosseous transcutaneous amputation prostheses using silanized fibronectin titanium alloy. Biomed Mater 2011;6(2):025008. 106.Gordon DJ, Bhagawati DD, Pendegrass CJ, Middleton CA, Blunn GW. Modification of titanium alloy surfaces for percutaneous implants by covalently attaching laminin. J Biomed Mater Res A 2010;94(2):586-93. 107.Middleton CA, Pendegrass CJ, Gordon D, Jacob J, Blunn GW. Fibronectin silanized titanium alloy: a bioinductive and durable coating to enhance fibroblast attachment in vitro. J Biomed Mater Res A 2007;83(4):1032-8. 108.Isackson D, Cook KJ, McGill LD, Bachus KN. Mesenchymal stem cells increase collagen infiltration and improve wound healing response to porous titanium percutaneous implants. Med Eng Phys 2012;08(002). 109.Cook SJ, Nichols FR, Brunker LB, Bachus KN. A novel vacuum assisted closure therapy model for use with percutaneous devices. Med Eng Phys 2014;36(6):768-73. 110.Fleckman P, Usui M, Zhao G, Underwood R, Maginness M, Marshall A, Glaister C, Ratner B, Olerud J. Cutaneous and inflammatory response to longterm percutaneous implants of sphere-templated porous/solid poly(HEMA) and silicone in mice. J Biomed Mater Res A 2012;100(5):1256-68. 41 111.Fukano Y, Usui ML, Underwood RA, Isenhath S, Marshall AJ, Hauch KD, Ratner BD, Olerud JE, Fleckman P. Epidermal and dermal integration into sphere-templated porous poly(2-hydroxyethyl methacrylate) implants in mice. J Biomed Mater Res A 2010;94(4):1172-86. 112.Kim H, Murakami H, Chehroudi B, Textor M, Brunette DM. Effects of surface topography on the connective tissue attachment to subcutaneous implants. Int J Oral Maxillofac Implants 2006;21(3):354-65. 113.Argenta LC, Morykwas MJ. Vacuum-assisted closure: a new method for wound control and treatment: clinical experience. Ann Plast Surg 1997;38(6):563-76; discussion 577. 114.Morykwas MJ, Argenta LC, Shelton-Brown EI, McGuirt W. Vacuumassisted closure: a new method for wound control and treatment: animal studies and basic foundation. Ann Plast Surg 1997;38(6):553-62. 115.Orgill DP, Bayer LR. Update on negative-pressure wound therapy. Plast Reconstr Surg 2011;127 Suppl 1:105S-115S. 116.Huang C, Leavitt T, Bayer LR, Orgill DP. Effect of negative pressure wound therapy on wound healing. Curr Probl Surg 2014;51(7):301-31. 117.Bollero D, Driver V, Glat P, Gupta S, Lazaro-Martinez JL, Lyder C, Ottonello M, Pelham F, Vig S, Woo K. The role of negative pressure wound therapy in the spectrum of wound healing. Ostomy Wound Manage 2010;56(5 Suppl):118. 118.Greene AK, Puder M, Roy R, Arsenault D, Kwei S, Moses MA, Orgill DP. Microdeformational wound therapy: effects on angiogenesis and matrix metalloproteinases in chronic wounds of 3 debilitated patients. Ann Plast Surg 2006;56(4):418-22. 119.Scherer SS, Pietramaggiori G, Mathews JC, Prsa MJ, Huang S, Orgill DP. The mechanism of action of the vacuum-assisted closure device. Plast Reconstr Surg 2008;122(3):786-97. 120.Wiegand C, White R. Microdeformation in wound healing. Wound Repair Regen 2013;21(6):793-9. 121.Tanaka T, Panthee N, Itoda Y, Yamauchi N, Fukayama M, Ono M. Negative pressure wound therapy induces early wound healing by increased and accelerated expression of vascular endothelial growth factor receptors. Eur J Plast Surg 2016;39:247-256. 122.Jacobs S, Simhaee DA, Marsano A, Fomovsky GM, Niedt G, Wu JK. Efficacy and mechanisms of vacuum-assisted closure (VAC) therapy in promoting 42 wound healing: a rodent model. J Plast Reconstr Aesthet Surg 2009;62(10):13318. 123.Orgill DP, Bayer LR. Negative pressure wound therapy: past, present and future. Int Wound J 2013;10 Suppl 1:15-9. 124.Liu D, Zhang L, Li T, Wang G, Du H, Hou H, Han L, Tang P. Negativepressure wound therapy enhances local inflammatory responses in acute infected soft-tissue wound. Cell Biochem Biophys 2014;70(1):539-47. 125.Nuutila K, Siltanen A, Peura M, Harjula A, Nieminen T, Vuola J, Kankuri E, Aarnio P. Gene expression profiling of negative-pressure-treated skin graft donor site wounds. Burns 2013;39(4):687-93. 126.Norbury K, Kieswetter K. Vacuum-assisted closure therapy attenuates the inflammatory response in a porcine acute wound healing model. Wounds 2007;19(4):97-106. 127.Glass GE, Murphy GF, Esmaeili A, Lai LM, Nanchahal J. Systematic review of molecular mechanism of action of negative-pressure wound therapy. Br J Surg 2014;101(13):1627-36. 128.Orgill DP, Manders EK, Sumpio BE, Lee RC, Attinger CE, Gurtner GC, Ehrlich HP. The mechanisms of action of vacuum assisted closure: more to learn. Surgery 2009;146(1):40-51. 129.Isackson D, McGill LD, Bachus KN. Percutaneous implants with porous titanium dermal barriers: an in vivo evaluation of infection risk. Med Eng Phys 2011;33(4):418-26. 130.Farrell BJ, Prilutsky BI, Ritter JM, Kelley S, Popat K, Pitkin M. Effects of pore size, implantation time, and nano-surface properties on rat skin ingrowth into percutaneous porous titanium implants. J Biomed Mater Res A 2014;102(5):1305-15. 131.Jeyapalina S, Beck JP, Agarwal J, Bachus KN. A 24-month evaluation of a percutaneous osseointegrated limb-skin interface in an ovine amputation model. J Mater Sci Mater Med 2017;28(11):179. 132.Holt BM, Betz DH, Ford TA, Beck JP, Bloebaum RD, Jeyapalina S. Pig dorsum model for examining impaired wound healing at the skin-implant interface of percutaneous devices. J Mater Sci: Mater Med 2013;24:2181-2193. 133.Pendegrass CJ, Gordon D, Middleton CA, Sun SN, Blunn GW. Sealing the skin barrier around transcutaneous implants: in vitro study of keratinocyte proliferation and adhesion in response to surface modifications of titanium alloy. J Bone Joint Surg Br 2008;90(1):114-21. 43 134.Chehroudi B, Gould TR, Brunette DM. Effects of a grooved epoxy substratum on epithelial cell behavior in vitro and in vivo. J Biomed Mater Res 1988;22(6):459-73. 135.Chehroudi B, Gould TR, Brunette DM. Effects of a grooved titaniumcoated implant surface on epithelial cell behavior in vitro and in vivo. J Biomed Mater Res 1989;23(9):1067-85. 136.Chehroudi B, Gould TR, Brunette DM. Titanium-coated micromachined grooves of different dimensions affect epithelial and connective-tissue cells differently in vivo. J Biomed Mater Res 1990;24(9):1203-19. 137.Chehroudi B, Brunette DM. Subcutaneous microfabricated surfaces inhibit epithelial recession and promote long-term survival of percutaneous implants. Biomaterials 2002;23(1):229-37. 138.Ketabi M, Deporter D. The effects of laser microgrooves on hard and soft tissue attachment to implant collar surfaces: a literature review and interpretation. Int J Periodontics Restorative Dent 2013;33(6):e145-52. 139.Dumas V, Rattner A, Vico L, Audouard E, Dumas JC, Naisson P, Bertrand P. Multiscale grooved titanium processed with femtosecond laser influences mesenchymal stem cell morphology, adhesion, and matrix organization. J Biomed Mater Res A 2012;100(11):3108-16. 140.Ricci JL, Grew JC, Alexander H. Connective-tissue responses to defined biomaterial surfaces. I. Growth of rat fibroblast and bone marrow cell colonies on microgrooved substrates. J Biomed Mater Res A 2008;85(2):313-25. 141.Grew JC, Ricci JL, Alexander H. Connective-tissue responses to defined biomaterial surfaces. II. Behavior of rat and mouse fibroblasts cultured on microgrooved substrates. J Biomed Mater Res A 2008;85(2):326-35. 142.Ricci JL, Terracio L. Where is dentistry in regenerative medicine? Int Dent J 2011;61 Suppl 1:2-10. 143.Weiner S, Simon J, Ehrenberg DS, Zweig B, Ricci JL. The effects of laser microtextured collars upon crestal bone levels of dental implants. Implant Dent 2008;17(2):217-28. 144.Nevins M, Kim DM, Jun SH, Guze K, Schupbach P, Nevins ML. Histologic evidence of a connective tissue attachment to laser microgrooved abutments: a canine study. Int J Periodontics Restorative Dent 2010;30(3):245-55. 145.Neiva R, Tovar N, Jimbo R, Gil LF, Goldberg P, Barbosa JP, Lilin T, Coelho PG. The effect of laser-etched surface design on soft tissue healing of two different implant abutment systems: an experimental study in dogs. Int J Periodontics Restorative Dent 2016;36(5):673-9. 44 146.Nevins M, Nevins M, Gobbato L, Lee HJ, Wang CW, Kim DM. Maintaining interimplant crestal bone height via a combined platform-switched, Laser-Lok implant/abutment system: a proof-of-principle canine study. Int J Periodontics Restorative Dent 2013;33(3):261-7. 147.Nevins M, Nevins ML, Camelo M, Boyesen JL, Kim DM. Human histologic evidence of a connective tissue attachment to a dental implant. Int J Periodontics Restorative Dent 2008;28(2):111-21. 148.Nevins M, Camelo M, Nevins ML, Schupbach P, Kim DM. Connective tissue attachment to laser-microgrooved abutments: a human histologic case report. Int J Periodontics Restorative Dent 2012;32(4):385-92. 149.Geurs NC, Vassilopoulos PJ, Reddy MS. Histologic evidence of connective tissue integration on laser microgrooved abutments in humans. Clin Adv Periodontics 2011;1(1):29-33. 45 Figure 1.1. Schematic of the components of the OPRA (Osseointegrated Prostheses for the Rehabilitation of Amputees) system. The prosthetic device consists of a fixture which is implanted into the residual bone of the amputee patient. An abutment then connects the fixture to an abutment screw which finally serves as a docking site for external prostheses. This relives some of the common problems associated with socket prosthetics since the load is transferred directly to the bone instead of the soft-tissue. Figure from Ref. 42. Reproduced with permission of The Licensor through PLSclear: Branemark R, Berlin O, Hagberg K, Bergh P, Gunterberg B, Rydevik B. A novel osseointegrated percutaneous prosthetic system for the treatment of patients with transfemoral amputation: A prospective study of 51 patients. Bone Joint J 2014;96-B(1):106-13. Copyright © 2014, The British Editorial Society of Bone and Joint Surgery. All rights reserved. 46 Figure 1.2. Schematic of epidermal downgrowth progression around percutaneous devices. The top panel depicts epidermal downgrowth around smooth percutaneous devices, in which the epidermis migrates adjacent to the device surface and once it reestablishes continuity with the opposite side, the device is extruded from the body. The bottom panel depicts epidermal downgrowth around a porous coated device with a solid core. The epidermis migrates inside the pores of the device before migrating proximally while lysing the immature connective tissue. Eventually, the epidermis reaches the bottom of the device and the device is extruded. Adapted from Ref. 36. Adapted with permission from John Wiley and Sons, Inc.: von Recum AF. Applications and failure modes of percutaneous devices: a review. J Biomed Mater Res 1984;18(4):323-36. Copyright © 1984 John Wiley and Sons, Inc. All rights reserved. 47 Figure 1.3. Application of negative pressure wound therapy for 4 weeks over percutaneous devices limited epidermal downgrowth in a guinea pig model. Hematoxylin and eosin stained skin-device cross-sections of porous coated titanium subdermal devices. Regardless of the surgical approach tested, NPWT treated peri-prosthetic tissues showed significantly reduced downgrowth compared to the untreated controls. The average percent downgrowth values for the groups are indicated within each image. The yellow lines indicate the exposed porous coating. Scale bar - 500 μm. Figure from Ref. 104. Reproduced with permission from John Wiley and Sons, Inc.: Mitchell SJ, Jeyapalina S, Nichols FR, Agarwal J, Bachus KN. Negative pressure wound therapy limits downgrowth in percutaneous devices. Wound Repair Regen 2016;24(1):35-44. Copyright © 2015 by the Wound Healing Society. All rights reserved. 48 Figure 1.4. Progression of epidermal downgrowth with increased device in situ time. Hematoxylin and eosin stained cross-sections of porous coated titanium bone-anchored devices implanted in an ovine model from 0-, 3-, 6-, 9- and 12month time points. White arrows indicate the 3-point junction (point of intersection of skin, device and air) in all the sections. Although the porous coating reduced the risk of infection and rate of epidermal downgrowth, it was unable to completely halt downgrowth. Sinus tract formation is observed as a gap between the downgrowing epidermis and the device, in sections b, c and d. Figure from Ref. 5. Reproduced with permission from John Wiley and Sons, Inc.: Holt BM, Bachus KN, Beck JP, Bloebaum RD, Jeyapalina S. Immediate post-implantation skin immobilization decreases skin regression around percutaneous osseointegrated prosthetic implant systems. J Biomed Mater Res A 2013;101(7):2075-82. Copyright © 2012 Wiley Periodicals, Inc. All rights reserved. CHAPTER 2 PERI-PROSTHETIC TISSUE REACTION TO DISCONTINUATION OF NEGATIVE PRESSURE WOUND THERAPY AROUND POROUS TITANIUM PERCUTANEOUS DEVICES 1 2.1 Abstract Negative pressure wound therapy (NPWT) has been reported to limit epithelial downgrowth, one of the failure mechanisms of percutaneous devices. In a previous study, when NPWT was applied for 4 weeks (NPWT Group) to porous coated titanium percutaneous devices, downgrowth (5 ± 4%; mean ± one standard deviation) was significantly reduced compared to untreated controls (Untreated Group) (16 ± 6%; p ≤ 0.01). However, it was unclear whether this beneficial effect was sustained when NPWT was discontinued. In order to test this, porous coated titanium percutaneous devices were implanted into 6 hairless guinea pigs. Postsurgery, animals received 4 weeks of NPWT treatment followed by 4 weeks of no treatment (Discontinued Group). At necropsy, the devices and surrounding tissues 1 Reprinted with permission from John Wiley and Sons, Inc.: Pawar DRL, Mitchell SJ, Jeyapalina S, Hawkes JE, Florell SR, Bachus KN. Peri-prosthetic tissue reaction to discontinuation of negative pressure wound therapy around porous titanium percutaneous devices. J Biomed Mater Res B 2018. May 7. doi: 10.1002/jbm.b.34148. Copyright © 2018 Wiley Periodicals, Inc. All rights reserved. 50 were harvested and processed. Quantitative downgrowth measurements and qualitative analyses of tissue characteristics were performed, and compared to historical controls (NPWT and Untreated Groups). The Discontinued Group, at 8 weeks, had significantly more downgrowth than the NPWT Group at 4 weeks (23 ± 3% vs 5 ± 4%; p ≤ 0.01). At 8 weeks, the Discontinued Group qualitatively appeared to exhibit reduced numbers of blood vessels and increased degree of fibrosis compared to the NPWT Group at 4 weeks. This study suggests that NPWT will only be an effective treatment for limiting downgrowth if used continuously. 2.2 Introduction Percutaneous osseointegrated prosthetics have many benefits over suspension-type prosthetics, the current standard of care for amputee patients.1,2 However, percutaneous devices are subject to various failure mechanisms including infection and epithelial downgrowth.3 Downgrowth, also known as either marsupialization or skin regression,3,4 is an important concern for these devices since downgrowth can lead to sinus tract formation and subsequent device exposure, creating a portal for bacterial invasion thereby increasing the risk of infection.5 Thus, downgrowth may compromise the function of the device and its longevity.6 Previously, investigators have attempted to limit epithelial downgrowth by stabilizing the peri-prosthetic soft-tissues either by altering implant surface topography,7,8 by applying surface coatings,9 or by modifying device design.10 Porous devices7,9,11 and grooved devices8 have been shown to limit epithelial 51 downgrowth and to promote skin-implant integration in animal models. In particular, porous devices have been shown to allow the in-growth of vascularized soft tissue within their pores, and have been reported as showing reduced epidermal migration rates in translational studies.3,7 However, the inability to completely eliminate downgrowth based solely on topography has motivated researchers to find alternative solutions.12 A recent study has now suggested that negative pressure wound therapy (NPWT) may limit epithelial downgrowth while being applied.13 This study measured downgrowth of NPWT treated tissues over a 4-week period to be 5 ± 4% (mean ± one standard deviation) compared with 16 ± 6% for untreated controls (p ≤ 0.01).13 However, the application of NPWT is clinically impractical as a long-term treatment option. Thus, there is a need to understand whether the application of a short course of NPWT can have longlasting consequences on epithelial downgrowth after it has been discontinued. Clinically, NPWT is used to accelerate healing in acute and chronic wounds.14,15 The mechanisms of action include mechanically drawing the wound margins together, inducing rapid formation of granulation tissue, and stimulating angiogenesis in the wound bed.14-18 Although, these mechanisms might have been responsible for the successful soft-tissue outcomes in the previous study,13 the effect of discontinuing NPWT on the peri-prosthetic soft-tissue morphology and epithelial downgrowth is currently unknown. NPWT is discontinued after wound closure has been achieved or when changes in wound margin measurements have plateaued.14,18 Although the duration of treatment varies, NPWT is rarely applied clinically for > 3 months.14 52 However, the use of NPWT in combination with percutaneous devices differs from typical clinical scenarios, in that the skin seal is permanently breached due to the percutaneous post, so the wound never attains full closure. Therefore, it is difficult to determine the exact moment when stabilization of the peri-prosthetic soft-tissues is achieved, and whether or not epithelial downgrowth will persist when the application of NPWT is discontinued. The goal of this study was to investigate the effect of discontinuing NPWT on epithelial downgrowth and peri-prosthetic tissue morphology. It was hypothesized that the limited epithelial downgrowth observed around the percutaneous devices after an initial, 4-week application of NPWT would be maintained when the therapy was discontinued. 2.3 Materials and Methods 2.3.1 Animal model The study followed an approved animal protocol from the Institutional Animal Care and Use Committee at the Comparative Medicine Center, University of Utah and by the Animal Care and Use Review Office, a component of the US Army Medical Research Materiel Command, Office of Research Protections. A total of six, 4-week old, female Institute Armand Fappier (IAF) hairless Guinea pigs (~200 g in weight; strain code 161; Charles River, Kingston, NY) were used. 53 2.3.2 Device design and surgical procedure Each animal was implanted with a historically successful percutaneous device.13 Briefly, the percutaneous devices consisted of a subdermal device and a percutaneous post (Figure 2.1), which were fabricated using titanium alloy (Ti6Al4V). The subdermal device was further treated with commercially pure titanium porous coating (K-coating, Thortex Inc., Portland, OR). The pore size distribution of the coating was reported by the manufacturer to be 230-500µm. Scanning electron microscopic imaging (Quanta 600 FEG, FEI, Hillsboro, OR; High Vacuum Mode; accelerating voltage: 15 kV) was also performed to visualize the topography of the porous coating. The devices were passivated according to the ASTM F86 standard and steam sterilized prior to implantation. A previously successful Two-Stage procedure was used for the implantation of the device.13 Briefly, the first part of the Two-Stage surgery entailed implantation of the subdermal device under the dorsal skin. During the Stage 1 surgery, the dorsal skin was prepared for aseptic surgery, and then a 4 cm longitudinal incision (0.5 cm left of the spinal column and in the cranial to caudal direction) was made to expose the underlying soft tissues. Blunt dissection was then used to create a subdermal pocket, into which the porous coated device was placed. The incision was sutured closed using 4-0 Vicryl sutures (Ethicon, Hamburg, Germany) and the animal was allowed to recover and heal for 3 weeks. During the Stage 2 aseptic surgery, the percutaneous post was attached to the subdermal device. After preparing the dorsal skin for aseptic surgery, the center of the subdermal disk was located and a 4-mm circular biopsy punch was 54 used to remove tissue over the subdermal tap hole to allow for insertion of the percutaneous post. The post was hand tightened to the device followed by cleaning of the site with sterile gauze soaked in saline. Before recovery, all animals received a base dressing consisting of sterile gauze which was placed over the incision and the percutaneous post. The gauze was then covered and secured into place using a semi-occlusive dressing (Tegaderm™ Film, Saint Paul, MN). A NPWT delivery line consisting of a tube, a neoprene rubber washer, and a stainless steel retaining ring was inserted into the base dressing. The tube was further secured into place by using semi-occlusive dressings to create an airtight seal. After surgery, animals were housed singly in custom-designed, clear polycarbonate cylindrical cages (Model#RFSCW22, CAMBRO, Huntington Beach, CA) with a wall-mounted water bottle and a food bowl placed inside the cage. The NPWT delivery tube was connected from the dressing to the removable lid, which was custom designed to allow the animal to have unrestrained movement inside the cage. 2.3.3 NPWT treatment Following recovery, the peri-prosthetic soft tissues were treated with NPWT using a vacuum pump (Renasys Go, Smith & Nephew Inc., St. Petersburg, FL) that was set to continuously apply 80-90 mmHg of vacuum. Pressures were monitored using an external pressure gauge (Model DPG-4000-15C-RM, Omega®, Stamford, CT) that was connected to the NPWT delivery line using a tconnector. Dressings were changed either at 48-h intervals, or earlier based upon 55 the condition of the dressing. During animal checks (which occurred twice a day), each dressing was visually inspected and the status of the dressings (i.e. intact, breached or completely detached from the site) as well as the pressure gauge reading was noted. These recordings allowed for the calculation of dressing longevity for each animal. The pressure gauge was also used as an objective measure to determine if the dressing was intact or breached. If the dressing was breached or the gauge did not reflect the target pressure then the dressing was changed while maintaining the animals under general isoflurane anesthesia. The NPWT treatment was applied for 4 weeks similar to previous reports,13 after which, the treatment was discontinued. Four weeks after discontinuation of the NPWT, the animals were sacrificed. In total, the animals remained in the study for 8 weeks after the Two-Stage surgery. This group is referred to as the “Discontinued Group”. The downgrowth measurements from these animals were compared to the downgrowth measurements of groups from a previously published study,13 in which the NPWT Group (N = 5) received 4 weeks of NPWT treatment and the Untreated Group (N = 5) received no treatment for 4 weeks. Both the NPWT and Untreated Groups underwent the Two-Stage surgical procedure. 