MCUB REGULATES THE MOLECULAR COMPOSITION OF THE MITOCHONDRIAL CALCIUM UNIPORTER CHANNEL TO LIMIT MITOCHONDRIAL CALCIUM OVERLOAD DURING STRESS
A Dissertation Submitted to The Temple University Graduate Board
In Partial Fulfillment of the Requirements for the Degree DOCTOR OF PHILOSOPHY
by Jonathan P. Lambert August 2019
Examining Committee Members:
Dr. John W. Elrod, Ph.D., Advisory Chair, Center for Translational Medicine Dr. Douglas Tilley, Ph.D., Center for Translational Medicine Dr. Servio Ramirez, Ph.D., Pathology and Laboratory Medicine Dr. Raj Kishore, Ph.D., Center for Translational Medicine Dr. Toren Finkel, M.D., Ph.D., External Member, Aging Institute of the University Pittsburgh Medical Center, University of Pittsburgh
ABSTRACT
2+ Mitochondrial Calcium (mCa ) overload is a central event in myocardial-ischemia reperfusion (IR) injury that leads to metabolic derangement as well as activation of the mitochondrial permeability transition pore (mPTP). mPTP activation results in necrosis and loss of cardiomyocytes which results in acute death in some individuals while survivors are prone to developing heart failure and are
2+ predisposed to recurrent infarction events. mCa has also long been known to
2+ activate cellular bioenergetics implicating mCa in the highly metabolically demanding state of cardiac contractility. The mitochondrial calcium uniporter channel (mtCU) is a multi-subunit complex that resides in the inner mitochondrial
2+ 2+ membrane and is required for mitochondrial Ca (mCa ) uptake. Mitochondrial
Calcium Uniporter B (MCUB, CCDC109B gene), a recently identified paralog of
2+ MCU, is reported to negatively regulate mCa uptake; however, its precise regulation of uniporter function and contribution to cardiac physiology remain unresolved. Size exclusion chromatography of mitochondria isolated from ventricular tissue revealed MCUB was undetectable in the high-molecular weight
(MW) fraction of sham animals (~700kD, size of functional mtCU), but 24 hours following myocardial ischemia-reperfusion injury (IR) MCUB was clearly observed in the high-MW fraction. To investigate how MCUB contributes to mtCU regulation we created a stable MCUB-/- HeLa cell line using CRISPR-Cas9n.
2+ MCUB deletion increased histamine-mediated mCa transient amplitude by
~50% versus Wild-Type (WT) controls (mito-R-GECO1). Further, MCUB deletion
2+ increased mtCU capacitance (patch-clamp) and rate of [mCa ] uptake. Fast
ii protein liquid chromatography (FPLC) fractionation of the mtCU revealed that loss of MCUB increased MCU incorporation into the high-MW complex suggesting stoichiometric replacement and overall increase in functional mtCU complexes. Next, we generated a cardiac-specific, tamoxifen-inducible MCUB mouse model (CAG-CAT-MCUB x MCM; MCUB-Tg) to examine how the
MCUB/MCU ratio regulates mtCU function and contributes to cardiac physiology.
2+ MCUB-Tg mice were infected with AAV9-mitycam (genetic mCa reporter) and
2+ adult cardiomyocytes were isolated to record [mCa ] transients during pacing
2+ using live cell imaging. Increasing the MCUB/MCU ratio decreased [mCa ] peak
2+ amplitude by ~30% and significantly reduced the [mCa ] uptake rate. FPLC assessment revealed MCUB was undetectable in the high-MW fraction of
MerCreMer controls, but enriched in MCUB-Tg hearts. MCUB incorporation into the mtCU decreased the overall size of the uniporter and reduced the presence of channel gatekeepers, MICU1/2. Immunoprecipitations suggest that MCUB directly interacts with MCU but does not bind MICU1/2. These results suggest that MCUB replaces MCU in the mtCU and thereby modulates the association of
MICU1/2 to regulate channel gating. Cardiomyocytes isolated from MCUB-Tg hearts displayed decreased maximal respiration and reserve capacity, which correlated with a severe impairment in isoproterenol-induced contractile reserve
(LV invasive hemodynamics). MCUB-Tg cardiac mitochondria were resistant to
Ca2+-induced mitochondrial swelling suggesting MCUB limits mitochondrial permeability transition. Further, MCUB-Tg mice subjected to in vivo myocardial
IR revealed a ~50% decrease in infarct size per area-at-risk suggesting
iii 2+ increased MCUB expression prevents mCa overload and limits cell death.
These data suggest that MCUB regulation of the mtCU is an endogenous
2+ compensatory mechanism to decrease mCa overload during ischemic injury, but this expression is maladaptive to cardiac energetic responsiveness and contractility.
iv
This dissertation is dedicated to all human beings.
v ACKNOWLEDGMENTS
First off, I have to acknowledge Temple University and in particular the Deans of the Biomedical Sciences program, Dr. Dianne Soprano and Dr. Scott Shore, for not only accepting me into the program but also for their unwavering support in my endeavor to attain a Ph.D. in Biomedical Science. Temple has been an outstanding training ground for me to become an actual scientist and I am extremely grateful to the university.
Next, I would like to thank my fellow classmates in particular the group of
Ph.D. students that began classes in the fall of 2014. This group of individuals from day one considered everyone as an equal and a friend and drove, challenged, and inspired each other to become the best scientist they could be.
To my brother, mother, and father I am eternally grateful for your love and support. Without you I would never be here today. Mom and Dad, you have absolutely supported every single aspect of my education. Thank you for investing the time, resources, and love into me so I could be the first Ph.D. of the family.
The Center for Translational Medicine has become my second family in life. I am extremely lucky to have been a member of a collective of individuals whom selflessly support each other in all of life’s challenges. When I move on after my Ph.D. I will proudly say I am a proud graduate of CTM first and foremost.
Everyone who worked on the 9th floor of MERB over the last 5 years impacted the work of this thesis and it is a product of its environment.
vi I have to particularly thank Dr. Walter Koch and Dr. Steven Houser for how their extensive leadership in the American Heart Association laid the groundwork for a young trainee like me to be guided and mentored to be a basic cardiovascular biologist. The AHA community has been instrumental in developing my career and I am fortunate to be considered a member of this community.
I could never really express my full gratitude to the members of my thesis research committee for taking an active interest in my work and for sacrificing their time to challenge me to become a best scientist I can possibly be. Dr.
Douglas Tilley has known me even longer than Dr. Elrod and has been one of the most thoughtful and positively inquisitive committee members you could ask for. Dr. Servio Ramirez always reminded me to think outside of the level of the cardiomyocyte and his open-door policy and honest and direct career advice has been very helpful to navigate the next stage of my career. Dr. Raj Kishore and I started at Temple on pretty much the same day and over the last 5 years his mentorship in the classroom in particular has allowed me to understand the fundamentals of stem cell therapy in heart failure and I can now understand and appreciate important developments in the field. Last, I would like to thank Dr.
Toren Finkel for taking the time to serve as my external thesis committee member.
Without Dr. John Elrod none of this work would have been possible. Dr.
Elrod has committed countless hours, almost limitless resources, and his expert knowledge to mentor my development into a young basic cardiovascular
vii biologist. Dr. Elrod’s absolute commitment to pure science and respect for scientific rigor and reproducibility are some of his qualities I admire most that he has in turn instilled in me. Dr. Elrod has challenged me to acquire advanced technical expertise, grant writing experience, scientific networking skills, and an inquisitive mind to investigate the unknown. I am extremely fortunate and grateful to have had such a terrific mentor whom has prepared me to embark on a successful scientific career.
I also have to acknowledge all the members of the Elrod lab for their assistance, expertise, and friendship. When I first started the lab consisted of just
Tim, Mary, and I and it has been exciting to see the lab grow and develop into what it is today. Tim, Dhanendra, Pooja, Alyssa, Andrew, Emma, Trevor, Joanne,
Alycia, Devin, Neil, and Nadina I wish you nothing but the best and appreciate your support.
Last but not least I have to thank the city of Philadelphia which has embraced me and turned me into one of its brothers. I cannot possibly thank all the people in this city whom have supported and befriended enough as I went down this difficult endeavor but all of you have made this an enjoyable journey. In particular I must thank my dearest and most supportive friends Jessica, Nora,
Tony, Betta, Sam, Jess, Luke, Noel, Andy, Zara, Tongan, Rabbo, Alex, Mike
Nash, Michael, Kim, Tim, CJ, Brian, Geoff, Jeff, Trevor, Khari, Yamil, Dom,
Maria, Berto, Andrew, Efi, Alana, Brett, Matt, Graham, Claudio, Ben, and Aaron.
viii TABLE OF CONTENTS
Page
ABSTRACT ...... ii
DEDICATION...... v
ACKNOWLEDGMENTS ...... vi
LIST OF TABLES ...... xiii
LIST OF FIGURES ...... xiv
LIST OF ABBREVIATIONS ...... xvi
CHAPTER
1. CALCIUM SIGNALING ...... 1
Overview of Calcium Signaling ...... 1
Calcium: The Ion and Electrochemical Gradient ...... 3
Intracellular Ca2+ Storage: The Sarcoplasmic Reticulum ...... 4
Protein Recognition of Ca2+ ...... 5
Calcium as a Signaling Molecule ...... 7
Histamine Induced Intracellular Ca2+ Signaling ...... 7
Calcium Signaling During the Cardiac Cycle...... 8
The Mitochondrial Calcium Uniporter Complex ...... 11
Evidence of Mitochondrial Ca2+ Uptake ...... 11
Genetic Identification of mtCU Members ...... 12
MICU1 ...... 13
MCU ...... 18
ix EMRE ...... 21
MICU2 and MICU3 ...... 23
MCUR1 ...... 25
MCUB ...... 26
2. MITOCHONDRIAL CALCIUM IN CARDIAC PATHOPHYSIOLOGY ...... 29
Overview of Cardiac Metabolism ...... 29
Fatty Acid Metabolism ...... 30
Glycolysis...... 31
Pyruvate Dehydrogenase Complex ...... 33
Tricarboxylic Acid Cycle ...... 34
Oxidative Phosphorylation ...... 36
Mitochondrial Calcium’s Influence of Cellular Bioenergetics ...... 38
Overview of Myocardial Ischemia Reperfusion Injury ...... 40
Prevalence of Ischemic Heart Failure ...... 40
Pathophysiology of Myocardial Ischemia Reperfusion Injury ...... 40
mPTP Driven Necrosis ...... 42
Molecular Components of mPTP ...... 43
Rationale and Objective of Current Study ...... 45
2+ Preventing mCa Induced mPTP as a Therapeutic for MI ...... 45
2+ Modulation of mCa exchange during Ischemic Heart Disease ...... 46
NCLX’s Role in Ischemic Cardiac Stress ...... 46
MCU During Ischemic Cardiac Stress ...... 48
MCUB’s Role in Ischemic Cardiac Stress ...... 48
x 3. MCUB REGULATES THE MOLECULAR COMPOSITION OF THE MITOCHONDRIAL CALCIUM UNIPORTER CHANNEL TO LIMIT MITOCHONDRIAL CALCIUM OVERLOAD DURING STRESS ...... 51
Abstract ...... 51
Introduction ...... 52
Experimental Procedures ...... 55
Generation of a stable MCUB-/- HeLa cell line ...... 55
Generation of a cardiac-specific MCUB gain-of-function transgenic mouse model ...... 56
Size exclusion chromatographic analysis of the high molecular weight mtCU complex ...... 57
qPCR mRNA Analysis ...... 58
Western Blot Analysis ...... 58
Echocardiography ...... 59
Isolation of ACMs ...... 60
2+ 2+ Evaluation of mCa Uptake, Content, and Ca retention capacity experiments ...... 60
Mitochondria Isolation and Swelling Assay ...... 61
2+ 2+ Adult mouse cardiomyocyte iCa and mCa transient recordings ...... 62
Mitoplast Patch-Clamp Analysis of MCU Current ...... 62
Oxygen Consumption Assays ...... 63
Invasive hemodynamic measurements ...... 63
Permanent myocardial infarction and myocardial IR Injury ...... 63
2+ 2+ cCa and mCa flux in HeLa cells ...... 64
Immunoprecipitations ...... 65
xi Statistics ...... 66
Results ...... 66
MCUB deletion alters mtCU formation, stoichiometry, and 2+ mCa uptake ...... 66
MCUB incorporates into the mtCU complex after injury ...... 69
Generation of a cardiac-restricted MCUB overexpression mouse model ...... 70
2+ Overexpression of MCUB reduces mCa -overload and cardiac IR Injury ...... 72
MCUB expression impairs mitochondrial energetics and contractility during stress...... 73
MCUB expression displaces MCU-subunits in the mtCU (high-MW complex) to modulate MICU1/2 gating ...... 75
