VOLTAGE-SENSITIVE CALCIUM CHANNELS IN BONE:
THE ROLE OF ELECTRICALLY EXCITABLE CALCIUM
CHANNELS IN A NON-EXCITABLE TISSUE
by
Case Gregory
A thesis submitted to the Faculty of the University of Delaware in partial fulfillment of the requirements for the degree of Master of Science in Biological Sciences
Summer 2020
© 2020 Case Gregory All Rights Reserved
VOLTAGE-SENSITIVE CALCIUM CHANNELS IN BONE:
THE ROLE OF ELECTRICALLY EXCITABLE CALCIUM
CHANNELS IN A NON-EXCITABLE TISSUE
by
Case Gregory
Approved: ______Randall A. Duncan, Ph.D. Professor in charge of thesis on behalf of the Advisory Committee
Approved: ______Velia M. Fowler, Ph.D. Chair of the Department of Biological Sciences
Approved: ______John A. Pelesko, Ph.D. Dean of the of College Arts and Sciences
Approved: ______Louis F. Rossi, Ph.D. Vice Provost for Graduate and Professional Education and Dean of the Graduate College ACKNOWLEDGMENTS
I would like to thank my advisor Dr. Randall Duncan for his guidance and mentorship during my time in his research lab. He has taught me what it means to be a scientific researcher and very many lessons about life outside of science as well. I can truly say that I have grown tremendously as a person as a result of the time I have spent in his lab. Throughout the years, our lab has persisted in the face of many obstacles—construction fires, multiple moves, funding difficulties, and more. None of this would have been possible without the support and friendship of the many great people in the Duncan lab. I would like to say a special thanks to Dr. Mary Boggs, who served as the heart, soul, and glue of the Duncan lab. Dr. Boggs consistently went above and beyond what was expected of her as mentor, a manager, and a friend. I would also like to thank Dr. Carlton Cooper for his advice and our many conversations in which he served as both a teacher and a friend. I am also grateful to the other graduate and undergraduate students who have been a part of the Duncan lab throughout the years. I would especially like to thank my fellow graduate student Nick Trompeter, who was pivotal as leader in the lab and Abigail DeLa Pez, whose tireless work as an undergraduate researcher was critical to my research. I would like to thank my committee members, Dr. Gary Laverty, Dr. Jessica Tanis, Dr. Liyun Wang, and Dr. William Cain for providing advice that was essential towards the completion of my research goals. I would also like to extend a thank you to Dr. Selva for her advice and assistance in surmounting research challenges and completing my degree. I would like to thank the Bridge to the Doctorate Program and
iii Dr. Michael Vaughan for providing me with the opportunity to conduct my research. Dr. Vaughan has provided invaluable guidance and mentorship throughout my time in the program, and his support has been unwavering. I would also like to say a special thanks to Betty Cowgill, who is a truly indispensable part of the Biology department. Her words of encouragement and help with organization have been a key part in the success of every student I have known throughout my time in the program. Thank you to everyone in the biology department for making this more than just a typical graduate school experience. I have made so many friends along the way. I wish you all the best, and I hope to stay in touch to see where life takes you. Finally, I would like to thank my parents, Donald and Danita Gregory, who have always pushed and supported me to do great things. I am eternally grateful for all that they have done to make me into the man I am today.
iv TABLE OF CONTENTS
LIST OF TABLES ...... vii LIST OF FIGURES ...... viii LIST OF ABBREVIATIONS ...... x ABSTRACT ...... xii
Chapter
1 INTRODUCTION ...... 1
1.1 Bone Tissue Dynamics and Function in the Body ...... 1 1.2 Bone Remodeling with Age, Osteoporosis, and Current Treatment Strategies ...... 9 1.3 Voltage-Sensitive Calcium Channels ...... 13 1.4 The L-VSCC in Bone ...... 21 1.5 The T-VSCC in Bone ...... 25 1.6 Mechanostransduction and VSCC-Mediated Responses in Bone ...... 26 1.7 Mechanoelectric Response in Bone ...... 32
2 MATERIALS AND METHODS ...... 36
2.1 Pharmacological Agents ...... 36 2.2 Cell Culture ...... 36 2.3 Proliferation Assays ...... 37 2.4 Annexin-V Cell Viability Assays ...... 38 2.5 Acridine Orange Cell Viability Assays ...... 41 2.6 Cell Cycle Assays ...... 43 2.7 Creating a PEMF Generator ...... 45 2.8 Proliferation Assays for Cells Exposed to PEMF ...... 47 2.9 Fluorescent Microscopy Assays for Cells Exposed to PEMF ...... 48
3 RESULTS ...... 51
3.1 The L-VSCC is necessary for proliferation of MC3T3-E1 cells ...... 51 3.2 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells ... 53 3.3 Inhibition of the L-VSCC causes cell cycle slowing ...... 56 3.4 T-VSCC inhibition reduces cell number of MC3T3-E1 in a dose- dependent manner ...... 59
v 3.5 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells ...... 60 3.6 VSCC activation is necessary for PEMF-induced increase in proliferation of MC3T3-E1 cells ...... 63
4 CONCLUSION AND DISCUSSION ...... 67
4.1 Discussion ...... 67 4.2 Future Directions ...... 80 4.3 Conclusion ...... 83
REFERENCES ...... 85
vi LIST OF TABLES
Table 2.1 Calculations of currents necessary to generate four different magnetic fields ...... 47
vii LIST OF FIGURES
Figure 1.1 Cellular lineage of osteoclasts, osteoblasts, and osteocytes: ...... 2
Figure 1.2 Overview of bone anatomy ...... 4
Figure 1.3 The stages of bone remodeling ...... 6
Figure 1.4 Generic model of a voltage-sensitive calcium channel and its various subunits ...... 14
Figure 1.5 Mechanism of mechano-coupling in bone via fluid shear ...... 29
Figure 2.1 Representative images of annexin-V assay gating ...... 40
Figure 2.2 Criteria for assessing cell viability via acridine orange assay ...... 42
Figure 2.3 Schematic of a PEMF bioreactor ...... 46
Figure 2.4 Representative images of quinacrine staining analyzed via fluorescent microscopy ...... 49
Figure 3.1 The L-VSCC is necessary for proliferation of MC3T3-E1 cells ...... 52
Figure 3.2 Inhibition of the L-VSCC reduces proliferation of MC3T3-E1 cells in a dose-dependent manner ...... 53
Figure 3.3 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells (as measured via an annexin-V assay) ...... 54
Figure 3.4 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells (as measured via acridine orange assay) ...... 55
Figure 3.5 Serum starvation induces synchronization of MC3T3-E1 cells in the G1/G0 phase: ...... 57
Figure 3.6 Inhibition of the L-VSCC causes cell cycle slowing ...... 58
Figure 3.7 T-VSCC inhibition reduces cell number in a dose-dependent manner ...... 60
viii Figure 3.8 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells (as measured via an annexin-V assay) ...... 61
Figure 3.9 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells (as measured via an acridine orange assays) ...... 62
Figure 3.10 VSCC activation is necessary for PEMF-induced increase in proliferation of MC3T3-E1 cells ...... 64
Figure 3.11 PEMF exposure induces extracellular release of ATP ...... 66
Figure 4.1 Proposed model of the role of calcium-mediated signaling in pre- osteoblast proliferation and survival ...... 78
ix LIST OF ABBREVIATIONS
α-MEM Minimum Essential Medium Eagle, alpha modification AO Acridine orange BMD Bone mineral density CaMKII Calcium/calmodulin-dependent protein kinase II CBP CREB-binding protein CREB cAMP element-binding protein DPBS Dulbecco’s phosphate-buffered solution DMSO Dimethyl sulfoxide DXA Dual-energy X-ray absorptiometry EB Ethidium bromide ECG Electrocardiogram ES Electrical stimulation FBS Fetal bovine serum FS media Fully supplemented media (10% fetal bovine serum) GAG Glycosaminoglycan HBSS Hank’s balanced salt solution HRT Hormone replacement therapy HVA channels High voltage-activated channels iPHAH Idiopathic pulmonary arterial hypertension LVA channels Low voltage-activated channels NFAT Nuclear factor of activated T-cells
x NNC NNC 55-0396 M-CSF Macrophage colony-stimulating factor PKA Protein kinase A PI Propidium iodide PPA’s Prototypical phenylalkylamines PTH Parathyroid hormone PEMF Pulsed-electromagnetic field PASMC’s Pulmonary artery smooth muscle cells RANK Receptor activator for nuclear factor κ B RANKL Receptor activator for nuclear factor κ B ligand
SA cation channels Stretch-activated cation channels VSCC Voltage-sensitive calcium channel
xi ABSTRACT
Bone is a dynamic, living tissue that serves many functions throughout the body, including locomotion, protection of vital organs, and mineral homeostasis. Central to these roles, is the ability of bone to adapt its mass, architecture, and strength according to its environment and the body’s needs (R. L. Duncan & Turner, 1995). These changes take place throughout one’s life and occur via bone remodeling, a process by which specialized cells called “osteoclasts” resorb old or damaged bone and allow for “osteoblasts” to form new bone in its place. Here, I seek to define the roles that specific calcium channels play within the osteoblast to help build new bone and explore novel means of therapeutically targeting these channels to address conditions of bone loss. Voltage-sensitive calcium channels (VSCC’s) are activated by membrane depolarization, allowing calcium influx into the cell to regulate a number of cellular processes. VSCC’s are found in both excitable and non-excitable tissues throughout the body, yet how these channels are activated in non-excitable tissue and how they alter cell function are unknown (Catterall, 2011). In bone, L-type and T-type VSCCs are found in all cells of the osteogenic lineage except for osteocytes, where the L- VSCC is lost (Shao & et al., 2005). The loss of the L-VSCC in the osteocyte parallels the loss of a proliferative phenotype, which leads me to postulate that the L-VSCC is essential for proliferation of osteoprogenitor cells and osteoblasts. The presence of the T-VSCC throughout the entire osteogenic lineage suggests that it may play a consistent and critical role in all bone cells. This is further supported by the unique
xii electrophysiological properties of the T-VSCC which allow it to become activated easily and remain activated at a steady state by low levels of stimulation. Together, these activation properties and expression patterns suggest that the T-VSCC serves a fundamental process that needs to be steadily maintained; this leads me to hypothesize that the T-VSCC is necessary for the survival of osteogenic cells. The role of VSCC’s is well-characterized in excitable tissues, and VSCC inhibitors have been successfully used in excitable tissues to address conditions such as angina, high blood pressure, and premature labor (Pontremoli, Leoncini, & Parodi, 2014; Songthamwat, 2018; Terry, 1982). However, VSCC’s inhibitors are deleterious to bone, and thus are not viable candidates for treating issues of bone loss, but rather are more akin to analogs for osteoporosis and other conditions of bone loss. VSCC inhibitors have been effective as research tools to help researchers better understand the role of these channels in bone, but there is still an urgent need for agents which can activate bone in a manner that will offset bone loss. The ability of VSCC’s to be activated by mechanical load poses limited therapeutic potential for those who are unable to exert the necessary mechanical strains to counteract bone loss in vivo. Pulsed electromagnetic field (PEMF) stimulation shows promising therapeutic potential as a non-invasive means of combating issues of bone loss, since evidence suggests that PEMF’s can be adjusted to selectively target specific VSCC’s within certain tissues (Buckner, Buckner, Koren, Persinger, & Lafrenie, 2015). Here, I explore pulsed electromagnetic fields as an alternative stimulus to mechanical stimuli in bone and hypothesize that pulsed electromagnetic fields can be used to activate VSCC-mediated processes within bone cells.
xiii Our lab has previously shown that both the L-VSCC and the T-VSCC are essential to the increase in bone formation in response to mechanical loading (J. Li, Liu, Ke, Duncan, & Turner, 2005; Owan, Ibaraki, Duncan, Turner, & Burr, 1999; Ryder & Duncan, 2001). I now show that these channels are necessary for proliferation and survival of osteoblasts even in the absence of mechanical load. Here, I demonstrate that T-VSCC inhibition, but not L-VSCC inhibition, results in reduced cell survival of pre-osteoblast-like MC3T3-E1 cells. Rather, inhibition of the L-VSCC reduces proliferation by slowing cell cycle progression. These studies, in combination with unpublished work from our lab, suggest that the L-VSCC works in conjunction with purinergic signaling to mediate cell cycle progression of MC3T3-E1 cells, and that this occurs through two distinct pathways that eventually converge within the nucleus (Jones, 2011). I further demonstrate that pulsed electromagnetic field (PEMF) stimulation of MC3T3-E1 cells has a proliferative effect in vitro that does not occur when the L-VSCC and the T-VSCC are blocked. Preliminary evidence indicates that this effect may be partially dependent on purinergic signaling, in a manner that is similar to that of mechanical stimulation (Jones, 2011). This provides valuable insight into the mechanisms by which PEMF technology has been used to treat bone fractures and supports the therapeutic potential of this approach. Furthermore, it suggests that we can expand upon and improve the way this approach is currently being used within the clinical setting by selectively targeting specific VSCC’s. In studying VSCC activation and function, we can better understand how we can treat abnormal bone loss and perhaps shed new light on their roles within other tissues.
xiv Chapter 1
INTRODUCTION
1.1 Bone Tissue Dynamics and Function in the Body
Bone is a rigid organ and a dense form of connective tissue that constitutes the vertebrate skeleton in animals (Raggatt & Partridge, 2010). Contrary to its inert appearance, bone is a complex living tissue that undergoes daily changes throughout an organism’s life. The dynamic characteristics of bone are crucial to its ability to respond to its environment and the body’s needs. These changes occur via bone remodeling, a process by which specialized cells—osteoclasts, osteoblasts, and osteocytes—work to coordinate the balance between bone formation and bone resorption. In order to understand how bone is remodeled, it is important to appreciate the cells that are involved in this process, including their cellular lineage, where they are located within bone, and how they affect bone composition. In the bone remodeling process, osteoclasts degrade bone by acidification and proteolytic digestion (Raggatt & Partridge, 2010). Osteoclasts are members of the monocyte/macrophage lineage (Figure 1.1A) and are derived from hematopoietic stem cell precursors (Soltanoff, Yang, Chen, & Li, 2009; Vaananen & Laitala-Leinonen, 2008; Yavropoulou & Yovos, 2008). In contrast, osteoblasts are derived from mesenchymal stem cells (Figure 1.1B) and form new bone by secreting osteoid into the resorption cavity to form the organic matrix of bone. Osteoid constitutes approximately 30% of all bone
1
Figure 1.1 Cellular lineage of osteoclasts, osteoblasts, and osteocytes: (A) The stages of osteoclast differentiation begin with hematopoietic stem cells in the bone marrow (Collins, Rios-Arce, Schepper, Parameswaran, & McCabe, 2017; Negishi-Koga, 2009; Soltanoff et al., 2009). These can potentially give rise to osteoclasts as they proceed down the myeloid lineage, starting with myeloid precursor cells. Macrophage colony stimulating factor (M-CSF) is a cytokine that is essential during the early stages of osteoclast development for the survival and proliferation of osteoclast precursors. Osteoclast precursors express receptor activator for nuclear factor κ B (RANK). RANK is activated by RANK ligand (RANKL), a membrane protein that is produced by osteoblasts and bone marrow stromal cells and is essential for activation of osteoclastogenesis. When several osteoclast progenitors fuse together, they form a multinucleated osteoclast precursor, which then becomes polarized in a manner which forms the ruffled membrane and the sealing zone that attaches to the bone surface for bone resorption. (B) Osteoblasts, chondrocytes, and osteocytes are all derived from a common mesenchymal stem cell precursor (Collins et al., 2017; Soltanoff et al., 2009). BMP, Wnt, b-Catenin, and RUNX2 are essential mediators of the osteoblast differentiation pathway, which is accompanied by changes in the expression level of many proteins. As mature or differentiated osteoblasts differentiate further, they transition into terminally differentiated osteocytes. The remaining subset of mature osteoblasts eventually undergo cell death via apoptosis or become inactive bone lining cells.
2 and is largely composed of type I collagen protein. Collagen proteins only comprise an estimated 10% of all bone mass, however, the elasticity of collagenous fibers plays a critical role in conferring flexibility and tensile strength to bone (Florencio-Silva, Sasso, Sasso-Cerri, Simões, & Cerri, 2015). These fibers are supplemented within the extracellular space by ground substance, the remaining organic component of bone which consists of water and various organic molecules, such as glycosaminoglycans (GAG’s), proteoglycans, and glycoproteins. The binding of the inorganic mineral salt calcium phosphate in the form of hydroxylapatite crystals to the organic portion of bone is an important factor that hardens bone and lends compressive strength to its structure (Yavropoulou & Yovos, 2008). As mature osteoblasts proliferate, they either differentiate into bone lining cells on the surface of bone or they become deeply embedded within the osteoid that they secrete and undergo a transition into terminally differentiated osteocytes. Within bone tissue, osteocytes are located within spaces called lacunae and have processes that extend outward from lacunae through canaliculi. Together, this system is termed the lacunocanalicular network, and it allows osteocytes to connect to one another as well as to blood vessels within the bone (Principles of Bone Biology - 2nd Edition, 2002). The connectivity of the lacunocanalicular network and the movement of interstitial fluid within it has led researchers to believe that the osteocytes can perceive mechanical loads exerted on the skeleton. They can in turn, communicate this signal to osteoblasts to increase bone formation in regions that experience mechanical stimuli. Figure 1.2 shows the location of osteocytes within lacunae below the bone surface, and in contrast, it also shows osteoblasts and osteoclasts, which lie on the surface itself (Le et al., 2011; Lopes, Martins-Cruz, Oliveira, & Mano, 2018; Principles of Bone Biology - 2nd Edition,
3 2002; Weatherholt, Fuchs, & Warden, 2012). Specifically, the image depicts a cross- section of trabecular bone, also known as spongy bone or cancellous osseous tissue. Trabecular bone constitutes approximately 20% of all bone in the body, while cortical bone, or compact bone, comprises the remaining 80%. The names of these bone types reflect the differences in their structure. Trabecular bone is arranged in a web-like,
Figure 1.2 Overview of bone anatomy Bone is comprised of trabecular bone, or cancellous osseous tissue, and compact bone, or cortical osseous tissue (Le et al., 2011; Lopes et al., 2018; Weatherholt et al., 2012). Trabecular bone is composed of a network of webbing structures called trabeculae. Cross-sections of trabecular bone can be seen above at three different levels of view. The most magnified level of view illustrates the location of osteoblasts and osteoclasts on the surface of trabeculae and shows that osteocytes, conversely, are located more deeply within trabeculae. Osteogenic are arranged in a similar pattern within compact bone, with osteoblast and osteoclasts on the surface of bone and osteocytes more deeply within bone; however, instead of trabeculae, these cells are located on the inside or outside of osteons. Osteons (not shown) are tightly packed cylindrical structures which lie in parallel to the long axis of the bone.
