SYNEMIN, the HEARTIEST SPICE a Dissertation Submitted to Kent State

Home , Synemin, Titin, Troponin I, Troponin T

SYNEMIN, THE HEARTIEST SPICE

A dissertation submitted to Kent State University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Bethany C. Prudner

December 2013

Dissertation written by Bethany C. Prudner B.S., Youngstown State University, 2004 Ph.D., Kent State University, 2013

Approved by

______, Chair, Doctoral Dissertation Committee Derek S. Damron, Ph.D.

______, Co-advisor, Doctoral Dissertation Committee Mary A. Russell, Ph.D.

______, Member, Doctoral Dissertation Committee Soumitra Basu, Ph.D.

______, Member, Doctoral Dissertation Committee J. Gary Meszaros, Ph.D.

______, Graduate Faculty Representative Hanbin Mao, Ph.D.

Accepted by

______, Director, School of Biomedical Sciences Eric Mintz, Ph.D.

______, Dean, College of Arts and Sciences Janis Crowther, Ph.D.

Table of Contents

LIST OF FIGURES ………………………………………………………………….. v

CHAPTER ONE: INTRODUCTION AND BACKGROUND …………………..... 1

1.1 Intermediate Filament ………………………………………………….. 2

1.2 Excitation-Contraction Coupling ………………………………………. 8

1.3 Protein Kinase A (PKA) ……………………………………………….. 11

1.4 A Kinase Anchoring Proteins (AKAP) ………………………………… 15

DISSERTATION RESEARCH FOCUS …………………………………………... 20

CHAPTER TWO: α- SYNEMIN BINDS TO THE M-BAND REGION OF TITIN 22

Abstract …………………………………………………………………….... 23

2.1 Introduction ……………………………………………………………… 24

2.2 Experimental Procedures ……………………………………………… 27

2.3 Results …………………………………………………………………... 34

2.4 Discussion ………………………………………………………………. 41

CHAPTER THREE: β-SYNEMIN IS A DUAL AKAP …………………………... 45

Abstract ……………………………………………………………………… 46

3.1 Introduction …………………………………………………………….. 47

3.2 Experimental Procedures ……………………………………………... 50

3.3 Results …………………………………………………………………... 57

3.4 Discussion ………………………………………………………………. 65

CHAPTER FOUR: IDENTIFYTING THE VOLTAGE-DEPENDENT ANION CHANNEL (VDAC) AS A SUBSTRATE FOR β-SYNEMIN ANCHORED PKA …………………………………………………………………………………. 74

Abstract …………………………………………………………………….. 75

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4.1 Introduction ……………………………………………………………. 76

4.2 Experimental Procedures ……………………………………………. 81

4.3 Results ………………………………………………………………… 89

4.4 Discussion ……………………………………………………………. 103

CHAPTER FIVE: CONCLUDING REMARKS ...... 114

REFERENCES ...... 117

List of Figures

CHAPTER ONE: INTRODUCTION AND BACKGROUND

Figure 1.1. Schematic representation of the intermediate filament …………… 5

Figure 1.2. Schematic representation of the mitochondria microdomain …….. 10

Figure 1.3. Schematic representation of AKAPs in the heart ………………….. 18

CHAPTER TWO: α-SYNEMIN BINDS TO THE M-BAND REGION OF TITIN

Figure 2.1. Schematic representation of α-synemin ……………………………. 36

Figure 2.2. Yeast two hybrid analysis ……………………………………………. 36

Figure 2.3. Blot overlay ……………………………………………………………. 37

Figure 2.4. GST pulldown assay …………………………………………………. 38

Figure 2.5. Localization of synemin at the M-band …………………………….. 38

Figure 2.6. In vivo interactions of titin and α-synemin …………………………. 39

CHAPTER THREE: β-SYNEMIN IS A DUAL AKAP

Figure 3.1. Sequence comparison of the RISR region between dual-AKAPs 57

Figure 3.2. PKA RI interacts with β-synemin …………………………… 60,61,62

Figure 3.3. Quantification of fluorescence in the FRET ………………………... 63

Figure 3.4. Co-immunoprecipitation of RI:β-synemin …………………………... 64

CHAPTER FOUR: IDENTIFYING THE VOLTAGE-DEPENDENT ANION CHANNEL (VDAC) AS A SUBSTRATE FOR β-SYNEMIN ANCHORED PKA

Figure 4.1. Schematic representation of intracellular Ca2+ dynamics of MAM 80

Figure 4.2. β-synemin anchored PKA phosphorylated substrates ……………. 90

Figure 4.3. Separation of β-synemin anchored PKA phosphorylated substrates by 2D …………………………………………………………………………………. 92

Figure 4.4. Effects of β-synemin anchored PKA on VDAC phosphorylation ... 95

Figure 4.5. Effects of β-synemin anchored PKA on VDAC phosphorylation under H-89 inhibition ……………………………………………………………………….. 96

Figure 4.6. Effects of β-synemin anchored PKA on VDAC phosphorylation under (Rp)-8-Cl-cAMPS treatment ……………………………………………………….. 97

Figure 4.7. Effects of β-synemin anchored PKA on VDAC phosphorylation under (Rp)-8-PIP-cAMPS treatment ………………………………………………………. 98

Figure 4.8. Sequence comparison of the mitochondria targeting sequence between known mitochondria AKAPs …………………………………………….. 99

Figure 4.9. Intracellular [Ca2+] before and after isoproterenol treatment ……. 102

Figure 4.10. Schematic representation of the proposed β-synemin anchored PKA in the MAM …………………………………………………………………… 113

I may not have ended up where I intended to go, but I think I have ended up where I intended to be Douglas Adams

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Chapter One

Introduction and Background

1.1 Intermediate Filaments (IF)

1.1.1 IF background

One of the three cellular cytoskeleton systems is intermediate filaments (IFs). The other two cytoskeleton systems are the actin-containing microfilaments (MF) and tubulin-containing microtubules (MT). IFs are a key determinant for cellular architecture and dynamics and they play a major role in the integration of structure and function in striated muscles. The IFs are formed when the canonical central α-helical rod domain of about 310 amino acids from two IF proteins forms coiled coil dimmers as the first step of assembly into filaments. In turn, this region is surrounded by a non-α-helical head and tail domains that vary in size and amino acid composition[1]. Six different types of

IFs have been distinguished and are categorized based on their primary sequences, the exon/intron structures of their genes, and on subtle sequence differences of the rod domain. IFs type I and II contain the keratins and are the largest IF group. Type III IFs include many of the muscle IFs, such as vimentin and desmin. Type IV IFs are expressed mainly in neuronal cells; and, the nuclear lamins comprise the type V IFs. Nestin, tanabin and synemin make up the newer class, type VI [2], which are large proteins that consist of a short head that is less than 20 amino acids with a conventional central rod about 310 amino acids and a very long tail over 1200 amino acids [3]. Synemin, however, could be further classified in a class of its own due to the expression of different mRNAs that are

3 generated by alternative splicing in different muscle types, its distinct intron-exon organization patterns and its identification as an AKAP.

IFs are the “stress-buffering” elements of cells compared to MTs and MFs.

IFs are rather flexible; in contrast to MTs and MFs, which break when subjected to shear stress, IFs become more viscoelastic [4]. IFs traditionally were thought to only be involved in mechanical scaffolding, but are now considered to have novel functions important in many cellular processes such as cell adhesion/migration, mechanical integration of all the contractile actions of a muscle fiber, cellular integrity, force transmission, mechanochemical signal, signal transduction, targeting of proteins and lipids and organization and function of subcellular organelles [5-7].

1.1.2 IFs in the heart

In striated muscles there is a highly organized mechanochemical signaling network between the contractile apparatus and the sarcolemma cytoskeleton as well as the sarcoplasmic reticulum, mitochondria and the nucleus [8, 9]. The IF cytoskeleton in muscle cells is composed predominantly of type III IFs [10], desmin and vimentin along with a non-muscle specific IF, paranemin (Fig.1.1)

[11, 12]. An additional non-muscle specific IF not of the type III family that makes up the cytoskeleton IF network is synemin. Studies using different IF transgenic mouse models indicate that the IFs of striated muscles serve a mechanical function in force transmission and in maintaining cell and tissue integrity. They also participate in signaling cascades, including those that regulate apoptosis, modulate protein and lipid targeting, and membranous organelle biogenesis [7,

13, 14]. Specifically, studies have demonstrated that desmin deficiency blocks myoblast fusion and myotube formation [15]. This indicates one function of the IF desmin in myogenesis or myoblast fusion.

1.1.3 IFs at the mitochondria

Mitochondria are linked directly or indirectly to microfilaments, microtubules, and IFs [7]. Recent studies with desmin-deficient mice revealed the link between the desmin cytoskeleton and the loss of proper distribution of the mitochondria, a decrease in the amount of mitochondria in the cell, and a change in their morphology and their function, such as mitochondrial respiration [16].

These alterations in the mitochondria distribution and morphology are associated with the development of dilated cardiomyopathy and heart failure [17]. In desmin- null muscles, the loss of proper mitochondrial positioning leads to the loss of the proximity of the mitochondria to other organelles such as the sarcoplasmic reticulum (SR) [18]. This leads to incorrect targeting or destabilization of proteins at the mitochondria-associated SR membranes (MAM) [19]. The MAM are compartments of SR Ca2+ release near the mitochondrial membranes. One important protein at this location is the voltage-dependent anion channel (VDAC).

This channel allows the mitochondria outer membrane to take up Ca2+ from the

MAMs [20]. Thus, desmin IFs and their associated proteins play a significant role in mitochondrial morphology, positioning and respiratory function in cardiac muscles.

1.1.4 Synemin as an IF protein

Synemin was first identified as an IF-associated protein because it co-localized and co-purified with the IF proteins desmin and vimentin and is incapable of forming homopolymeric IFs [10, 11]. Synemin was eventually characterize as an IF when a canonical IF rod domain was identified within its cDNA. It was also shown that, while it is unable to form homopolymeric IFs, it can form heteropolymeric IFs with the type III IFs desmin and vimentin [21, 22].

Synemin is recognized as being a very large, unrelated member of the IF proteins. The structure of synemin contains the typified central 310 amino acid conserved α-helical rod domain but is unique in being flanked by a very short 10 amino acid N-terminal head domain and a very long ≈1000 amino acid C-terminal tail domain [21, 23]. Synemin is expressed as at least two splice variants, α- synemin and β-synemin [24]. The difference between α- and β-synemin is a 312 amino acid insert present within α-synemin. This 312 α-synemin specific insert

(ASI) is a result of alternative mRNA splicing that inserts 936 bp between the two terminal exons of the mRNA [24].

Synemin has previously been demonstrated to mainly localize at the

Z-disk [10, 11, 21, 22, 24-27] as well as at intercalated disks and the sarcolemma

[28, 29]. This is supported by its ability to form heteropolymeric IF structures with desmin and vimentin. This association leads to the suspected role of synemin to play a part in desmin related myopathies, myofibrillar myopathy (MM). A limited number of patients that suffer from MM are caused by a mutation in the IF

7 desmin or in the αB-crystallin gene, but a mutation within synemin has not yet been identified in these diseases. However, in all the patients that exhibit these diseases, each one did express an abnormally high level of synemin; they also displayed differences in the morphological patterns of synemin [24, 30]. For example, focal and subsarcolemmal aggregates in the cytoplasm of muscle fibers were formed, as well as diffused deposits of synemin throughout the total muscle fiber [26]. This indicates the involvement of synemin in the cellular pathological process seen by desminopathies, which leads to disruption of the Z- bands and an abnormal, disorganized desmin filament network [26, 31-33].

Most IF proteins represent differentiation markers due to their tissue- or cell-type specific expression however this is not the case for synemin. Although it was first discovered within muscle cells [10], It has been shown to be expressed in a broad range of non-muscle cell types and in some types of cancer. Such as vertebrate erythrocytes [34], lens cells [35], normal and malignant astrocytes [36,

37], glia and neurons [38], hepatic stellate cells [39], human hepatoma cells [40,

41], human malignant bilary epithelial cells [42], mouse spinal cord [43], and myoepithelial cells of breast carcinomas [44], this expansive expression of synemin and particularly its expression in cancer cells, suggest that synemin has important fundamental cellular functions.

Synemin interacts with an array of proteins that are IF and non-IF proteins, this further suggests that it functions in both maintaining the cell structure integrity and in stabilizing the cytoskeleton in muscle cells during repeated cycles of cell contraction and relaxation. Currently, the list of proteins that synemin has

8 been shown to bind include α-actinin, vinculin, talin, plectin-1, α-dystobrevin, desmin, nebulin, zyxin, metavinculin, dystophin, and protein phosphatase 2A [2,

21, 27, 29, 45-49]. One last important feature that sets synemin apart from the other IFs is its ability to bind protein kinase A (PKA) [29], establishing its role as not just an IF, but as an AKAP as well.

1.2 Excitation-Contraction Coupling

Efficient contraction of the heart requires coordinated handling of cAMP and

Ca2+ signaling events in cardiomyocytes. The membrane depolarization initiates contraction in a process known as excitation-contraction coupling (EC). The EC events are propagated by the sympathetic nervous system. This enhances contractile force (inotropy), heart rate (chronotropy), and myocardial relaxation

(lusitropy) through activation of the β-adrenergic receptors (β-AR) located on the sarcolemma of cardiomyocyctes. This occurs when an action potential depolarizes the membrane triggering a transient rise in intracellular Ca2+ levels by opening voltage-gated L-type Ca2+channels (LTCC) in the transverse tubules.

This induces a local increase in cytosolic Ca2+ concentrations. Which, in turn, activates the release of Ca2+ from intracellular stores through the ryanodine receptor 2 (RyR2) at the SR. The release of Ca2+ from the SR produces a global increase in Ca2+ concentrations that drives contraction of the cardiomyoctyes

[50]. The contraction takes place at the sarcomere, where troponin C binds Ca2+ ions and induces a conformational change in the troponin complex that exposes binding sites for myosin on the actin filaments and allows for the contraction to occur [51]. Subsequent relaxation occurs by removal of Ca2+ from the cytosol via

9 four main pathways; (1) Removal of Ca2+ occurs either through transport back into the SR lumen by SR Ca-ATPase (SERCA). (2) Extrusion via the Na+/ Ca2+ exchanger (NCX) and (3) the sarcolemma Ca-ATPase [50]. The fourth avenue of

Ca2+ sequestration potentially involves the mitochondria, which are equipped with an efficient machinery for Ca2+ transport and are capable of storing large amounts of Ca2+ [52-57].

The contraction and relaxation events can be enhanced by the β-adrenergic pathway, which activates protein kinase A (PKA) using the ubiquitous second messenger cyclic adenosine monophosphate (cAMP). PKA directly or indirectly regulates the phosphorylation and, thus, activity of proteins controlling Ca2+ cycling and sarcomere contraction [58]. The β-adrenergic stimulus elevates the cardiac workload by increasing the rate and amplitude of cytosolic Ca2+, which in turn accelerates ATP consumption by Ca2+ cycling proteins and the myofilaments. This leads to the mitochondria accumulating Ca2+ through several uptake mechanisms [52, 59]. The accumulation of Ca2+ in the mitochondria stimulates the electron transport chain (ETC), the F1F0-ATP synthase and the aspartate-glutamate shuttle to secure relatively constant rations of ATP/ADP during cardiac workload [60, 61]. Given this central role of Ca2+ for regulating mitochondrial energetics, the mechanisms and kinetics of mitochondrial Ca2+ uptake are of the utmost importance for understanding energy supply and demand in the heart (Fig. 1.2).

Figure 1.2 – Schematic representation of the detailed aspects of a mitochondria Ca2+ microdomain. MICU1: mitochondrial Ca2+ uptake; mNHE: mitochondrial Na+/H+ exchanger; IMM and OMM: inner and outer mitochondrial membranes; VDAC: voltage-dependent anion channel. Kohlhaas M , and Maack C Cardiovasc Res 2013;cvr.cvt032 Published on behalf of the European Society of Cardiology. All rights reserved. © The Author 2013. For permissions please email: [email protected].

1.3 Protein Kinase A (PKA)

1.3.1 PKA overview

PKA functions as a broad-specificity serine/threonine protein kinase. It is responsible for the physiologic action of many hormones, neurotransmitters and growth factors. PKA is important in the heart because cAMP signaling regulates both calcium dynamics and the rate and force of the contraction. Regulation by cAMP occurs by binding and activating the PKA enzyme. Enzymatic activity of

PKA is extensively involved in normal cardiac myoctyes function. When PKA activity is perturbed and phosphorylation is decreased on phospholamban, myosin binding protein C, and troponin I, this accompanies remodeling of the heart, leading to heart failure [62, 63]. PKA phosphorylation is also important in the RyR, and LTCC Ca2+ function. Both of these receptors are within the dyad of the myocyte. The dyad is the area where the t-tubules are in close proximity to the SR membrane. The RyR is the Ca2+ release channel and is located on the

SR, and the LTCC channels are located on the sarcolemma, primarily at the t- tubules. This close proximity of these two channels in the dyad allows for the

Ca2+ influx from the LTCC to trigger the neighboring RyR, stimulating for more

Ca2+ to be released from the SR. This is referred to as calcium release calcium reduced (CICR). The dyad insures that CICR events occur in a timely manner.

[64]

The PKA holoenzyme (R2C2) is a heterotetramer that consists of two regulatory subunits (R) bound to two catalytic subunits (C). Each of the R

12 subunits contains an N-terminal D/D (dimerization/docking) domain followed by a flexible linker that contains an inhibitor site and two-tandem cAMP binding domains. The D/D domain is an antiparallel four-helix bundle that provides the docking surface for the AKAP helix [65, 66]. The R subunits are the major receptors in the cell for cAMP. They bind the active C subunits holding them in an inactive state until cAMP levels increase to the appropriate levels [67]. In the absence of cAMP the two R subunit dimers bind the two C subunits forming the inactive R2C2. The inhibitor site of the R subunits is docked to the active site cleft on the C subunit. The cooperative binding of cAMP to the R subunit allows for release of the active C subunits [68, 69], which can then phosphorylate serine

(S) or threonine (T) residues, typically within the sequence –R-R-X-S/T-X [70].

There are two classes of PKA subunits: type I and type II. Each class contains two functionally non-redundant R subunits. Type I contains RIα or RIβ subunits and type II contains RIIα or RIIβ subunits [71]. The two classes are distinguished by their ability to be autophosphorylate. RI subunits have a pseuodophosphorylation site with an Ala at the P-site while RII subunits have a

Ser, which allows for autophospohrylation [72]. RIα and RIIα are the predominant isoforms and are expressed in most tissues, whereas RIβ and RIIβ are found mostly in central nervous systems and reproductive tissues. There are three C subunit genes (Cα, Cβ and Cγ) that have been characterized [73]. Cα represent the isoform found in most tissues, while the highest levels of Cβ have been observed in the human prostate, intestine, brain, and testis and Cγ has only been detected in human testis [74, 75]

1.3.2 PKA isoforms

PKA holoenzyme exists in two forms: a cytoplasmic type I PKA and an exclusively particulate type II PKA [73]. PKA type II is typically localize to discrete sites in the cell, such as the plasma membrane, mitochondria, cytoskeleton, and centrosomes [76], where as PKA type I tends to be more diffusely located throughout the cell but can be dynamically recruited to a specific site. Early studies indicated that AKAPs only bind the RII subunits with high affinity (KD = 1-

5 nM) [77, 78], but more recent data has characterized dual AKAPs, which are able to bind to both the RI and RII subunits (Kd = 2 and 48, respectively) [79].