Discussion ...... 77
4. CONCLUSIONS AND SIGNIFICANCE ...... 94
Extended Discussion and Future Directions ...... 94
Paralog Interference ...... 94
Ischemic cardiac injury: Hif1-alpha ...... 100
Ischemia reperfusion injury: AP-1 ...... 102
Translational Significance ...... 103
Conclusion ...... 104
BIBLIOGRAPHY ...... 109
xii LIST OF TABLES
Table Page
3.1 Antibody Information…………………………………………….……………...59
xiii LIST OF FIGURES
Figure Page
1-1. Molecular Makeup of the Mitochondrial Calcium Uniporter Complex (mtCU)……………………………………….……………………………………. 28
2-1. Mitochondrial Calcium Control of Cellular Bioenergetics ……………… 50
3-1. MCUB deletion alters mtCU formation, stoichiometry, and 2+ mCa uptake……………….……………………………………………………... 82
3-2. mRNA expression in HeLa cells, Fura-2 calibration, size exclusion chromatography of known protein standards for generation of a standard 2+ curve of molecular weights, and mCa ………………………………….……...84
3-3. MCUB incorporates into the mtCU complex after injury warranting the generation of a MCUB conditional mutant mouse model………………...85
3-4. Assessing leakiness of the MCUB transgenic mouse, exponential 2+ 2+ decay rates for mCa transients and cCa transients……………………….. 86
2+ 3-5. Overexpression of MCUB reduces mCa -overload and cardiac IR-injury ……………………………………….………………………….87
3-6. Echocardiographic assessment of parameters of cardiac function, Kaplan Meier survival curve of I/R injury 1 month post tamoxifen, and representative images of EBD/TTC staining in mouse models subjected to I/R injury 1 month post tamoxifen.……….………………………………….. 88
3-7. MCUB expression impairs mitochondrial energetics and contractility during stress.………………………………….………………………………….. 89
3-8. Expression levels of total pyruvate dehydrogenase E1α.……………… 90
3-9. MCUB expression displaces MCU-subunits in the mtCU (high-MW complex) to modulate MICU1/2 gating…………………………….. 91
3-10. Endogenous immunoprecipitations confirm MCUB interacts with MCU and not MICU1……………………………….……………………………. 92
3-11. Compensatory mechanisms take place 1 month following tamoxifen administration restoring cellular bioenergetics and reversing the cardiac phenotype.……………………………………….………………………………...93
xiv 4-1. Graphical depiction of how the increase in MCUB affects cellular processes 2+ dependent on mCa signaling in the context of cardiac (patho)physiology..105
4-2. Graphical depiction of how the increase in MCUB seen during IR injury 2+ alters the composition of mtCU to limit mCa overload..………………...……106
4-3. Computer models of MCU and MCUB in monomeric and tetrameric form.……………………………………….……………………………………….107
4-4. Promoter regions of MCUB possibly targeted by transcription factors associated with IR injury.……………………………………….………………..108
xv LIST OF ABBREVIATIONS
AAR area-at-risk
Acetyl-CoA acetyl coenzyme A
Acyl-CoA acyl coenzyme A
ACMs adult cardiomyocytes
α-KDH α-ketoglutarate dehydrogenase
ATP adenosine triphosphate
BH4 tetrahydrobiopterin
CI Complex I; NADH:ubiquinone reductase
CII Complex II; succinate:quinone oxidoreductase
CIII Complex III; Coenzyme Q:cytochrome c reductase
CIV Complex IV; cytochrome c oxidase
CV Complex V; F1FO ATP synthase
Ca2+ calcium
Ca45 calcium-45
CAT carnitine:acylcarnitine translocase
CD36 fatty acid translocase
CICR calcium induced calcium release
CO2 carbon dioxide
CPTI carnitine palmitoyltransferase I
CPT2 carnitine palmitoyltransferase 2
CVD cardiovascular diseases
CsA cyclosporine A
xvi Cyp D cyclophilin D
DAG diacylglycerol
E1 thiamin diphosphate dependent enzyme
E2 dihydrolipoamide acetyltransferase enzyme
E3 dihydrolipoamide dehydrogenase eNOS endothelial nitric oxide synthase
ER endoplasmic reticulum
ETC electron transport chain
FA fatty acid
FABPpm plasma membrane fatty acid binding protein
FADH2 flavin adenine dinucleotide
FATB fatty acid transport protein
FABP3 heart specific fatty acid binding protein fatty acyl-CoA fatty acyl coenzyme A
FMN flavin mononucleotide
H+ hydrogen
HCN hyperpolarization-activated, cyclic nucleotide gated channels
HIF-1α hypoxia inducible factor-1α
HR1-4 histamine receptors 1-4
2+ iCa intracellular calcium
IDH isocitrate dehydrogenase
IMS intermembrane space
IMM inner mitochondrial membrane
xvii
IP3 inositol 1,4,5-triphosphate
IP3R inositol 1,4,5-triphosphate receptors
IR ischemia reperfusion injury
L-type long-lasting voltage dependent Ca2+ channels
2+ mCa mitochondrial calcium
MCU mitochondrial calcium uniporter
MCU cKO MCU conditional knockout mouse model
MI myocardial Infarction
MICU1 mitochondrial calcium uptake 1
MICU2 mitochondrial calcium uptake 2
MICU3 mitochondrial calcium uptake 3
Mg2+ magnesium mtCU mitochondrial calcium uniporter complex mPTP mitochondrial permeability transition pore
MW molecular weight
NADH nicotinamide adenine dinucleotide
NCLX mitochondrial Na+/Ca2+ exchanger
NCLX cKO NCLX conditional knockout mouse model
NCX Na+/Ca2+ exchanger
NFAT nuclear factor of activated t cells
O2 molecular oxygen
OMM outer mitochondrial membrane
OxPhos oxidative phosphorylation
xviii
PDH pyruvate dehydrogenase
PDK pyruvate dehydrogenase kinase
PDP1 pyruvate dehydrogenase phosphatase subunit 1
Pi inorganic phosphate
PLC phospholipase C
PKC protein kinase C
PMCA plasma Ca2+ ATPase
P/O ratio phosphate/oxygen ratio
PP2C pyruvate phosphatase 2C
Ppif-/- Ppif knockout mouse
ROS reactive oxygen species
SA sino-atrial node
SEC size exclusion chromatography
SERCA sarcoendoplasmic reticulum Ca2+-ATPase
SR sarcoplasmic reticulum
RyR ryanodine receptors
TALEN transcription activator-like effector nuclease technology
TCA cycle tricarboxylic acid cycle
TRE-NCLX NCLX gain-of-function mouse model
T-type transient voltage dependent Ca2+ channels
T-tubules transverse tubules
WT wild-type
xix CHAPTER 1
CALCIUM SIGNALING
Overview of Calcium Signaling
Calcium’s (Ca2+) significance to human physiology is evident in the fact that it is the 5th most abundant element in the human body. When life first evolved in the
Precambrian period over 3.5 billion years ago the early oceans were likely alkaline containing very low concentrations of Ca2+, reflected by the fact that intracellular free Ca2+ levels of all cellular organisms are also kept quite low in the
100 nM range suggesting early life formed in a low calcium environment(Plattner
& Verkhratsky, 2015). Ocean acidification, water runoff from land, geothermal activity, and Ca2+ release from basalts over time slowly increased the Ca2+ concentration in the oceans to levels seen today of roughly 10 mM(Jozef,
Stephan, & Barbara, 2013). This means cellular evolution was not only concurrent to increases in extracellular Ca2+ but influenced by it as well. The evolutionarily setup of this large electrochemical gradient across the plasma membrane as well as the calcium ions intrinsic chemical properties capable of regulating protein function has allowed Ca2+ to become a ubiquitous second messenger signaling molecule utilized by a plethora of cellular processes that span the entire lifetime of organisms.
Ca2+ signaling is triggered at the moment of conception as sperm fertilization introduces a sperm specific phospholipase C-ζ into the oocyte causing waves of Inositol 1,4,5-triphosphate (IP3) induced intracellular Ca2+
2+ (iCa ) transients that last for hours resulting in egg activation, completion of
1 meiosis, and the initiation of the cell cycle allowing for embryonic development
(Berridge, Lipp, & Bootman, 2000; Whitaker, 2006). In the process of embryonic development cellular differentiation leads to the development of specific cell and tissue types that utilize the same core set Ca2+ signaling machinery in a variety of different cellular functions. Ca2+ is heavily integrated in the process of synaptic transmission in the brain. In neurons Ca2+ influx triggers the depolarization and propagation of an action potential which upon reaching the synapse activates voltage dependent Ca2+ channels causing them to open allowing further Ca2+ influx triggering vesicular fusion and neurotransmitter release(Brini, Cali, Ottolini,
& Carafoli, 2014). In the heart Ca2+ not only triggers the depolarization of cardiomyocytes but also directly activates myofilaments implicating Ca2+ in cardiac contraction and relaxation(Eisner, Caldwell, Kistamás, & Trafford, 2017).
Many other cellular processes are influenced by Ca2+ including gene transcription, cell motility, and bioenergetics(Rizzuto & Pozzan, 2006). Ca2+’s influence even extends to cellular demise as it is a key activator of apoptosis and necrosis implicating Ca2+’s role in pathophysiology(Kruman & Mattson, 1999).
Ca2+’s ubiquitous influence in cellular biology and essential role from the conception of life to initiation of cellular death mechanisms necessitates the understanding of this powerful force in biology. As Otto Loewi, who received the
Nobel Prize in Physiology or Medicine in 1936, exclaimed “Ja, Kalzium ist nicht alles!” “Yes, Calcium is everything!”(Bush, 2010)
2 Calcium: The Ion and Electrochemical Gradient
Ca2+’s intrinsic chemical properties make this divalent cation an ideal second messenger. The primary coordination sphere of the Ca2+ ion is capable of accommodating four to twelve oxygen atoms while biologically proteins typically use the carboxylate and carbonyl oxygen atoms of amino acids to coordinate
Ca2+ within six to eight oxygen atoms(Williams, 2006). Ca2+ exchanges water molecules very rapidly, in comparison 103 times faster than Magnesium (Mg2+), enabling it to diffuse and thus signal in a rapid manner(Williams, 2006). Ca2+’s chemistry alone is not what makes this ion a powerful signaler, the actual environment of the cell also favors rapid transmission of signaling.
The intracellular concentration of free Ca2+ in the cytosol is kept quite low in the ~100 nanomolar range while the extracellular Ca2+ concentration is ~2 millimolar generating a 20,000 fold difference in [Ca2+] along the plasma membrane establishing a powerful electrochemical gradient strongly favoring cellular Ca2+ influx(Clapham, 2007). Maintenance of the electrochemical gradient is integral to allowing Ca2+ to act as a second messenger and this is achieved by transporting Ca2+ out of the cell, compartmentalizing Ca2+ in intracellular storage sites, and buffering Ca2+ within the cell.
Along the plasma membrane two main modes of Ca2+ export serve to keep Ca2+ out of the cell; the Ca2+ pump and the Ca2+ exchanger. The Ca2+ pumps are a family of high affinity, low capacity plasma Ca2+ ATPase (PMCA) that transport one Ca2+ ion against its electrochemical gradient to extracellular space for every adenosine triphosphate (ATP) consumed thus this process is
3 energy intensive(Bruce, 2018). The second mode of Ca2+ export is via the low affinity, high capacity Na+/Ca2+ exchanger (NCX) which uses the energy efficient
Na+ gradient down the plasma membrane to transport Ca2+ up its electrochemical gradient from the cytosol to the extracellular environment(Philipson et al., 2002).
One Ca2+ ion is exchanged for three Na+ ions(Jeffs, Meloni, Bakker, & Knuckey,
2007). Both systems operate together but due to NCX’s high capacitance it
2+ largely operates when iCa levels are in the μM range while PMCA’s generally operate in the nM range(Guerini, Coletto, & Carafoli, 2005).
Intracellular Ca2+ Storage: The Sarcoplasmic Reticulum
2+ 2+ Ca is also compartmentalized within the cell itself not only to keep iCa levels
2+ low but also to give cells the flexibility to stimulate release of iCa stores themselves for further signaling. The endoplasmic reticulum (ER, also known as the sarcoplasmic reticulum (SR) in skeletal and cardiac cells), is the organelle that serves as the main Ca2+ storage site within the cell. The SR’s total Ca2+ levels are dynamic, but generally range between 100 to 800 μM, thus to uptake
Ca2+ from the cytosol the SR must transport Ca2+ against an electrochemical gradient(Samtleben et al., 2013). SR Ca2+ uptake is mediated via a family of sarcoendoplasmic reticulum Ca2+-ATPases (SERCAs) in which 2 Ca2+ ions are transported into the SR for every ATP hydrolyzed making this an energy intensive process(Stammers et al., 2015). High capacity, low affinity Ca2+ binding proteins within the SR, such as calsequestrin and calreticulun, serve to keep free
SR Ca2+ levels low by buffering Ca2+ thus reducing Ca2+’s electrochemical
4 gradient and promoting SR Ca2+ uptake(Song et al., 2007). This buffering
2+ capacity also makes the SR an iCa storage site which is targeted by signal
2+ transduction in order to mobilize iCa stores for its use as a second messenger.
