Cellular and molecular mechanisms of human aortic valve calcification

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Cellular and molecular mechanisms of human aortic valve calcification

Thesis for the degree of Philosophiae Doctor (PhD) Maria Bogdanova

Division of Physiology

Department of Molecular Medicine Institute of Basic Medical Sciences

Faculty of Medicine University of Oslo

2019

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© Maria Bogdanova, 2019

Series of dissertations submitted to the Faculty of Medicine, University of Oslo

ISBN 978-82-8377-514-3

reproduced or transmitted, in any form or by any means, without permission.

Cover: Hanne Baadsgaard Utigard.

Print production: Reprosentralen, University of Oslo.

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Acknowledgements

I would like to start by thanking professor Jarle Vaage who gave me opportunity to do my phD project at University of Oslo. When I just came to study and work in Oslo from Russia I felt unconfident in my skills and knowledge, I hardly even spoke English. But you trained me again and again to feel confident to give a presentation on public, to improve tremendously my writing skills and to develop scientific way of thinking. Working with you as a mentor I got unavailable experience which I am sure will make huge impact on my future life and career. I will never forget your wise advices, long adventure stories and jokes. Thank you for sharing with me your kindness, optimism in any situation and wisdom.

I am very grateful to my principal supervisor Arkady Rutkovskiy for everything what he did for me from the beginning of my life in Oslo. Arkady, you are smart and very ambitious, but also the person who is talented to behave with people on an equal footing likes a brother.

Please never change! I believe that it is an exceptional case to have a supervisor with whom I could discuss literally everything from professional problems to deeply personal. A year after we started to work together, you got medical doctor position and left our group. We did not see each other every day at work, but you were always available for me by phone. Thank you for all those long conversations, for listening me and understanding. Few times you pulled me out from the state of despair and loss of faith in myself and my phD project. You helped me to gain self-confidence and made me believe in the fact that I can achieve much more then I think. I sincerely consider myself lucky that you are my supervisor and a good friend.

No words can truly express how grateful I am to my co-supervisor Anna Malashicheva who is also supervised me during my Master and Bachelor program at the Saint-Petersburg State University and in Almazov Federal Medical Research Centre in Saint-Petersburg, Russia. It was you who introduced me first time to the heart physiology group at University of Oslo Anna, you are example of heroic women in science for me. I could never understand how you manage to take care of three kids, to work in science supervising few students and to do many additional things. I am impressed and inspired by your strong character and faith in the best despite the family tragedy and all difficulties you faced. Thank you for sharing with me your professional and personal experience, for your belief in me and for everything what you did for me.

I would like to thank the leader of heart physiology group and my co-supervisor, professor Kåre-Olav Stensløkken, who always with rationalism and justice helped me to resolve issues related to my phD project. Thank you for giving me the words of support in right moment, reassuring me that I will finally get my phD degree, that meant a lot for me. I am also grateful for your help to decide my administrative questions and for giving me the opportunity to travel and present my project at many national and international conferences. The most my brightest memory is our trip to the conference in Argentina, so much fun and impressions!

And I will always remember all your rational life advises during lunch conversations and skiing lessons.

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8 My research group, I am so happy and proud that I worked with all of you. I will miss you a lot and I hope we will always keep in touch.

Arsenii Zabirnyk, I know you for long time since we work together in Almazov Federal Medical Research Centre in Saint-Petersburg. This time in Oslo as previous one we were

“partners in crime” sharing the same project. Thank you for teaching me to different methods of cellular and molecular biology and for your scientific approach to clearly understand the essence of every method as well as for new ideas and help with experiments that significantly improved my PhD thesis. And thank you for your friendship and support.

May-Kristin Torp, we came through almost all way of phD life together and we shared the same office from the beginning. I could hardly imagine who else if not you could listen and answer me on all my questions every day with such patience. What would I do without you?

Thank you also for organizing “fun” in our lab and for your delicious cakes.

Chrisina Heiestad, I am happy that you joined our group few years ago. You are smart, fun and interesting person and I always like to talk to you. Thank you for your psychological support, especially on that conference in Geilo where we shared the same room. I believe that your phD will be finished successfully as well.

Torun Flatebø, thank you for all your help in the laboratory and thank you that you always asked how I feel and how I am doing.

I would also like to thank to my previous colleague Katharina Zihlavnikova Enayati, we started to work on this project together and she made significant contribution to it, especially in establishing the methods. We presented our first results together on ISHR conference in Bordeaux, France in 2015.

I am very grateful to professor Gareth Sullivan who was listening to my ideas, discussing all my methodological problems during my phD project, revising my papers and helping me to answer on the question of reviewers. Thank you, Gareth, also for giving me curiosity and inspiration and for you career advices.

I thank all my co-authors who kindly have contributed to my phD project. Special thanks to professor Arnt Fiane, doctor Mari-Liis Kaljusto and doctor John-Peder Escobar Kvitting for helping me to harvest aortic valves from patients, without your help this project will be impossible to perform.

I am thankful to my colleagues who was part of my research group in the beginning of my phD life and helped me to adapt to the new surroundings and to feel welcome in the lab: Lars Henrik Mariero, Anton Baysa, Mark Scott and Yuchuan Li. Also thanks to the colleagues from neighboring lab for nice company and chatting during lunch time and rest breaks at work:

Mari Falck, Elke Maes, Maria Skytioti, professor Maja Elstad and to phD student from Spain Patricia Sanchez who was visiting our lab last year.

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9 I am grateful to executive officer Gjøril Seierstad who was listening to all my stories about

“what is happening in my phD life” while I stopped by administration desk on my way to cafeteria, and of course for helping me to solve all administrative problems.

I would like to acknowledge Anna Kostareva, head of the Institute of Molecular Biology and Genetics in Almazov Federal Medical Research Centre in Saint-Petersburg, Russia where I worked before and all members of great research team at the Institute that you did not forget about me when I moved to Oslo and keep interested in my scientific life, sending me supportive and inspiring mails from time to time. Special thanks to Aleksandra Kostina and Daria Semenova who are also working on the project related to aortic valve calcification and shared their results and experience during our common meetings in Oslo and in Saint- Petersburg.

I would like to thank my family for care, warmth and support: to my mom who is always worried about me, to my dad who always believe in me and reassures my mom, to my sister Olga and to my cousin Julia, who are far away but still stay close. I am thankful to my friends with whom I could emotionally detach from my work after long days in the lab. Finally, I thank my French fiancé Hugues, whom I met in the middle of my phD life but who gave me incredible support in the most difficult period. Thank you, that you flew from France to Norway every weekend during whole year to see me, that you forced me sleep when I really needed it, that you cooked a lot of food for me that I could stay in the lab long hours without being hungry, that you dealt with all my tantrums and all my mood swings.

The phD program gave me incredible experience of self-growth and self-discovery. There was everything: ups and downs, happy days and moments of despair, laugh and tears. But I am happy that I went throw it and met many great people on my way. Now I am ready to turn the page to the next chapter of my life.

