Adipose Tissue Extract for Wound Healing : Operating room preparation, in vitro and clinical use

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Adipose Tissue Extract for Wound Healing

Operating room preparation, in vitro and clinical use

JENNY LOPEZ

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Tampere University Dissertations 324

JENNY LOPEZ

Adipose Tissue Extract for Wound Healing

Operating room preparation, in vitro and clinical use

ACADEMIC DISSERTATION To be presented, with the permission of the Faculty of Medicine and Health Technology

of Tampere University,

for public discussion in the Auditorium F114 of the Arvo Building, Arvo Ylpön katu 34, Tampere,

on 13 November 2020, at 12 o’clock

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ACADEMIC DISSERTATION

Tampere University, Faculty of Medicine and Health Technology Finland

Responsible

supervisor Docent, Professor h.c.

Hannu Kuokkanen Tampere University Finland

Supervisors Docent Ilkka Kaartinen Tampere University Finland

Professor Timo Ylikomi Tampere University Finland

Pre-examiners Professor Salvatore Giordano University of Turku

Finland

Docent Kristo Nuutila University of Helsinki Finland

Opponent Docent Susanna Kauhanen University of Helsinki Finland

Custos Associate professor (tenure track) Ville Mattila

Tampere University Finland

The originality of this thesis has been checked using the Turnitin OriginalityCheck service.

ISBN 978-952-03-1729-4 (print) ISBN 978-952-03-1730-0 (pdf) ISSN 2489-9860 (print) ISSN 2490-0028 (pdf)

http://urn.fi/URN:ISBN:978-952-03-1730-0

PunaMusta Oy – Yliopistopaino Tampere 2020

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To God.

To my family.

To all that inspired this research.

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ACKNOWLEDGEMENTS

My sincerest gratitude to Prof. Hannu Kuokkanen, MD, PhD for welcoming me into a life of research, for being a true mentor and for supporting me whilst supervising my work throughout the years. Your experience and feedback proved to be very valuable in achieving our joint research goals.

Ilkka Kaartinen, PD, PhD, thank you for supervising my studies and dedicating the time and effort that made them what they are. Thank you for collaborating and sharing your innovative ideas, statistical assistance and for providing the equipment required for the clinical studies. Your role in this journey was crucial and I cannot thank you enough.

The studies were carried out in the Department of Cell Biology and the Finnish Center for Alternative Methods (FICAM), in the School of Medicine, at the University of Tampere. The human study was performed in the Department of Musculoskeletal Diseases, Plastic Surgery Unit, at Tampere University Hospital. I would like to express my appreciation to all the dedicated staff and these two locations for making these studies possible. Thanks to the Finnish Technology and Innovation Agency TEKES, and Competitive State Research Financing of the Expert Responsibility area of Tampere University Hospital for their support in funding the studies.

My deepest gratitude to Prof. Timo Ylikomi, MD, PhD, for his guidance throughout experiments and for welcoming me into his incredible team that expanded my curiosity and passion in the area of tissue engineering. Your guidance and experience helped envision the studies, with you adding your valuable perspective every step of the way.

Riina Sarkanen, MSc. PhD, no words can ever express my gratitude towards you.

You have been the fuel behind these studies. Without your vision, your knowledge, experience and time, none if these studies would have been possible. Thank you for your valuable guidance at every step of the way.

A special thanks to Outi Huttala, MSc. PhD, for being a very treasured part of the team. Your persistent presence and aid with laboratory studies were crucial to achieving our study goals. Your incredible academic research and statistical experience was a key factor in finalizing the studies.

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Thanks to Minna Kääriäinen, MD, PhD Antti Mikkola, MD, Juha Kiiski, MD, PhD, Kristiina Hietanen, MD, PhD and Anne Koskela, MD for their amazing help in patient recruitment and follow-up. You went out of your way to make this study happen, thank you for this and for always being so supportive. Kristiina, you have been my thesis motivator, through the good times and the bad. You have also been a treasured colleague and unconditional friend and for this I am very grateful to you.

Thanks to Elina Nieminen, MD for helping organizing time off work to culminate this project.

A deep thank you to Hilkka Mäkinen, laboratory technician. So many hours together in the laboratory grew into a beautiful friendship that I will always cherish.

Thank you for your patience and guidance with all laboratory-related work.

A special thanks to Petri Välisuo for his valuable help in processing and measuring the spectrophotocutometric images.

Thanks to my family and friends for supporting me in this journey.

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ABSTRACT

With the number of acute and chronic wounds rapidly increasing worldwide, there is a crucial need for improved treatment methods. The mainstream treatment includes wound dressings, mechanical or chemical debridement and surgery.

However, to date, no single method has yielded optimal wound healing. Some products are chemically enhanced, but most are bioactively inert. Most tissue engineered treatments are based on cell therapy, but their methods of preparation are expensive, lengthy, require training and are not without potential adverse effects.

Wounds heal thanks to a cascade of orchestrated mechanisms resulting in cell proliferation, chemotaxis, migration, angiogenesis and extracellular matrix production. Key to wound healing, growth factors (GFs) are secreted by most cells within the zone of injury, i.e. platelets, endothelial cells, macrophages, lymphocytes, fibroblasts, adipocytes and epithelial cells. These proteins attach to membrane receptors and stimulate nuclear transcription, leading to wound closure. Attempts have been made to bioengineer single recombinant GFs for clinical use, but a combination of GFs is known to better mimic physiological healing. Bone marrow, blood and adipose tissue are known autologous sources of GFs. The use of platelet- rich plasma (PRP), has not gained much popularity due to varying clinical results in randomized controlled trials. Adipose tissue can easily be harvested and contains adipocytes and stromal vascular cells. Adipose tissue extract (ATE), is a cell-free, GF-rich secretome with proven adipogenic and angiogenic properties. The aim of the current study was to describe the effects of ATE within the wound healing setting.

The first study aimed at developing a new operating-room method to obtain ATE. Adipose tissue was obtained from the lipoaspirates of 27 healthy female donors and processed at different incubation time periods, temperatures, and employing different filter types. We compared the protein and GF values produced in the operating room to those in the laboratory. The second study focused on discovering the in vitro effects of ATE (13 samples) and PRP (11 samples) on the proliferation and migration of cells involved in wound healing: keratinocytes, fibroblasts, adipose stem cells and endothelial cells. We also measured the GF content of these samples. The third study was dedicated to unravelling ATE’s clinical

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effects. In patients with an indication for split-thickness skin graft reconstruction, two or more grafts were harvested. Autologous ATE or PRP was topically applied to one donor site while the other was left untreated. 15 ATE and 13 PRP-treated donor sites were studied. Wound re-epithelialization was evaluated with digital photography on postoperative days 5, 7, 10 and 15. Scars were measured using spectral imaging and scored with the Vancouver-Manchester modified scar scale on days 30 and 60.

In the first study we found that the ATE prepared in the operating theatre incubated for 30 min at room temperature and employing a 0.2 µm pore syringe filter recovered protein and GFs comparable to ATE produced in a laboratory setting.