SR Ca2+ extrusion is largely mediated via two families of proteins, the Ryanodine
Receptors (RyR) and the Inositol 1,4,5-triphosphate receptors (IP3Rs)(Lanner,
Georgiou, Joshi, & Hamilton, 2010).
Protein Recognition of Ca2+
2+ Protein recognition of alterations in iCa levels are an important step in transmitting the Ca2+ signal to cellular processes. Ca2+ binding proteins consist of large families of proteins that affect a plethora of cellular processes though modulating many different signaling pathways. There are two major Ca2+ binding motifs that exist which bind Ca2+ in a selective, reversible manner with fast kinetics allowing for rapid signal transduction(Drake, Zimmer, Kundrot, & Falke,
1997).
The most common Ca2+ binding motif is the EF-hand motif which are normally found adjacent to each other within a protein in at least pairs(Nakayama
& Kretsinger, 1994). EF-hands are helix-loop-helix motifs that consists of two α- helices separated by a short loop of ~12 amino acids(Yáñez, Gil-Longo, &
Campos-Toimil, 2012). In the canonical EF-hand oxygen atoms of carboxylates or carbonyls at amino acid position’s 1, 3, 5, and 12 within the loop region are usually responsible for coordinating Ca2+(Grabarek, 2006). Hundreds of proteins contain EF-hands and over 66 subfamilies have been identified(Day, Reddy,
5 Shad Ali, & Reddy, 2002). Some of the most well-known EF-hand containing
2+ 2+ proteins are; Parvalbumin which buffers free cytosolic Ca (cCa ) after muscle contraction playing a role in relaxation(Asp, Sjaastad, Siddiqui, Davis, & Metzger,
2016); Calmodulin in which Ca2+ binding causes a conformational change allowing it to bind and regulate over 300 different proteins impacting diverse cellular processes such as cardiac contraction by activating myosin light chain kinase(Dewenter, von der Lieth, Katus Hugo, & Backs, 2017); the S100 proteins which are the largest EF-hand protein family and act as multifactorial, diverse signal transducers regulating differentiation, proliferation, apoptosis, metabolism, among other cellular processes(Donato et al., 2013; Sedaghat & Notopoulos,
2008); and Calcineurin, a serine/threonine phosphatase, which is activated by
Ca2+ allowing dephosphorylation of its target nuclear factor of activated t cells
(NFAT) causing translocation of this transcription factor to the nucleus which then alters gene expression(Hogan, Chen, Nardone, & Rao, 2003).
The second most common Ca2+ binding domain is the C2 domain which was originally identified as the domain in Protein Kinase C (PKC) that was responsible for Ca2+ dependent cellular membrane trafficking(Shin, Han, Wang,
& Sudhof, 2005). Further research found similar amino acid sequences present in a diverse set of proteins involved in cellular trafficking, membrane fusion, or signal transduction. The C2 domain is made up of roughly 130 amino acids that form a pair of 4 stranded β-sheets(Corbalan-Garcia & Gómez-Fernández, 2014).
They can bind 2 or 3 Ca2+ ions and variations in amino acid sequences of C2 domains allow for different sensitivity and specificity to the Ca2+ signal(Nalefski &
6 Falke, 1996). Protein families that contain C2 domains include; PKC in which
Ca2+ binding causes translocation to the cellular membrane(Nishizuka, 1988);
Phospholipase C (PLC) which when activated cleaves Phosphatidylinositol 4,5- bisphosphate (PIP2) into diacylglycerol (DAG) and IP3 resulting in activation of
2+ 2+ iCa stores(Kadamur & Ross, 2013); and synaptotagmins which upon Ca binding dimerize or bind vesicular proteins leading to vesicular transport and release(T. Wang et al., 1999).
Calcium as a Signaling Molecule
The eukaryotic cell through extrusion and compartmentalization has thus kept its
2+ iCa concentrations low, but has also established large electrochemical gradients along the plasma membrane and SR to allow external stimuli to utilize
Ca2+ as a powerful secondary messenger. Receptors are now able to transmit
2+ 2+ internal signals through Ca influx or by the stimulation of the release of iCa stores. This coupled with the ability of Ca2+ to diffuse throughout the cell and proteins abilities to coordinate and bind Ca2+ to affect their function gives Ca2+ its second messenger capabilities. This signal can be Ca2+ itself or some other type
2+ of cellular signaling that elicits an iCa response.
Histamine Induced Intracellular Ca2+ Signaling
The agonist histamine binds to histamine receptors (HR1-4) which are G protein coupled receptors. Stimulated H1 receptors linked to Gq subunits activate PLC which cleaves the phospholipid PIP2 into the second messengers DAG
7 (Sedaghat & Notopoulos) and IP3(Tabarean, 2016). IP3 is soluble and diffuses to its receptor the IP3R, located on the ER, and upon binding Ca2+ is released from this intracellular storage site into the cytosol(Fukui et al., 2017). HRs are expressed in many non-excitable cell types used for loss- and gain-of-function studies such as HeLa cells. Cell biologists can stain cells with permeable Ca2+ fluorescent dyes or express genetically encoded Ca2+ reporters and use
2+ fluorescent microscopy to investigate iCa dynamics upon agonist stimulation in these non-excitable cell types.
Calcium Signaling During the Cardiac Cycle
The cardiac cycle of the mammalian heart is the process in which the four chamber heart contracts and relaxes to pump blood systemically throughout the body and Ca2+ signaling is heavily involved in the entire process(Fukuta & Little,
2008). The action potential of the cardiac cycle begins when pacemaker cells within the sino-atrial (SA) node located in the apex of the right atrium hyperpolarize to -60 millivolts causing induction of a funny current where hyperpolarization-activated, cyclic nucleotide gated (HCN) channels activated by this hyperpolarization slowly conduct Na+ into the cell beginning cell depolarization(Altomare et al., 2003). During depolarization as the membrane potential reaches -50 millivolts transient voltage dependent Ca2+ channels (T- type) open allowing Ca2+ to travel down its electrochemical gradient further depolarizing the cell(Perez-Reyes, 2003). Once the membrane potential reaches roughly -40 millivolts long-lasting voltage dependent Ca2+ channels (L-type) open
8 depolarizing the cell to the point where the threshold for an action potential is reached(Bodi, Mikala, Koch, Akhter, & Schwartz, 2005). The action potential travels throughout the atria at 0.5 meters/second (m/s) by cell-to-cell conduction causing coordinated atrial contraction and pumping of blood into ventricles(Valderrábano, 2007). At the atrioventricular node the action potential slows to 0.05 m/s to allow for proper atrial depolarization and ventricular filling of blood(Pinnell, Turner, & Howell, 2007). The action potential then reaches the base of the ventricle and travels down the Bundle of His in the interventricular septum through a system of Purkinje fibers that conduct the action potential throughout the ventricle for uniform ventricular contraction ejecting blood to the pulmonary system (right ventricle) and throughout the body via the aorta (left ventricle)(Temple, Inada, Dobrzynski, & Boyett, 2013).
At the level of the cardiomyocyte the action potential sweeps into deep invaginations of the sarcolemma called transverse tubules (t-tubules) which can be 200 to 300 nms wide(T. Hong & Shaw, 2016). Depolarization from -90 mVs to
20 mVs opens L-type Voltage gated Ca2+ channels (Cav1.2) causing Ca2+ influx into a specialized area called the dyadic cleft where t-tubules are juxtaposed in very close proximity (15 nms) to the SR(Eisner et al., 2017). In this microdomain when L-type Ca2+ channels open it causes Ca2+ levels to rise from ~100 nM to greater than 10 μM(Hake & Lines, 2008). This induces a process known as calcium induced calcium release (CICR) in which RyRs become activated by this local Ca2+ signal and further release SR-Ca2+ stores into the cytosol(Perez-
2+ 2+ Reyes & Bers, 1999). The Ca ions now diffuse globally and raise the total cCa
9 levels to ~1 μM and Ca2+ now interacts with contractile machinery of the cardiomyocyte(Marks, 2003).
Cardiomyocytes contain contractile units called sarcomeres. Sarcomeres are composed of myofilaments where myosin forms the thick filament and actin forms the thin filament that associates with tropomyosin and the regulatory troponin complex which is a hetero-trimer made up of Troponin-C, Troponin-I, and Troponin-T(Fearnley, Roderick, & Bootman). Ca2+ binds to the EF-hands of
Troponin-C which causes a conformational change in the inhibitory subunit,
Troponin-I, allowing tropomyosin to slide and reveal myosin binding sites on actin(Parmacek & Solaro, 2004). Myosin, with an ADP and inorganic phosphate
(Pi) bound to it, binds to actin and generates the power stroke by pulling actin and in the process ADP and phosphate are released from myosin. ATP binds myosin causing the cross bridge to detach allowing myosin to return to its recovery position and ATP then disassociates into ADP and Pi(Tang et al., 2017).
If Ca2+ remains bound to troponin-C the cycle will repeat. Thus to cause cardiomyocyte relaxation Ca2+ must now be removed from the cytosol which is mediated by SR-mediated Ca2+ uptake through SERCA or removal from the cell via NCX.
Ca2+’s importance to cardiac contraction is not yet fully explained and to date new discoveries into how Ca2+ is further involved in the highly coordinated system of the cardiac cycle are being made. Ca2+ signaling to the mitochondria and how this is implicated in cardiac (patho)physiology will be the focus of this work.
10 The Mitochondrial Calcium Uniporter Complex
Evidence of Mitochondrial Ca2+ Uptake
In 1961 researchers exposed purified rat kidney mitochondria to radiolabeled calcium-45 (Ca45) and observed that mitochondria were able to uptake large quantities of Ca2+(Deluca & Engstrom, 1961). Subsequently, it was found that almost all vertebrate tissue types and some invertebrate animals exhibited
2+ 2+ 2+ mitochondrial Ca (mCa ) uptake(Bygrave & Ash, 1977). Initially, mCa uptake was just thought to be an active process until the discovery of the large electrochemical gradient of -180 mVs along the inner mitochondrial membrane
(IMM)(Mitchell, 1966). It was then realized Ca2+ transport would be very energetically favorable so there must be a channel or carrier responsible for Ca2+ transport(Rottenberg & Scarpa, 1974). Further, it was found that ruthenium red
2+ inhibited mCa uptake(Vasington, Gazzotti, Tiozzo, & Carafoli, 1972). It should be noted that ruthenium red is poorly cell permeable and non-selective, also inhibiting SR-Ca2+ uptake thus it is not suitable to be used as pharmaceutical
2+ agent. Though the identity of mCa uptake remained elusive researchers coined this mechanism the Mitochondrial Calcium Uniporter (MCU)(Gunter & Pfeiffer,
1990). Patch-clamping experiments of mitoplasts (mitochondria stripped of the outer mitochondrial membrane (OMM)) conclusively determined that MCU was a channel, not a carrier, that was highly selective for Ca2+ where conductance was only observed at largely negative membrane potentials of -160 millivolts(Y.
Kirichok, G. Krapivinsky, & D. E. Clapham, 2004). This means mitochondria must
11 be depolarized for channel conductance and that the driving force for Ca2+ entry is established by the electron transport chain.
Genetic Identification of mtCU members
2+ Evidence for mCa uptake goes back all the way to the 1960’s but the actual molecular identity of the uniporter remained elusive until the key observation that ruthenium red sensitive mitochondrial calcium uptake is evolutionarily conserved in eukaryotes and kinetoplastids but is lacking in S. cerevisiae(Baughman et al.,
2011; D. De Stefani, Raffaello, Teardo, Szabo, & Rizzuto, 2011; Perocchi et al.,
2010). Researchers took advantage of these insights into the evolutionary history
2+ of mCa uptake and employed comparative physiology to identify possible genetic components of mtCU. For instance, genes targeted to the mitochondria were evaluated to see which were present in eukaryotes and kinetoplastids but absent in S. cerevisiae as these were ideal candidates to be the channel.
Identified candidates were then targeted with genetic gain- and loss-of-function studies to evaluate if they contributed to the uniporter and to discover how they contributed to the function and regulation of the uniporter. Here each bona fide channel constituent’s discovery will be characterized and experimental evidence for how each member precisely contributes to the regulation or function of the uniporter along with each member’s physiological relevance will be detailed.
12 MICU1
MICU1 (mitochondrial calcium uptake 1, formerly known as CBARA1) was the first genetic component of the mitochondrial calcium uniporter complex (mtCU) identified through comparative physiology and evolutionary genomics(Perocchi et al., 2010). This gene was selected as a possible constituent because it was listed in the Mitocarta database (an inventory of mammalian genes likely targeted to the mitochondria) that had paralogs present in eukaryotes and kinetoplastids but absent in S. cerevisae. Following its identification numerous groups employed genetic gain- and loss-of-function studies to describe MICU1’s precise regulation
2+ on mCa uptake reaching divergent conclusions largely due to the different model systems utilized and variation amongst the experimental procedures employed.