While some dual AKAPs and specific RI AKAPs do bind the RI subunit, it is at a lower affinity than most AKAPs bind RII [80].

The RII D/D domain is organized as an antiparallel dimer, forming an X- type, four-helix bundle. This dimeric protein module contains a hydrophobic docking groove that associates with the AKAPs amphipathic helix [66, 81]. RI subunits dimerize in a similar way, although their D/D domain is larger, incorporating an additional 16-residue helix-turn-helix segment [82, 83]. Studies suggest that the N-terminus of the RI dimer folds back on the four-helix bundle to alter the shape and presentation of the AKAP binding determinants on the RI dimer.

Benedetto et al. compared the RI and RII compartments within a cell by fusing the docking domains to the Epac1-cAMPs FRET sensor [84]. Epac proteins (Epac1 and Epac2) are cAMP-dependent guanine-nucleotide-exchange

14 factors for the small GTPases Rap1 and Rap2 and are important mediators of cAMP signaling. Their regulation is somewhat analogous to the regulation of PKA by cAMP except that instead of the regulatory subunit dissociating from the catalytic subunit upon cAMP binding, Epac has a regulatory domain that functions as an auto-inhibitory domain. The auto-inhibitory domain acts to holds the protein in an inactive folded form so when cAMP binds to the regulatory domain, this acts to relieves inhibition, and thus allows for the unfolding the protein and for its catalytic domain to surface [85]. This makes Epac an exceptional FRET sensor for detecting the levels of cAMP. The Epac sensor was placed between a cyan fluorescent protein (CFP) and yellow fluorescent protein

(YFP). Upon cAMP binding, the protein unfolds, consequently increasing the distance between the two FRET pairs, quenching the signal. Their studies gave a better understanding between the two classes of PKA. For example, previously the RI subunit was thought to be solely cytosolic, these studies showed that RI is also anchored to AKAPs in myocytes, though the affinity is lower as compared to

RII subunits. Also, RII was originally thought to be the only isoform activated by isoproterenol (a β-AR agonist). Their studies showed that while the RII-Epac sensor was selectively activated by isoproterenol and inhibited by phosphodiesterase 4 (PDE4), the RI-Epac was also activated by isoproterenol.

This sensor was also activated by agonists for the prostaglandin E1 (PGE1), glucagon and GLP-1 receptors and preferentially inhibited by PDE2. These results give insight into the multiple cAMP compartments within the same organelle in a cell.

1.4 A Kinase Anchoring Proteins (AKAPs)

1.4.1 AKAP background

Early experiments demonstrated that hormone mediated stimulation of cAMP synthesis by different agonists induced distinctive physiological outcomes within the same cells [86]. For instance, a stimulus activating the β−adrenergic pathway was shown to selectively activate a distinct pool of PKA associated with a particulate fraction of cardiomyoctyes. In contrast, PGE1 stimulation activated a separate cytosolic pool of PKA within the same cells, each inducing a different physiological effect [87]. Realizing that cAMP was not uniformly distributed throughout the cell gave rise to the hypothesis that the opposing actions of adenylyl cyclases (AC) and PDEs generated intracellular gradients and compartmentalized pools of cAMP [88, 89]. A variety of protein-protein interactions screens identified a set of molecules that interact with the R subunits of the PKA holoenzyme [77, 90]. These proteins function to sequester signaling enzymes into discrete sub-cellar compartments to maintain localized pools of kinase activity and were named A-kinase anchoring proteins (AKAPs). The role of AKAPs in the cell is to anchor PKA and other cAMP responsive enzymes in proximity to their substrates eliciting a spatial and temporal control over substrate phosphorylation, depending on the hormonal stimulation [62, 76, 91].

AKAPs are a diverse family of proteins that vary greatly in structure, localization and the proteins that they scaffold. The feature that solely characterizes an AKAP is its ability to bind the regulatory subunits of PKA. This

16 binding occurs through a 14-18 amino acid sequence that forms an amphipathic

α-helix with hydrophobic residues that align at the NH2-terminus of the R-subunit

[92]. Most AKAPs that have been identified bind PKA RII subunit but several dual-function anchoring proteins have been identified. They bind PKA RI subunit as well as PKA RII [93, 94]. Very recently there have been AKAPs that have been identified as solely binding PKA RI [80, 95].

Many AKAPs are able to insure the correct spatial and temporal control of

PKA action through the formation of multimolecular complexes that includes a specific upstream activator, select PKA substrates, and other signaling enzymes that participate in pathway cross talk, such as kinases other than PKA, phosphatases, PDEs, ACs, and GTPases [96]. By anchoring and coordinating the activity of the signal, AKAPs are able to affect the location and dynamics of signal transduction by tethering these specific signaling molecules to particular locations in the cell [97].

1.4.2 AKAPs in the heart

The heart is a dynamic organ that responds to physiological demands made by the body’s fight or flight response. This sympathetic innervation coordinates the release of catecholamines, which elicit rapid increases in the heart rate, stroke volume, and cardiac output [98]. β–adrenergic stimulus impacts Ca2+ handling by increasing the force of contraction and by accelerating the rate of relaxation [99]. Distinct AKAP complexes contribute to each phase of

EC by directing PKA to modulate the contractile dynamics through

17 phosphorylation of accessory proteins and ion channels that comprise the contractile machinery in myocytes [50].

Multiple AKAPs associate with specific components of the cardiac EC coupling machinery and, along with other signaling proteins, facilitate modulation of the cardiac cycle, for instance, AKAPs are found in association with each of the Ca2+ transporters (LTCC, RyR2 and SERCA2). Interestingly, it has been shown that all three are associated with more than one AKAP, depending on the location of the channel and the signaling pathway involved [50, 100, 101]. For example, AKAP18α was originally thought to be the only AKAP to associate with

LTCC, but recent studies have shown that AKAP79 also binds. One possible reason that two AKAPs are found in the same subcellular location is so that each may facilitate different signaling pathways by binding two different subsets of proteins [102].

To date there have been eighteen different AKAPs identified in cardiac myoctytes [62]. These are AKAP18α [102], AKAP18δ [103], AKAP79 [104],

AKAP250 [105], Yotiao [106], muscle AKAP (mAKAP) [107], AKAP-Lbc [108],

AKAP220 [109], dual-specific (D)-AKAP-1 and -2 [93, 94, 110], AKAP95 [111], microtubule-associated protein 2 [112], Brefeldin A-inhibited guanine nucleotide- exchange protein 2 [113], ezrin [114], sphingosine kinase type 1-interacting protein (SKIP) [115, 116], gravin [105], myospryn [117], troponin T [118] and synemin [29] (Fig. 1.2).

Figure. 1.3 - AKAPs coordinate PKA signaling in the heart. In cardiac myocytes, multiple AKAPs organize physiological signaling events. Michael D. Kritzer , Jinliang Li , Kimberly Dodge-Kafka , Michael S. Kapiloff AKAPs: The architectural underpinnings of local cAMP signaling Journal of Molecular and Cellular Cardiology Volume 52, Issue 2 2012 351 – 358 http://dx.doi.org/10.1016/j.yjmcc.2011.05.002

1.4.3 Synemin as an AKAP

Russell et al. was the first to identify that the IF protein synemin was also an AKAP [29]. By screening a human heart cDNA library with the RII regulatory subunit of PKA, synemin was identified as a novel AKAP. Synemin was further characterized as an AKAP through amino acid sequence analysis, which predicted a RII binding region within synemin. When this binding region was mutated it resulted in the inability of the RII subunit to anchor to synemin. Their studies also showed that endogenous synemin bound and localized RII in cardiac myocytes at the Z-disks with desmin in adult hearts. The RII:synemin complex was also shown by co-IP studies in adult cardiac myocytes. Lastly they showed that in failing human hearts there was an increase in synemin protein levels [29]. This study was the first to identify synemin as an AKAP and is the basis of on going experiments in our lab.

Dissertation Research Focus

Synemin is a very large and unique cytoplasmic IF protein that also functions as an AKAP in cardiomyocytes. Whether α- and β-synemin have distinct subcellular localizations and/or functions is unclear. The goal in these studies was to better understand the role of synemin as an AKAP and to attempt to tease apart the roles of the two large isoforms of synemin expressed in the heart. Our findings lay the groundwork for a clearer picture of the functional impact of synemin as an AKAP and the potential role of each isoform in the cell.

Additionally, these results contribute a great deal of novel findings to the synemin field as described below.

First, we revealed the novel localization of α-synemin to the M-band of the sarcomere. Through yeast-two-hybrid screening, binding assays and co- immunoprecipitation studies we concluded that the 312 α-synemin specific insert

(ASI) that distinguishes α-synemin from β-synemin, binds the most C-terminal immunoglobulin-like domain of titin termed the M10 region.

Second, we report by yeast-two hybrid, FACS-FRET analyses and co- immunoprecipitation that PKA type I anchors to β-synemin, characterizing it for the first time as a dual AKAP. Additionally, we find that the binding of RI to β- synemin is dependent upon stimulation of the β-adrenergic pathway.

Lastly, through the use of two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) and mass spectrometry, we have identified for the

21 first time a phosphorylated target of PKA-anchored β-synemin. The identified substrate is the voltage dependent anion channel (VDAC) located at the outer membrane of the mitochondrion (OMM). Furthermore, fluorescent calcium probe studies indicate a role for β-synemin anchored PKA in calcium handling upon activation of the β-adrenergic pathway.

Chapter Two

α-Synemin binds to the M-band Region of Titin

Abstract

Synemin, an intermediate filament (IF) protein found predominately in the heart, has been implicated in protein-kinase signaling, and previously localized to the sarcomeric Z-disk. Here, we reveal the localization of α-synemin to the M- band of the sarcomere. The 312 amino acid α-synemin specific insert (ASI) that distinguishes α-synemin and β-synemin binds the most C-terminal domain of titin at the M10 region. We report herein a direct interaction of α-synemin with the sarcomeric protein titin by protein-protein interaction assays. The last 103 amino acids of the ASI region of α-synemin are shown to bind to the most C-terminal portion, M10, of titin, through in vitro, ex vivo assays. α-Synemin also co-localizes with obscurin (a well known M-band protein) at the M-band of sarcomeres within myocytes. Our results suggest that the principal role of M-band-localized α- synemin might be as a biomechanical signaling molecule (as an A kinase anchoring protein (AKAP)), rather than as a cell shape integrity molecule (as an

IF).

2.1 Introduction

Intermediate filaments (IFs) are filamentous cytoskeletal polymers approximately 10 nm in diameter. In mammalian cells, IFs provide crucial structural support; they also provide regulation of mechanical stress and force transduction [119-121]. In addition to the traditional mechanical scaffolding role, recent studies have indicated that IFs may have novel functions in many important cellular processes such as cell adhesion and migration, signal transduction, targeting of proteins and lipids, organization of subcellular organelles and as A kinase anchoring proteins (AKAPs) [5, 6].

Synemin is a very large, unique member of the IF protein superfamily

[122], expressed in skeletal, smooth and cardiac muscle. The molecular structure of the synemin protein is characteristic of a type IV IF, with a canonical, 10 amino acid N-terminal IF rod domain and an extended, 1000 amino acid C-terminal tail domain [21, 23]. Synemin is not able to self-assemble into homopolymeric filaments in vivo [21, 24, 123] and is dependent upon its ability to associate with the major type III IF proteins desmin and vimentin via their conserved α-helical rod domains to form heteropolymeric IFs [122]. Synemin also interacts with the non-IF proteins α-actinin [21, 22], vinculin [22, 47], talin [47], α-dystrobrevin [45], dystrophin/utrophin [48], and plectin-1 [27]. Thus, synemin appears capable of linking the heteropolymeric synemin/desmin or synemin/vimentin IFs to subcellular structures via interaction with non-IF protein binding partners. Our lab has shown that protein kinase A (PKA) is one of the non-IF binding partners of synemin [29], this characterizes synemin as not only an IF but an AKAP as well.

Synemin is expressed as at least two splice variants, α-synemin and β- synemin [24]. The difference between α- and β-synemin is a 312 amino acid insert present within α-synemin. This 312 α-synemin specific insert (ASI) is a result of alternative mRNA splicing that inserts 936 bp between the two terminal exons of the mRNA [24]. In previous studies, Sun et al. has shown that vinculin, metavinculin and talin interact with the 312 amino acid insert within α-synemin

[46, 47, 124] but β-synemin does not. Having different binding partners sheds light on possible divergent functional roles for the α- and β-synemin isoforms.

To investigate the possibility of different functional roles for α- and β- synemin, a yeast two-hybrid screen was used to identify additional interacting proteins for α-synemin. The ASI region of α-synemin was used as the bait in a screen of a human heart cDNA library. The rational for this experiment was that having unique binding partners would be a basis of distinct functional roles of α- and β-synemin; and, the logical region of interaction for binding partners unique to α-synemin is ASI. The C-terminal immunoglobulin (Ig)-like region of titin, M10, was identified as a strong binding partner for the ASI region of α-synemin.

Titin is one of the most abundant proteins in striated muscle and forms a continuous filament system in the myofibrils, with a single protein spanning from the sarcomeric Z-disk to the M-band [125, 126]. Titin guides myofbrillogenesis, it maintains the sarcomeric structure during muscle contraction and aids in distributing the force across sarcomeres [127]. The titin domains located at the sarcomeric Z-disc and M-band regions sense the mechanical status of the

26 sarcomere and transmit information from signaling pathways regulating muscle function[128-130].

The M-band portion of titin has surfaced as a hotspot for autosomal dominant and recessive mutations. So far, four different phenotypes that have been associated with mutations in the titin M10 region. Two of these are tibial muscular dystrophy (TMD) and limb-girdle muscular dystrophy type 2J (LGMD2J)

[131]. These disorders result from a mutation in the last exon of titin referred to as the FINmaj mutation. This mutation leads to the replacement of the four amino acids EITW by VKEK. TMD is is an autosomal dominant late-onset distal myopathy [132]. LGMD2J is is an autosomal recessive early-onset proximal muscular dystrophy [133-135].

Synemin has previously been shown to be present at the sarcomeric Z- disk where it binds to desmin [29], vimentin [10] and α-actinin [21]. Our in vitro assays suggest an additional sub-cellular location for α-synemin at the sarcomeric M-band, where it acts as a titin binding partner. We further demonstrate that the M10 region of titin specifically binds to the final 103 amino acids of ASI (ASIc) within α-synemin. Our collaborators further localized α- synemin with the well characterized M-band protein, obscurin. In addition to the in vitro assays, endogenous α-synemin and titin were also found to interact within intact cells via co- immunoprecipitation analysis from HL-1 cells. Our data suggests that α-synemin is not localized solely to the Z-disk, as previously suggested, but is also found at the M-band region of the sarcomere.

2.2 Experimental Procedures

2.2.1 Yeast strains, media and constructs

The two-hybrid reporter yeast strains used in this study were AH109

(MATa, trp1-901, leu2-3, 112, ura3-52, is3-200, gal4Δ, gal80Δ,

LYS2::GAL1UASGAL1TATA-HIS3,GAL2UAS-GAL2TATA-ADE2,URA3::MEL1UAS-

MEL1TATA-lacZ, MEL1) and Y187 (MATα, ura3-52, his3-200, ade2-101, trp1-901, leu2-3 112, gal4Δ, met, gal80Δ, MEL1, URA3::GAL1UAS-GAL1TATA-LacZ)

(Clontech, Mountain View, CA, USA). Yeast cells were cultured at 30°C in YEPD plus adenine (1% yeast extract, 2% bacto peptone, 2% dextrose and 1% adenine) or SD medium (0.67% yeast nitrogen base without amino acids and 2% dextrose, supplemented with appropriate amino acids and bases). Media was solidified with 4% agar and 40 µg/mL X-α-gal was added for detection of α- galactosidase reporter gene expression. All media reagents were obtained from

Clontech (Mountain View, CA, USA).

The titin prey construct pGADT7-M10titin was purified using the

Yeastmaker™ yeast plasmid isolation kit (Clontech, Mountain View, CA, USA).

The prey protein chosen for further experiments spanned the C-terminal 69 amino acids of human titin, thus including the final class II repeat, a Ig-like domain, (M10) of this very large protein. cDNA encoding other Ig-like regions of titin were obtained from Dr. Kerr and Dr. Bloch (University of Maryland, School of

Medicine) and then subcloned into pGADT7, generating pGADT7-titinM Ig 1,2, pGADT7-titinZ Ig 1,2, and pGADT7-titinZ Ig 4,5 to use as negative controls.

To generate the α-synemin bait constructs pGBKT7-ASI (expressing amino acids 1151-1462 (312 aa)), pGBKT7-ASIa (expressing amino acids 1151-

1243 (92 aa)), pGBKT7-ASIb (expressing amino acids 1244-1358 (114 aa)) and pGBKT7-ASIc (expressing amino acids 1359-1462 (103 aa)) the appropriate regions of the α-synemin cDNA were PCR-amplified from the original pCDNA3-

α-synemin, kindly provided by Dr. Li (Biologie Moléculaire de la Différenciation,

Université Denis-Diderot Paris 7) [24], and sub-cloned into TOPO® TA vector using the TOPO® TA Cloning® Kit (Life technologies, Grand Island, NY, USA) and transferred to pGBKT7 using the restriction sites Sfi I and Bam H III.

2.2.1 Yeast two-hybrid screening assay

To find novel interacting partners for the ASI region of α-synemin, we cloned the region corresponding to amino (1151-1462) into the yeast two- hybrid DNA binding domain vector pGBKT7. The clone (pGBKT7-ASI) was subsequently used as a bait to screen a human heart cDNA library. Yeast two- hybrid screening was performed using to the MatchmakerTM Pretransformed libraries kit, following manufacturer’s protocol (Clontech, Mountain View, CA,

USA). Briefly, 10 µg of the pGBKT7-ASI plasmid was transformed into the yeast strain AH109 via the Yeastmaker yeast transformation system (Clontech,

Mountain View, CA, USA). An overnight culture of the bait strain (pGBKT7-ASI transformed into AH109) was prepared per manufacture’s protocol. The pelleted bait strain was diluted and 5 mL of the bait culture was combined with a 1 mL aliquot of human heart cDNA library supplied by the manufacture. The two strains were allowed to mate at 30°C for 20-24 h, with gentle agitation (30-50

29 rpm). The mating was allowed to continue until zygotes were present when viewed under 10X magnification. The cells were then pelleted at 1,000xg for 10 m at room temperature. The pelleted cells were plated at 200 µL per 150 mm plate, using QD/-Ade/-His/-Leu/-Trp/X- α-gal (quadruple drop out media containing X-α-gal) agar. Plates were incubated at 30°C for 3-8 days. Colonies exhibiting a positive β-galactosidase reaction (dark blue color) were selected for further analysis using colony PCR. The PCR products were purified via a Cycle

Pure kit or isolated via a gel extraction kit (Omega bio-tek, Norcross, GA, USA) and sequenced (Genomics Core Facility Lerner Research Institute, Cleveland

Clinic Foundation, Cleveland OH) using appropriate primers. Analyses of the

DNA sequence data were performed using BLASTx, provided by the NCBI server at the National Institute of Health.