Maria Bogdanova 07.04.2019

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List of included articles:

Article I: Methods for Isolation, Characterization and Investigation of Human Valvular Interstitial and Endothelial Cells

M Bogdanova, K Zihlavnikova Enayati, A Malashicheva, A Kostareva, J Vaage, K Larsen Sand, A Zabirnyk, K-O Stensløkken, A Fiane, J-Ø Moskaug, R Siller, G Sullivan*, A Rutkovskiy*. To be submitted.

Article II: Inflammation and Mechanical Stress Stimulate Osteogenic Differentiation of Human Aortic Valve Interstitial Cells

M Bogdanova, A Kostina, K Zihlavnikova Enayati, A Zabirnyk, A Malashicheva, K-O Stensløkken, G J Sullivan, M-L Kaljusto, J-P E Kvitting, A Kostareva, J Vaage*, A Rutkovskiy*. Frontiers in Physiology. 9, 1635, doi:10.3389/fphys.2018.01635 (2018).

Article III: Interstitial Cells in Calcified Aortic Valves Have Reduced Differentiation Potential and Stem Cell-Like Properties

M Bogdanova, A Zabirnyk, A Malashicheva, K Zihlavnikova Enayati, T A Karlsen, M-L Kaljusto, J-P E Kvitting, E Dissen, G J Sullivan, A Kostareva, K-O Stensløkken, A Rutkovskiy*, J Vaage*. Scientific Reports. 2019. doi:10.1038/s41598-019-49016-0.

Article IV: SNF472, a Novel Anti-Crystallization Agent, Inhibits Induced Calcification in an In Vitro Model of Human Aortic Valve Calcification

A Zabirnyk, M D Ferrer, M Bogdanova, M M Pérez, C Salcedo³, M-L Kaljusto, J-P E Kvitting, K-O Stensløkken, J. Perello, J Vaage. Vascular Pharmacology. 2019.

doi:10.1016/j.vph.2019.106583.

* These authors have a shared last authorship

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Abbreviations

ACAN – aggrecan ACTA2 – actin alpha 2

ARBs – angiotensin II-receptor blockers AVR – aortic valve replacement

BAV – bicuspid aortic valve

BMPs – bone morphogenetic proteins BMP2 – bone morphogenetic protein 2 CAVD - calcific aortic valve disease

CEBPA – CCAAT enhancer binding protein alpha CFD – complement factor D

CNN1 – calponin 1

COL2A1 – collagen type II alpha 1 chain CTNNB1 – catenin beta 1

DAMPs – damage-associated molecule patterns DMEM – Dubecco’s modified Eagle’s medium ECM – extracellular matrix

EMT – endothelial to mesenchymal transition FBS – fetal bovine serum

HMBG1 – high mobility group protein B1 ICAM1 – intracellular adhesion molecule 1 IL-1β – interleukin-1 beta

IL-6 – interleukin-6

LDL – low density lipoproteins LPL – lipoprotein lipase

LPS – lipopolysaccharide

MACS – magnetic-activated cell sorting MGP – matrix Gla protein

MMPs – matrix metalloproteinases mRNA – messenger RNA

NF-kB – nuclear factor-kappa B NO – nitric oxide

OPG – osteoprotegerin

OxLDL – oxidized low density lipoproteins OxPL – oxidized phospholipids

PAMPs – pathogen-associated molecule patterns PBS – phosphate buffered saline

PECAM1 – platelet endothelial cell adhesion molecule 1 (CD31) POSTN – periostin

PPARG – peroxisome proliferator-activated receptor gamma RANKL – receptor activator of nuclear factor kappa-B ligand RNA – ribonucleic acid

RT-qPCR – quantitative reverse transcription polymerase chain reaction RUNX2 – runt-related transcription factor 2

SMC – smooth muscle cells TAGLN – transgelin (SM22) TAV– tricuspid aortic valve

TAVR – transcatheter aortic valve replacement TGF-β1 – transforming growth factor beta-1

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14 THBS1 – thrombospondin 1

TIMPs – tissue inhibitor metalloproteinases TLRs – toll-like receptors

TNF-α – tumor necrosis factor alpha

VCAM – vascular cell adhesion molecule 1 VEGF – vascular endothelial growth factor VECs – valve endothelial cells

VICs – valve interstitial cells vWF – von Willebrand factor αSMA – alpha smooth muscle actin

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1. Introduction

1.1 Aortic valve biology

The aortic valve opens and closes approximately 100.000 times a day. The valve allows blood flow from the left ventricle to the aorta during systole and prevents regurgitation during diastole (1-4). All parts of the aortic valve move in a coordinated manner thanks to precise geometry of the valve and to the complex mechanisms regulating aortic valve functions (1).

1.1.1 Human aortic valve anatomy and development

The normal human aortic valve leaflet is around one millimeter in thickness, there are three of them and they have semilunar shape (4-8). They are named according to their location relative to the coronary arteries as the left coronary, right coronary and noncoronary leaflet (5, 7, 8). They are attached to the fibrous annulus which is a supporting structure (1, 5, 7). Together with the annulus the leaflets form the aortic root (7). All leaflets are composed of three layers: lamina fibrosa, lamina spongiosa and lamina ventricularis (Figure 1) (2, 3, 6, 9- 11). Every layer has its own specific extracellular matrix that provides important biomechanical functions during cardiac cycle (3, 8, 9, 11, 12). Lamina fibrosa (represents 45%

of the thickness) is located on the aortic side of the valve and consists predominantly of circumferentially arranged collagen I and collagen III fibers. This layer confers strength, stiffness and flexibility to the valve leaflets. Lamina spongiosa (represents 35% of the valve thickness) is the middle layer and it is rich in proteoglycans with scattered collagen fibers.

Due to the ability to compress, lamina spongiosa helps resist hemodynamic forces and facilitate movements between the fibrosa and ventricularis. Lamina ventricularis (represents 20% of the thickness) is located on the ventricular side of the valve and its outside is washed by the blood flow during systole. This layer mostly consists of radially oriented elastin fibers and relatively fewer collagen fibers. The elastin fibers allow the valve leaflets to undergo a high level of deformation while it opens and closes (1-3, 9, 11).

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16 Figure 1. Simplified structure of human aortic valve. On the left is a schematic cross section through the noncoronary leaflet of the aortic valve. On the right the blowup shows the trilayed organization of extracellular matrix and the localization of the aortic valve endothelial (VECs) and interstitial cells (VICs). Figure adapted from Rutkovskiy et al. (6).

The nervous system is also represented in aortic valve leaflets, predominantly on the ventricular side. It has been suggested that nerves are required for regulation of valve function in normal state as well as playing a role in diseased valves (13, 14). However, this topic is outside the scope of the current study. The valve leaflets are surrounded by oxygenated blood, additionally, limited microvasculature delivers oxygen within valve leaflets. Blood vessels were found within the spongiosa layer in the outmost third of the valve leaflet. The valve leaflets are thin enough to permit that valvular interstitial cells get oxygen by passive diffusion from the blood stream (15, 16). Markers of lymphatic vessels have been found within aortic valve leaflets; however, existence of a lymphatic system within the valve remains questionable (17).