Although PRP had overall higher GF measurements, we found that the ATE was a rich source of keratinocyte GF. This reflected positively in vitro (Study II) showing improved keratinocyte proliferation on culture day 6. The ATE also accelerated fibroblast and adipose stem cell migration. ATE and PRP promoted keratinocyte migration. In the clinical study (Study III), we found that ATE stimulated early re- epithelialization on days 5 (p 0.003) and 7 (p 0.04) while PRP promoted healing on day 7 (p 0.001), compared with controls. The ATE-treated sites demonstrated improved scar scores on days 30 and 60 (p 0.0005 and 0.02, respectively).

Angiogenesis and re-epithelialization are key events in wound healing. The current study encourages further investigation to better understand the mechanisms behind ATE’s healing potential. This study opens new doors in the area of tissue engineering and unveils important findings: an acellular extract is effective in concentrating the adequate combination of GFs to promote healing, and that wound repair does not require extremely high GF values to occur.

Key words: Adipose tissue extract, growth factors, wound healing, cell proliferation, cell migration, re-epithelization.

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TIIVISTELMÄ

Haavojen määrän kasvaessa nopeasti maailmanlaajuisesti tarvitaan parempia keinoja akuuttien ja kroonisten haavojen hoitamiseksi. Hoitoon kuuluvat keskeisesti haavanhoitotuotteet, mekaaninen tai kemiallinen puhdistus sekä kirurgia.

Kuitenkaan tähän mennessä mikään yksittäinen keino ei ole johtanut optimaaliseen haavan paranemiseen. Jotkut tuotteista toimivat kemiallisesti, mutta suurin osa ei reagoi bioaktiivisesti. Suurin osa kudosteknologiaan liittyvistä hoidoista perustuu soluterapiaan, mutta solujen hankkiminen vie aikaa, on kallista ja vaatii harjoittelua.

Hoitoihin saattaa liittyä myös haittavaikutuksia.

Haavan paraneminen on seurausta peräkkäisistä tapahtumista, jotka johtavat solujen jakautumiseen, viestintään, liikkumiseen, verisuonimuodostukseen ja solunulkoisen aineen tuottamiseen. Haavanparanemisen avainasemassa ovat kasvutekijät, joita erittyy suurimmasta osasta kudosvaurion alueella olevista soluista ja verihiutaleista kuten endoteelisoluista, makrofageista, lymfosyyteistä, fibroblasteista, rasvasoluista ja epiteelisoluista. Nämä valkuaisaineet kiinnittyvät solukalvon reseptoreihin ja stimuloivat proteiinisynteesiä johtaen haavan sulkeutumiseen. Yksittäisiä rekombinanttikasvutekijöitä on yritetty valmistaa kliiniseen käyttöön, mutta kasvutekijöiden yhdistelmän tiedetään paremmin matkivan fysiologista paranemista. Elimistön omia kasvutekijöiden lähteitä ovat luuydin, veri ja rasvakudos. Rasvakudos sisältää rasvasoluja ja strooman verisuonisoluja, joita voidaan kerätä helposti. Verihiutalerikkaan plasman (platelet- rich plasma, PRP) käyttö ei ole yleistynyt johtuen vaihtelevista kliinisistä tuloksista satunnaistetuissa kontrolloiduissa tutkimuksissa. Rasvakudosuute on soluton runsaasti kasvutekijöitä sisältävä seos, jolla on todistetusti rasvasolujen ja verisuonien muodostumista lisääviä ominaisuuksia. Tämän tutkimuksen tarkoitus oli kuvailla rasvakudosuutteen vaikutuksia haavan paranemiseen.

Ensimmäisessä tutkimuksessa pyrittiin löytämään uusi leikkaussaliin sopiva keino rasvakudosuutteen keräämiseen. Rasvakudosta kerättiin vesisuihkuavusteisella rasvaimulla 27 terveeltä naiselta. Valmisteluun liittyviä muuttujia olivat inkubaatioaika (15-45 minuuttia), lämpötila ja erilaiset suodatintyypit. Leikkaussalissa tuotettuja proteiini- ja kasvutekijäarvoja verrattiin laboratoriossa tuotettuihin.

Toisessa tutkimuksessa keskityttiin havaitsemaan rasvakudosuutteen (13 näytettä) ja

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PRP:n (11 näytettä) in vitro -vaikutuksia haavan paranemiseen osallistuvien solujen, kuten keratinosyyttien, fibroblastien, rasvakudoksen kantasolujen ja endoteelisolujen jakautumiseen ja liikkumiseen. Rasvakudosuute- ja PRP-näytteistä mitattiin myös kasvutekijäpitoisuuksia. Kolmannessa tutkimuksessa eriteltiin rasvakudosuutteen vaikutuksia ihmisillä. Ihosiirreleikkaukseen tulevilla potilailla otettiin kaksi tai useampia ihosiirteitä, joista yhteen ihosiirteen ottokohtaan laitettiin pinnallisesti joko omaa rasvakudosuutetta tai PRP:ta, ja toiseen satunnaisesti valittuun kohtaan asetettiin pelkkä haavasidos. Tutkimuksessa seurattiin 15 rasvakudosuutteella ja 13 PRP:llä hoidettua ihonottokohtaa. Haavan re-epitelisaatiota arvioitiin digitaalisilla valokuvilla 5., 7., 10. ja 15. leikkauksen jälkeisenä päivänä. Arvet mitattiin spektrocutometrialla ja pisteytettiin modifioidulla Vancouver-Manchester- arpiasteikolla 30. ja 60. päivänä.

Tutkimuksessa havaittiin, että leikkaussalissa valmistettu rasvakudosuute, jota inkuboitiin 30 minuuttia huoneenlämmössä ja jonka valmistuksessa käytettiin 0,2 µm reiällistä ruiskusuodatinta, sisälsi yhtä paljon proteiineja ja kasvutekijöitä laboratoriossa valmistettuun rasvakudosuutteeseen verrattuna. Vaikka PRP:ssä oli kaiken kaikkiaan enemmän kasvutekijöitä, tutkimuksessa havaittiin, että rasvakudosuute oli hyvä keratinosyyttikasvutekijöiden lähde. Tällä oli positiivinen vaikutus keratinosyyttienjakautumiseen kuudentena viljelypäivänä. Rasvakudosuute myös kiihdytti fibroblastien ja rasvan kantasolujen liikkumista. Rasvakudosuute ja PRP lisäsivät keratinosyyttien liikkumista. Kuten edellä, rasvakudosuutteen havaittiin stimuloivan varhaista re-epitelisaatiota 5. (p 0,003) ja 7. (p 0,04) päivänä, kun taas PRP:n havaittiin edistävän paranemista 7. (p 0,001) päivänä kontrolleihin verrattuna.

Rasvakudosuutteella hoidetuilla ihonottokohdilla oli paremmat arpipisteet 30. (p 0,0005) ja 60. päivänä (p 0,02).