In the original study identifying MICU1, HeLa cells expressing a mitochondrial targeted Ca2+ sensor, aequorin (mt-AEQ), were treated with MICU1 targeted shRNA to achieve knockdown and then stimulated with the agonist
2+ 2+ histamine to initiate an IP3 mediated iCa response to observe mCa uptake. In
2+ this initial study MICU1 knockdown ablated mCa uptake initially indicating
2+ MICU1 was required for mCa uptake(Perocchi et al., 2010). MICU1 was thus hypothesized to either be the pore-forming subunit of the channel or possibly be a channel gatekeeper. The subsequent identification of MCU as the actual pore- forming subunit of mtCU discounted the former hypothesis(D. De Stefani et al.,
2011). Additional studies provided evidence of MICU1’s true functional role as acting as a channel gatekeeper. Intact HeLa cells stained with the mitochondrial
13 sensitive dye Rhod-2 exhibited higher baseline fluorescence intensity with
2+ knockdown of MICU1 indicating increased basal mCa levels but upon histamine
2+ stimulation no alteration in the mCa uptake rate was observed(Mallilankaraman et al., 2012). Additionally, this study found that in a permeabilized cell system
2+ 2+ MICU1 knockdown cells exhibited increased mCa uptake at lower Ca concentrations than WT counterparts. This data provided the first evidence that
2+ 2+ 2+ indicates that MICU1 inhibits mCa uptake al low Ca levels by setting a Ca threshold for entry at ~2 μM(Mallilankaraman et al., 2012). This gatekeeping
2+ functionality is critical because it means mCa uptake is inhibited at physiological
2+ levels of iCa (~200 nM).
Another layer of complexity was then added to MICU1’s functional role when it was further confirmed that MICU1 not only set a Ca2+ threshold for entry but additionally imparted cooperativity. In permeabilized cells exposure to various
2+ 2+ doses of [Ca ] revealed that silencing MICU1 resulted in less efficient mCa uptake at increasing Ca2+ boluses (greater than 3 μM) indicating MICU1 not only
2+ sets a threshold for entry but also cooperatively enhancing mCa uptake rates at increasing Ca2+ concentrations(Csordas et al., 2013). It is noteworthy that this cooperativity was not observed by the Madesh group but was observed in the
Csordas study which was found to be due to the absence or presence of the divalent ion Mg2+ in the buffer. Mg2+ is another divalent ion that has been shown to allosterically activate uniporter activity(Kroner, 1986). Cooperative activation was only observed in experiments containing Mg2+ further suggesting a role for
Mg2+ in uniporter functionality. Overexpressing MICU1 in HeLa cells further
14 2+ increases mCa transients compared to WT upon exposure to the same strength of histamine stimulation further complementing the idea that increased MICU1 in the uniporter promotes cooperativity of channel activation(Patron et al., 2014;
Vecellio Reane et al., 2016).
After considering the differences in findings due to experimental inconsistencies and the results of each of the above studies it is the general
2+ 2+ consensus that MICU1 inhibits mCa uptake at low Ca concentrations (baseline
2+ conditions) and that upon stimulation of iCa stores MICU1 senses elevations in
2+ 2+ cCa levels to activate the uniporter and induce mCa uptake. A closer inspection of MICU1’s protein domains further resolves its functionality.
MICU1 encodes for a 54 kilodalton (kD) protein (~48 kD when the mitochondrial targeting sequence is removed) that was initially characterized as an IMM protein due to an incorrectly identified transmembrane domain, but most studies now agree that MICU1 is localized to the intermembrane space
(IMS)(Csordas et al., 2013; Patron et al., 2014; L. Wang et al., 2014). MICU1 contains two highly conserved canonical EF-hand motifs which coordinate Ca2+,
2+ thus it is hypothesized that these EF-hands may sense changes in iCa levels to
2+ activate mCa uptake. Their functional role was revealed in rescue experiments where either cDNA encoding WT MICU1 or MICU1 with point mutations in the
EF-hands (EF1:D231A,E242K; EF2:D421A,E432K) (rendering them unable to coordinate Ca2+) were expressed in MICU1 knockdown cells. Expression of WT
MICU1 completely restored the normal Ca2+ phenotype while expression of
MICU1 EF-hand mutants displayed a Ca2+ phenotype where a Ca2+ threshold for
15 2+ entry was observed but cooperativity of enhanced mCa uptake with increasing
Ca2+ concentrations was no longer present(Csordas et al., 2013). This indicates that the EF-hand motifs have a role in cooperativity but not in setting the initial
Ca2+ threshold for entry. MICU1 forms ~100 kD homodimers through disulfide bonds at a highly conserved cysteine residue (MICU1C465)(Patron et al., 2014).
Through this cysteine residue MICU1 also forms ~95 kD heterodimers with its gene paralog MICU2 and also forms heterodimers with MICU3(Payne, Hoff,
Roskowski, & Foskett, 2017). These interactions are thought to alter and fine tune the Ca2+ threshold for entry and cooperativity. Additionally, MICU1 contains a polybasic sequence (98-KKKKR-103) near its C-terminus that interacts with a polyaspartate tail of EMRE (100-EDDDDDD) to help anchor MICU1 within the mtCU(Tsai et al., 2016). Interestingly, MICU1 interacts with EMRE in the absence of MCU with the reversal also true that MICU1 interacts with MCU in the absence of EMRE(Tsai et al., 2016). This suggests MICU1 also interacts with
MCU directly and it was shown that (aa443-KQRLMRGL-451, now named the
DIME interacting domain) are critical for MICU1 binding to the DIME domain of
MCU through the formation of salt bridges as point mutations within this region
2+ reduces MICU1’s ability to interact with MCU and alters mCa uptake by lowering the threshold for entry similar to knockout of MICU1 alone(Paillard et al., 2018).
MICU1’s physiological role was revealed through evaluation of human disease and with the generation of MICU1 deficient knockout mice. Humans that display mutations associated with a loss-of-function of MICU1 develop skeletal muscle myopathies and brain disorders in an autosomal recessive manner in a
16 similar fashion to known mitochondrial disorders suggesting MICU1 functionality is crucial to human physiology(Logan et al., 2013). Genomic deletion of MICU1 in a mouse model showed normal embryogenesis but upon birth perinatal lethality where only one out of seven MICU1 knockout mice survived to 1 week of age.
Interestingly, MICU1 knockout mice that survived past 1 week displayed defects associated with skeletal muscle myopathy and neuropathies similar to what is seen in the human MICU1 loss-of-function phenotype(Liu et al., 2016). Further, physiologic functionality of MICU1 was revealed when MICU1’s mRNA and protein expression levels were noted to be tissue specific. For example, MICU1 at the protein level is more expressed in liver than the heart and most highly expressed in the skeletal muscle(Paillard et al., 2017). In the Paillard et al study it was then experimentally proven that tissues with high expression of MICU1 had a higher threshold for Ca2+ entry and increased cooperativity while tissues with less abundance of MICU1 had a lower threshold for Ca2+ entry and decreased cooperativity. This has led to the hypothesis that MICU1 expression allows different tissue and cell types to decode tissue specific Ca2+ signaling further indicating the actual molecular composition of the uniporter reflects physiologic function.
Overall MICU1’s functional role is to impart gatekeeping activity. This serves to prevent calcium overload at low Ca2+ concentrations but allows for
2+ 2+ mCa uptake upon stimulation of iCa stores implicating MICU1’s role in the
2+ 2+ mCa dependent regulation of cellular bioenergetics and mCa overload. In mice
17 and humans, loss of MICU1 function leads to a distinct clinical phenotype of skeletal muscle myopathy and neurological defects.
MCU
After identification of the regulator of the uniporter, MICU1, two groups independently discovered the next member of the elusive mitochondrial calcium uniporter, MCU(Baughman et al., 2011; D. De Stefani et al., 2011). Mootha’s group used integrative genomics to predict if any other genes were functionally
2+ related to MICU1 and required for mCa uptake. Using computational methods all mammalian genes were ranked based on their phylogenetic profile to MICU1, co-similarities in their RNA expression profile with MICU1 in 81 different mouse cell types or tissues, as well as a score for mitochondrial protein expression amongst 14 different mouse tissues. All three computational methods identified an unknown protein, CCDC109A, which is now called the mitochondrial calcium uniporter, MCU, because it was further revealed to be the bona fide pore-forming subunit of mtCU(Baughman et al., 2011). In this seminal study, MCU (a 40 kD protein that is ~34 kD when the mitochondrial localization is cleaved), was found to localize to the IMM, form oligomers, and interact with the regulator MICU1 to form a high molecular weight (high-MW) complex ranging from ~450 kD to 700 kD as identified by blue native page(Baughman et al., 2011). Silencing MCU
2+ expression in HeLa cells or in the liver ablated mCa uptake. This group also identified two transmembrane domains within MCU and computational biology suggested that the N and C termini of MCU were situated in the mitochondrial
18 matrix while a short loop within the transmembrane domains faced the IMS.
Within this short loop a highly evolutionarily conserved set of amino acids was identified and termed the DIME motif as mutations of these acidic amino acids
2+ (E257A, D261A, or E264A) was associated with a failure to rescue mCa uptake in MCU knockdown cells(Baughman et al., 2011).
Rizzuto’s group set out to find possible mtCU channel constituents by identifying genes within the inventory of the MitoCarta database expressed in most tissue types, that contained one or two transmembrane domains and had paralogs present in kinetoplastids but were absent in S. cerevisiae(D. De Stefani et al., 2011). Like Mootha’s group, they identified CCDC109A now named MCU.
2+ They found that silencing MCU led to decreased mCa uptake upon agonist stimulation in intact HeLa cells and additionally found that overexpressing MCU actually increased mitochondrial Ca2+ transients and made cells more sensitive to apoptotic stimuli resulting in increased cell death. They also showed that MCU
2+ imparted the long known biophysical properties of mCa uptake by expressing
MCU cDNA into a lipid bilayer and performing electrophysiology experiments confirming a channel with low affinity and high capacitance(D. De Stefani et al.,
2+ 2011). With the identification of the pore-forming subunit responsible for mCa uptake the race was on to establish the physiological function of this gene by establishing genetic loss-of-function transgenic mouse models but differences in approach have led to differences in the physiological role of this gene.
Pan et al developed the first MCU-/- mouse using a gene trap strategy by integrating a retrovirus based gene trapping vector into intron 1 of MCU(Pan et
19 al., 2013). The Pan et al MCU KO mouse was not found to be embryonically lethal and developed normally but was slightly smaller by body weight. Skeletal muscle mitochondria, cardiac mitochondria and MEFS from the Pan et al MCU-/-
2+ mouse exhibited no mCa uptake. In skeletal muscle tissue this led to increased phosphorylation levels of pyruvate dehydrogenase at its inhibitory site (Serine-
293) and the Pan et al mouse had decreased muscle strength activity. MCU KO mitochondria of the Pan et al mouse were also resistant to mitochondrial swelling suggesting resistance to mPTP but in an ex vivo model of ischemia reperfusion
(IR) injury there was no difference in infarct size in the Pan et al mouse(Pan et al., 2013).
Subsequently, two independent labs found that germline deletion of MCU was actually embryonically lethal(Kwong et al., 2015; Luongo et al., 2015).
Luongo et al generated a tamoxifen inducible, cardiac specific MCU conditional
2+ knockout (MCU cKO) mouse and found Mcu deletion completely ablated mCa uptake. This was found to impair cellular bioenergetics limiting contractile reserve under β-adrenergic stress signaling. Mcu ablation also made cardiac mitochondria resistant to permeability transition and provided cardioprotection in an in vivo model of myocardial IR injury(Luongo et al., 2015). Kwong et al also generated a cardiac specific, tamoxifen inducible MCU cKO mouse model and
2+ further confirmed that Mcu deletion ablated mCa uptake negatively impacting cellular bioenergetics but also providing resistance to mPTP during IR injury with cardioprotection being displayed(Kwong et al., 2015). Overall these results suggest that MCU germline deletion is embryonically lethal but in the adult
20 mouse cardiac specific deletion of MCU is dispensable for homeostatic cardiac function although is required for stress responsive signaling.