During embryogenesis the aortic valve develops from the primordial heart tube at around two weeks of gestation (18). At this time the heart tube consists of endocardial cells and cardiomyocytes in an acid-rich substance called the “cardiac jelly” (1, 3, 9, 12, 19, 20).

The development of the aortic valve begins with endothelial-to-mesenchymal transition (EMT) where endocardial cells transform to mesenchymal cells, migrate into the cardiac jelly, proliferate and form local swellings called the cardiac cushions. The cardiac cushions further develop into the valve leaflets (1, 3, 9, 12, 19, 20). Many signaling pathways regulate aortic valve development; the main ones are transforming growth factor beta-1 (TGF-β1), bone morphogenetic protein (BMP), Notch and WNT (1, 9, 19). Remodeling of aortic valves

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17 continues during fetal and postnatal development (21, 22). Interestingly, the aortic valve development has similarities to bone, cartilage and tendon development in relation to regulatory genes and structural proteins involved in the process (3). For example, BMP2 signaling (including BMP2 and its downstream target genes: Sox and Aggrecan) is required for both chondrogenesis in cartilage and endocardial cushion formation in valve leaflets.

Another example: activation of receptor activator of nuclear factor kappa-B ligand (RANKL), its receptor RANK and downstream targets nuclear factor of activated T-cells, cytoplasmic 1 (NFATc1) and cathepsin K are essential for osteoclast lineage development in bone and at the same time they contribute to ECM modeling during valve leaflet formation (3).

1.1.2 Cellular structure of aortic valve

Both surfaces of the valve leaflet are covered by a monolayer of valve endothelial cells and all three layers inside the leaflet are populated by valve interstitial cells (VICs) (1, 6). In addition small populations of smooth muscle cells (23, 24) and bone marrow-derived stem cells (25) have been described to exist in healthy aortic valves.

1.1.2.1 Valve endothelial cells (VECs)

The main functions of valve endothelial cells (VECs) are to maintain valve homeostasis and to form a barrier between circulating blood and the underlying microstructure of the valve (1, 9, 11). The VECs have similar functions and morphology to endothelial cells from most other locations (1). However, some features of VECs remain unique. For instance, the valve endothelial cells are different from vascular endothelial cells regarding their orientation to the blood flow (26). Whereas the long axis of the majority of cells within the monolayer of vascular endothelial cells aligns parallel to the direction of blood flow, the valve endothelial cells align perpendicularly (26). This property is related to the differences in cytoskeleton and mechanotransduction pathways (27). Farivar et al. (27) compared transcriptional profile of untreated endothelial cells from the porcine valve and the aorta, showing that valve endothelial cells had higher transcriptional activity and proliferated more rapidly than vascular endothelial cells. Fifty-five transcriptionally activated genes were common for porcine vascular and valvular endothelium, whereas another 14 genes were unique in vascular and 34 in valvular endothelial cells. Vascular cell adhesion molecule 1 (VCAM) and vascular endothelial cell growth factor (VEGF) were characterized as markers of vascular, but not valvular endothelium (27). VECs express endothelial specific markers

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18 such as PECAM1 (CD31), platelet-derived growth factor receptor (PDGF-R) and von Willebrand factor (vWF) (27, 28).

The phenotypic profile of VECs also depends on age (29) and the side of the leaflet (30-33). The differences between gene expressions on different sides of the valve can possibly be explained by exposure to different types of mechanical stress: mostly oscillatory shear stress on the aortic side of the valve and mostly laminar shear stress on the ventricular side (31, 33, 34). This might be related to the reason why VECs on the ventricular side are more prone to have a protective function against calcification (30, 35).

The study by Bischoff and Aikawa (36) suggested that VECs contain a certain population of progenitor cells that can replenish VICs by re-activating the mechanism of EMT during the postnatal period. EMT can also be activated during disease and in response to mechanical stress (36).

1.1.2.2 Valve interstitial cells (VICs)

The matrix of aortic valve leaflets is populated by valve interstitial cells that maintain physiological valve structure and function. VICs are the predominant cell population in aortic valves (6, 8, 37). VICs communicate with each other through gap junctions, desmosomes and adherens junctions (38). They express cell-cell junction molecules such as N-cadherin, connexin-26 and -43 and desmoglein (38).

In healthy valves the VICs have fibroblast-like morphology (37). In addition, VICs have multipotent properties similar to mesenchymal stem cells and can upon the appropriate stimulation in vitro differentiate into osteoblasts, adipocytes, or chondrocytes (39).

Furthermore, VICs have a different phenotype compared to pericardial cells and dermal fibroblasts (40) as well as to mesenchymal and vascular cells (37).

Specific markers that define healthy VICs are not completely clarified. Firstly, it is important to take into consideration species differences (6). Secondly, markers that were described for VICs isolated from healthy and calcified human valves may be similar, however, the level of expression could be different. For example, alpha smooth muscle actin (ɑSMA) and desmin have much higher expression in human VICs isolated from calcified valves compared to healthy (6). This may denote cells that have drifted away from the original VIC phenotype.

VICs seem to be a heterogeneous population of cells. The study by Brands et al. (41) demonstrated that human VICs from healthy valves express a variety of proteins typical for cardiac and skeletal muscles (troponins, β-MHC, myogenin) as well as non-muscle proteins

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19 (non-muscle isoforms of α- and β- tropomyosins). Consequently, it was suggested that VICs include a population of myofibroblast-like cells with the ability to contract (41). Liu et al. (37) described five distinct phenotypes of VICs: quiescent VICs embryonic progenitor endothelial/mesenchymal cells, progenitor VICs, activated VICs and osteoblastic VICs.

Quiescent VICs are considered to be the population of cells that are responsible for maintenance of healthy valve physiology. Embryonic progenitors endothelial/mesenchymal cells are derived from endothelial cells undergoing EMT at the earliest stage of valve development. EMT may also reappear in diseased adult valves; consequently, these types of cells may only exist in diseased valves. Liu et al. (37) suggested that progenitor VICs, activated VICs, and osteoblastic VICs exist only in calcified valve. Progenitor VICs are a proliferative population of cells that plays a role in valve repair, whereas activated VICs (myofibroblast-like cells) and osteoblastic VICs (osteoblast-like cells) represent the populations of pathologically differentiated VICs (37). Pathological differentiation of VICs will be reviewed in chapter 1.2.5.6.

1.1.2.3 Other types of cells found in the aortic valves

Different phenotypes of VIC appear to glide in one another, but even in healthy human and porcine valves some cells are set so far apart that they can be defined as either fibroblasts, myofibroblasts or smooth muscle cells (SMC), the latter expressing both αSMA and smoothelin (24). While myofibroblasts and fibroblasts are well described and in most of the studies considered to belong to VICs, the origin of SMC is not fully understood or described (24). SMC represent only a small percentage of cells in heathy human aortic valves, but their population increases in calcified valves (23). This type of cells is located on the ventricular side of the valve leaflet and can be identified by expression of several markers including ɑSMA, calponin, SM22 (transgelin), desmin, h-caldesmon, smoothelin and smooth muscle myosin, heavy chain. The latter four markers distinguish SMC from myofibroblasts (23).