4 interconnective network that leaves spaces interspersed throughout. The spatial complexity of trabecular bone’s structure confers maximal strength with minimum mass and is optimal for resisting mechanical loads generated by functional activities such as running or jumping. Cortical bone consists of multiple microscopic columns called osteons. These columns are tightly packed together in a manner gives strength to the structure of bone and makes it ideal for protective and supportive functions within the body ("Bone Remodeling," 2010). Osteogenic tissue is able to maintain its structural integrity through its ability to remodel itself, which allows for the removal and replacement of microdamaged, old bone as well as bone that has been fractured due to injury (Owen & Reilly, 2018).
Bone remodeling is ultimately a coordinated balance between two distinct forces: catabolism (tissue breakdown) and anabolism (tissue formation) (Schindeler, 2008). The process by which these forces achieve bone remodeling (Figure 1.3) consists of five stages: quiescence or resting, resorption, reversal, formation and mineralization (Feng & Teitelbaum, 2013; Hienz, Paliwal, & Ivanovski, 2015; Kumar, 2011). During the quiescent or resting stage, bone is inactive, and transitions into the resorption stage when it senses and responds to events such as injury, mechanical loading, or low calcium levels. Activation itself is sometimes viewed as its own separate phase and occurs when osteoclast precursors are recruited and activated by molecular cues including receptor activator of nuclear kappa-B ligand (RANKL) and macrophage colony-stimulating factor (M-CSF). Consequentially, the resorption process begins as pre-osteoclasts become active, multinucleated osteoclasts, which attach to the bone surface. Carbonic anhydrase is utilized within osteoclasts to generate hydrogen ions, which are then secreted via ion pumps to demineralize bone
5
Figure 1.3 The stages of bone remodeling Osteoclasts and osteoblasts are specialized cells that lie upon the surface of bone where they perform bone remodeling by degrading and building bone, respectively ("Bone Remodeling," 2010; Owen, 2018). The bone remodeling process ultimately consists of five total phases— the resting state (quiescence), resorption, reversal, formation, and mineralization—which take place at the surface of bone.
by degrading hydroxyapatite. Additionally, several enzymes, such as acid phosphatase, cathepsin K, and matrix metalloproteinase, are produced within the osteoclast and released via matrix vesicles to act upon the bone surface where they digest the organic matrix. Osteoclasts then undergo apoptosis and are cleared away by macrophages along with other cellular debris. The reversal phase is poorly understood but is thought to aid in the coupling bone resorption to bone formation. In the bone formation stage, osteoblasts deposit osteoid to fill the cavity of the resorption pit left by the osteoclasts. Finally, during the mineralization phase, this organic portion of bone is supplemented with inorganic calcium and phosphate from matrix vesicles that
6 are secreted by the osteoblasts (Michigami, 2019). The process of fracture healing is very similar to that of typical bone remodeling, but there are several additional steps (Beamer, 2009; Wang, 2017). Following fracture, a hematoma forms around the site of injury. Within several days, a soft callus forms and angiogenesis occurs to bring blood supply from surrounding blood vessels into the site of injury. At approximately two weeks post-injury, the soft callus becomes mineralized as hypertrophic chondrocytes apoptose. Finally, at three to four weeks post-injury bone remodeling occurs, converting bone back to a fully repaired state and returning the blood supply back to normal. These processes allow bone to maintain a healthy state, through which it can continue to serve many important functions in the body.
Bone has the ability to change and adapt its structure ultimately allowing it to serve mechanical, synthetic, and metabolic functions throughout the body ("Bone Remodeling," 2010; Principles of Bone Biology - 2nd Edition, 2002). For example, bone acts to shield our vital organs, such as the brain, the heart, and the spinal cord from damage. Bone also supports the weight and locomotion of our body by working in conjunction with skeletal muscles, tendons, ligaments, and joints. These mechanical roles of bone—protection, support, and movement—all depend on the ability of bone to maintain its structural integrity by replacing old or damaged bone with new bone via the bone remodeling process. In its synthetic role, bone is responsible for producing blood cells within the bone marrow, in a process called hematopoiesis. Again, bone remodeling is crucial for this process, as osteoblasts provide niches for hematopoietic stem cells and osteoclasts induce mobilization of stem and progenitor cells through stress signaling (Kollet, Dar, & Lapidot, 2007). Bone remodeling is particularly relevant to bone’s large set of metabolic roles. Bone serves as an
7 important storage site for growth factors, oils, and minerals ("Bone Remodeling," 2010; Principles of Bone Biology - 2nd Edition, 2002). Growth factors such as insulin- like growth factor (IGF), bone morphogenic protein (BMP), and transforming growth factor (TGF) are stored in mineralized bone matrix, whereas fatty acids are stored within yellow bone marrow (Korkusuz, 2016). Bone is the storage site for important minerals such as calcium. Blood calcium levels need to be especially tightly regulated and kept within a range of 8.5 to 10.5 mg/dL (Goldstein, 1990). When blood calcium levels are low, parathyroid hormone (PTH) is released by the parathyroid glands and stimulates osteoclasts to break bone down and release calcium into the blood stream ("Bone Remodeling," 2010; Principles of Bone Biology - 2nd Edition, 2002).
Conversely, when blood calcium levels are too high, calcitonin is released from the thyroid and works to increase calcium uptake and storage from the blood by inhibiting processes of bone breakdown and stimulating processes of bone formation. Similarly, bone also helps to regulate acid-base balance by buffering the blood against excessive pH changes and absorbing and/or releasing alkaline salts. Finally, bone can also aid in detoxification by storing heavy metals and other foreign elements so that they can be removed from the blood and then gradually and safely released for excretion at a later time. The diversity of the roles bone plays throughout the body speaks to the importance of maintaining the proper balance in the bone remodeling process. When bone remodeling becomes abnormal, this can upset the ability of bone to perform these roles and can have drastic long-term effects on the health of the body as a whole. Furthermore, since bone health is inextricably tied to the health of the body as a whole, conditions of bone disease can often be difficult to correct. Treatment options which affect bone often have secondary effects on the rest of the body that may not be
8 desirable. Thus, it is crucial that we better understand the diseases that affect bone and develop solutions that address these issues in a more targeted manner.
1.2 Bone Remodeling with Age, Osteoporosis, and Current Treatment Strategies
Bone remodeling is a continuous process that occurs throughout life, with the entire skeleton being replaced approximately every ten years (Bone Health and Osteoporosis: A Report of the Surgeon General, 2004). The skeleton typically reaches peak bone mass for humans between 26 to 30 years of age (Weaver et al., 2000). Peak bone mass is marked by a shift in the balance between bone resorption and bone formation. In the earlier years of life, bone formation predominates the bone remodeling. However, as aging continues, there is a progressive decline in overall bone mass due to an imbalance in which bone resorption exceeds bone formation during remodeling. This shift towards resorption can significantly reduce bone mass over time, and if severe enough, can lead to osteoporosis. Osteoporosis, or porous bones, is defined by the World Health Organization as a reduction in bone mineral density (BMD) that is 2.5 standard deviations or more below the mean BMD value for that particular gender and age group (J. A. Kanis, 2007). Osteoporosis is characterized by reduced bone mass and compromised bone strength, which predisposes those afflicted to a heightened risk of fractures (Kanis, 2007; J. Melton, et al., 2009; L. J. Melton, Chrischilles, E. A., Cooper, C., Lane, A. W., Riggs, B. L., 1992; "Osteoporosis and Smoking Raise Fracture Risk in People With HIV," 2016). Osteoporosis poses a major public health problem and results in more than 8.9 million fractures each year worldwide. This is equivalent to one fracture every three seconds due to osteoporosis alone. In the United States, 54 million Americans are affected or at
9 high risk for osteoporosis, which amounts to $22 billion per year in national healthcare costs. A single hip fracture is estimated to cost $50,508 annually per patient in direct medical expenses alone, and the estimated lifetime risk for suffering a hip fracture is one-in-six within individuals over the age of 50. For the same subset of the population, the International Osteoporosis Foundation estimates that one out of every three women will experience at least one fracture due to osteoporosis, as will one out of every five men ("International Osteoporosis Foundation," 2017; Kanis, 2007; L. J. Melton, Chrischilles, E. A., Cooper, C., Lane, A. W., Riggs, B. L., 1992). Furthermore, in women over the age of 45, osteoporosis results in more days spent in the hospital than any other due to the limits they impose on patient mobility (L. J. Melton, Chrischilles,
E. A., Cooper, C., Lane, A. W., Riggs, B. L., 1992). Bedridden patients can lose up to 15% of their bone density in just three months and typically lose approximately 1% bone density per month on average ("International Osteoporosis Foundation," 2017; Zerwekh, 2009). Immobilization not only weakens bone, but also affects proper fracture healing, which perpetuates a vicious cycle of bone deterioration and injury (Zhong & et al., 2019). When compared to the average normal person, the risk of refracture is 3-5 times higher in individuals who have already suffered a fracture due to osteoporosis (Zhong & et al., 2019). Following a hip fracture, many patients never regain the ability to walk independently again and for those who do, two thirds of patients do not recover the functional status they had prior to the injury (Hannan, Magaziner, Wang, & al., 2001). Moreover, this diminished mobility indirectly leads to a host of health complications which further reduce the quality of life, such as pressure ulcers, urinary tract infections, blood-clotting, and a range of cardiac complications (Hannan et al., 2001). As a sum of these comorbidities, one meta-analysis indicates
10 that women and men who suffer hip fractures experience 5-fold and 8-fold increases, respectively, in the relative likelihood of death within the first three months of injury (Schnell, Friedman, Mendelson, Bingham, & Kates, 2010). A separate study found that the postoperative mortality rate for patients following a hip fracture increased to 27.3% within the first year and to 79.0% within 9 years (Panula et al., 2011). There are currently 158 million individuals worldwide at the age of 50 years or older who are at high risk for fractures due to osteoporosis, and this number is projected to double by 2040 (Odén, McCloskey, Kanis, Harvey, & Johansson, 2015). The overall financial burden of osteoporosis worldwide is projected to reach $240 billion by the same year (Odén et al., 2015). The elderly population is the fastest growing population in the world, which raises the economic and human cost of osteoporosis on a daily basis. This in turn, demands a need for improved therapies and a better understanding of bone (Nowycky, Fox, & Tsien, 1985). Abnormal bone remodeling in osteoporosis is, fundamentally, the result of an imbalance between the catabolic and anabolic processes of bone remodeling. Many current treatments for osteoporosis are antiresorptive agents, such as bisphosphonates, parathyroid hormone (PTH), and hormone replacement therapy (HRT), which attempt to address the bone loss issue by reducing the number and/or activity of osteoclasts
(Gambacciani & Levancini, 2014). HRT treatments, such as selective estrogen receptor modulators (SERMs), seek to specifically address bone loss due to decreased estrogen in menopausal women. Normally, estrogen induces stromal cells to secrete osteoprotegerin, a decoy receptor for RANK-L that prevents it from binding to its receptor, RANK, on osteoclast precursor cells (Feng & Teitelbaum, 2013). The secretion of osteoprotegerin prevents the stromal cells from differentiating into mature
11 osteoclasts, thus reducing bone resorption. HRT is typically effective at reducing bone turnover and increasing bone mineral density. However, HRT is associated with an elevated risk for serious adverse effects, including vaginal bleeding, angina, myocardial infarction, pulmonary embolism, ovarian cancer, and endometrial breast cancer (L. J. Melton, Chrischilles, E. A., Cooper, C., Lane, A. W., Riggs, B. L., 1992). The potential risk of these long-term complications may outweigh the benefits of HRT, particularly in individuals aged 60 years and older (Gambacciani & Levancini, 2014). For this reason, HRT is not regarded as a first-option treatment for osteoporosis. Aside for health complications, another major pitfall of HRT as a treatment option for osteoporosis is that it is specifically targeted to estrogen-deficient, post-menopausal women and fails to address osteoporosis in men. PTH treatment of osteoporosis also has many drawbacks and is characterized by an undesirable treatment time profile: PTH treatment is dose- and time-dependent, with low intermittent doses increasing bone formation, while high, chronic doses induce resorption (Carpinteri et al., 2010; Diez-Perez & et al., 2012) (Diez-Perez & et al., 2012). For bisphosphonate treatment, selection of the appropriate bisphosphonate is an important factor in treatment success, as are dosage and timing. Bisphosphonates are analogs of pyrophosphate that inhibit activation of enzymes that use pyrophosphate
(Fleisch, 2005). Bisphosphonates preferentially bind to calcium, and thus are drawn to bone, where they are released by osteoclasts. Once bisphosphonates have entered into osteoclasts, they disrupt the ability of osteoclasts to use carbonic anhydrase to resorb bone, as this enzyme relies on pyrophosphate for activity. This not only inhibits resorption but can also induce early apoptosis in osteoclasts. Evidence indicates that the adverse risks of bisphosphonates treatment may include osteonecrosis of the jaw
12 (ONJ) and atypical femur fractures (Khosla, 2012). The major limitation of bisphosphonates and other current antiresorptive bone treatments is that they focus primarily on reducing bone resorption and fail to adequately address the bone-building processes of bone remodeling. Osteoporosis often goes undiagnosed until a fracture occurs, which often leads to increased immobility and further risk of fracture. This speaks to the need for therapeutic interventions at the preventative level to address osteoporosis. Mechanical load has been studied as a potential means offsetting bone loss, since bone loss has been shown to be exacerbated with loss of load (Zhong & et al., 2019). It has been demonstrated previously that the VSCC’s play a pivotal role in cellular response to loss of load (J. Li, Liu, Ke, Duncan, & Turner, 2005; Owan,
Ibaraki, Duncan, Turner, & Burr, 1999; Ryder & Duncan, 2001). This demonstrates the importance of this channel in regulating bone structure and provides a candidate to study for better understanding and potential treatment of osteoporosis.
1.3 Voltage-Sensitive Calcium Channels
Voltage-sensitive calcium channels (VSCC or Cav) are expressed in both excitable and non-excitable tissues ubiquitously throughout the body, where they mediate calcium influx in response to membrane depolarizations (Catterall, 2011). These non-selective ion channels are heteromeric protein complexes, typically consisting of five subunits, α1, α2, β, δ, and γ. Variations of different subunits determine channel properties and sensitivities to certain drugs, which when taken
13
Figure 1.4 Generic model of a voltage-sensitive calcium channel and its various subunits The model depicts a typical VSCC with a magnified view of the α1 subunit structure (Iftinca, 2011; Petersohn, 2015). The α1 subunit contains the voltage-sensing element and forms the ion channel pore for all VSCC’s. The auxiliary subunits, α2, β, δ, and γ, are also shown and modify electrophysiological and pharmacological properties of VSCC’s. together, divide the VSCC’s into five distinct subtype classifications: L-, T-, N-, P/Q-, and R-type voltage-sensitive calcium channels. The basis of these classifications is inextricably tied to specific operative properties, thus comprehension of the distinctions between VSCC subtypes is critical for understanding how these channels can be manipulated with respect to their roles in biological processes as well as disease progression (Catterall, 2011; Feng, 2018). An example of a typical VSCC is illustrated in Figure 1.4. All high-voltage activated channels (HVA’s) were initially considered to be one channel, however, experimental studies in the 1970’s distinguished the L-type VSCC from the N-, P/Q-, and R-type VSCC’s due to its pharmacological sensitivity to dihydropyridines (Catterall, 2011). VSCC’s typically contain five subunits—�1, �2, β, γ, and �—though the γ subunit has not been
14 confirmed in all tissues, including bone (W. R. Thompson et al., 2011). Throughout all
VSCC subtypes the α1 subunit forms the channel pore and contains the voltage- sensitive element, while the remaining four subunits are thought to be primarily regulatory in function (Catterall, 2011). The β subunit and the �2�1 subunit have both been associated with trafficking of the channel to the membrane (Buraei & Yang,
2010). As demonstrated by our lab, the �2�1 subunit of the T-VSCC also helps to regulate purinergic signaling in osteocytes (W. R. Thompson et al., 2011). For
VSCC’s the α1 subunit is the most relevant in determining channel properties, as it contains the conducting pore and gating machinery for the channel and the majority of sites for regulation by drugs, toxins, and second messengers (Buraei & Yang, 2010;
Catterall, 2011). The α1 subunits of VSCC’s are composed of four transmembrane domains, which in turn, are each composed of six helices. Each domain is connected through cytoplasmic loops, and each domain contains re-entrant pore-forming loops (P-loops) between transmembrane segments S5 and S6. Negatively charged amino acids (glutamate or aspartate) on each of the four P-loops create the ion-selectivity filter. Voltage-dependent movement of positively charged amino acids on the S4 transmembrane segment allow it to serve as a voltage sensor that opens and closes the ion pore of the �1 subunit (Buraei & Yang, 2010). Altogether, this amounts to a size of
190 to 250 kDa for the α1 subunit (Buraei & Yang, 2010). This range in α1 subunit size can be attributed to the existence of a variety of α1 subunit isoforms.