Verisuonimuodostus ja re-epitelisaatio ovat keskeisiä tapahtumia haavan paranemisessa. Tämä tutkimus puoltaa lisätutkimuksia rasvakudosuutteen parantavan vaikutuksen taustalla olevien mekanismien ymmärtämiseksi paremmin.

Tutkimus avaa uusia ovia kudosteknologian alalla ja paljastaa tärkeät löydökset:

soluton uute on tehokas konsentroimaan riittävän ja sopivan kasvutekijöiden yhdistelmän paranemisen edistämiseksi ja haavan paraneminen ei vaadi erityisen korkeita kasvutekijäpitoisuuksia.

Avainsanat: rasvakudosuute, kasvutekijät, haavan paraneminen, solujakautuminen, solujen liikkuminen, re-epitelisaatio.

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Figure 7. Adipose tissue extract (ATE) and platelet-rich plasma (PRP) application on donor sites. PRP gel clot and remaining liquid spread over the donor site (left). The intradermal injections of ATE initially performed (center). Separation of experimental and control donor sites with hydrocolloid dressing followed by coverage with transparent semi-occlusive film (right).

Figure 8. Image J wound processing. A) Original image (left), B) Image filter (center), C) Selection of the non-epithelialized areas within the donor site wound (right).

Figure 9. Spectrophotometry. Original images (left) and the masking performed around the donor site area along with a circle of normal surrounding skin (right) for analysis.

Figure 10. Adipose tissue extract (ATE) protein content within the study timepoints.

Figure 11. Protein concentrations of the ATE samples within the different studies.

1-OR: study I, operating-room samples, 1-Lab: study I, laboratory samples, 2-ATE:

study II ATE samples.

Figure 12. Growth factor (GF) concentrations within the Adipose tissue extract (ATE) and platelet-rich plasma (PRP) samples (Study II).

Figure 13. Growth factors (GFs) recovered from the different study arms.

Figure 14. The effect of donor characteristics on GF concentrations of Adipose tissue extract (ATE) samples (Study II).

Figure 15. The influence of Adipose tissue extract (ATE) and platelet-rich plasma (PRP) on cell proliferation.

Figure 16. The influence of 10 and 20% platelet-rich plasma (PRP) on cell proliferation.

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Figure 17. Percentage of wound closure after performing scratch assays on studied cell lines.

Figure 18. Microscopy digital images of scratch wound assays.

Figure 19. The release of Adipose tissue extract (ATE) proteins from common wound dressings, fibers and gels.

Figure 20. Image progression of donor site wound healing after adipose tissue extract (ATE) and platelet-rich plasma (PRP) application. Above ATE-treated donor sites at post-op days 5, 10 and 14, and 2 other sites at days 3, 5, 10, 14 and 30. Below PRP-treated sites on days 5, 7, 10, 14 and 30. (A) Experimental donor sites (B) Homologous control donor site.

Figure 21. Re-epithelialization of ATE- (above) and PRP- (below) treated donor sites and their homologous controls. ATE significantly stimulated wound closure on days 5 and 10 compared with controls (p 0.003 and 0.04, respectively). PRP significantly stimulated wound closure on day 7 compared with controls (p 0.001).

Figure 22. Comparison of platelet-rich plasma (PRP) and adipose tissue extract (ATE) -treated donor site re-epithelialization.

Figure 23. Donor site fluid sample GF measurements from ATE-treated sites and their homologous control sites.

Figure 24. Spectral imaging analysis of hemoglobin and melanin content in ATE and PRP-treated donor sites, expressed in estimated concentration changes (ECC). Hbo (oxygenated hemoglobin), Hbdo (deoxygenated Hb), Hb (total Hb), Fmel (melanin).

ATE-treated donor sites demonstrated lower oxygenated hemoglobin values compared with their controls (p 0.05).

Figure 25. Adipose tissue extract (ATE) (left) and platelet-rich plasma (PRP) (right) scars on days 30 and 60. A) ATE day 30 (left) and day 60 (right). B) Control day 30 (left) and control day 60 (right). Note the highly vascularized scar begins to fade due to dermal epidermal thickening and vascular regression.

Figure 26. The relationship between Adipose tissue extract (ATE) GF measurements and donor site wound healing. The ‘control’ columns describe the ATE donor sites that showed a similar healing pattern to control donor sites

Figure 27. Correlation between in vitro proliferation and patient healing after ATE treatment.

Figure 28. Percentage of gap closure (cell migration) and clinical wound closure after ATE treatment.

Figure 29. Percentage of gap closure (cell migration) and in vivo wound closure after platelet-rich plasma (PRP) treatment.

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List of Tables

Table 1. Growth factors and their cellular effects.

Table 2. Growth factors and cytokines secreted by adipose tissue and their function.

Table 3. Growth factors employed in wound healing.

Table 4. Frequently employed scales for scar assessment.

Table 5. Patient demographics

Table 6. The Vancouver-Manchester modified scar scale.

Table 7. Adipose tissue extract (ATE) growth factor values in the operating room (OR) and laboratory (LAB) settings represented as mean (SD)

Table 8. Adipose tissue extract (ATE) growth factor measurements represented as mean values (SD).

Table 9. Platelet-rich plasma (PRP) growth factor comparison.

Table 10. Adipose tissue extract (ATE) and adipose liquid extract (ALE) protein (mg/ml) and GF (ng/ml) concentration represented as mean (SD).

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ABBREVIATIONS

ADSC adipose derived stem cells

Ang-1 angiopoetin 1

Ang-2 angiopoetin 2

ANOVA analysis of variance

ALE adipose liquid extract ATE adipose tissue extract

BJ human foreskin fibroblasts

BMI body mass index

BMP-2 bone morphogenetic protein 2 BMP-4 bone morphogenetic protein 4

BSA bovine serum albumin

CTGF/CCN2 connective tissue growth factor

DFU Diabetic foot ulcer

DMEM Dulbecco’s Modified Eagle’s Medium

DMEM/F12 Dulbecco’s Modified Eagle’s Medium: nutrient mixture F12 EBM-2 endothelial cell basal medium-2

ECCs estimated concentration changes

ECM extracellular matrix

EGF epidermal growth factor

EGM-2 endothelial cell growth medium -2 ELISA enzyme-linked immune absorbent assay

FBS fetal bovine serum

FGF fibroblast growth factor bFGF basic fibroblast growth factor

G-CSF granulocyte colony stimulating factor

GF growth factor

GM-CSF granulocyte macrophage colony stimulating factor

GTP guanosine triphosphate

HaCaT immortalized human keratinocyte cell line hASC human adipose stromal cells

Hb total hemoglobin (Hb)

Hbo oxygenated hemoglobin

Hbdo deoxygenated Hb

HGF hepatocyte growth factor

HIF-1 hypoxia inducible factor-1

Hr hour(s)

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HUVEC human umbilical vein endothelial cells ICAM-1 intercellular adhesion molecule-1