A population of bone morrow-derived cells has been shown in the mouse model to be recruited into normal aortic valves and to replenish the population of VICs. This type of cells was characterized by expression of CD45 and Hsp47 (25). Further studies are required to prove the existence of bone marrow-derived cells in human valves.

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1.2 Calcific aortic valve disease

Calcific aortic valve disease (CAVD) is a severe disorder that starts with thickening of aortic valve leaflets (aortic valve sclerosis) and leads to calcification (aortic valve calcification) and obstruction of blood flow from left ventricle to the aorta (aortic valve stenosis) (10, 42). To date there are no effective therapies to prevent or slow the progression of CAVD (43). The only solution for patients with aortic valve stenosis due to calcification is surgical or transcatheter aortic valve replacement.

For many years CAVD was considered a passive process of calcium deposition in aortic valve leaflets related to aging (2, 10, 11). In recent years it became clear that it is an active cellular process occurring within the valve leaflet and involving inflammation, lipoprotein deposition and pathological differentiation of cells (2, 10, 11).

1.2.1 Epidemiology of calcific aortic valve disease

CAVD is the third leading cardiovascular disease after hypertension and ischemic heart disease, and it is the valvular heart disease which is most commonly invasively treated worldwide. CAVD itself is a strong risk factor for cardiovascular mortality, myocardial infarction and heart failure (44). The mortality rate of untreated patients with aortic valve calcification after the onset of symptoms is 80-90% within 10 years (45) and for patients with additional heart failure symptoms it is 50% within one year (42). Symptoms at the late stage of the disease include abnormal heart sound (heart murmur), shortness of breath and other symptoms of heart failure, angina pectoris and sometimes syncope (46). The patients with disease at early stages often do not have any symptoms, and first seek medical help when the disease already has progressed to severe aortic stenosis (42). CAVD with aortic stenosis is diagnosed by clinical examination and echocardiography (42). High resolution computer tomography (CT) scanning is helpful to detect the disease at early stages (47, 48).

The prevalence of CAVD increases exponentially with age (10, 49) and varies depending on the sample age and geographical distribution. In Euro Heart Survey 33.9 % of the European population with valve diseases had aortic stenosis with 81.9 % accompanied by dystrophic calcification, and 54% of them were older than 70 years (50). In the cohort of people considered to be healthy in Helsinki Ageing Study, 53% of people in the age range 75- 86 had signs of aortic valve calcification (51). In the American Cardiovascular Health Study, 29% of the overall healthy cohort over 65 years old had aortic sclerosis, and 2% had aortic stenosis. For the cohort aged over 75 years old the percentage increased to 37% for aortic

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21 sclerosis and to 2.5% for aortic stenosis (52). In a population-based study on the prevalence of CAVD in Mediterranean area, 45.4% of the population over 65 years of age had aortic sclerosis and 3% of them developed aortic stenosis. For the subpopulation over 85 years old these numbers were 73.5% and 7.4% respectively (53). In Norway the incidence of aortic stenosis with calcification in a population over 60 years old was estimated to be 4.9 per 1000 people per year (54). Furthermore, during the last years the prevalence of CAVD is significantly increasing in North America and Europe (49, 51, 52, 54, 55). It has been suggested that the prevalence of CAVD may double during the next 50 years because of an ageing population (43, 56).

1.2.2 Risk factors of calcific aortic valve disease

The major risk factors of developing CAVD are old age (over 65 years), male gender, high plasma low density lipoproteins, hypertension, smoking, and diabetes (including metabolic syndrome) (48, 49, 52, 53, 57). The first three risk factors have the strongest correlation with disease and age is the leading one (57). Elevated serum phosphate levels could be an additional risk factor of CAVD (58). Patients with chronic renal failure are observed to develop aortic valve calcification as well (59). This association was also demonstrated in apolipoprotein E knock out mice with additional chronic renal deficiency and high activity of protease cathepsin S. These mice had accelerated development of aortic valve calcification (60). Interestingly, aortic valve calcification shares risk factors with atherosclerosis (60, 61). A recent study in a large Swedish and Danish cohort found an association of genetically determined elevated level of lipoprotein (a) with the development of aortic valve calcification (47). Although many risk factors for CAVD have been described, the cause of the disease is still elusive. It is important to take into consideration that only some patients with aortic sclerosis will develop aortic stenosis with calcification.

Consequently, some risk factors between these between aortic sclerosis and aortic stenosis may be different (43). More studies to identify predictors as well as new diagnostic strategies of CAVD are needed (43).

1.2.3 Biomarkers of calcific aortic valve disease

Beckmann et al. in his review divided possible biomarkers of CAVD into four groups:

markers of endothelial dysfunction, markers of inflammation, markers of lipid deposition, markers of osteoblastic differentiation and markers of clinical progression (42). The most promising candidates for biomarkers of CAVD appeared to be asymmetric dimethylarginine,

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22 fetuin-A, calcium-phosphorus product, natriuretic peptides and osteopontin. Unlike other markers natriuretic peptides were shown not only to reflect the presence and progression of the disease, but also to predict the prognosis of CAVD. However, specificity of natriuretic peptides for CAVD is low since they are also well-established markers for heart failure (42).

Identification of new biomarkers is required for prediction and/or early diagnosis of the disease (62).

1.2.4 Calcification of tricuspid, bicuspid and unicuspid aortic valves

The majority of aortic valves are tricuspid (TAVs, contain three valve leaflets; Figure 2). Bicuspid aortic valves (BAVs, contain only two valve leaflets) are an independent and powerful risk factor of aortic valve calcification. Even though the prevalence of bicuspid aortic valves (BAVs) is only 0.5-2%, almost all bicuspid aortic valves become calcified before the patient turns 50 (63). Patients with bicuspid valves develop calcific nodules one or two decades earlier than the ones with tricuspid valves (10, 64). It is also known that BAVs occur three times more frequently in males than in females (63).

A US study compared the prevalence of TAVs, BAVs and unicuspid valves (contain only one leaflet) in a cohort of 932 patients with aortic valve calcification aged 26 to 91 years.

Forty-nine per cent of these patients had BAV, 45% had TAV, 5% had unicuspid aortic valves and 1% had valves of an undetermined type (65).

Figure 2. Healthy tricuspid aortic valve (left), calcified tricuspid aortic valve (middle) and calcified bicuspid aortic valve (right). Figure modified from Mathieu et al. (63).

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23 Bicuspid and unicuspid aortic valves are congenital malformations, the nature of which is not fully understood (some aspects of the genetics of BAVs are reviewed in chapter 1.2.5.2).