Researchers have identified ten different isoforms of the Cavα1 subunit associated with high-voltage-activated (HVA) channels and three different isoforms associated with low-voltage-activated channels. The Cavα1 subunit mediates the majority of the channel’s electrophysiological and pharmacological properties, and
15 thus, differences among Cavα1 subunit isoforms are key to the behavior and function of the channel as a whole (Catterall, 2011; Rossier, 2016). These differences are reflected in the nomenclature, which distinguishes VSCC’s according to the isoform of the α1 subunit that their genetic sequence encodes (Catterall, 2011). The L-VSCC
�1-subunits are Cav1.1, Cav1.2, Cav1.3, and Cav1.4, and these are encoded for by the CACNA1S, CACNA1C, CACNA1D, and CACNA1F genes, respectively. T-VSCC channel subtypes are also defined by distinct �1-subunit isoforms, Cav3.1, Cav3.2, and
Cav3.3, which correspond to specific genes—CACNA1G, CACNA1H, and
CACNA1I. Pore loops between S5 and S6 are highly conserved among Cavα1 isoforms (Rossier, 2016). However, there is more divergence between isoform sequences that encode for intracellular loops, and these can significantly alter channel properties. Differences in the cytoplasmic loop linking domains I and II are particularly relevant, as this loop serves as the gate blocking mechanism for the pore
(Rossier, 2016). T-VSCC Cavα1 isoforms are characterized by a much longer intracellular loop between domains I and II, and larger conduction pore than HVA’s (M. Li, Hansen, Huang, Keyser, & Taylor, 2005; Rossier, 2016). These differences may underlie certain unique electrophysiological properties of the T-VSCC when compared with HVA’s (Catterall, 2011).
The L-type VSCC forms the largest family of VSCC’s and is prevalent throughout the body (Feng, 2018). The L-VSCC is generally characterized by high voltage activation, large single channel conductance, slow voltage-dependent inactivation, noted upregulation by cAMP-dependent protein phosphorylation pathways, and specific inhibition by Ca2+ antagonist drugs, such as dihydropyridines, phenylalkylamines, and benzothiazepines (Catterall, 2011). In contrast, the T-type
16 voltage-sensitive calcium channel (T-VSCC) is distinguished by relatively low voltage-activation, rapid inactivation, slow deactivation, and small single channel conductance. The currents induced by activation of different T-VSCC isoforms of the
α1 subunit are nearly identical to one another, however, they can be distinguished by differential expression patterns within various tissues and varying pharmacological sensitivities. For example, the Cav3.2 demonstrates a higher sensitivity to inhibition by nickel than the other T-VSCC isoforms (Rossier, 2016). The typical roles of VSCC’s include regulation of contraction, secretion, neurotransmission, and gene expression. Certain tissues tend to be dominated by particular types of VSCC’s, which are more tightly linked to the proper functioning of those tissues. In addition to bone, the T-VSCC is expressed in most excitable tissues including the heart, the central and peripheral nervous systems, endocrine tissues, and smooth muscles (Korkusuz, 2016). Expression of the L-VSCC is observed in a wide variety of cells, including bone, smooth muscle, ventricular myocytes, skeletal muscle, and dendritic cells (Feng, 2018). It has been reported that that distinct activation properties of the T-VSCC allow it to serve unique functions within tissues (Catterall, 2011). For example, activation of the T-VSCC plays a critical role in mediating the pace-making activity of the SA-node in the heart by allowing for the production of continuous rhythmic bursts. The repetitive firing of action potentials is crucial for cardiac cells and other cell types with rhythmic firing patterns such as thalamic neurons within the brain. The L-VSCC has many physiologic roles including regulation of the cardiac pace-maker, auditory transduction, visual transduction, aldosterone secretion from endocrine cells, and excitation-contraction coupling in skeletal, smooth, and cardiac muscle (Catterall, 2011). Atypical functioning of
17 VSCC’s is also associated with a wide range of disorders, including epilepsy, migraines, Lambert-Eaton Myasthenic Syndrome, and several long QT syndromes. Abnormal function and abnormal expression of VSCC’s, provide clues to their roles within the body. Abnormal overexpression of the T-VSCC in distal pulmonary artery smooth muscle cells (PASMC’s) of the lungs is linked to idiopathic pulmonary arterial hypertension (iPAH), which is characterized by obstructive hyperproliferation and apoptosis resistance (Sankhe, 2017). Within this paradigm, elevated T-VSCC expression is associated with changes in cell cycle progression and elevated expression of anti-apoptotic proteins. Overexpression of the T-type channel is also associated with a variety of cancers and may underlie increased proliferation and survival of these cells. Although the exact mechanisms of action for these processes remain unclear, abnormal increases in expression or increased kinetics of VSCC’s is often linked to increased malignant potential, as indicated by their role in cell proliferation, migration, differentiation, and survival (Fiske, 2006; Kazuyuki & et al., 2002; W. Li, Duncan, Karin, & Farach-Carson, 1997; Ohkubo & Yamazaki, 2012; C. e. a. Wang, 2015). Maintaining normal expression levels of VSCC’s is crucial for their ability to function correctly in non-cancerous tissue. Improper functioning of the L- VSCC due to mutation can have adverse cardiac and neurological effects (Zhang,
Chen, Qin, Wang, & Zhou, 2018). A G-to-A mutation in the CACNA1C gene encoding for the Cav1.2 isoform of the L-VSCC, was found in individuals with Timothy Syndrome, a disease characterized by developmental abnormalities, neurological disorders, and severe cardiac arrythmias. Small glycine residues normally confer flexibility to the I-II loop of the L-VSCC channel, however, the G-to-A mutation switches glycine to arginine, which is bulkier and impedes the I-II loop from
18 inactivating the channel as quickly as it normally would after depolarization. The subsequent prolonged action potential manifests as a prolonged QT interval on patient EKG’s, an underlies the poor survival outcomes for this disease. Other mutations in the L-VSCC have similar effects, resulting in conditions such as Brugada syndrome and short QT syndrome. The mechanisms for how these mutations alter channel properties and exert adverse effects, reveals the critical nature of proper channel kinetics with regards to specific functions within tissues. Since VSCC’s serve as a key regulator of calcium influx into electrically stimulated cells, they are a main target for therapeutic drugs for disorders including, but not limited to, pain, hypertension, and epilepsy (Belardetti & Zamponi, 2012). The first VSCC inhibitor to be used clinically was verapamil, a phenylalkylamine VSCC antagonist capable of inhibiting both the L- and T-VSCC in bone (Bergson, 2010). Verapamil exhibits a 10-fold higher affinity for binding sites on open L-VSCC channels compared to closed channels (Bergson, 2010). This state-dependency is less pronounced in T-VSCC’s with a less than 2-fold increase between closed and open channels. This may be due to differences between the conformational changes that occur with respect to each channel upon their opening and inactivation (Bergson, 2010). Data is limited on how verapamil binds to and blocks the T-VSCC, however, evidence for the L-VSCC suggests that verapamil and other prototypical phenylalkylamines (PPA’s) likely pass through the plasma membrane in their uncharged state and bind to the inner pore, blocking the channel from the inside (Bergson, 2010). This is different from the mechanism of action for dihydropyridine- derived VSCC antagonists and benzothiazepine VSCC antagonists. Most dihydropyridines, such as nifedipine, are thought to bind to a portion of S6 extending
19 into the channel pore on domains I, III, and IV (Opie, 1997). In contrast, evidence suggests that the binding site for benzothiazepines, such as diltiazem, is located on the linker region between the S5 and S6 on the IV domain (Watanabe, 1993). Mibefradil, the most widely used T-VSCC antagonist, was implemented clinically for a time as an antihypertensive agent but was eventually withdrawn as experts began to observe fatal interactions with other drugs. Initially, mibefradil was believed to be selective for the T-VSCC, but it was later found to impact HVA’s as well (L. Huang, 2004). HVA’s are not blocked by mibefradil itself, but rather, by an active metabolite of mibefradil produced by intracellular hydrolysis. In 2004, researchers worked to develop three non-hydrolysable analogues of mibefradil: NNC 55-0395, NNC 55-0396, and NNC
55-0397 (Huang, 2004). All three analogs demonstrated inhibitory effects on the T-
VSCC. At 100 μM concentrations, the highest dosage level used in the study, NNC 55-0395 and NNC 55-0397 showed inhibitory effects on HVA channels, but NNC 55- 0396 did not demonstrate a significant inhibitory effect on HVA’s at this concentration. Thus, NNC 55-0396 is characterized as an inhibitor with high selectivity for the T-VSCC, with an IC50 of 6.8 μM for the T-VSCC Cav3.1 isoform and an IC50 of > 100 μM for HVA’s in INS-1 cells. Since its development, NNC 55- 0396 has largely been used to target T-VSCC’s in cancerous tissues (Sallán, 2018).
The roles of the L-VSCC have been extensively studied within excitable tissues, and this has enabled the successful use of L-VSCC inhibitors in treating conditions such as angina, high blood pressure, and premature labor (Pontremoli et al., 2014; Songthamwat, 2018; Terry, 1982). T-VSCC inhibitors are less extensively studied, but currently show promising therapeutic potential for treating epilepsy and certain types of cancers (W. Huang, Lu, Wu, Ouyang, & Chen, 2015; Powell, Cain,
20 Snutch, & O'Brien, 2014). In 1989, Guggino et al. demonstrated that the secretion of bone matrix proteins by osteoblast-like cells is increased by BAY-K-8644, a dihydropyridine calcium channel agonist and decreased by phenylalkylamine calcium channel antagonists (Guggino, 1989). That same year, our lab observed the presence of a dihydropyridine-sensitive Ba2+ conducting channel in osteoblast-like cells, which resembled L-type Ca2+ conducting channels in other tissues (R. Duncan & Misler, 1989). Since then, there have been extensive studies on the effects of VSCC inhibitors in bone tissue. Clinical studies caution that administration of VSCC inhibitors such as verapamil, nifedipine, and diltiazem to pregnant mothers is linked to increased incidence of skeletal defects in newborn infants (Shao & et al., 2005). In animal models, VSCC inhibitors are also associated with reductions in osteogenesis, increased vertebral defects, decreased mineral apposition rates, and abnormal bone formation in animal models (Duriez, Flautre, Blary, & Hardouin, 1993; J. Li, Duncan, Burr, Gattone, & Turner, 2003; Ridings, Palmer, Davidson, & Baldwin, 1996). Overall, the deleterious effects of VSCC inhibitors with regards to bone tissue is well- established, however, there are limited therapeutic options for agonists that can activate VSCC’s. Moreover, VSCC agonists such as BAY-K-8644, are largely problematic because they do not selectively target L-VSCC’s within bone and can have drastic effects throughout the body where other L-VSCC’s are expressed. There is a need for a better understanding of how these VSCC’s operate within non-excitable tissue, and how they can be selectively activated to address issues of bone loss.
1.4 The L-VSCC in Bone As mature osteoblasts proliferate, the remaining population of older osteoblasts either apoptose or else becomes more deeply embedded within the bone matrix where
21 they terminally-differentiate into osteocytes. As the proliferative capacity of mature osteoblasts decreases, an increase occurs in markers of bone mineralization, such as bone sialoprotein, osteopontin, osteocalcin, and alkaline phosphatase (G. S. Stein & Lian, 1993). The L-VSCC is expressed in all cells of the osteogenic line, except for osteocytes, in which the channel is lost (Shao & et al., 2005). Immunohistochemical analysis has demonstrated that the L-VSCC Cav1.2 isoform is highly expressed in osteoblasts and chondrocytes in the growth plate throughout skeletal development. However, immunocytochemical studies have demonstrated that the L-VSCC isoforms
(Cav1.1, Cav1.2, Cav1.3, and Cav1.4) are not expressed in MLO-Y4 osteocyte-like cells (Shao & et al., 2005). The loss of the L-VSCC in the shift from proliferative, mature osteoblasts to non-proliferative osteocytes, leads us to hypothesize that the L- VSCC is necessary for proliferation of osteogenic cells. This is further supported by evidence indicating that inhibition of the L-VSCC in MC3T3-E1 cells leads to increased alkaline phosphatase production, suggesting that the loss of L-VSCC activation promotes differentiation into a non-proliferative state (Nishiya, Kosaka, Uchii, & Sugimoto, 2002).
Our lab has previously demonstrated that the Cav1.2 is critical to the anabolic response of bone to mechanical stimulation at both the cell and tissue levels (Fiske,
2006). Continued research into the role of the L-VSCC in mechanosensation, which will be discussed more fully in chapter 1.6 of this thesis, prompted our lab’s investigation of the roles of VSCC’s at resting conditions of stimulation. Unpublished data from our lab has indicated that the L-VSCC is essential for proliferation of osteoblasts even in the absence of mechanical stimulation (Jones, 2011). The L-VSCC selective inhibitor, nifedipine, was found to reduce osteoblast proliferation in a manner
22 that was dependent upon the time and dose of treatment. Importantly, the effect of nifedipine varied with cell density, suggesting that the reductive effect of nifedipine might be mitigated at high densities by autocrine and/or paracrine factors in the extracellular environment, such as ATP. ATP, a mitogen, has been previously shown to increase proliferation in osteoblastic cells (Jones, 2011). However, when cells were treated with nifedipine, the addition of extracellular ATP was unable to rescue proliferation, suggesting that the L-VSCC does not mediate proliferation solely through ATP release and works through a separate pathway from purinergic signaling. This establishes an important relationship between the roles of VSCC’s and purinergic signaling at rest. Additional findings from our lab indicate that these pathways could potentially play similar roles at rest as they do in response to mechanical stimulation and other types of stimulation yet to be explored, such as PEMF stimulation. Thus, further exploration of the relationship between these pathways is warranted. The calcineurin and nuclear factor of activated T-cells (NFAT) pathway represents a potential mechanism by which purinergic signaling regulates osteoblast proliferation. The calcineurin-NFAT pathway is associated with proliferation and is known to be mediated by calcium influx (Jones, 2011). Inhibition of calcineurin in MC3T3-E1 cells reduces cell proliferation while calcineurin activity is increased in the presence of ATP suggesting that ATP release may trigger the activation of the calcineurin-NFAT pathway (Jones, 2011). Parallel to purinergic signaling, it was proposed that the L-VSCC might mediate proliferation through a calmodulin pathway. Inhibition of calmodulin and inhibition of calmodulin-dependent kinase II (CaMKII) both reduce proliferation of MC3T3-E1 cells at similar levels, suggesting that the effects are potentially exerted through one shared pathway. Mechanically-induced
23 increases in calcium-calmodulin binding is blocked in MC3T3-E1 cells in the presence of the L-VSCC inhibitor, nifedipine, indicating that mechanically-induced calcium- calmodulin binding occurs through L-VSCC-mediated calcium influx. Altogether, these findings suggest that pre-osteoblast proliferation is mediated by the ability of the L-VSCC and purinergic signaling to activate two converging calcium-mediated signaling pathways. However, it is necessary for additional studies to confirm specific sets of data and for new studies to expand upon these to determine how these pathways ultimately achieve a concerted effect on proliferation. Preliminary evidence indicated that L-VSCC blockade could cause detachment of adherent MC3T3-E1 cells in culture (Jones, 2011). Thus, it was necessary to confirm that reductions in MC3T3-E1 cell number seen with nifedipine treatments were not due to removal of detached cells within the collection process. In an attempt to address the potential issue of cell detachment induced by nifedipine, the collection protocol was modified in the current study. Another limitation of previous studies was a lack of understanding of the effect of VSCC’s on cell viability and cell cycle progression. Thus, cell viability assays were performed to confirm that nifedipine was not exerting its effect on MC3T3-E1 cell number through inducing cell death. These were conducted within the current study and include secondary cell viability assays for additional validation of results. Finally, it was critically important to perform cell cycle studies as a follow-up to previous studies to determine what cellular processes were ultimately affected as a result of the convergence of the L-VSCC and purinergic signaling pathways.