2.3.4 Sample collection and processing At necropsy, each device with the surrounding soft-tissue was harvested, fixed in Karnovsky’s fixative, dehydrated with ascending grades of ethanol, and then embedded in poly (methyl) methacrylate (PMMA) using an established procedure.19 After polymerization, ~2 mm transverse sections through the post, in 56 the medial-lateral direction, were obtained using a precision saw (LS10 Lapidary Slap Saw, Lortone Inc. Mukilteo, WA). These sections were then ground to ~5070 µm thickness and polished to an optical finish (Buehler Metaserv™ 250, Lake Bluff, IL). The slides were stained with modified Trichrome.20 Briefly, slides were stained with Modified Weigert’s Hematoxylin for 8 min and then blued in running lukewarm tap water for 20 min. The slides were then immersed in 1% Neutral Red solution (Thermo-Fisher Scientific, Waltham, MA) for 10 minutes. Finally, they were stained with 1% Light Green (Sigma-Aldrich, St. Louis, MO) solution for 5 min. Excess stain was removed using lint-free disposable Kimwipes and 70% ethanol. Skin samples from two separate guinea pigs were collected away from the device and analyzed histologically for baseline measurements. These slides were stained using a standard Hematoxylin & Eosin (Sigma-Aldrich, St. Louis, MO) staining procedure. The skin-device interface was analyzed using a transmitted light microscope (Nikon Instruments INC, Melville, NY) with an attached DS-Vi1 Color digital camera (MQA12010, Nikon Instruments INC, Melville, NY). Still images were captured using an imaging software package (NIS Elements, version 4.0, Nikon Instruments, Inc., Melville, NY) and processed using Photoshop (Version 2015, Adobe Systems Inc., San Jose, CA). 2.3.5 Histological analysis Using three histological landmarks, downgrowth was measured following an established method.13 The first landmark was the attachment point of the post 57 and the subdermal device. The second landmark was the attachment point of the epidermis to the device (i.e. 3-point junction) and the third landmark marked the edge of the subdermal portion of the device. The total length of the porous coating was measured by tracing the outer profile of the porous coating from the first to the third landmarks. The exposed coating length was measured from the first to the second landmarks. Downgrowth was calculated as the percentage of the exposed coating length to the total length of the porous coating. Downgrowth measurements were obtained from both sides of the post in each sample and averaged. Downgrowth data from the Discontinued Group at 8 weeks were compared to the previously reported downgrowth measurements from the NPWT Group and Untreated Group at 4 weeks.13 Histopathological analyses were performed on sections from all three groups: NPWT, Untreated and Discontinued Groups. The peri-prosthetic tissues were blindly assessed by a dermatopathologist, who was asked to record the number of blood vessels, tissue morphology, and presence of inflammatory infiltrate in each sample. In addition, the dermatopathologist provided quantitative assessment of the epidermal thickness for normal skin and the epidermis at the 3point junction. The number of blood vessels was qualitatively assessed at three locations: peri-prosthetic tissues adjacent to the 3-point junction (i.e. within the granulation tissue), within the porous coating along the slope of the device and at the vertical edge of the device (Figure 2.2). The tissue morphology for each treatment group was qualitatively characterized based on the number of blood vessels within the granulation tissue and the degree of fibrosis. Granulation tissue 58 was identified as a hypercellular region with unorganized matrix. For each sample, quantitative analysis of epidermal thickness was performed, in which, the length of the thickest point of the epidermis was measured from the stratum basal layer to the granular layer. Measurements from each group were averaged. Inflammation was said to be present if inflammatory infiltrates were seen at the 3-point junction adjacent to the exposed parts of the device. 2.3.6 Statistical analysis Independent, two-tailed Student t-Tests were performed (Microsoft Office, Excel, 2013) to compare groups and quantify significance in epithelial downgrowth and epidermal thickness measurements. The level of significance was set at 5% for all statistical analyses. All measures were reported as mean ± one standard deviation. 2.4 Results In general, administration of the NPWT treatment was tolerated by all animals without complication or abnormal changes in animal behavior. On average, animals maintained each dressing for 41 ± 6 h. The percutaneous exit sites did not show any clinical signs of infection, redness, or discharge over the duration of the study. One animal died for unknown reasons at 30 days post second surgery and was excluded from the study. N = 5 samples were histologically processed. One side of a sample slide in the Discontinued Group contained a processing artifact and the whole of this slide was excluded from our 59 histological analyses. Therefore, N = 4 total samples were used for histological analyses of the Discontinued Group. After 4 weeks of NPWT followed by 4 weeks of no treatment, the Discontinued Group showed 23 ± 3% downgrowth compared to the NPWT Group at 4 weeks (5 ± 4%; p ≤ 0.001).13 Compared to the Untreated Group at 4 weeks (16 ± 6%),13 the Discontinued Group did not show any significant difference in downgrowth (p = 0.09). Overall, upon discontinuation of NPWT, the tissues appeared to continue to undergo downgrowth. Qualitatively, there were differences in the peri-prosthetic soft-tissues for all three groups. Over the device as a whole, the Discontinued Group showed decreased vascularization (i.e. number of blood vessels) when compared to the NPWT Group, but was similar to the Untreated Group. Within the Discontinued and Untreated Groups, the peri-prosthetic tissues at the edge of the device appeared to have the highest number of small blood vessels [Figure 2.3(I,G)]. The tissues adjacent to the 3-point junction and within the porous coating along the slope of the device displayed a similar number of blood vessels [Figure 2.3(A,C,D,F)]. Within the NPWT samples, the tissues adjacent to the 3-point junction showed a relatively higher number of small blood vessels followed by the tissues at the edge of the device while the tissues along the slope showed the least number of vessels [Figure 2.3(B,E,H)]. In addition, the peri-prosthetic tissues for the Discontinued and Untreated Groups showed relatively more fibrosis within the granulation tissue while the NPWT treated tissues appeared to be less fibrotic. Overall, the tissues in the Discontinued and Untreated Groups showed less granulation tissue with 60 reduced numbers of blood vessels and increased fibrosis compared to the NPWT Group. All three groups appeared to have acanthosis at the 3-point junction (Figure 2.4; Table 2.1). While the NPWT and Untreated Groups showed significantly more acanthosis compared to normal epidermis (p ≤ 0.003 and p ≤ 0.0002, respectively), the Discontinued Group did not show significant acanthosis compared to normal epidermis (p = 0.12). Comparisons within the 3 groups showed no statistical difference in the degree of acanthosis between the Untreated and the NPWT Groups (p = 0.18), NPWT and Discontinued Groups (p = 0.44), and Discontinued and Untreated Groups (p = 0.93). External to the 3-point junction, a neutrophil-rich hemorrhagic crust was present in all the groups (Figure 2.5). 2.5 Discussion In this study, we hypothesized that upon discontinuation of NPWT, the reduced epithelial downgrowth observed around a percutaneous device after 4 weeks of NPWT would be maintained. The data from this study suggest that epithelial downgrowth continued after termination of NPWT. Therefore, the results of our study failed to support hypothesis. Moreover, our data suggest that 4 weeks after discontinuing NPWT, the amount of epithelial downgrowth was comparable to the Untreated Group after 4 weeks, without NPWT. The data available suggest that NPWT only limited downgrowth while being applied. While the mechanism of action for this outcome is unclear, there are some clues from the literature. Previously, studies have shown that in a typical wound 61 (i.e. without a device), NPWT induces wound shrinkage through macrodeformations and by mechanically pulling the edges together.14-16 In contrast, our model is an atypical wound, where the percutaneous post permanently protrudes through the skin, which physically prevents the wound edges from converging. Even in this atypical wound, macro-deformations and mechanical forces due to NPWT might be responsible for limiting epithelial downgrowth by preventing the wound edges from migrating proximally. However, after discontinuing NPWT, the normal trajectory of the epithelial cells simply resumed, leading to the progression of downgrowth. Our study also confirmed some histological parallels and differences between the current and previous data. Our histology data showed that the NPWT treated peri-prosthetic tissues have the characteristics of an actively healing wound with relatively more granulation tissue, increased numbers of blood vessels [Figure 2.3(B,E,H)], and decreased fibrosis. This is similar to the results of a study which showed that NPWT changes the micro-environment of a wound into an acute wound.21 Further, the tissue features of the NPWT Group are similar to results in clinical wounds which have shown that NPWT increases blood flow and stimulates granulation tissue formation, essential for wound healing.15,22 An increased blood supply to the peri-prosthetic tissues might have been one of the reasons why the NPWT Group showed the least amount of epithelial downgrowth. Blood supply is important for bringing vital nutrients and oxygen to support wound healing.18,23 Moreover, Erba et al. showed that the formation of increased blood vessels, in NPWT treated wounds compared to untreated wounds, was associated 62 with improved wound closure.17 In the case of percutaneous devices treated with NPWT, increased blood vessels in combination with the mechanical forces pulling the wound edges towards the post might have led to limited epithelial downgrowth. Taken together, these results offer some insight into possible mechanisms of action of NPWT on peri-prosthetic tissues and its ability to limit downgrowth in percutaneous devices while being applied. In contrast, qualitative analysis of the tissues in both the Discontinued and Untreated Groups showed reduced granulation tissue with relatively fewer numbers of blood vessels, and increased fibrosis when compared to the NPWT Group (Figure 2.3). These tissue characteristics are similar to those of a chronic wound. An inadequate blood supply and the presence of a percutaneous device18 are some of the many reasons why a wound might deviate from the expected trajectory for healing. It is possible that the morphological changes that occurred during NPWT application in the Discontinued Group were not sustained after its termination, which instead led to the development of fibrosis. Alternatively, matrix remodeling that occurred with increased healing time (i.e. 4 weeks) could also have resulted in increased levels of fibrosis. Fibrosis around percutaneous devices can reduce the longevity of the implants by increasing their susceptibility to infection and epithelial downgrowth.24 In addition, fibrosis could also lead to implant extrusion3 which may explain increased device exposure and epithelial downgrowth. Our qualitative data indicated that discontinuing NPWT resulted in a reduced number of blood vessels compared to the NPWT Group, but a similar 63 number of blood vessels compared to the Untreated Group. This suggests that blood vessel formation, induced by NPWT application, diminished upon removal of NPWT. The literature on vascularization has indicated that more blood vessels are present in NPWT treated tissues compared to untreated tissues.17 However, there have been no previous studies regarding the morphological changes in the tissues following discontinuation of NPWT. Therefore, our data have yielded some insights into the changes in vascularity occurring in the peri-prosthetic tissues after discontinuation of NPWT. External to the 3-point junction, and adjacent to the implant surface, a hemorrhagic neutrophilic crust was apparent in most samples (Figure 2.5). Similar findings have been reported for other percutaneous devices, such as dental implants25 and bone anchored hearing aids.26 These studies reported that the inflammatory neutrophilic crust was present for prolonged durations.25,26 In dental implants, inflammatory infiltrates consisting of neutrophils have been associated with tissue destruction,27 and this could explain the destruction of the periprosthetic tissues observed with our implants, which might have led to epithelial downgrowth. At the 3-point junction, thickened epidermis (acanthosis/hyperplastic) was present in most samples (Figure 2.4). Significant differences in epidermal thickness were observed between Untreated and NPWT samples at 4 weeks compared to normal epidermis. In open human wounds, the epidermis at the wound edge thickens within 24 h of injury.28 However, by 48 h the epidermis at the leading wound edge thins and becomes migratory in nature to close the defect.29 64 On the other hand, a study showed that in rodents undergoing wound healing by contraction, the epidermis at the wound edge thickens and remains thick throughout the process of wound healing.30 Collectively, this suggests that the thickened epidermis (acanthosis) found in the NPWT and Untreated Group at 4 weeks might be due to the inherent contractile wound healing mechanisms of a guinea pig around the percutaneous post. Interestingly, no significant difference in epidermal thickness was found between the Discontinued Group at 8 weeks and normal epidermis. This might have been due to the increased healing time which led to thickening followed by eventual thinning of the epidermis, which then returned to baseline levels. Between the three groups, there were no significant differences in epidermal thickness. From our data, we have concluded that it is the innate wound healing mechanism of guinea pigs elicited by the presence of percutaneous post, rather than treatment with NPWT, that may have resulted in acanthosis. Our study had several limitations. First, there are physiological differences in wound healing between guinea pigs and humans.31 Guinea pigs have a panniculus carnosus muscle which is absent in humans. Therefore, the primary mechanism for wound healing in guinea pigs is contraction rather than reepithelialization.32 Despite this, use of a guinea pig model has provided some insights into ways of limiting epithelial downgrowth that could be further validated in more translatable animal models such as pigs.33 Second, our study did not include an 8-week untreated control or 8 week NPWT control groups. Although inclusion of these groups would have been ideal, we decided that this would have 65 resulted in an unnecessary use of animals based on previous experimental outcomes.34 Additionally, the purpose of the current study was to study the effects of discontinuation of NPWT on epithelial downgrowth and not to determine the efficacy of NPWT over longer durations. Third, use of PMMA for embedding our soft-tissue/metal specimens made it difficult to perform full histological analysis due to the inability of immunohistochemical stains to penetrate the plastic sections at room temperature. Investigation of new embedding techniques will be needed to elucidate the molecular mechanisms responsible for epithelial downgrowth and the tissue response to NPWT, but fall outside the scope of this study. Lastly, the percutaneous device was not anchored in bone. Studies have indicated that the downgrowth phenomena has been observed in both bone-anchored,34 and subdermal barrier only percutaneous implants.35 While differences in interfacial forces between bone-anchored and subdermal barrier only percutaneous implants might influence the overall rate of epithelial downgrowth, the literature implies that regardless of the fixation method, the downgrowth observed around the percutaneous devices is due to epithelial migration associated with fundamentals of wound healing.3,9 Since, the goal of this study was to investigate the utility of a short-term treatment of NPWT to limit epithelial downgrowth, it was considered reasonable to use the subdermal barrier only percutaneous implants. In summary, our study suggests that the positive effects of NPWT in limiting epithelial downgrowth can only be maintained when used continuously. This is clearly a major drawback of NPWT application for percutaneous devices. Our study also examined some of the histological changes associated with the 66 application and termination of NPWT. While NPWT was in use, the tissues had the appearance of an actively healing wound, but after its discontinuation, the tissues changed their morphological appearance to resemble a chronic or non-actively healing wound. Although a previous study has shown that NPWT is effective in limiting epithelial downgrowth,13 the continuous use of NPWT is impractical. However, further investigations of NPWT as a means of assisting the integration of percutaneous devices during the early and critical periods after implantation may be of value. 2.6 Acknowledgements The views, opinions, and/or findings presented are those of the authors and should not be construed as an official position, policy or decision of any of these funding sources unless so designated by other documentation. This publication made use of the University of Utah Shared facilities of the Surface Analysis and Nanoscale Imaging group sponsored by the College of Engineering, Health Sciences Center, Office of the Vice President for Research, and the Utah Science Technology and Research (USTAR) Initiative of the State of Utah. In conducting research using animals, the investigators adhered to the Animal Welfare Act Regulations and other Federal statutes relating to animals and experiments involving animals and the principles set forth in the current version of the Guide for Care and Use of Laboratory Animals, National Research Council. All authors confirm that there is no potential conflict of interest including employment, stock ownership, consultancies, honoraria, paid expert testimony, 67 and patent applications/registrations influencing this work. 2.7 References 1.Baars EC, Dijkstra PU, Geertzen JH. Skin problems of the stump and hand function in lower limb amputees: a historic cohort study. Prosthet Orthot Int 2008;32(2):179-85. 2.Frossard L, Hagberg K, Haggstrom E, Gow DL, Branemark R, Pearcy M. Functional outcome of transfemoral amputees fitted with an osseointegrated fixation: temporal gait characteristics. J Prosthet Orthot 2010;22(1):11-20. 3.von Recum AF. Applications and failure modes of percutaneous devices: a review. J Biomed Mater Res 1984;18(4):323-36. 4.Winter GD. Transcutaneous implants: reactions of the skin-implant interface. J Biomed Mater Res 1974;8(3):99-113. 5.Fleckman P, Olerud JE. Models for the histologic study of the skin interface with percutaneous biomaterials. Biomed Mater 2008;3(3):034006. 6.Heaney TG, Doherty PJ, Williams DF. Marsupialization of percutaneous implants in presence of deep connective tissue. J Biomed Mater Res 1996;32(4):593-601. 7.Jeyapalina S, Beck JP, Bachus KN, Williams DL, Bloebaum RD. Efficacy of a porous-structured titanium subdermal barrier for preventing infection in percutaneous osseointegrated prostheses. J Orthop Res 2012;30(8):1304-11. 8.Chehroudi B, Gould TR, Brunette DM. Effects of a grooved epoxy substratum on epithelial cell behavior in vitro and in vivo. J Biomed Mater Res 1988;22(6):459-73. 9.Pendegrass CJ, Goodship AE, Blunn GW. Development of a soft tissue seal around bone-anchored transcutaneous amputation prostheses. Biomaterials 2006;27(23):4183-91. 10.Pendegrass CJ, Gordon D, Middleton CA, Sun SN, Blunn GW. Sealing the skin barrier around transcutaneous implants: in vitro study of keratinocyte proliferation and adhesion in response to surface modifications of titanium alloy. J Bone Joint Surg Br 2008;90(1):114-21. 11.Isackson D, Cook KJ, McGill LD, Bachus KN. Mesenchymal stem cells increase collagen infiltration and improve wound healing response to porous titanium percutaneous implants. Med Eng Phys 2013;35(6):743-53. 68 12.Stynes G, Kiroff GK, Morrison WA, Kirkland MA. Tissue compatibility of biomaterials: benefits and problems of skin biointegration. ANZ J Surg 2008;78(8):654-9. 13.Mitchell SJ, Jeyapalina S, Nichols FR, Agarwal J, Bachus KN. Negative pressure wound therapy limits downgrowth in percutaneous devices. Wound Repair Regen 2016;24(1):35-44. 14.Huang C, Leavitt T, Bayer LR, Orgill DP. Effect of negative pressure wound therapy on wound healing. Curr Probl Surg 2014;51(7):301-31. 15.Orgill DP, Manders EK, Sumpio BE, Lee RC, Attinger CE, Gurtner GC, Ehrlich HP. The mechanisms of action of vacuum assisted closure: more to learn. Surgery 2009;146(1):40-51. 16.Argenta LC, Morykwas MJ. Vacuum-assisted closure: a new method for wound control and treatment: clinical experience. Ann Plast Surg 1997;38(6):56376; discussion 577. 17.Erba P, Ogawa R, Ackermann M, Adini A, Miele LF, Dastouri P, Helm D, Mentzer SJ, D'Amato RJ, Murphy GF and others. Angiogenesis in wounds treated by microdeformational wound therapy. Ann Surg 2011;253(2):402-9. 18.Bollero D, Driver V, Glat P, Gupta S, Lazaro-Martinez JL, Lyder C, Ottonello M, Pelham F, Vig S, Woo K. The role of negative pressure wound therapy in the spectrum of wound healing. Ostomy Wound Manage 2010;56(5 Suppl):118. 19.Emmanual J, Hornbeck C, Bloebaum RD. A polymethyl methacrylate method for large specimens of mineralized bone with implants. Stain Technol 1987;62(6):401-10. 20.Betz DH, Epperson RT, Holt BM, Bloebaum RD, Jeyapalina S. A new trichrome technique for PMMA embedded percutaneous implants for the study and characterization of epithelial integration. Journal of Histotechnology 2012;35(4):164-170. 21.Lalezari S, Lee CJ, Borovikova AA, Banyard DA, Paydar KZ, Wirth GA, Widgerow AD. Deconstructing negative pressure wound therapy. Int Wound J 2016. 22.Morykwas MJ, Argenta LC, Shelton-Brown EI, McGuirt W. Vacuumassisted closure: a new method for wound control and treatment: animal studies and basic foundation. Ann Plast Surg 1997;38(6):553-62. 23.Tonnesen MG, Feng X, Clark RA. Angiogenesis in wound healing. J Investig Dermatol Symp Proc 2000;5(1):40-6. 