End stage aortic valve calcification seems to have a lot of similarities between BAVs and TAVs based on the studies comparing genomic profile between them (66, 67). At the same time, there are differences in molecular and cellular mechanisms that underlie calcification of BAVs and TAVs at an early stage (68).

The calcific deposits in aortic valves consist mainly of calcium and phosphorus, with a high content of CO32- (carbonate) akin to bone matrix (69). Besides calcific fibers and compact calcification that might be associated with mature lamellar bone formation in human aortic valves, Bertazzo at al. discovered spherical particles in the aortic valves that are structurally different from bone matrix and are composed of highly crystalline hydroxyapatites (Ca5(PO4)3) (70). Because these particles were present at all stages of CAVD the authors suggested that they may play role in initiation and/or disease progression.

The process of mineralization in aortic valves is gradual, stepwise. Interestingly, it was demonstrated that calcified BAV contain lower amounts of calcium and phosphate and have greater variations in calcium/phosphate ratio compared to calcified TAVs, although BAVs are characterized by heavier calcification then TAVs (69).

1.2.5 Cellular and molecular mechanisms of aortic valve calcification 1.2.5.1 Phases of the disease progression

With or without the underlying genetic predisposition (71), inflammation and mechanical stress have been suggested to trigger the pathophysiology of CAVD (72).

Development of CAVD includes highly complex, strictly regulated and interconnected series of processes (73). A complete and detailed discussion may be beyond the scope of this work, but the basics are outlined below.

It is convenient to distinguish between three phases of disease progression, where the first phase is believed to be analogous to atherosclerosis (73-76) (Figure 3), while the other two are not. This probably explains why the medication used to treat atherosclerosis does not improve the situation in the valves (more on that in chapter 1.4.2.1)

The initiation phase (1) begins with disruption of the endothelial layer on the aortic side of the valve leaflet that facilitates infiltration of inflammatory cells and lipid deposition.

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24 Activated inflammatory cells produce pro-osteogenic factors that at the propagation phase (2) stimulate pathological differentiation of VICs into myofibroblastic and osteogenic pathways (73-76). Inflammatory cells also may induce activation of the complement system in the leaflets (77). Enhanced inflammation and pathological differentiation of cells induce extracellular matrix remodeling that leads to leaflet thickening before the formation of nucleation sites for calcium deposition (microcalcification) (73-76). Once leaflet thickening has occurred, the valve may be on a “road of no return”. Thickening of the leaflets causes intraleaflet hypoxia (78) and stimulates neovascularization (79). Hypoxia in the leaflets influences further leaflet remodeling (78). Changes in extracellular matrix elasticity also can contribute to VIC differentiation (73-76). Calcification itself increases mechanical stress and endothelial injury inducing further calcification. The final, advanced phase (3) is characterized by progressive calcium deposition (macrocalcification) (73-76).

Figure 3. Three phases of calcific aortic valve disease progression: (1) initiation phase, (2) propagation phase, (3) last phase. VIC - valve interstitial cell, LDL - low density lipoproteins, ECM - extracellular matrix. Figure adapted from Garcia-Rodriguez et al. (76).

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25 Contemporary knowledge regarding potential triggers of the disease and mechanisms that are involved in first and second phases of disease progression are reviewed below.

1.2.5.2 Genetic factors

Bicuspid aortic valve is a congenital abnormality characterized by fusion (or rather lack of separation) of two aortic valve leaflets (80). Unlike TAVs, as already mentioned, the BAVs, are predisposed to becoming calcified. The genetic etiology of BAVs is not clarified (80). Whole genome sequencing studies of human BAVs concluded that there is no single disease-causing mutation and the development of BAVs is characterized by involvement of more than one genetic variant (81). Heterogeneous nature of BAVs partly explains their different anatomical phenotypes, for instance, which of the leaflets are fused together (right coronary, left coronary or noncoronary) (80). One of the well-described causes of human BAV development is a number of mutations in Notch1 gene that encodes transmembrane receptor NOTCH1. NOTCH1 is a component of Notch signaling pathway that is, among innumerable other things, involved in the development of aortic and pulmonary valves.

Early signaling events including Notch-dependent mechanisms that are responsible for the initiation of aortic valve calcification are suggested to be different between human BAVs and TAVs (68). Garg et al. (71) found that familiar clustering of BAVs with calcification was associated with mutations in Notch1. In vivo studies demonstrated that 30% of mice with Notch1 deletion in VECs developed BAVs and had characteristics of human aortic valve stenosis including thickening of aortic valve leaflets and calcification (82). In addition, several mouse models with a knockout of genes involved in cardiac development were shown to develop BAVs. For example, endothelial nitric oxide synthase knockout mice (83) and mice with knockout of Gata5 gene (encodes transcription factor Gata 5, involved in valve development) (84) in separate studies developed BAVs with specific anatomical phenotype (fusion of the noncoronory and right coronary valve leaflets). Interestingly, a study comparing microRNA (non-coding RNAs that regulate gene expression at the translational level) profile between human BAVs and TAVs discovered that 34 microRNA were differentially regulated between them (85). No single microRNA has been linked, though, to the development of CAVD. Further studies are required to dissect the role of these genes and their signaling pathways in human aortic valves.

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26 1.2.5.3 Inflammation

CAVD is universally considered to be an inflammatory disease (10). It was suggested that both innate and adaptive immune responses are involved in the pathophysiology of CAVD (86, 87). The cell types involved in chronic inflammation in human calcified aortic valves include monocytes, macrophages, mast cells, CD4+ lymphocytes and CD8+

lymphocytes (86), whereas normal aortic valves do not contain inflammatory cells except for a few macrophages (88). Recruitment of inflammatory cells probably happens at the initiation phase of the disease since they were shown to accumulate in early aortic valve lesion in a mouse model of CAVD (89). Inflammatory cells secrete several cytokines such as tumor necrosis factor alpha (TNF-ɑ), interleukin-1 beta (IL-1β), interleukin-6 (IL-6) and receptor activator of nuclear factor-kappa B ligand (RANKL) (89-94). In VICs these cytokines stimulate expression of matrix metalloproteinases (MMPs), which can degrade extracellular matrix proteins leading to thickening of aortic valve leaflets as well as promote neovascularization (88, 93, 95). In addition, TNF-ɑ was shown to directly stimulate osteogenic differentiation with calcification of VICs through the activation of nuclear factor- kappa B (NF-kB) pathway (90, 92). Inflammatory cells may also secrete C-reactive protein – potential activator of compliment system (a cascade system of proteolytic enzymes that leads to cell lysis or inflammatory activation of target cells (77). The thickening of the valve leaflet that occurs in the propagation phase also seems to lead to the hypoxia in the valve tissue (78).

Hypoxia in leaflets increases expression of hypoxia inducible factor-1 alpha (HIF-1α) activating the innate immune system via toll-like receptors (TLRs) and nuclear factor-kappa B (NF-kB) (96).