2.2.2 Yeast two-hybrid mating analysis of protein-protein interactions between α- synemin and titin

To verify the results of the initial screen, the plasmid that encoded the final

69 amino acids of titin, pGADT7-M10titin, was purified from yeast and transformed into Escherichia coli for amplification, and then retransformed into native AH109. The interaction between ASI and titin M10 region was confirmed in yeast using the bait construct pGBKT7-ASI. The region of interaction on ASI was further refined using three smaller fragments of ASI, ASIa, ASIb, and ASIc cloned into pGBKT7 (pGBKT7-ASIa, pGBKT7-ASIb or pGBKT7-ASIc). To ensure specificity of the affinity of ASI for the M10 Ig-like domain, three other class II repeats within titin were used as negative controls, pGADT7-Titin M Ig 1,2,

30 pGADT7-Titin Z Ig 1,2, and pGADT7-Titin Z Ig 4,5. The experiments were carried out with the mating strategy as described in the Clontech Yeast Protocols

Handbook, with the bait constructs in AH109 and prey constructs in the Y187 strain. Briefly, competent yeast strain Y187 was transformed with plasmids pGBKT7-ASI, pGBKT7-ASIa, pGBKT7-ASIb or pGBKT7-ASIc so as to generate four different bait strains, and AH109 was transformed with pGADT7-M10Titin, pGADT7-Titin M Ig 1,2, pGADT7-Titin Z Ig 1,2, and pGADT7-Titin Z Ig 4,5 to generate four different prey strains, via the Yeastmaker yeast transformation system (Clontech, Mountain View, CA, USA). An overnight culture of each of the eight strains (4 bait and 4 prey) were prepared per manufactures protocol. The pelleted strains were diluted and 5 mL of the bait culture and prey culture were allowed to mate at 30°C for 20-24 h, while agitating (30-50 rpm). Each possible combination of bait and prey were used (Fig. 2). The cells were then pelleted at

1,000xg for 10 m at room temperature. The pelleted cells were plated at 100 µL per 100 mm plate, using quadruple drop out media containing X-α-gal. Plates were incubated at 30°C for 3 days. Colonies exhibiting a positive β-galactosidase reaction (dark blue color) were selected as positive for protein-protein interaction

2.2.3 In vitro blot overlay assay

To confirm the yeast two-hybrid results, our colleagues (J. Kerr and R.J.

Bloch from the University of Maryland, School of Maryland) carried out blot overlay assays as described [136] using the last 100 bp of the C-terminal of titin

(TTN-CT) subcloned into the GST expression vector pGEX4T-1 (GST-M10titin)

31 and ASI, ASIa, ASIb, and ASIc subcloned into the maltose binding protein (MBP) expression vector pMAL c2x (MBP-ASI, MBP-ASIa, MBP-ASIb, and MBP-ASIc).

2.2.4 Expression of GST-titinM10 and MBP-ASI, ASIa, ASIb, and ASIc proteins

Each expression construct, GST-M10titin, MBP-ASI, MBP-ASIa, MBP-

ASIb, MBP-ASIc (kind gifts from Dr. Kerr University of Maryland, School of

Maryland) was obtained from our collaborators and used to transform chemically competent expression host cells Escherichia coli BL21 (DE3) bacterial cells (New

England Biolabs, Ipswich, MA, USA) for further analysis in our lab. Positive transformants were selected on an LB-agar plate supplemented with carbomycin

(VWR, Radnor, PA, USA). A single colony picked from the agar plate was inoculated in 2 mLs of Luria-Bertani broth (LB, BD, Franklin Lakes, New Jersey,

USA) supplemented with carbomycin and kept at 37 °C with continuous agitation

(350 rpm). The overnight culture was then used to inoculate terrific broth (TB, BD,

Franklin Lakes, New Jersey, USA) supplemented with carbomycin and kept at

37 °C with agitation until an OD600 of 0.5-0.08 was reached. Protein expression was then induced by addition of isopropyl-b-D-1- thiogalactopyranoside (IPTG,

Sigma-Aldrich, St. Louis, MO, USA) to a final concentration of 1 mM. Growth was monitored over 4 hrs. A non-induced control for each protein was made using cultures supplemented with 50 lg/mL carbomycin and 0.5% glucose and no IPTG.

Cells were spun down at 500 x g for 5 m and used for further analysis.

2.2.6 In vitro GST pull down assays

Protein purification of the GST-M10titin protein was conducted according to the manufacturer’s instructions (Pierce® GST Spin Purification Kit, Thermo

Scientific, Rockford, IL, USA) with slight modifications. Briefly, all cell pellets were lysed using B-PER protein extraction reagent (Thermo Scientific, Rockford, IL,

USA). First the GST-M10titin protein extract was added to the glutathione-S- transferase (GST) columns and incubated for one hour at room temperature. The columns were then washed extensively. Then either MBP-ASI, MBP-ASIa, MBP-

ASIb, or MBP-ASIc protein extract was added to the M10titin bound GST column.

The two protein extracts (one containing the titin M10 fusion protein and one containing a synemin fusion protein) were then allowed to incubate overnight at

4°C while rotating. The columns were then washed and the proteins were eluted per manufacture’s protocol. Each of the four protein elutions were pooled and concentrated using Amicon Ultra-2 mL centrifugal filters (Millipore, Billerica, MA,

USA) and the entire volume of each sample was subjected to 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–PAGE) and analyzed by western blotting with anti-MBP monoclonal antibody, 1:15,000 (New England

Biolabs, Ipswich, MA, USA) or anti-GST, 1:1000, negative control (data not shown, Cell signaling, Boston, MA, USA). Secondary anti-mouse was used,

1:10,000 (Santa Cruz Biotechnology Inc., Dallas, Texas, USA)

2.2.7 Immunostaining of skeletal muscle

In order to visualize α-synemin at the M-band our colleagues prepared cryostat sections of mouse tibialis anterior (TA) as previously described [137] and double immunostained the sections using anti-synemin (antibody that recognizes both isoforms of synemin) and anti-M-band obscurin antibodies.

2.2.8 Co-immunoprecipitation of endogenous titin and α-synemin

HL-1 cells were grown to 95% confluence in a 100 mm plate and lysed using 500 µL of lysis buffer (10 mM Tris-HCL pH 7.5, 300 mM NaCl, 1 % Triton-X

100, 2 mM EDTA, 1X Halt protease inhibitor and 1X Roche phosphatase inhibitor) as follows; cells were vortexed twice for 10 s and incubated on ice for

10 m Cells were vortexed once more for 10 s and centrifuged at 14,000 rpm for

20 m at 4°C. Whole cell lysate (500 µL) were placed on a rotatory shaker overnight with 5 µg of anti-α-synemin antibody (J. Kerr and R.J Bloch from the

University of Maryland, School of Maryland) at 4°C. The cell lysate containing the antibody-antigen complex was then incubated with Protein A/G PureProteomeTM magnetic beads (EMD Millipore, Bellerica, MA, USA) on a rotatory shaker for 1 h at 25°C. The antibody-antigen-bead complex was washed eight times with 10 column volumes of PBS. The protein was then eluted with 60 µLof 6X sample buffer (Thermo Scientific, Rockford, IL, USA) heated, at 95°C for 10 m and analyzed for titin with the M10-1 polyclonal antibody, 1:500, a generous gift from

Prof. Bjarne Udd [138] secondary anti-mouse, 1:5,000, (Santa Cruz

Biotechnology Inc., Dallas, Texas, USA) by western blotting. The samples were subjected to NuPAGE® Novex® 3-8% Tris-Acetate Gel (Life technologies,

Grand Island, NY, USA). NuPAGE Electrophoresis was performed at 90 V for

190 m Transfer of the proteins to nitrocellulose was set at 30 V overnight with constant buffer circulation. Reciprocal experiments were done using anti-titin E-2 antibody (Santa Cruz Biotechnology Inc., Dallas, Texas, USA)for IP and anti-α- synemin antibody for western blot analysis, anti-myc antibody was used as a negative control (Santa Cruz Biotechnology Inc., Dallas, Texas, USA). The

34 nitrocellulose membranes were developed using SuperSignal West Femto

Chemiluminescent Substrate (Thermo Scientific, Rockford, IL, USA).

2.2.9 Cell Culture

HL-1 cardiomyocytes were kindly provided by Dr. William C. Claycomb of the Department of Biochemistry and Molecular Biology, Louisiana State

University, New Orleans, LA, USA [139]. Cells were maintained in Claycomb medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% FBS

(Sigma-Aldrich, St. Louis, MO, USA), 0.1 mM norepinephrine (Sigma-Aldrich, St.

Louis, MO, USA), 100 units/mL penicillin-streptomyocin (Life technologies, Grand

Island, NY, USA), and 2 mM L-glutamine (Life Technologies, Grand Island, NY,

USA). The medium was changed approximately every 24 hours. Cells were grown at 37°C in 5% CO2 and 95% humidity

2.3 Results

2.3.1 Titin Interacts with the ASI region of α-synemin in the yeast two-hybrid system.

To identify proteins interacting specifically with α-synemin, a yeast two- hybrid interaction screen was carried out on a human heart cDNA library using the ASI region as bait (pGBKT7-ASI). Screening of 5.5 million colonies yielded

45 interacting prey clones, representing approximately 10 different known proteins. Of these 45 clones, 17 of them encoded 8 variations of the M10 portion of titin, differing slightly in length. The cDNA in these clones ranged from encoding the final 37 to the final 83 amino acids of M10 (the final Ig domain of titin). This region is coded by exon Mex 6 and is found in all full-length titin

35 isoforms, including the cardiac isoforms N2B and N2BA. One prey plasmid encoding the final 69 amino acids of titin was purified from the yeast and used for further yeast two-hybrid and in vitro assays.

2.3.2 Titin M10 interacts with the ASIc in the ASI region of α-synemin

To further identify the binding region within ASI for M10, additional yeast two-hybrid studies were performed with 3 bait plasmids covering the ASI region using methods described earlier (pGBKT7-ASIa, pGBKT7-ASIb, and pGBKT7-

ASIc; see Fig. 2.1). The strongest interaction was obtained between ASIc and titinM10 (Fig. 2.2) There does appear to be some low affinity binding between

ASIa and ASIb with titinM10, however, it was not as strong as that of the interaction between ASIc and titinM10 as denoted by the differing levels of growth and intensity of the blue color on the quadruple drop out plates containing

X-α-gal (Fig. 2.2). Additionally, in order to ensure specificity of the interaction between α-synemin with the M-band region of titin, M10, three control constructs that contain other Ig domains of titin were also used in yeast mattings as bait with the ASI as prey, no interaction was detected.

To corroborate the yeast two-hybrid interaction results, two different in vitro binding assays; blot overlay assays (Fig. 2.3) and GST pulldown assays

(Fig. 2.4) were carried out. In both the blot overlay assay and the GST pulldown, titin was shown to interact only with the ASIc region of α-synemin. These results are consistent with the yeast two-hybrid results and confirms that α-synemin interacts with titin and that the region of interaction encompasses the final 69 amino acids of the M10 region of titin and the final 103 amino acids of the 312

insert unique to the larger synemin isoform. (J. Kerr and R.J. Bloch, University of

Maryland, School of Medicine generously provided blot overlay data)

PKA binding domain

ASI !-synemin 1563 amino acids

!-synemin C-terminal tail 1243 amino acids

Rod Domain !-synemin specific insert (ASI) 312 amino acids

ASIa 92 amino acids

ASIb 114 amino acids

ASIc 103 amino acids

Figure. 2.1 - Schematic of human !-synemin. The C-terminal tail begins with aa 322. The RII binding domain is within the C-terminal tail (aa 635-653), Alpha synemin contains a 312 aa insert (aa 1151-1462) that is lacking in "-synemin. The ASI peptide was subdivided into ASIa, (aa 1151-1243), ASIb (aa 1244-1358), and ASIc (aa 1359-1462) for use in experiments as described in material and methods.

Prey Bait Section (BD -fusion) (AD-fusion) Interaction A ASI Titin M10 ++

B ASI a Titin M10 +/- C ASI b Titin M10 +/-

D ASI c Titin M10 ++ E ASI Titin M Ig 1,2 -

F ASI Titin Z Ig 1,2 - G ASI Titin Z Ig 4,5 - Figure 2.2 - Yeast two hybrid! analysis of the interaction between ASI and titin M10. The BD-fusion plasmids encoding full length ASI or ASI fragments were used as prey with AD-fusion plasmids encoding the final sixty nine amino acids of M10 region of cardiac titin. Strong interaction was observed (blue color) between ASI and titin M10 and ASIc and titin M10 on QDO medium containing X-a -gal. Weak interaction was observed both ASIa and ASIb with titin M10. As a negative control, BD-fusion plasmids expressing ASI were used as prey with AD-fusion bait plasmids expressing other regions of titin structurally similar to M10; no interaction between these pairs of peptides was observed.

Figure 2.4 - The !-synemin specific insert interacts with the M10 Ig domain of titin. Titin-GST was immobilized onto a glutathione S-transferase column. Induced MBP-ASI E.coli cell lysates were ran over the GST-titin immbolized column. This GST pull down assay was carried out to confirm the interaction between titin and !-synemin. MBP-ASIa, MBP-ASIb, and MBP-ASIc were also used in order to narrow down the interaction domain of synemin with titin. Only the last 103 amino acids of the !-synemin insert (MBP-ASIc) were capable of binding to titin-GST in this in vitro pull-down assay.

A B

Figure 2.5 - !-Synemin localizes to both the Z-disc and M-band in skeletal muscle. (A) Longitudinal sections of mouse TA muscle were immunostained for synemin (using antibody that recognizes both isoforms) and the M-line epitope of obscurin. Synemin is found predominantly at the Z-disc (overlay, red), and also localizes to the M-band (overlay, yellow). Scale bar = 10 mm. (B) Single fibers from the mouse flexorum digitorum brevis muscle were immunostained with the a synemin specific antibody. Staining appears at both the Z-disc (thick green band) and the M-band (thin green bands, white arrows). The specificity of this antibody was previously presented (99). Scale bar = 10mm. This data were generously provided by J. Kerr and R.J. Bloch (University of Maryland, School of Medicine)

2.2.3 α-Synemin is localized at the M-band in skeletal muscle sections

Confocal microscopy of longitudinal sections from stretched and immersion-fixed mouse muscles showed that α-synemin is at the Z-disc, as expected, and also localized to the M-band as confirmed in the overlay with a well-defined M-band protein, obscurin. Single fibers were then immunostained with a newly-generated, α-synemin specific antibody. These results revealed that

(Fig. 2.5) α-synemin localizes to both locations, the Z-disc where it is known to form IFs with desmin, and the M-band (This data generously provided by J. Kerr and R.J. Bloch, University of Maryland, School of Medicine). This data visually confirms that α-synemin is at the M-band in myocytes as predicted by our in vitro studies.

2.3.4 Co-immunoprecipitation confirms the interaction of α-synemin with titin

To examine the interactions of α-synemin and titin , we performed co-IP of endogenous proteins expressed in HL-1 cells. As shown in figure 2.6A titin was efficiently recovered when synemin was used to IP figure 2.6B shows that synemin was efficiently recovered when titin was used to IP, but neither were detected when the negative control antibody,Endogenous anti-myc, was used. Results These findings ! indicate that endogenous α-synemin interacts with endogenous titin in intact cells.

-Synemin bound to immobilized titin! !!"#$%&#!!!'(#&!!!!!)*+,%-! !!"#$%&#!!!!!!!!!!'(#&!!!!!!!!!)*+,%-!

"#$%&#!!!'(#&!!!!!!!)*+,%-! -Titin bound to !!!"#$%&#!!!!!!!!!!'(#&!!!!!!!!!!!!)*+,%-! immobilized synemin!

Figure 2.6 – In vivo interactions of titin and !-synemin in HL-1 cells. A. Endogenous synemin was co-immunoprecipitated from HL-1 cell lysates by anti-titin antibody, as seen in the lanes marked as elution in the control section. B. Endogenous titin was immunoprecipitated from HL-1 cells lysates by anti-synemin antibody, as seen in elution. In control experiemnts, synemin or titin did not co-immunoprecipitate with anti-myc antibody, as seen in elution.

2.4 Discussion

Here we identified an interaction between α-synemin and the sarcomeric protein titin. In the co-immunoprecipitation experiments, endogenous α-synemin was precipitated with endogenous titin from HL-1 cells, indicating close association of synemin with titin within living cells. Using the ASI region of α- synemin in a yeast two-hybrid screening of a human heart cDNA library yielded

17 clones for the M10 region of titin. This is further evidence of binding between these two regions (i.e., ASI of α-synemin and the M10 region of titin) was verified using a confirmatory yeast two-hybrid assay. Subsequent protein-protein interaction assays using GST and MBP tagged regions of titin and synemin, respectively, allowed us to refine the specific region within ASI that interacts with the titin M10 region. The last 103 amino acids of the ASI region of α-synemin were shown to bind to the M10 region of titin with high affinity.

The results of the confocal microscopy studies provided additional evidence for interaction between α-synemin and titin at the M-band in skeletal muscle cells. Synemin and obscurin co-localized at the M-band region of titin.

In the future, it would be of interest to determine if synemin at the M-band can be linked to the pathomechansims of a M-band titin-linked hereditary myopathy, such as TMD or a form of LGMD.

Calpain 3 is another M-band binding partner of titin and has been shown to be responsible for muscle disease, LGMD2A [140]. Calpain 3 is a multi- substrate calcium-dependent cysteine protease whose function is not yet fully understood. It binds titin at interdomain 7 (is7) which is the region (a region

41 between the IG domains) just upstream the M10 region. Titin is thought to stabilize calpain 3 and prevent autolytic degradation. Calpain 3 levels are reduced in the titin-induced myopathy LGMD2J and in some TMD patients [141], while synemin has been shown to be increased in failing hearts [29]. Studies by

Bilak et al. illustrated that synemin is the most calpain-labile substrate of the cytoskeletal proteins [142]. This finding reveals a pivotal link between the two titin binding proteins, calpain and α-synemin, and gives rise to the hypothesis that synemin is an important factor in the titin-M10 protein complex. It is currently not known how calpain 3 functions and how its dysfunction causes disease, but we postulate that because of its unsupervised activity due to a mutation in titin, it is unable to regulate α-synemin, leading to an increase in the synemin protein. The fact that synemin is more susceptible to calpain degradation than desmin led

Bilak et.al. to speculate that synemin has a more direct role in myopathies than previously thought [143].

We suggest that, in an M-band titin-linked hereditary myopathy, such as

TMD or a form of LGMD, mutations in the M10 region render titin unable to stabilize the autolytic degradation of calpain, thereby leading to the accumulation of synemin within cells. Further in support of this theory, preliminary data from our lab indicate that when HL-1 cells are treated with calyculin A (a PP1 and

PP2A phosphatase inhibitor) the level of synemin protein is much higher than in cells not treated with the inhibitor (data not shown). This result could arise from one of two potential mechanisms; (1) the inhibition of the phosphatases prevents phosphorylated synemin from becoming dephosphorylated, which would then

42 prevent calpain from degrading synemin or (2) the inhibition of PP2A renders it unable to dephosphorylate calpain, which then prevents calpain from becoming active and degrading synemin. In support of the second hypothesis, PP2A has been shown to bind directly to synemin [49]; it also has been shown to dephosphorylate calpains [144].