69 24.Isackson D, McGill LD, Bachus KN. Percutaneous implants with porous titanium dermal barriers: an in vivo evaluation of infection risk. Med Eng Phys 2011;33(4):418-26. 25.Broggini N, McManus LM, Hermann JS, Medina R, Schenk RK, Buser D, Cochran DL. Peri-implant inflammation defined by the implant-abutment interface. J Dent Res 2006;85(5):473-8. 26.Asma A, Ubaidah MA, Hasan SS, Wan Fazlina WH, Lim BY, Saim L, Goh BS. Surgical outcome of bone anchored hearing aid (baha) implant surgery: a 10 years experience. Indian J Otolaryngol Head Neck Surg 2013;65(3):251-4. 27.Broggini N, McManus LM, Hermann JS, Medina RU, Oates TW, Schenk RK, Buser D, Mellonig JT, Cochran DL. Persistent acute inflammation at the implant-abutment interface. J Dent Res 2003;82(3):232-7. 28.Odland G, Ross R. Human wound repair. I. Epidermal regeneration. J Cell Biol 1968;39(1):135-51. 29.Patel GK, Wilson CH, Harding KG, Finlay AY, Bowden PE. Numerous keratinocyte subtypes involved in wound re-epithelialization. J Invest Dermatol 2006;126(2):497-502. 30.Lemo N, Marignac G, Reyes-Gomez E, Lilin T, Crosaz O, Dohan Ehrenfest DM. Cutaneous reepithelialization and wound contraction after skin biopsies in rabbits: a mathematical model for healing and remodelling index. Veterinarski arhiv 2010;80(5):637-652. 31.Rittie L. Cellular mechanisms of skin repair in humans and other mammals. J Cell Commun Signal 2016;10(2):103-20. 32.Charlton CA, Higton DI, James DW, Nicol AR, Stewart JO. Wound contraction in the guinea-pig. Br J Surg 1961;49:96-102. 33.Sullivan TP, Eaglstein WH, Davis SC, Mertz P. The pig as a model for human wound healing. Wound Repair Regen 2001;9(2):66-76. 34.Holt BM, Bachus KN, Beck JP, Bloebaum RD, Jeyapalina S. Immediate post-implantation skin immobilization decreases skin regression around percutaneous osseointegrated prosthetic implant systems. J Biomed Mater Res A 2013;101(7):2075-82. 35.Holt BM, Betz DH, Ford TA, Beck JP, Bloebaum RD, Jeyapalina S. Pig dorsum model for examining impaired wound healing at the skin-implant interface of percutaneous devices. J Mater Sci: Mater Med 2013;24:2181-2193. 70 Figure 2.1. (A) A photograph showing the porous coated subdermal and smooth device percutaneous post. (B) A representative image of scanning electron micrograph showing the topographical features of the porous coating (accelerating voltage: 15 kV; magnification: X100). Pore size ranges from ~230-500 µm (manufacturer’s specification). White scale bar= 1 mm. 71 Figure 2.2. A representative photomacrograph showing the three regions used for qualitative analyses. The region within the white solid box indicates the granulation tissue adjacent to the 3-point junction (point of epidermal attachment to device), the dashed white box along the slope of the device represents tissue within the pores and the vertical small dash white box represents tissue at edge of the device. White scale bar= 500 µm. 72 Figure 2.3. Qualitative differences in blood vessels were seen between the groups. The Untreated Group showed relatively lower number of blood vessels adjacent to the 3-point junction (A), along the length of the device (D) and at the edge of the device (G). The NPWT Group (B, E, H) showed a relatively higher number of blood vessels compared to the Untreated and Discontinued Groups. The Discontinued Group revealed a similar number of blood vessels (C, F, I) compared to the Untreated Group. All images are at the same magnification of X10. White scale bar= 200 µm. White arrows in each image represent location of a subset of blood vessels. 73 Figure 2.4. A representative set of photomicrographs showing (A) normal epidermal thickness and epithelial tissues at the 3-point junction for the (B) Untreated Group, (C) NPWT Group, and (D) Discontinued Group. White arrows depict the thickness of the healthy/acanthotic region. All images are at the same magnification of X10 with black and white scale bar= 200 µm. 74 Figure 2.5. A representative photomicrograph of the neutrophilic inflammatory crust (area within dash line) present externally at the 3-point junction (marked by *) in all the groups adjacent to the device and percutaneous post. Image at magnification of X20 with white scale bar = 100 µm. 75 Table 2.1. Epidermal thickness measurements near the 3-point junction and healthy epithelium. Data showed no significant difference in epidermal thickness within the three groups at the 3-point junctions. However, there were significant differences in thickness at the 3-point junction in the Untreated and NPWT Groups compared to healthy epidermis (p ≤ 0.0002 and p ≤ 0.003, respectively), indicating presence of a hyperplastic (acanthosis) epithelium at 3-point junction in these groups. Treatment Thickness (µm) Normal Epidermis 70 ± 30 Untreated 180 ± 20 NPWT 230 ± 70 Discontinued 180 ± 120 CHAPTER 3 EVALUATION OF SOFT-TISSUE RESPONSES AROUND LASER MICROGROOVED TITANIUM PERCUTANEOUS DEVICES 3.1 Abstract Percutaneous devices are prone to epidermal downgrowth and sinus tract formation, which can serve as a nidus for bacterial colonization and increase the risk of infection. Laser-Lok® microgrooved topography has been shown to limit gingival epithelial downgrowth around dental implants. However, the efficacy of the microgrooved topography to limit epidermal downgrowth around non-gingival percutaneous devices is yet to be investigated. In order to bridge this gap, devices with a percutaneous post and a porous coated subdermal component were designed. The proximal 2 mm section of the post was textured with either microgrooved, porous or left untextured (smooth). The porous and smooth topographies served as controls. The devices were tested in a guinea pig back model, where 18 hairless guinea pigs were randomly assigned into three groups with each group receiving one topography design (N = 6/group). Four weeks postimplantation, the devices with surrounding soft-tissues were harvested and processed for histological analyses. Results indicated that the microgrooved topography failed to prevent epidermal downgrowth (23 ± 4%) around 77 percutaneous devices in this model. Furthermore, no significant difference (p = 0.70) in epidermal downgrowth was present between the three topographies, with all the groups exhibiting similar degrees of downgrowth. Overall, these findings suggest that the microgrooved topography may be unable to halt downgrowth around percutaneous devices for dermal applications. 3.2 Introduction Percutaneous devices are prone to many types of failure, which includes epidermal downgrowth.1 Epidermal downgrowth can be attributed to the reepithelialization phase of wound healing and is defined as the apical migration of epidermal cells along the device surface, leading to device exposure, sinus tract formation and increased susceptibility to infection.1-3 In order to achieve long-term clinical success of permanent percutaneous devices, it is important to establish an infection-free, stable, skin-device interface with limited downgrowth. To establish a skin-device seal, porous surface topography has been used in translational animal studies4-7 and in early European human clinical trials.8,9 The results from the animal studies showed potential for promoting skin-device integration,7,10 limiting the rate of epidermal downgrowth4 and decreasing the risk of infection.5,7 In a 9-month study, Holt et al. showed that percutaneous osseointegrated devices with a porous coated subdermal barrier significantly reduced downgrowth compared to smooth percutaneous devices in an ovine model.4 However, a 2-year follow-up study reported that while these porous coated percutaneous devices did show fibrous tissue-device integration, the porous 78 topography was unable to completely halt downgrowth over an extended period of time.6 Results from the early clinical trials suggested that while the incorporation of a porous surface topography reduced epidermal downgrowth, it caused softtissue irritation at the skin-device interface, leading to infection.11 Since then, the device design has been modified to remove the abrasive porous coating at the skin-device interface, leading to a reduction in the rate of infection.11-13 Currently, European prosthetic groups utilize a smooth surface topography at the skin-device interface in percutaneous osseointegrated devices.11,13,14 Utilizing a smooth topography offers benefits such as a relatively easy cleaning regime and reduced soft-tissue irritation caused by device-induced mechanical abrasion.8,11 Additionally, it has been theorized that inclusion of a smooth percutaneous surface increases the rate of epidermal downgrowth and allows the epidermis to attach to the underlying bone, which presumably creates a biological seal with the bone.11 However, the use of smooth surfaces in percutaneous osseointegrated devices has been reported to result in a sinus tract formation possibly due to the inability to establish a robust skin-device seal to the smooth surface. Sinus tract formation may subsequently lead to a high rate of infection. While superficial infections are the most common and can be typically treated with antibiotics,15,16 there is a risk of developing deep infections which can result in the removal of the entire prosthetic system.15,17 Thus, the inability of the smooth and porous topographies to establish a robust skin-device seal motivates the investigation of alternate topographies to halt epidermal downgrowth around percutaneous devices. 79 Dental implants, which are an example of percutaneous osseointegrated devices, are challenged by gingival epithelial downgrowth.18,19 The lack of softtissue-implant integration contributes to the gingival epithelial downgrowth and can allow for the accumulation of bacterial and inflammatory cell infiltrates at the skinimplant interface, leading to peri-implantitis.20 Peri-implantitis can further result in the apical loss of soft-tissue and bone, and eventual device exposure with increasing implant in situ time.21 Therefore, establishing a robust epithelial seal at the tissue-implant interface is important to enhance the efficacy of dental implants.18 Recent studies have shown that application of a microgrooved topography, known as Laser-Lok®, has shown a high success rate in limiting gingival epithelial downgrowth around dental implants.22,23 The Laser-Lok® topography is developed on the surface of titanium dental implants, by a laser ablation technique that etches three-dimensional microgrooves with microscale texturing within the grooves.22 Studies have demonstrated that 8 μm sized microgrooves, placed circumferentially over a 2 mm wide band on the abutment portion of the dental implants, are effective in directing epithelial cell migration within the grooves, resulting in limited gingival epithelial downgrowth.23-25 While the Laser-Lok® microgrooved topography has shown success in restricting gingival epithelial downgrowth around dental implants, it is yet to be tested in other non-gingival percutaneous devices. Intrinsic differences between gingival epithelial cells and normal skin epidermal cells26 may influence the ability of the Laser-Lok® topography to limit epidermal downgrowth. Therefore, there is a need to evaluate the microgrooved topography in normal skin. 80 The goal of this study was to evaluate the efficacy of percutaneous devices with Laser-Lok® topography to limit epidermal downgrowth. The microgrooved percutaneous device was tested in an established subdermal guinea pig model.27,28 Porous and smooth topographies were used as controls. We hypothesized that similar to the Laser-Lok® dental implants, the microgrooved topography would limit epidermal downgrowth around percutaneous devices implanted in epidermal tissues. Additionally, the microgrooved devices would show the least amount of downgrowth compared to the porous and the smooth devices. 3.3 Materials and Methods 3.3.1 Implant fabrication Percutaneous devices consisted of two design components: a cylindrical percutaneous post and a subdermal device (Thortex Inc., Portland, OR). The percutaneous post contained either a smooth (Figure 3.1C), microgrooved (Figure 3.1A) or porous topography (Figure 3.1B). To obtain the smooth topography, the percutaneous posts were machined to obtain a smooth finish. To obtain the LaserLok® technology, referred to as the “microgrooved topography” from this point forward, smooth posts were first manufactured and then laser-etched to obtain circumferential, 8 μm wide microgrooves over a 2 mm wide band at the proximal end of the post (Resonetics, Nashua, NH). The microgrooves in this study were placed on the percutaneous post in order to mimic the position of the microgrooves in dental implants. To obtain the porous topography, a porous coating was applied to a 2 mm region at the proximal end of the post (K-coating, Thortex Inc., Portland, 81 OR). The subdermal components for all the devices consisted of a Ti6Al4V core with a commercially pure titanium porous coating (K-coating, Thortex Inc., Portland, OR). All devices were passivated according to the ASTM F86 standards. Following passivation, the devices were steam sterilized prior to implantation. 3.3.2 Implant surface characterization Scanning electron microscopic imaging (Quanta 600 FEG, FEI, Hillsboro, OR; High Vacuum Mode; accelerating voltage: 15 kV) was performed to visualize the surface topographies. Surface chemical composition was characterized by Xray photoelectron spectroscopy (XPS). The XPS survey and high-resolution spectra were recorded using a Kratos Axis Ultra DLD system (Kratos Analytical, Manchester, UK) using a monochromatic Al Kα (hν = 1486.6 eV) X-ray source (8 mA emission current, 15 kV anode tension) with a power of 120 W. Scan analyses were carried out over a measurement area of 700 x 300 µm. Survey spectra and high-resolution spectra were collected using pass energies of 160 eV and 40 eV, respectively, with an energy step of 1 eV and dwell time of 300 ms. All binding energy values were charge corrected to C 1s at 284.8 eV. Relative concentrations (atomic %) of the detected elements were calculated from peak intensities in the high-resolution spectra recorded for each element recognized in the survey spectra. 82 3.3.3 Study design The study followed an approved animal protocol from the Institutional Animal Care and Use Committee at the George E. Wahlen Department of Veterans Affairs Medical Center. A total of 18, 6-week old, female Institute Armand Fappier (IAF) hairless guinea pigs (~350 g in weight, strain code 161, Charles River, Kingston, NY) were randomly assigned to three equal groups. These hairless guinea pigs were chosen because they are euthymic (immunocompetent) and are used for wound healing studies due to the lack of hair (https://www.criver.com/products-services/find-model/iaf-hairless-guineapig?region=3611). Animals were implanted with a single percutaneous device: microgrooved (N = 6), porous (N = 6) or smooth (N = 6). Post-implantation, all animals remained in the study for a duration of 4 weeks. 3.3.4 Surgical procedure A previously successful One-Stage aseptic surgical procedure was used.27 Briefly, the dorsum region of the animal was prepared for aseptic surgery. A 4 cm longitudinal incision, 0.5 cm left of the spinal column in the cranial to caudal direction, was made to expose the underlying soft-tissues. Following incision, a subcutaneous pocket was created by blunt dissection. Using a 4-mm skin biopsy punch, a percutaneous access portal was created in the skin. A fully assembled device was then inserted into the pocket and the percutaneous post was made to protrude through the access portal. The skin incision was sutured close (4-0 Vicryl suture, Ethicon, Hamburg). Prior to recovery, a subcutaneous injection of 83 carprofen (4 mg/kg, Lake Forest, IL) was administered to provide post-operative analgesic treatment. After cleaning the surgical site with sterile saline water, all animals received a gauze dressing around the post and the site was covered with semi-occlusive dressings (Tegaderm™ Film, Saint Paul, MN). Following recovery, all animals were housed in custom designed housing units which adhered to the guidelines in the “Guide for the Care and Use of Laboratory Animals”. During the study period, the animals were monitored three times daily. During animal checks, the condition of each dressing (intact or detached from the site) was recorded. If the dressing was detached from the site and the percutaneous post was visible, then the dressing was changed while the animal was under the influence of general isoflurane anesthesia. Dressings were applied to limit the infiltration of exogenous agents at the percutaneous sites. 3.3.5 Sample collection and processing Four weeks post-implantation, all animals were euthanized according to an institutionally approved protocol. The device with the surrounding soft-tissue was harvested, fixed in 10% Neutral Buffered Formalin, dehydrated with ascending grades of ethanol and then embedded in poly (methyl) methacrylate (PMMA) using an established procedure.29 Upon polymerization, using a custom high-speed water-cooled saw equipped with a diamond-coated blade (LS10 Lapidary Slap Saw, Lortone Inc., Mukilteo, WA), ~2 mm transverse sections through the post were obtained from each specimen. These sections were ground to a thickness of ~50 to 70 µm and polished to an optical finish with a semi-automatic grinding unit 84 (Ecomet™ 300, Buehler, Lake Bluff, IL). Following sectioning and polishing, the slides were stained with Hematoxylin and Eosin (H&E). Briefly, the slides were immersed in pre-heated (50 to 55oC) Mayer’s Hematoxylin (Sigma Aldrich, St. Louis, MO) for 15 min and blued-in with running lukewarm tap water for 30 min. They were then immersed in 25% acidified Eosin solution (Thermo Fisher Scientific, Waltham, MA) for 45 s and finally rinsed with 100% ethanol. 3.3.6 Histological analysis Histological slides were analyzed using a transmitted light microscope (Nikon Eclipse Ni, Nikon Instruments Inc., Melville, NY) with an attached DS-Vi1 color digital camera (MQA12010, Nikon Instruments Inc., Melville, NY). In order to quantify downgrowth and perform histological analysis, still images were captured using an imaging software package (NIS Elements, version 4.0, Nikon Instruments Inc., Melville, NY). Pictures for publication were compiled using commercially available software (Photoshop, Adobe Systems Inc., San Jose, CA). Downgrowth was measured using a previously published method.28 Briefly, three histological landmarks were noted with the first landmark being the attachment point of the percutaneous post and the subdermal device. The second landmark was denoted as the point at which the epidermis, the air and the device met, defined as the 3-point junction. The third landmark marked the edge of the subdermal device. The total length of the porous coating was measured from the first to the third landmarks and the exposed coating length was measured from the first to the second landmarks. Downgrowth was calculated as the percentage of 85 the exposed coating length to the total length of the porous coating. Downgrowth measurements were obtained from both sides of the post in each sample and reported as an average. 3.3.7 Statistical analysis To compare the average dressing longevity and epidermal downgrowth amongst the three groups, a one-way analysis of variance (ANOVA) was used. To quantify significance in epidermal downgrowth, independent, two-tailed Student tTests were performed (Microsoft Office, Excel, 2013). The level of significance was set at 5% for all statistical analyses. All measures were reported as mean ± one standard deviation. 3.4 Results Clinically, no signs of infection were observed around the percutaneous post throughout the study. During the post-implantation period, all animals in the three groups exhibited healthy tissue with no redness at the interface. On average, each dressing was maintained for 50 ± 8 h for animals in the Microgrooved Group, 48 ± 15 h for animals in the Porous Group and 45 ± 12 h for animals in the Smooth Group. Statistically, there was no significant difference (p = 0.80) in dressing longevity between the three groups. XPS was used to analyze the surface elemental composition of the titanium devices. The data revealed signals for O, Ti, N, C and Al on the surfaces of all the samples. On the microgrooved and smooth surface, Si was also detected. The 86 relative atomic concentrations of the detected elements are summarized in Table 3.1. Overall, the data indicated that the chemical composition was relatively similar for all surfaces regardless of fabrication techniques used to obtain the desired topographical features. In all three groups, gross photographs (Figure 3.2) and photomacrographs (Figure 3.3) of the skin-device interfaces indicated that the epidermis attached to the subdermal device but not to the percutaneous post. After 4 weeks postimplantation, the Microgrooved Group showed 23 ± 4% downgrowth compared to the Porous Group (20 ± 4%; p = 0.18). Compared to the Smooth Group (21 ± 9%), both the Microgrooved Group and the Porous Group did not show any significant differences in downgrowth (p = 0.63 and p = 0.79 respectively). Overall, at the end of 4 weeks, the peri-prosthetic soft-tissues appeared to undergo epidermal downgrowth regardless of the topographical features (Figure 3.3). Histological analysis of the skin-device interface in all the samples revealed that at the 3-point junction, the epidermis terminated at the subdermal surface with the epidermal cells contacting the porous coating of the subdermal device. All samples showed the presence of a hyperplastic epidermis leading up to the 3-point junction. Additionally, a hyper-cellular granulation tissue was observed underneath the hyperplastic epidermis at the 3-point junction (Figures 3.4C, 3.5C). Proteinaceous crusts consisting of neutrophil-rich debris and remnants of previous epidermal attachments were found to be present external to the 3-point junction and along the exposed surface of the device in all the groups (Figures 3.4A, 3.4B and 3.5A, 3.5B). 87 Two types of epidermal responses were observed at the 3-point junction with some samples exhibiting a double epidermis (Figure 3.4C) and some samples exhibiting a single epidermis (Figure 3.5C). A double epidermis was characterized by the presence of the main epidermal tongue and a smaller epidermal tongue which formed as an offshoot from the main epidermal tongue (Figure 3.4C). Looking at individual sides of the skin-device cross sections in the Microgrooved Group, a double epidermis was present in 75% of sides, while 25% of sides showed a single epidermis. In the Porous Group, 42% of sides showed a double epidermis and 58% of sides showed a single epidermis. Lastly, in the Smooth Group, 67% of sides showed a double epidermis and 33% of sides showed a single epidermis. All histological slides demonstrated fibrous tissue ingrowth into the pores of the coating, along the slope of the subdermal device. Tissue ingrowth was characterized by the presence of fibrovascular tissue inside the pores. A thin fibrous capsule, oriented parallel to the implant surface, was observed around the subdermal device in all the samples. 