A study by Nagy and Bäck suggested that VICs can trigger inflammation by transforming into immune-like cells and secreting leukotrienes (97). VICs from calcified valves have increased expression of gene encoding leukotriene-synthesizing enzyme 5- lipoxygenase and a decreased DNA methylation in the promoter of this gene, signifying its transcriptional activity. Thus the epigenetic modifications of the VIC genome may be involved in the activation of inflammation leading to CAVD (97).

Non-sterile inflammation

Chlamydophila pneumoniae, Mycoplasma pneumoniae, Cytomegalovirus and Epstein- Barr virus were found in aortic valve leaflets of patients with aortic valve calcification (98- 101). For example, in the study of Bayram et al. eight (24.2%) out of 33 patients with aortic valve calcification were positive for Chlamydophila pneumoniae, three (9.1%) patients were

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27 positive for Mycoplasma pneumoniae, seven (21.2%) were positive for Cytomegalovirus and one (3%) patient was positive for Epstein-Barr virus (100). The link between bacterial and viral infection and aortic valve calcification is not established yet; therefore it would be tempting to know whether this infection may initiate calcification or accelerate the process (101). Interestingly, Chlamydophila pneumoniae was demonstrated in aortic valves a long time before patients were in need of surgery (101). It was further shown on an in vivo rabbit model that injection of bacteria from the oral cavity induced calcification in aortic valves with endothelial surface injured beforehand (102).

Bacteria and viruses are identified by innate immune system as bearers of pathogen- associated molecular patterns (PAMPs), recognized by toll-like receptors (TLRs) on the cell surface (76). TLR4 and TLR2 are sensitive for lipopolysaccharides (LPS), a component of the outer membrane of gram-negative bacteria. Viral molecules in turn are recognized by TLR3 and TLR9 (76). Meng et al. were the first who showed expression of TLR2 and TLR4 in VICs (103). Subsequent studies demonstrated increased expression of these receptors in calcified aortic valves (103-105). In human VICs from healthy valves, LPS through binding with TLR4 and TLR2 receptors triggers activation of NF-kB signaling that induced expression of proinflammatory cytokines and adhesion molecules (103). Furthermore, it was shown that stimulation with LPS activates expression of osteogenic markers such as BMP2, RUNX2 and alkaline phosphatase (103, 106). LPS also promotes calcification of VICs after stimulation with osteogenic medium (104, 105, 107). Remarkably, VICs from calcified valves are more sensitive to LPS stimulation compared to VICs from healthy valves (104, 107, 108). Using in vivo mouse model it was demonstrated that TLR2 and TLR4 inactivation prevents aortic valve thickening (107). TLR2 and TLR4 also recognize peptidoglycans which are components of the outer membrane of gram-positive bacteria. Stimulation of human VICs with synthetic lipopeptide Pam3CSK4 that mimics the amino terminus of lipoproteins of both gram-negative and gram-positive bacteria, also stimulated expression of osteogenic markers and enhanced calcification (104, 105, 107). Interestingly, the treatment of human VICs from calcified valves with an anti-inflammatory cytokine interleukin-37 (IL-37) suppressed the osteogenic gene expression and calcification that were induced by the stimulation of cells with LPS or Pam3CSK4 in combination with osteogenic medium (107). Exposure of human VICs to agonists mimicking viral patterns activates the TLR3 receptor through the NF-kB pathway and promotes osteogenic differentiation of VICs (105, 109).

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28 Sterile inflammation

Sterile inflammation is pathogen-free inflammation induced by endogenous molecules that are released upon tissue injury (76). These molecules are called damage-associated molecule patterns (DAMPs). Several DAMPs that are shown to be associated with CAVD are listed below.

Sterile inflammation in CAVD may be induced by circulating lipids (110). As mentioned before, elevated levels of lipoprotein(a) are a genetically associated risk factor of CAVD (47). Oxidized phospholipids (OxPL) and apolipoptotein(a) that form lipoprotein(a) have been found to be attached to injured endothelium (111-113). Mechanical stress is suggested to facilitate delivery of lipoproteins into the valve (114). Levels of lipoprotein(a) and OxPL are increased in explanted valves from patients with CAVD (115, 116). Most OxPL in humans bind to apolipoptotein(a), but the remaining ones bind to apolipoprotein B (114). Recently, increased levels of both lipoprotein(a) and OxPL/apolipoprotein B were linked to progression of aortic stenosis with severe calcification (117). The oxidized form of lipoproteins that include apolipoprotein B and phospholipids in their structure is called oxidized low-density lipoprotein OxLDL. Elevated level of OxLDL is found in plasma and valves of patients with aortic stenosis (118, 119) and is associated with ECM remodeling in calcified valves (119, 120). In in vitro studies the treatment of VICs with OxLDL alone mimicked the effect of LPS to stimulate osteogenic differentiation through the activation of TLR2, TLR4 receptors, and NF-κB signaling (107, 121). Also there are data suggesting that oxLDL initiates activation of compliment system (77).

The other DAMPs that were linked to sterile inflammation in CAVD are high mobility group protein B1 (HMGB1, proinflammatory cytokine), matrilin-2 and galectin-3 (extracellular matrix proteins) and cyclooxygenase 2 (Cox2, proinflammatory enzyme). In separate studies the overexpression of HMGB1 (122), biglycan (123), matrilin-2 (124) galectin-3 (125) and Cox2 (126) were found in patients with CAVD. In vitro experiments showed increased expression of inflammatory and osteogenic markers after stimulation of VICs by recombinant forms of these proteins (124-130), suggesting a mechanistic link.

1.2.5.4 Role of mechanical stress

The aortic valve is exposed to substantial mechanical stress (defined as force per valve area) during systole and diastole (34, 131). Two types of mechanical stress that are involved in the biomechanics of the aortic valve are axial stress that is perpendicular to the valve area

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29 and shear stress that is parallel to the valve area (34). During systole, the ventricular side of the valve leaflet is exposed to laminar shear stress when the blood flows past the leaflet. At the same time the aortic side of the valve leaflet is exposed to oscillatory shear stress, created by the blood behind the valve leaflets pushed along with the main flow after valve opening.

The normal axial stress (or axial pressure) occurs during diastole when the valve is closed as a result of transvalvular pressure gradient (34, 131).

Since the valve is an elastic structure, the mechanical forces applied to it result in the valve deformation, which affects the cellular environment within the valve. This deformation is usually called strain, and it is measured as a change in length and in relative angle of different structures. Bäck et al. distinguished between bending strain (changes in the leaflet curvature) and tensile stain (elongation of valve leaflet) (34) (Figure 4).

Figure 4. Types of mechanical stress and strain that the aortic valve leaflet experiences during systole and diastole. During systole, the ventricular side of the valve is exposed to laminar stress and the aortic side of the valve is subjected to oscillatory shear stress (dotted violet arrows) that result in bending strain (curved solid blue arrows). During diastole, axial pressure on the aortic side of the valve leads to tensile strain (straight solid green arrows). At the same time in diastole the aortic side of the valve is exposed to oscillatory and laminar shear stress (dotted violet and yellow arrows). Figure modified from Magnus Back et al. (34).