The novel finding of an interaction between α-synemin and titin lead us to inquire about potential cellular functions for synemin in this location. While titin is primarily known as a structural protein crucial for sarcomere assembly, structure, elasticity and integrity, it has also been thought to orchestrate several signaling pathways [145]. For example, titin has been shown to play an important role in the excitation/contraction mechanism that is carried out by obscurin tethered at the longitudinal sarcoplasmic reticulum around the myofibrils [146-148]. Now that synemin has been identified, as a protein in the titin-M10 complex, the next question is whether it is principally functioning as an IF or as an AKAP in this location. The down regulation of the synemin protein using siRNA in cardiac myocytes, therefore from this complex, resulted in no disruption of the distribution of M-band proteins such as titin and obscurin [28]. These results suggest that the loss of α-synemin does not have an immediate consequence on sarcomere stability and myofibrial order, indicating that α-synemin may play more of an

AKAP role than an IF at this location. Although there has been an AKAP identified in this complex, myospryn [149],this does not rule out the possibility that another AKAP is present,.

The results presented in this study are the first evidence that the ASI region within α-synemin contains a binding site for titin, localizing it to the M- band. Differential localization of the two isoforms is thought to further delineate the roles between α-synemin and β-synemin within the cell. Our studies have placed α-synemin at the M-band in adult cardiac myocytes. Although there are elements of experimental evidence for these proteins to be involved in a pathological state, much remains to be studied to learn how they coordinate together and their communal roles in a signaling pathway.

Chapter Three

β-Synemin is a Dual AKAP

Abstract

While our lab previously established that the intermediate filament protein

β-synemin is an A-kinase anchoring protein (AKAP) for PKA type II, which plays a role in maintaining the specificity of the protein kinase A (PKA) signaling pathway; we recently found that β-synemin also binds PKA type I. Classically

PKA type I was thought to be soluble due to low affinity for AKAPs. However, recent work has shown that several AKAPs are capable of binding PKA type I.

Here we report yeast-two hybrid and FACS-FRET analyses and co- immunoprecipitation data that demonstrate the binding of PKA type I to β- synemin. Additionally, this binding is dependent upon stimulation of the β- adrenergic pathway. We hypothesize that β-synemin functions as a dual AKAP in which PKA type II is bound during resting conditions and activated following β- adrenergic stimulation. The regulatory subunit of PKA type I binds only upon β- adrenergic stimulation. Though the physiological relevance of the intersection of the two types of PKA at β-synemin remains unclear, due to differences in biochemical characteristics including downstream substrates, the binding of PKA type I to β-synemin following stimulation of the β-adrenergic pathway would allow for a highly coordinated spatial and temporal response to β-adrenergic stimulation.

3.1 Introduction

Cells are able to receive information from their outside environment through a vast array of receptors and ligands, but the number of second messengers responding to these external stimuli is much more limited. Even with this limitation, each message is conveyed to the precise downstream protein(s) resulting in the desired functional response [150]. One of these second messengers, cyclic 3'5' adenosine monophosphate (cAMP), is synthesized and degraded by two types of enzymes, adenylyl cyclases (AC) and phosphodiesterases (PDEs). AC facilitates cAMP synthesis from ATP upon stimulation of G-protein-coupled receptors [151], while PDEs catalyze the conversion of cAMP to 5’AMP and are involved in termination of the signaling pathway [152, 153]. cAMP diffuses throughout the cell to activate protein kinase

A (PKA), as well as other effector proteins [154]. Through this activation of PKA, cAMP is able to exert a broad influence inside the cell. Activation of PKA occurs due to cooperative binding of cAMP to two sites on each regulatory (R) subunit, resulting in a decrease of the binding affinity of the R subunits for the catalytic (C) subunits. This allows for dissociation of the two active C subunits from the tetrameric complex and results in the phosphorylation of substrates in the vicinity of the active kinase [155-157].

PKA is classified as either type I or type II on the basis of the composition of the R subunits (RI or RII) [154]. Four genes encode R subunits (RIα, RIβ, RIIα, and RIIβ) and all four are expressed in cardiac myocytes. These subunits have distinct physical properties and affinities for cAMP. It has become evident that the

47 subcellular localization of specific PKA isoforms governs the duration and intensity of the cAMP signaling response across various subcellular compartments [153]. The specificity of the subcellular location of PKA is mediated through interaction with AKAPs .

The role of AKAPs is to act as a modifier rather than direct transducer of the cellular signal. They are capable of doing this by affecting the location, duration and intensity of this signal through the binding of both PKA and other signaling proteins simultaneously. This creates a separate signaling nanodomain termed a signalosome [158, 159]. A large number of proteins have been shown to bind to AKAPs. For example, mAKAPβ binds PKA, PDE4D3, adenylyl cyclase-

5 (AC5) and EpaI [100, 160]. In this signalsome the increasing levels of cAMP activate PKA, which then activates PDE4D3 in a negative feedback loop. The activation of PDE4D3 leads to the degradation of cAMP, and brings the signal to a halt [161, 162]. The ability of mAKAPβ to bind all of the machinery required for regulation of cAMP levels is important because it allows for spatial and temporal modulation of cAMP signaling pathways.

Conventional AKAPs contain a 14-18 amino acid sequence that forms an amphipathic helix that binds with high affinity to the regulatory subunit of PKA type II. As the understanding of AKAPs has evolved, it has become evident that some also tether PKA type I, which classifies them as dual specificity AKAPs.

While duel-specific AKAPs 1 and 2 (D-AKAP 1 and 2) were the first dual AKAPs to be identified [79, 93], in time more AKAPs initially thought to bind only PKA RII

48 were discovered to also bind PKA RI, this includes cardiac troponin T (cTNT),

PAP7, ezrin and merlin [155, 163].

To identify proteins capable of interacting with β-synemin, we used the entire C-terminal tail domain of human β-synemin as the bait in a yeast two- hybrid screen of a human cardiac cDNA library; PKA-RIα was identified as a binding partner of synemin. The non-invasive technique, Förster resonance energy transfer (FRET) and fluorescence activated cell sorting (FACS) was used to further demonstrate the interaction between β-synemin and RIα. We demonstrate that direct binding between RI and β-synemin does occur.

Additionally, we verified direct binding of RII and β-synemin [29].

Mutations of two amino acid residues in the RII binding region of β- synemin (β-synemin-P) resulted in a significant decrease in binding affinity of this mutant protein for RI. Similar results were obtained between β-synemin and RII. -

These results suggest that β-synemin is a dual-AKAP and that the RI binds either in the same region as RII or that RII binding is needed for RI to bind.

Finally, we demonstrated through co-immunoprecipitation that RI is in a complex with β-synemin. Interestingly, RI only forms a complex with β-synemin upon β-adrenergic stimulation. Our results suggest that synemin can now be classified as a dual-AKAP binding RII, as previously identified and RI, but only upon β-adrenergic stimulation. The exact mechanism of how or why RI binds to

β-synemin is not clear and is under current investigation

3.2 Experimental Procedures

3.2.1 Generation of expression constructs and site-directed mutagenesis

The human β-synemin cDNA, cloned into the eukaryotic expression vector pCDNA3, was kindly provided by Dr. Li (Biologie Moléculaire de la

Différenciation, Université Denis-Diderot Paris 7) [24]. The β-synemin fragment was subcloned in-frame into the mammalian expression vector pECFP-N1

(Clontech, Mountain View, CA, USA), using Hin dIII and Bam HI restriction sites

(ECFP- β-synemin-WT). The QuikChange® site-directed mutagenesis kit (Agilent

Technologies, Santa Clara, CA, USA) was used to introduce mutations into the

PKA binding region of full-length β-synemin cDNA. This resulted in cDNA- directed expression of a non-RII binding β-synemin with two proline substitutions

(I640P and L645P), referred to as ECFP- β-synemin-P [29].

3.2.2 Culture and transfection of HL-1 cardiomyocytes

HL-1 cardiomyocytes were kindly provided by Dr. William C. Claycomb of the Department of Biochemistry and Molecular Biology, Louisiana State

University, New Orleans, LA, USA [139]. Cells were maintained in Claycomb medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% FBS

(Sigma-Aldrich, St. Louis, MO, USA), 0.1 mM norepinephrine (Sigma-Aldrich, St.

Louis, MO, USA), 100 units/mL penicillin-streptomycin (Life technologies, Grand

Island, NY, USA), and 2 mM L-glutamine (Life Technologies, Grand Island, NY,

USA). The medium was changed approximately every 24 hrs. Cells were grown at 37°C in 5% CO2 and 95% humidity. .

Cells were seeded one day prior to transfection in complete medium at the density of 2.5 x 104 onto 0.02% gelatin/0.00125% fibronectin-coated 60-mm dishes. At the time of transfection the complete medium was exchanged with norepinephrine-free medium. Cells were transfected using Mirus 2000 (Mirus,

Madison, WI, USA) via manufacturer’s protocol.

3.2.3 Sequence analysis

The amino acid sequences of the dual specificity AKAPs ezrin

(NP_003370), merlin (NP_000259), PAP7 (NP_073573), D-AKAP1 (CAA6000),

D-AKAP2 (O88845) and MTG (075081) were aligned with synemin (this region is common to both synemin isoforms, but the specific sequence used was from

β-synemin ,CAC838581) using the multiple sequence alignment feature of

MacVector 12.7.1 (MacVector Inc, Cary, NC, USA). MacVector settings included

ClustalW multiple sequence alignment; matrix: blossom, opengap penalty: 40, extended gap penalty: 0.05, pairwise alignment mode: slow, delay divergent:

50% gap separation distance: and 8, with residue specific and hydrophilic penalties. MEME software was used for consensus sequence generation. MEME setting included one motif per sequence, and a motif length of 24 amino acids was specified [155, 164, 165].

3.2.4 Yeast strains and media

The two-hybrid reporter yeast strains used in this study were AH109

(MATa, trp1-901, leu2-3, 112, ura3-52, is3-200, gal4Δ, gal80Δ,

LYS2::GAL1UASGAL1TATA-HIS3,GAL2UAS-GAL2TATA-ADE2,URA3::MEL1UAS-

MEL1TATA-lacZ, MEL1) and Y187 (MATα, ura3-52, his3-200, ade2-101, trp1-901,

leu2-3 112, gal4Δ, met, gal80Δ, MEL1, URA3::GAL1UAS-GAL1TATA-LacZ)

(Clontech, Mountain Veiw, Ca, USA). Yeast cells were cultured at 30°C in YEPD containing adenine (1% yeast extract, 2% bacto peptone, 2% dextrose and 1% adenine) or SD medium (0.67% yeast nitrogen base without amino acids and 2% dextrose, supplemented with appropriate amino acids and bases). Media was solidified with 4% agar and 40 µg/mL X-α-gal was added for detection of α- galactosidase reporter gene expression. All media reagents were obtained from

Clontech.

3.2.5 Yeast two-hybrid screening

Yeast two-hybrid screening was performed using to the MatchmakerTM

Pretransformed libraries kit, following manufacturer’s protocol (Clontech). Briefly,

10 µg of the pGBKT7-β-synemin plasmid was transformed into yeast strain

AH109 via the Yeastmaker yeast transformation system (Clontech). An overnight culture of the bait strain (pGBKT7-β-synemin transformed into AH109) was prepared per manufactures protocol. The pelleted bait strain was diluted and 5 mL of the bait culture was combined with 1 mL of the human heart cDNA library supplied by the manufacture. The two strains were allowed to mate at 30°C for

20-24 hrs, with agitation (30-50 rpm). The mating was allowed to continue until zygotes were present when viewed under 10X magnification. The cells were then pelleted at 1,000xg for 10 m at room temperature. The pelleted cells were plated at 200 µL per 150 mm plate, using QD/-Ade/-His/-Leu/-Trp/X- α-gal (quadruple drop out media containing X-α-gal) agar. Plates were incubated at 30°C for 3-8 days. Colonies exhibiting a positive β-galactosidase reaction (dark blue color)

52 were selected for further analysis using colony PCR. The PCR products were purified either directly using a Cycle Pure kit or run on an agarose gel and isolated via a gel extraction kit (Omega bio-tek, Norcross, GA, USA).In each case, the PCR products were sequenced (Genomics Core Facility Lerner

Research Institute, Cleveland Clinic Foundation, Cleveland OH) using appropriate primers. Analyses of the DNA sequence data were performed using

BLASTx, provided by the NCBI server at the National Institute of Health.

3.2.6 Activation of PKA through stimulation of the β-adrenergic pathway

In order to activate PKA, cells were treated with the β-adrenergic agonist isoproterenol (10 µM) (Cell signaling, Boston, MA, USA) for 10 m 48 h after transfection. To prevent dephosphorylation , the phosphatases in the cells were inhibited with a PP1 and PP2A inhibitor, calyculin A (Cell signaling, Boston, MA,

USA), for 20 m prior to isoproterenol treatment. Cells were then fixed and permeabilized and used for FACS-FRET analysis or lysed for immunoprecipitation studies.

3.2.7 FRET measurement by FACS

The FRET technique is based upon the transfer of energy from an excited donor flurophore to a close-by acceptor flurophore, resulting in enhanced fluorescence emission of the acceptor. This only occurs when the distances between the donor and acceptor is within10 nm [166-168]. Performing FRET based experiments using microscopy for analysis prohibits the analysis of a population of cells in a high throughput screening fashion [169]. Therefore, in

53 order to be able to use this method on a large population of cells fluorescence

FACS was utilized to quantify the FRET signal [166, 170].

HL-1 cells were transfected with either ECFP-β-synemin-WT or ECFP-β- synemin-P and were serum-starved and stimulated with isoproterenol as indicated above. Cells transfected with empty vector pECFP-N1 served as controls. The cells were trypsinized and washed in PBS before aliquoting into 5 mL BD Falcon™ round bottom tube (BD Biosciences, Franklin Lake, New Jersey,

USA) at 50 X 104 cells/tube. They were immediately fixed in 2% formaldehyde in phosphate buffer saline (PBS, Sigma-Aldrich, Saint Louis, MO, USA) for 10 m at

37°C and then chilled on ice for 1 m. They were washed with PBS three times and spun at 300xg for 5 m each. The cells were permeabilized in 0.1% Triton X-

100/PBS (Sigma-Aldrich, Saint Louis, MO, USA) for 5 m at room temperature, followed by three wash steps as described above. Cells were centrifuged and resuspended in 3% BSA/PBS (Sigma-Aldrich, Saint Louis, MO, USA) blocking solution for 15 m at room temperature. Primary antibody, recognizing either

EGFP,PKA-RI or PKA-RII (Santa Cruz Biotechnolog Inc., Dallas, Texas, USA) was diluted 1:50 in the 3% BSA/PBS solution and incubated on a rotatory shaker overnight at 4°C. The cells were washed three times with ice cold PBS and incubated with anti-goat or anti-rabbit Alexa Fluor® 488 or Alexa Fluor® 568 (Life technologies, Grand Island, NY, USA) conjugated secondary antibody diluted

1:200 on a rotatory shaker for 60 m at 25°C. This was followed by another series of wash steps as described. Samples were resuspended in PBS and stored at

4°C in the dark until analyzed by FACSAria, equipped with 488-nm and 633-nm wavelength using FACSDiva software (BD Bioscience, Franklin Lakes, NJ, USA).

Data was analyzed with FlowJo software (Tree Star Inc., Ashland, OR,

USA). Cells were gated according to forward and sideward scatter (FSC/SSC)..

All samples were excited with 488 nm wavelength and emission was measured with either the fluorescein (FITC) or the phycoerythrin-TexasRed (PE-TXRed) channel with standard filter 530/30 and 610/20, which allowed for detection of secondary antibody Alexa Fluor® 488 or secondary antibody Alexa Fluor® 568, respectively. The samples stained with only one antibody, Alexa-568 conjugated secondary antibody allowed for measurement of the background emission for the dye when excited with the 488 nm wavelength. The samples stained with only one antibody, Alexa-488 conjugated secondary antibodies allowed for the removal of the bleed-through fluorescence of the donor (Alexa-488) in the FRET channel allowing for specific assess of the FRET pairs in samples stained with two antibodies (i.e., FRET pairs of antibodies) [166, 171].

3.2.8 Co-Immunoprecipitation studies

Six 60 mm plates transfected with β-synemin-WT or β-synemin-P were serum-starved and either left untreated or stimulated with isoproterenol as indicated above. Cells transfected with empty vector pECFP-N1 served as controls. After treatment, cells were lysed in cell lysis buffer (10 mM Tris-HCL pH

7.5, 300 mM NaCl, 1 % Triton-X 100, 2 mM EDTA, 1X Halt protease inhibitor and

1X Roche phosphatase inhibitor). Cells were vortexed twice for 10 s and incubated on ice for 10 m. Cells were vortexed once more for 10 s and

55 centrifuged at 14,000 rpm for 20 m at 4°C. The protein concentration was measured using Direct Detect (Millipore, Billerica, MA, USA) and 300 ug of protein was incubated on a rotatory shaker overnight with 5 µg of anti-EGFP antibody (Santa Cruz Biotechnology Inc., Dallas, Texas, USA) at 4°C. The cell lysate containing the antibody-antigen complex was then incubated with Protein

G PureProteomeTM magnetic beads (EMD Millipore, Bellerica, MA, USA) on a rotatory shaker for 1 h at 25°C. The antibody-antigen-bead complex was washed eight times with 10 column volumes of PBS. The protein was then eluted with 60

µL of 6X sample buffer (Pierce, Rockford, IL, USA) heated, at 95°C for 10 m and analyzed for anti-PKA-RI (Santa Cruz Biotechnology Inc., Dallas, Texas, USA) in the complex by Western blotting.

3.2.9 Western Blotting

Samples for Western blotting were prepared in 6X sample buffer previously described above. Gel electrophoresis of proteins was performed on 4-

15% gradient Mini-PROTEAN® TGX™ Precast Gels (Bio-rad, Hercules, CA,

USA) following manufacture’s procedure. Separated proteins were then transferred onto nitrocellulose membrane at 100 V for 20 m. Nonspecific protein binding sites on the nitrocellulose blots were blocked with 10 % nonfat dry milk in

TBST (25 mM Tris, .15M NaCl, .05% Tween-20, pH 7.5) for one hour at room temperature. The primary antibody recognizing the RI subunit of PKA was diluted

1:1000 in 5% nonfat milk-TBST and incubated with the membrane for 1 h at room temperature. After three 15 m washes in TBST, blots were incubated with HRP conjugated-goat anti-rabbit (Santa Cruz Biotechnolog Inc., Dallas, Texas, USA)

56 diluted 1:10,000 in 5% nonfat milk-TBST. Following the washes in TBST and one final wash in TBS, the peroxidase activity of the nitrocellulose-bound-secondary antibody was detected using SuperSignal West Pico Chemiluminescent

Substrate (Pierce, Rockford, IL, USA).