24 1.5 The T-VSCC in Bone As previously described, the T-VSCC is expressed throughout the entire osteogenic lineage and remains expressed in terminally differentiated osteocytes while expression of the L-VSCC is lost (Shao & et al., 2005). Osteocytes, the mechanosensory cells of bone, are responsible for receiving mechanosensory signals and communicating these signals to other cells throughout the bone (Bonewald, 2007). Osteocytes are still poorly understood because they are more difficult to access for scientific study, however, they are approximately ten times more abundant than osteoblasts in bone (Shao & et al., 2005). Preliminary evidence has shown that knockout of the T-VSCC isoform Cav3.2 gene in mice results in a mild skeletal phenotype and significantly reduces the anabolic response of bone to mechanical load
(Kronbergs, 2011). In addition to the Cav3.2 isoform the Cav3.1 is also expressed in osteocytes; however, knockout of the Cav3.1 gene in mice results in fatality. As reviewed in chapter 1.6 of this thesis, current research supports the role of the T- VSCC in osteogenic mechanosensation; however, the differences between the roles of the L-VSCC and the T-VSCC in mechanosensation are not fully understood. Furthermore, current research does not address the role of the T-type channel in the absence of mechanical load. To gain a better understanding of the specific roles of the T-VSCC, we must first understand the unique electrophysiological properties of the T- VSCC. As with excitable cells, such as cardiac and neuronal cells, these properties may allow the T-VSCC to serve specific functions in non-excitable cells, like osteoblasts and osteocytes. The distinctive channel characteristics of the T-VSCC may play an important role in long-term cell survival by keeping a portion of T-VSCC’s incompletely activated at a nearly constant rate during resting conditions in osteogenic cells
25 (Catterall, 2011). While the L-, N-, P-, Q-, and R-types all require strong depolarizations for activation, T-VSCC’s are activated by weak depolarizations. Following weak membrane depolarization, the T-type channels are only activated for a short period of time; the “T” within the T-type channel’s name denotes this transient activation. After activation, T-VSCC’s undergo incomplete activation resulting in a “window current,” which is a sustained inward Ca2+ current carried by a portion of channels that were not completely inactivated. In osteogenic cells, the unique gating and electrophysiological properties of the T-VSCC may enable Ca2+ influx to occur at small variations of membrane depolarization across the membrane, which could sequentially activate other classes of high-voltage-activated channels (Catterall, 2011;
Shao & et al., 2005). The easy-activation and incomplete activation properties of the T-VSCC suggest that it is well-suited to play specific roles in osteogenic cells, such as mechanosensation and survival, and is crucial to the ability of osteocytes to become easily activated while deeply embedded within the bone matrix. Specifically, the unique features of the T-VSCC may allow it to serve various important cellular roles according to differences in its active states: firstly, a role in cell survival through incomplete deactivation of some channels at rest and secondly, a role in mechanosensation through complete activation under conditions of greater stimulation. This theory is supported by evidence indicating that mechanical load is necessary not only for bone growth in vivo, but also for basic bone survival ("International Osteoporosis Foundation," 2017; Zerwekh, 2009).
1.6 Mechanostransduction and VSCC-Mediated Responses in Bone The ability of bone to adapt its mass, architecture, and strength in response to mechanical forces, known as Wolff’s law, has been studied since the 19th century.
26 However, the mechanism of mechanotransduction still remains poorly understood (Chow & et al., 1993; Randall L. Duncan, 1998; Lanyon, 1982; Rubin & Lanyon, 1984; Turner, Akhter, Raab, Kimmel, & Recker, 1991). Mechanotransduction is the process by which cells detect and convert the energy from a mechanical stimulus into a biochemical signal to produce a response within the cell, and ultimately, a response at the tissue level (Randall L. Duncan, 1998). Mechanotransduction is divided into four distinct processes: (1) Mechano-coupling, which is the process whereby the energy from a physical stimulus acts upon a tissue and is converted into a form that is detectable by the cell; (2) Biochemical coupling, which is the process whereby a detectable mechanical signal is received by the cell and converted into a biochemical signal within the cell; (3) Signal transmission, which is the cascade of events whereby a biochemical signal is converted into a signal that stimulates the effector cell; (4) Effector cell response, which is how the signal-induced actions of the effector cell alter tissue form or function (Randall L. Duncan, 1998). Mechanosensitive channels exist in a variety of organisms—from mammals to bacteria—and operate within a range of diverse tissues including both excitable and non-excitable tissues. Mechanosensitive channels include stretch-activated channels and stretch-inactivated channels. In 1989, our lab observed the presence of a mechanosensitive Ba2+ conducting channel in osteoblast-like cells (R. Duncan & Misler, 1989). This mechanosensitive-sensitive channel exhibited low-voltage activation and slow inactivation, properties which closely resembled stretch-activated calcium channels in other tissues, such as neuroblastoma cells, choroid plexus epithelial cells, and endothelial cells. In the latter two types of tissues, these channels can be activated through cell swelling due to hypotonic challenge. At that time,
27 evidence suggested that stretch-activated (SA) cation channels in skeletal myocytes responded to increased membrane tension (Sachs, 1991). Polymerization of the cytoskeleton was known to affect cell stiffness, and in 1992, our lab demonstrated that the stretch-activated cation response in bone was focused by the cytoskeleton; this fit the patterns previously observed within other tissues (Randall L. Duncan, 1998). Alteration of the polymerization of the actin cytoskeleton was found to generate a six- to tenfold increase in the activity of SA cation channels. Our lab proceeded to find evidence indicating that the development of stress fibers and subsequent anchoring to the membrane at focal adhesion points was critical for the mechano-response of MC3T3-E1 cells to fluid shear (Pavalko et al., 1998). Through subsequent studies and collaborations, it was established that fluid shear, rather than substrate strain, was the main mechanism of mechano-coupling in bone (Owan, Ibaraki, Duncan, Turner, & Burr, 1999). This supported existing data, which indicated that osteoblasts required a supraphysiological amount of strain (greater than 10,000 μstrain (µɛ)) to induce an anabolic response in vitro. Typical levels of strain experienced by humans in vivo during vigorous exercise are far less (2,000 µɛ), which suggests that fluid shear is a more likely candidate for the mechano-coupling process in bone (Owan et al., 1999). Figure 1.5 illustrates the mechano-coupling process by which mechanical strain is converted into fluid shear within bone (R. L. Duncan & Turner, 1995). Under bending loads, a spatial gradient is created within bone, with compressive and tensile stresses on opposite sides; this in turn, produces a pressure gradient within the canaliculae of bone that drives the flow of interstitial fluid from regions of compression to regions of tension. As this process takes effect, fluid-shear forces are exerted upon the cell membranes of the osteocytes that are housed within the canaliculae. Downstream of
28 load
Bending force
fluid flow
c - o + - + t m - + e p - + n r - + s e - + i s - + o s - + n i - + o n canaliculi osteocyte bone lining cell
Bone at Rest Bone Bending Under Load
Figure 1.5 Mechanism of mechano-coupling in bone via fluid shear Bending forces exerted upon bone produce a spatial gradient between compressive and tensile stresses, which further yield a pressure gradient (Chen et al., 2000; R. L. Duncan & Turner, 1995). Movement of interstitial fluid is induced towards the side of tensile stress due to the pressure gradient. As the fluid moves within the canaliculae, it passes over the osteocytes embedded within, exerting forces of fluid shear across the membranes of the cells. mechano-coupling, our lab has shown that fluid shear induces cytoskeletal reorganization and increases expression of c-fos and COX-2 in a manner dependent on
2+ [Ca ]i in MC3T3-E1 cells (Chen et al., 2000). The proteins c-fos and COX-2 have both been associated with the anabolic response of bone to mechanical load in vivo
(Chen et al., 2000). Our lab later showed that both fluid shear and mechanical loading (at supraphysiological levels) can induce increases in COX-2 expression and intracellular calcium in MC3T3-E1 cells, and that these changes could be further increased by treating cells with PTH (Ryder & Duncan, 2014). Both fluid shear and
2+ PTH resulted in a rapid increase in [Ca ]i, and both stimuli were known to modulate the kinetics of mechanosensitive calcium channels (MSCC’s). It was therefore
29 2+ postulated that PTH increases the [Ca ]i response of osteoblasts to fluid shear by increasing the sensitivity of MSCC’s. It was found that the MSCC inhibitor
2+ gadolinium significantly reduced the [Ca ]i response of osteoblasts to fluid shear alone and to fluid shear with PTH pre-treated cells. L-VSCC inhibition with nifedipine
2+ also reduced the [Ca ]i response of osteoblasts to fluid shear, however, it did not reduce the response to fluid shear alone (Ryder & Duncan, 2001). This suggests that the L-VSCC plays a role in integrating these two stimuli together to achieve their
2+ combined effect on [Ca ]i. PTH is known to regulate the protein kinase A (PKA) pathway, and it was further found that treatment with a PKA-specific agonist
2+ increased the [Ca ]i response of osteoblasts to fluid shear (Ryder & Duncan, 2001).
2+ This suggests that PTH-induced increases in the [Ca ]i response of osteoblasts to fluid shear occur via PKA modulation of the L-VSCC and the MSCC (Ryder & Duncan, 2001). In a subsequent in vivo study, our lab demonstrated that mechanical load enhanced bone formation in rats and that PTH further enhanced load-induced bone formation (Fiske, 2006). However, load-induced bone formation was suppressed in rats given an oral dose of the L-VSCC inhibitor verapamil, and verapamil also suppressed the synergistic effect of PTH on load induced-bone formation. This confirms previous in vitro studies and indicates that the L-VSCC is essential for the anabolic response of bone to mechanical loading, and that this channel mediates the ability of PTH to enhance this anabolic response (J. Li et al., 2003). Several subsequent studies allowed for a better understanding of the signal transmission phase of mechanotransduction in response to fluid shear. It was demonstrated that mRNA expression of the Cav1.2 subunit is increased by mechanical stimulation and fluid shear in MC3T3-E1 cells (Puente, 2003). ATP secretion was
30 discovered to be essential for prostaglandin release, which in turn proved to be necessary for the fluid shear-induced proliferative response of osteoblasts (D. Genetos, Geist, Liu, Donahue, & Duncan, 2009). It was established that fluid shear alters gene expression in osteoblasts, in part by activating NF-κB and triggering its translocation into the nucleus. Our lab determined that the P2X7 and P2Y6 purinergic receptors are essential to fluid shear-induced degradation of the NF-κB inhibitor IκBα and nuclear accumulation of NF-κB (D. C. Genetos, Karin, Geist, Donahue, & Duncan, 2011).
This is supported by data indicating that mice with a null mutation of the P2X7 receptor exhibit significantly weaker load-bearing bones (J. Li et al., 2005). It was found that the purinergic signaling through the P2X7 receptor is essential for load- induced release of prostaglandins and the subsequent anabolic response in bone (J. Li et al., 2005). Prostaglandin E2 mediates this response in MC3T3-E1 cells by modulating F-actin stress fiber remodeling in response to fluid shear in a manner that is dependent upon PKA activation (Gong, 2013). It further established that purinergic signaling and NF-κB translocation do not involve ERK1/2 signaling within this pathway (D. C. Genetos et al., 2011). Instead, it was determined that ERK1/2 signaling is associated with the mechanoresponse of the T-VSCC channel. Evidence indicates that the T-VSCC associates with the α2δ1 subunit and that it aids in trafficking of the channel to the membrane, which is critical to ATP release and ERK1/2 activation in MLO-Y4 osteocyte-like cells in response to membrane stretch (Raggatt & Partridge, 2010). Overall, current data suggests that the L-VSCC and T- VSCC play specific roles within bone and mediate signal transmission through separate pathways that both involve purinergic signaling.
31 1.7 Mechanoelectric Response in Bone In addition to the mechanisms of mechanotransduction discussed above, researchers have sought to determine how mechanical load influences electrical fields within bone and how this contributes to the anabolic response in bone. The term “animal electricity” was coined by Luigi Galvani in 1792 to describe the ability of animals and humans to generate endogenous electrical signals (Isaacson & Bloebaum, 2010). Since then, it has been found that electromagnetic fields affect a range of organisms, from bacteria to mammals, and greatly influence tissue growth and healing (Isaacson & Bloebaum, 2010). Thus far, EMF’s have been used to treat a variety of conditions, including headaches, fibromyalgia pain, insomnia, arthritis pain, and post- surgical pain (Colbert, 2009; Strauch, Herman, Dabb, Ignarro, & Pilla, 2009). At the tissue level, EMF stimulation has been found to facilitate vasodilation and angiogenesis and is known to assist in various types of wound healing, including bones fractures (Strauch et al., 2009). Despite some therapeutic success, it is difficult to establish direct correlations between EMF treatment and its effects. There has been a lack of consistency in EMF parameters being used by researchers, and this parallels a lack of consistency in results. This has led some experts to remain skeptical on the benefits of EMF use in healthcare. In order to address the doubts around EMF use, further research must be conducted to establish which EMF parameters consistently achieve effects and to elucidate the mechanisms underlying these effects. By studying the effects of EMF’s on bone tissue, we can achieve both a better understanding of bone and a better understanding of how EMF’s can be implemented therapeutically throughout the body. Bone tissue responds to electromagnetic stimulation, and this ability works in concert with mechanotransduction (Isaacson & Bloebaum, 2010). When bone is
32 deformed by mechanical stimulation, the bone tissue generates an internal electrical field (Isaacson & Bloebaum, 2010). The ability of bone to produce stress-generated potentials, termed piezoelectricity, has been well-established and dates back to the work of Eiichi Fukada and Iwao Yasuda in the 1950’s (Fukada, 1957, 1964; Yasuda, 1977a, 1977b). Human tibia bone has shown a piezoelectric response as high as 300 mV in vivo while walking, however, piezoelectricity is known to vary depending on hydration, nutritional factors, age, gender, and the type of bone (Isaacson & Bloebaum, 2010). In the 1980’s, it was postulated that stress-generated potentials may underlie the anabolic response of bone to mechanical deformation. However, the more current consensus is that a combination of both mechanical strain and endogenous electrical currents are necessary for fully successful bone growth. It has been theorized this effect is achieved through the ability of mechanical forces to induce fluid flow, and the ability of fluid flow in turn, to produce streaming potentials in bone (Pollack, Salzstein, & Pienkowski, 2011). Streaming potentials are potentials generated when liquid is forced through a porous membrane or capillary. As the liquid passes through, oppositely charged layers of the electrical double layer are separated, which results in the development of potential between the two sides of the membrane. Thus, streaming potential can be viewed as the reverse process of electroosmosis: In electroosmosis fluid flow occurs due to applied potential, whereas, in streaming potential, the potential is generated due to the flow of fluid. It is possible to measure streaming potentials using reversible electrodes, and there is evidence from such studies to suggest that stress-generated streaming potentials occur in bone (Pollack et al., 2011). In this manner, bone is thought to generate electrical fields in response to mechanical forces (Spadaro, 1998). Until recently, the therapeutic potential of
33 electrical stimulation in bone was viewed with skepticism by the medical community for the same reasons listed above: a lack of consistency in trial methods, inadequately sized subject pools, and wide variation in results. Some skepticism remains today, however, electrical stimulation (ES) has begun to be more widely implemented and has seen some success. For example, a recent study found that electrical stimulation can increase union rates for non-union fractures (Kooistra, Jain, & Hanson, 2009). There is still limited data regarding the mechanisms for how electrical stimulation produces an anabolic response in bone, but current evidence suggests that electrical stimuli increase proliferation and differentiation of osteoblasts and increase paracrine and autocrine growth factors in osteoblasts in a manner dependent on calcium signaling (Kooistra et al., 2009). Studies conducted in other tissues further support the theory that this effect may be exerted through activation of VSCC’s. If we can better understand the processes that govern the anabolic response of bone to ES, it will pose massive therapeutic potential in bone, as it has been recently used in other tissues to selectively target specific VSCC’s (Buckner, Buckner, Koren, Persinger, & Lafrenie, 2015). It has been postulated that stimulation via electromagnetic fields (EMF’s), represents a vastly untapped resource in healthcare that could potentially rival that of pharmacological interventions (Buckner et al., 2015). By altering the intensity, frequency, duration, and pattern of EMF’s, we can manipulate EMF’s in a manner that is similar to modifying the molecular structure of a pharmacological agent (Buckner et al., 2015). Like pharmacological agents, EMF stimulation can be modified to target specific cells within specific tissues to achieve specific outcomes. A recent study concluded that exposure of cancerous cells to time-varying electromagnetic fields can
34 inhibit cancerous cell growth through inhibition of T-VSCC’s (Buckner et al., 2015). Exposure to Thomas-EMF—a low-intensity, frequency-modulated (25-6 Hz) EMF pattern—was found to inhibit cell proliferation without inducing cell death, and of the cell lines tested this effect was only observed in malignant cancer cell lines. The anti- proliferative effect likely occurred via the T-type channel since Thomas-EMF exposure also blocked calcium influx through the T-VSCC; this became reduced when the T-VSCC was blocked via pre-treatment with channel-specific inhibitors, but not when the L-VSCC was blocked. Further data indicated that the EMF-induced changes in cytoplasmic Ca2+ via the T-VSCC, were linked to changes in cell cycle progression and cyclin expression, which is consistent with their normal roles in other tissues
(Buckner et al., 2015). These studies suggest that specific VSCC’s can be targeted within tissues, to selectively stimulate specific functions and better achieve therapeutic outcomes. Similar techniques may be utilized within bone, as we expand our understanding of the roles VSCC’s play in bone. As previously discussed, evidence suggests that common patterns may exist between the pathways VSCC’s utilize at rest in bone and the pathways they utilize in response to mechanical stimulation in bone; electrical activation of VSCC’s may also share these common pathways and may allow for the selective targeting of specific functions based upon the distinct activation properties of VSCC’s. Thus, it may be possible to activate specific VSCC’s within bone to achieve specific outcomes without affecting VSCC’s in neighboring tissues.
35 Chapter 2
MATERIALS AND METHODS
2.1 Pharmacological Agents
Nifedipine (Sigma-Aldrich, St. Louis, MO) and NNC55-0396 (Sigma-Aldrich) were used at the following concentrations: 1 μM nifedipine, 2.5 μM nifedipine, and 5 μM nifedipine (from 5 mM stock in ethanol), 1 μM NNC, 5 μM NNC, and 10 μM NNC (from 10 mM stock in dimethyl sulfoxide; DMSO).