More recently the architecture of the pore has been revealed by cyro- electron microscopy and nuclear magnetic resonance evaluation of fungal
MCU(Fan et al., 2018; Oxenoid et al., 2016; Yoo et al., 2018). These three independent studies have determined that MCU’s oligomerize into tetramers in a dimer of dimers configuration. The N and C termini of MCU face the mitochondrial matrix space while two transmembrane domains form the pore with a highly evolutionarily conserved DIME motif facing the IMS. The second transmembrane domain is hydrophilic and forms the inner pore responsible for channel capacitance(Oxenoid et al., 2016). The DIME motif forms the narrowest portion of the pore allowing for Ca2+ selectivity(Yoo et al., 2018). The first transmembrane domain forms the outer portion of the channel and interacts with the second transmembrane of the next MCU monomer(Fan et al., 2018). To date the structure of only fungal MCU’s has been solved due to the limitation of this technology as it is not yet feasible to solve the structure of large macromolecular complexes(Yu, 1999). Much more needs to be done to determine mtCU’s true stoichiometry and how its macromolecular composition relates to its function.
EMRE
The next channel constituent was identified using the method, stable isotope labeling by amino acids, in which MCU overexpressing cells were grown in medium containing amino acid isotopes. Protein lysates were MCU-
21 immunoprecipitated to affinity purify the high-MW mtCU complex before samples were processed for mass spectrometry. This method revealed that MCU bound an uncharacterized mitochondrial protein named C22of32 (which is now named essential MCU regulator, EMRE)(Sancak et al., 2013). In this seminal study,
EMRE (~10 kD) was detailed as a single transmembrane protein that was required for uniporter activity as transcription activator-like effector nuclease
2+ (TALEN) technology EMRE deleted HEK-293 cells were incapable of mCa uptake. Others have suggested that the minimal metazoan uniporter consists of both EMRE and MCU, even though some EMRE-lacking protists are fully
2+ capable of mCa uptake(Kovács-Bogdán et al., 2014). EMRE’s topology was further detailed by using direct tagging methods and it was found that the N terminus faces the mitochondrial matrix while the C terminus faces the IMS. This study also found that the transmembrane domain region of EMRE (aa64-aa92) interacted with the first transmembrane domain of MCU while its C-terminal polyaspartate (EDDDDDD) tail interacts with the polybasic region (KKKKKR) within MICU1(Tsai et al., 2016). EMRE’s full physiological relevance is yet to be elucidated as no EMRE knockout mouse model has been published but EMRE’s role in the complex appears to be multifactorial in that it forms the functional pore through MCU interactions and helps promote MICU1’s interaction with MCU by also interacting with MICU1.
22 MICU2 and MICU3
After identifying MICU1, Mootha’s group additionally used genomic sequence analysis to discover two additional paralogs EFHA1 and EFHA2 which are now named MICU2 and MICU3(Plovanich et al., 2013). These genes likely arose from gene duplication events of MICU1 and their mRNA and protein expression has also been shown to be tissue dependent. Thus like their ancestor, MICU1, their tissue dependent expression levels also reflect tissue and cell type specific sensing and decoding of cytoplasmic Ca2+ signals(Paillard et al., 2017). Each paralog also contains very similar EF-hand motifs. MICU2 contains EF-hands at amino acid positions 172-207, 227-262, 293-328, 362-397. MICU3 contains EF- hands at amino acid positions 232-267, 401-436, and 470-505.
MICU2 at the protein level shares 41% amino acid similarity with MICU1.
In the Plovanich et al study it was noted that silencing MICU1 with shRNA in both
HeLa and HEK-293 cells reduced MICU2 expression with the opposite being true as well, that silencing MICU2 lead to reduced MICU1 expression. This suggested that the protein levels of these gene paralogs were dependent on each other. It was further determined that MICU2 localized to the IMS and formed heterodimers with a disulfide bond at cysteine-410 with MICU1 but interestingly was not found to homodimerize(Patron et al., 2014). It was also discovered that while MICU1 is able to interact with MCU, MICU2 is unable to and must incorporate into the high-MW mtCU by heterodimerizing with MICU1(Vecellio
Reane et al., 2016). MICU2’s functional role on channel activity was determined by deleting MICU2 in HEK-293T cells. MICU2 knockout cells had an altered Ca2+
23 threshold for entry at ~0.5 μM Ca2+ while wild-type (WT) cells had a Ca2+ threshold for entry of ~1.5 μM Ca2+(Payne et al., 2017). Additionally, Payne et al found, MICU2 knockout cells also displayed reduced cooperativity suggesting that MICU2 functionally fine-tunes the Ca2+ threshold for entry and cooperativity of channel function. This is important because it indicates that the tissue and pathological dependent expression of MICU2 could alter channel function and activation. In human physiology, MICU2 loss-of-function mutations have resulted in abnormal neurodevelopment leading to cognitive impairments(Shamseldin et al., 2017). A global MICU2 knockout mouse has also been developed using a gene trap strategy and these mice are born at mendelian ratios but do have less longevity which was attributed to the development of diastolic heart disease(Bick et al., 2017)
MICU3’s expression is largely restricted to the skeletal muscle and the central nervous system(Plovanich et al., 2013). At the protein level MICU3 shares
34% amino acid similarity with MICU1 and 47% amino acid similarity with MICU2 with the main difference from its other paralogs being the addition of a 50 amino acid stretch centrally located in the sequence(Patron, Granatiero, Espino, Rizzuto,
& De Stefani, 2019). Patron et al found that MICU3 formed heterodimers with
MICU1 via disulfide bonds via it’s cysteine residue at amino acid 515 but was unable to form ~100 kD heterodimers with MICU2.
This means the MICU family of proteins exist in three possible conformations MICU1 homodimers, MICU1-MICU2 heterodimers, or MICU1-
MICU3 heterodimers. Patron et al overexpressed MICU1, MICU2, or MICU3 alone
24 2+ or in combination in HeLa cells and evoked mCa uptake using histamine as an
2+ agonist. MICU3 addition was found to increase mCa transients the most suggesting MICU3 maybe the most potent activator of the MICU family. No relevant MICU3 knockout cell line or MICU3 knockout mouse exists to date so the true physiological function of MICU3 remains unknown.
MCUR1
2+ Mallilankaraman et al set out to find novel regulators of mCa uptake by performing an RNA interference, RNAi, screen for 45 predicted inner mitochondrial membrane proteins in HEK-293 cells. Following knockdown of each candidate cells were exposed to Ionomycin at levels that enhance plasma
2+ 2+ membrane Ca permeability to induce mCa uptake. Knockdown of mitochondrial calcium uniporter regulator-1, MCUR1 (formerly a protein of unknown function referred to as coiled-coil domain containing 90A, CCDC90A),
2+ inhibited mCa uptake(Mallilankaraman et al., 2015). MCUR1 encodes for a ~40 kD protein with two transmembrane domains where topologically the C-terminus faces the cytosol and the N-terminus faces the mitochondrial matrix.
Mallilankaraman et al found that MCUR1 knockdown lead to diminished cellular bioenergetics suggesting a physiological role for MCUR1. MCUR1 interacts with both MCU and EMRE but not MICU1(Adlakha et al., 2019; Tomar et al., 2016).
Tomar et al additionally found that MCUR1 works as a scaffold factor for proper uniporter assembly as deletion of MCUR1 lead to less high-MW mtCU formation as assessed by blue native page and size exclusion chromatography (SEC).
25 Tomar et al generated a cardiac specific MCUR1 knockout mouse and found physiologically these mice were viable but died 3 weeks after birth. Overall it is thought that MCUR1 is a scaffold factor whose presence is necessary for proper uniporter formation.
MCUB
Sequence analysis revealed that the pore-forming subunit of the channel, MCU, also has a gene paralog named MCUB (originally referred to as CCDC109B gene). This gene encodes for a protein with two similar transmembrane domains joined by an evolutionarily conserved DIME motif and that MCUB shares 50% amino acid homology with MCU overall. shRNA knockdown of MCUB in HEK-
2+ 293 cells was found to increase mCa transients upon histamine stimulation and the expression of MCUB alone in a lipid bilayer resulted in a channel that did not display Ca2+ capacitance leading to the hypothesis that MCUB exerts a dominant negative effect on channel function(Raffaello et al., 2013). MCUB was found to localize with MCU and form oligomers with MCU in a lipid bilayer. MCU and
MCUB’s expression are tissue dependent with the mRNA expression of MCUB in murine samples being highest in the lung, heart, and brain while expression is low in skeletal muscle. It has also been reported that the current density of the channel, a measure of the charge moving across the mitochondrial membrane, is also tissue dependent and is 30 times smaller in mitoplasts derived from cardiac tissue than in mitoplasts derived from skeletal muscle; supporting the notion that
MCUB expression may be a major regulator of uniporter capacity(Fieni, Lee, Jan,
26 & Kirichok, 2012). While this early study of MCUB suggests it may be a powerful way to regulate channel activity how this subunit precisely contributes to channel function and its physiological relevance remain unknown making the study of this gene of great scientific interest.
27
Figure 1-1. Molecular makeup of the Mitochondrial Calcium Uniporter Complex (mtCU). mtCU is a macromolecular channel consisting of many subunits whose precise contribution to the regulation of channel function and activity remain to be fully elucidated. Channel opening largely depends on the cytosolic calcium concentration and the driving force for Ca2+ entry relies on the large electrochemical gradient of -180 mV established across the inner mitochondrial membrane (IMM) by the electron transport chain. Over the last few years the genetic components of the mtCU have been identified and numerous papers have been published describing how each subunit contributes to the function or regulation of mtCU activity. The channel consists of: MCU (blue), which encodes the inward-rectifying pore-forming subunit of the channel. Our lab has shown that the absence of MCU completely ablates all channel function 2+ resulting in no acute mCa uptake; Essential MCU Regulator (EMRE, green) is required for complex formation and the minimal uniporter is thought to consist of MCU and EMRE; Mitochondrial Calcium Uptake 1 and 2 (MICU1, light green, and MICU2, pink) are considered gatekeepers of the complex that modulate open probability via their Ca2+-binding EF-hand motifs positioned in the intermembrane space (IMS); Mitochondrial Ca2+ Uniporter Regulator 1 (MCUR1, purple) is thought to be required for proper channel functionality and is thought to add stability to mtCU formation. Mitochondrial Calcium Uniporter B (MCUB, yellow, formerly CCDC109B gene), was recently identified as a paralog of MCU 2+ and is reported to directly bind MCU and have a negative impact on mCa uptake.
28 CHAPTER 2
MITOCHONDRIAL CALCIUM IN CARDIAC PATHOPHYSIOLOGY
Overview of Cardiac Metabolism
Metabolism is the set of chemical processes that occur within organisms to maintain the fabric of their being. The heart is a highly efficient pump that delivers blood to the body via the circulatory system in order to meet cellular metabolomic demand for oxygen and nutrients. Like almost all eukaryotic cells cardiomyocytes use highly coordinated pathways whereby enzymes and ion channels convert metabolites into chemical energy products which are in turn used at level of the sarcomere to generate the mechanical force of cardiomyocyte contraction in order to effectively deliver blood to the body. The heart’s metabolic needs cannot be understated. It is estimated that on a daily average the human heart pumps
7,200 liters of blood by metabolizing greater than 20 grams of carbohydrates and
30 grams of fat while consuming over 35 liters of oxygen to generate enough
ATP via oxidative phosphorylation (OxPhos) to support human life(Taegtmeyer et al., 2016). The heart indeed has the highest metabolic demand of all organs and cardiac contractility uses 70% of ATP generated in the cardiomyocyte, while the remaining 30% is largely used to maintain ionic gradient(Gibbs, 2003). Roughly,
70% of ATP is generated by OxPhos(Houten & Wanders, 2010), while 30% is derived from glycolysis or other sources such as ketone bodies and amino acids(Doenst, Nguyen, & Abel, 2013). The main metabolites utilized by the heart are fats and carbohydrates. Fatty acid oxidation and glycolysis produce the common end product acetyl coenzyme A (acetyl-CoA), which further undergoes
29 oxidation in the tricarboxylic acid (TCA) cycle in order to generate reducing equivalents (in the form of the nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH2)) which donate their hydrogen’s to the electron transport chain (ETC) to establish a proton-motive force across the IMM responsible for generating ATP through OxPhos(Mailloux et al., 2007). mtCU
2+ mediated mCa uptake is a critical regulator of key steps in metabolism thus
2+ linking mCa with cellular bioenergetics and cardiac contractility.