3.3 Results

3.2.1 PKA-RI was identified as a synemin binding protein by a yeast two-hybrid screening using the β-synemin C-terminal tail as bait

Some AKAPs have been shown to bind, not only PKA-RII, but also a large array of other proteins. To identify additional cellular proteins that interact with β- synemin, the entire tail domain (amino acids 322-1253) was used as the bait in a yeast two-hybrid screen of a human cardiac cDNA library. A total of approximately 1.5 x 106 transformants were screened with a mating efficiency of

6.52%, resulting in 37 positive clones that were restreaked and grown on SD/-

Ade/-His/-Leu/-Trp/ X-α-gal agar plates as further confirmation of positivity. Eight of the 37 positive clones contained the prey plasmids encoding the RIα subunit of

PKA.

3.2.2 Identification of an additional R-binding motif in dual specificity AKAPs

Following identification of PKA-RIα as a putative binding partner of β- synemin by yeast two-hybrid assay, sequence alignment analyses were performed. The data suggest that, in addition to the well-characterized RII- binding amphipathic helix domain, synemin also contains the PKA binding region called the RI Specifier Region (RISR) (Fig. 3.1) [155].

!"!"#!$%&'%()*+,'-./(-+57

!"!"#!$%&'%()*+,'-./(-+ A. A.

RISR PKA RI/RII binding domain

ASI !-synemin 1251 amino acids!-synemin 1563 amino acids

!-synemin C-terminal !-synemin C-terminal tail 1243 amino acids ,01,+ ,0#,002345+ 931 amino acids RodRod Domain Domain ,01,+ ,0#,002345+ B.

Figure 3.1 - Sequence comparison of RISR region among dual-AKAPs and !-synemin. A, Schematic diagram of the putative RI specifier region (RISR) in !-synmein (202-231 aa). This is the region proposed to augment the binding of RI subunit of PKA type I. The PKA D/D domain which has been shown to bind PKA type II (elsewhere) and PKA type I (here within) is locatedat aa 556-707. B, Sequence alignment of the RISR region in dual AKAPs azrin, merlin, pap7, D-AKAP1 and D-AKAP-2 with synemin (this sequence is common to both the "- and !-syneminisoforms). Functionally conserved Figure 3.1 – Sequence comparison of RISR region between dual-AKAPs. A. schematic diagram of RI residues among the AKAPs are boxed in green. specifier region (RISR) located at 202-231 aa of synemin. This is the region proposed to augment PKA RI binding. PKA RI and PKA RII binding domain located at 556-707 aa. This domain is known to bind PKA RII and hypothesized to bind PKA RI. B. Sequence alignment of the RISR region in dual specific AKAPs ezrin, merlin, pap7, D-AKAP-1, D-AKAP-2 and synemin. Functionally conserved residues among the AKAPs in the green boxes.

3.2.3 FRET-FACS analysis confirms the hypothesis of PKA-RI binding to β- synemin

On the basis of the preliminary in vivo and in silico data, we assessed the interactions of RI and synemin via FRET, which was performed in HL-1 cells and analyzed by FACS. Cells were transfected with plasmids directing expression of

ECFP-β-synemin-WT or ECFP-β-synemin-P. H L-1 cells transfected with empty vector (pECFP-N1) were used as controls. The methodology used to detect steady state FRET using FACS analysis depends on increased fluorescence of the acceptor following donor excitation, without direct excitation of the acceptor itself.

Control cells transfected with EGFP-β-synemin-WT in Fig. 3.2A show that

Alexa-488 can be excited and that there is a 39% increase of fluorescence emission (Fig. 3.3), as compared to unstimulated cells. This indicates that Alexa

488 bound to EGFP-β-synemin-WT can be excited. The emissions from cells treated with only one antibody (conjugated to Alexa-568) that recognizes either

RI or RII and that were expressing EGFP-β-synemin-WT measured below the limit of detection (0-1%) (Fig. 3.2B and Fig. 3, control samples) as compared to unstimulated cells. This indicates that Alexa 568 bound to RI or RII is unable to be excited without its FRET pair . When Alexa-488 bound to EGFP-β-synemin-

WT is also within the cells containing Alexa-568 bound to RI or RII a 7-8% shift is detected in the acceptor emission profile of Alexa-568 due to stimulation of the donor emission profile of Alexa-488. This indicates that interaction occurs between EGFP-β-synemin-WT and RI (Fig. 3.2C and 3.3) and also between

EGFP-β-synemin-WT and RII (Fig 3.2D and 3). Analysis of the acceptor emission profiles shows that stimulation of Alexa-488 bound to EGFP-β-synemin-P did not excite Alexa-568 bound to RI or RII (Fig. 3.2E, 3.2F and 3.3)

C and D.

E and F.

Figure 3.2 – PKA RI interacts with !-synemin. FRET analysis was performed by FACS as described in experimental procedures section. Panels on the left are histograms taken at 488 nm, donor emission wavelength, while the panels on the right are histograms of the same cells under the same conditions taken at 568 nm, acceptor emission wavelength. A, Histogram of the donor fluorescence measured by FACS in cells only expressing EGFP-!-synemin (single positive control for Alexa-488 emissions) and non-excited cells (control for baseline). B. Histogram of cells only treated with antibodies recognizing endogenous RI (top) or RII (bottom) and not treated with antibodies recognizing EGFP, thus the donor (Alexa 488) was not able to be excited. C and D. Histograms of the fluorescence measured by FACS from cells expressing EGFP-synemin- WT due to interaction with endogenous PKA RI (C) or PKA RII (D). The histograms on the right shows that the acceptor fluorescence is excited by the donor fluorescence, indicating close proximity of the two fluorophores, and thus that !-synemin WT and PKA RI or PKA RII are within 10 nm of each other, i.e. binding partners. E and F. Histograms of fluorescence measured by FACS in cells expressing EGFP-synemin-P. The histograms on the right show that the acceptor fluorescence is not excited by the donor fluorescence, indicating that the two fluorophores are not within 10 nm of each other, and thus that !-synemin-P is not a binding partner of PKA RI or PKA RII.

'#" '!" &#" &!" %#" %!" !"#$%&" ()*+,"'--" $#" $!" ()*+,"#.-" #" !"

Figure 3.3 – Quantification of fluorescence in the FRET channels measured by FACS in HL-1 cells transfected with !-synemin-WT or !-synemin-P and labeled with acceptor-dye-conjugated antibody Alexa 488 (donor) or Alexa 568 (acceptor). Values are the means +/- s.e.m. from three independent experiments.

3.2.4 Co-immunoprecipitation results in a unique binding feature of PKA-RI to β- synemin.

To further confirm the binding between synemin and RI, co-IP assays using ECFP-β-synemin-WT or ECFP-β-synemin-P transfected HL-1 cell lysates revealed that β-synemin-WT interacts with endogenously expressed RI (Fig. 3.4).

However, β-synemin-P does not. Interestingly, RI bound to β-synemin only upon

β-adrenergic stimulation.

Figure 3.4 – Co-immunoprecipitation assay in HL-1 trasfected cell to test PKA-RI interaction with !- synemin. ECFP- !-synemin-WT or ECFP- !-synemin-P were transfected into HL-1 cells. Fourty-eight hours after transfection cells were lysed and 300 µg of total protein was incubated with anti goat EGFP ab and immobilized on Protein G magnetic beads. Precipitated PKA-RI was deteceted with anti rabbit PKA RI ab. PKA RI was detected specifically in !-synemin-WT samples upon isoproterenol stimulation (WT+I) and was unable to be detected in control cells -/+ isoproterenol (C -/+ I), !-synemin-P -/+ isoproterenol (P -/+I) and the !-synemin-WT without isoproterenol (WT-I).

3.4 Discussion

The organization of the signaling machinery into discrete compartments is increasingly recognized as a critical feature for the specificity of the cAMP/PKA signaling pathway in cardiomyoctyes [172]. Our studies demonstrate for the first time that β-synemin is not only an AKAP for PKA type II but also a dual-AKAP binding both type II and type I PKA. Using the tail domain of human β-synemin in a yeast two-hybrid screen of a human heart cDNA library yielded eight clones containing cDNA sequences encoding PKA-RIα. To determine if synemin contained a putative RISR region, sequence analysis was performed. It revealed that synemin does contain a presumed RISR region between residues 202-231.

The addition of the RISR region provides a mechanism for multi-site contact with the RI subunit that is thought to stabilize the formation of the anchored PKA type

I complex [155]. The addition of the RISR region leads to a more dynamic binding between the AKAP-PKA type I holoenzyme than previously thought.

The RISR region is thought to augment RI binding to the AKAP. Bioinformatics analysis of synemin aligned with dual AKAPs shown to have the RISR motif suggests that synemin does contain this additional binding determinant upstream of the amphipathic helix at residues 202-231 (Fig.1). Jarnaess et al. showed that the RISR acts synergistically with the amphipathic helix in dual specificity anchoring proteins to enhance anchoring of PKA type I [155]. Further biochemical studies are needed to confirm the role of the putative RISR region of

β-synemin in RI binding. These include site-directed mutagenesis within the

RISR motif to see if this inhibits RI binding, but not RII binding, to β-synemin.

Conducting experiments using β-synemin with the mutated RISR region under β- adrenergic stimulation will allow us to examine if the RISR region has a role in binding β-synemin RI upon stimulation. Also, plasmids could be constructed that express the putative β-synemin RISR region alone to see if this region is able to bind RI in an overlay assay as Jarnaess et al. has previously shown with PKA binding region of ezrin [155]. This will lead to a better understanding of the dynamic interaction between β-synemin and the two isoforms of the regulatory subunit of PKA. This should, in turn, lead to a better understanding of the functional role of β-synemin as an AKAP.

The in vivo assays FACS-FRET and co-immunoprecipitation analysis indicate that the interaction of β-synemin with RI was specific and not due to a false positive in the yeast two-hybrid system. Since the percent of FRET is a direct measurement of the proximity of two proteins inside a cell and is a strong proof of direct interaction between two proteins we conclude that β-synemin directly binds to RI. It also provides insight into the manner in which RI binds to

β-synemin; one of two things could be occurring, either RI is binding the same region of β-synemin as RII or RII must be bound to β-synemin for RI to bind. Our data does not distinguish between these two possible mechanisms.

To further investigate the mechanism of RI and RII binding to EGFP-β- synemin-WT in the future, specific intracellular localization of the complexes could be analyzed by FRET experiments using confocal microscopy under non- stimulated conditions (the experiments in this study were done under stimulated conditions). These experiments can be done in a time dependent fashion [171] to

67 see if non-stimulated cells would alter either the localization or the binding, via

FRET confocal microscopy, of RI or RII to EGFP-β-synemin-WT as shown to occur in co-immunoprecipitation assays.

Currently, it is unclear if α-synemin binds RI in the same manner as β- synemin, these studies have not been done. The 312 amino acid insert may have impact on other regions within the protein, for instance, Sun et al. [46] showed that the protein zyxin was unable to bind α-synemin with the same affinity as it does to β-synemin, due to the insert, even though its binding region is outside of this region. Our previous findings have shown that α-synemin is predominantly at the M-band in cardiomyocytes and that β-synemin is at the Z-disk; indicating that they have two separate roles within the cell and so it is possible that RI is not needed to bind for α-synemin to carry out its function, adding further divergence to their functional roles.

Not only do these results represent the first evidence that β-synemin is a dual AKAP but it also gives an organization to the interaction between the two isoforms anchored to β-synemin with respect to the β-adrenergic pathway. How the individual PKA isoforms serve to deliver a specific response remains undetermined sense they possess different structural features and biochemical characteristics, which results in a non-redundant function within the cell [35] .

This would lead to the assumption that they are anchored to β-synemin under certain conditions to elicit a very specific response. Biochemically PKA RII is more sensitive to lower levels of cAMP than PKA RI and so it could be postulated that PKA type II is the first to respond to a cAMP signal. However, the

68 mechanism by which different PKA isoforms expressed in the same cell manage to perform distinct functions upon activation by the same soluble intracellular messenger, cAMP, remains to be elucidated.

Previous work form our lab has shown that the RII subunit binds to β- synemin in the absence of β-adrenergic stimulation while the present studies show that the RI subunit binds to β-synemin only upon β-adrenergic stimulation.

This reveals a possible mechanism by which PKA type I and type II, may respond to temporally distinct but spatially restricted cAMP signals: first by binding PKA type II to the dual-AKAP β-synemin under resting conditions and then, upon stimulation of the β-adrenergic pathway, binding PKA type I.

Benedetto et al. showed that after treatment with isoproterenol, AC generated a restricted pool of cAMP that selectively activated AKAP-anchored

PKA-RII [84, 89]. We have shown here that stimulation of the β-adrenergic pathway causes RI to bind to β-synemin. But whether or not cAMP binds to PKA type I before or after it anchors to β-synemin is currently unclear.

The ability to utilize two FRET-based probes RI_epac and RII_epac sensors would allow us to assess if PKA-RI and PKA-RII are selectively and independently activated by confined pools of cAMP elicited in response to β- adrenergic stimulation. The sensors allow for analysis of selective targeting of the

RI or RII subunits. When cAMP binds to either of the sensor’s cAMP binding domains, it will result in a conformational change that causes an increase in the distance between the cyan florescent protein and the yellow fluorescent protein moieties on the PKA isoform, decreasing the FRET signal. This would mimic the

69 response of endogenous PKA isoforms to cAMP signals upon β-adrenergic stimulation [84, 173]. This would allow us to identify if the RII subunit is released from β-synemin after β-adrenergic stimulation or if it remains bound. It would also allow us to analyze if cAMP binds to the RI subunit before it is anchored to β- synemin or if it anchors and then cAMP binds.

Another technique that would be useful to study the role of the RI and RII isoforms with respect to their binding to β-synemin upon activation of the β- adrenergic pathway would be bioluminescence resonance energy transfer

(BRET). Prinz et al. established this isoform specific protein kinase A subunit interaction assay as a tool to unravel signal transduction events in a living cell.

They tagged RI-α or RII-α and the C subunits with Renilla luciferase (Rluc) as the bioluminescent donor or with green fluorescent protein as the energy acceptor.

This allows for direct probing of PKA subunits and their interaction within living cells allowing for the study of side-by-side PKA type I versus type II holoenzyme dynamics [174].

BRET occurs when luminescence energy, generated by Rluc catalyzed substrate oxidation, is transferred to the GFP in close proximity (1-10 nm). The

BRET signal is determined by measuring the ratio of green (acceptor, 515 nm) over blue (donor, 410 nm) light [174, 175]. This differs from FRET based-sensors because the detection in BRET occurs between direct interactions of fusion proteins without a florescent energy donor excited by an external light source.

Through this method we could quantify the PKA subunits in response to PKA agonists and antagonists as well as to substances interfering with the binding of

PKA to AKAPs thereby affecting the local PKA activity in response to β- adrenergic stimulation in the intact cell.

The advantages of this technique is that it would allow us to compare

BRET signals between β-synemin-WT and β-synemin-P transfected cells. If PKA type I is activated before binding to β-synemin then the BRET signal should be increased in both sets of cells. If it is only activated once bound to β-synemin, then the BRET signal should only decrease in the β-synemin-WT cells., indicating that PKA type I must be bound to cAMP and β-synemin before it can release the catalytic subunit.

One possible explanation for the sequential binding of PKA type II then

PKA type I to β-synemin upon stimulation of the β-adrenergic pathway is that while PKA type II is playing an active role, PKA type I is playing an inhibitory role.

This conclusion is based on the assumption that PKA type I binds cAMP before anchoring to β-synemin, and once anchored it becomes active and elicits its response well after the response of PKA type II. A recent study characterized a

RIα-RegA complex, which participates in signal transduction. RegA is a PDE-like protein. Its responsibility upon binding to RIα is to mediate active dissociation of cAMP from R subunits, thus hastening PDE catalysis, which leads to signal termination and inactivation of PKA . This study concluded that RIα is an activator of PDEs and, in this way, RIα acts to terminate the signaling pathway, demonstrating that PDEs play a bigger role in signal termination than previously thought [153].

We therefore hypothesize that the responsibility of RIα in the synemin complex may be to recruit a PDE to the complex. Upon β-adrenergic stimulation an increase in local cAMP near PKA type II anchored β-synemin would bind to and activate this isoform, allowing for it to phosphorylate its downstream substrates. Since PKA RI is not as sensitive to cAMP levels as PKA RII, PKA RI would not bind cAMP until the local concentration is much higher. The binding of cAMP to RI would then recruit the proposed complex of RIα-PDE to β-synemin.

Once bound to β-synemin, PKA type I would become active. Our proposed local substrate, PDE would then be, activated causing a decrease the level of cAMP in the microdomain. This would allow for the signal termination and for the inactivation of PKA type II to be facilitated. Although we have not attempted to detect PDEs in the β-synemin complex, other AKAPs have been shown to associate with PDEs [161, 162].

PKA type I activation is not based solely on cAMP levels, it is also based on the availability of downstream substrates. One study showed that at high cAMP levels, the C subunit remains inactive and bound to the R dimer as a holoenzyme when the substrate availability was low. They showed that PKA type

I did not become active until the substrate was available [176]. This study supports our hypothesis that PKA type I does not become active until it has been anchored by β-synemin even though cAMP has bound to the R subunit., Since the activity of a PDE is not required until it is time to shut off the signaling pathway by degrading cAMP, its activity is not needed until later in time. Thus, its

72 activity needs to be regulated temporally as would be achieved with the proposed sequential binding to β-synemin and then activation upon binding to this AKAP.

PDE4 has been shown to be the isoform prevalent in cardiomyoctyes and to be the contributor to the temporal cAMP signal in the signalsome [177]. PDE4 is also the major isoform responsible for the regulation of PKA mediated phosphorylation upon β-adrenergic stimulation. To determine if there actually is a

PDE in the β-synemin-PKA type I complex, a global proteomics approach with

HL-1 cell lysates can be carried out. In co-IP experiments synemin can be immobilized and the HL-1 cell lysate used for the protein source to capture all proteins that bind to β-synemin. The proteins bound to β-synemin could then be identified via mass spectrometry using methods similar to those in section 4.2..

In summary, β-synemin had been previously identified as a classical

AKAP binding RIIα subunit. Here we have expanded on its role as an AKAP identifying it as a dual-AKAP binding RIIα as well as RIα. Evidence through bioinformatics mapping shows that β-synemin contains the upstream RISR region that is similar in most dual AKAPs. FACS-FRET data showed that PKA-

RIα binding is dependent upon the PKA-RIIα domain and this recruitment of RIα only occurs upon β-adrenergic stimulation. Based on our data, we propose the following model: under resting conditions PKA type II is bound to β-synemin.

Upon β-adrenergic stimulation, PKA type II is activated. The increasing levels of cAMP bind to PKA type I anchoring it to β-synemin. The bound cAMP anchored

PKA type I is now primed to become active. One possibility of the sequential binding between the two isoforms is that PKA type I is responsible for recruiting

PDE to the signalsome. This recruitment would not be necessary until after PKA type II fulfilled its responsibility of generating a response to the β-adrenergic pathway. PKA type I would then be responsible for terminating the signal by activating PDE.