IFNγ interferon gamma

IGF-1 insuline-like growth factor-1

IL interleukin

Jak/STAT Janus Kinase/Signal Transducer and Activator of Transcription

KGF keratinocyte growth factor LEDs light-emitting diodes

MCP-1 monocyte chemoattractant protein-1

Mel melanin

MIF macrophage migration inhibitory factor

Min minutes

MIP-1 macrophage inflammatory protein-1

MMP matrix metalloproteinase

MSP macrophage stimulating protein

MSS Manchester scar scale

NGF nerve growth factor

NT neurotrophin

PAI-1 plasmin activator inhibitor 1 PBS phosphate buffered saline PDAF platelet derived angiogenic factor PDEGF platelet derived endothelial GF PDGFb platelet-derived growth factor BB PES polyethylene sulfone

PGC-1 proliferator-activated receptor gamma coactivator POSAS patient and observer scar assessment scale

PPP platelet-poor plasma

PRP platelet-rich plasma

RANTES regulated upon activation T-cell expressed and secreted CCL5(CC chemokine ligand 5)

RAS renin-angiotensin system RCT randomized control trial

RT room temperature

SBSES Stony Brook scale

SDF-1 stromal derived factor -1

SREBP1 sterol regulatory element-binding protein 1 STSG split-thickness skin graft

SVF stromal vascular fraction

TGFb tumor growth factor beta

Tie tyrosine kinase with Ig and epidermal growth factor homology domain

TIMP tissue inhibitor of metalloproteinase

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TNFα tumor necrosis factor alfa

Tsp thrombospondin

VCAM-1 vascular cell adhesion protein 1 VEGF vascular endothelial growth factor VMM Vancouver Manchester Modified Scale

VSS Vancouver Scar Scale

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ORIGINAL PUBLICATIONS

Publication I Lopez, J., Huttala, O., Sarkanen, J.R., Kaartinen, I., Kuokkanen, H.

and Ylikomi, T., 2016. Cytokine-rich adipose tissue extract production from water-assisted lipoaspirate: methodology for clinical use. BioResearch Open Access, 5(1), pp.269-278.

Publication II López, J.F., Sarkanen, J.R., Huttala, O., Kaartinen, I.S., Kuokkanen, H.O. and Ylikomi, T., 2018. Adipose tissue extract shows potential for wound healing: in vitro proliferation and migration of cell types contributing to wound healing in the presence of adipose tissue preparation and platelet rich plasma. Cytotechnology, 70(4), pp.1193-1204.

Publication III Lopez, J., Mikkola, A., Sarkanen, J.R., Kaartinen, I., Kuokkanen, H.

and Ylikomi, T., 2019. Adipose tissue extract is an alternative source of growth factors with the potential to promote wound healing: a human study of split- thickness skin graft donor sites. Journal of Wound Care (accepted for publication).

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1 INTRODUCTION

There is an increasing need for tissue engineered products in the area of wound healing. Globally, it is estimated that 305 million acute wounds are treated in an emergency setting every year (Global Wound Dressings Market 2018-2022 report;

World Cancer Fact Sheet, January 2014). Chronic non-healing wounds have a prevalence of 1-2% in developed countries (Heyer et al. 2016, Guest et al. 2015).

Proportional to the rise in diabetic, obese, vasculopathic and aging populations, ulcers are expected to increase by 2% annually (Chronic Wound Epidemic, 2016).

Worldwide, wounds are responsible for 1-3% of a nation’s health expenditure, not only due to wound and health-related costs, but also because of prolonged absences from work and psychosocial issues (Nussbaum et al. 2018, Järbrink et al. 2017). In the USA, chronic wounds affect 6.5 million people and costs exceed USA $25 billion annually (Sen et al. 2009, Chandan 2019). In the UK, they have a 6% prevalence and an expenditure of approximately 5.5% (Phillips et al. 2016), while in Scandinavia, this expenditure is 2–4% (Sen et al. 2009). In Finland, 1.3-3.6% of the population will develop a lower limb chronic wound in their lifetime and approximately Euro 5000-7000 will be spent on each patient per year (Coco and Okker-Tikkunen 2016).

Wounds pose a challenge to the treating clinician, demanding multidisciplinary care for lengthy periods and potentially culminating in radical surgery, like extremity amputation (Järbrink et al. 2017). This represents an economic and social burden where the quality of life is hindered. The causes of wound treatment failure are multifactorial and no single wound care protocol has proven to be superior (Guest et al. 2017, Dangwal et al. 2015).

Led by North America and Europe, the global wound healing market is predicted to grow at a rate of 4.8% annually from 2017 to 2025 and will surpass USA

$15-20 billion by the year 2022 (Wound Healing Market, 2017-2025). Therapeutic strategies include the use of ointments, gels, dressings, negative-pressure therapy, surgical debridement and reconstruction. The ideal treatment should comprise optimal chemical and physical properties, accelerate closure, replace soft tissue

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volume, and reduce pain, whilst being accessible and inexpensive (Rezvani Ghomi et al. 2019, Amato et al. 2015). However, no single product can offer all these characteristics (Frykberg and Banks 2015, Hurlow et al. 2015). Considerable funding is being invested in new technologies targeted at wound repair (Zarrintaj et al. 2017, Jatoi et al. 2019, Qu et al. 2019, Jordan et al. 2018). Scars compose a different research area dedicated to improving scarring once wound closure is complete (Marshall et al. 2018). There is a need for a tissue engineered product able to promote healing by inducing accelerated re-epithelization, extracellular matrix formation and angiogenesis. This is why this topic has instigated research into the underlying pathological mechanisms and treatment of wounds (Duscher et al. 2016, Sen et al.

2009).

Wound healing is commonly divided into three phases: inflammation, proliferation and remodeling (Nelligan 2013). However, to summarize such a complex process into three separate phases represents a very simplistic view (Yamaguchi and Yoshikawa 2001). These events overlap in time, and cells are actively signaling each other through the expression of growth factors (GFs) to induce cell proliferation, differentiation, migration and angiogenesis. Because GFs play a physiological role in cell signaling for wound closure, attention has been placed on the use of two autologous sources, blood and adipose tissue (Lindley et al. 2016, Dinh et al. 2015, Han and Ceilley 2017, Minutti et al. 2017, Park et al. 2017, Etulain 2018, Park et al. 2017, Shingyochi et al. 2015). Platelet-rich plasma (PRP), a concentrate rich in GFs, has been studied in numerous areas of medicine for cutaneous, tendinous, bone and muscle repair and regeneration (Martinez-Zapata et al. 2016, Marx 2004, Lacci and Dardik 2010, Sanchez et al. 2003, Foster et al. 2009, Sommeling et al. 2013). However, there are conflicting views over the use of PRP, mainly due to the different methods of preparation, activation and application (Hussain et al. 2017, Wroblewski et al. 2010, de Vos et al. 2014). In contrast, very few studies have considered adipose tissue as an alternative source of GFs.