3.5 Discussion This study was designed to investigate the efficacy of a microgrooved topography to limit epidermal downgrowth around non-gingival percutaneous devices. The data obtained 4 weeks post-implantation in a guinea pig back-model showed that the tested hypothesis was not proven. Results indicated that application of the microgrooved topography to the percutaneous post alone did not 88 halt epidermal downgrowth. Additionally, when the microgrooved topography was compared to the porous and the smooth topographies, no significant difference in downgrowth was found. Overall, the data demonstrated that regardless of the topographical features tested, epidermal downgrowth occurred around percutaneous devices (Figures 3.2, 3.3). Histologically, all the three groups exhibited similar soft-tissue morphologies at the skin-device interface. Observations indicated the presence of a proteinaceous debris external to the 3-point junction, a double or single epidermis and an area of granulation tissue at the 3-point junction. The proteinaceous debris consisted of remnants of epidermis which might indicate previous epidermal attachments (Figures 3.4B, 3.5B). This observation suggests that downgrowth along the device might have followed a mechanism which included serial delamination and re-epithelialization processes. More specifically, repeated epidermal attachment to the device followed by repeated epidermal delamination (split) at the basement membrane might have led to the progression of downgrowth. Previous studies with percutaneous devices in an ovine model have similarly reported the continuous “split and re-epithelialization process”, indicating a history of previous epidermal attachments and subsequent detachment as downgrowth progresses.4,6 Additionally, the histological observation of a double or single epidermis at the 3-point junction (Figures 3.4C, 3.5C) appeared to validate the postulation of the “split and re-epithelialization process”.4,6 A double epidermis might indicate that a new epidermal layer was in the process of forming while the older attachment was in the process of delamination at the basement membrane. 89 A third histological observation was the presence of granulation tissue at the 3point junction. It is well known that granulation tissue is remodeled into scar tissue during the wound maturation phase.30,31 Thus, the presence of granulation tissue after 4 weeks of implantation (Figures 3.4C, 3.5C) may indicate that wound healing in peri-prosthetic tissues was unresolved and ongoing. During cutaneous wound healing, epidermal cells at the wound edges, supported by the underlying granulation tissue, undergo proliferation and migration in order to cover the defect.32,33 Around percutaneous devices, proliferation and migration of epidermal cells due to unresolved and ongoing wound healing may contribute to epidermal downgrowth.1,19,34 Thus, the histological characteristics observed in this study, including the presence of a double epidermis and the granulation tissue, may provide evidence that the peri-prosthetic epidermal tissue was in a state of continuous wound healing contributing to the observed downgrowth. The present study was the first to investigate the influence of the microgrooved topography (Laser-Lok® technology) to limit epidermal downgrowth around percutaneous devices implanted in normal skin. While the microgrooved topography has shown success in limiting gingival epithelial downgrowth around dental implants,24,25 it was unable to halt epidermal downgrowth around percutaneous devices implanted in normal skin (Figures 3.2, 3.3). One possible reason why the microgrooved topography failed to halt epidermal downgrowth in the present study may be due to intrinsic differences between the gingival epithelial cells and epidermal cells.26 Pendegrass et al. reported that the in vitro hemidesmosome expression in gingival cells was three times greater compared to 90 that of epidermal keratinocyte cells seeded on titanium surfaces.35 Furthermore, studies have suggested that gingival epithelial cells attach to the dental implant surfaces by means of multiprotein junctional complexes, known as hemidesmosomes, which enforce stable adhesion of gingival epithelial cells to the underlying titanium surface.35-37 Thus, future studies should investigate methodologies to increase hemidesmosome expression which may allow epidermal keratinocytes to attach more strongly to microgrooved devices, leading to limited downgrowth. Another plausible reason why the microgrooved topography did not show success in limiting downgrowth may be due to the unanchored nature of the devices.38 The microgrooved topography has been successful in limiting downgrowth around bone-anchored dental implants. Bone-anchored percutaneous devices are relatively stable, with reduced tissue movement and shear forces at the skin-device interface.2,4,39 In the present study, the microgrooved topography was tested in an unanchored percutaneous device which may have increased relative tissue movement, preventing stable attachment of the epidermis to the microgrooves.40 The continuous disruption of the epidermal-device seal may have exacerbated the amount of epidermal downgrowth. Some evidence of this is seen by the remnants of the epidermis on the microgrooved and porous posts (Figures 3.4B, 3.5B), which may be a result of continuous interfacial forces disrupting previous epidermal-device attachments. Further studies may be needed to evaluate the efficacy of the microgrooved topography in limiting downgrowth around bone-anchored percutaneous devices 91 in normal skin. The limitations to this study were two-fold. First, this study did not test the microgrooved topography on the subdermal component of the device. To mimic the design of the dental implants in which the microgrooved topography is applied to the abutment which comes in contact with the gingival epithelium, the devices in the present study were designed such that the microgrooves were applied to the percutaneous post which comes in contact with the epidermis. However, the percutaneous post might not provide a sufficient surface area to allow for tissue integration and stabilization during the initial healing phase. Future studies should explore the application of the microgrooved topography to the subdermal component to provide an increased surface area for soft-tissue attachment in order to limit downgrowth. Second, use of PMMA for embedding the soft-tissue specimens made it difficult to perform in depth histological and molecular analyses. Investigation of new embedding techniques to allow for use of molecular techniques may improve understanding of the peri-prosthetic tissue environment and elucidate the mechanisms responsible for epidermal downgrowth. In summary, our study suggests that the microgrooved topography, which has previously shown success in limiting gingival epithelial downgrowth around dental implants, might not limit downgrowth in percutaneous devices implanted in normal skin. Inherent differences between gingival tissues and epidermal tissues as well as interfacial forces at the skin-device interface due to the unanchored nature of the device might be possible reasons for the occurrence of downgrowth. Further investigations are needed to evaluate the application of the microgrooved 92 topography to the subdermal component of the device. Additionally, supplementing topographical features with chemical or mechanical therapies may help stabilize the peri-prosthetic soft-tissues in order to limit downgrowth. 3.6 Acknowledgements This work was supported in part by the US Army Medical Research and Materiel Command under contract #W81XWH-15-C-0058, by the Department of Defense under grant #W81XWH-11-1-0435, by the United States Department of Veterans Affairs Rehabilitation Research and Development Service under Merit Review Awards #I01RX001217 and #I01RX001246, by the Department of Orthopaedics, University of Utah School of Medicine, Salt Lake City, Utah and by the LS-Peery Program in Musculoskeletal Restoration. The views, opinions and/or findings presented are those of the authors and should not be construed as an official position, policy or decision of any of these funding sources unless so designated by other documentation. This publication made use of the University of Utah Shared facilities of the Surface Analysis and Nanoscale Imaging group sponsored by the College of Engineering, Health Sciences Center, Office of the Vice President for Research, and the Utah Science Technology and Research (USTAR) Initiative of the State of Utah. The authors would like to express their gratitude to Brian Roy Van Devener for his help with XPS analysis. The authors would like to thank Richard Tyler Epperson and Brooke Kawaguchi (Bone and Joint Research Laboratory, Department of Veterans Affairs, 93 Salt Lake City, UT) for their help in preparing samples for histological analysis. The authors would also like to acknowledge Kelli Hafer for her assistance during the animal study. 3.7 References 1.von Recum AF. Applications and failure modes of percutaneous devices: a review. J Biomed Mater Res 1984;18(4):323-36. 2.Pendegrass CJ, Goodship AE, Blunn GW. 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Int J Periodontics Restorative Dent 2010;30(3):245-55. 26.Turabelidze A, Guo S, Chung AY, Chen L, Dai Y, Marucha PT, DiPietro LA. Intrinsic differences between oral and skin keratinocytes. PLoS One 2014;9(9):e101480. 27.Mitchell SJ, Jeyapalina S, Nichols FR, Agarwal J, Bachus KN. Negative pressure wound therapy limits downgrowth in percutaneous devices. Wound Repair Regen 2016;24(1):35-44. 28.Pawar DRL, Mitchell SJ, Jeyapalina S, Hawkes JE, Florell SR, Bachus KN. Peri-prosthetic tissue reaction to discontinuation of negative pressure wound therapy around porous titanium percutaneous devices. J Biomed Mater Res B Appl Biomater 2018. 29.Emmanual J, Hornbeck C, Bloebaum RD. A polymethyl methacrylate method for large specimens of mineralized bone with implants. Stain Technol 1987;62(6):401-10. 30.Barrientos S, Stojadinovic O, Golinko MS, Brem H, Tomic-Canic M. Growth factors and cytokines in wound healing. Wound Repair Regen 2008;16(5):585-601. 31.Enoch S, Leaper DJ. Basic science of wound healing. Surgery (Oxford) 2008;26(2):31-37. 32.Landen NX, Li D, Stahle M. Transition from inflammation to proliferation: a critical step during wound healing. Cell Mol Life Sci 2016;73(20):3861-85. 33.Werner S, Krieg T, Smola H. Keratinocyte-fibroblast interactions in wound healing. J Invest Dermatol 2007;127(5):998-1008. 34.Holt BM, Betz DH, Ford TA, Beck JP, Bloebaum RD, Jeyapalina S. Pig dorsum model for examining impaired wound healing at the skin-implant interface of percutaneous devices. J Mater Sci Mater Med 2013;24(9):2181-93. 35.Pendegrass CJ, Lancashire HT, Fontaine C, Chan G, Hosseini P, Blunn GW. Intraosseous transcutaneous amputation prostheses versus dental implants: 96 a comparison between keratinocyte and gingival epithelial cell adhesion in vitro. Eur Cell Mater 2015;29:237-49. 36.Rezaei Esfahrood Z, Kadkhodazadeh M, Gholamin P, Amid R, Passanezi E, Hosein Zadeh H. Biologic width around dental implants: an updated review. J Dent Mater Tech 2016;5(2):68-81. 37.Schupbach P, Glauser R. The defense architecture of the human periimplant mucosa: a histological study. J Prosthet Dent 2007;97(6 Suppl):S1525. 38.Nevins M, Nevins M, Gobbato L, Lee HJ, Wang CW, Kim DM. Maintaining interimplant crestal bone height via a combined platform-switched, Laser-Lok implant/abutment system: a proof-of-principle canine study. Int J Periodontics Restorative Dent 2013;33(3):261-7. 39.Grosse‐Siestrup C, Affeld K. Design criteria for percutaneous devices. J Biomed Mater Res 1984;18(4):357-382. 40.Jansen JA, van der Waerden JP, van der Lubbe HB, de Groot K. Tissue response to percutaneous implants in rabbits. J Biomed Mater Res 1990;24(3):295-307. 97 Figure 3.1. A set of photographs showing the three topographies used for this study: A) microgrooved, B) porous and C) smooth (machine finish). The topographies were only applied to the proximal region of the percutaneous post while the porous coated subdermal device was the same in all the groups. Corresponding scanning electron microscope images (D, E and F) show a magnified view of the three topographies (magnification: 100x; accelerating voltage: 15 kV). 98 Figure 3.2. A representative set of photographic images of the skin-device exit sites at 4 weeks post-implantation: A) Microgrooved, B) Porous and C) Smooth Groups. Tissues in all three groups had undergone downgrowth with the epidermis attaching to subdermal device. The black arrow indicates exposed subdermal device in all the three groups. 99 Figure 3.3. A representative set of H&E stained photomacrographs: A) Microgrooved, B) Porous and C) Smooth Groups. White arrows show the position of tissue attachment with respect to the base of the percutaneous post (white dotted line). No significant difference in epidermal downgrowth was found between the three groups. White scale bar – 1000 µm. 100 Figure 3.4. A representative H&E stained photomicrograph showing the skindevice interface of the microgrooved device (A). Photomicrographs (B) and (C) show the magnified regions within the yellow solid and dashed box in (A), respectively. The photomicrograph (B) shows proteinaceous debris, which suggests a history of previous epidermal attachments (denoted by white arrows) on the microgrooved percutaneous post (P). The photomicrograph (C) shows double epidermis (DE) and granulation tissue (GT) at the skin-device interface which may indicate unresolved and ongoing wound healing. Image A was taken at 4x magnification, scale bar- 500 µm. Images B and C taken at 10x magnification, scale bar- 200 µm. 101 Figure 3.5. A representative H&E stained photomicrograph showing the skindevice interface of the porous device (A). Photomicrographs (B) and (C) show magnified regions within the yellow solid and dashed box in (A). Image (B) shows proteinaceous debris which suggests a history of previous epidermal attachments (denoted by white arrows) on the porous percutaneous post (P). Image (C) shows single epidermis (SE) and underlying granulation tissue (GT) at the skin-device interface, which may indicate unresolved and ongoing wound healing. Image A was taken at 2x magnification, scale bar- 1000 µm. Image B was taken at 4x magnification, scale bar- 500 µm and Image C at 10x magnification, scale bar- 200 µm. 102 Table 3.1. Elemental composition (atomic %) of device surfaces after cleaning and sterilization, determined by XPS analysis. Component O Ti N C Si Al Microgrooved Post 36.93 10.88 1.23 48.58 0.95 1.41 Porous Post 38.17 14.61 2.21 43.03 - 1.98 Smooth Post 40.49 16.30 1.64 36.14 1.21 4.22 Subdermal Device 36.75 14.11 1.15 46.55 - 1.43 CHAPTER 4 INFLUENCE OF NEGATIVE PRESSURE WOUND THERAPY ON PERIPROSTHETIC TISSUE VASCULARIZATION AND INFLAMMATION AROUND POROUS TITANIUM PERCUTANEOUS DEVICES 2 4.1 Abstract Negative pressure wound therapy (NPWT) has been shown to limit downgrowth around percutaneous devices in a guinea pig model. However, the influence of NPWT on peri-prosthetic tissue characteristics leading to limited downgrowth is still unclear. In order to investigate this, 12 CD hairless rats were assigned into 2 groups, NPWT and Untreated (n=6/group). Each animal was implanted with a porous coated titanium percutaneous device and was dressed with a gauze and semi-occlusive base dressing. Post-surgery, animals in the NPWT Group received a regimen of NPWT treatment (-70 to -90 mmHg). After 4 weeks, tissue was collected over the device and stained with CD31 and CD68 to quantify blood vessel density and inflammation, respectively. The device with the 2 Reprinted with permission from John Wiley and Sons, Inc.: Pawar DRL, Jeyapalina S, Hafer K, Bachus KN. Influence of negative pressure wound therapy on peri-prosthetic tissue vascularization and inflammation around porous titanium percutaneous devices. J Biomed Mater Res B 2019. doi: 10.1002/jbm.b.34302. Copyright © 2019 Wiley Periodicals, Inc. All rights reserved. 104 surrounding tissue was also collected to quantify downgrowth. NPWT treatment led to a 1.6-fold increase in blood vessel densities compared to untreated tissues (p < 0.05). NPWT treatment also resulted in half the downgrowth as the Untreated Group, although not statistically significant (p = 0.19). Additionally, the results showed a trend towards increased CD68 cell densities in the NPWT Group compared to the Untreated Group (p = 0.09). These findings suggest that NPWT may influence wound healing responses in percutaneous devices by increasing blood vessel inflammation. densities, Overall, limiting NPWT downgrowth may enhance and potentially tissue increasing vascularity around percutaneous devices, especially in patients with impaired wound healing. 4.2 Introduction Downgrowth, commonly observed around many percutaneous devices, is marked by the migration of the epidermal layer alongside the implant in order to re-establish continuity in the epidermis.1-3 As a result, the underlying device becomes exposed which increases the risk of infection. Although the exact mechanism of downgrowth is not entirely clear, factors associated with wound healing, such as vascularization and inflammation in the peri-prosthetic tissues, might be involved.4,5 Moreover, the importance of vascular supply to the periprosthetic tissues may be ascertained from percutaneous dental implant literature. Studies have suggested that a reduced blood supply can be one of the main reasons for a lack of healing in gingival soft-tissues surrounding dental implants, which may ultimately lead to tissue downgrowth and device failure.6,7 Since the 105 epidermal tissue is avascular, the underlying vasculature within the connective tissue is an essential source of oxygen and nutrients necessary for wound healing.8 Thus, improving the vascularity of the peri-prosthetic tissues may play an important role for the healing outcomes of the epidermis and dermis, and consequently reduce downgrowth. With regards to inflammation, studies have indicated that while all implanted materials induce some degree of foreign body reaction and subsequent inflammatory response, persistent and unmitigated inflammation can cause tissue destruction, which may negatively affect wound healing outcomes.9,10 In addition to wound healing cues, micro-movement (i.e., shear forces) at the skin-implant interface may also lead to disruption of the softtissue, which further exacerbates downgrowth rates.1 Thus, strategies to improve wound healing outcomes by inducing vascularization, mitigating inflammation and reducing mechanical forces within the peri-prosthetic tissues may benefit the longevity and functionality of percutaneous devices. One such strategy is to use negative pressure wound therapy (NPWT). In a translational animal model, NPWT has been shown to improve downgrowth outcomes around percutaneous devices.11 This result was not surprising since NPWT is used to facilitate the healing of chronic and “hard to heal” wounds such as diabetic ulcers.12 Typically, these wounds are large defects. Wound site preparation for application of NPWT involves the use of an interface material (commonly a polyurethane foam) and a semi-occlusive dressing. The polyurethane foam, which is sized to fit the wound, is believed to bring the wound margins together during the application of NPWT.12 Alternatively, studies have 106 indicated that a gauze based interface material can transmit the negative pressure as efficiently as the foam.13,14 It has been suggested that, gauze based dressings are easier to use for atypical wounds receiving NPWT and result in relatively less pain and discomfort to the patients.14 Moreover, a range of negative pressures have been used to treat wounds.14 A pressure magnitude of -125 mmHg has been shown to improve clinical wound healing outcomes,15,16 and is commonly used in clinical practice.17,18 However, many studies have challenged this value and shown that a pressure of -80 mmHg may be optimal for wound contraction.14,19,20 Clinically, NPWT is commonly used to improve wound healing outcomes by mechanically drawing the wound edges together, increasing angiogenesis and modulating inflammation.21 Studies have shown that wounds treated with NPWT show increased blood vessel densities and are associated with improved wound closure compared to untreated wounds.17,22 These studies concluded that NPWT provides a favorable wound healing environment by increasing vascularization of the wound bed. While the efficacy of NPWT in inducing vascularity is well established, there is ambiguity about the effects of NPWT on inflammation. Many studies suggest that NPWT application results in a decrease of the inflammatory response,18,23,24 while some suggest that NPWT actually increases inflammation in the wound.25,26 Although, there is a disagreement about the mechanism of action of NPWT on inflammation, studies show that NPWT leads to good wound healing outcomes compared to untreated wounds.22,27 Despite clinical success in open wounds, there is a relative lack of data regarding the use of NPWT with percutaneous devices. One recent study 107 suggested that NPWT has the ability to reduce epidermal downgrowth over percutaneous devices in a guinea pig model.11 Mitchell et al. showed that a negative pressure treatment of -80 to -100 mmHg applied through gauze and semiocclusive dressings over percutaneous devices was able to limit downgrowth.11 One shortcoming of the study was that apart from performing downgrowth measurements, the effects of NPWT on wound healing characteristics such as blood vessels and inflammation within peri-prosthetic tissues were neither studied, nor characterized. Overall, there is a need to better understand the utilization of NPWT with percutaneous devices and to gain insight into the possible mechanisms by which NPWT facilitates the healing outcomes of peri-prosthetic tissues. Therefore, the goal of this study was to investigate the effects of NPWT on the vascularization and inflammation of peri-prosthetic soft-tissues and to reproducibly limit epidermal downgrowth around percutaneous devices. Unlike the previous study which used a guinea pig model to investigate the effects of NPWT on peri-prosthetic tissues,11 this study utilized a rat model due to the availability of molecular-level analysis tools. This study tested two hypotheses. First, based on the established effects of NPWT in open wounds, this study hypothesized that NPWT would increase vascularization within the peri-prosthetic tissues compared to untreated tissues. Second, NPWT treated peri-prosthetic tissues would exhibit limited epidermal downgrowth around the percutaneous devices compared to untreated tissues. In addition, this study also sought to characterize the effects of NPWT on inflammation within the peri-prosthetic tissues. 