Chapter Four

Identifying the Voltage-Dependent Anion Channel (VDAC) as a substrate for β-synemin anchored PKA

Abstract

A-Kinase anchoring proteins (AKAP) immobilize and concentrate protein kinase A (PKA) isoforms at specific subcellular compartments where they preferentially phosphorylate target substrates. Intracellular targeting of the PKA holoenzyme elicits rapid and efficient phosphorylation of target proteins, thereby increasing the sensitivity of downstream effectors to cAMP action. Through the use of two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) and mass spectrometry, we have identified a phosphorylated target of PKA-anchored to β- synemin. The identified substrate is the voltage dependent anion channel

(VDAC) located at the outer membrane of the mitochondria (OMM). In silico studies suggest the localization of β-synemin to the mitochondria where VDAC is localized. We propose that β-synemin anchors PKA to the outer mitochondrial membrane (OMM), which focuses and enhances cyclic AMP signaling in key microdomains associated with Ca2+ transport between the mitochondria and sarcoplasmic reticulum (SR). PKA tethering between the two organelles allows it to phosphorylate VDAC, which leads to binding of tubulin and closing of the channel. Fluorescent calcium probe studies indicate a role for β-synemin- anchored PKA in calcium handling. As the closing of VDAC increases its affinity for Ca2+, we can postulate that β-synemin anchored PKA plays a notable role in

Ca2+ buffering homeostasis and excitation-contraction coupling within cardiomyocytes.

4.1 Introduction

Excitation-contraction (EC) coupling in cardiac myocytes consumes vast amounts of energy in the form of adenosine triphosphate (ATP) that needs to be efficiently replenished by oxidative phosphorylation from the mitochondria. During a single heartbeat, approximately 2% of the cellular ATP is consumed and the whole ATP pool of cardiac myocytes is turned over within less than a minute

[178, 179]. Two of the most important regulatory factors that orchestrate oxidative phosphorylation in response to the constant change in work by the heart are Ca2+ and ADP [180, 181].

2+ 2+ The rise in cytosolic free Ca concentration ([Ca ]c) causes the activation of the contractile proteins. The cytosolic Ca2+ influx is due to Ca2+ entering the cytosol from two sources, one is the pool of Ca2+ that enters via the sarcolemma

L-type Ca2+ channels (LTCC), the other is the Ca2+ release from the SR, these events are referred to as calcium induced calcium release (CICR). The relaxation occurs by Ca2+ reuptake into the intracellular store by SR Ca2+ pumps (SERCA) and efflux by sarcolemma Na+/ Ca2+ exchange. The SR is not the only organelle

2+ that plays a role in reducing the [Ca ]c, the mitochondria is also implicated in

2+ 2+ Ca signaling by buffering the [Ca ]c . The buffering properties of the mitochondria indicate the importance it has in regulation during Ca2+ dependent processes.

The effectiveness of the mitochondria in these events relies on its localization at sites of Ca2+ release, such as the ryanodine receptor (RyR), inositol 1,4,5-triphosphate receptor (IP3) and LTCC receptors. This allows for

77 spatial regulation to occur and for efficient signaling within a cardiac myocyte.

2+ 2+ The areas within a cell where high [Ca ]c is produced during Ca release, due to these receptors, is referred to as microdomains [182]. The mitochondria comprises a third of the cell volume and are aligned regularly along the ATP- consuming myofilaments. They are tightly connected to the Ca2+ stores of the cell, the SR and are highly co-localized in the vicinity of the dyads. The dyads are where activation of LTCC triggers Ca2+ release from the junctional SR (jSR)

[183].

During β-adrenergic stimulation the mitochondria takes up Ca2+ rapidly,

2+ 2+ with mitochondrial Ca ( [Ca ]m) transients peaking slightly earlier than cytosolic

Ca2+ transients [184], leading to an increase in the concentration of Ca2+ within the inter mitochondrial membrane (IMM) and thus an increase in ATP production

2+ [185, 186]. The increase in [Ca ]m of the IMM is added to the microdomain shared between the SR and the mitochondria. The RyRs on the SR release Ca2+ early during CICR when Ca2+ levels in this microdomain are very high compared to the Ca2+ within the bulk of the cytosol; this is what allows for an

2+ earlier [Ca ]m transient than in the cytosol [186]. These studies support earlier studies that conclude that rapid mitochondrial Ca2+ uptake shapes cytosolic Ca2+ transients [187]. In studies done by Mackenzie et al., inhibition of SERCA or inhibition of the mitochondria Ca2+ uptake led to an increase in the propagation of

2+ 2+ cytosolic Ca in the central regions of the cell, thus increasing [Ca ]c [188].

These studies further supported the idea that the mitochondria could serve as a

78 spatial buffer and thus, affect the magnitude and propagation of cytosolic Ca2+ transients.

Voltage-dependent anion channel (VDAC), a well-characterized Ca2+ channel in the OMM governs Ca2+ transmission from the SR to the mitochondria

[189]. The VDAC is physically linked to the SR Ca2+ release channel, inositol

1,4,5-triphosphate receptor (IP3R), by the tethering protein glucose-regulated protein 75 (grp75) [20]. VDAC has been shown to also co-localize with RyR2 in the SR-mitochondrial microdomain [190]. Min et al. were unable to conclude if the interaction between RyR and VDAC was direct or if it is mediated by grp75 or by an as yet unidentified tethering protein. The tethering protein would help create the microdomain between the SR and OMM. The areas of the OMM that are in close contact to the SR are also the areas within the mitochondria that are closest to the IMM, these areas also have the highest abundance of VDAC [191].

These results suggest that the described contact points function as anchorage sites for the SR-mitochondrial physical coupling, and that close coupling of the

SR, OMM, and IMM is likely to provide a favorable spatial arrangement for local

RyR2-mitochondrial Ca2+ signaling.

Our data suggests that there is another tethering protein within this microdomain in the form of the AKAP β-synemin. Classically β-synemin is characterized as an intermediate filament (IF). A key role of IFs is to mediate the mechanochemical link between the contractile apparatus and the mitochondria, among other organelles. This linkage of all the membranous structures to the contractile apparatus occurs mainly through the Z-disks. This linkage supports

79 energy demands for the working myocyte to be able to withstand the EC coupling requirements [121]. Synemin is co-expressed and forms copolymers with desmin, a well-characterized muscle-specific IF protein [12]. Studies in desmin null cardiomyocytes lead to the loss of proximity of the mitochondria to other organelles such as the SR [16]. The compartment where these two organelles interact is referred to as the mitochondria associated ER membrane (MAM), and this is where the Ca2+ microdomains decribed above exist [192] (Fig. 4.1). It is not known whether desmin binds to the mitochondria directly or if it is linked through a desmin associated protein such as β-synemin. IFs do appear to have a structural function in positioning the mitochondria in areas of high-energy demand and Ca2+ cycling. The ubiquitous presence of these cytoskeletal proteins at such key locations clearly leaves open the possibility that they might take on additional roles in important signaling cascades.

The characterization of β-synemin as an IF and an AKAP would make it an appropriate candidate to be positioned at the MAM. Through phosphorylation studies we show that β-synemin anchored PKA does phosphorylate VDAC, indicating its close proximity to the mitochondria. In silico studies suggest a putative mitochondrial targeting sequence within the protein, supporting its association with the mitochondrial membrane. Finally our studies show that β- synemin anchored PKA plays a role in calcium handling within HL-1 cells. This establishes a functional role for β-synemin within the Ca2+ microdomain at the

MAM.

P SERCA T P t C C K s L L H i P P

n C

o A D g V K

A C BR Gq IP3R SERCA D 3 NT C P I 6p 6Shc A

IP3R ROS ETC PML PP2a 2+ KT S Ca PMCA A grp75 IM

2+ a IP3R VDAC CU 2+ C M e a l C c LX y NC U1 C MIC + VOC s a + b N 2 e Ca r SERCA K RyR M1 SERCA ET + L H ROC 2+ Ca grp75 AKT 2+ Ca C A D IP3R V ix atr T 2+ M a + N C NCX H A C + A a A D N C P + V S 2 n ERp44 M grp75 + 2 IP3R a C PLASMA MEMBRANE C - + i A olg D G tus BIP/grp78 V V - + ara m pp 180 - + A Sig-1R M - OM - + - + MFN-2 - MFN-1 + - +

ER MAMs MITOCHONDRION

Figure 4.1 – Schematic representation of intracellular Ca2+ dynamics and MAMs proteins involved in ER- mitochondria Ca2+ cross-talk. A series of protein localized in MAM (VDAC, grp-75) regulate Ca2+ release from the ER and an efficient mitochondria Ca2+ uptake, resulting in different functional outcomes. Mitochondrial surface directly interacts with the ER through contact sites defining hotspot Ca2+ signaling units. Mitochondria Ca2+ (red dots) import and export occur from the matrix. Allowing Ca2+ levels to return to resting conditions through a series of channels and pumps. Patergnani S, Suski JM, Agnoletto C, Bononi A, Bonora M, De Marchi E, Giorgi C, Marchi S, Missiroli S, Poletti F et al: Calcium signaling around Mitochondria Associated Membranes (MAMs). Cell communication and signaling : CCS 2011, 9:19 [58]

4.2 Experimental Procedures

4.2.1 Generation of expression constructs and site-directed mutagenesis

The human β-synemin cDNA, cloned into the eukaryotic expression vector pCDNA3, was kindly provided by Dr. Li (Biologie Moléculaire de la

Différenciation, Université Denis-Diderot Paris 7) [24]. The β-synemin fragment was subcloned in-frame into the mammalian expression vector pECFP-N1

(Clontech, Mountain View, CA, USA), using Hind III and Bam HI restriction sites

(ECFP- β-synemin-WT). The QuikChange® site-directed mutagenesis kit (Agilent

Technologies, Santa Clara, CA, USA) was used to introduce mutations into the

PKA binding region of full-length β-synemin cDNA. This resulted in cDNA- directed expression of a non-RII binding β-synemin with two proline substitutions

(I640P and L645P), referred to as ECFP- β-synemin-P [29].

4.2.2 Culture and transfection of HL-1 cardiomyocytes

HL-1 cardiomyocytes were kindly provided by Dr. William C. Claycomb of the Department of Biochemistry and Molecular Biology, Louisiana State

University, New Orleans, LA, USA [139]. Cells were maintained in Claycomb medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% FBS

(Sigma-Aldrich, St. Louis, MO, USA), 0.1 mM norepinephrine (Sigma-Aldrich, St.

Louis, MO, USA), 100 units/mL penicillin-streptomycin (Life technologies, Grand

Island, NY, USA), and 2 mM L-glutamine (Life Technologies, Grand Island, NY,

USA). The medium was changed approximately every 24 hrs. Cells were grown at 37°C in 5% CO2 and 95% humidity. .

Madison, WI, USA) via manufacturer’s protocol.

4.2.3 Sequence analysis

The amino acid sequences of mitochondrial AKAPs D-AKAP1

(NP_003479.1) and SKIP were aligned with β-synemin (this region is common to both synemin isoforms, but the specific sequence used was from β-synemin

,CAC838581) using the multiple sequence alignment feature of MacVector 12.7.1

(MacVector Inc, Cary, NC, USA). MacVector settings included ClustalW multiple sequence alignment; matrix: blossom, open gap penalty: 40, extended gap penalty: 0.05, pairwise alignment mode: slow, delay divergent: 50% gap separation distance: and 8, with residue specific and hydrophilic penalties.

MEME software was used for consensus sequence generation. MEME setting included one motif per sequence, and a motif length of 24 amino acids was specified [155, 164, 165].

4.2.4 Activation of PKA through stimulation of the β-adrenergic pathway

In order to activate PKA, cells were treated with the β-adrenergic agonist isoproterenol (10 µM, Cell signaling, Boston, MA, USA) for 10 m 48 h after transfection. To prevent dephosphorylation , the phosphatases in the cells were inhibited with a PP1 and PP2A inhibitor, calyculin A (Cell signaling, Boston, MA,

USA), for 20 m prior to isoproterenol treatment. Cells were either lysed and used

83 for two-dimensional (2D) analysis or plated on a 96-well plate and used for Fluo-

4 Direct™ Calcium Assay.

4.2.5 PKA inhibition

In order to inhibit both PKA type I and type II cells were treated with PKA type I and type II inhibitor N-[2-(p-bromocinnamylamino)ethyl]-5- isoquinolinesulfonamide (H89, 20 µM, Cell signaling, Boston, MA, USA) for 30 m

48 h after transfection. To inhibit PKA type I, cells were treated with 8- chloroadenosine-3’, 5’-cyclic monophosphorothioate ((Rp)-8-Cl-cAMPS), 10 µM,

Axxora, Farmingdale, NY, USA) for 30 m 48 h after transfection. To inhibit PKA type II, cells were treated with 8-Piperidinoadenosine-3’,5’-cyclic monophosphorothioate ((Rp)-8-PIP-cAMPS), 90 µM, Axxora, Farmingdale, NY,

USA) for 30 m 48 h after transfection. PKA pathway was activated as stated above with isoproterenol. Cells were either lysed and used for phosphorylation studies via Western blot or plated on a 96-well plate, prior to transfection and treatment, and used for Fluo-4 Direct™ Calcium Assay (Life Technologies,

Grand Island, NY, USA).

4.2.6 Analysis of phosphorylation substrates of β-synemin anchored PKA, via

Western blot analysis

Six 60 mm plates transfected with β-synemin-WT or β-synemin-P were serum-starved for 4 h prior to treatment. Cells were treated with a PKA or cAMP inhibitor (H89, (Rp)-8-PIP-cAMPS or (Rp)-8-Cl-cAMPS) and either not stimulated or stimulated with isoproterenol as indicated above. Cells transfected with empty vector pECFP-N1 served as controls. After treatment, cells were lysed in cell

84 lysis buffer (10 mM Tris-HCL pH 7.5, 300 mM NaCl, 1 % Triton-X 100, 2 mM

EDTA, 1X Halt protease inhibitor and 1X Roche phosphatase inhibitor). Cells were vortexed twice for 10 s and incubated on ice for 10 m. Cells were vortexed once more for 10 s and centrifuged at 14,000 rpm for 20 m at 4°C. The protein concentration was measured using Direct Detect (Millipore, Billerica, MA, USA)

Western blotting samples were prepared in 6X sample buffer (Pierce, Rockford,

IL, USA). Gel electrophoresis of proteins was performed on 4-15% gradient Mini-

PROTEAN® TGX™ Precast Gels (Bio-rad, Hercules, CA, USA) following manufacture’s procedure. Separated proteins were then transferred onto nitrocellulose membrane at 100 V for 20 m. Nonspecific protein binding sites on the nitrocellulose membranes were blocked with 10 % nonfat dry milk in TBST

(25 mM Tris, .15M NaCl, .05% Tween-20, pH 7.5) for one hour at room temperature. The membrane was washed three times for 30 minutes with TBST.

The primary antibody that detects PKA phosphorylated substrates that contain a phospho-Ser/Thr residue with an arginine at the -3 and -2 positions (Phospho-

PKA-Substrate, Cell signaling, Boston, MA, USA) was diluted 1:1000 in 5% bovine serum albumin-TBST (BSA, Sigma-Aldrich, St. Louis, MO, USA) and incubated with the membrane overnight at 4°. After three 15 m washes in TBST, membranes were incubated with HRP conjugated-goat anti-rabbit (Santa Cruz

Biotechnolog Inc., Dallas, Texas, USA) diluted 1:10,000 in 5% nonfat milk-TBST.

Membranes were incubated for 1 h at room temperature. The incubation was followed by three washes in TBST and one final wash in TBS, the peroxidase

85 activity of the nitrocellulose-bound-secondary antibody was detected by Pierce

ECL Western Blotting Substrate (Pierce, Rockford, IL, USA).

Statistical analysis: Collected data were analyzed using Microsoft Excel

2011 and Stat Plus. For an evaluation of statistical significance a Student’s t-test was used. A probability of p<0.05 was considered to represent a significant difference. Protein bands were normalized to GAPDH and differences compared by one-tailed T-test. Error bars represent means +/- s.e.m

4.2.7 Co-immunoprecipitation studies for two-dimensional (2D) gel analysis

Thirty 100 mm plates transfected with β-synemin-WT were serum-starved for 4 h prior to treatment. Cells were stimulated with isoproterenol as indicated above. After treatment, cells were lysed in 500 µL of cell lysis buffer (10 mM Tris-

HCL pH 7.5, 300 mM NaCl, 1 % Triton-X 100, 2 mM EDTA, 1X Halt protease inhibitor and 1X Roche phosphatase inhibitor). Cells were vortexed twice for 10 s and incubated on ice for 10 m. Cells were vortexed once more for 10 s and centrifuged at 14,000 rpm for 20 m at 4°C. The protein concentration was measured using Direct Detect (Millipore, Billerica, MA, USA). Anti-phospho-PKA- substrate antibody (specifically made in PBS, Cell signaling, Boston, MA, USA) was covalently coupled to Dynabeads® following the manufacture’s protocol using the Dynabeads® Co-Immunoprecipitation Kit (Life Technologies, Grand

Island, NY, USA) The elutes were pooled and concentrated and the buffer was exchanged with rehydration/sample buffer (ReadyPrep™ 2-D Starter Kit, Bio-rad,

Hercules, CA, USA) using Amicon Ultra-2 mL centrifugal filters (Millipore,

Billerica, MA, USA) and analyzed by 2D.

4.2.8 2D analysis

Proteins phosphorylated by PKA, which were co-IPed as described above

(250 µg/strip), were diluted in ReadyPrep™ 2-D Starter Kit rehydration/sample

Buffer (Bio-rad, Hercules, CA, USA) and loaded onto 11-cm immobilized pH gradient (IPG) strips, pH 3-11 (ReadyStrip™ IPG strip, Bio-rad, Hercules, CA,

USA) and rehydrated over night at room temperature. The first-dimension separation was performed in a Protean isoelectric focusing cell (Bio-rad) at 250 V for 15 minutes, followed by an increase to 8000 V for 30kVh. After the first- dimension separation, gel strips were rinsed and equilibrated according to the manufacturer’s protocol. IPG strips were then loaded onto a 4-15% Criterion TGX precast gel (Bio-rad), and proteins were subjected to electrophoresis at 200 V for

45 minutes. Gels were used for either protein spot analysis or for Western blotting. Gels for protein spot analysis were stained with Pierce Silver Stain Kit for Mass Spectrometry (Pierce, Rockford, IL, USA) per manufacturer’s protocol.

Gels for Western blot analysis followed same protocol as stated above using the anti-Phospho-PKA substrate antibody. The selected spots were excised from the silver-stained gels and prepared for analysis using liquid chromatography tandem mass spectrometry (LC-MS/MS).

4.2.9 LC-MS/MS sample preparation

The excised 2D spot gel pieces were destained following the manufacturer’s protocol. Pieces were washed four times with water, once with

200 mM ammonium bicarbonate, and twice with 100 mM ammonium bicarbonate in 50% acetonitrile, then dried under vacuum using a Savant SpeedVac

(Instruments Inc., Farmingdale, NY, USA). To each gel piece, 200 µL of 100 mM ammonium bicarbonate (pH 8.0) containing 0.5 µg of trypsin (Promega, Madison,

WI, USA) was added and incubated overnight at 37°C. Peptides in each gel piece were extracted with three washed of 70% acetonitrile and 0.1% formic acid.