2.2 Cell Culture The mouse calvarial osteoblast-like cell line, MC3T3-E1 (subclone 14) was purchased from American Type Culture Collection (Manassas, VA). Cells were cultured in minimum essential medium, alpha modification (α-MEM, Sigma-Aldrich) supplemented with 10% fetal bovine serum (FBS; Lot # F12C18D1; Atlas Biologicals, Fort Collins, CO), 100 μg/ml streptomycin, 100 IU/mL penicillin (Hyclone, Logan,
UT), and 0.22% w/v sodium bicarbonate and maintained at 37ºC in a 5% CO2 environment. Cells were passaged at 80% confluence. Following seeding, cells were allowed to incubate for 48 hours prior to experimentation. Prior to experimentation, all seeding surfaces were coated with type I collagen from rat tail (Sigma-Aldrich). For this process, a thin layer of 100 µg/mL collagen in 0.02 N acetic acid was applied to completely cover the seeding surface. After an incubation period of 1 hour at room temperature, collagen was removed, and surfaces were exposed to UV light overnight. Following UV exposure, surfaces were rinsed with Dulbecco’s phosphate-buffered
36 solution without calcium or magnesium (DPBS; Corning, Corning, NY) to remove any residual acid. Unless otherwise noted, incubation conditions for all experiments were maintained at the same cell culture conditions as listed above, and all cells were maintained in the fully supplemented (FS) media as listed above.
2.3 Proliferation Assays For the generation of growth curves of MC3T3-E1 pre-osteoblasts, cells were seeded at a density of 2500 cells/cm2 on 6-well plates (Corning) coated with collagen as previously described and were treated with antagonists. All treatments—5 μM nifedipine, 10 μM NNC, and combined treatment of 5 μM nifedipine and 10 μM NNC —were added via FS media. FS media was also included as the negative control treatment condition. All conditions were replaced with fresh treatment every 24 hours, and cell number was measured at the 24 hour, 48 hour, 72 hour, and 96 hour time points. At the appropriate time points of cell collection, the supernatant from each well was collected via serological pipettes and saved in conical tubes at room temperature from later use. To induce cell detachment, 200 μL of cell-harvesting solution was added: 0.25% trypsin, 0.1% EDTA in Hank’s Balanced Salt Solution (HBSS) without calcium, magnesium, and sodium bicarbonate (Corning). Cells were allowed to completely detach (for approximately 5 minutes) before the addition of 1000 μL of FS media to neutralize cell-harvesting solution. Harvested cells were pooled with the supernatant from their respective samples, and the pooled solution was centrifuged at 1000 rpm for 5 minutes to isolate cell pellets. Cell pellets were resuspended in FS media, and cell number was subsequently determined by counting via a hemocytometer. This experiment was repeated in triplicate with four separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare
37 means in order to determine statistical significance between experimental treatment groups and time-matched negative controls.
For dose response assessment of the effects of nifedipine (1 μM, 2.5 μM, and 5 μM) and NNC (1 μM, 5 μM, and 10 μM) on proliferation of MC3T3-E1’s, cells were prepared, collected, and assessed in the manner described above. This experiment was repeated in triplicate with three separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in order to determine statistical significance between experimental treatment groups and time-matched negative controls.
2.4 Annexin-V Cell Viability Assays For the measurement of cell viability, MC3T3-E1 pre-osteoblasts were seeded at a density of 2500 cells/cm2 on 100 mm tissue culture plates (Stellar Scientific, Baltimore, MD) coated with 100 μg type I collagen as previously described. Following seeding, cells were allowed to incubate for 48 hours prior to administering experimental treatments. For nifedipine dose-response trials, cells were collected and analyzed at the 24 hour and 48 hour treatment time points for each of the following treatment conditions: negative control (FS media), 1 μM nifedipine, 2.5 μM nifedipine, and 5 μM nifedipine. All conditions were added via FS media and were refreshed daily. For cell collection, supernatant was collected and saved prior to the collection process. Then cells were rinsed once with 200 μL DPBS without calcium or magnesium, and residual solution was pooled with the respective supernatant. To induce cell detachment, 1.5 mL of cell-harvesting solution—0.25% trypsin, 0.1% EDTA in HBSS without calcium, magnesium, and sodium bicarbonate—was added to each dish and cells were allowed to completely detach, for approximately 5 minutes.
38 An additional 1.5 mL of FS media was added to each dish, and the resulting solution was then pooled with the respective supernatant previously collected. The pooled solutions were centrifuged to isolate cell pellets. Following centrifugation, cells were re-suspended in 2.0 mL microcentrifuge tubes (Thermo Fisher Scientific, Waltham, MA) tubes via a solution of DPBS with calcium and magnesium and containing the following staining agents: 108 μL 1X-annexin-V binding buffer (Thermo Fisher
Scientific) with 1 μL of PI (100 μg/mL) (Thermo Fisher Scientific) and 1 μL of Alexa- Fluor 488 annexin-V conjugate (Thermo Fisher Scientific). Cells were allowed to incubate in staining solution in the dark at room temp for 15 minutes. Then samples were placed on ice. Immediately prior to cytometric analysis, 400 μL of 1X-annexin-V binding buffer was added to each sample. Samples were gently mixed via micropipette and were then pipetted into round-bottom polystyrene Falcon® test tubes through the
35 μM nylon mesh of the accompanying strainer caps (Corning). Percentages of viable, early apoptotic, late apoptotic, and necrotic cells were determined via flow cytometry (Acea Bioscience Novocyte Flow 2060R). A positive control for apoptosis,
2000 μM H2O2, was also utilized during the set-up process at each time point in order to establish proper gating parameters during cytometric analysis. Prior to analysis of experimental conditions, gating parameters were set using the following control conditions: Negative control unstained, negative control stained with PI, negative control stained with annexin-V, negative control stained with PI and annexin-V, positive control unstained, positive control stained with PI, positive control stained with annexin-V, positive control stained with PI and annexin-V. All negative and positive controls used for gating were added via FS media and were refreshed daily throughout the treatment time course. The data shown (Fig. 2.1A-B) are representative
39 images cytometric analysis of cells, where Figure 2.1A depicts cells treated with negative control conditions and stained for both PI and annexin-V, whereas Figure 2.1B shows cells treated with positive control conditions and stained for both PI and annexin-V. This experiment was repeated in triplicate with three separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in Back to Table of Contents (Link) order to determine statistical significance between experimental treatment groups and
time-matched negative controls.
A B
FigureFigure 2.1. Representative 2.1 Representative images images of annexin of annexin-V -assayV assay gating gating A-B: RepresentativeA- B:images Representative of Annexin ima-Vges and of AnnexinPI staining-V and measured PI staining via measured flow cytometry via at 24 hours. For gating, quadrantflow cytometry 3 (Q2 at-3) 24 represents hours. For cellsgating, negative quadrant for 3 (Q2 annexin-3) represents-V and PI fluorescence cells negative for annexin-V and PI fluorescence (viable cells), quadrant (viable cells), quadrant 4 (Q2-4) represents cells negative for PI fluorescence and positive for 4 (Q2-4) represents cells negative for PI fluorescence and positive for annexin-V fluorescenceannexin (early-V fluorescence apoptotic (earlycells), apoptotic quadrant cells), 2 (Q2 quadrant-2) represents 2 (Q2-2) cells positive for PI fluorescence and positiverepresents for cells annexin positive-V fluorescencefor PI fluorescence (late andapoptotic positive cells), for annexin and quadrant-V 1 (Q2- 1) represents cells fluorescencenegative for (late annexin apoptotic-V fluorescence cells), and qua anddrant positive 1 (Q2- for1) represents PI fluorescence cells (necrotic cells). negative for annexin-V fluorescence and positive for PI fluorescence (necrotic cells).
40
For the measurement of the effects of 10 μM NNC on MC3T3-E1 cell viability at 24 and 48 hours, cells were prepared, collected, and assessed in the manner described above. This experiment was repeated in triplicate with three separate passages of cells. All data are reported as mean ±SEM. Student’s t tests were used to compare means in order to determine statistical significance between experimental treatment groups and time-matched negative controls.
2.5 Acridine Orange Cell Viability Assays As a secondary measurement of cell viability, MC3T3-E1 pre-osteoblasts were seeded at a density of 5000 cells/cm2 on 100 mm tissue culture plates (Stellar Scientific) coated with 100 μg type I collagen as previously described. A higher seeding density was utilized to ensure enough images would be acquired of cells during microscopy. Following seeding, cells were allowed to incubate for 48 hours prior to experimentation. For examination of the effects of L-VSCC inhibition on cell viability, cells were treated with either 5 µM nifedipine or negative control (FS media) and cell viability was assessed at 24 at 48 hour time points. The nifedipine treatment condition was added via FS media, and all conditions were refreshed daily throughout the treatment time course. To induce apoptosis in positive control conditions, cells were incubated at 43°C for 90 minutes and then incubated at 37°C for 24 hours. Prior to the collection of treated cells at each time point, the supernatant was collected and saved to be pooled later with cells harvested from its respective sample. Cells were resuspended in 50 μL DPBS with calcium and magnesium and stained with 4 μL ethidium bromide/acridine orange (EB/AO) stain (100 µg/mL EB and 100 µg/mL AO in DPBS with calcium and magnesium). Cells were incubated for 5 minutes at room temperature in staining solution, then 10 µL of each cell suspension was added to a
41 Back to Table of Contents (Link) criteria (Fig. 2.2) were used to assess cell viability, which was tabulated by two independent observers and averaged to yield the results shown.
microscope slide for analysis. Analysis was achieved via fluorescent microscopy utilizing a VWR® Inverted Fluorescent Microscope at the 20x objective, with a laser
wavelength of 488 nm for excitation. Both fluorescent and morphological criteria
Live (Round and Apoptotic uniformly (Membrane green) blebbing)
Apoptotic Apoptotic (Red dotting) (Chromatin condensation)
Necrotic (Intense red Apoptotic staining (fragmentation) throughout)
Figure 2.2 Criteria for assessing cell viability via acridine orange assay Representative images of the morphological and fluorescent criteria used Figure 2.2. Representativeto assess cell images viability of acridineof MC3T3 orange-E1 cells assay stained criteria with foracridine cell viability: orange Commented [2]: I need to re-inert this figure so that its Representative imagesand ethidium of the morphological bromide. and fluorescent criteria used to assess cell viability no longer blurry of cells stained with acridine orange and ethidium bromide. (Figure 2.2) were used to assess cell viability, which was tabulated by two independent observers and averaged to yield the results shown. This experiment was repeated in triplicate with three separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in order to determine statistical significance between experimental treatment groups and time-matched
negative controls. For the measurement of the effects of 1 μM NNC on MC3T3-E1
42 cell viability at 24 hours, cells were prepared, collected, and assessed in the manner previously described above. In order to ensure sufficient cells would remain for analysis, a lesser dose of NNC, 1 μM, was administered in this assay compared to that of the annexin-V assay, 10 μM, and treatment time was limited to 24 hours. This experiment was repeated in triplicate with three separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in order to determine statistical significance between experimental treatment groups the negative control.
2.6 Cell Cycle Assays For the measurement of cell cycle progression of MC3T3-E1 pre-osteoblasts, cell cycle studies were first performed in order to establish serum starvation as an effective positive control for G1/G0 phase arrest in MC3T3-E1. Subsequent studies were then performed using serum starvation to pre-synchronize MC3T3-E1 cells in the G1/G0 phase prior to treatment with VSCC antagonists. For the validation of serum starvation as an effective positive control for G1/G0 phase arrest, MC3T3-E1 cells were seeded at a density of 2500 cells/cm2 on 100 mm tissue culture plates (Stellar Scientific) coated with 100 μg type I collagen as previously described. Following seeding, cells were allowed to incubate for 48 hours in FS media prior to any treatment. A subset of cells was fixed for analysis at this time point to establish the cell cycle profile prior to synchronization. For the next 72 hours, all remaining cells underwent serum starvation in basal medium 0% FBS, 100 U/ml of penicillin, 100 μg/ml of streptomycin, and 0.22% w/v sodium bicarbonate. Cells were harvested at the 24 hour, 48 hour, and 72 hour time points of treatment, and media was replaced with fresh basal media for any remaining samples. At the appropriate time
43 points of cell collection, the supernatant from each well was collected via serological pipettes and saved in conical tubes at room temperature from later use. Then cells were rinsed once with 200 μL DPBS without calcium or magnesium (Corning), and residual solution was pooled with the respective supernatant. To induce cell detachment, 1.5 mL of cell-harvesting solution—0.25% trypsin, 0.1% EDTA in HBSS without calcium, magnesium, and sodium bicarbonate (Corning)—was added to each dish and cells were allowed to completely detach, for approximately 5 minutes. An additional 1.5 mL of FS media was added to each dish, and the resulting solution was then pooled with the respective supernatant previously collected. The pooled solutions were centrifuged to isolate cell pellets, and following centrifugation, cells were re- suspended in 1 mL of fully supplemented media containing DNAse (1 μg/mL) for approximately 15-20 minutes. Cells were centrifuged at 5,000 rpm for 5 minutes and re-suspended in 500 μL DPBS without calcium and magnesium, prior to the addition of 4.5 mL of 70% ethanol to achieve fixation. Cells were fixed for 4 to 8 days at -20 degrees Celsius, with gentle vortexing daily to prevent cell clumping. Following fixation, cells were washed twice with DPBS with calcium and magnesium, centrifuged, and re-suspended in 500 μL DNA staining solution, containing the following in DPBS with calcium and magnesium: RNAse (200 μg/mL), propidium iodide (PI) (20 μg/mL), and Triton X-100 (0.03% v/v). Cells were stained in DNA staining solution for 30 minutes at room temperature, then kept on ice to await analysis via flow cytometry (Acea Bioscience Novocyte Flow 2060R). A minimum of 50,000 cells were assessed for each treatment condition, and samples were analyzed via NovoExpress software utilizing the Watson pragmatic model to determine cell cycle distribution profiles. This experiment was repeated in triplicate with three
44 separate passages of cells. All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in order to determine statistical significance between experimental treatment groups the negative control. For the measurement of VSCC effects on cell cycle progression of MC3T3-E1 pre-osteoblasts, cells were seeded at a density of 2500 cells/cm2 on 100 mm tissue culture plates (Stellar Scientific) coated with 100 μg type I collagen as previously described. Following seeding, prior to any pre-treatment or experimental treatments, cells were allowed to incubate for 48 hours in FS media. A subset of cells was fixed at this time point, and then the process of pre-synchronization was initiated for all remaining cells via serum starvation in basal media. Cells were treated, harvested, and analyzed as described above to document successful pre-synchronization in G1/G0 phase prior to experimental treatment. After 72 hours of serum starvation treatment, remaining cells subsequently underwent 48 hours additional hours of experimental treatment. Cells were treated with either FS media, as a negative control, or 5 μM nifedipine added in FS media; treatments were refreshed every 24 hours. To observe the process of desynchronization, cells were collected for cell cycle distribution analysis at the following treatment time points: 3 hours, 6 hours, 12, hours, 18 hours, 24 hours, and 48 hours (data not shown for 3-6 hour time points). This experiment was conducted once in triplicate with three separate passages of cells. As such, statistical analysis has not been performed on the results.
2.7 Creating a PEMF Generator MC3T3-E1 pre-osteoblast cells were exposed in vitro to a PEMF bioreactor, which was developed using a 33220 arbitrary waveform generator (Agilent, Santa Clara, CA) as a voltage source and a pair of 3B Scientific 51000611 Hemholtz coils to
45 create the EMF (3B Scientific, Tucker, GA). An Intuilink waveform editor application (Agilent) was used to create the specific pulsed waveform necessary, while a Lepai® LP2020A+ high stereo amplifier was used to amplify the source signal, and a PCE
Figure 2.3 Schematic of a PEMF bioreactor An arbitrary waveform generator, digital amplifier, and a pair of Helmholtz coils were used to stimulate MC3T3-E1 via pulsed electromagnetic fields. To receive PEMF stimulation, cells were placed upon the platform between the Helmholtz coils. MFM 3000 AC/DC magnetic meter (PCE Instruments, Jupiter, FL) was used to assess the strength of the resulting magnetic field. The pulsed waveform was made to have a burst width of 5 ms, a pulse width of 0.3 ms, a pulse wait of 0.02 ms, and a burst wait of 15 ms, repeated at a frequency of 45 Hz. A platform, designed in Solidworks, was made to hold a 96-well cell culture plate between the pair of Helmholtz coils in an equidistant manner (Figure 2.3). The platform and its support frame were made from acetal plastic using machines in the University of Delaware’s College of Engineering Student Machine Shop. The magnitude of the uniform magnetic field between Hemholtz coils was defined using a derivation of the Biot-Savart law, shown in Equation 1. Equation 2 shows how Equation 1 has been rearranged to solve for the
46 Back to Table of Contents (Link) colorimetric changes, and absorbance values were subsequently assessed using a Synergy™ H1 plate reader from Biotek.