108 4.3 Materials and Methods 4.3.1 Study design The study followed an approved animal protocol from the Institutional Animal Care and Use Committee. A total of 12, 8-week old, female CD hairless rats (~ 200 g in weight, strain code 184, Charles River, Wilmington, MA) were randomly assigned to two equal groups. These hairless rats were chosen because they are euthymic (immunocompetent) and are commonly used for wound healing studies due to the lack of hair (https://www.criver.com/products-services/findmodel/cd-hairless-rat?region=3616). Post-surgery, six animals were subjected to NPWT treatment through a gauze and semi-occlusive dressing (NPWT Group). The remaining six animals received only the gauze and semi-occlusive dressing in order to minimize the environmental contamination to the site and served as controls (Untreated Group). 4.3.2 Device design Each animal was implanted with a historically successful percutaneous device.11 Briefly, each device consisted of a machined smooth titanium alloy (Ti6Al4V) percutaneous post, and a subdermal device consisting of a Ti6Al4V core with a commercially pure titanium porous coating (K-coating, Thortex Inc., Portland, OR). The devices were passivated according to the ASTM F86 standards. Following passivation, the devices were steam sterilized prior to implantation. 109 4.3.3 Surgical procedure Each animal was implanted with one percutaneous device using a previously successful One-Stage aseptic surgical procedure.11 Briefly, after preparing the dorsum region for aseptic surgery, a 4 cm longitudinal incision (0.5 cm left of the spinal column and in the cranial to caudal direction) was made to expose the underlying soft-tissues. A subcutaneous pocket was created by blunt dissection on the left side of the incision line, followed by the creation of a percutaneous access portal using a 4-mm skin biopsy punch. A fully assembled device was then inserted into the pocket and the percutaneous post was made to protrude through the skin. The skin incision was sutured closed (4-0 Vicryl, Ethicon, Hamburg, Germany). A subcutaneous injection of carprofen (4 mg/kg, Lake Forest, IL) was administered to provide post-operative analgesic treatment. As described in previous studies,11,28,29 the implant exit site was covered with a sterile gauze and a semi-occlusive base dressing in all the animals. Briefly, post-surgery the skin was cleaned with sterile saline and gauze. A liquid adhesive (Mastisol®, Fendale Laboratories, Inc., Ferndale, MI) was applied to the skin circumferentially, ~1 cm from the percutaneous device, creating an outline for the placement of the semi-occlusive dressing. Sterile gauze was cut into a circular shape and a small hole was created in the center of the gauze in order to allow the percutaneous post to protrude through. Further, a layer of gauze was placed on top of the percutaneous post to cover the entire device. Finally, semi-occlusive dressings (Tegaderm™ Film, Saint Paul, MN) were applied over the gauze and the adhesive to achieve an airtight seal. Application of the adhesive allowed for the 110 Tegaderm to stay leak proof for longer. For animals in the NPWT Group, a NPWT delivery line (consisting of a tube, a neoprene rubber washer, and a stainless steel retaining ring) was inserted into the base dressing (Figure 4.1A). Four smaller semi-occlusive dressings were used to secure the NPWT delivery tube to the base dressing and to achieve an airtight seal. This tube allowed for the application of the negative pressure treatment directly to the gauze over the implantation site. To limit physical stress resulting from the NPWT delivery set-up, custom designed cages based upon previously reported studies, were used (Figure 4.1B).11,28,29 Briefly, the cages were designed with a rotary L-fitting (Model RHL ¼-N2U, PISCO Inc., Bensenville, IL) and a straight connector (Model PCF ¼-N2U, PISCO Inc., Bensenville, IL), which allowed the animal unrestricted access to the entire cage. Additionally, in order to limit tensile or compressive forces to the NPWT application site, the length of the NPWT delivery tube (Model SVEB20 clear vinyl tubing, Watts, Andover, MA) was cut longer than the distance between the lid and the base of the cage to allow for slack in the delivery tube. 4.3.4 Post-operative management Following recovery, the animals were returned to their cages and the periprosthetic tissues were treated with NPWT set to continuously apply -70 to -90 mmHg using a vacuum pump (Model 7314P-D-EXF, DeVilbiss Healthcare, Port Washington, NY). Previous studies have shown that the application of negative pressure within this range was tolerated well by the animals.11,29 Pressures were monitored using an external pressure sensor (Model QPSL-AP-42, ProSense®, 111 Cumming, GA) that was connected proximal to the NPWT delivery pump using a t-connector (Model 47334, Dorman Auto Grade Connectors, www.oreillyauto.com). Based upon previous studies, it was noted that animals maintained their dressings for ~48 h.11,29 Thus, dressings were changed either at 48 h intervals or earlier based upon the condition of the dressing. During the duration of the study, animal checks were performed three times daily to inspect the dressing and monitor for leaks in the negative pressure delivery system. For the animals in the Untreated Group, the condition of each dressing (intact or detached from the site) was visually inspected and noted. If the dressing was detached from the percutaneous device, then the dressing was changed. For the animals in the NPWT Group, pressure sensor readings were noted in addition to visual observations. If the pressure sensor reading was less than -70 mmHg, the dressing was considered breached and was changed. 4.3.5 Sample collection and processing Four weeks after implantation, all animals were euthanized following an institutionally approved protocol. Following euthanasia, a strip of soft-tissue (~2 mm thick, 6 mm in length) was collected from over the subdermal device in the cranial-caudal direction from each animal. One skin section (5 x 5 mm) was collected from the opposite side of the spine from each animal to serve as an internal control and will be referred to as “distal tissue” (Figure 4.2). The spleen and liver were also collected to serve as positive controls for immunohistochemical (IHC) staining. All tissue specimens were fixed in 10% neutral buffered formalin 112 (NBF) for 24 to 48 h, embedded in paraffin, sectioned at 3 µm thickness and mounted on slides for evaluation. Additionally, the device with the surrounding soft-tissue was harvested, fixed in 10% NBF, dehydrated with ascending grades of ethanol, and then embedded in methyl methacrylate (Technovit® 9100, Heraeus Kulzer, Wehrheim, Germany) according to the manufacturer’s protocols (https://www.emsdiasum.com/microscopy/technical/datasheet/14655.aspx). After complete polymerization, ~2 mm transverse sections through the post, in the medial-lateral direction, were obtained from each specimen using a precision saw (IsoMet™ 4000 linear precision saw, Buehler, Lake Bluff, IL). These sections were ground to ~50 to 70 µm sections and polished to an optical finish (Ecomet™ 300, Buehler, Lake Bluff, IL). The slides were stained with Hematoxylin and Eosin. Briefly, the slides were immersed in pre-heated (50 to 55oC) Mayer’s Hematoxylin (Sigma Aldrich, St. Louis, MO) for 10 min and blued-in with running tap water for 15 min. They were then immersed in 1% Eosin for 10 s. 4.3.6 Immunohistochemistry All IHC staining was performed in the Ventana Discovery automated staining system (BenchMark™ Discovery, Ventana Medical Systems, Tucson, AZ) using Ventana reagents for the entire procedure. Paraffin-embedded sections were placed in the automated staining system, deparaffinized and pretreated with cell conditioning buffer for 60 min at 95oC for antigen retrieval. Slides were then incubated for 1 h at 37°C with a rabbit anti-rat CD31 primary antibody (1:50 dilution, 113 ab28364, Abcam, Cambridge, MA) for labelling blood vessels and a rabbit anti-rat CD68 primary antibody (1:200 dilution, ab125212, Abcam, Cambridge, MA) for labelling monocyte-macrophage lineage cell densities. Then, secondary antibody (goat anti-rabbit immunoglobulin, Sigma Aldrich, St. Louis, MO) was applied at dilution 1/100 for 32 min at 37°C. Detection of slides was carried out using the IView DAB detection kit, which is a Streptavidin-HRP system, utilizing DAB (3-3’ diaminobenzidine) as the chromogen. The slides were counterstained with hematoxylin (Ventana Medical Systems, Tucson, AZ) followed by bluing. The spleen and liver were used for both positive and negative controls. For positive controls, CD31 and CD68 staining was performed on the spleen and liver sections in the same manner as the tissue samples described above. For negative controls, the spleen and liver were incubated with just the secondary antibody without the primary. Absence of any stain indicated that the secondary antibody did not bind non-specifically. 4.3.7 Histological analysis All sections were histologically analyzed using a transmitted light microscope (Nikon Eclipse Ni, Nikon Instruments Inc., Melville, NY) with an attached DS-Vi1 color digital camera (MQA12010, Nikon Instruments Inc., Melville, NY). Still images were captured using an imaging software package (NIS Elements, version 4.0, Nikon Instruments Inc., Melville, NY), and publication quality pictures were compiled using commercially available software (Photoshop, Adobe Systems Inc., San Jose, CA). 114 4.3.7.1 Downgrowth measurements The sections obtained from the methacrylate embedded samples were used for downgrowth measurements. Downgrowth was measured following an established method.11 Briefly, downgrowth was calculated as the percentage of the exposed coating length to the total available porous coating length for both sides of the device. The exposed porous coating length was measured from the base of the post [dashed line; Figure 4.3(C,D)] to the point where epidermis first touches the subdermal device [white arrows; Figure 4.3(C,D)]. It must be noted that in some sections, the epidermis at the interface was breached. In these sections, the attachment point of the underlying tissue with the device was noted. Downgrowth measurements were obtained from both sides of the post in each sample and expressed as mean ± one standard deviation for each group. 4.3.7.2 Blood vessel and macrophage density Slides, which were stained with CD31 and CD68, were used for quantification of blood vessel and macrophage densities, respectively. Quantification was performed, blinded to the treatment group, in two regions within each sample, at the wound edge and at a region distal to the wound edge denoted as “tissue over device” (Figure 4.2 Inset). Each sample was oriented lengthwise along the horizontal axis of the field of view. For each sample, the wound edge was defined as the region enclosed by a semi-circle of 500 µm radius placed at the termination point of the epidermis. Using the Nikon imaging software, a circular grid of 500 µm radius was centered on the end of the epidermis and the area of 115 the tissue within the circular grid was traced and measured. Within this region, the number of blood vessels and CD68 positive cells were counted. Vessel and cell densities were calculated as the number of structures counted per unit area. A structure was counted as a vessel if: (1) there was a positive CD31 stain and (2) there was an identifiable lumen. Similarly, a structure was counted as a CD68 cell if there was a positive brown stain. For density counts in the tissue over device, a square grid (627 x 627 µm) was overlaid on the tissue. Three squares were used for measurement. A random number generator was used to determine the column for each square with respect to the grid and each square was positioned to encompass the region below the epidermis for quantification of blood vessel densities. The densities measured from the three squares were then averaged and used as a count for the tissue over device. Blood vessel density counts in the distal tissue were measured and calculated in the same manner as the tissue over device. Densities were expressed as the mean number of structures/mm2 ± one standard deviation for each group. 4.3.7.3 Qualitative analysis of peri-prosthetic tissue morphology Tissues were analyzed qualitatively to determine epidermal responses at the three-point junction, the point at which the environment, tissue and implant interact. If the epidermal response contained the formation of a direct epidermal tissue-implant contact, it was characterized as a “sealed interface”. If the epidermis failed to contact the implant surface and terminated on the dermal bed, it was referred to as a “breached interface”. 116 4.3.8 Statistical analysis Independent, two-tailed Student t tests were performed to compare the NPWT and Untreated Groups and quantify significance in downgrowth measurements, blood vessel densities and CD68 positive cell densities. To compare the wound edge and the tissue over device densities with distal tissue densities within each animal, Paired t tests were performed. Differences were considered statistically significant at p < 0.05 for all analyses. All measures were reported as mean ± one standard deviation. 4.4 Results 4.4.1 Clinical observations Clinically, no signs of infection were observed around the percutaneous post throughout the study. Daily checks confirmed that none of the animals (both groups) showed any signs of physical stress, complications or abnormal behavior changes. Animals in the Untreated Group exhibited healthy tissue with no redness. On the other hand, all animals receiving the NPWT treatment exhibited skin redness at sites where the Tegaderm dressing was in direct contact with the underlying skin. Gross photographs indicated that there was a limited exposure of the underlying subdermal porous coating at the end of 4 weeks for both the Untreated and NPWT Groups [Figure 4.3(A,B)]. 117 4.4.2 Dressing longevity On average, each dressing was maintained for 36 ± 13 h for animals in the Untreated Group and for 33 ± 8 h for animals in the NPWT Group over the 4-week period. Statistically, there was no significant difference in dressing longevity between the two groups (p = 0.16). 4.4.3 Quantitative assessment of downgrowth Two sides from the Untreated Group and two sides from the NPWT Group contained processing artifacts and were not used for quantitative downgrowth analyses. Thus, 10 sides from the Untreated Group and 10 sides from the NPWT Group were used for downgrowth measurements. At 4 weeks post-implantation, the Untreated Group showed 5.3 ± 5.1% downgrowth and the NPWT Group showed 2.6 ± 3.5% downgrowth. No statistical difference in downgrowth was measurable between the Untreated and NPWT Groups (p = 0.19) [Figure 4.3(C,D)]. 4.4.4 Quantitative assessment of blood vessel density One untreated tissue sample was lost during histological processing thus, five tissue sections from Untreated Group and six tissue sections from NPWT Group were analyzed for blood vessel densities. The wound edge of the NPWT Group showed significantly increased (1.6x) blood vessel density compared to the wound edge of the Untreated Group (p < 0.05) (Figures 4.4, 4.5). The wound edges in both the NPWT and Untreated Groups showed increased blood vessel densities 118 (6.1x and 3.4x respectively) when compared to the distal tissue from each animal (p < 0.001 and p < 0.05 respectively). The distal tissues in the NPWT and Untreated Groups did not show any significant difference in blood vessel densities (p = 0.40). Within the NPWT treated tissue, the wound edge showed significantly increased blood vessel density (2.9x) compared to the tissue over device (p < 0.001). Likewise, within the Untreated Group, the wound edge showed relatively high blood vessel density (3.5x) compared to the tissue over device (p < 0.05). The tissue over device in the NPWT Group showed relatively high blood vessel density (1.9x) compared to the tissue over device in the Untreated Group (p < 0.05). Overall, tissues treated with NPWT had increased blood vessel densities compared to the untreated tissues. In addition, the wound edges in both groups had increased blood vessel densities compared to the tissue over device and distal tissues. 4.4.5 Quantitative assessment of CD68 positive cell density The wound edge in the Untreated Group showed a mean of 702 ± 598 cells/mm2 CD68 positive cell counts and the NPWT Group showed a mean of 1250 ± 372 cells/mm2 CD68 positive cell counts. Tissues at the wound edge in the Untreated Group showed variable responses with 40% of the samples showing minimal CD68 positive cell densities (Figure 4.6A) and 60% of the samples showing a relatively higher degree of inflammatory response (Figure 4.6B). However, no significant difference in CD68 positive cell densities was found to be 119 present between the wound edges of the Untreated and NPWT Groups (p = 0.09) (Figure 4.7). The wound edges in both the groups showed significantly increased CD68 positive cell densities than the tissue over device (p < 0.05 for both). The tissue over device in the NPWT Group showed increased CD68 positive cell density compared to the tissue over device in the Untreated Group (p < 0.05). 4.4.6 Qualitative assessment of peri-prosthetic tissue morphology In the samples, both breached and sealed epidermal responses to the presence of device were observed. The Untreated Group exhibited variable responses in which 40% of the sides showed a sealed interface with the epidermis terminating at the implant interface (Figure 4.8A) whereas 60% of the sides reflected a breached interface. In the NPWT Group, 100% of the skin-implant interface sides exhibited a breached interface (Figure 4.8B). The region between the epidermis and the implant, adjacent to the three-point junction was filled with proteinaceous debris in all of the untreated samples. Regardless of the termination point of the epidermis with respect to the device, all the Untreated and NPWT samples showed a thinning epidermal tongue closer to the three-point junction. The thin epidermis became hyperplastic in the direction away from the three-point junction (Figure 4.8). 4.5 Discussion This study tested two hypotheses. First, peri-prosthetic tissues treated with NPWT would exhibit increased blood vessel densities compared to the Untreated 120 Group. The data supported this hypothesis. Second, peri-prosthetic tissues treated with NPWT would exhibit limited downgrowth compared to untreated tissues in a rat model. Although the data did not statistically support this hypothesis, the NPWT treated tissue showed 50% of the downgrowth (2.6 ± 3.5%) compared to the untreated tissue (5.3 ± 5.1%). Additionally, this study also sought to characterize the effect of NPWT on inflammation in peri-prosthetic soft-tissues. Monocyte and macrophage marker, CD68, was used as a quantification tool for inflammation. Although there was no significant difference in macrophages/monocyte densities between the NPWT and Untreated Groups at the wound edge, NPWT treatment appeared to recruit more CD68 positive cells to the sites. In the tissue over device, significantly increased inflammation (CD68 positive cell density) was found in the NPWT Group compared to the Untreated Group. Our results are in alignment with the established literature in dermal clinical wounds, which show that application of NPWT leads to an increase in blood vessel densities compared to untreated wounds.17,22 Based on the new quantitative data, which have not been published previously, the NPWT treated dermal periprosthetic tissues reflected increased blood vessel densities compared to untreated tissues (Figures 4.4, 4.5). While the mechanism of action for this outcome is unclear, some answer might be ascertained from the clinical wound literature which suggests that NPWT application increases blood vessel densities by inducing an increase in pro-angiogenic factors and endothelial cell proliferation markers.27,30 In the present study, in order to determine the base-level blood vessel density, unwounded skin tissues from the opposite side of the spine were analyzed 121 from each animal. These distal tissues from the two groups showed similar amounts of baseline blood vessel densities, indicating that changes in vascularity in the peri-prosthetic tissues may have been due to the NPWT treatment or lack thereof rather than the inherent differences in animals. In the case of percutaneous devices, it is crucial to facilitate vascularization in peri-prosthetic tissues in order to stimulate wound healing and maintain the health of soft-tissues.6 Since, establishing a blood supply is necessary for bringing vital nutrients and oxygen to support tissue repair site,6 an increase in blood vessel densities in peri-prosthetic tissues due to NPWT, may improve implant longevity.31 Although our results showed a significant increase in blood vessel densities in the NPWT Group compared to the Untreated Group, no significant difference in downgrowth was found between the groups (Figure 4.3). The results indicated that, in both groups, variations in downgrowth between the animals were high. However, NPWT resulted in improved downgrowth outcomes (2.6 ± 3.5%) with values less than half of the Untreated Group (5.3 ± 5.1%). One of the reasons for reduced epidermal downgrowth in the NPWT Group compared to the Untreated Group might have been due to the increase in blood vessel densities in the NPWT treated peri-prosthetic tissues. We observed that the NPWT Group reflected 1.6 times more blood vessel density and 0.5 times the downgrowth as the Untreated Group. Although percutaneous dental implant literature suggests that a lack of sufficient blood supply can be one of the key reasons for impaired soft-tissue healing,6 there is a lack of studies which investigate the association between vascular supply and downgrowth in dermal applications. In this study, we were unable to demonstrate 122 a correlation between epidermal downgrowth and blood vessel density. Although only a single time point was investigated in this study, preliminary data presented here appeared to indicate that the application of NPWT did influence the blood vessel density within the peri-prosthetic soft-tissue regions. Further time-course studies are needed to understand the relationship between