Adipose tissue was thought to be limited to energy storage functionality, but it constitutes a metabolically active system and a rich source of GFs with wound healing potential (Kershaw and Flier 2004, Fortuño et al. 2003, Trayhurn and Wood 2004, Nakagami et al. 2005, Lombardi et al. 2019, Fromer et al. 2018). Although stem cells have attracted most of the attention, this treatment involves high costs, burdensome preparation and antigenicity (Medvedev et al. 2010).

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This thesis consists of determining the effect adipose tissue extract (ATE) has on cell function in vitro and on human donor site wound healing, expanding on the research foundation laid down by Sarkanen et al. on ATE (Sarkanen et al. 2012 a,b,c). Up to this point, ATE has been the product of solid adipose tissue processing (mincing, centrifuging, incubating and filtering) to yield a cell-free, cytokine-rich bioactive substance. Through extensive in vitro and animal experimental studies, ATE was found to possess angiogenic and adipogenic properties. These studies were performed in a laboratory setting, so the next logical step was to describe an operating-room ready extraction method along with in vitro and clinical studies specifically directed at ATE’s wound healing potential.

Responding to the crucial need for a bioactive substance capable of enhancing wound healing through its GF content, ATE is an encouraging therapeutical option in tissue engineering. Its simple preparation and proven angiogenic properties make ATE an excellent product for accelerating wound healing and improving final scarring.

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2 LITERATURE REVIEW

2.1 Normal Wound Healing

2.1.1 Structure and function of the skin

Considered the largest organ in the body, the skin is the basis for complex dynamic processes leading to regeneration and repair following injury. Structurally, it is divided into three layers, epidermis, dermis and hypodermis, where different cells coexist harmoniously to protect against environmental noxa (Nelligan 2013). The epidermis is a stratified epithelium that encompasses five distinct layers of keratinocytes, the lowermost named the stratum basale, responsible for continuous skin renewal through its resident stem and basal cells. The epidermis is also composed of Merkel cells, melanocytes and Langerhans cells. The function of this layer is not only to provide a protective, semipermeable barrier but also contribute to the immune response (Langerhans cells), ultraviolet light absorption and skin pigmentation (melanocytes) and perception of light touch through mechanoreceptors (Merkel cells) (Soutor et al. 2013, Yousef et al. 2019). The dermis constitutes the basic substance providing a foundation for the epidermis and its many structures. The fibroblasts in this layer are in charge of producing extracellular matrix (ECM), collagen and elastin fibers. Dermal leukocytes actively phagocytoze debris and present antigens, while mast cells secrete heparin and histamine (Yanez et al. 2017). The dermis is also home to hair follicles, eccrine and apocrine glands (for temperature regulation) while specialized nerve receptors aid in light touch, pressure and vibration perception (Rittié, et al. 2013). Within the dermis lies a complex network of blood vessels and lymphatics that are crucial for oxygenation and nutrition. Finally, the hypodermis is mainly for energy storage, but also protects against trauma and cold temperatures (Wong, et al. 2016, Driskell et al. 2014).

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2.1.2 Wound healing

Wound healing is a beautifully orchestrated cascade of events often divided into three overlapping phases of inflammation, proliferation and remodeling to achieve the final goal of re-epithelialization and soft tissue replacement. Immediately after injury, vasoconstriction occurs and coagulation pathways are activated for hemostasis (Gonzalez, et al. 2016, Smith et al. 2015). Platelets play a vital role in this period, by adhering to each other and the endothelium to form the initial clot. In response to injury platelets liberate their alfa granules, secreting the GFs contained within them. GFs are milestone proteins signaling the up-regulation of crucial events such as cell chemotaxis, migration, proliferation and differentiation crucial for wound healing (Wang et al. 2018, Martins-Green et al. 2013). After a brief period of vasoconstriction, vasodilation occurs during the inflammatory phase. This welcomes the arrival of neutrophils, macrophages, monocytes and other immune cells in response to circulating cytokines (Strbo et al. 2014). The cellular and humoral immune responses rid the wound of pathogens (Ellis et al. 2018). During this period, complementary GFs are released from macrophages, endothelial cells or keratinocytes to stimulate migration and proliferation of endothelial cells, fibroblasts and keratinocytes (Park et al. 2017, Li et al. 2003, Ucuzian et al. 2010). In general, this phase completes itself in one week.

During the second wound healing phase (fibroproliferation) fibroblasts are protagonists for approximately two weeks and appear as early as 48 hr post injury.

The hemostatic clot is degraded and replaced by a new granulation matrix rich in GFs, integrins, collagen type III, elastin and glycosaminoglycans (Caley et al. 2015).

Fibroblasts also secrete matrix metalloproteinases (MMPs) and their inhibitors (TIMPs), which regulate ECM deposition and degradation (Rohani and Parks 2015).

In response to signaling GFs, angiogenesis is a key event for the newly formed tissue to survive and achieve closure (Tonnesen et al. 2000). New vessels are formed, which allows the entry of new GFs, oxygen and nutrients into the zone of injury. Finally, the process of re-epithelialization occurs within hours after injury (Fig 1).

Keratinocytes migrate from the wound edges and the adnexal structures, such as the pilosebaceous units (Rittié et al. 2013). Wound contraction contributes to scar formation by aid of specialized fibroblasts with contractile function (Nelligan 2013, Thorne 2007, Fahs et al. 2015, Mulholland et al. 2017).

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The final phase of wound repair, remodelation, can last up to 18-24 months post injury (Velnar et al. 2009). A stronger foundational extracellular matrix mainly composed of organized collagen type I is deposited, while the previous is eliminated (Nelligan, 2013). Eventually, the scar’s content of inflammatory cells, glycosaminoglycans, water and blood vessels decrease (Burge 2014). Scar tissue can form as a fine line or be widespread, atrophic, hypertrophic, keloid or contractured (Erickson and Echeverri 2018). Peak scar tensile strength is reached at 60 days and achieves 80% of its original strength (Rippa, et al. 2019, Neligan 2013). There is a continuous cycle of ECM degradation and renewal by collagenases, forming the mature scar.

Figure 1. Phases of wound healing. Hemostatic/inflammatory (above) and re- epithelialization phases (below) of wound healing after acute injury.

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2.1.3 Growth factors

The first ever described GF was discovered by Rita Levi-Montalcini and Stanley Cohen in 1952, which granted the former a Nobel Prize. Historically, GFs were defined as peptides (2-50 amino acid residues) or proteins (greater than 50 residues) that affect cell growth and survival. However, it is current knowledge that GFs are bioactive proteins or hormones that initiate complex cell signaling pathways to promote or inhibit specific cell functions (Stone and Varacallo 2019). Signaling mechanisms can occur between neighboring cells (paracrine/juxtacrine), distantly (endocrine) or a cell can signal itself (autocrine/intracrine) (Zarei and Soleimaninejad 2018).

Through the process of cell transduction, GFs bind to membrane receptors and transfer their signals to the intracellular cytoplasm, ultimately modifying gene expression to codify a specific function, like cell proliferation, migration or differentiation (Naderali et al. 2018). The exact series of events are unknown, but signaling begins when the extracellular protein binds to the ligand and activates transmembrane amino acids (Ullrich and Schlessinger 1990, Ganapathy et al. 2012).