Back to Table of Contents (Link)
colorimetric changes, and absorbance values were subsequently assessed using a Synergy™ H1
plate reader from Biotek.
current, I, needed to generate specific magnetic fields. A current of 0.520 A was calculated for the magnetic field, 2.2 mT, used in this study. Additionally, three different currents have been calculated for different applied magnetic fields to be used
in future studies, shown in Table 2.1. Figure 2.3. Schematic of a PEMF generator
! # = !" ##$% (Equation 2.1) " &
Figure 2.3. Schematic of a PEMF generator'& & = ! (Equation 2.2) $ " ) $ % # ! # = !" ##$% (Equation 2.1) " &
Table 2.1. Calculations of currents necessary'& to generate four different magnetic fields & = ! (Equation 2.2) $ " ) $ Table 2.1 Calculations of currentsB = 1mT% # necessaryB to= 2.4mTgenerate four different magnetic fields I = 0.236A I = 0.567A
Table 2.1. Calculations of currentsB = necessary 2.2mT to generateB = 3mT four different magnetic fields I = 0.520A I = 0.709A B = 1mT B = 2.4mT I = 0.236A I = 0.567A
B = 2.2mT B = 3mT I = 0.520A I = 0.709A
2.8 Proliferation Assays for Cells Exposed to PEMF
MTS assays were used to measure the activity of the mitochondrial reductase
enzyme, as represented by increased formation of formazan product in MC3T3-E1 cells. Increased absorbance due to formation of formazan in viable cells enabled colorimetric sensitive quantification of cells, which was used as a measure of proliferation. MC3T3-E1 cells were seeded at a density of 2,500 cells/cm2 on type I
47 collagen-coated 96-well plates (Corning). Cells were treated with Nifedipine, NNC, or negative control (FS media) and cell proliferation was assessed at 24, 48, 72, and 96 hour time points. Antagonist treatment conditions were added via FS media, and all conditions were refreshed daily prior to PEMF exposure. For conditions of PEMF treatment, cells were exposed to a PEMF at a magnitude of 2.2 ± 0.5mT for two hours per day across the four-day time course. For each condition, 16 wells were assessed at each time point. The CellTiter96™ AQueous One Cell Proliferation assay (Promega, Madison, WI) was used to produce colorimetric changes, and absorbance values were subsequently assessed using a Synergy™ H1 plate reader from (Biotek, Winooski, VT). This experiment was repeated in triplicate with five separate groups of cells; however, the same passage number was used for all cell groups, as there is evidence that the proliferative effect of PEMF can be altered by the maturation stage of osteoblasts (Diniz, 2002). All data are reported as mean ± SEM. Dunnett’s tests were used to compare means in order to determine statistical significance between experimental treatment groups the time-matched negative controls.
2.9 Fluorescent Microscopy Assays for Cells Exposed to PEMF Fluorescent microscopy assays were used to measure the release of ATP by MC3T3-E1 cells in response to PEMF exposure. MC3T3-E1 cells were seeded at a density of 4000 cells/mL on 35 mm MatTek™ petri dishes with 14 mm microwells coated with type-1 collagen (Thermo-Fisher Scientific). Cells were incubated for two days in FS media, then treated with 25 µM quinacrine for 30 minutes. Cells were rinsed twice with HBSS with calcium and magnesium (Sigma-Aldrich). Prior to treatment, 1 mL of HBSS with calcium and magnesium was added to each dish. Then cells were exposed to PEMF at a magnitude of 2.2 ± 0.5mT for 0, 1, 2, 3, 4, or 5
48 Back to Table of Contents (Link)
A
B
Figure 2.4 Representative images of quinacrine staining analyzed via C fluorescent microscopy Representative images showing quinacrine-stained MC3T3-E1 cells (A) with no PEMF stimulation, (B) with 1 minute of PEMF exposure (2.2mT ± 0.5mT). minutes. Negative control cells were not stained with quinacrine and were not exposed to PEMF. Following PEMF exposure, cells were imaged via a Zeiss Axio Scope A1 microscope at the 20x objective, using 476 nm wavelength for excitation. The fluorescent intensities were subsequently analyzed with respect to grayscale using a procedure in Image J developed at the University of Chicago by Christine Labno (Labno, 2017). Representative images of are shown for quinacrine-stained MC3T3-E1 cells without PEMF stimulation (Figure 2.4A) and with PEMF stimulation (2.4B).
49 This experiment was conducted once in triplicate with three separate passages of cells. As such, statistical analysis has not been performed on the results.
50 Chapter 3
RESULTS
3.1 The L-VSCC is necessary for proliferation of MC3T3-E1 cells
Previous unpublished findings established that the L-VSCC was central to proliferation of MC3T3-E1 cells at resting conditions (Jones, 2011). However, these studies did not include NNC as a comparative measure to nifedipine. Furthermore, these studies implemented a collection process that potentially failed to account for detachment of cells due to nifedipine exposure. Upon re-examination of these studies with new protocols in place (see chapter 2.3), the following was found: Over four days of treatment for each of condition—negative control, 5 μM nifedipine, 10 μM NNC, or 5 μM nifedipine and 10 μM NNC—results showed that osteoblast proliferation was attenuated by daily treatment with all antagonist conditions compared to the negative control (Figure 3.1). At days 24 hours, 48 hours, 72 hours, and 96 hours after the initial treatment with nifedipine, osteoblasts exhibited significant reductions in proliferation compared to the control, showing an average reduction in cell number of approximately 45%, 68%, 80%, 76% respectively. Osteoblast number was also significantly reduced with NNC treatment at all time points, however, cell number was reduced below the initial seeding number within the first day of treatment with NNC, suggesting that cell death was induced. Cell number was reduced to a similar level to that of NNC treatment when treated with a combination of nifedipine and NNC. It was further found that nifedipine also inhibits proliferation in a dose-dependent manner
51
Figure 3.1 The L-VSCC is necessary for proliferation of MC3T3-E1 cells Proliferation of MC3T3-E1 cells in the presence of 5 μM nifedipine, 10 μM NNC, 5 μM nifedipine, and 10 μM NNC, or control media at 24, 48, 72 and 96 hours. (as compared to time-matched negative controls: ap≤0.05, cp≤0.005, ep≤0.0005, fp≤0.0001)
52 Back to Table of Contents (Link)
A B
C D
Figure 3.2 Inhibition of the L-VSCC reduces proliferation of MC3T3-E1 cells in FigureFigure 3.2.3.2 InhiInhibitiona bitiondose- dependentof theof the L- VSCCL- VSCCmanner reduces reduces proliferation proliferation of MC3T3 of MC3T3-E1 -cellsE1 cells in a dosein - dependent manner:a dose -Dosedependent-dependent manner: effects Dose of nifedipine-dependent on effectsnumber of of nifedipine MC3T3-E1 on cells at (A) Dose-dependent effects of nifedipine on number of MC3T3-E1 cells at 24 hours, (B) 48number hours, of (C) MC3T3 72 hours,-E1 andcells (D) at 96(A) hours. 24 hours, (n=3) (B) (as 48 compared hours, (C) to time72 hours,-matched negative controls:and(A) b (D)p24≤0. hours,9601) hours. (B) (as 48 compared hours, (C) to 72time hours,-matched and negative (D) 96 hours.controls: (as compared b btop≤ time0.01)- matched negative controls: p≤0.01)
(Figure 3.2). Overall, this suggests that the L-VSCC plays a role in proliferation
whereas the T-VSCC potentially drives cell survival.
3.2 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells
Following proliferation studies, an annexin-V assay was performed in order to confirm that the reduction in cell number64 observed with nifedipine treatment in proliferation studies was not due to reductions in cell viability. Cells were stained with propidium iodide and annexin-V, each conjugated to fluorophores, and then fluorescence was measured via flow cytometry. Cells positive for propidium iodide
53
A
B
Figure 3.3 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells (as measured via an annexin-V assay) MC3T3-E1 cells were treated with either negative control (media) or NNC 10 μM. Cells were collected at 24 and 48 hours, and cell viability was assessed via annexin-V assays and cytometric analysis. Cell viability levels are represented as the percentage of 50,000 total cells negative for annexin-V and PI fluorescence at 24 hour (A) and 48 hour time points (B), when normalized to the time-matched negative control.
(PI) indicate that the cell membrane has become compromised and suggest cell death, since PI cannot normally pass through the membranes of healthy, viable cells. Cells positive for annexin-V, indicate early stages of apoptosis are occurring, since annexin-
V binds to phosphatidylserine which becomes66 externalized on the membrane during
54 the early stages of apoptosis. Late apoptosis is indicated for cells positive for both annexin-V and PI, whereas viable cells are indicated by cells negative for annexin-V and PI. When treating for 24 and 48 hours with three doses of nifedipine, 1 μM, 2.5 μM, and 5 μM, there were no changes in the percentage of viable cells compared to control at any dose (Figure 3.3A-B). These results were further confirmed through an acridine orange/ethidium bromide apoptosis assay. Cells were stained with acridine orange and ethidium bromide and analyzed via fluorescence microscopy to determine cell viability based on fluorescent and morphological criteria (see chapter 2.5 for details).
Figure 3.4 L-VSCC inhibition does not reduce cell viability of MC3T3-E1 cells (as measured via acridine orange assay) MC3T3-E1 cells were treated with either negative control (media), nifedipine 5 μM, or positive control (heat-induced apoptosis). Then, acridine orange assays were used to quantify levels of cell viability, which are represented here as the percentage of total MC3T3-E1 cells negative for fluorescent and morphological changes associated with apoptosis and necrosis. All viability percentages were normalized to those of the time-matched negative control. (as compared to the negative control: ep≤0.0005)
55 Viable cells were categorized as those that were negative for fluorescent and morphological characteristics of necrosis and apoptosis. Treatment with 5 μM nifedipine for 24 hours (Figure 3.4) and 48 hours (data not shown) demonstrated no change in the percent of viable cells compared to the control; however, the positive control demonstrated a significant difference between the negative control as well as nifedipine. Both cell viability assays are consistent with one another in their results and demonstrate that nifedipine does not induce cell death, suggesting that the reduction in cell number observed in proliferation assays is due to alternative mechanisms.
3.3 Inhibition of the L-VSCC causes cell cycle slowing Previous research from our lab has demonstrated that the L-VSCC and purinergic signaling likely regulate proliferation through two pathways which converge within the nucleus (Jones, 2011). However, the ultimate effects of these pathways beyond their convergence in the nucleus was not established, and the mechanism through which cellular proliferation is altered is still unknown. In order to test if L-VSCC inhibition affects proliferation through altering cell cycle progression, cells were stained for DNA content with propidium iodide (PI), and fluorescence was measured via flow cytometry to determine cell cycle distribution in each phase. Prior to experimental treatment, MC3T3-E1 cells were synchronized in G1/G0 phase via serum starvation in MEM-α media with 0% FBS for 72 hours. Representative images (Figure 3.5A) of cell cycle distribution profiles and quantification (Figure 3.5B) of the percentage of cells in G1/G0 phase, indicate that cells were successfully synchronized in the G1/G0 phase. Approximately 75% of cells were synchronized in the G1/G0 phase before treatment conditions were implemented. Cell cycle distribution was
56 Back to Table of Contents (Link)
exposed to 5 μM nifedipine remained synchronized in G1/G0 phase for the first 12 hours of
recovery. At 18 hours, cells began to recover and desynchronize from the G1/G0 phase in both
conditions, however, the 18 hour time window shows that a larger proportion of cells remained
synchronized
A
B
Figure 3.5 Serum starvation induces synchronization of MC3T3-E1 cells in the Figure 3.4 SerumG1/G0 starvationphase: induces synchronization of MC3T3-E1 cells in the G1/G0Cells were phase: treated Cells with were basal treated media with (0% basal FBS) media across (0% 24, FBS) 48, and across 72 hour24, 48,time and points. 72 hour At each time timepoints. point, At eachcells timewere point, collected cells and were fixed collected for and fixedsubsequent for subsequent cytometric cytometric analysis. analysis.Cell cycle Cell distribution cycle distribution was measured was via measuredflow cytometry via flow for cytometrya minimum for of a 50,000 minimum per ofsample. 50,000 Representative per sample. Representativeimages of the cell images cycle of profiles the cell from cycle individual profiles from samples individual are shown samples along arewith shown the per alongcentage with of the cells percentage in the G1/G0 of cells phase in thefor G1/G0each respective phase for each respectivesample (A ).sample The average (A). The percentage average percentage of cells in theof cells G1/G0 in thephase G1/G0 was phasequantified was forquantified each treatment for each condition treatment (B). condition (as compared (B). (as tocompared negative to negativecontrol at control 0 hours: at f0p ≤hours:0.0001) fp ≤0.0001) measured via flow cytometry for a minimum of 50,000 cells per sample. In subsequent studies, serum starvation was utilized as a pre-treatment to synchronize cells in the
7157 Back to Table of Contents (Link)
Figure 3.5. Serum starvation induces synchronization of MC3T3-E1 cells in the G1/G0 phase: Cells were treated with basal media (0% FBS) across 24, 48, and 72 hour time points. At each time point, cells were collected and fixed for subsequent cytometric analysis. Cell cycle distribution was measured via flow cytometry for a minimum of 50,000 per sample. Representative images of the cell cycle profiles from individual samples are shown along with the percentage of cells in the G1/G0 phase for each respective sample (A). The average percentage of cells in the G1/G0 phase was quantified for each treatment condition (B). (n=3) (as compared to negative control at 0 hours: fp≤0.0001)
A
B
FigureFigure 3.5 3.6 InhibitionInhibition of of the the L L-VSCC-VSCC causes causes cell cell cycle cycle slowing slowing Prior to experimental treatments, MC3T3-E1 cells were synchronized in G1/G0 phase via serum starvation in MEM-α media with 0% FBS for 72 hours. Following synchronization, cells were allowed to recover in negative control (media) or 5 μM nifedipine, both of which were supplemented with 10% FBS. At each time point, 3, 6, 9, 12, 24, and 48 hours (data not shown for 3 and 6 hours) cells were collected and fixed for subsequent cytometric analysis. Cell cycle distribution was measured via flow cytometry for a minimum of 50,000 cells per sample. Representative images of the cell cycle profiles from individual samples are shown along with the percentage of cells in the G1/G0 phase for each respective sample (A). The average percentage of cells in the G1/G0 phase was quantified for each treatment condition (B).
7358 G1/G0 phase prior to experimental treatments. To assess whether nifedipine slows the cell cycle of MC3T3-E1 cells, cells were synchronized via serum starvation and then allowed to recover in treatment conditions. Representative images (Figure 3.6A) and quantification (Figure 3.6B) of the percentage of cells in G1/G0 phase, show cell cycle recovery after synchronization in the presence of either nifedipine treatment or negative control. Both negative control cells and cells exposed to 5 μM nifedipine remained synchronized in G1/G0 phase for the first 12 hours of recovery. At 18 hours, cells began to recover and desynchronize from the G1/G0 phase in both conditions, however, the 18 hour time window shows that a larger proportion of cells remained synchronized compared to the negative control. Both conditions eventually recovered fully from synchronization after 48 hours. These data indicate that nifedipine slows cell cycle progression at the G1/S boundary, which suggests that the L-VSCC mediates proliferation through regulation of cell cycle progression.
3.4 T-VSCC inhibition reduces cell number of MC3T3-E1 in a dose-dependent manner
Following experimentation with the L-VSCC, it was necessary to explore the relationship between the T-VSCC and MC3T3-E1 cell proliferation. Cell number losses were observed in initial studies with the T-VSCC antagonist NNC (Figure 3.1), and I expand upon these here to demonstrate that this effect is dose-dependent. Cell number was quantified daily via hemocytometer count over four days of treatment for the following conditions: 1 μM NNC, 5 μM NNC, and 10 μM NNC (Figure 3.7A-D). By 24 hours of treatment, all three NNC treatments showed a significant reduction in cell number compared to the control. At each time point, cells treated with 10 μM
59 Back to Table of Contents (Link)
A B
C D
Figure 3.6 T-VSCC inhibition reduces cell number in a dose-dependent manner Figure 3.7 DoseT-VSCC-dependent inhibition effects reduces of NNC cell on thenu mbernumber in of a MC3T3dose-dependent-E1 cells at manner (A) Figure 3.7. T-VSCCDose inhibition-dependent reduces effects cell of number NNC on in thea dose number-dependent of MC3T3 manner:-E1 cells at (A) Dose-dependent24 effects hours, of (B)NNC 48 on hours, the number (C) 72 of hours, MC3T3 and-E1 (D) cells 96 at hours (A) 24 (as hours, compared (B) 48 to 24 hours, (B) 48 hours, (C) 72 ahours, andb (D) 96c hours (ase compared to time-matched negative controls:a p≤b0.05, p≤c0.01, p≤e0.005, p≤f0.0005, hours, (C) 72 hours, and (D) 96 hours. (n=3) ( p≤0.05,a p≤0.01, bp≤0.005, cp≤0.0005, ep≤0.0001) ftime-matched negative controls: p≤0.05, p≤0.01, p≤0.005, p≤0.0005, p≤0.0001). fp≤0.0001).
NNC exhibited a significant reduction in cell number compared to the control. At 72 hours, all three NNC conditions showed a significant difference compared to one another, suggesting that the effects of NNC are dose-dependent.
3.5 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells
3.5 TAs-VSCC reported inhibition in chapter reduces 3.1, proliferation cell viability assays of MC3T3 indicated-E1 cells that high doses of NNC As reported above, proliferation assays indicated that high doses of NNC caused a loss of MC3T3-1 cells below that of the initial seeding number, suggesting caused a loss of MC3T3-1 cells below that of the initial seeding number, suggesting that cell death might occur when the T-VSCC is blocked. Thus, an annexin-V assay was performed in order to determine if the reduction in cell number observed with
76
60
A
B
Figure 3.8 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells (as measured via an annexin-V assay) MC3T3-E1 cells were treated with either negative control (media) or 10 μM NNC. Cells were collected at 24 and 48 hours, and cell viability was assessed via annexin-V assays and cytometric analysis. Cell viability levels are represented as the percentage of 50,000 total MC3T3-E1 cells negative for Annexin-V and PI fluorescence at 24 hour (A) and 48 hour time points (B) when normalized to the negative control for that time point (when compared to the time-matched negative control: ep≤0.001, fp≤0.0001).