Fatty Acid Metabolism
Fatty acids (FAs) are the primary fuel source of cardiomyocytes for
ATP generation and are delivered to the heart either as free FAs bound to albumin or as FAs released from triacylglycerols of circulating lipoproteins. FA uptake is facilitated along the plasma membrane by fatty acid transport protein
(FATP), fatty acid translocase (CD36) or the plasma membrane fatty acid-binding protein (FABPpm)(Glatz, van Bilsen, & van der Vusse, 2000). FAs are also able to traverse the plasma membrane themselves. In the cytosol FAs are esterified to fatty acyl-CoA which is further esterified to triglyceride for storage or is shuttled by heart specific fatty acid binding protein (MFABP or FABP3) to the OMM where carnitine palmitoyl transferase I (CPTI) converts fatty acyl-CoA to long-chain acylcarnitine which can now be shuttled into the mitochondria(Boord, Fazio, &
Linton, 2002). Acylcarnitine is then exchanged for carnitine across the IMM via the carnitine:acylcarnitine translocase (CAT) thus transporting acylcarnitine into the mitochondrial matrix(Longo, Amat di San Filippo, & Pasquali, 2006). In the
30 mitochondrial matrix carnitine palmitoyl transferase 2 (CPT2) converts acylcarnitine back to long chain acyl-CoA which now enters the FA β-oxidation pathway which is a set of 4 classical reactions that convert acyl-CoA’s to acetyl-
CoA(Deschauer, Wieser, & Zierz, 2005). In the first step an acyl-CoA-ester is dehydrogenated by acyl-CoA dehydrogenase to yield a trans-2-enoyl-CoA which is then hydrated at the double bond by enoyl-coenzyme A hydratase. In the third step the newly formed L-3-hydroxy-acyl-CoA is dehydrogenated to 3-keto-acyl-
CoA which then undergoes thiolytic cleavage to produce acetyl-CoA with the fatty
Acyl-CoA now being 2 carbons shorter(Houten & Wanders, 2010). Every β- oxidation cycle produces one NADH, one FADH2, an acyl-CoA that is two carbons shorter, and an acetyl-CoA that can now enter the TCA cycle. The fatty acyl-CoA can now further undergo β-oxidation until all carbons are converted to acetyl-CoA.
Glycolysis
Glycolysis is the metabolic process by which one glucose molecule is broken down into two pyruvate molecules. Glucose uptake in cardiomyocytes is facilitated by glucose transporters with GLUT4 being the most abundant transporter in the heart(Abel, 2004). Once in the cytosol glucose enters glycolysis which is a 10 step set of enzymatic reactions that can be separated into two main phases, the preparatory phase where ATP is consumed and the pay-off phase during which ATP is produced(Berg, Stryer, & Tymoczko, 2002).
31 In the first step an ATP is consumed whereby hexokinase phosphorylates glucose to form the negatively charged glucose 6-phosphate which is no longer cell permeable and is trapped intracellularly(Gibb & Hill, 2018). Glucose-6- phosphate is then isomerized by phosphoglucose isomerase into fructose 6- phosphate(Bahaji et al., 2015). In the next step fructose 6-phosphate is phosphorylated by phosphofructokinase consuming another ATP to produce fructose 1,6-bisphosphate(Dunaway, 1983). Fructose 1,6-bisphosphate is then split by aldolase into the two three carbon molecules, dihydroxyacetone phosphate and glyceraldehyde 3-phosphate, which marks the beginning of the pay-off phase of glycolysis(Chang, Yang, Tien, Yang, & Hsiao, 2018).
Dihydroxyacetone phosphate is isomerized by triose phosphate isomerase into glyceraldehyde 3-phosphate(Wierenga, Kapetaniou, & Venkatesan, 2010). The two glyceraldehyde 3-phospates are converted into 1,3-bisphosphoglycerate by glyceraldehyde 3-phosphate dehydrogenase and as a byproduct NAD+ is converted into NADH(Kosova, Khodyreva, & Lavrik, 2017). 1,3- bisphosphoglycerate is then converted into 3-phosphoglycerate by phosphoglycerate kinase generating one ATP as a byproduct(Bowler, 2013). 3- phosphoglycerate is converted by phosphoglycerate mutase into 2- phosphoglycerate(Dhar et al., 2010). Enolase converts 2-phosphoglycerate into phosphoenolpyruvate(Ji et al., 2016). In the final step phosphoenolpyruvate is converted to pyruvate by pyruvate kinase generating an ATP as a byproduct of this reaction(Israelsen & Vander Heiden, 2015). During the preparatory phase 2
ATPs are consumed but during the pay-off phase 2 ATPs are generated per
32 three carbon source thus 2 net ATPs are produced per glucose molecule.
Through glycolysis one glucose is catabolized into two pyruvates whereby 2 net
ATP and 2 NADH’s are produced(Berg et al., 2002).
The committed, irreversible steps of glycolysis are mediated by hexokinase, phosphofructokinase, and pyruvate kinase. Phosphofructokinase also mediates the most rate limiting step of glycolysis(Berg et al., 2002). The two pyruvates and 2 NADH further enter other energy producing pathways. The
NADH molecules are used as hydrogen donors to maintain the proton electrochemical gradient of the inner mitochondrial membrane established by the electron transport chain which is critical for generating ATP by OxPhos. The two pyruvate molecules now enter the mitochondria and are translocated across the
IMM by the pyruvate carrier and are converted into acetyl-CoA by the pyruvate dehydrogenase complex(X.-B. Li, Gu, & Zhou, 2015).
Pyruvate Dehydrogenase Complex
The pyruvate dehydrogenase (PDH) complex is the central component that links glycolysis to OxPhos by decarboxylating pyruvate into acetyl-CoA and carbon dioxide (CO2) also generating one NADH in the process. The mammalian PDH complex is a massive complex with a molecular weight of ~9 Megadaltons that consists of multiple copies of three catalytic enzymes referred to as E1, E2, and
E3(M. S. Patel & Roche, 1990). E1 is the thiamin diphosphate dependent enzyme called pyruvate dehydrogenase, E2 is a dihydrolipoamide acetyltransferase enzyme, and E3 is the dihydrolipoamide dehydrogenase enzyme(Reed, 2001).
33 Proposed models suggest as many as 60 copies of E2 and 12 copies of the E3 binding protein form the core of the PDH complex which also bind 20-30 heterotetramers of E1 and 6-12 homodimers of E3 although the exact stoichiometry of the mammalian PDH complex has yet to be revealed(Mulchand
S. Patel, Nemeria, Furey, & Jordan, 2014). E1 decarboxylates pyruvate to CO2 and forms a C2α-hydroxyethylidene enamine intermediate while also acetylating lipoyl groups attached to the E2(Bunik, Tylicki, & Lukashev, 2013). E2 transfers the acetyl group to CoA producing acetyl-CoA. E3 facilitates the transfer of electrons of the dihydrolipoyl moieties of E2 to FADH which are then transferred to NAD+ reducing it to NADH(Harris, Bowker-Kinley, Huang, & Wu, 2002).
In humans the PDH complex is tightly regulated through phosphorylation.
When serine’s at amino acid sites 203, 264, and 273 of the E1 subunit are phosphorylated by pyruvate dehydrogenase kinases (PDK, isoforms 1-4) PDH complex activity is inhibited(Korotchkina & Patel, 2001). Pyruvate dehydrogenase phosphatases (pyruvate dehydrogenase phosphatase subunit 1 (PDP1) also known as protein phosphatase 2C (PP2C)) dephosphorylate these sites to activate PDH activity(Lander et al., 2018).
Tricarboxylic Acid Cycle
FAs and glucose, the hearts major fuel sources, have thus been catabolized to acetyl-CoA which is the metabolic entry point for the TCA cycle (also known as the citric acid cycle or the Krebs cycle) within the mitochondrial matrix. The TCA cycle is a set of metabolic processes in which 8 enzymes oxidize acetyl-CoA to
34 CO2 while also generating energy products in the form of ATP, NADH, and
FADH2 through each turn of the cycle.
In the first step citrate synthase forms citrate by transferring the acetyl group of acetyl-CoA to oxaloacetate(Wiegand & Remington, 1986). Next, aconitase isomerizes citrate into isocitrate(Lushchak, Piroddi, Galli, & Lushchak,
2014). Isocitrate dehydrogenase (IDH) then oxidizes and decarboxylates isocitrate into α-ketoglutarate generating NADH in the process(Al-Khallaf, 2017).
α-ketoglutarate dehydrogenase (α-KDH) decarboxylates α-ketoglutarate into succinyl-CoA generating NADH in the process(Vatrinet et al., 2017). Succinyl-
CoA synthetase converts succinyl-CoA into succinate by substrate level phosphorylation generating GTP/ATP in the process(Furuyama & Sassa, 2000).
Succinate dehydrogenase oxidizes succinate into fumurate generating
FADH2(Hederstedt & Rutberg, 1981). Fumarase then hydrates fumarate into malate(Yogev et al., 2010). Malate dehydrogenase then oxidizes malate into oxaloacetate generating NADH(Minarik, Tomaskova, Kollarova, & Antalik, 2002).
Oxaloacetate is now regenerated and if acetyl-CoA is present the TCA cycle can begin again. For each turn of the TCA cycle 3 NADH’s, 1 FADH2, and 1
GTP/ATP has been generated(Akram, 2014). NADH and FADH2 are further oxidized by the ETC where the transfer of electrons are coupled to the translocation of protons across the IMM generating an electrochemical gradient which drives synthesis of ATP in a process known as OxPhos.
35 Oxidative Phosphorylation
The metabolism of fats, carbohydrates, and proteins generate the reducing equivalents NADH and FADH2 which donate electrons to the ETC which passes the electrons onto molecular oxygen (O2) releasing a large amount of free energy to synthesize ATP in a process termed OxPhos. The ETC consists of Complexes
I-IV which are all located in the inner mitochondrial membrane and the two mobile electron carriers ubiquinone (also known as Coenzyme Q) and cytochrome c. At the level of Complex I (CI, also known as the NADH:ubiquinone reductase), NADH first donates two electrons to flavin mononucleotide (FMN) reducing it to FMNH2 which in turn is oxidized and electrons are donated to an
Iron-sulfur clusters before being transferred to ubiquinone forming ubiquinol(Hirst, 2013). This facilitates the transfer of 4 H+ from the matrix to the
IMS(Mourier & Larsson, 2011). Through Complex II (CII, also known as succinate dehydrogenase or succinate:quinone oxidoreductase) succinate donates electrons to FAD reducing it to FADH2 which then transfers electrons to quinone(Cecchini, 2003). CII is the only ETC member that does not directly transfer electrons to the IMS but it is unique because it also catalyzes the oxidation of succinate to fumurate so it contributes to the TCA cycle as well as
OxPhos(Y. R. Chen & Zweier, 2014). At Complex III (CIII, also known as the
Coenzyme Q:cytochrome c reductase) two electrons are transferred from ubiquinol to cytochrome c and two electrons are passed to quinone reducing it to quinol ultimately resulting in four H+ being translocated to the intermembrane space(Bleier & Dröse, 2013). At Complex IV (CIV, also known as cytochrome c
36 oxidase) four electrons are transferred from four cytochrome c’s to O2 resulting in four H+ being translocated from the mitochondrial matrix to the IMS(Y. Li, Park,
Deng, & Bai, 2006). CI-IV has thus established a proton motive electrochemical gradient where the H+ concentration has increased in the IMS resulting in the -
180 mV membrane potential established along the IMM(Mitchell, 1966). Complex
V (CV, also known as the F1FO ATP synthase) utilizes this electrochemical gradient to generate ATP. The FO component acts as an ion channel and allows
+ H to re-enter the mitochondrial matrix while the hydrophilic F1 component works as a rotary motor to actually catalyze ATP generation(Jonckheere, Smeitink, &
Rodenburg, 2012). The translocation of nine to twelve H+ cause one complete
turn of the F1FO ATP synthase generating ~3 ATP(Mailloux et al., 2007).
OxPhos generates the most amount of ATP in aerobic organisms and 26 of the 30 ATPs generated during full glucose oxidation are generated by
OxPhos(Berg et al., 2002). The phosphate/oxygen (P/O) ratio refers to the amount of ATP produced by two electrons moving through the ETC. The oxidation of NADH and FADH2 by the ETC generates ATP at a physiological P/O ratio where ~2.5 ATP are produced for every NADH oxidized and ~1.5 ATP are produced for every FADH2 oxidized(Hinkle, 2005). One glucose molecule that has undergone glycolysis has generated net 2 ATP, 2 NADH, and two pyruvates.
The two pyruvates generated by glycolysis undergo decarboxylation into acetyl-
CoA by the PDH complex making one NADH for each pyruvate. The catabolism of each pyruvate in the TCA cycle generates 1 ATP, 3 NADH, and 1 FADH2.
After NADH and FADH2 donate their electrons to the ETC to generate ATP one
37 molecule of fully oxidized glucose has produced ~30 ATP. In the case of FAs the
16 carbon palmitate is the most predominant fatty acid energy source of the heart and when palmitate fully undergoes β-oxidation it will produce 8 acetyl-CoA’s that feed into the TCA cycle and generate ~106 ATP through OxPhos(Schönfeld &
Reiser, 2013).
Mitochondrial Calcium’s Influence on Cellular Bioenergetics
2+ Mitochondrial energetics are regulated by mCa at multiple control points. The
PDH complex is regulated by phosphorylation and the PDH phosphatase responsible for dephosphorylation of these sites is activated by Ca2+ directly
2+ implicating mCa in regulating the link between the glycolytic pathway and
OxPhos(Huang et al., 1998). Our lab has indeed shown that cardiac specific
2+ deletion of Mcu (ablating mCa uptake) results in increased phosphorylation status of PDH even when hearts are stimulated with isoproterenol suggesting
2+ mCa dependent activation of PDH is directly implicated in β-adrenergic stress signaling(Luongo et al., 2015).