The extracts were then dried. Peptides were resuspended in 20 µL of 6 M guanidine-HCl in 5 mM potassium phosphate and 1 mM DTT (pH 6.5) was added to each dried sample and subsequently sonicated. Peptides were then extracted using a C18 ZipTip (Millipore, Billerica, MA, USA) and subjected to LC-MS/MS analysis by Lerner Research Institute Mass Spectrometry Laboratory for protein sequencing at The Cleveland Clinic Foundation [193].

LC-MS/MS Spectrometry: The LC-MS system was a Finnigan LTQ-

Orbitrap Elite hybrid mass spectrometer system. The HPLC column was a

Dionex 15 cm x 75 µm id Acclaim Pepmap C18, 2µm, 100 Å reversed- phase capillary chromatography column. Five µL volumes of the extract were injected and the peptides eluted from the column by an acetonitrile/0.1% formic acid gradient at a flow rate of 0.25 µL/min were introduced into the source of the mass spectrometer on-line. The microelectrospray ion source is operated at 2.5 kV.

The digest was analyzed using the data dependent multitask capability of the instrument acquiring full scan mass spectra to determine peptide molecular weights and product ion spectra to determine amino acid sequence in successive instrument scans. This mode of analysis produces approximately 15000 collisionally induced dissociation (CID) spectra of ions ranging in abundance over several orders of magnitude.

Data analysis: The data was analyzed using all CID spectra collected in the experiment to search the NCBI non-redundant database with the search program Mascot using a human taxonomy filter. All matching spectra were verified by manual interpretation. The interpretation process was aided by additional searches using the programs Sequest and Blast as needed.

4.2.10 Calcium indicator assay

HL-1 cells were seeded into black-walled clear-bottomed 96-well plate at a density of 20,000 cells/well. Cells transfected with β-synemin-WT or β-synemin-P were serum-starved for 4 h prior to treatment. Cells were treated with a PKA inhibitor ((Rp)-8-PIP-cAMPS or (Rp)-8-Cl-cAMPS) and either left unstimulated or stimulated with isoproterenol as indicated above. Cells transfected with empty vector pECFP-N1 served as controls. Before initiating the screen, the medium was removed and the cells were washed twice with assay buffer containing 2 mM

CaCl2, 138 mM NaCl, 5.33 mM KCl, 0.34 mM Na2HPO4, 0.44 mM KH2PO4, 4.17 mM NaHCO3, 5.56 mM d-glucose, and 20 mM HEPES at pH 7.4. The cells were then incubated in assay buffer supplemented with 100 µl of Fluo-4 no-washTM calcium assay dye solution (Life technologies, Grand Island, NY, USA) for 30 min at 37°C according to the manufacturer's protocol. After incubation at 37°C, the cells were allowed to equilibrate to room temperature for 10 min before initiation of the assay. Changes in Fluo-4 fluorescence in response to treatment with inhibitors and/or isoproterenol were measured at 538 nm using a Beckman

Coulter DTX880 multimode detect (Beckman Coulter, Brea, CA, USA) with excitation at 485 nm and emission cut-off at 530 nm to minimize background

89 fluorescence. Baseline fluorescence was measured for 20 s before the addition of agonist and isoproterenol to the assay plate, and the peak fluorescence was measured 10 m after addition of the isoproterenol.

Data Analysis: Measurements are given in relative fluorescent units

(RFU) as the maximum response minus the minimum response divided by the minimum response.

4.3 Results

4.3.1 Phosphorylation substrates of β-synemin anchored PKA

To evaluate the phosphorylation substrates of β-synemin anchored PKA, whole cell lysates from cells treated with and without isoproterenol were subjected to Western blot analysis. A phospho-PKA substrate antibody that specifically recognizes residues that contain a phospho-Ser/Thr with an arginine at the -3 and -2 positions which is the phosphorylation site specific to PKA

(RRXS*/T*) was utilized. To verify the specificity of the phospho-PKA substrate antibody for proteins phosphorylated via the β-adrenergic pathway, a variety of proteins of different molecular weights were identified upon isoproterenol treatment compared with untreated cells, for example phospholamban at ~8 kDa, troponin I at ~ 24 kDa and myosin binding protein C at ~142 kDa (Fig. 4.2 lane 1 versus lane 2, cells are transfected with empty vector and serve as controls).

These proteins are the major substrates of PKA in cardiomyocytes commonly seen in phosphorylation studies when the β-adrenergic receptor is stimulated.

To determine if β-synemin anchored PKA affected the phosphorylation of

any phosphos-proteins, cells were transfected with β-synemin-P (lanes 3 and 4)

or with β-synemin-WT (lanes 5 and 6). Protein phosphorylation patterns were

then compared to untreated (lanes 3 and 5) and treated (lanes 4 and 6) and

between β-synemin-P to β-synemin-WT, to observe if there were any differences.

Four bands were identified to have differences of phosphorylation due to β-

synemin anchored PKA. The bands were identified at molecular weights of 15,

35, 37 and 130 kDa.

A. B.

142 – cMyBP-C 130

37 35 24-TNI

15 8-PLB

C-I C+I P-I P+I WT-I WT+I C-I C+I P-I P+I WT-I WT+I

GAPDH

Figure 4.2 – Identification of !-synemin anchored PKA phosphorylated substrates. To determine if !- synemin anchored PKA had an affect on the phosphorylation of PKA substrates. HL-1 cells were transfected with !-synemin-WT (WT) or !-synemin-P (P). PKA was activated via isoproterenol (+I) and compared to cells where PKA was not activated (-I). Comparing bands from phosphor proteins expressing !-synemin-P (PKA is not anchored) to those cells expressing !-synemin-WT (PKA is anchored) revealed that the phosphorylation state of four proteins were effected. Compare the bands in P+I to WT+I at 15,35,37, and 130 kDa. Cells transfected with empty vector and either not treated or treated (C-I and C+I, respectively) served as controls to show that PKA was activated, notice the protein bands at phospholamban (PLB) 8 kDa, troponin I (TNI) 24 kDa, and myosin binding protein (cMyBP-C) 142 kDa. GAPDH was used to verify equal loading. A. is a 3 minute exposure and B. is a 7 min exposure

4.3.2 Identification of phosphorylated substrates of β-synemin anchored PKA

To identify the four proteins suspected of being phosphorylated by β- synemin anchored PKA, the phospho-PKA substrate antibody was used to IP the phosphorylated proteins from whole cell lysates transfected with β-synemin-WT and treated with isoproterenol. The IP was followed by 2D electrophoresis and the proteins were detected by silver staining and Western blot analysis.

The four protein spots were excised from the silver stain (Fig 4.3) and analyzed by LC-MS/MS. They were identified as plakoglobin and desmoplakin

(Desmosomes, 130 kDa), voltage-dependent anion channel 1 (VDAC, 31 kDa), voltage-dependent anion channel 2 (VDAC, 32 kDa) and superoxide dismutase

[Cu-Zn] (15 kDa).

4.3.3 β-Synemin modulates PKA phosphorylation of VDAC

Previously we had shown that β-synemin is a dual AKAP, capable of anchoring both PKA type I and PKA type II. Therefore we used several PKA inhibitors to examine whether the phosphorylation of VDAC was due to the action of PKA type I or PKA type II. Under normal conditions VDAC phosphorylation increases above basal levels only when PKA is bound to β-synemin (Fig. 4.4, lane 5 verses lane 6). This is in contrast to a slight decrease in phosphorylation upon stimulation when PKA is not bound to β synemin (Fig. 4.4, lanes 3 verses

4).

H89 has been reported to be a highly specific PKA inhibitor with little effect on the activity of protein kinase C and other protein kinases [194]. The inhibitory

210 kDa

130

37 35

10 kDa Acidic Basic 3 11

Figure 4.3 – Separation of !-synemin anchored PKA substrates using two-dimensional gel effect electrophoresis (2D-PAGE). HL-1 cells transfected with !-synemin-WT and treated with isoproterenol were lysed and used in a immunoprecipitation assay utilizing the anit-phospho-PKA substrate antibody. Proteins were separated based on their charge and molecular weights. Arrows indicate the 4 unique spots that are phosphorylated by !-synemin anchored PKA. Protein spots were excised and identified by LC-MS/MS. 15 kDa – superoxide dismustase [Cu-Zn], 35 – voltage dependent anion channel 2 (VDAC), 37 – voltage dependent anion channel 3 (VDAC), 130 – plakoglobin and desmoplakin.

93 of H-89 is due to its competitive binding to the ATP pocket on the kinase catalytic subunit [195]. When cells are pretreated with H89 before treatment with isoproterenol, phosphorylation of VDAC is blocked (Fig. 4.5). Confirming that

PKA is necessary to phosphorylate VDAC.

To attempt to determine which class of PKA is involved in phosphorylation of VDAC (based on our knowledge that β-synemin is a dual

AKAP and thus anchors PKA type I and type II), we employed two potent membrane-permeable and metabolically stable inhibitors of PKA type I and type

II, (Rp)-8-Cl-cAMPS and (RP)-8-PIP-cAMPs. These inhibitors occupy specific cAMP-binding sites on the regulatory subunit of PKA thus preventing the holoenzyme dissociation and, consequently, kinase activation. (Rp)-8-Cl-cAMPS has been previously reported to function as a site-selective competitive inhibitor of PKA with preference for type I PKA [196, 197]. Wojtal et al. showed that the

(Rp)-8-Cl-cAMPS inhibitor prevents dissociation of the RI/C holoenzyme induced by forskolin, whereas it does not prevent forskolin stimulated dissociation of RII/C holoenzymes.

In our studies, cells were pretreated with (Rp)-8-Cl-cAMPS before treatment with isoproterenol (PKA type II is active and PKA type I is inhibited),

When PKA type I was inhibited the increase in phosphorylation of VDAC in the β- synemin-WT cells (Fig. 4.6 lanes 5 and 6) was similar to cells with both classes of PKA active (no inhibition) (Fig 4.4, lanes 5 and 6). This indicates that VDAC could be phosphorylated upon isoproterenol stimulation when only PKA type II is active.

To analyze the effects of phosphorylation of PKA type I while PKA type II was inhibited, cells were pretreated with (Rp)-8-PIP-cAMPS before treatment with isoproterenol. The phosphorylation of VDAC in the cells expressing β- synemin-WT (Fig. 4.7 lanes 5 and 6) did not increase when PKA was activated upon isoproterenol stimulation. As in the experiments in which PKA type I was inhibited (and type II was active), phosphorylation occurred in cells expressing β- synemin-P upon stimulation (Fig. 4.7 lanes 3 and 4).

'#$" '" &#$" +4.34 -.875 +1.19 &" %#$" * ""/" %" * " !#$" VDAC-P/GAPDH !"

Relative densitometry units ()*" (+*" ,)*" ,+*" -.)*" -.+*"

VDAC-P

GAPDH

Figure 4.4 – Effects of !-synemin anchored PKA on VDAC phosphorylation. HL-1 cells were transfected with either empty vector (C), !-synemin-P (P) or !-synemin-WT (WT) and treated without (-I) or with (+I) isoproterenol. VDAC phosphorylation was detected using phospho-specific PKA phosphorylation specific antibody The data shows that VDAC is phosphorylated by PKA upon isoproterenol stimulation (C-I compared to C+I). When PKA is not anchored to !-synemin phosphorylation of VDAC is slightly decreased upon isoproterenol treatment (P-I compared to P+I). In cells expressing !-synemin-WT VDAC is phosphorylated above the basal level seen in control cells at rest (WT-I compared to C-I) and upon isoproterenol is phosphorylated to the same degree as control (WT+I compared to C+I). The fold change in phosphorylation upon stimulation is shown above each set of cells (+4.34, -.875, +1.19). GAPDH was used as loading control and densitometry analysis on three independent experiments. A probability of p<0.05 was considered to represent a significant difference (*). Protein bands were normalized to GAPDH and differences compared by one-tailed T-test. Error bars represent means +/- SEM

VDAC

C-I C+I P-I P+I WT-I WT+I

GAPDH

Figure 4.5 – Effect of !-synemin anchored PKA on VDAC phosphorylation under H-89 inhibition. HL- 1 cells were transfected with either empty vector (C), !-synemin-P (P) or !-synemin-WT (WT) and treated without (-I) or with (+I) isoproterenol. VDAC phosphorylation was detected using phospho- specific PKA phosphorylation specific antibody.This demonstrates that phosphorylation of VDAC is dependent on PKA. The band at 37 kDa is no longer visible under these conditions (as compared to figure 4.2 B), indicated by the arrow. To support this 50 µg of protein was loaded instead of the standard 15 µg. The blot was also exposed for 30 minutes. GAPDH was used as loading control.

'#$" '" &#$" +1.26 +1.3 +1.52 &" %#$" ""/" /" %" /" !#$" VDAC-P/GAPDH !" ()*" (+*" ,)*" ,+*" -.)*" -.+*" Relative densitometry units

VDAC-P RII active/RI inhibited

GAPDH

Figure 4.6 - Effects of !-synemin anchored PKA on VDAC phosphorylation with (Rp)-8-Cl-cAMPS treatment (RII is active and RI is inhibited). HL-1 cells were transfected with either empty vector (C), !- synemin-P (P) or !-synemin-WT (WT) and treated without (-I) or with (+I) isoproterenol. VDAC phosphorylation was detected using phospho-specific PKA phosphorylation specific antibody. The significant increase in phosphorylation in control cells upon stimulation demonstrates that VDAC phosphorylation by PKA upon isoproterenol stimulation can occur when only PKA type II is active (C+I vs. C-I) in the system. When PKA is not anchored to !-synemin, phosphorylation of VDAC is increased upon isoproterenol treatment (P-I compared to P+I). In cells transfected with !-synemin-WT, VDAC is phosphorylated upon isoproterenol treatment (WT-I compared to WT+I). The fold change in phosphorylation upon stimulation is shown above each set of cells (+1.26, +1.30, +1.52). GAPDH was used as loading control and densitometry analysis on three independent experiments. A probability of p<0.05 was considered to represent a significant difference (*). Protein bands were normalized to GAPDH and differences compared by one-tailed T-test. Error bars represent means +/- SEM

'#$" '" &#$" +2.70 +1.67 +0.90 &" /" %#$" /" %"

VDAC-P/GAPDH !#$" !"

Relative densitometry units ()*" (+*" ,)*" ,+*" -.)*" -.+*"

VDAC-P RI active/RII inactive

GAPDH

Figure 4.7 - Effects of !-synemin anchored PKA on VDAC phosphorylation with (Rp)-8-PIP-cAMPS treatment (RI is active RII is inhibited). HL-1 cells were transfected with either empty vector (C), !- synemin-P (P) or !-synemin-WT (WT) and treated without (-I) or with (+I) isoproterenol. VDAC phosphorylation was detected using phospho-specific PKA phosphorylation specific antibody. The significant increase in phosphorylation upon stimulation demonstrates that VDAC is phosphorylated by PKA upon isoproterenol stimulation when only PKA type I is active (C-I vs C+I). When PKA is not anchored to !-synemin, phosphorylation of VDAC is increased upon isoproterenol treatment (P-I compared to P+I). In cells transfected with !-synemin-WT, there is no change in VDAC phosphorylation upon isoproterenol treatment (WT-I compared to WT+I). The fold change in phosphorylation upon stimulation is shown above each set of cells (+2.70, +1.67, +0.90). GAPDH was used as loading control and densitometry analysis on three independent experiments. A probability of p<0.05 was considered to represent a significant difference (*). Protein bands were normalized to GAPDH and differences compared by one-tailed T-test. Error bars represent means +/- SEM

4.3.4 Identification of a mitochondrial signaling motif within β-synemin

Following the identification (through 2D SDS-PAGE and LC-MS/MS

analysis) of VDAC as a substrate of β-synemin anchored PKA, protein sequence

alignment analyses were performed. Since VDAC is a mitochondrial protein it is

proposed that β-synemin is also localized to the mitochondria. AKAPD1 and

SKIP are two AKAPs known to be concentrated at the mitochondria membrane.

Aligning the mitochondrial targeting sequences of both of these proteins with β-

synemin allowed us to identify a putative mitochondrial targeting sequence at

amino acids 330-360 (Fig. 4.8).

Figure 4.8 - Sequence comparison with !-synemin of the mitochondrial targeting sequence between known mitochondrial targeted AKAPs, AKAP1 and SKIP. The proposed mitochondrial targeting domain of synemin is located at 330-360 aa. Blocked amino acids are conserved among the sequences.

4.3.5 Determination of internal calcium in respect to β-synemin anchored PKA

Fluo-4 AM is a fluorescent Ca2+ indicator dye used to measure changes

in intracellular Ca2+ in response to a variety of experimental treatments, including

agonist-stimulation and antagonist-inhibition. Upon binding to Ca2+, Fluo-4

fluorescence is enhanced up to 100-fold, making it easy to track intracellular

calcium mobilization in dye-loaded cells. Fluo-4 AM freely crosses over the cell

100 membrane, as the AM moiety in ester-linkage renders the Fluo-4 molecule uncharged. Once inside the cell, however, endogenous esterases hydrolyze the

AM ester linkage, yielding free Fluo 4, a fluorescent dye bearing negatively- charged carboxyl groups that both trap it within the cell, and allow it to bind to free cytosolic Ca2+ [198].

To evaluate if the β-synemin anchored PKA plays a role in the regulation

2+ of internal calcium ([Ca ]i) levels, HL-1 cells were transfected with either β- synemin-P or β-synemin-WT. The cells were allowed to incubate and then

2+ treated with Fluo-4 AM, [Ca ]i and were monitored 10 m after the addition of

2+ isoproterenol. Activation of PKA by isoproterenol resulted in a rise in [Ca ]i in control cells transfected with empty vector (Fig. 4.9, blue columns 1 and 2),

2+ whereas the [Ca ]i was decreased upon stimulation in β-synemin-P cells (Fig.

2+ 4.9 blue columns 3 and 4). The increase in [Ca ]i upon stimulation was similar to control cells in cells expressing β-synemin-WT transfected (Fig. 4.9 blue columns

5 and 6).

To specifically evaluate whether PKA type I or PKA type II bound to β-

2+ synemin had an effect on the [Ca ]i levels, HL-1 cells were pretreated with (Rp)-

8-Cl-cAMPS or (Rp)-8-PIP-cAMPS before isoproterenol stimulation. Cells in which only PKA type II was active and PKA type I was inhibited exhibited an

2+ increase in [Ca ]I upon isoproterenol treatment similar to the control without inhibition, but to a much lesser degree (Fig. 4.9, red columns 1 and 2 compared to blue columns 1 and 2). When PKA type II was not bound to β-synemin (cells

2+ expressing β-synemin-P) and PKA type I is inhibited, [Ca ]I are elevated upon

101 isoproterenol stimulation (Fig. 4.9, red columns 3 and 4). When PKA type II was bound to β-synemin (cells expressing β-synemin-WT) and PKA type I is inhibited,

2+ [Ca ]i decrease significantly upon isoproterenol stimulation (Fig.4.9, red columns 5 and 6).