vascularization and downgrowth around percutaneous devices over a longer wound healing period. In the present study, the results showed a trend towards increased CD68 positive cell density at the wound edge in the NPWT Group compared to the Untreated Group (Figures 4.6, 4.7). Additionally, there was significantly increased CD68 positive cell density in the tissue over device in the NPWT Group compared to the Untreated Group. Increased CD68 positive cell densities might have been one of the reasons for the relatively reduced downgrowth in the NPWT Group. At the 4-week time point, the NPWT treated peri-prosthetic tissues might have been arrested in the inflammatory phase of the wound healing cascade. The wound healing cascade involves an interplay between the overlapping phases: hemostasis, inflammation, re-epithelialization/proliferation and remodeling. Reepithelialization involves the proliferation and migration of keratinocytes over the wound bed in order to re-establish continuity.32 In the literature, a full thickness rat wound is reported to complete re-epithelialization and remodeling within 10 days.33 However, due to the presence of the percutaneous post, wound healing around percutaneous devices is complex and thus the inflammatory and reepithelialization phases may be prolonged. Based on the current knowledge, epidermal downgrowth around percutaneous devices is attributed to the re- 123 epithelialization phase of wound healing.3,34 Thus, unresolved inflammation, as seen by the increased CD68 positive cell densities, may have arrested the progression of the wound healing cascade within the NPWT treated Group, which could have subsequently delayed re-epithelialization.25,35 The 4-week end point used in this study may have also been an insufficient follow-up period to allow for resolution of inflammation around the percutaneous devices in the rat model. Further studies evaluating longer time points may be necessary to elucidate the relationship between NPWT, inflammation and downgrowth. In addition to showing relatively increased CD68 positive cell densities and limited downgrowth, the NPWT Group also exhibited breached interfaces: i.e., lack of epidermal-to-implant attachment (Figure 4.8B). The presence of breached interfaces may have allowed for the penetration of exogenous pathogens and contaminants into the peri-prosthetic tissues,36,37 resulting in the accumulation of CD68 positive cells to aid in the clearance of local debris and contamination. Alternatively, the application of NPWT itself, may have resulted in an increase in CD68 positive cells. At the 4-week time point, the presence of inflammation could have delayed the re-epithelialization phase and thus arrested epidermal migration towards the implant. However, at a longer time point, resolution of inflammation could be possible and allow for the progression of re-epithelialization. Once reepithelialization occurs, the epithelium may contact the implant leading to subsequent downgrowth.34 At longer time points, downgrowth would be detrimental since it could lead to the creation of a sinus tract, which increases the risk of implant infection.1,3 Therefore, based on this data alone, one cannot 124 determine the merits of downgrowth compared to breached interfaces and a longer time-series study is needed. While the results of this study showed a trend towards increased inflammation in the NPWT treated peri-prosthetic tissues compared to the untreated tissues, the literature is ambiguous regarding the influence of NPWT on inflammation in clinical wounds. Studies report that the application of NPWT can either increase,25,26,35 or decrease18,38 the expression of pro-inflammatory chemokines. When using CD68 as a marker for inflammation, one study found that the application of NPWT reduced the expression of CD68 in muscle flaps,18 while another study detected an increasing trend in CD68 positive cells in NPWT treated venous leg ulcers.39 Although CD68 staining can be used as one of the ways to assess inflammation.18,40 CD68 is a general marker for cells from the monocytemacrophage lineage and cannot reveal whether the wound environment is proinflammatory or anti-inflammatory. In fact, macrophages can exist along a spectrum between M1 (pro-inflammatory) which are associated with antigen presentation and killing of intracellular pathogens and M2 (anti-inflammatory) which promote tissue repair and constructive tissue remodeling.41,42 While our results indicate that NPWT may increase inflammation as evidenced by the presence of a relatively large number of CD68 positive cells, CD68 detects all types of macrophages without differentiating the various macrophage phenotypes. Thus, the percentage of M1 or M2 macrophage phenotypes needs to be further investigated by immunohistochemical staining for specific cell surface markers for each phenotype. Future studies are necessary to elucidate the role of NPWT in 125 modulating macrophage phenotypes towards either the pro-, or the antiinflammatory state. Our study had limitations. One limitation of this study was the small sample size. The sample size used in the present study (n = 6) was based on the epithelial downgrowth data around percutaneous devices from Mitchell et al.11 and blood vessel density data from a NPWT study in open wounds,43 since blood vessel densities around percutaneous devices have not been previously quantified. Lastly, to the authors’ knowledge, there are no previously published NPWT based animal studies which quantify CD68 positive cell densities; therefore, a similar effect size as blood vessel densities was assumed. Future studies can use the data from this work, as a basis for power analysis, to investigate correlations between NPWT and inflammation or vascularity. Furthermore, it would be also beneficial to explore the effect of NPWT on other inflammatory markers, such as lymphocytes and macrophage phenotypes and their influence on the healing cascades present at the peri-prosthetic interfaces. A second limitation is that there are physiological differences in wound healing between rats and humans.44 Rats have a panniculus carnosus muscle which allows them to heal by contraction rather than re-epithelialization, the primary wound healing mechanism in humans. Interestingly, a recent study in human amputee patients without percutaneous implants, incorporated NPWT into socket-prosthetic caps and showed promising results to treat stump wounds,45 which highlights the opportunity for wider clinical translatability of NPWT. Thus, the results of this study need to be validated in a porcine model, which heals by re-epithelialization similar to humans,44 in order to 126 better understand the mechanism of action of NPWT. Lastly, although this study showed a significant increase in blood vessel densities due to the application of NPWT, the nature of these blood vessels (mature vs. premature) is unknown. Future studies should investigate the quality of the blood vessels via pericyte markers to determine blood vessel maturity. In summary, wound healing responses in percutaneous devices including epidermal downgrowth, vascularity and inflammation with and without NPWT were evaluated at 4 weeks in a hairless rat model. Animals, in both the NPWT and Untreated Groups, received a gauze and semi-occlusive base dressing and the NPWT Group received -70 to -90 mmHg of negative pressure treatment. While NPWT is known to induce vascularization in clinical wounds without a device, our results are the first to demonstrate that NPWT can increase blood vessel densities in peri-prosthetic tissues by nearly two-fold compared to untreated tissues. Establishing a blood supply to the peri-prosthetic tissues is important in maintaining the health of the tissue and thus, NPWT application may increase the longevity of percutaneous implants. Although no significant difference in downgrowth was found between the groups, the NPWT application resulted in reduced downgrowth compared to the Untreated Group. Additionally, while no significant difference in CD68 positive cell density was found between the groups at the wound edge, NPWT treatment appeared to recruit more CD68 positive cells to the sites. In the tissue over device, significantly increased CD68 positive cell density was found in the NPWT Group compared to the Untreated Group, which may suggest that NPWT modulates inflammation around percutaneous devices. 127 Overall, these findings may be important in developing strategies that improve the quality of the soft-tissues around implants especially in patients with impaired wound healing and insufficient vascularity such as diabetic or chronic wounds. 4.6 Acknowledgements This work was supported in part by the US Army Medical Research and Materiel Command under contract #W81XWH-15-C-0058, by the Department of Defense under grant #W81XWH-11-1-0435, by the United States Department of Veterans Affairs Rehabilitation Research and Development Service under Merit Review Awards #I01RX001217 and #I01RX001246, by the Department of Orthopaedics, University of Utah School of Medicine, Salt Lake City, Utah and by the LS-Peery Program in Musculoskeletal Restoration. The views, opinions, and/or findings presented are those of the authors and should not be construed as an official position, policy or decision of any of these funding sources unless so designated by other documentation. The authors would like to thank Gregory Stoddard for statistical consultation and Sheryl Tripp at ARUP laboratories (Salt Lake City, UT), for her support in handling histopathologic staining of tissues. 4.7 References 1.Pendegrass CJ, Goodship AE, Blunn GW. Development of a soft tissue seal around bone-anchored transcutaneous amputation prostheses. Biomaterials 2006;27(23):4183-91. 2.Holt BM, Bachus KN, Beck JP, Bloebaum RD, Jeyapalina S. Immediate post-implantation skin immobilization decreases skin regression around 128 percutaneous osseointegrated prosthetic implant systems. J Biomed Mater Res A 2013;101(7):2075-82. 3.von Recum AF. Applications and failure modes of percutaneous devices: a review. J Biomed Mater Res 1984;18(4):323-36. 4.Holt BM, Betz DH, Ford TA, Beck JP, Bloebaum RD, Jeyapalina S. Pig dorsum model for examining impaired wound healing at the skin-implant interface of percutaneous devices. J Mater Sci Mater Med 2013;24(9):2181-93. 5.Degidi M, Artese L, Scarano A, Perrotti V, Gehrke P, Piattelli A. Inflammatory infiltrate, microvessel density, nitric oxide synthase expression, vascular endothelial growth factor expression, and proliferative activity in periimplant soft tissues around titanium and zirconium oxide healing caps. J Periodontol 2006;77(1):73-80. 6.Wang Y, Zhang Y, Miron RJ. Health, maintenance, and recovery of soft tissues around implants. Clin Implant Dent Relat Res 2016;18(3):618-34. 7.Saghiri MA, Asatourian A, Garcia-Godoy F, Sheibani N. The role of angiogenesis in implant dentistry part I: Review of titanium alloys, surface characteristics and treatments. Med Oral Patol Oral Cir Bucal 2016;21(4):e514525. 8.Tonnesen MG, Feng X, Clark RA. Angiogenesis in wound healing. J Investig Dermatol Symp Proc 2000;5(1):40-6. 9.Broggini N, McManus LM, Hermann JS, Medina R, Schenk RK, Buser D, Cochran DL. Peri-implant inflammation defined by the implant-abutment interface. J Dent Res 2006;85(5):473-8. 10.Esposito M, Thomsen P, Ericson LE, Sennerby L, Lekholm U. Histopathologic observations on late oral implant failures. Clin Implant Dent Relat Res 2000;2(1):18-32. 11.Mitchell SJ, Jeyapalina S, Nichols FR, Agarwal J, Bachus KN. Negative pressure wound therapy limits downgrowth in percutaneous devices. Wound Repair Regen 2016;24(1):35-44. 12.Orgill DP, Bayer LR. Update on negative-pressure wound therapy. Plast Reconstr Surg 2011;127 Suppl 1:105S-115S. 13.Malmsjo M, Ingemansson R, Martin R, Huddleston E. Wound edge microvascular blood flow: effects of negative pressure wound therapy using gauze or polyurethane foam. Ann Plast Surg 2009;63(6):676-81. 129 14.Glass GE, Nanchahal J. The methodology of negative pressure wound therapy: separating fact from fiction. J Plast Reconstr Aesthet Surg 2012;65(8):989-1001. 15.Argenta LC, Morykwas MJ. Vacuum-assisted closure: a new method for wound control and treatment: clinical experience. Ann Plast Surg 1997;38(6):56376; discussion 577. 16.Morykwas MJ, Argenta LC, Shelton-Brown EI, McGuirt W. Vacuumassisted closure: a new method for wound control and treatment: animal studies and basic foundation. Ann Plast Surg 1997;38(6):553-62. 17.Greene AK, Puder M, Roy R, Arsenault D, Kwei S, Moses MA, Orgill DP. Microdeformational wound therapy: effects on angiogenesis and matrix metalloproteinases in chronic wounds of 3 debilitated patients. Ann Plast Surg 2006;56(4):418-22. 18.Eisenhardt SU, Schmidt Y, Thiele JR, Iblher N, Penna V, Torio-Padron N, Stark GB, Bannasch H. Negative pressure wound therapy reduces the ischaemia/reperfusion-associated inflammatory response in free muscle flaps. J Plast Reconstr Aesthet Surg 2012;65(5):640-9. 19.Borgquist O, Ingemansson R, Malmsjo M. Wound edge microvascular blood flow during negative-pressure wound therapy: examining the effects of pressures from -10 to -175 mmHg. Plast Reconstr Surg 2010;125(2):502-9. 20.Borgquist O, Ingemansson R, Malmsjo M. The influence of low and high pressure levels during negative-pressure wound therapy on wound contraction and fluid evacuation. Plast Reconstr Surg 2011;127(2):551-9. 21.Orgill DP, Bayer LR. Negative pressure wound therapy: past, present and future. Int Wound J 2013;10 Suppl 1:15-9. 22.Erba P, Ogawa R, Ackermann M, Adini A, Miele LF, Dastouri P, Helm D, Mentzer SJ, D'Amato RJ, Murphy GF and others. Angiogenesis in wounds treated by microdeformational wound therapy. Ann Surg 2011;253(2):402-9. 23.Glass GE, Murphy GF, Esmaeili A, Lai LM, Nanchahal J. Systematic review of molecular mechanism of action of negative-pressure wound therapy. Br J Surg 2014. 24.Norbury K, Kieswetter K. Vacuum-assisted closure therapy attenuates the inflammatory response in a porcine acute wound healing model. Wounds 2007;19(4):97-106. 25.Nuutila K, Siltanen A, Peura M, Harjula A, Nieminen T, Vuola J, Kankuri E, Aarnio P. Gene expression profiling of negative-pressure-treated skin graft donor site wounds. Burns 2013;39(4):687-93. 130 26.Liu D, Zhang L, Li T, Wang G, Du H, Hou H, Han L, Tang P. Negativepressure wound therapy enhances local inflammatory responses in acute infected soft-tissue wound. Cell Biochem Biophys 2014;70(1):539-47. 27.Jacobs S, Simhaee DA, Marsano A, Fomovsky GM, Niedt G, Wu JK. Efficacy and mechanisms of vacuum-assisted closure (VAC) therapy in promoting wound healing: a rodent model. J Plast Reconstr Aesthet Surg 2009;62(10):13318. 28.Cook SJ, Nichols FR, Brunker LB, Bachus KN. A novel vacuum assisted closure therapy model for use with percutaneous devices. Med Eng Phys 2014;36(6):768-73. 29.Pawar DRL, Mitchell SJ, Jeyapalina S, Hawkes JE, Florell SR, Bachus KN. Peri-prosthetic tissue reaction to discontinuation of negative pressure wound therapy around porous titanium percutaneous devices. J Biomed Mater Res B Appl Biomater 2018. 30.Scherer SS, Pietramaggiori G, Mathews JC, Prsa MJ, Huang S, Orgill DP. The mechanism of action of the vacuum-assisted closure device. Plast Reconstr Surg 2008;122(3):786-97. 31.Hugate R, Clarke R, Hoeman T, Friedman A. Transcutaneous implants in a porcine model: the use of highly porous tantalum. Int J Adv Mater Res 2015;1(2):32-40. 32.Pastar I, Stojadinovic O, Yin NC, Ramirez H, Nusbaum AG, Sawaya A, Patel SB, Khalid L, Isseroff RR, Tomic-Canic M. Epithelialization in wound healing: a comprehensive review. Adv Wound Care (New Rochelle) 2014;3(7):445-464. 33.Gal P, Toporcer T, Vidinsky B, Mokry M, Novotny M, Kilik R, Smetana K, Jr., Gal T, Sabo J. Early changes in the tensile strength and morphology of primary sutured skin wounds in rats. Folia Biol (Praha) 2006;52(4):109-15. 34.Jeyapalina S, Beck JP, Agarwal J, Bachus KN. A 24-month evaluation of a percutaneous osseointegrated limb-skin interface in an ovine amputation model. J Mater Sci Mater Med 2017;28(11):179. 35.Labler L, Rancan M, Mica L, Harter L, Mihic-Probst D, Keel M. Vacuumassisted closure therapy increases local interleukin-8 and vascular endothelial growth factor levels in traumatic wounds. J Trauma 2009;66(3):749-57. 36.Isenhath SN, Fukano Y, Usui ML, Underwood RA, Irvin CA, Marshall AJ, Hauch KD, Ratner BD, Fleckman P, Olerud JE. A mouse model to evaluate the interface between skin and a percutaneous device. J Biomed Mater Res A 2007;83(4):915-22. 131 37.Holgers KM, Thomsen P, Tjellstrom A, Bjursten LM. Immunohistochemical study of the soft tissue around long-term skin-penetrating titanium implants. Biomaterials 1995;16(8):611-6. 38.Lalezari S, Lee CJ, Borovikova AA, Banyard DA, Paydar KZ, Wirth GA, Widgerow AD. Deconstructing negative pressure wound therapy. Int Wound J 2017;14(4):649-657. 39.Dini V, Miteva M, Romanelli P, Bertone M, Romanelli M. Immunohistochemical evaluation of venous leg ulcers before and after negative pressure wound therapy. Wounds 2011;23(9):257-66. 40.Bridges AW, Whitmire RE, Singh N, Templeman KL, Babensee JE, Lyon LA, Garcia AJ. Chronic inflammatory responses to microgel-based implant coatings. J Biomed Mater Res A 2010;94(1):252-8. 41.Badylak SF, Valentin JE, Ravindra AK, McCabe GP, Stewart-Akers AM. Macrophage phenotype as a determinant of biologic scaffold remodeling. Tissue Eng Part A 2008;14(11):1835-42. 42.Brown BN, Londono R, Tottey S, Zhang L, Kukla KA, Wolf MT, Daly KA, Reing JE, Badylak SF. Macrophage phenotype as a predictor of constructive remodeling following the implantation of biologically derived surgical mesh materials. Acta Biomater 2012;8(3):978-87. 43.Shou K, Niu Y, Zheng X, Ma Z, Jian C, Qi B, Hu X, Yu A. Enhancement of bone-marrow-derived mesenchymal stem cell angiogenic capacity by NPWT for a combinatorial therapy to promote wound healing with large defect. Biomed Res Int 2017;2017:7920265. 44.Summerfield A, Meurens F, Ricklin ME. The immunology of the porcine skin and its value as a model for human skin. Mol Immunol 2015;66(1):14-21. 45.Wise J, White A, Stinner DJ, Fergason JR. A unique application of negative pressure wound therapy used to facilitate patient engagement in the amputation recovery process. Adv Wound Care (New Rochelle) 2017;6(8):253260. 132 Figure 4.1. A) A magnified photograph of the gauze and semi-occlusive dressing with the NPWT delivery tube inserted onto the dressing and further secured with semi-occlusive dressings to obtain an airtight seal. B) A photograph showing the NPWT treatment delivery set-up: (a) a pump, (b) a NPWT delivery line attached to the lid, (c) animal cage and (d) a NPWT tube attachment to the dressing. The negative pressure administered was monitored through an externally connected pressure sensor (not shown in the photo). 133 Figure 4.2. A photograph of the animal dorsum indicating the tissue collection sites. The dotted circle indicates the outline of the underlying subdermal device and a top view of the percutaneous post (P). In each animal, tissue was collected over the subdermal device, sectioned and used for immunohistochemical (IHC) staining. The inset image shows a cross-section of the soft-tissue in which tissue within the solid semi-circle was defined as the “wound edge” and the solid box denoted “tissue over device”. The dotted line indicates the incision line close to the spine. The dotted box indicates “distal tissue” collected from each animal to serve as an internal control for density measurements. 