Phosphorylation of amino acids by protein kinases or phosphatases transmits the messenger signals to trigger nuclear transcriptional factors (Ardito et al. 2017). For instance, in the case of the epidermal GF (EGF) receptor, phosphorylation activates several signaling pathways specific to a given function (Ganapathy et al. 2012). The AKT pathway signals for cell survival by activating specific phosphorylates, while the AKT, RAS and MEK pathways induce cell proliferation, migration and apoptosis. GFs can also stimulate the receptor-ligand cascade through guanosine triphosphate (GTP) binding proteins, which also elicit transduction (Ardito et al.

2017). A third proposed mechanism by which GFs activate receptors is through the action of proto-oncogenes and the transcription of genes like C-FOS and C-MYC (Ardito, et al. 2017). This regulation is complex, as multiple types of receptors can respond to a single GF to ‘up-’ or ‘down-’ regulate a specific cellular action.

Interestingly, various cells can secrete the same GF and these can have similar or distinct effects on other cell types (pleiotropism). On the other hand, different GFs may perform a similar target function (Babensee et al. 2000). Most GFs are not stored in latency, but quite the opposite and are briefly secreted and activated in response to injury and have momentary half-lives (Babensee et al. 2000). In wound healing, GFs are key in regulating cell division, chemotaxis, migration, adhesion, angiogenesis, apoptosis and ECM synthesis.

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Hundreds of GFs have been identified and classified according to different systems (Bafico and Aaronson 2003). In relation to their biochemical composition, GFs are divided into two broad groups, proteins and hormones (Babensee et al.

2000). Secreted by endocrine glands, estrogens, androgens, and progestogens are considered GF hormones because of their influence over cell growth and differentiation. Lipid-soluble steroid hormones can traverse the cell membrane, avoiding the first step of binding to the membrane receptor and, instead, bind to an intracellular one (Ardito et al. 2017). However, most GFs involved in wound healing such as vascular endothelial GF (VEGF), EGF and platelet-derived GF beta (PDGFb) are proteins (Bafico and Aaronson 2003). According to their receptor location, GFs have also been categorized into four main classes: on the cell surface (Class 1), cell surface hormone receptors (Class 2), intracellular signal transmitters (Class 3) or nuclear transcription factors (Class 4) (Ségaliny et al. 2015). GFs can also be grouped according to their function in inflammation, homeostasis, angiogenesis, amongst others (Rees et al. 2015).

The terms GF and cytokine are commonly used interchangeably by the scientific community. While the term “GF” includes proteins and hormones directly affecting cell differentiation and proliferation, cytokines are associated with the hematopoietic and immune systems (Atanasova, and Whitty 2012). The majority of cytokines are secreted by leukocytes and trigger humoral or cellular immune responses. Cytokines originating from monocytes or macrophages are termed “monokines” while those derived from lymphocytes are “lymphokines”. The latter are also known as interleukins (ILs) because they originate and have function over leukocytes (García Morán et al. 2013). Chemokines are cytokines that stimulate cell migration or chemotaxis. The addition of the “kine” suffix to cytokine, helps understand the origin story. In this sense, “adipokines” are cytokines secreted by the adipose tissue,

“hepatokines” by the liver, “myokines” by muscle, and so forth. However, these cytokines are not exclusively produced by these tissues. The most common classification divides GFs into 11 families according to their function and structure (Table 1).

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Table 1. Growth factors and their cellular effects.

Growth

factor Cell of origin Target cell Main function Author PDGF -AA, -

AB, -BB (platelet- derived GF)

Platelets, macrophages, endothelial cells, keratinocytes, fibroblasts, vascular smooth muscle cells, fibroblasts

Fibroblasts,

macrophages Chemoattracts and

activates macrophages

and fibroblasts,

fibroblast and

osteoblast proliferation and ECM production, collagenase production, angiogenesis

Uutela et al.

2004, Lederle et al. 2006.

EGF (Epidermal

GF)

Platelets, macrophages, keratinocytes, fibroblasts

Fibroblasts,

keratinocytes Collagenase

production, fibroblast

and keratinocyte

proliferation and differentiation,

granulation tissue

Shiraha et al.

1999, Schultz et al. 1991, Brown et al.

1988, Brown et al. 1986.

TGF -a,-b (tumor GF alfa, beta)

Platelets, macrophages, lymphocytes, keratinocytes, fibroblasts, osteocytes, hepatocytes

Keratinocytes, fibroblasts, endothelial cells, leukocytes

TGFa:

Chemoattractant, induces proliferation of keratinocytes and fibroblasts, granulation tissue, ECM production and angiogenesis TGF-b (1 and 2):

chemoattracts

leukocytes, angiogenic,

stimulation of

fibroblasts and

osteoblasts, ECM

production

TGF-b 3 (as opposed to TGF-b 1 and 2): scar- less wound healing (fetal)

Lee et al.

1997, Rolfe et al. 2007.

a, b FGF (fibroblast

GF a,b)

Macrophages, mast cells, lymphocytes, hypophysis, fibroblasts, endothelial cells, smooth muscle cells,

chondrocytes

Fibroblasts, endothelial cells, keratinocytes

Chemoattracts

endothelial cells and leucocytes, angiogenesis, granulation,

fibroblast and

keratinocyte proliferation

(epithelialization) and

migration, ECM

production, wound contraction

Powers et al.

2000, Ornitz 2000.

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IGF-1 (insulin-like

GF 1)

Platelets, Hepatocytes, fibroblasts

Fibroblasts Chemotaxis fibroblasts,

ECM production,

fibroblast proliferation, osteosynthesis

VEGF A-E (Vascular endothelial

GF A-E)

Endothelial cells Endothelial

cells Angiogenesis

(endothelial cell proliferation and migration, creation of

vessels lumen),

chemotaxis leukocytes, vasodilation

Suzuma et al.

1998, Pepper et al. 1992.

KGF (FGF- (Keratinocyte 7)

GF)

Fibroblasts Keratinocytes Keratinocyte

proliferation, migration, differentiation

Ceccarelli et al. 2007, Wu et al. 1996 TNFa

(Tumor necrosis factor alfa)

Macrophages, neutrophils, mast cells, eosinophils, and natural killer lymphocytes

Fibroblasts Fibroblast proliferation, macrophage-derived angiogenesis

Mast and

Schultz 1996.

IFNg (interferon

gamma)

Lymphocytes,

fibroblasts Fibroblasts Inhibits fibroblast

proliferation and ECM production

IL

(interleukins) Macrophages Fibroblasts, keratinocytes, leukocytes (pro-

inflammatory)

IL-1 Chemoattracts Fibroblasts,

keratinocytes and neutrophils, fibroblast proliferation

IL-4 Fibroblast

proliferation and differentiation, ECM production

IL-8 Chemoattracts

fibroblasts and

neutrophils

Gallucci et al.

2004, Sato et al. 1999.