In the opposite experiments where PKA type I was active and PKA type II was inhibited, identical results were observed in the control cells as in the other two experiments (Fig. 4.9 green columns 1 and 2). When PKA type I was not

2+ bound to β-synemin (and PKA type II was inhibited) [Ca ]I were elevated upon isoproterenol stimulation (Fig. 4.9, red columns 3 and 4). When PKA type I is

2+ bound to β-synemin (PKA type II was inhibited) [Ca ]i a decrease is observed upon isoproterenol stimulation but to a lesser extent than in cells in which only

PKA type II was active (Fig.4.9, red columns 5 and 6).

!"#$"%#&"'"# 102

.)*#

54# .# * * ()-# 6$78/-/!9/:;<2=#62>;# '?7@#00#":A'B@C2>;#'?7@#0# (),# * !"#$ ADEAFA'@&8# 6$78/-/202/:;<2=#62>;# ()+# * * '?7@#0#":GB@H#2>;#'?7@#00# * * * * ADEAFA'@&8# ()*#

(# !/0# !10# 2/0# 210# 34/0# 3410#

Figure 4.9 – Intracellular [Ca2+] before and after isoproterenol treatment (-I/+I) in HL-1 transfected cells with either empty vector (C), !-synemin-P (P), or !- synemin-WT (WT). Before treatment with isoproterenol cells were either not treated (blue columns), or treated with (Rp)-8-Cl-cAMPS (red columns), or (Rp)-8- PIP-cAMPS (green columns). Measurements are given in relative fluorescent units (RFU) as the maximum response minus the minimum response divided by the minimum response. For an evaluation of statistical significance a Student’s t- test was used and a probability of p<0.05 was considered to represent a significant difference (*). Error bars represent means +/- SEM. (n=9 independent experiments). RFU (relative florescence units) = (max florescence – min florescence)/ min florescence.

103

4.4 Discussion

The primary finding of this study is the identification of VDAC as a substrate of β-synemin anchored PKA. Phosphorylation studies indicate that PKA type II phosphorylates VDAC, while PKA type I may play a role in the dephosphorylation of this protein. In silico studies identify an internal mitochondrial targeting sequence within β-synemin, indicating its localization at the mitochondrial outer membrane. Within this targeting sequence there is also a tubulin binding motif. Calcium handling experiments suggest that β-synemin anchored PKA plays an important role in the function of VDAC and its ability to shuttle Ca2+ across the OMM from the SR-mitochondria microdomain.

While the ability of β-synemin anchored PKA to phosphorylate VDAC indicates that there may be a direct binding between these two proteins, we cannot be sure of this at this time. However, based on our in silico analysis we hypothesize that β-synemin is located at the mitochondria outer membrane of the

MAM. Though the putative mitochondrial targeting sequence identified is within the sequence of β-synemin rather than at the commonly recognized N-terminal location, as most targeting and translocation signals of mitochondrial proteins are located [199, 200], it is not unheard of to have the sequence in this location.

About 30% of mitochondrial proteins lack they canonical N-terminal mitochondrial targeting sequences. For example Tom22 and heme lyase proteins both contain an internal mitochondrial targeting sequence that resembles the accepted N- terminal sequences [201, 202]. Further studies would need to be carried out to

104 incontrovertibly declare the 330-360 amino acid region of β-synemin as an actual internal mitochondrial targeting sequence.

To characterize this proposed localization sequence, a vector expressing just the mitochondrial targeting motif along with a GFP tag could be constructed and transfected into HL-1 cells to see if it localized to the mitochondria. The expressed protein could then be visualized using immunofluorescence coupled with confocal microscopy, as compared to a mitochondrial marker (Mitotraker).

To determine if this motif is necessary and/or sufficient to target the mitochondria, a GFP-synemin(Δ 330-360) containing GFP-tagged full length β- synemin lacking the 30 residues postulated to be the localization signal and (330-

360)synemin-GFP containing only the 30 mitochondrial localizing residues of β- synemin fused to GFP could be constructed and transfected into HL-1 cells [93].

We would hypothesize that the GFP-synemin(Δ 330-360) would lose it’s mitochondrial targeting and the (330-360) synemin-GFP would show characteristics of mitochondrial patterning.

Mutations in genes encoding Z-disk IFs or their interacting proteins have been identified as being responsible for cardiomyopathy [203]. An IF protein identified to play a significant role in these diseases is desmin, which categorizes most of these diseases as desminopatheis. The pathology of these diseases is characterized by myofibril disruption that initiates at the Z-disk. The absence of desmin in cardiomyocytes leads to dilated cardiomyopathy and heart failure characterized by mitochondrial defects [7, 16, 17]. Some of the mitochondrial abnormalities that occur are misdistribution throughout the cell, the number of

105 mitochondria available to the cell, as well as the mitochondrial morphology and disruptions in their functionality [16]. The binding of synemin to desmin is a well established interaction and thus supports the theory that β-synemin can be located at the mitochondria [10]. If desmin/synemin interactions are necessary for the proper function of synemin at the mitochondria, then a lack of desmin (as seen in desminopathies) will inevitably lead to a disruption of mitochondrial localized synemin activity. This raises the possibility that the symptoms of some desminopathies might proximally arise due to a loss of synemin function (or the function of a complex in which synemin is a member) with desmin itself, potentially playing what might be termed a ‘secondary,’ (but nonetheless, essential) scaffolding role.

This hypothesis is further supported with data from Maloyan et al. showing that mutations within αB-Crystalline (CryAB), a heat shock protein, has the ability to cause desmin-like cardiomyopathy [204]. CryAB is present in high concentrations in cardiac muscle and binds to desmin as well as VDAC. Recently a direct linkage between BAG3, a member of the Bcl-2-associated athanogene family, and CryAB has been shown to prevent protein aggregations and cell death in myofibrillar myopathies [205]. In a yeast two-hybrid screening we identified that β-synemin interact with both CryAB and BAG3 (data not shown), further supporting β-synemin as a key modulator of the mitochondria function.

Mutational studies within the β-synemin binding domain of desmin could lead to a better understanding of the role β-synemin plays in cardiomyopathies. Although loss of desmin function is an essential component of the pathology, it has not

106 explained these diseases completely. We can therefore postulate that a complex that includes BAG3, CryAB and VDAC is regulated by β-synemin anchored PKA in mitochondrial functions within cardiomyocytes. A possible role for this complex would be to buffer cytosolic Ca2+ during EC events.

Ca2+ is a vital intracellular second messenger that governs countless cellular functions. To maintain basal levels of cytoplasmic Ca2+ under various and ever-changing conditions, cells have evolved mechanism to carefully regulate

Ca2+ entry and removal [206, 207]. The sensitivity and response of cells to various stresses is dependent on the ability of cells to adequately sequester Ca2+ into their internal stores. Ca2+ is stored in intracellular organelles such as the SR and mitochondria. The transient increase in cytosolic Ca2+ concentration, which triggers the contractile response, has been intensively studied, but little is known about the dynamic changes in free Ca2+ concentration inside the SR and mitochondria, or in the microdomain they share. However, in recent years the understanding of this phenomenon is becoming less mysterious due to more sensitive equipment and more elegant experimental design.

There are three main pathways that elevate internal calcium concentrations; (1) an external calcium influx through the LTCC, which increase the sarcoplasmic calcium levels, (2) then, due to LTCC calcium influx subsequently triggering CICR events, the Ca2+ within the calcium stores of the

SR are released via the RyR, increasing the sarcoplasmic levels even further and

(3) the inability of the mitochondria to uptake calcium from the SR-mitochondrial microdomain through VDAC, will lead to increased sarcoplasmic calcium [208].

107

The studies presented here focused on the last of the these mechanisms.

The main roles of the mitochondria during EC events are; (1) to produce ATP

2+ 2+ and, (2) to buffer the elevated [Ca ]c, by allowing Ca into the matrix of the mitochondria through the IMM, via the OMM [209, 210]. This cytosolic Ca2+ buffering mechanism is possible due to the positioning of the mitochondria in close proximity to the cell membrane of the SR junctions. The microdomain between SR and mitochondria and its roles in Ca2+ transfer between the two membrane systems in the heart has been demonstrated [20, 211] but the functional linkage between the SR and mitochondria Ca2+ uptake remains to be resolved.

VDAC phosphorylation by PKA increases the on-rate of the C-terminal tail of tubulin binding to the channel with high specificity by two orders of magnitude.

This binding changes the channels preference for cations over anions [212-214].

The bound state favors cations with an anion-to-cation permeability ratio of 1:4, compared with the anion selectivity of the VDAC open state with a ratio of 7:3

[212]. This bound state allows for Ca2+ be transported across the OMM and makes it impermeable to ATP [215].

Due to the change in mechanics of VDAC upon phosphorylation, due to the binding of tubulin, it would be assumed that dephospohrylation of the channel is necessary for an on/off mechanism. Das et al. showed that VDAC phosphorylation is also modulated by glycogen synthase kinase (GSK-3β) [216].

They proposed that a possible mechanism of phosphorylation of VDAC by GSK-

3β is that it regulates the phosphatases that dephosphorylate VDAC, in other

108

words PKA phosphorylation results in β-tubulin binding to VDAC and an concomitant increase in affinity for Ca2+ allowing C2+a to pass through the channel; this is opposed by GSK-3β activity which phosphorylates and activates a phosphatase that dephosphorylates VDAC causing β-tubulin to dissociate and results in a decrease in affinity of the channel for Ca2+ This was shown in studies where GSK-3β was inhibited, and VDAC phosphorylation was decreased due to increased activity of phosphatases.

Excitingly, GSK-3β has been shown to be phosphorylated by PKA on serine 9 rendering it inactive [217]. This purposed activity for GSK-3β gives insight into our phosphorylation studies. These studies indicate that VDAC is phosphorylated when PKA is anchored to β-synemin due to the fact that phosphorylation increased upon stimulation in control cells and cells expressing wild type β-synemin but this was lost in cells where PKA was not anchored by β- synemin. We can also conclude that the phosphorylation of VDAC appears to be primarily due to the action of PKA type II. This is revealed in inhibition studies in which PKA type II is active and PKA type I is inactive in which a significant increase in phosphorylation of VDAC occurs upon stimulation when PKA type II is bound to β-synemin, but not when PKA type II is inhibited.

It appears that PKA type I could regulate dephosphorylation rather than phosphorylation of VDAC. When PKA type I was active and PKA type II was inhibited in cells expressing β-synemin-P, VDAC phosphorylation increased upon stimulation, even though PKA RI was not anchored by β-synemin and PKA type

109

II was not active. This suggests a possible role of PKA type I in regulating the phosphorylation state of VDAC. Our data indicate that PKA type I bound to β- synemin appears to either stay inactive until PKA type II also joins the complex or that PKA type I activates a phosphatase that acts on of VDAC. These data support that theory the enzymatic activity of type II PKA is the key isoform phosphorylating VDAC. Whether PKA type I is bound to β-synemin to regulate the phosphorylation of PKA type II on VDAC is still under investigation.

We can hypothesize that since; (1) GSK-3β is phosphorylated by PKA and

(2) that GSK-3β plays a role in the VDAC phosphorylation, that the phosphorylation of GSK-3β is carried out by PKA type I. This phosphorylation renders GSK-3β inactive and thus phosphatases active. This is concluded from the inhibition experiments in which either PKA type I or PKA type II were inactive.

In both sets of experiments, phosphorylation of VDAC occurred in the β-synemin-

P cells, but did not occur in the set of cells where both isoforms were active. In the studies in which only PKA type II was active, these results could be contributed to the fact that PKA type I was inhibited and so unable to phosphorylate GSK-3β, thus phosphatases were not active and not able to dephosphorylate VDAC. In the studies in which PKA type I was active, the results could be contributed to the fact that PKA type I was not bound to β-synemin and so was still unable to phosphorylate GSK-3β. This would leave the kinase phosphorylated and consequently leave phosphatases inactive thereby allowing

VDAC to remain phosphorylated. Further characterization of the phosphorylation sites of VDAC will be necessary to resolve this matter. Using antibodies specific

110 to VDAC and GSK-3β total protein levels and their phosphorylated sites would allow for better interpretation of these results. Also a more involved series of inhibition studies that include GSK-3β could reveal the mechanism of phosphorylation on VDAC by PKA and by GSK-3β.

Robert et al. showed that the mitochondria Ca2+ oscillations are modulated in frequency and amplitude by β-adrenergic stimulation. This demonstrates that mitochondrial Ca2+ transport is responsive to the most important stimuli that controls inotropic and chronotropic inputs. They went on to show that an increase of mitochondrial diastolic Ca2+ concentrations occurs when the extracellular calcium concentrations are increased allowing for a buffering of Ca2+ in the cell at rest, because of this it is possible that the mitochondria Ca2+ uptake and release might modify EC coupling and, thus, the mechanical activity of the cardiomyocyte

[209].

To evaluate the effect of β-synemin anchoring of PKA on calcium handling in HL-1 cells, the fluorescent calcium probe, Fluo-4 AM, was used to detect

2+ 2+ changes in intracellular free Ca ([Ca ]i) upon isoproterenol stimulation. The data indicates that β-synemin anchored PKA does play a role in modulating the levels of internal calcium during an EC event due to a β-adrenergic response.

2+ When β-synemin can not anchor PKA [Ca ]I are elevated at rest compared to

2+ control cells. Upon isoproterenol stimulation the [Ca ]I decrease. Relating this data with the phosphorylation data of VDAC, we can postulate that when PKA is unable to bind to β-synemin and phosphorylate VDAC, the pore stays in the

111

“open” conformation (i.e., low affinity for Ca2+). This prevents Ca2+ from being taken up into the mitochondria. The calcium data correlates with our phosphorylation studies with respect that VDAC phosphorylation decreases upon isoproterenol stimulation in cells expressing β-synemin-P, thus decreasing the ability of VDAC to take up Ca2+. This is the reason that in these cells, at rest,

2+ 2+ there is an elevated level of [Ca ]I and upon isoproterenol stimulation the [Ca ]I levels drop due to the mitochondria being unable to buffer the cytosolic calcium levels.

2+ In the calcium studies when either PKA type I or type II is inhibited, [Ca ]I increased slightly in the cells expressing β-synemin-P (thus PKA is not anchored to its AKAP). These result correlate to the phosphorylation studies in which

VDAC is phosphorylated upon isoproterenol stimulation, due to the increased phosphorylation and the “closing” of the pore, the mitochondria are able to uptake Ca2+ from the cytosol.

The data from the cells expressing β-synemin-WT cells are not as easy to

2+ interpret. For instance, when PKA type I was inhibited the [Ca ]I decreases even though the phosphorylation studies showed VDAC was phosphorylated (i.e. PKA type II active and PKA type I inactive). In cells in which PKA type II was active

2+ cells there is a dramatic decrease in [Ca ]I. As for the cells in which PKA type I

2+ was active, there was a decrease in [Ca ]I but not to the same extent as seen when the PKA type II was active. We can conclude that both PKA type I and PKA type II need to be anchored to β-synemin to have a response similar in magnitude as control cells at rest as well as upon isoproterenol stimulation.

112

Studies show that when the mitochondria are blocked from taking up Ca2+

, the cytosolic Ca2+ levels increase [181, 218]. This finding is parallel with Min et al. whose studies that showed that in HL-1 cells when VDAC is down-regulated

2+ there is an increase in diastolic [Ca ]i [190]. This leads to the theory that VDAC regulates mitochondrial as well as cytosolic Ca2+ levels in the heart. A study by

Subedi et al. also showed in HL-1 cells that an increased Ca2+ spark intensity, width and duration occurs when VDAC is knocked down [219]. We believe that the calcium concentrations are directly related to VDAC phosphorylation and that it is mainly due to the Ca2+ within the SR-mitochondria microdomain, but to further expand our exploratory results the source of the internal calcium increase and decrease needs to be confirmed. This can be done through organelle- specific Ca2+ sensitive indicators.

Our data point to the importance of PKA anchored to β-synemin in phosphorylating VDAC (Fig 4.10). It is clear that PKA type II is the isoform that initiates the phosphorylation of the channel, while PKA type I may initiates the dephosphorylation. Whether this has an acute effect on cardiomyoctye mechanics, such as influencing Ca2+ uptake into the mitochondria, is not clear, based upon the present data. As a better understanding of β-synemin’s effects on myocyte contractility promises to provide new insight into cardiac function in both healthy and diseased states, the study of this unique IF/AKAP remains an intriguing subject for future investigations.

113

Figure. 4-10 – Schematic representation of the proposed PKA anchored !-synemin in the mitochondria-associated SR membranes (MAM) Ca2+ microdomain. VDAC: voltage-dependent anion channel; CK:creatine kinase; IFs: intermediate filaments; IP3R; 1,4,5-triphosphate receptor; M: mitochondria; MAM: mitochondria-associtated ER membrane; PLB: phospholamban; RYR: ryanodine receptor; serca: sarcoplasmic reticulum calcium ATPase; SR: sarcoplasmic reticulum. Yassemi Capetanaki , Robert J. Bloch , Asimina Kouloumenta , Manolis Mavroidis , Stelios Psarras Muscle intermediate filaments and their links to membranes and membranous organelles Experimental Cell Research Volume 313, Issue 10 2007 2063 – 2076 http://dx.doi.org/10.1016/j.yexcr.2007.03.033

Chapter Five

Concluding Remarks

114

115

Synemin was identified as an AKAP in 2006, since then the function of it as an AKAP has been unclear. In these studies we were able to reveal the functional relevance of synemin as an AKAP and to tease apart the roles of each isoform of synemin within a multimolecular complex in cardiomyocytes.

Specifically, we show the novel localization of α-synemin to the M-band of the sarcomere through protein-protein interaction with the giant protein titin. The role of α-synemin bound to titin may support a signal transduction pathway specific for excitation/contraction at this location.

As for the role of β-synemin in cardiomyocytes, we have identified a substrate specific for β-synemin anchored PKA, VDAC. For the first time we are able to determine a specific purpose for β-synemin as an AKAP. Previously it was known that VDAC is phosphorylated by PKA, and that when VDAC becomes phosphorylated the on-rate of tubulin binding to the channel increases. This binding of tubulin changes the channels preference for cations over anions, which allows for the transport of Ca2+ across the OMM. This transport of Ca2+ across the OMM allows for the cell to maintain the Ca2+ concentration at a normal level during an EC event. Our findings have added to this previous work by identifying VDAC as a substrate of β-synemin anchored PKA. We have revealed that β-synemin is a dual AKAP that can bind PKA type I and PKA type II. We therefore propose a mechanism in which each PKA isoform participates in the phosphorylation state of VDAC.

116

The model we deduce here for the phosphorylation of VDAC by each isoform of

PKA anchored to β-synemin is as follows:

• VDAC regulates the Ca2+ homeostasis during the EC events

during contraction when Ca22+ levels are high

o β-synemin anchored PKA type II phosphorylates VDAC

o VDAC is able to take up the subsidiary Ca2+ into the

mitochondria, allowing for the maintained elevated levels of

Ca2+ in the cytosol needed for contraction.

• During relaxation when Ca2+ levels are dissipated

o β-synemin anchored PKA type I aids in the

dephosphorylation of VDAC

o VDAC is unable to take up Ca2+ into the mitochondria,

allowing for the maintained levels of Ca2+ at rest.

Our studies here, in conjunction with all the previous studies of synemin in

the heart, truly points to the idea that it is the heartiest spice.

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