2+ 2+ The TCA cycle is also regulated by mCa . Increased mCa lowers the Km of both α-KDH and IDH making these enzymes more accessible to their respective substrates(Rutter & Denton, 1988). In eukaryotes IDH exists as an octamer and 2 Ca2+ are thought to bind each octamer although the exact Ca2+ binding site is unknown(Tarasov, Griffiths, & Rutter, 2012). α-KDH, similar to the
PDH complex, consists of multiple copies of three subunits (similarly referred to
2+ as E1, E2, and E3) and it has been suggested that the E1 subunit is Ca
38 sensitive(Lawlis & Roche, 1981). Interestingly, Ca2+ only activates IDH and α-
2+ KDH from mitochondria of organisms that exhibit mtCU mCa uptake where as
2+ IDH and α-KDH from plants or insects that do not exhibit mtCU mediated mCa
2+ 2+ uptake are insensitive to Ca suggesting mtCU mediated mCa makes the TCA cycle susceptible to Ca2+ sensitivity(James G. McCormack, Daniel, Osbaldeston,
Rutter, & Denton, 1992);(J. G. McCormack & Denton, 1981).
Ca2+ also influences cellular bioenergetics through the modulation of
2+ OxPhos. Ca activation of the F1FO ATP synthase increases its maximal velocity but it is unclear how this actually occurs because there is no evidence for direct
2+ interaction of Ca with the F1FO ATP synthase (Territo, Mootha, French, &
Balaban, 2000). It is thought that Ca2+ activation could be dependent on post- translational modifications as phosphorylation of the γ-subunit has been shown to
2+ be sensitive to mCa (Hopper et al., 2006). Even more recent work has shown that the Ca2+ sensitive EF-hand containing S100A1 protein can interact and
2+ activate with the F1-ATP synthase and this maybe how Ca influences the maximal velocity(Boerries et al., 2007).
2+ To summarize mCa influences energy production by increasing the amount of fuel to feed the TCA cycle by activating the PDH complex, by increasing the activity of TCA cycle dehydrogenases thus stimulating the TCA cycle, and by directly activating ATP generation through modulation of the F1FO
ATP synthase.
39 Overview of Myocardial Ischemia Reperfusion Injury
Prevalence of Ischemic Heart Failure
Cardiovascular diseases (CVD) are a group of disorders of the circulatory system and the heart which contribute to 31% worldwide mortality making CVD the number one cause of death worldwide(Benjamin et al., 2017). Ischemic heart disease is the leading contributor of CVD mortality being the primary cause of death for approximately 9.4 million people every year(Vos et al., 2017). Due to its prevalence CVD is major health concern and the goal of the World Health
Organization is to reduce the number of premature deaths from CVD by 25% by
2025(Allen, 2017). In order to meet this goal new therapies are needed to treat ischemic heart disease and in order to do so we must understand the pathological signaling that contributes to ischemic heart disease.
Pathophysiology of Myocardial Ischemia Reperfusion Injury
Atherosclerosis is a coronary artery disease caused by hyperlipidemia where fats accumulate within arteries and become oxidized resulting in the formation of plaques(Hennekens & Gaziano, 1993). Over time a fibrous layer of smooth muscle cells forms over the plaque and hardens narrowing arteries. Vulnerable plaques erode or rupture resulting in thrombosis, occlusion of the coronary arteries, and disruption of blood flow subjecting downstream myocardium to ischemia(Rafieian-Kopaei, Setorki, Doudi, Baradaran, & Nasri, 2014).
The ischemic zone of the myocardium is hypoxic which depresses
OxPhos causing a loss of ATP production leading to metabolic
40 derangement(Kubasiak, Hernandez, Bishopric, & Webster, 2002). In order to generate ATP to continue cardiac contraction and to maintain ionic gradients cardiomyocytes switch their main substrate source from FAs to glucose which is now metabolized through anaerobic glycolysis resulting in excess production of lactate and H+(Monassier, 2008). H+ is exchanged for extracellular Na+ along the plasma membrane via the Na+/H+ exchanger increasing intracellular Na+ levels which in turn makes the plasma membrane NCX operate in reverse mode,
2+ further increasing iCa levels(S. Chen & Li, 2012).
The most effective form of therapy for the treatment of myocardial infarction (MI) is reperfusion of the occluded artery by balloon angioplasty but rapid restoration of oxygenated blood greatly augments oxidative stress through the production of reactive oxygen species (ROS). ROS are generated during reperfusion through several mechanisms. Xanthine Oxidase and NADPH oxidase have been shown to generate ROS following IR injury(Granger & Kvietys, 2015).
Tetrahydrobiopterin (BH4) depletion during ischemia leads to endothelial nitric oxide synthase (eNOS) uncoupling which further augments ROS generation because monomeric eNOS produces superoxide(Barr et al., 2017; Elrod John et al., 2006). The largest source of ROS generation following I/R injury comes from the reactivation of aerobic metabolism itself which increases OxPhos activity and thus mitochondrially produced ROS. It has even been suggested that dysregulated conformations of Complex I of the ETC due to ischemic injury may even further augment ROS production(Maklashina et al., 2002).
41 ROS further contributes to I/R pathogenesis. ROS causes lipid peroxidation which leads to membrane permeability and cellular swelling(Catalá
& Díaz, 2016). ROS inhibits the Na+/K+ ATPase increasing intracellular Na+
2+ levels further increasing the activity of reverse mode NCX augmenting iCa levels(Inserte, Garcia-Dorado, Hernando, & Soler-Soler, 2005). ROS also inhibits
2+ SERCA which further elevates iCa levels(Sharov, Dremina, Galeva, Williams, &
Schoneich, 2006). Mitochondria are exposed to these pathogenic increases in
2+ 2+ iCa levels and become overloaded with Ca .
mPTP Driven Necrosis
2+ I/R injury generates ROS and elevates iCa levels which have both been long known to be key activators of inner mitochondrial membrane permeability
2+ transition. mCa overload results in superoxide generation and activation of a large non-specific channel (called the mitochondrial permeability transition pore, mPTP) along the IMM that freely allows ions and solutes as big as 1.5 kD to move across the IMM(Bernardi et al., 2006). mPTP causes matrix swelling which collapses the mitochondrial membrane potential and the loss of the electrochemical gradient along the IMM means OxPhos can no longer generate
ATP(A. P. Halestrap, Clarke, & Javadov, 2004). In fact the matrix environment actually now promotes ATP hydrolysis by the F1FO ATP synthase(Andrew P.
Halestrap & Pasdois, 2009). This results in the shift to glycolysis seen in cardiomyocytes during I/R injury and if ATP generation can no longer keep up
42 with cellular demand to maintain cardiac contraction and ionic gradients cells will undergo necrotic cell death(Lambert, Nicholson, Amin, Amin, & Calvert, 2014).
Necrosis leads to infarcted tissue and loss of viable myocardium. The infarct size is determined by the size of the area at risk (AAR), duration of occlusion of the artery, and the amount of residual collateral blood flow and dysfunction of the coronary microvasculature(Kalogeris, Baines, Krenz, &
Korthuis, 2012). Infarcted hearts also display reduced cardiac function which is a major risk for sudden death. Individuals that undergo reperfusion treatment may survive but are predisposed to recurrent MI’s and the further development of heart failure due to pathological remodeling of the heart(Sutton Martin & Sharpe,
2000). To date there is no effective therapy for the treatment of MI beyond reperfusion of the occluded artery although research is highly focused on therapies that replace the lost cardiomyocytes (stem cells) or therapies that prevent necrotic cell death due to mPTP activation.
Molecular Components of mPTP
2+ Because mCa induced mPTP activation results in necrotic cell death the molecular components of the mPTP have long been sought after in order to target them for therapeutic development for the treatment of MI and have been the subject of much debate over the years. Cyclophilin D (encoded by the Ppif gene) has been the only putative member identified as belonging to the mPTP that functions in gating of the pore. Cyp D is a mitochondrially localized enzyme belonging to the peptidyl cis-trans isomerase family(Griffiths & Halestrap, 1991).
43 PPIs have protein folding capacities and catalyze peptide bond isomerization from cis to trans conformation immediately before proline residues.(Hazelton,
Petrasheuskaya, Fiskum, & Kristián, 2009). Cyp D was discovered as a mPTP member because the immunosuppressant Cyclosporine A (CsA) was found to inhibit Cyp D and in turn prevent calcium induced opening of the mPTP(Crompton, Ellinger, & Costi, 1988; A. P. Halestrap & Davidson, 1990).
The generation of a Ppif knockout mouse (Ppif-/-) allowed for determination of
Cyp D’s exact physiological role. Mitochondria from Ppif-/- mice were resistant to
Ca2+ induced swelling but eventually did undergo permeability transition at very high Ca2+ levels (greater than 2 mM Ca2+) suggesting Cyp D regulates the pore but is not the pore itself(Baines et al., 2005). Additionally, this same group found that the peptidyl cis-trans isomerase activity of Cyp D was critical in regulating mPTP as a prolyl isomerase-deficient mutant form of Cyp D was unable to restore Ca2+ induced mPTP in Ppif-/- mitochondria.
Various other possible genetic members of the mPTP have been proposed such as ANT, VDAC, SPG7, and the c-ring of the F1FO ATP synthase but all studies have indicated that mitochondria or cells lacking these members are still able to undergo permeability transition suggesting they only modulate activity of the pore but are unlikely the actual molecular identity of the pore forming subunit(Bernardi, Rasola, Forte, & Lippe, 2015).
44 RATIONALE AND OBJECTIVE OF CURRENT STUDY
2+ Preventing mCa Induced mPTP as a Therapeutic for MI
With the realization that CsA regulated permeability transition by inhibiting Cyp D and with the generation of the Ppif-/- mouse it offered the study of both genetic and pharmacologic inhibition of mPTP in vivo allowing for investigation into
2+ potential ways to limit mCa induced necrosis in the setting of disease such as
MI or IR injury. Ppif-/- mice subjected to IR injury exhibited reduced infarct size compared to controls suggesting Cyp D is an attractive target for the treatment of ischemic heart disease(Nakagawa et al., 2005). Sanglifehrin-A, another Cyp D inhibitor, given at the time of reperfusion reduced infarct size in rats following cardiac IR injury(Hausenloy, Duchen, & Yellon, 2003). CsA and NIM-811 (a more specific Cyp D inhibitor) treatment limited infarct size in rabbit models of IR injury compared to control treatment(Argaud et al., 2005). These basic science studies provided strong evidence that targeting Cyp D could be a powerful way to limit
2+ mCa induced mPTP activated necrotic cell death during IR injury. CsA was already approved as an immunosuppressive drug by the FDA so human clinical trials soon followed to establish if CsA could be used to target mPTP for the treatment of myocardial IR injury(Hallworth, 2014). These clinical trials have shown mixed results with some showing that a single bolus of CsA given at the time of reperfusion can reduce infarct size in patients with ST-elevation myocardial infarction compared to placebo controls (Piot et al., 2008) while other studies show no improvement in function between CsA treatment and placebo groups following acute MI(Cung et al., 2015; Rahman et al., 2018).
45 One of the pitfalls of CsA treatment is that it is immunosuppressive, has a biphasic dose response curve which decreases its ability to be combined with dual therapies, and is a non-selective inhibitor which also inhibits Calcineurin which maybe problematic in the heart because calcineurin contributes to the physiological remodeling of the heart following MI(Øie, Bjørnerheim, Clausen, &
Attramadal, 2000). This makes CsA treatment problematic and its use in clinical trials has now stopped but novel mPTP inhibitors that are more selective for the
2+ actual pore or novel ways to regulate mCa mediated mPTP activation are very attractive areas of research for therapeutics.
2+ Modulation of mCa Exchange during Ischemic Heart Disease
While much work has been focused on preventing mPTP by directly inhibiting the pore the recent discovery of the genetic identities of mtCU components and the
2+ confirmation of the discovery by our lab of the genetic identity of the mCa efflux pathway, NCLX, has spurred investigation into understanding how the
2+ modulation of mCa exchange is implicated in cardiac pathophysiology.
NCLX’s Role in Ischemic Cardiac Stress
The mitochondrial Na+/Ca2+ exchanger (NCLX encoded by the SLC8B1 gene) is
2+ the primary mechanism for mCa efflux. Our lab was the first to investigate how genomic deletion of NCLX impacted cardiac physiology by generating a floxed
NCLX mouse, Slc8b1fl/fl, which we subsequently crossed to the αMHC-
MerCreMer mouse model so we could conditionally delete NCLX using tamoxifen
46 in a cardiac specific manner (NCLX cKO)(Luongo et al., 2017). Interestingly, we found that 87% of NCLX cKO mice died within 2 weeks of tamoxifen delivery due to fulminant heart failure suggesting NCLX is critical to maintain homeostasis and viability. We additionally found that NCLX cKO cardiac mitochondria were pre- swollen and when subjected to a large Ca2+ bolus NCLX cKO mitochondria swelled at a greater rate and to greater extent than control mitochondria suggesting they were susceptible to permeability transition.