CSGF (colony stimulating

GF)

Fibroblasts, lymphocytes, endothelial cells

Leukocytes G-CSF: granulocyte

proliferation

GM-CSF: granulocyte

and macrophage

proliferation

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2.1.4 Role of growth factors in wound healing

In wound healing, GFs are considered signaling focal points, particularly in the first two phases. In response to injury, interleukine-1 (IL-1) is the first GF to be released by local keratinocytes, warning the microenvironment of the skin barrier compromise (Zarei and Soleimaninejad 2018, Barrientos et al. 2008). The incoming platelets degranulate, releasing EGF, PDGFb, TGFb, bFGF, VEGF, IGF-1.

Chemokines such as IL-1, CD4, rantes, MIP-1, IL-8 are also secreted early on (Werner and Grose 2003). Thanks to the effects of PDGFb and IL-1, neutrophils are chemoattracted to the injury site and decontaminate the wound. TGFb actively converts peripheral monocytes into macrophages that also clear the wound of detritus by phagocytosis (Snyder et al. 2016). Within the injury microenvironment macrophages secrete b-FGF, EGF, TGFb, PDGFb and inflammatory cytokines.

Angiogenesis is the formation of new vessels from pre-existing vasculature and is classified as sprouting and intussusceptive (DiPietro 2016). In the first, new buds of endothelial cells are added to existing vessels in response to VEGF and bFGF, and secondarily to IGF-1, TGFb and IFNγ (Tonnesen et al. 2000, Mongiat et al. 2016, Khan et al. 2017). Intussusception occurs when stromal tissue invades and “splits” the existing vessels, duplicating the vessel.

Fibroblast proliferation and migration results in the creation of organized ECM, initiated by TGFb, bFGF and PDGFb. Re-epithelialization occurs by virtue of EGF, TGFa, bFGF, and KGF. Finally, tissue remodeling is mediated by TGFb and PDGFb (Barrientos et al. 2008, Grayson et al. 1993, Dvonch et al. 1992, Vogt et al.1998, Arnold et al. 1987, Grellner et al. 2000). As opposed to acute wounds, chronic wounds demonstrate a persistent inflammatory response and an uninhibited metalloproteinase action, where GFs are degraded and cannot promote healing (Landén, et al. 2016). In fetal wound healing, EGF, bFGF and PDGFb levels may be increased immediately after injury but have accelerated clearance. While IGF-1 is decreased during early healing, TGFb 3 is increased. In the remodelling phase, TGFb, PDGFb, and bFGF regulate ECM deposition and stimulate contraction.

TGFb also increases ECM deposition via collagenase activity. (Davis 2016)

Undoubtedly, successful wound healing demands the harmonious coordination and timely action of GFs (Kiritsy and Lynch 1993, Lynch et al.1987).

GFs originating from distant organs, also play a role in wound repair. For example, hepatocyte-derived angiopoietins, bFGF, IGF-1 and HGF have shown positive healing effects by inducing cell migration, proliferation, and angiogenesis while

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promoting ECM degradation and renewal (Werner and Grose 2003, Zuliani-Alvarez and Midwood 2015, Youssef et al. 2017, Werner 2011).

In wound healing, IL-1α, IL-1β, IL-6 and IL-8 mediate the early inflammatory response while IL-10 is present subsequently. TNFα takes part in the acute phase reaction of inflammatory responses, inducing fever, inhibiting viral replication and inducing apoptosis (Larouche et al. 2018, Ritsu et al. 2017).

Granulocyte-macrophage colony stimulating GF (GM-CSF) stimulates the recruitment of inflammatory cells, myofibroblast function and epidermal proliferation. Connective tissue GFs CCN2/CTGF induce cell proliferation, chemotaxis, angiogenesis, and deposition of ECM (Wang et al. 2003, Henshaw, et al. 2015). Bone morphogenetic proteins (BMP), particularly BMP-6, are secreted by keratinocytes and fibroblasts and regulate repair. Nerve GF (NGF) promotes nerve regeneration and stimulates keratinocyte proliferation and migration (Pincelli 2000.) Finally, leptin is believed to enhance cutaneous wound healing by stimulating keratinocyte proliferation and migration, and angiogenesis (Tadokoro et al. 2015).

Earlier it was thought that the clinical effect of GFs was directly proportional to their concentration, but we now know that this is a complex relationship that does not necessarily correlate. In vitro, higher concentrations of GFs decelerated cell proliferation, differentiation and migration (Kakudo et al. 2008, Graziani et al. 2006, Berndt et al. 2019, Lucarelli et al. 2003, Tavassoli-Hojjati et al. 2016). In addition, unregulated or unlimited GF production is seen in diseased states. For instance, VEGF is up-regulated in carcinogenic tumors, leading to angiogenesis, tumor growth and spread (Witsch et al. 2010, Kareva et al. 2016). Similarly, in the remodeling phase, an excess of GFs, particularly of TGFb, PDGFb and activin have been implicated in pathological scarring conditions, such as keloids and hypertrophic scars (Berman et al. 2017).

2.2 Sources of growth factors

Autologous sources of GFs originate in whole blood, bone marrow and adipose tissue. These seem to provide certain advantages by reducing antigenicity and costs, however, their preparation can be complicated and time-consuming.

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2.2.1 Platelet-rich plasma

Since the early 1970s platelet-rich plasma (PRP) has been investigated in vitro and in clinical settings. PRP is formed when a sample of anticoagulated blood is centrifuged to concentrate platelets from 1.5-12 times their baseline plasma values (Lacci and Dardik 2010, Foster et al. 2009). When PRP is activated, the platelets release their granule contents, resulting in a gel–like bioactive substance (Mussano et al. 2016, Fréchette et al. 2005, Everts et al. 2006). This gel is rich in a combination of concentrated GFs, mainly PDGF-B, FGF, TGFb, VEGF and IGF, known to stimulate cell division, migration, organization and hemostasis (Landesberger et al.

1998, Weibrich et al. 2004). The combination of biomarkers originating from PRP simulates platelet degranulation in an injury microenvironment, which is considered an advantage compared with the administration of single GF preparations.

PRP can be prepared in an artisanal fashion or by employing commercial kits, such as RegenPRP-Kit® (RegenLab, Mollens-VD, CH); Fibrinet® (Cascade Medical Enterprises LLC, Wayne, NJ, USA); Plateltex® (Bratislava, Slovakia), Symphony II® (Which Medical Device, USA), Vivostat System® (Vivostat A/S, Alleroed, Denmark), SmartPrep® (Terumo Harvest, USA), PRP GLO Pro kit (Glofinn Oy, Salo, Finland), GPS (Biomet Biologics, USA), Magellan (Arteriocyte Medical Systems, USA), and ACP (Device Technologies, Arthrex) (Middleton et al.

2012, Weibrich and Kleis 2002). After extraction, blood is centrifuged to form three layers, from bottom to top there is a red blood cell layer (rich in erythrocytes and white blood cells), a fine white buffy coat (rich in platelets and white blood cells) and a top plasma level (Fitzpatrick et al. 2017, Salamanna et al. 2015, Panda et al.