The aortic valve is exposed the pathological mechanical stress as a consequence of aortic stenosis and aortic valve calcification (34, 131). At the advanced stage of aortic stenosis, the calcified aortic valve leaflets cannot open or close properly and they are subjected to

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30 abnormally high blood velocity and pressure gradient between the aorta and the left ventricle (34). With progression of aortic stenosis more blood flow becomes turbulent, which may cause damage to blood cells and to the endothelial layer of the valve (131).

It is believed that abnormal mechanical stress and strain play a role in initiation and at the early, subclinical stages of CAVD. Firstly, transvalvular pressure gradient increases considerably in aortic valves with advancing age as a consequence of raising blood pressure (132). Moreover, aortic valve becomes thicker and stiffer due to ageing, thereby reducing its overall strain At different time points calcific aortic valve disease may be associated with both decreased and increased strain (34). This is a sort of a chicken-and-egg question, as excessive strain being a result of increased mechanical stress leads to compensatory thickening of the valve, so that the potential for strain (deformation) is reduced. Changes in mechanical stress and strain may potentially lead to aortic valve calcification. Mechanical stress and strain also contribute to initiation of CAVD when malformations of the valve take place, for example, in bicuspid aortic valves (34). Non-physiological geometry of BAVs leads to abnormally high shear stress (133) and increased bending strain (134) that can cause valve calcification. It could be one of the reasons why BAVs are prone to calcification at an early age (34).

Numerous in vitro and ex-vivo studies simulated different types of physiological and non-physiological mechanical stress and demonstrated their effect on gene and protein expression in valve tissue (135-140), VECs (26, 141, 142) and VICs (143-145). For example, non-physiological shear stress increases expression of inflammatory markers on the aortic side of the leaflet and not on the ventricular side (138). Abnormally low shear stress upregulates EMT- and inflammation-related genes in VECs (142). Abnormally high cyclic strain triggers expression of osteogenic markers in human VICs (143) and induces calcification in porcine aortic valves (135).

1.2.5.5 Cross-talk between aortic valve endothelial and valve interstitial cells

Recent studies of the molecular and cellular mechanisms of CAVD have emphasized the importance of VECs-VICs interaction. Butcher’s lab developed an in vitro 3D co-culture models of VECs and VICs to better understand their interaction (146). They demonstrated that porcine VICs reduce expression of myofibroblastic gene αSMA when grown in co- culture with VECs, implying that VECs are necessary to regulate and maintain the physiological VIC phenotype (myofibroblasts are considered a stage in pathological differentiation). Subsequent studies confirmed that VECs inhibited myofibroblastic or

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31 osteogenic differentiation of VICs in co-culture (35, 147, 148) and this effect was prevented by nitric oxide synthase blocker L-NG-nitroarginine methyl ester (L-NAME). As the VECs produce nitric oxide, it was suggested that VEC-derived nitric oxide might be the potential inhibitor of the early phases of valve calcification (35, 147, 148). Interestingly, in human valves the VECs had a higher expression of nitric oxide synthase on the ventricular side of the valve, where calcification does not develop, compared to the aortic side where calcification takes place (35).

As mentioned above, VECs undergo endothelial-to-mesenchymal transition (EMT) during valve development (chapter 1.1.1). This process could play an important role in the pathogenesis of CAVD (149). In a study by Farrar et al. it was shown that VECs seeded on top of the collagen gel and stimulated by tumor necrosis factor alpha (TNF-α), migrated inside the gel and underwent EMT (150). The transformed VECs expressed markers of VIC, and were characterized by low expression of nitric oxide synthase (150), but increased intracellular oxidative stress and secretion of hydrogen peroxide (151). A similar study confirmed that the VECs undergo EMT after stimulation by TGF-β1 and osteogenic medium (152). In addition, it was suggested that the VICs inhibit EMT of VEC in co-culture (152).

EMT seems to recapitulate the embryonic development, where the VICs are the descendants of invading endothelial cells.

Recently White et al. found that physiological shear stress activates Notch 1 signaling in human VECs, which may lead to the downregulation of osteogenic genes in VICs and prevent calcification (141). A study in mice showed that Notch 1 signaling activates TNF-α expression in VECs which is necessary for post-EMT development of aortic valves.

Overexpression or deletion of Notch 1 gene in VECs therefore led to development of defective aortic valves (82).

To summarize, the cross talk between VECs and VICs plays an important role in the pathogenesis of CAVD, where in healthy aortic valves VEC-VIC interaction may prevent EMT of VECs and/or pathological differentiation of VICs. The disruption of the endothelial barrier may thus be a trigger event in the pathophysiology of CAVD.

1.2.5.6 Pathological differentiation of valve interstitial cells

Most researchers agree that the key mechanism that underlies CAVD is pathological differentiation of VICs into myofibroblast- and osteoblast-like cells (Figure 5). The relative contribution of these two pathways during disease progression remains questionable (6, 153,

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32 154). Only one of the studies suggests the approximate percentage ratio: 13% of total valves excised during valve replacement surgery (256 aortic, 91 mitral) were described to contain mature lamellar bone with osteoblasts and osteoclasts, 1% - endochondral bone formation, whereas 83% had dystrophic calcification that might be related to myofibroblast-like cells (155). It is not clear whether these two pathways of calcification occur simultaneously or sequentially in the cells (154). The other question is the origin of osteoblast-like cells. Some studies claim that myofibroblastic and osteoblastic pathways are independent and parallel (6, 37, 154), whereas others suggest that osteoblast-like cells are derived by transdifferentiation from myofibroblast-like cells (76, 149, 156, 157). There is also possibility that VICs de- differentiate into multipotent progenitor-like state before differentiation down to osteogenic pathway (149).

Figure 5. Pathological differentiation of quiescent valve interstitial cells (qVICs) into myofibroblast-like cells (by means of TGF-β1) or osteoblast-like cells (by osteogenic factors) (the collage created using www.smart.servier.com). Myofibroblast-like cells are characterized by the presence of stress fibers and by increased expression of alpha-smooth muscle actin (αSMA), calponin and transgelin (SM22). Osteoblast-like cells are characterized by increased expression of bone morphogenetic protein 2 (BMP2), runt-related transcription factor 2 (RUNX2), alkaline phosphatase (ALP) and some other markers. Myofibroblast-like cells cause dystrophic calcification through apoptosis whereas osteoblast-like cells cause calcification by the process similar to lamellar bone formation.

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33 Myofibroblastic differentiation of VIC

Myofibroblasts are defined as fibroblasts that have some properties of smooth muscle cells and are characterized by the presence of stress fibers (composed mainly of alpha smooth muscle actin, αSMA) which lend them the ability to contract the extracellular matrix (158). It seems that myofibroblast-like cells in CAVD play an important role in extracellular matrix remodeling (37) (reviewed in chapter 1.2.5.7). αSMA at high expression levels is a well described marker of myofibroblasts (158). The expression of αSMA is increased in calcified human aortic valves compared to the healthy valves (159). αSMA, calponin and SM22 are the established markers to detect myofibroblast-like cells (23, 68, 160). The VICs differentiated into myofibroblasts are also functionally characterized by their ability of gel contraction (161, 162).