134 Figure 4.3. Top panel - A representative set of photographic images of skinimplant exit site at 4 weeks post-implantation. (A) Untreated controls. (B) Treated with a regimen of NPWT. Tissues in both groups had limited subdermal device exposure. The bottom panel - A representative set of photomacrographs showing the position of tissue attachment (white arrows) with respect to the base of the post (white dotted line). There was no significant difference in downgrowth (p=0.19) between the Untreated (C) and NPWT (D) Groups. White scale bar – 1000 µm. 135 Figure 4.4. A set of representative photomicrographs of peri-prosthetic tissues showing the immunohistochemical staining for CD31 and counterstained with hematoxylin. Top panel – Untreated Group. Bottom panel – NPWT Group. Images show an overall increase in blood vessel density (black arrows) in tissues treated with NPWT (D and F) compared to the untreated tissues (C and E). The red semicircles and green dotted boxes in A and B denote the regions used for quantification of blood vessels at the wound edge and tissue over device respectively. Blood vessels at the wound edge were quantified within the union of a semi-circle of radius 500 µm (not drawn to scale) centered at the end of the epidermis (*) and the tissue. Regions within the red semi-circles and green dotted boxes are magnified in C, D, E, and F. Tissue distal from device, on the opposite of the spine was collected to serve as an internal control for each animal (G and H). Images A and B were taken at 4x magnification, scale bar- 500 µm. All other images were taken at 20x magnification, scale bar - 100 µm. 136 Figure 4.5. A bar-chart showing the blood vessel densities quantified in 3 regions. Overall, tissues treated with NPWT showed increased blood vessel densities compared to the untreated tissues. Comparing just the wound edges, NPWT treated tissue exhibited relatively high blood vessel density than Untreated (p<0.05). Within both the groups, region specific differences in blood vessel densities were observed with the wound edges showing increased blood vessel densities compared to the tissue over device (NPWT Group, p<0.001; Untreated Group, p<0.05). The tissue over device in the Untreated Group did not show any significant difference in blood vessel densities compared to the distal tissue (p=0.88). Values are expressed as mean ± SD. 137 Figure 4.6. A set of photomicrographs of Untreated and NPWT treated periprosthetic tissues at the wound edge showing immunohistochemical staining for CD68. The positive cells are stained in brown. The Untreated Group showed variable responses with some samples showing reduced CD68 positive cell densities at the wound edge (A) while other samples showed markedly increased cell densities (B). Although the NPWT Group showed (C) a trend towards increased CD68 positive cell densities at the wound edge, no significant differences were observed in CD68 positive cell densities in the wound edges between the Untreated and NPWT Groups (p=0.09). All images were taken at 20x magnification, scale bar- 100 µm. 138 Figure 4.7. A bar-chart showing the CD68 positive cell densities quantified in 2 regions. Comparing just the wound edges, NPWT treated tissue showed a trend towards increased CD68 positive cell densities compared to the Untreated Group. However, no significant difference in CD68 positive cell densities was found (p=0.09). In the tissue over device, significantly increased CD68 cell density was found in the NPWT group compared to the Untreated Group (p<0.05). Within both the groups, region specific differences in CD68 positive cell densities were observed with the wound edges showing increased CD68 positive cell densities compared to the tissue over device (p<0.05 for both the groups). Values are expressed as mean ± SD. 139 Figure 4.8. A representative set of photomicrographs showing the soft tissuedevice interfaces: (A) Untreated Group showing a sealed interface with the epidermis contacting the subdermal device, denoted by the white arrow. Proteinaceous debris was present adjacent to the tissue-implant interface (marked by white dotted circle). (B) NPWT Group showing a breached interface with the epidermis ending within the dermis (marked by the white arrow), with no contact to the device. In both images, the double asterisks (**) indicate hyperplastic region of the epidermal tissue which becomes thinner (*) closer to the 3-point junction. P denotes percutaneous post. All images were taken at 4x magnification with white scale bar -250 µm. CHAPTER 5 SUMMARY, CONCLUSIONS AND FUTURE WORK 5.1 Summary and Conclusions Percutaneous devices are used in a variety of medical applications. While percutaneous devices are being developed worldwide, there is a constant risk of device failure due to soft-tissue complications, which need to be addressed before their widespread use. Implantation of percutaneous devices disrupts the protective skin barrier and elicits wound healing responses. The epidermis surrounding the device responds to the break in the skin barrier by proliferating and migrating along the surface of the device, in an attempt to re-establish continuity. This mechanism is referred to as epidermal downgrowth and can lead to sinus tract formation and exposure of the underlying device which can harbor bacteria and provide a route to infection. To improve the long-term residence of percutaneous devices, there is a need to establish a robust skin-device seal and limit epidermal downgrowth. This dissertation investigated the use of negative pressure wound therapy (NPWT) and device surface topography as strategies to limit epidermal downgrowth around percutaneous devices. Chapter 2 evaluated the ability of NPWT to limit epidermal downgrowth after its discontinuation. Chapter 3 evaluated the effect of microgrooved surface topography to limit downgrowth around percutaneous 141 devices implanted in normal skin. Chapter 4 evaluated the effects of NPWT on peri-prosthetic soft-tissue characteristics which may limit epidermal downgrowth. In Chapter 2, the effect of discontinuing NPWT on epidermal downgrowth around percutaneous devices was investigated. Percutaneous devices were implanted in a guinea pig model, which received 4 weeks of NPWT treatment followed by 4 weeks of no NPWT (Discontinued Group). This group was compared to two historical controls where one group received NPWT treatment continuously for 4 weeks (NPWT Group) and the second group received no NPWT for 4 weeks (Untreated Group). Quantitative downgrowth analysis showed that the Discontinued Group exhibited approximately five times more downgrowth than the NPWT Group. Furthermore, the amount of downgrowth in the Discontinued Group was comparable to that of the Untreated Group. This indicated that NPWT limits epidermal downgrowth during its period of application and that downgrowth resumes after NPWT is discontinued. Qualitative analysis showed that the periprosthetic tissues treated with NPWT exhibited morphological differences compared to the Discontinued and Untreated Groups. From these results, we can infer that NPWT induced morphological changes in the peri-prosthetic tissues during its period of application which may have limited epidermal downgrowth. However, after the discontinuation of NPWT, these morphological changes reverted to that of untreated tissue and downgrowth resumed. Taken together, this study showed that a short-term application of NPWT may not have long-term effects on limiting downgrowth around percutaneous devices. In Chapter 3, device surface topography, specifically the microgrooved 142 topography, was evaluated as a strategy to limit epidermal downgrowth around percutaneous devices. The microgrooved topography effectively halts gingival tissue downgrowth around dental implants without assistance from other therapies such as NPWT. This study was the first to evaluate the microgrooved topography in limiting epidermal downgrowth around percutaneous devices implanted in normal skin. In order to test this, three percutaneous devices were designed to evaluate different topographies— microgrooved, porous and smooth. The porous and smooth topographies served as controls. The devices were implanted into three groups of guinea pigs for 4 weeks. Histological analyses revealed that the epidermis did not form a stable seal with the microgrooves, although evidence of previous epidermal attachments adjacent to the microgrooved post was found. This finding suggests that the epidermis may have initially formed attachments to the post, but over time, the microgrooved topography was unable to maintain these attachments and epidermal downgrowth continued. It was postulated that the microgrooved topography may have been unable to limit epidermal downgrowth due to intrinsic differences between gingival tissue and normal skin. Furthermore, the porous and the smooth topographies also showed a comparable amount of downgrowth to the microgrooved topography. Collectively, the findings suggest that topographical alterations may be inadequate to halt epidermal downgrowth and may need to be supplemented with other approaches to establish a robust epidermal-device seal. Chapter 4 aimed to evaluate the characteristics exhibited by peri-prosthetic tissues undergoing limited downgrowth. Towards this goal, downgrowth, 143 vascularity and inflammation were analyzed in a rat model, at a 4-week time point. One group of rats received NPWT and a second group served as untreated controls. A rat model was used due to the increased availability of antibodies for investigating vascularization and inflammation as opposed to that available for the guinea pig. Although not statistically significant, the NPWT Group showed half the amount of downgrowth as the Untreated Group. With regards to vascularity, the NPWT Group showed significantly increased blood vessel densities compared to the Untreated Group, which may have contributed to limited downgrowth around the devices. CD68 was used as a general marker for inflammation. The NPWTtreated tissues exhibited a trend towards increased CD68 positive cell densities compared to the Untreated Group. However, it is unknown what percentage of the CD68 cells were polarized towards an M1 (pro-inflammatory) or M2 (antiinflammatory) macrophage phenotype. Therefore, further investigations into the macrophage phenotypes are needed to gain insight on how NPWT modulates inflammation, which may clarify its role in limiting downgrowth. Overall, the results of this study showed that tissues treated with NPWT exhibited reduced downgrowth, showed significantly increased vascularity and showed a trend towards increased CD68 positive cell densities. These findings may be important towards developing strategies which influence morphological characteristics leading to improved wound healing outcomes, including limited downgrowth around percutaneous devices. 144 5.2 Challenges and Future Work 5.2.1 Challenges and future work based on Chapter 2 results In Chapter 2, we found that NPWT limited epidermal downgrowth around percutaneous devices during its period of application. Following the discontinuation of NPWT, epidermal downgrowth resumed, which suggested that the effects of NPWT dissipate after it is discontinued. To better understand the effects of NPWT termination on downgrowth, it may prove beneficial to compare the downgrowth measurements from the 8-week Discontinued Group to a group of animals that do not receive any NPWT for 8 weeks and a group that receives NPWT continuously for 8 weeks. If the amount of downgrowth at 8 weeks in the Untreated Group is significantly increased compared to the Discontinued Group, it would indicate that the effects of NPWT do not rapidly attenuate following its discontinuation and are moderately sustained. If the amount of downgrowth at 8 weeks following continuous NPWT is considerably greater than at 4 weeks, then it could potentially indicate that even continuous application of NPWT does not limit the progression of downgrowth over time. Instead of complete discontinuation, a future study could investigate the influence of a supplemental regimen of NPWT application on epidermal downgrowth. For example, one experimental approach could be that after the initial 4-week application, NPWT is applied once, twice, or three times a week for the remaining 4 weeks. If the resulting downgrowth from the experimental regimens is significantly reduced compared to the Discontinued Group, then that would help develop a potential follow-up therapy protocol. This would, in turn, increase the 145 potential for translation since the patient would not have to continuously receive NPWT for extended periods of time. Instead, NPWT could either be administered during periods of rest or during clinical visits. Additionally, another study could be performed in which NPWT would be administered periodically instead of continuous application over the initial 4 weeks. In open wounds, Scherer et al. found that application of NPWT for 4 hours every other day resulted in similar wound closure compared to continuous NPWT for 7 days, in a mouse model.1 Similarly, in percutaneous devices, application of NPWT for 6, 12 or 18 hours every other day may result in an equivalent or reduced amount of epidermal downgrowth compared to continuous application of NPWT. This may help determine if there is a reduced duration of NPWT application that can limit downgrowth, which would in turn make it more practical and potentially increase its translatability. In clinical wounds, depending on the size, type and location, NPWT is discontinued either after full wound closure is attained, the wound becomes superficial or when the wound has been optimized for secondary closure.2 After discontinuation of NPWT, if the wound has not completely closed, then it is either surgically closed or transitioned to another treatment modality such as moist wound healing dressings2 or hydroconductive dressings.3 Around percutaneous devices, similar adjunct treatment therapies could be used following the discontinuation of NPWT in order to maintain limited downgrowth. 146 5.2.2 Challenges and future work based on Chapter 3 results In Chapter 3, we found that the microgrooved topography was unable to limit epidermal downgrowth around percutaneous devices implanted in normal skin. Although the microgrooved topography halts gingival epithelial downgrowth around dental implants, there are environmental and anatomical differences between the gingival tissue and normal skin, which may influence its translation to percutaneous devices in normal skin applications. Differences include thicker gingival epithelium,4 faster wound repair with an increased re-epithelialization rate and reduced scar formation compared to skin wounds.4-6 Saliva also contains cytokines, growth factors and protease inhibitors which may impart better wound healing qualities to the gingival tissue.7 In order to elucidate factors that allow gingival epithelial cells to form attachments to devices resulting in limited downgrowth, in vitro studies are needed. Based on an in vitro study, Pendegrass et al. indicated that gingival epithelial cells exhibited improved attachment on titanium substrates by upregulating hemidesmosome expression compared to epidermal cells.8 Increased hemidesmosome expression may be one of the reasons why gingival epithelial cells show reduced downgrowth around microgrooved dental implants. To conduct future in vitro experiments, human gingival epithelial cells and human epidermal cells would be isolated from the gum and skin, respectively. These cells would then be seeded onto titanium substrates to analyze their responses to the microgrooved, porous or smooth topographies. Cell morphology and attachment would be assessed using immunofluorescence at various time points following 147 seeding. These studies may provide insights into ways to enhance epidermal cell attachment to devices and limit downgrowth. To encourage epidermal cell adhesion to device topographies, future studies should investigate the incorporation of coatings such as hydroxyapatite (HA) or laminin-5. HA is the main constituent of bone and teeth, and enhances the attachment and integration of soft-tissues to HA-coated titanium devices.9,10 Hemidesmosome-based cellular adhesion is regulated by laminin-5, which is considered crucial for adhesion of cells on titanium substrates.11 Therefore, coating the device with HA or laminin-5 may enhance epidermal cell attachment to the device and subsequently reduce epidermal downgrowth.12,13 However, as is the case with any coating, the long-term stability and potential for degradation will have to be evaluated prior to widespread use. In the present study, the microgrooves were placed over a 2 mm thick band at the distal end of the percutaneous post. The device was designed to mimic dental implants in which microgrooves are placed over the abutment region which comes in contact with the gingival epithelium. However, the microgrooves on the percutaneous post may not have provided enough surface area for tissue interaction, attachment and stabilization. Furthermore, studies have shown that dermal tissue attachment and integration play an important role in limiting epidermal downgrowth.14-16 Therefore, it would be useful to evaluate the application of the microgrooved topography to the subdermal component which will increase the available surface area for soft-tissue attachment. Lastly, it would be beneficial to incorporate non-invasive methods to monitor 148 downgrowth around percutaneous devices. This would allow for the determination of the initial position of the skin around the post immediately after surgery and for measurements to be made periodically during the duration of the study. In turn, this would allow for the quantification of the rate of downgrowth which may reveal periods where downgrowth progresses rapidly. Previously, Chehroudi et al. attempted to characterize the rate of downgrowth around percutaneous devices using vinyl silicone dental impression materials.15 However, a disadvantage of using this technique is that the impression material may compress the soft-tissue around the post and result in inaccurate measurements. Additionally, the presence of a surface coating on the device may also lead to inaccurate measurements. Utilizing digital measuring systems such as photography,17 electronic wound measurement device18 and laser analysis19 will enable accurate and reliable downgrowth measures, and help overcome the challenges associated with manual measurement methods (impression technique, rulers, vernier caliper). 5.2.3 Challenges and future work based on Chapter 4 results While this study reported that NPWT treated peri-prosthetic tissues showed significantly increased blood vessel densities (CD31), future studies should investigate the quality of the blood vessels to determine blood vessel maturity. Development of mature vessels is vital because insufficient maturity may lead to edema, hemorrhage and reduced transport of nutrients and oxygen required for wound closure.20 Previous studies have demonstrated that NPWT promotes vessel maturation in wounds without devices, which increases blood flow and 149 accelerates the speed of wound healing.21,22 Therefore, it would be useful to assess if NPWT has a similar effect on vessel maturity in peri-prosthetic tissues. To investigate blood vessel maturation, double staining immunohistochemistry (IHC) using antibodies for CD31 and alpha-smooth muscle actin (α-SMA) can be performed. The α-SMA is typically used as a marker for pericytes which wrap around endothelial cells in mature blood vessels. Thus, co-localization of the markers for endothelial cells and pericytes can indicate the number of mature blood vessels present in the tissue.23 If NPWT application is found to induce immature blood vessels, further studies might investigate pro-angiogenic therapies to promote long-term maturation and stabilization of vessels.20 While increased blood vessel densities may have been one of the possible reasons for limited downgrowth, it is unclear if a correlation exists between the two factors. Future studies should investigate if downgrowth may be linked to periprosthetic tissue vascularity by evaluating NPWT and no-NPWT using a pig model. The pig model exhibits downgrowth,24,25 undergoes similar wound healing as humans (re-epithelialization) and has a similar dermal blood supply as humans.26 If an association is found to exist, further studies could be performed to assess the direct effect of vascularity in limiting downgrowth around percutaneous devices. One study would be to implant percutaneous devices in pigs, with one group of animals receiving a pro-angiogenic treatment and a control group receiving no treatment. The expected result would be that the group receiving treatment will exhibit increased vascularity and reduced downgrowth compared to the control group. Angiogenic therapy such as the application of topical vascular endothelial 150 growth factor (VEGF) has shown potential to improve wound healing by enhancing vascularization in open wounds.27,28 However, the dosage and potential side effects of VEGF therapy, such as increased permeability of vessels and scar formation, will need to be considered to maximize treatment efficacy.27,28 The NPWT Group showed a trend towards increased CD68 cell densities at the wound edge and showed significantly increased CD68 cell densities in the tissue away from the wound edge, compared to the Untreated Group. One possible reason for the increased CD68 positive cell densities in the NPWT Group could have been due to the application of NPWT. Since there is a lack of studies quantifying CD68 in peri-prosthetic tissues treated with NPWT, further studies in other animal models, such as pigs, are needed to validate our results. In this study, high variability in the CD68 positive cell densities at the wound edge in the Untreated Group was observed. Micro-motion at the skin-implant interface could have been one possible reason for the variability in the amount of CD68 positive cell densities. Some animals in the Untreated Group may have experienced increased micro-motion compared to the other animals, leading to increased skin irritation and, as a result, higher CD68 positive cells. On the other hand, mechanical forces due to the application of NPWT may have mitigated the effects of micro-motion, leading to reduced variability in the NPWT Group. Finally, the presence of microbes at the skin-device interface may have increased CD68 positive cell densities.29 In order to limit the pathogenic bio-load, dressings were applied over the device in both the groups and no clinical signs of infection were observed around the skin-device exit site. However, studies have reported that 151 Staphylococcus aureus and coagulase-negative staphylococci, such as Staphylococcus epidermidis, are commonly found at the skin-device interface and account for most infections associated with percutaneous osseointegrated devices.30,31 Future studies could include swab cultures taken from the skin-device exit site to determine the extent of pathogenic load present.30,32 Quantitative molecular methods such as PCR could be employed to enhance the detection of organisms and markers. A study found a positive correlation between TNF‐α and MMP-8 expression and the presence of Staphylococcus aureus in infected percutaneous osseointegrated devices, using swab cultures and PCR.32 Studies have also indicated that changes in inflammatory markers such as C-reactive protein (CRP), IL-6, polymorphonuclear leukocytes, procalcitonin and erythrocyte sedimentation rate can be used to detect infections.33-35 For example, CRP greater than 120 mg/L, evaluated using blood tests, has been associated with bacterial infection in prosthetics.36 Future studies can include blood tests (CRP, erythrocyte sedimentation rate, IL-6), blood cultures (presence of micro-organisms) and histopathological analysis (polymorphonuclear leukocytes) to determine the levels of various markers present around percutaneous devices to evaluate infection. While CD68 is used as a marker for cells from the monocyte-macrophage lineage,37,38 in vitro studies have indicated that other cell types such as endothelial cells, basophils and neutrophils may also express CD68.39-41 Therefore, future experiments should perform either double or triple IHC staining with additional markers to classify the CD68 positive cells. For example, cells that show positive staining for CD68 and either neutrophil elastase or myeloperoxidase would be 152 regarded as neutrophils.39 Cells that are positive for CD68 but negative for neutrophil elastase or myeloperoxidase would be regarded as monocytemacrophage cells. Furthermore, macrophages can acquire a spectrum of phenotypes. Macrophages at either end of the spectrum are referred to as M1 (proinflammatory) and M2 (anti-inflammatory and pro-remodeling).42 IHC staining using markers such as CCR7, CD86 for M1 phenotypes and CD206, CD163 for M2 phenotypes could be used to classify the macrophages.38,43 Phenotype identification may provide insights into the role of NPWT in inducing either a proor anti-inflammatory wound environment around percutaneous devices. Based on this knowledge, strategies could be introduced to modulate inflammation for improved wound healing.44 All the NPWT treated tissues and 60% of the untreated tissues exhibited breached interfaces, that is, lack of epidermal-device attachment. One possible reason for the occurrence of a breached epidermal response could be due to micro-motion at the skin-device interface. The rat model inherently has a thinner epidermis compared to the guinea pig or the pig,45 which may make the rat epidermis more susceptible to disruption due to micro-motion. In our study, mechanical forces may have disrupted any thin epidermal attachments to the device, resulting in a breached interface. There is a lack of studies that report whether epidermal responses to percutaneous devices are breached or sealed. It is difficult to state whether this is due to the absence of breached interfaces in other animal models or if this factor is overlooked. Therefore, future percutaneous device studies, especially those utilizing a rat model, should evaluate the 153 epidermal response and consider whether this factor will influence their experiment. 