2012, Anitua et al. 2005). For PRP to be clinically effective, there is a consensus that platelets should exceed 1-2 x10⁹ platelets/ml (Leitner et al. 2006, Kakudo et al. 2009).

Nonetheless, studies have shown adequate clinical results with lower platelet counts, confirming that there is no direct correlation between platelet and GF content (Smith and Roukis 2009, Roberts and Sporn 1993, Marx 2001, El Bagdadi et al. 2017, Rutkowski et al. 2008, Zimmermann et al. 2008, Leitner et al. 2006). Activating PRP by adding thrombin, collagen and/or calcium (calcium chloride or calcium gluconate) recovers greater GFs and results in a gel that serves as a scaffold over which cell migration and wound closure can occur (Cavallo et al. 2016). Once activated, approximately 60-70% of GFs are released within 15 min and the rest within 2 hr of degranulation (Foster et al. 2009). Small quantities of GFs continue to be released from the platelets until their senescence (8-12 days) (Borzini and Mazzucco 2007). Activation is not essential because it naturally occurs when PRP is

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in contact with collagen, endothelial cells, and calcium components within a wound.

Inactivated PRP has the advantage of remaining in a liquid form that is easy to inject.

A major obstacle in PRP methodology is that there is no standardized form of preparation. Variations are seen in the amount of blood extracted, centrifugation settings, number of spin cycles, whether PRP activation is performed or not and the type of activating agent used. These will influence the final volume of PRP, platelet and GF yield, as do intrinsic factors like age, gender, BMI, comorbidities, ethnicity and habits (Mazzucco et al. 2009, Salamanna et al. 2015, Kushida et al. 2014). Even so, among similar patients, GF yield may significantly vary (Kuffler 2018, Salamanna et al. 2015, Weibrich and Kleis 2002).

Opponents to the use of PRP suggest that there is a wide margin of GF variation, and that overexposure could result in continuous activation of different cell lines that may not differentiate properly, potentially promoting carcinogenesis (Hussain et al. 2017, Kia et al. 2018, McNamee et al. 2018, Andrade et al. 2017).

Neoplastic tissues overexpress a number of GFs that happen to be present in PRP.

Nonetheless, no cases of this phenomenon have been reported. PRP GFs have a short half-life (hours to days) and act locally; it would take extremely high doses with continuous administration for oncogenesis to occur. (Martinez-Gonzalez et al.

2002). Conventional PRP GF concentrations are considerably low (Marx 2001, Schmitz and Hollinger 2001, Akhundov et al. 2012).

Adverse side effects after PRP use are local pain, erythema mild swelling and infection. A single case of serious coagulopathy mediated by antibodies has been reported after activating PRP with bovine thrombin (Salamanna et al. 2015). Also, a case of severe allergy has recently been described (Latalski et al. 2019). Previously it was believed that, because GF effect could not be regulated, patients with cancer should be excluded from PRP use to avoid carcinogenesis. Nonetheless, it is now believed that PRP has a very limited local effect and that activated GFs have such a short half-life that this risk is low (Vilar et al. 2017).

2.2.1.1 In vitro effects of platelet-rich plasma

Roubelakis et al. studied the effects of PRP on dermal fibroblast and mesenchymal stem cell proliferation and migration. They discovered that PRP enhanced cell proliferation and migration (Roubelakis et al. 2014). Another study found that PRP- treated fibroblasts only exhibited improved proliferation rates in short-term cultures while long term cultures showed greater vimentin, fibronectin and actin, which

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suggests that PRP facilitates the conversion of fibroblasts to myofibroblasts in the scar remodelling phase (Ramos-Torrecillas et al. 2014).

A study on cell proliferation of human adipose stem cells and fibroblasts with 1%, 5%, 10% and 20% PRP demonstrated that 5% PRP showed improved proliferation, while 20% decelerated it (Kakudo et al. 2008). Graziani et al. studied the in vitro effects of 2.5, 3.5 and 5.5x PRP in osteoblast and fibroblast cell cultures.

The greatest proliferation was achieved at a 2.5x concentration while higher PRP concentrations decreased it (Graziani et al. 2006). Repeatedly, studies have found the same phenomenon after PRP treatment (Lucarelli et al. 2003, Tavassoli-Hojjati et al. 2016). This dose-dependent effect seems to follow a bell curve up to a point where higher PRP concentrations no longer stimulate proliferation (Berndt et al.

2019). In a study by Xian et al., human fibroblast and keratinocyte cells from healthy donors were seeded in separate cell culture inserts combined with 5, 10 and 20%

PRP followed by proliferation and scratch assays. Higher concentrations of PRP seemed detrimental to keratinocyte proliferation but promoted inflammation and collagen storage. On the other hand, 10% PRP stimulated keratinocyte migration.

Nonetheless, PRP seemed to decelerate fibroblast migration (Xian et al. 2015). Other studies, however, support the wound healing effects of PRP in the migration of different cell lines (Roubelakis et al. 2014, Ranzato et al. 2008, Ranzato et al. 2009, Ranzato et al. 2009).

2.2.1.2 Clinical effects of platelet-rich plasma

Although PRP is gathering increasing attention, there is insufficient scientific evidence to support its use in clinical practice. Many PRP trials are isolated, poorly designed, biased or underpowered, thus rendering low-quality evidence. In the field of orthopaedics, for example, PRP has been employed for the treatment of acute and chronic tendinous, osseous, cartilaginous, muscular and ligamentous injuries (Griffin et al. 2012, Patel et al. 2013, Gosens et al. 2011, Moraes et al. 2014, Foster et al. 2009). However, evidence to support its standardized use in this field is lacking (Martinez-Zapata et al. 2016, Nourissat et al. 2013, Le et al. 2018, Sheth et al. 2018, de Vos RJ et al. 2010, Schepull et al. 2011, de Jonge et al. 2011). Other surgical fields such as maxillofacial surgery (Stähli et al. 2018, Franchini et al. 2019, Albanese et al.

2013, Anitua et al. 2015, Del Fabbro, et al. 2018), cardiothoracic and vascular surgery (Patel et al. 2016, Kirmani et al. 2017), ophthalmology (Alio et al. 2017), spinal/neurosurgery (Akeda et al. 2019, Teymur et al. 2017, Mohammed and Yu 2018), gynecology and urology (Dawood and Salem 2018, Epifanova et al. 2020)

Adipose Tissue Extract for Wound Healing : Operating room preparation, in vitro and clinical use

Adipose Tissue Extract for Wound Healing

Operating room preparation, in vitro and clinical use

JENNY LOPEZ

JENNY LOPEZ

Adipose Tissue Extract for Wound Healing

Operating room preparation, in vitro and clinical use

ACKNOWLEDGEMENTS

ABSTRACT

TIIVISTELMÄ

CONTENTS

ABBREVIATIONS

ORIGINAL PUBLICATIONS

1 INTRODUCTION

2 LITERATURE REVIEW

2.1 Normal Wound Healing

2.2 Sources of growth factors