61 NNC treatment was due to changes in cell viability. Treatment with 10 μM NNC demonstrated a significant reduction in cell viability comparedBack to to the Table negative of Contents control (Link) at the 24 hourFigure and 483.8. hourT-VSCC time inhibition points reduces (Figure cell 3.viability8). By of 48 MC3T3 hours,-E1 cellcells (asviability measured had via annexin-V assays): MC3T3-E1 cells were treated with either negative control (media) or 10 μM dropped to lessNNC than. Cells half were of collected the viability at 24 and observed48 hours, and in cell the viability negative was assessed control via coannexinndition.-V assays and cytometric analysis. Cell viability levels are represented as the percentage of 50,000 This data wastotal supported MC3T3-E1 through cells negative fluorescent for Annexin microscopy-V and PI fluorescence with acridine at 24 hour (A)orange and 48 and hour time points (B), when normalized to the time-matched negative control. (ep≤0.001, fp≤0.0001) ethidium bromide staining (Figure 3.9). Treatment with 1 μM NNC for 24 and 48 hours demonstrated a significant reduction in the percentage of viable cells compared
to the control at each time point respectively. Both cell viability assays demonstrate
Figure 3.9 T-VSCC inhibition reduces cell viability of MC3T3-E1 cells (as measuredFigure 3.9. T- VSCCvia an inhibition acridine reduces orange cell viability assay ofs) MC3T3 -E1 cells (as measured via MC3T3acridine orange-E1 cells assays were): MC3T3 treated-E1 cells with were either treated negativewith either negative control control (med (media),ia), 1 10μM μM NNC, or positive control (heat-induced apoptosis). Then, acridine orange assays were NNCutilized, toor quantify positive cell viability,control which (heat are-induced represented apoptosis). here as the percentage Then, acridineof total MC3T3 orange- assaysE1 cells negativewere used for fluorescent to quantify and morphological levels of cechangesll viability, associated which with apoptosis are and representednecrosis. All viability here perceas thentages percentage were normalized of total to those MC3T3 of the time-E1-matched cells negative for control. (n=3) (cp≤0.005, ep≤0.001) fluorescent and morphological changes associated with apoptosis and necrosis. All viability percentages were normalized to those of the negative control. (in comparison to the negative control: cp≤0.005, ep≤0.0005)
62 that NNC reduces the viability of MC3T3-E1 cells, suggesting that T-VSCC is necessary for survival.
3.6 VSCC activation is necessary for PEMF-induced increase in proliferation of MC3T3-E1 cells
VSCC’s have demonstrated an ability to mediate osteoblast proliferation, sustaining basal levels of proliferation at resting conditions and driving increased levels of proliferation in response to mechanical stimuli. We now investigate PEMF as an alternative means of activating VSCC’s and the functions they mediate within osteoblasts. The effects of PEMF stimulation on osteoblast proliferation were examined through supervision of a series of experiments that were largely performed by Abigail Dela Paz, an undergraduate within our lab (Dela Paz, 2019). Through the use of Hemholtz coils to generate and expose MC3T3-E1 cells to PEMF stimulation, exposure was observed to significantly increase proliferation at 48 hours, 72 hours, and 96 hours compared to the negative control (Figure 3.10). However, the effect of PEMF was abrogated with nifedipine treatment, suggesting that PEMF stimulation induces an increase in proliferation through activation of VSCC’s. NNC treatment was also observed to block the effect of PEMF treatment; however, the results from previous proliferation assays (chapter 3.1 and 3.4) and cell viability assays (chapter
3.5) suggest that this effect was likely exerted through reductions in cell viability rather than through reductions in proliferation. Overall, these results suggest that PEMF stimulation induces increased proliferation of MC3T3-E1 cells and that this effect is exerted through activation of the L-VSCC.
63
Figure 3.10 VSCC activation is necessary for PEMF-induced increase in proliferation of MC3T3-E1 cells Proliferation of MC3T3-E1 cells in response to control, nifedipine, and NNC conditions +/- PEMF treatment, as measured by relative absorbance values of MTS assay at 24 hours, 48 hours, 72 hours, and 96 hours with respect to the recorded baseline at Day 0. PEMF exposure was defined as 2 hours of repeated PEMF exposure (2.2mT ± 0.5mT) (as compared to time-matched negative controls (no antagonists): ap≤0.05, cp≤0.005, ep≤0.0005, fp≤0.0001).
3.7: PEMF exposure induces extracellular release of ATP The L-VSCC and purinergic signaling have been shown to mediate the proliferative response of MC3T3-E1 cells to mechanical stimuli, as well as the ability of MC3T3-E1 cells to proliferate at resting conditions (Genetos et al., 2009; Genetos et al., 2011; Li et al., 2003; Li et al., 2005; Puente et al., 2003). The current results provide further support for the importance of the L-VSCC in proliferation of MC3T3-
64 E1 cells at resting conditions (chapter 3.1) and now show the importance of the L- VSCC in the proliferative response induced by PEMF stimulation (chapter 3.6). Due to the similarities between the proliferation pathways of MC3T3-E1 cells—at resting conditions, under mechanical stimulation, and under PEMF stimulation—it was necessary to determine if the proliferative effect of electromagnetic stimulation involved purinergic signaling. As with the aforementioned PEMF study (chapter 3.6), the results presented here can be largely attributed to the work of Abigail Dela Paz (Dela Paz, 2019). In order to ascertain whether PEMF stimulation induced ATP release from osteoblasts, MC3T3-E1 cells were stained with quinacrine, stimulated with PEMF, and analyzed via fluorescent microscopy. Quinacrine fluoresces when bound to ATP, however, fluorescence of quinacrine is lost when ATP is released extracellularly due to the pH of the extracellular environment. Release of ATP, as indicated by a reduction in fluorescence, was measured for the following conditions: no PEMF exposure without quinacrine, no PEMF exposure with quinacrine, 1, 2, 3, 4, and 5 minutes of PEMF exposure (2.2mT ± 0.5mT). MC3T3-E1 cells exposed to PEMF stimulation exhibited decreased light intensity when compared to cells that were not were not stimulated by PEMF, suggesting that PEMF induces ATP release (Figure 3.11). These preliminary results suggest that the L-VSCC likely mediates
PEMF-induced increases in proliferation, in part, through activation of purinergic signaling; this in turn, suggests similarities may exist between the pathways by which L-VSCC’s maintain basal levels of osteoblast proliferation at rest, mediate increased levels of proliferation in response mechanical stimulation, and induce increased proliferation when exposed to PEMF stimulation.
65
Figure 3.11 PEMF exposure induces extracellular release of ATP Fluorescent intensities of MC3T3-E1 cells in response to treatments: 0, 1, 2, 3, 4, and 5 minutes of PEMF exposure (2.2mT ± 0.5mT) with quinacrine staining. Fluorescent values represent quinacrine staining of intracellular ATP.
66 Chapter 4
CONCLUSION AND DISCUSSION
4.1 Discussion
There is a crucial need for therapeutic solutions targeted towards the bone- building side of bone remodeling in order to help grow and maintain bone in conditions of bone loss. Within bone tissue, the survival and proliferation of pre- osteoblasts is imperative to establishing a critical number of bone-forming mature osteoblasts on the surface of bone. As mature osteoblasts differentiate into osteocytes, becoming fully embedded within the bone matrix, they lose their proliferative capacity but retain the ability to survive for many years (Shao & et al., 2005; Tate, 2004). This shift in proliferative capacity is accompanied by decreases in markers of bone mineralization and a loss of the L-VSCC (G. Stein, et al., 1993). Osteocytes retain the T-VSCC as well as the ability to survive and sense mechanical stimulation, even while deeply embedded within bone. The loss of the L-VSCC along with the proliferative phenotype within the osteocyte suggests the L-VSCC is necessary for proliferation, whereas the continued presence of the T-VSCC within osteocytes suggests a role in survival. Here we examine the processes of proliferation and survival in pre-osteoblast cells and demonstrate that VSCC’s play distinct roles to mediate these processes within bone. Proliferation and survival are calcium-mediated processes. Our lab has shown that calcium influx is necessary for load-induced bone formation in vivo and essential to signal amplification and gene expression in osteoblasts in response to mechanical
67 and hormonal stimuli (R. L. Duncan & Turner, 1995; Fiske, 2006; J. Li et al., 2003; W. Li et al., 1997) . Within the anabolic response to mechanical load, ATP release occurs, and it has been shown that the L-VSCC is crucial to this process. Researchers within our lab further postulate that the L-VSCC is necessary for basal ATP release and proliferation of MC3T3-E1 cells even in the absence of mechanical load. Unpublished research from our lab supports this, demonstrating that the L-VSCC and ATP mediate basal levels of proliferation together through two distinct pathways (Jones, 2011). Firstly, our research has shown that calcium influx through the L-VSCC triggers ATP release, activating a purinergic pathway that induces the translocation of NFAT to the nucleus. It was further shown that calcium influx through the L-VSCC activates a separate calmodulin pathway, culminating in the translocation of CREB to the nucleus. These studies shed valuable light on the cellular pathways associated with the L-VSCC and proliferation, however, they do not provide an understanding of how these pathways achieve an effector cell response following the changes made within the nucleus. Thus, it was necessary to explore how L-VSCC inhibition and activation might ultimately affect the cellular processes of pre-osteoblasts to achieve a response at the tissue level; for example, changes in cell attachment, cell viability, or cell cycle. Furthermore, it was necessary to study the T-VSCC as well, in order to delineate the difference between the roles that the L-VSCC and the T-VSCC play in bone. Findings from our lab demonstrate that inhibition of Cav1.2, an isoform of the L-VSCC �1 subunit, leads to significantly reduced responses to mechanical load at both the tissue and the cellular level (Fiske, 2006; J. Li et al., 2003; W. Li et al., 1997; Puente, 2003; Ryder & Duncan, 2014). Preliminary data has indicated that murine knockout of the
Cav3.2, an isoform of the T-VSCC �1 subunit, results in a mild skeletal phenotype and
68 a significantly reduced response to mechanical load. In contrast, murine knockout of the Cav3.1 isoform of the T-VSCC �1 subunit results in fatality, suggesting that the two isoforms of the T-VSCC may play separate roles of survival and mechanosensation within bone. VSCC inhibitors have been used within other tissues to treat a wide variety of conditions (Pontremoli et al., 2014; Songthamwat, 2018; Terry, 1982). Inhibitors and knockout mice have helped to understand the cellular pathways of VSCC’s in bone, however, there is a glaring lack of knowledge on how VSCC’s can be activated within bone to treat conditions of bone loss. Evidence shows that the difference in the activation properties of VSCC’s can allow for selective targeting of specific VSCC’s within tissues, which suggests that researchers can potentially select for specific therapeutic outcomes in bone if we understand the difference in the roles that VSCC’s play within bone tissue (Buckner et al., 2015; Catterall, 2011; Jones, 2011). Here, we explore the difference in the roles that the L-VSCC and the T-VSCC play within bone cells and test the effects of PEMF stimulation on these roles in vitro. Prior to this current study, unpublished data from our lab indicated that inhibition of the L-VSCC attenuated proliferation of MC3T3-E1 cells across four days of treatment compared to the negative control. In the current study, the procedure was modified to address potential issues of cell loss within L-VSCC antagonist conditions, and the treatments were expanded to include T-VSCC antagonists as well. When treated with nifedipine, the doubling time MC3T3-E1 cells was slowed to approximately 33 hours. The typical doubling time for sub-confluent MC3T3-E1 cells is approximately 20 hours, which is reflected in the growth pattern of the negative control cells for this study (Figure 3.1, Figure 3.2) (Amagai, Iijima, & Kasai, 1983). It is likely that the negative control cells
69 in Figure 3.2D, have reached confluency. At confluency, MC3T3-E1 cells begin to differentiate, losing the L-VSCC and the proliferative phenotype (Shao & et al., 2005). Highest dose of nifedipine showed greater significance in the reduction of cell number compared to the negative control suggesting that the effect of nifedipine on proliferation is dose-dependent (Figure 3.2D). Overall this data, which was achieved with a revised cell collection protocol, reaffirms the previous data (Jones, 2011) and suggests that nifedipine reduces cell number by affecting proliferation. In comparison to the nifedipine treatment, the NNC treatment and the combined NNC and nifedipine treatments show a higher decline in cell numbers. However, the reduction seen with administration of the T-VSCC antagonist shows that cells had been lost, since observed numbers were reduced below the number of cells originally seeded. Thus, rather than suggesting that NNC reduces proliferation, the data instead suggests that NNC reduces the viability of MC3T3-E1 cells. This supports our hypothesis and the underlying data for the role of the T-VSCC in osteogenic cells and other cell types, suggesting that it plays an important role in cell survival. In certain cell types and tissues, the L-VSCC is known to play a role in regulating both cell viability and cell proliferation. Thus, it was necessary to confirm that the reductions in cell number observed in nifedipine-treated cells were due to changes in proliferation of MC3T3-E1 cells rather than changes in cell viability. The current study shows that nifedipine does not alter cell viability in MC3T3-E1 cells, as confirmed through two separate viability assays. A dose response trial of nifedipine at 3 doses—1 µM, 2.5 µM, and 5 µM— showed no significant differences in viability compared to the negative control in an Annexin-V cell viability assay. The doses used in this assay correspond to those used in previous proliferation assays. Unlike the
70 proliferation assays, not even the highest dose of nifedipine shows a significant difference from the control at any time point. This is supported by results from an acridine orange viability assay, which shows that when cells are treated with the highest dose of nifedipine, 5 µM, they exhibit no significant reduction in viability compared to negative control cells. Apoptosis was induced in positive control treated cells, which show a significant reduction in viability when compared to the negative control as well as cells treated with 5 µM nifedipine. This confirms the data from the Annexin-V assay and suggests that the differences in cell number observed between nifedipine treatments and the negative control in the proliferation assays, were not due to changes in cell viability, but rather to changes in proliferation.
The process of proliferation is governed by the cell cycle, and calcium is universally necessary for cell cycle progression in all cell types (Hickie, 2011; C. Kahl, et al., 2003; C. R. Kahl & Means, 2004). Timing and progression of the cell through phases of the cell cycle is controlled by checkpoints that occur at the G1/S boundary and the G2/M boundary. Checkpoint progression relies on the presence of stimulatory signals such as growth factors and also depends on the absence of adverse features such as DNA damage and misalignment of chromosomes at the mitotic spindle. Calcium is necessary for cell cycle progression, because it activates cyclins which in turn activate cyclin-dependent kinases. It has been established that extracellular calcium and intracellular calcium stores are both necessary for proper proliferation of cells. When extracellular calcium concentration is lowered by a factor of ten, mammalian cells show a significant reduction in proliferation (Hickie, 2011; C. Kahl, et al., 2003). Furthermore, human fibroblasts have been shown to cease proliferation and arrest in the G1/G0 phase when placed in media containing low Ca2+;
71 interestingly, when transferred to media with normal Ca2+ levels fibroblast cells recovered within hours and underwent DNA synthesis. The importance of extracellular calcium in cell cycle progression is well established, but extracellular calcium simply provides a source to initiate the progression process. There is much that remains unknown about how calcium regulates cell cycle progression, such as the specific channels through which calcium influx occurs to regulate this process and the pathways by which their downstream targets affect it. In the current study, we postulated that L-VSCC inhibition would cause cell cycle arrest at the G1/S boundary. However, initial results showed no significant difference between nifedipine-treated samples and negative controls. Rather than testing for complete cell cycle arrest, a new experimental protocol was implemented in order to determine if inhibition of the L- VSCC induced cell cycle slowing via delayed progression at the G1/S boundary. A common pre-treatment technique for cell cycle studies is serum starvation, which is used to pre-synchronize cells in the G1/G0 phase. Here, we demonstrate that this method can be used successfully to synchronize MC3T3-E1 cells in the G1/G0 phase. When recovering from G1/G0 pre-synchronization, nifedipine-treated cells showed slower recovery time compared to the negative control cells, with delayed progression at the G1/S boundary. Positive controls (data not shown) for G1/G0 phase arrest and
G2 phase arrest at the 48 hour time point further indicate that nifedipine does not cause complete cell cycle arrest in any phase. Hydrogen peroxide, the positive control used to achieve G2 arrest in this study, has been shown in other studies to reduce proliferation of MC3T3-E1 cells and induce cell cycle arrest in the G2 phase with reduced expression levels of cyclin B (M. Li et al., 2009). The 48 hour time point indicates that cells treated with hydrogen peroxide (data not shown) become
72 synchronized in the G2 phase. The data further indicates that cells which remain in serum starvation conditions (data not shown) following initial pre-synchronization in the G1/G0 phase remain synchronized in this phase throughout treatment. Altogether, these positive controls provide context and suggest that nifedipine treatment does not arrest the cell cycle at any phase, but rather delays cell cycle progression at the G1/S boundary. This is consistent, with the role and timing of calcium signaling in cell cycle progression in other cell types, which links the many downstream targets of the L-VSCC pathway with the G1/S boundary. The cellular pathway established by our lab includes calmodulin, calcium/calmodulin-dependent protein kinase II (CaMKII), calcineurin, and NFAT signaling as downstream targets of calcium influx through the L-VSCC (Jones, 2011). All of these downstream signaling targets have been tied to similar sets of changes in cell cycle progression in various cell types. Research indicates that the calmodulin selective inhibitor KN-93 results in G1/G0 cell cycle arrest in a variety of cell types (C. R. Kahl & Means, 2004). In fibroblast cells, G1/G0 arrest via KN-93 has been shown to occur through prevention of cdk4/cyclin D1 activation (C. R. Kahl & Means, 2004). Similarly, the calcineurin/NFAT pathway has been linked to hyperproliferative properties in breast cancer cells, and when calcineurin is inhibited in breast cancer cells it results in delayed cell cycle progression at the G1/S boundary (Goshima, 2019). This is observed to occur through increased cyclin D1 degradation via inhibition of dephosphorylation at T286. During calcineurin inhibition, it is possible to partially rescue normal G1/S phase progression through overexpression of cyclin D1, which identifies cyclin D1 as an important cofactor downstream of calcineurin. Calcineurin inhibition has also been shown to reduce Rb phosphorylation, which is a
73 key step in cell cycle progression across the G1/S boundary. In mammalian cells, the G1/G0 phase is characterized first by activation of cyclin/cdk4 complexes, which phosphorylate retinoblastoma (pRb), a tumor suppressor protein. This is followed by cyclin E/cdk2 complexes, which sequentially phosphorylate pRb after cyclin D/cdk4 phosphorylation. Once it is hyperphosphorylated, pRb activates E2F transcription factors, which subsequently regulate expression of genes required for S phase progression (C. Kahl, et al., 2003). Cyclin A/cdk2 complexes are activated within S phase and remain activated in G2 phase. Finally, in the advanced stages of G2 phase cyclin B/cdc complexes are activated, and cells progress into mitosis (C. Kahl, et al., 2003). Calcium plays a key role in activating cyclins within each step of the cell cycle process; however, intracellular calcium pools may extend the time window by which cells can proceed without extracellular calcium influx. Even in low concentrations of calcium, addition of fresh calf serum alone has been found to trigger initial the re-start of cell cycle machinery in fibroblast cells that are arrested in G1/G0 phase; however, if extracellular calcium is not raised to normal levels within 10 hours, the cell cycle will return to an inactive state. This suggests that the growth factors contained within serum play an important role in reaching the advanced stages of pre-replicative development within the G1/G0 phase, but that extracellular calcium influx is necessary for the final steps in crossing the G1/S boundary (Boynton, 1976). This is consistent with the data from the current study, which demonstrates that MC3T3-E1 cells arrested in G1/G0 phase begin to show signs of cell cycle re-entry and G1/S boundary progression between 12-18 hours of treatment in the negative control condition.