5.2.4 Challenges and future work based on overall results Although animal models yield some promising results, there are inherent differences that should be considered before extrapolating the results from the animal based studies to humans. The skin in guinea pigs and rats is loosely connected to the underlying subcutaneous connective tissue.45 Additionally, both these animals have a panniculus carnosus muscle which contributes to wound contraction during wound healing.26 On the other hand, human and pig lack the panniculus carnosus muscle and their skin is tightly attached to the connective tissue.45 Therefore, while the results from studies involving guinea pigs and rats provide important basic knowledge, the physiological and anatomical differences between these animals and humans may limit reproducibility, clinical relevance and interpretation of results. Thus, future studies should utilize a more translational model, such as the pig, to validate the results and assess wound healing responses, including epidermal downgrowth, around percutaneous devices. All three studies involved the use of metal percutaneous devices. In order to perform histological analyses such as downgrowth measurements, the devices had to be embedded in polymethylmethacrylate (PMMA), which served as a medium for sectioning, grinding and polishing the metallic devices and the surrounding soft-tissues.46 While classical histological stains such as hematoxylin and eosin can be performed on the resulting PMMA sections, IHC staining is 154 limited due to the inability of the antibodies to penetrate the plastic sections and high polymerization temperatures. In Chapter 4, the author attempted to utilize a low temperature methylmethacrylate (MMA)-based embedding system (Technovit® 9100, Heraeus Kulzer, Germany) with the goal to perform IHC staining. However, due to technical difficulties during the polymerization process, the author was unable to do so. Future studies should develop a protocol for performing IHC using PMMA, by optimizing temperature and pressure ranges, catalyst ratio and polymerization rates.47,48 Future studies should also utilize a more sensitive and robust molecular technique such as real-time quantitative polymerase chain reaction (PCR) to characterize the molecular mechanisms and cues involved in epidermal cell migration, leading to downgrowth. To do so, peri-prosthetic tissue should be collected off of the device. The region of interest can be isolated using a precise technique such as laser capture microdissection.49 Following isolation, PCR would then be performed to evaluate gene expression, which would provide knowledge regarding genes implicated in downgrowth. Lastly, the percutaneous devices investigated in this dissertation were not bone-anchored. While the downgrowth phenomenon has been reported to occur in both bone-anchored50 and subdermal barrier only percutaneous devices,25 the author acknowledges that interfacial forces at the device exit site may exacerbate the rate of downgrowth and impact stable soft-tissue integration around the devices. This may affect the interpretation of the results obtained from unanchored devices to anchored devices. Therefore, follow-up studies would be beneficial to 155 validate the results using a bone-anchored model. 5.3 References 1.Scherer SS, Pietramaggiori G, Mathews JC, Orgill DP. Short periodic applications of the vacuum-assisted closure device cause an extended tissue response in the diabetic mouse model. Plast Reconstr Surg 2009;124(5):1458-65. 2.Bollero D, Driver V, Glat P, Gupta S, Lazaro-Martinez JL, Lyder C, Ottonello M, Pelham F, Vig S, Woo K. The role of negative pressure wound therapy in the spectrum of wound healing. Ostomy Wound Manage 2010;56(5 Suppl):118. 3.Sood A, Granick MS, Tomaselli NL. Wound dressings and comparative effectiveness data. Adv Wound Care (New Rochelle) 2014;3(8):511-529. 4.Boink MA, Roffel S, Breetveld M, Thon M, Haasjes MSP, Waaijman T, Scheper RJ, Blok CS, Gibbs S. Comparison of advanced therapy medicinal product gingiva and skin substitutes and their in vitro wound healing potentials. J Tissue Eng Regen Med 2018;12(2):e1088-e1097. 5.Glim JE, van Egmond M, Niessen FB, Everts V, Beelen RH. Detrimental dermal wound healing: what can we learn from the oral mucosa? Wound Repair Regen 2013;21(5):648-60. 6.Turabelidze A, Guo S, Chung AY, Chen L, Dai Y, Marucha PT, DiPietro LA. Intrinsic differences between oral and skin keratinocytes. PLoS One 2014;9(9):e101480. 7.Zelles T, Purushotham KR, Macauley SP, Oxford GE, Humphreys-Beher MG. Saliva and growth factors: the fountain of youth resides in us all. J Dent Res 1995;74(12):1826-32. 8.Pendegrass CJ, Lancashire HT, Fontaine C, Chan G, Hosseini P, Blunn GW. Intraosseous transcutaneous amputation prostheses versus dental implants: a comparison between keratinocyte and gingival epithelial cell adhesion in vitro. Eur Cell Mater 2015;29:237-49. 9.Larsson A, Andersson M, Wigren S, Pivodic A, Flynn M, Nannmark U. Soft tissue integration of hydroxyapatite-coated abutments for bone conduction implants. Clin Implant Dent Relat Res 2015;17 Suppl 2:e730-5. 10.Larsson A, Wigren S, Andersson M, Ekeroth G, Flynn M, Nannmark U. Histologic evaluation of soft tissue integration of experimental abutments for bone anchored hearing implants using surgery without soft tissue reduction. Otol Neurotol 2012;33(8):1445-51. 156 11.Shiraiwa M, Goto T, Yoshinari M, Koyano K, Tanaka T. A study of the initial attachment and subsequent behavior of rat oral epithelial cells cultured on titanium. J Periodontol 2002;73(8):852-60. 12.Gordon DJ, Bhagawati DD, Pendegrass CJ, Middleton CA, Blunn GW. Modification of titanium alloy surfaces for percutaneous implants by covalently attaching laminin. J Biomed Mater Res A 2010;94(2):586-93. 13.El-Ghannam A, Starr L, Jones J. Laminin-5 coating enhances epithelial cell attachment, spreading, and hemidesmosome assembly on Ti-6A1-4V implant material in vitro. J Biomed Mater Res 1998;41(1):30-40. 14.Pendegrass CJ, Goodship AE, Blunn GW. Development of a soft tissue seal around bone-anchored transcutaneous amputation prostheses. Biomaterials 2006;27(23):4183-91. 15.Chehroudi B, Brunette DM. Subcutaneous microfabricated surfaces inhibit epithelial recession and promote long-term survival of percutaneous implants. Biomaterials 2002;23(1):229-37. 16.Chehroudi B, Gould TR, Brunette DM. The role of connective tissue in inhibiting epithelial downgrowth on titanium-coated percutaneous implants. J Biomed Mater Res 1992;26(4):493-515. 17.Chang AC, Dearman B, Greenwood JE. A comparison of wound area measurement techniques: visitrak versus photography. Eplasty 2011;11:e18. 18.Hammond CE, Nixon MA. The reliability of a handheld wound measurement and documentation device in clinical practice. J Wound Ostomy Continence Nurs 2011;38(3):260-4. 19.Pavlovcic U, Diaci J, Mozina J, Jezersek M. Wound perimeter, area, and volume measurement based on laser 3D and color acquisition. Biomed Eng Online 2015;14:39. 20.Zhao J, Chen L, Shu B, Tang J, Zhang L, Xie J, Qi S, Xu Y. Granulocyte/macrophage colony-stimulating factor influences angiogenesis by regulating the coordinated expression of VEGF and the Ang/Tie system. PLoS One 2014;9(3):e92691. 21.Ma Z, Shou K, Li Z, Jian C, Qi B, Yu A. Negative pressure wound therapy promotes vessel destabilization and maturation at various stages of wound healing and thus influences wound prognosis. Exp Ther Med 2016;11(4):1307-1317. 22.Ma Z, Li Z, Shou K, Jian C, Li P, Niu Y, Qi B, Yu A. Negative pressure wound therapy: regulating blood flow perfusion and microvessel maturation through microvascular pericytes. Int J Mol Med 2017;40(5):1415-1425. 157 23.Bergers G, Song S. The role of pericytes in blood-vessel formation and maintenance. Neuro Oncol 2005;7(4):452-64. 24.Hugate R, Clarke R, Hoeman T, Friedman A. Transcutaneous implants in a porcine model: the use of highly porous tantalum. Int J Adv Mater Res 2015;1(2):32-40. 25.Holt BM, Betz DH, Ford TA, Beck JP, Bloebaum RD, Jeyapalina S. Pig dorsum model for examining impaired wound healing at the skin-implant interface of percutaneous devices. J Mater Sci: Mater Med 2013;24:2181-2193. 26.Sullivan TP, Eaglstein WH, Davis SC, Mertz P. The pig as a model for human wound healing. Wound Repair Regen 2001;9(2):66-76. 27.Johnson KE, Wilgus TA. Vascular endothelial growth factor and angiogenesis in the regulation of cutaneous wound repair. Adv Wound Care (New Rochelle) 2014;3(10):647-661. 28.Wietecha MS, DiPietro LA. Therapeutic approaches to the regulation of wound angiogenesis. Adv Wound Care (New Rochelle) 2013;2(3):81-86. 29.Holgers KM. Characteristics of the inflammatory process around skinpenetrating titanium implants for aural rehabilitation. Audiology 2000;39(5):253-9. 30.Tillander J, Hagberg K, Hagberg L, Branemark R. Osseointegrated titanium implants for limb prostheses attachments: infectious complications. Clin Orthop Relat Res 2010;468(10):2781-8. 31.Zaborowska M, Tillander J, Branemark R, Hagberg L, Thomsen P, Trobos M. Biofilm formation and antimicrobial susceptibility of staphylococci and enterococci from osteomyelitis associated with percutaneous orthopaedic implants. J Biomed Mater Res B Appl Biomater 2017;105(8):2630-2640. 32.Lenneras M, Tsikandylakis G, Trobos M, Omar O, Vazirisani F, Palmquist A, Berlin O, Branemark R, Thomsen P. The clinical, radiological, microbiological, and molecular profile of the skin-penetration site of transfemoral amputees treated with bone-anchored prostheses. J Biomed Mater Res A 2017;105(2):578-589. 33.Markanday A. Acute phase reactants in infections: evidence-based review and a guide for clinicians. Open Forum Infect Dis 2015;2(3):ofv098. 34.Berbari E, Mabry T, Tsaras G, Spangehl M, Erwin PJ, Murad MH, Steckelberg J, Osmon D. Inflammatory blood laboratory levels as markers of prosthetic joint infection: a systematic review and meta-analysis. J Bone Joint Surg Am 2010;92(11):2102-9. 158 35.Nishikawa H, Shirano M, Kasamatsu Y, Morimura A, Iida K, Kishi T, Goto T, Okamoto S, Ehara E. Comparative usefulness of inflammatory markers to indicate bacterial infection-analyzed according to blood culture results and related clinical factors. Diagn Microbiol Infect Dis 2016;84(1):69-73. 36.Tornero E, Garcia-Oltra E, Garcia-Ramiro S, Martinez-Pastor JC, Bosch J, Climent C, Morata L, Camacho P, Mensa J, Soriano A. Prosthetic joint infections due to Staphylococcus aureus and coagulase-negative staphylococci. Int J Artif Organs 2012;35(10):884-92. 37.Barros MH, Hauck F, Dreyer JH, Kempkes B, Niedobitek G. Macrophage polarisation: an immunohistochemical approach for identifying M1 and M2 macrophages. PLoS One 2013;8(11):e80908. 38.Brown BN, Londono R, Tottey S, Zhang L, Kukla KA, Wolf MT, Daly KA, Reing JE, Badylak SF. Macrophage phenotype as a predictor of constructive remodeling following the implantation of biologically derived surgical mesh materials. Acta Biomater 2012;8(3):978-87. 39.Amanzada A, Malik IA, Blaschke M, Khan S, Rahman H, Ramadori G, Moriconi F. Identification of CD68(+) neutrophil granulocytes in in vitro model of acute inflammation and inflammatory bowel disease. Int J Clin Exp Pathol 2013;6(4):561-70. 40.Gottfried E, Kunz-Schughart LA, Weber A, Rehli M, Peuker A, Muller A, Kastenberger M, Brockhoff G, Andreesen R, Kreutz M. Expression of CD68 in nonmyeloid cell types. Scand J Immunol 2008;67(5):453-63. 41.Pulford KA, Sipos A, Cordell JL, Stross WP, Mason DY. Distribution of the CD68 macrophage/myeloid associated antigen. Int Immunol 1990;2(10):97380. 42.Alvarez MM, Liu JC, Trujillo-de Santiago G, Cha BH, Vishwakarma A, Ghaemmaghami AM, Khademhosseini A. Delivery strategies to control inflammatory response: modulating M1-M2 polarization in tissue engineering applications. J Control Release 2016;240:349-363. 43.Faulk DM, Londono R, Wolf MT, Ranallo CA, Carruthers CA, Wildemann JD, Dearth CL, Badylak SF. ECM hydrogel coating mitigates the chronic inflammatory response to polypropylene mesh. Biomaterials 2014;35(30):8585-95. 44.Krzyszczyk P, Schloss R, Palmer A, Berthiaume F. The role of macrophages in acute and chronic wound healing and interventions to promote pro-wound healing phenotypes. Front Physiol 2018;9:419. 45.Summerfield A, Meurens F, Ricklin ME. The immunology of the porcine skin and its value as a model for human skin. Mol Immunol 2015;66(1):14-21. 159 46.Emmanual J, Hornbeck C, Bloebaum RD. A polymethyl methacrylate method for large specimens of mineralized bone with implants. Stain Technol 1987;62(6):401-10. 47.Yang R, Davies CM, Archer CW, Richards RG. Immunohistochemistry of matrix markers in Technovit 9100 New-embedded undecalcified bone sections. Eur Cell Mater 2003;6:57-71; discussion 71. 48.Quentin T, Poppe A, Bar K, Sigler A, Foth R, Michel-Behnke I, Paul T, Sigler M. A novel method for processing resin-embedded specimens with metal implants for immunohistochemical labelling. Acta Histochem 2009;111(6):538-42. 49.Fend F, Raffeld M. Laser capture microdissection in pathology. J Clin Pathol 2000;53(9):666-72. 50.Holt BM, Bachus KN, Beck JP, Bloebaum RD, Jeyapalina S. Immediate post-implantation skin immobilization decreases skin regression around percutaneous osseointegrated prosthetic implant systems. J Biomed Mater Res A 2013;101(7):2075-82. APPENDIX XPS ANALYSIS OF DEVICE SURFACE 161 Prior to implantation, all devices were cleaned according to the ASTM F86 standards using nitric acid treatment. Subsequently, the devices were packaged in sterilization pouches and steam sterilized. In industry, it is common practice to use nitric acid, as referenced in ASTM-F86, to clean metallic surgical implants manufactured from iron, cobalt, titanium and tantalum base materials. ASTM-F86 lists that nitric acid treatment provides passivation by surface oxidation and is able to remove certain foreign material such as iron or other exogenous materials from the surface, which may be present from manufacturing operations. In 2000, it was reported that Sulzer Orthopaedics experienced high revision rates of acetabular shell implants in lots that were manufactured without the nitric acid passivation step.1 It was reported that the principle cause of implant failure was due the omission of the nitric acid treatment step.2 To conform to the current standard specifications, the studies performed in this dissertation followed the protocol listed in ASTM-F86. The devices were immersed in 35% nitric acid at room temperature for a minimum of 30 minutes. Our research lab has been cleaning Ti-based implants using the nitric acid method and has not faced any adverse events such as infections or the loss of animals. Previous studies have utilized XPS to assess the chemical composition of titanium substrates passivated with nitric acid.3,4 Studies report that the surface of devices treated with nitric acid consists of a titanium concentration between 1926%, oxygen concentration between 50-54% and carbon concentration between 19-26%.3,4 In the present study, XPS analysis revealed the titanium concentration to be between 11-16%, oxygen concentration to be between 37-40% and carbon 162 concentration to be between 36-49%. Carbon concentrations of 20-30% are typical for even well-cleaned titanium surfaces that have been exposed to air.5 Every titanium surface adsorbs hydrocarbons or carbon-oxygen containing species within microseconds of being exposed to ambient conditions. One possible reason for the increased amount of carbon on our devices could be due to packaging. In this work, we used standard sterile packaging pouches (Duo-Check® sterilization pouches, Crosstex, Maumee, OH) which are used for packaging clinical instruments and are manufactured from medical grade paper and polyester cast polypropelene. However, contact of the devices with the pouches may modify the surface, leading to higher carbon concentrations.6 Future studies may be beneficial to determine the effect of the sterile pouches on the percentage of carbon present on the device surface by analyzing surface carbon concentrations before and after placing the device in a sterile package. A second possible reason for the high carbon concentration may have been due to the use of steam sterilization.4,5 Steam sterilization may increase the concentration of hydrocarbon species, resulting in the detection of increased carbon on the device surface.5,7 Further studies should determine carbon concentrations before and after steam sterilization and also consider the use of other cleaning techniques such as plasma treatment to reduce organic species on the surface of the devices.5 For all four device components, high-resolution XPS spectra were obtained at O 1s, Ti 2p, C 1s, N 1s and Al 2p (Figures A.1, A.2, A.3, A.4 and A.5). Highresolution spectra for Si 2p were obtained for the smooth and microgrooved percutaneous posts (Figure A.6). In order to determine the specific molecular 163 species present, the high-resolution spectra have to be deconvoluted into subpeaks and curve fitting needs to be performed. Although curve fitting analysis was not performed in this work, studies have demonstrated the binding energies that are associated with various functional groups.8 In this study, based upon the position of a peak at 285 eV, the high-resolution C 1s spectra suggests the presence of a hydrocarbon functional group (C-H, C-C). A secondary peak at approximately 288 eV may suggest the presence of carbon species with a carbonyl group (C=O). Thus, the surface of the devices contained organic species such as hydrocarbons or carboxylates, whose concentration may depend on sterilization, packaging and the exposure time to ambient conditions.9 The high-resolution spectra of Ti 2p indicated two peaks at binding energies of 459 eV and 464 eV. These peaks represent Ti 2p1/2 and Ti 2p3/2 of TiO2. Additionally, a peak at binding energy of approximately 454 eV is indicative of Ti metal. The high-resolution spectra of O 1s for all the four device components indicated a peak at approximately 530 eV binding energy which is representative of O 1s originating from a titanium oxide film. Additionally, the microgrooved post also displayed a peak between 532-533 eV which may be indicative of the presence of oxygencontaining species such as hydroxides (C-OH). Si was detected on the surface of the smooth and microgrooved posts. The Si 2p peak at approximately 102 eV binding energy may be indicative of the presence of organosilicon residues.10 However, from XPS data alone, it is not possible to determine the origin of this chemical species. Si is a common surface contaminant, and has been found on the surface of commercially available implants as well.10 164 Comparing the surface elemental composition of the devices in Chapter 3 to the results of XPS analysis performed on commercially available titanium implants, the findings indicate that the elemental composition values are comparable.11 Massaro et al. analyzed the surface elemental composition of commercially available dental implants (Branemark, TPS, SLA, ICE, Osseotite).10 For example, the average atomic % of titanium in the five implants was reported to be 9.8% which is comparable to the average reported in Chapter 3 of 13.98%. Similarly, the average atomic % carbon was reported to be 45.8% and the average atomic % oxygen was reported to be 41.2%. In our study, the average atomic % of carbon was 43.58% and atomic % oxygen was 38.09%. Although Massaro et al. and other studies refer to the surface of the commercially available implants as consisting of a titanium dioxide layer (TiO2),3,8,10 quantitatively, the XPS results suggest that majority of the surface contains a high percentage of carbon. Overall, the relatively high amount of carbon present on the surfaces of the titanium devices reported in this study, and on the devices reported in the literature, indicates that carbon is one of the primary elements which interacts with the host cells after implantation. Future studies should be careful to use appropriate terminology when referring to the surface of the devices made up of bulk titanium. References 1.Spiegelberg S, Deluzio K, Muratoglu O. Extractable residue from recalled inter-op acetabular shells. 2003; 49th Annual Meeting of Orthopaedic Research Society, New Orleans. 2.Bonsignore LA, Goldberg VM, Greenfield EM. Machine oil inhibits the osseointegration of orthopaedic implants by impairing osteoblast attachment and spreading. J Orthop Res 2015;33(7):979-87. 165 3.Sittig C, Textor M, Spencer ND, Wieland M, Vallotton PH. Surface characterization of implant materials c.p. Ti, Ti-6Al-7Nb and Ti-6Al-4V with different pretreatments. J Mater Sci Mater Med 1999;10(1):35-46. 4.Wieland M. Experimental determination and quantitative evaluation of the surface composition and topography of medical implant surfaces and their influence on osteoblastic cell-surface interactions: ETH Zurich; 1999. 5.Brunette DM, Tengvall P, Textor M, Thomsen P. Titanium in medicine: material science, surface science, engineering, biological responses and medical applications. Springer Science & Business Media; 2012. 6.Kasemo B, Lausmaa J. Biomaterial and implant surfaces: on the role of cleanliness, contamination, and preparation procedures. J Biomed Mater Res 1988;22(A2 Suppl):145-58. 7.Baier RE, Meyer AE, Akers CK, Natiella JR, Meenaghan M, Carter JM. Degradative effects of conventional steam sterilization on biomaterial surfaces. Biomaterials 1982;3(4):241-5. 8.Kang BS, Sul YT, Oh SJ, Lee HJ, Albrektsson T. XPS, AES and SEM analysis of recent dental implants. Acta Biomater 2009;5(6):2222-9. 9.Lausmaa J. Surface spectroscopic characterization of titanium implant materials. J Electron Spectros Relat Phenomena 1996;81(3):343-361. 10.Massaro C, Rotolo P, De Riccardis F, Milella E, Napoli A, Wieland M, Textor M, Spencer ND, Brunette DM. Comparative investigation of the surface properties of commercial titanium dental implants. Part I: chemical composition. J Mater Sci Mater Med 2002;13(6):535-48. 11.Castilho GA, Martins MD, Macedo WA. Surface characterization of titanium based dental implants. Brazilian Journal of Physics 2006;36(3B):10041008. 166 Figure A.1. High-resolution O 1s spectra obtained from XPS for all device components. 167 Figure A.2. High-resolution Ti 2p spectra obtained from XPS for all device components. 168 Figure A.3. High-resolution C 1s spectra obtained from XPS for all device components. 169 Figure A.4. High-resolution N 1s spectra obtained from XPS for all device components. 170 Figure A.5. High-resolution Al 2p spectra obtained from XPS for all device components. 171 Figure A.6. High-resolution Si 2p spectra obtained from XPS for smooth and microgrooved devices.