It is believed that the differentiation of cells into myofibroblasts requires TGF-β1 signaling (158, 163, 164). Increased TGF-β1 concentration was found in human calcified aortic valve leaflets (163); in many studies the in vitro differentiation of human VICs into myofibroblast-like cells was achieved by TGF-β1 stimulation (148, 164-167). At the same time, several studies on animal cells claim that TGF-β1 is not necessary for myofibroblastic differentiation (168, 169).

It is believed that myofibroblastic differentiation leads to cell contraction, aggregation of multiple cells and the detachment of aggregates from underlying ECM, followed by apoptosis. The multicellular aggregates that undergo apoptosis serve as a substrate for calcium accumulation (8, 153). Chen and Simmons suggested that apoptosis during myofibroblastic differentiation of VICs is a result of anoikis (a form of programmed cell death) (8). It is important to emphasize that calcification via myofibroblastic differentiation has only been established in animal cells (148, 154, 168, 170). There are no studies in human cells to date that would confirm these data.

Osteogenic differentiation of VIC

Osteoblast- and osteoclast-like cells have been identified histologically in calcified aortic valves (155), but never in healthy aortic valves. Many markers that are attributed to osteoblasts cells have been found in valves of patients with CAVD: bone morphogenetic protein 2 (BMP2), bone morphogenetic protein 4 (BMP4) (155), runt-related transcription factor 2 (RUNX2) (156, 171, 172), β-catenin (CTNNB1) (171), alkaline phosphatase (ALP) (156, 172), osteonectin (ON) (172), thrombospondin 1 (THBS1) (172), thrombospondin 2

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34 (THBS2) (172), osteopontin (OPN) (156, 173), bone sialoprotein (BSP) (156, 173), osteocalcin (OCN) (156) and osteoprotegerin (OPG) (173). Most of these markers are also expressed by VICs differentiated into osteoblast-like cells in vitro (174-176). The most established method for osteogenic differentiation of VICs is stimulating them with osteogenic medium containing ascorbic acid, dexamethasone and β-glycerophosphate (103, 174, 176, 177). The only study aimed at comparing the phenotype of VICs, pre-osteoblasts and mature osteoblasts differentiated by osteogenic medium found some differences between these cell types in the magnitude of ALP, OCN and αSMA expression (154). Downregulation of αSMA in VICs and not in the other cell types under the stimulation with osteogenic medium was emphasized in this study as a possible indicator that VICs can differentiate directly to osteoblast-like cells without progressing through the myofibroblastic stage (154). However, a limitation of this study is that different cell lines were obtained from different species.

Calcification through osteogenic differentiation of VICs has similarities to physiological osteogenesis in bone tissue. Chiefly lamellar bone formation (for the most part occurring by intramembranous ossification) was found in human calcified aortic valves, however, signs of endochondral bone formation (bone development from cartilage typical for the axial skeleton and the extremities) were also observed (155, 156). In addition, Egan et al.

identified the presence of bone marrow-derived circulating osteogenic precursor cells characterized by co-expression of osteogenic marker type 1 collagen and hematopoietic marker CD45 in valves of patients with aortic stenosis (178). They suggested that this cell population may contribute to osteogenic differentiation in CAVD. Thus, there remains a possibility that osteogenesis in valves is actually “metastatic” rather than intrinsic to the diseased valve.

1.2.5.7 Extracellular matrix remodeling

Organization, composition and biomechanical properties of extracellular matrix (ECM) in aortic valves undergo dramatic changes in the pathogenesis of CAVD (8). It is important to note that calcification occurs predominantly in the fibrosa layer, exclusively on the aortic side of the valve leaflet (8, 112, 113). The reason for that is unknown, but it probably is a consequence of unique biomechanical-, biochemical- cellular, and extracellular matrix - mediated factors (179). Remodeling of ECM during CAVD is characterized by simultaneous ECM protein synthesis and degradation, a change in composition, and ultimately disruption of the tri-layer structure and thickening of the valve leaflet (8).

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35 ECM degradation in aortic valves during disease progression is associated with increased level of matrix metalloproteinases (MMPs): MMP-1, MMP-2, MMP-3 and MMP-9, their tissue inhibitors (TIMPs): TIMPs-1, TIMPs-2 (180-185) and cathepsins S, K, V and G (60, 186, 187), secreted by myofibroblast-like cells (differentiated VICs) and inflammatory cells. Due to the activity of these proteins, collagen fibers in fibrosa layer become disorganized (188), and elastin fibers on the ventricularis become disrupted and fragmented (12). At some point, the ECM is enriched with bone-specific proteins such as osteocalcin, osteopontin and bone sialoprotein, which further promote osteogenic differentiation of VICs (156, 173, 189, 190).

CAVD progression is further characterized by increased content of proteoglycans and glycosaminoglycans - components of spongiosa layer (12, 191). Their contribution to disease development is possibly the accumulation of lipoprotein and sequestration of growth factors (192) and cytokines, thus making them more available for cells (193). Glycoproteins such as chondromodulin I and tenascin also contribute to the process of ECM remodeling.

Downregulation of chondromodulin I leads to the neovascularization of valve tissue that may promote calcification by delivering osteogenic factors to the cells (194). Elevated expression of tenascin may induce expression of MMPs in VICs (185).

In addition to extracellular matrix degradation, there is ample evidence of valve fibrosis, where new, but biomechanically incompetent matrix is synthetized. The central role in the regulation of fibrotic processes inside the valve is attributed to renin-angiotensin- system (RAS) (195). Angiotensinogen is a peptide produced by the liver that is cleaved to angiotensin I by the enzyme named renin, and angiotensin I in turn is cleaved to angiotensin II by the angiotensin-converting enzyme (ACE) (195). ACE is co-localized with apolipoprotein B in lesions of calcified aortic valves and is present on plasma LDL. Therefore it is suggested that ACE is likely delivered into the aortic valve by LDL (196). Angiotensin II is a ligand that bounds to angiotensin II type 1 (AT1R) receptor and may mediate the fibrotic process in aortic valves, for example, through activation of TGF-β1 which increases collagen synthesis (197).

Thickening of the valve leaflet due to the remodeling of ECM results in reduced elasticity. Elasticity of the valve leaflet defines its biomechanical function, but also influences the fate of resident cells, including VICs. For example, several studies demonstrated that the VICs differentiated into myofibroblast-like cells when they were cultured on stiffened collagen gel (compared to the compliant gel) and into osteoblast-like cells when they were cultured on even stiffer collagen substrate (198, 199). These data echoe the known mechanical regulation of osteogenic differentiation (200).

Cellular and molecular mechanisms of human aortic valve calcification