74 It is likely that calcium entry through the L-VSCC, and the subsequent calmodulin and calcineurin pathways that are activated, work to facilitate cell cycle progression across the G1/S boundary in a manner similar to those observed in other cell types. Once calcium ions bind to calmodulin, it in turn binds to and activates calcineurin, which dephosphorylates specific serine residues of NFAT (Jones, 2011). Dephosphorylated NFAT undergoes a conformational change in the cytoplasm that externalizes its nuclear localization signal (NLS), which causes its translocation to the nucleus (Mognol, 2016). It has been theorized that, once in the nucleus, NFAT binds to DNA and works in conjunction with other nuclear partner proteins activated by separate signaling pathways. Calcineurin activity, and subsequent NFAT activity, has been shown to play a major role in mediation of the cell cycle. NFAT2-α and NFAT1- C both promote transcription of c-Myc and cyclin A2. The protooncogene c-Myc drives cell cycle progression in the early G1/G0 phase and cyclin A2 complexes with CDK2 to drive the cell cycle in the late stages of S phase. NFAT2-/- T and B cells from mice exhibit reduced proliferation in comparison to wild-type mice and it has been demonstrated that NFAT2 upregulates expression of cyclin A2, D1, and D3. Cyclins D1 and D3 complex with CDK4 and CDK6 to drive cell cycle progression in the G1/G0 phase, and CDK4 and CDK6 expression is increased by NFAT1. Within a cancer setting, ectopic expression of NFAT1 induces chromatin formation on p15, a CDK inhibitor, which silences p15 expression and results in increased expression of cyclin D, CDK4, and CDK6. Furthermore, it has been observed that NFAT works in conjunction with STAT3 to promote CDK6 expression and increase proliferation in pancreatic cancer cells. The relationship between NFAT family proteins and the cell cycle is complex since NFAT family proteins and their isoforms demonstrate a wide
75 range of pro-proliferative and anti-proliferative effects. Researchers believe that protein partners play an important role in modulating the function of NFAT proteins to coordinate specific effects. CREB, cAMP response element-binding protein, is a likely candidate for this interaction within the context of NFAT and proliferation in bone. In addition to activating the phosphatase calcineurin, calmodulin also activates CaMKII as a result of L-VSCC calcium influx in osteoblasts. CaMKII activation in osteoblasts has been shown to lead to the phosphorylation of CREB in a manner that influences the production of bone specific markers such as, receptor activator of nuclear kappa-B ligand (RANKL) and osteoprotegerin (OPG) (Farach-Carson, Bergh, & Xu, 2004). CREB-binding protein (CBP) may allow for NFAT proteins and CREB to achieve synergistic activation of proliferation pathways. CBP, initially recognized as a factor binding to CREB, has also been found to bind to NFAT1, NFAT2, and NFAT-3 (Karamouzis & et al., 2007; Mognol, 2016). Researchers theorize that CBP may function as a physical bridge between gene-specific transcription factors and basal transcriptional machinery to stabilize the transcription complex. Alternatively, CBP may serve as a scaffold, laying a foundation for multi-component complexes comprised of transcription factors and co-factors (Karamouzis & et al., 2007). In the case of osteoblasts, it is likely CBP plays some role in integrating the calmodulin and purinergic signaling pathways to achieve a combined proliferative effect. This is supported by the strong interplay between these pathways upstream of this event. Parallel regulation of proliferation may serve a specific purpose in allowing energy to be conserved in times of stress by re-directing purinergic signaling to purposes other than proliferation. In such times, cells would be able to proliferate using the L-VSCC- mediated calmodulin pathway alone, whereas, in times of optimal health and energy
76 both pathways could be utilized together in a synergistic manner to maximize proliferation. It is likely that CREB and NFAT are integrated by CBP within the nucleus to upregulate cyclin d and CDK 4 in a manner that induces cell cycle progression across the G1/S boundary (Figure 4.1). The T-VSCC has shown a role in cell survival in many tissues, however, in certain cell types the channel has been implicated for roles in proliferation as well. Evidence from our lab has indicated that the T-VSCC plays a role in the anabolic response to mechanosensation within the osteoblast and the osteocyte (Raggatt & Partridge, 2010; W. Thompson et al., 2011). However, the role of the T-VSCC in proliferation and survival at resting levels of stimulation remains to be explored. In this study, we demonstrated that inhibition of the T-VSCC decreased cell number compared to the negative control in a manner that was dose-dependent and time- dependent. In certain instances when treating with NNC, the reduction in cell number was observed to decrease below the original seeding number. This demonstrated a loss of cells and suggested that inhibition of the T-VSCC reduced cell survival. Dose- testing within proliferation assays identified 10 μM NNC as an effective dose for inducing cell loss below the initial seeding number, and thus, a likely candidate for inducing apoptosis. Annexin-V assays confirmed that cells treated with 10 μM NNC exhibited significant reductions in cell viability compared to negative control cells. Further testing through acridine orange assays revealed that even one tenth the dose of
NNC, at 1 μM, resulted in significantly reduced cell viability compared to the negative control. Annexin-V assays within the current study have indicated that inhibition of the T-VSCC lead to elevated levels of phosphatidylserine externalization, an event associated with the early stages of apoptosis. In many cell types, such events do not
77 Back to Table of Contents (Link)
and purinergic signaling pathways to achieve a combined proliferative effect. This is supported
by the strong interplay between these pathways upstream of this event. Parallel regulation of
proliferation may serve a specific purpose in allowing energy to be conserved in times of stress
by re-directing purinergic signaling to purposes other than proliferation. In such times, cells
would be able to proliferate using the L-VSCC-mediated calmodulin pathway alone, whereas, in
times of optimal health and energy both pathways could be utilized together in a synergistic occur immediately and are instead preceded by cell cycle arrest at cell cycle manner to maximize proliferation. It is likely that CREB and NFAT are integrated by CBP checkpoints.within the nucleusIt has been to upregulate noted bycyclin researchers d and CDK that4 in athere manner may that be induces a direct cell cyclelink between
progression across the G1/S boundary (Fig. 3.12)
Figure 4.1 Proposed model of the role of calcium-mediated signaling in pre- osteoblast proliferation and survival Activation of the L-type calcium channel in the pre-osteoblasts triggers the influx of calcium into the cell. We propose that this calcium influx has multiple roles: among them, to trigger ATP release from the cell and to activate calmodulin, which then triggers activation of downstream kinases (such as CaMKII) that can lead to transcription of genes involved in proliferation. ATP release, in turn, can bind to P2 receptors on the osteoblast surface, leading to a further increase in cell calcium. Stimulation of these purinergic receptors can lead to activation of calcineurin, which can also be activated by calmodulin. When calcineurin is activated through these pathways it will dephosphorylate NFAT and this may lead to transcription of genes associated with proliferation. We postulate that CREB-binding protein coordinates the action of CREB and NFAT within the nucleus to upregulate cell cycle proteins associated with progression across the G1/S boundary (such as cyclin D1 and
78 CDK4/6). The T-VSCC’s role is unique from the proliferative role that the L-VSCC serves at resting conditions, however, the cellular pathway remains to be further explored. apoptosis and the cell cycle, as evidenced by the similarities in their processes (Pucci, 2000). In both instances, adherent cells detach from the substrate, become shrunken and rounded in shape, display chromatin condensation, and exhibit membrane blebbing (Pucci, 2000). Genes that regulate apoptosis are also linked to cell cycle progression. Particularly in the case of cancer progression, certain tumor-related genes, such as p53, Myc, and pRb, code for proliferative proteins which researchers theorize might sensitize cells to apoptosis. Thus, it is theorized that the relationship between cell cycle progression and apoptosis may be largely dependent on the cellular context in which events occur. Studies suggest that inhibition of the T-VSCC can induce G1/G0 or G2 phase cell cycle arrest in both cancer and non-cancerous cell types (Antal & Martin-Caraballo, 2019; Dziegielewska, 2013; W. Huang et al., 2015; Rim & et al., 2012; Sankhe, 2017). It would be beneficial to conduct cell cycle distribution assays in future studies to analyze early time points of NNC treatment and determine if T-VSCC inhibition causes cell cycle arrest. Regardless of whether cell cycle arrest occurs, it is likely that these assays would reveal elevated levels of events accumulated in the sub-G1 phase, a hallmark of apoptosis that represents DNA fragmentation. PEMF stimulation has been utilized in experimental and clinical settings to induce VSCC activation in various tissue types, including bone (Buckner et al., 2015; Kooistra, Jain, & Hanson, 2009). Although PEMF stimulation has shown promising therapeutic potential for fracture healing, inconsistent stimulation parameters between studies have produced indeterminant results that have deterred more widespread clinical use of the technique. This has been exacerbated by a lack of understanding
79 regarding the cellular mechanisms that underlie PEMF-induced osteogenesis. The current study has demonstrated a significant increase in proliferation of MC3T3-E1 cells in response to daily 2-hour exposure to PEMF when compared to cells that were not exposed to PEMF treatment. Utilizing quinacrine staining and fluorescent microscopy, our current pre-liminary studies have further shown that ATP release occurs in MC3T3-E1 cells in response to PEMF stimulation, suggesting that the proliferative increase seen with PEMF treatment may occur through L-VSCC activation of purinergic signaling. This is a logical conclusion, as previous studies from our lab have demonstrated that L-VSCC activates purinergic signaling as part of a proliferative response to mechanical stimulation (J. Li, Liu, Ke, Duncan, & Turner,
2005; Owan, Ibaraki, Duncan, Turner, & Burr, 1999; Ryder & Duncan, 2001). In vivo, the anabolic response of bone is dependent upon the ability of mechanical force to generate an internal electrical field within bone tissue (Pollack et al., 2011; Spadaro, 1998). Thus, it would be likely that mechanical stimulation and electromagnetic stimulation occur through similar pathways in bone.
4.2 Future Directions Future studies must be conducted to explore the cellular pathways of the L- VSCC. We postulate that CREB and NFAT are integrated by CBP within the nucleus to upregulate cyclin d and CDK 4 in a manner that induces cell cycle progression across the G1/S boundary. Western blots will be conducted in future work to confirm the link between these specific molecular changes, and L-VSCC-mediated proliferation at rest within the osteoblast. Future directions will also include exploration of additional downstream targets of purinergic signaling. In addition to CaMKII phosphorylation of CREB, PKA has been shown to phosphorylate and
80 activate CREB at Ser133 (Martin & Arthur, 2014). Our lab has previously shown that PKA is critical to the proliferative response of MC3T3-E1 cells to fluid shear that occurs through purinergic signaling pathways (Ryder & Duncan, 2001). The involvement of PKA in the proliferative response to fluid shear suggests that it may also be a downstream target of purinergic signaling at rest. Thus, in future studies it would be beneficial to explore whether basal levels of purinergic signaling activates both calcineurin and PKA-mediated pathways to work in conjunction with the L- VSCC-mediated calmodulin pathway to achieve proliferation at rest. Furthermore, cell cycle checkpoints are not only linked to proliferation, but also are strongly linked apoptotic processes (Pucci, 2000). An understanding of the role the T-VSCC plays in the cell cycle could shed light on the mechanisms by which it maintains cell viability in bone. As such, cell cycle distribution studies should be conducted to determine if T- VSCC inhibition induces cell cycle arrest prior to inducing apoptosis in osteoblast cells. Further exploration of the specific pathways by which T-VSCC inhibition induces cell death must be conducted, including analysis of changes in caspase expression. Since the T-VSCC still persists in osteocytes as the L-VSCC is lost, future directions should extend to include new explorations of the role of the T-VSCC in osteocytes at resting conditions (Shao & et al., 2005).
The effects of PEMF on osteogenic cell viability also warrant further examination, first in vitro and then in vivo, using VSCC knockout mice. An immediate future direction of the project is to determine whether PEMF stimulation can be utilized to increase cell viability, and whether this increase is dependent upon T-VSCC activation in vitro. This approach would demonstrate a distinction between the roles of the L-VSCC and the T-VSCC in PEMF-induced increases in proliferation
81 and cell viability, respectively, and would set the stage for PEMF as a means of achieving targeted therapeutic outcomes in bone. In preliminary trials with 30 minutes of PEMF exposure (data not shown), stimulated cells failed to show a significant increase in proliferation compared to the control; however, the total treatment time course was limited to 4 days. In future studies, daily treatment with 30 minutes of PEMF exposure will be extended over the course of several weeks to ascertain whether an increase in proliferation occurs. Once PEMF exposure and timing parameters are optimized, future studies will include western blotting experiments to determine if treatment increases cell cycle progression proteins associated with the G1/S boundary, such as cyclin d and CDK4 and whether this corresponds to an increase in osteoblastic proliferation markers such as osteocalcin and osteopontin. Further studies should include in vivo implementation of PEMF stimulation on Cav3.2 knockout mice. Unpublished research Dr. Ying Shao indicates that the Cav3.1 α1G
-/- subunit compensates for the loss of the Cav3.2 α1H in Cav3.2 mice. By isolating
-/- primary osteocytes from Cav3.2 mice and studying the effects of PEMF stimulation on cell viability in the presence and absence of the T-VSCC inhibitor NNC, we will be able to determine if the Cav3.1 regulates cell viability in osteocytes. We postulate that
PEMF stimulation will increase cell viability through activation of the Cav3.1. Past researchers have established that exposure to PEMF can at least partially reduce bone loss due to disuse in hindlimb unloaded mice; however, the effect of PEMF has been limited due to a lack of optimization in exposure parameters. Optimal activation of specific VSCC-mediated functions can only be validated by testing and understanding the effects the lie downstream of PEMF-induced activation. By identifying the specific roles played by VSCC channels vitro, we can then explore selective targeting of
82 functions based off of differences in electrophysiological criteria of VSCC channels in vivo. Thus, future directions of this project include the construction and implementation of a wearable device capable of exposing a mouse hindlimb to PEMF stimulation in vivo. Following in vivo treatment with PEMF, the mechanical changes in bone at the tissue level can be quantified through three-point bending tests and compression tests, which can be supported by biochemical analysis of serum markers for bone formation and resorption. The long-term future goals of this project are the optimization of PEMF stimulation parameters to selectively target the T-VSCC to maintain cell viability in conditions of bone loss in vivo, or alternatively, the targeting of the L-VSCC and T-VSCC together to promote conditions of bone growth and healing.
4.3 Conclusion In summary, these findings demonstrate distinct roles for the L-VSCC and the T-VSCC at resting conditions for the proliferation and survival, respectively, of pre- osteoblasts. Preliminary findings further characterize the calmodulin-mediated and purinergic-mediated pathways downstream of the L-VSCC and suggest that they converge within the nucleus in the form of transcription partners, CREB and NFAT, to promote cell cycle progression across the G1/S boundary. These same pathways may underlie PEMF-induced increases in proliferation of pre-osteoblasts, which the current study shows is dependent upon VSCC activation. This research sheds further light on the roles of VSCC’s at resting condition in bone and provides a much-needed understanding of PEMF-induced changes at the cellular level. Ultimately, these findings reveal the potential for more precise therapeutic interventions against issues of bone loss, including those which can be applied in instances ranging from
83 bedridden patients, to elderly osteoporotic individuals, to astronauts in the microgravity of space.
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