2+ 2+ Because deletion of the mCa efflux pathway enhanced mCa induced
2+ mPTP activation we next postulated that mCa efflux may prevent permeability
2+ transition by clearing mitochondria of pathogenic increases in mCa . To investigate this line of questioning we generated a NCLX gain-of-function mouse model by cloning human SLC8B1 into a plasmid containing tetracycline responsive promoter (TRE-NCLX). We crossed the TRE-NCLX mouse to the
αMHC-tTA (cardiomyocyte-restricted doxycycline-off) transgenic mouse so we could conditional overexpress NCLX by feeding mice doxycycline chow. We found that in isolated adult cardiomyocytes (ACMs) NCLX overexpression
2+ increased the rate of mCa efflux by 38% and very importantly provided resistance to Ca2+ induced permeability transition. We next evaluated if NCLX overexpression could prevent mPTP during I/R injury by subjecting mice to 40 minutes of ischemia and 24 hours of reperfusion. NCLX overexpression reduced infarct size by 43% compared to control. We additionally found that NCLX overexpression preserved cardiac function following a permanent model of MI by reducing cardiomyocyte cell death.
47 2+ 2+ Our study suggests that enhancing mCa efflux prevents mCa induced mPTP mediated necrosis during ischemic cardiac stress and provides further
2+ rationale for investigating how modulation of mCa exchange influences ischemic cardiac stress.
MCU During Ischemic Cardiac Stress
Our lab found that germline deletion of MCU was embryonically lethal so we generated an MCUfl/fl mouse which we crossed to the αMHC-MerCreMer mouse model so we could conditionally delete MCU using tamoxifen in a cardiac specific manner (MCU cKO)(Luongo et al., 2015). We found MCU deletion ablated acute
2+ mCa uptake and did not lead to a baseline cardiac phenotype. MCU cKO
2+ cardiac mitochondria were resistant to mCa induced permeability transition.
When MCU cKO mice were subjected to IR injury they displayed a ~45% reduction in infarct size compared to controls. MCU deletion did come at a cost though as we observed that MCU cKO mice lacked contractile responsiveness to isoproterenol induced cardiac contractility due to a diminishment of cellular
2+ bioenergetics. These data strongly suggest that reducing mCa uptake can
2+ prevent mCa induced mPTP driven necrosis in ischemic cardiac injury.
MCUB’s Role in Ischemic Cardiac Stress
MCUB is suggested to exert a dominant negative effect on channel activity although it’s role in cardiac pathophysiology has never been investigated. Also, because MCU deletion was found to come at the cost of limiting cellular
48 bioenergetics it is of immense interest to understand precisely how MCUB exerts its negative function on mtCU and determine whether there are any physiological differences in its regulatory activity compared to MCU deletion alone.
49
Figure 2-1. Mitochondrial Calcium Control of Cellular Bioenergetics. 2+ Increases in cytosolic calcium levels activate mtCU mediated mCa uptake. 2+ mCa is a central control point at multiple levels of cellular bioenergetics by influencing the regulation of enzymatic activity. Pyruvate dehydrogenase phosphatase 1 (PDP1) is directly activated by Ca2+ binding and PDP1 dephosphorylates Serine 203, Serine 264, and Serine 273 of pyruvate dehydrogenase (E1). This activates E1 and begins the catalysis of decarboxylation of pyruvate into acetyl coenzyme A. Acetyl-CoA enters the 2+ 2+ tricarboxylic acid cycle. The TCA cycle is also regulated by mCa as Ca lowers the Km of isocitrate dehydrogenase and α-ketoglutarate dehydrogenase making 2+ these TCA cycle members more accessible to their substrates. mCa also binds to EF hands of the S100A protein. Activated S100A1 in turn activates the F1 subunit of the ATP synthase (CV) increasing its maximal velocity.
50 CHAPTER 3
MCUB REGULATES THE MOLECULAR COMPOSITION OF THE
MITOCHONDRIAL CALCIUM UNIPORTER CHANNEL TO LIMIT
MITOCHONDRIAL CALCIUM OVERLOAD DURING STRESS
Jonathan P. Lambert1, Timothy S. Luongo1, Dhanendra Tomar1, Pooja Jadiya1,
Erhe Gao1, Xueqian Zhang1, Anna M. Lucchese1, Devin Kolmetzky1, and John
W. Elrod1
1Center for Translational Medicine, Temple University School of Medicine,
Philadelphia, PA, 19140
Abstract
Background: The Mitochondrial Calcium Uniporter (mtCU) is a macromolecular channel located at the inner mitochondrial membrane and is necessary for
2+ mitochondrial calcium (mCa ) uptake. Here we detail the contribution of MCUB, a paralog of the pore forming subunit - MCU, in mtCU regulation and function. Methods and Results: Using genetic gain- and loss-of-function approaches we show that MCUB expression displaces MCU from the high- molecular weight complex and thereby decreases the binding of MICU1 and
MICU2 to the mtCU to alter channel gating. This results in a loss of MICU1/2
2+ cooperative activation of the uniporter and decreases mCa uptake. Employing mice with conditional cardiomyocyte-specific expression of MCUB we show that
51 MCUB incorporation into the mtCU is a stress-responsive system to limit mitochondrial calcium overload during cardiac injury. Indeed, overexpression of
MCUB is sufficient to decrease infarct size following ischemia-reperfusion injury.
However, we also show that increased MCUB incorporation into the mtCU does come at a cost, as MCUB-mediated downregulation of mtCU function impairs mitochondrial energetics during stress. Conclusions: In summary, we detail a new regulatory mechanism to modulate mtCU function and mitochondrial calcium uptake. Our results suggest that the stoichiometry of the subunits associated with the uniporter is critical to its function and that MCUB is a prominent regulator of uniporter composition.
Introduction
The mitochondrial calcium uniporter (mtCU) is an inward-rectifying, high- molecular weight (MW) channel that resides in the inner mitochondrial membrane and is necessary to facilitate calcium (Ca2+) uptake into the mitochondrial matrix(Baughman et al., 2011). Ca2+ flux through the mtCU is driven by the large electrochemical gradient (-180 mV) across the inner mitochondrial membrane that is generated by the electron transport chain (Vos et al.; Zhao et al., 2013).
Physiologically, the mtCU is required for the activation of energetic pathways involved in oxidative phosphorylation. In addition, during stress the mtCU
2+ mediates mCa overload, which is a prominent activator and sensitizer of the mitochondrial permeability transition pore (MPTP) resulting in cell death. Our lab
2+ has previously shown that genetic modulation of either mCa uptake(Luongo et
52 al., 2015) or efflux(Luongo et al., 2017) is a powerful way to limit cardiomyocyte death in the context of ischemia-reperfusion (IR) injury and heart failure and therefore understanding the molecular regulation of these pathways holds therapeutic promise.
The mtCU consists of multiple components each with a unique role in channel regulation and function. MCU encodes the pore-forming subunit of the channel and is also the target of the MCU-inhibitor, ruthenium red, a pharmacological inhibitor long known to inhibit mitochondrial Ca2+ uptake
2+ (mCa )(Yuriy Kirichok, Grigory Krapivinsky, & David E. Clapham, 2004). Loss of
2+ MCU completely ablates all channel function resulting in no acute mCa uptake(Luongo et al., 2015). MCU is thought to form homooligomers with a highly-conserved ‘DIME’ sequence motif serving as the ion selectivity filter and the second transmembrane domains contributing to the formation of a hydrophilic pore spanning the IMM(Oxenoid et al., 2016). Mitochondrial Calcium Uptake 1
(MICU1) was the first component of the complex identified through comparative physiology and evolutionary genomic approaches(Perocchi et al., 2010). MICU1 contains two Ca2+-binding EF-hands and is reported to be a channel gatekeeper
2+ by maintaining closure of the mtCU at low cCa levels (>2 µM)(Paillard et al.,
2017). MICU1 also contributes to the cooperative activation of the channel by
2+ increasing uptake rates at high cCa levels(Csordas et al., 2013). MICU2 is a gene paralog of MICU1 and is reported to calibrate MICU1-mediated regulation of the uniporter complex(Payne et al., 2017). SMTD1 encodes for Essential MCU
Regulator (EMRE), which also appears to be required for mtCU function,
53 suggesting that the minimal functional channel may consist of MCU and
EMRE(Sancak et al., 2013). EMRE is reported to be necessary for
MICU1/MICU2 interactions with MCU and therefore is also a contributor of linking channel gating with activity(Sancak et al., 2013);(Vais et al., 2016). MCUR1
(Mitochondrial Calcium Uniporter Regulator 1) has been identified as another component that modulates mtCU function, possibly serving as a scaffold for channel formation(Tomar et al., 2016). MCUB was recently put forth as a possible channel constituent because of its close sequence similarity to MCU, encoding for a protein that shares 50% aa homology and contains two similar transmembrane domains linked by an identical DIME motif(Raffaello et al., 2013).
MCUB was found to form hetero-oligomers with MCU and expression of MCUB alone in a lipid-bilayer resulted in no Ca2+ exchange across an artificial membrane(Raffaello et al., 2013). This early study of MCUB strongly suggests it’s a bona fide mtCU constituent, but its unclear whether MCUB directly influences Ca2+ permeation of the pore or regulates the mtCU through some other unknown mechanism. Also, MCUB’s physiological relevance remains unresolved. Here using genetic loss- and gain-of-function approaches we report
2+ that MCUB modulates the stoichiometry of mtCU subunits to fine-tune mCa uptake. Further, MCUB is a prominent regulator of cardiac function by
2+ significantly impacting mitochondrial energetics and reducing mCa overload
(mPTP activation) during stress.
54 Experimental Procedures
Generation of stable MCUB-/- HeLa cell line
We generated a stable MCUB knockout HeLa cell line using CRISPR/Cas9n technology following the Nature protocol developed by the Zhang lab(Ran et al.,
2013). We obtained the pSpCas9n-2A-Puro (PX462) developed by the Zhang lab from Addgene (plasmid #48141) and created guide RNA’s using the optimized
CRIPSR Design Tool at crispr.mit.edu to target exon 1 of MCUB (Chromosome
4: 109,560,205-109,688,726) using a double nickase strategy (single strand cuts) to avoid off- target genomic editing. guide RNA’s were ordered as oligomers from
Eurofins genomics. Guide RNA’s utilized, scored with no off target sites when used for CRISPR/Cas9n:
Nick 1 (581 nts downstream from start of MCUB gene)
5’ CACCGAGCTTGGGCGGGCTCACGGG 3’
5’ AAACCCCGTGAGCCCGCCCAAGCTC 3’
Nick 2 (604 nts downstream from start of MCUB gene)
5’ CACCGCCGCGGCGCGCTGAGCGAGC 3’
5’ AAACGCTCGCTCAGCGCGCCGCGGC 3’
Oligomers were annealed using T4 Polynucleotide kinase, ligated into the Cas9n plasmid via a BbsI restriction enzyme site, and the ligation reaction was treated with a PlasmidSafe exonuclease according to the manufacturer’s instructions
(Lucigen Plasmid-Safe ATP-Dependent DNase, E3101K). Product was PCR cleaned, transformed into electrocompetent DH10B bacteria, and grown on ampicillin resistance plates for selection. Colonies were picked, grown overnight
55 in LB media with 100 μg ml-1 of ampicillin, and DNA was purified according to manufacturer’s instructions (Qiagen, QIAprep Spin Minprep Kit). Successful insertion of guide RNA’s was confirmed via an inability of the generated plasmid to be digested by BbsI due to the elimination of the restriction enzyme site. The plasmids generated for each nick site were simultaneously transfected into HeLa cells using the Invitrogen Lipofectamine 3000 transfection kit according to the manufacturer’s instructions, selected under puromycin resistance at 5μgrams/mL for 96h, and single cell clonal isolations were performed to generate multiple
MCUB knockout clonal HeLa cell lines. In order to confirm if MCUB was genetically deleted we then evaluated gene and protein expression by both qPCR and western blot analysis. MCUB-/- L2 was used for all experiments in the manuscript. qPCR primers designed to confirm deletion of MCUB.
Forward
5’ GTGTGAAGCTGTGTGGAAATG 3’
Reverse
5’ CAAGGGAAGGCCATGTCTATAA 3’
MCUB Antibody used to confirm KO
Santa Cruz-CCDC109B (S-15) (sc-163985, Lot #A1515)
Generation of a cardiac-specific MCUB gain-of-function transgenic model
To generate a MCUB conditional, gain-of-function mutant mouse for in vivo experimentation we employed a flox-stop strategy where mouse Mcub cDNA
56 (CCDS 17839.1) was cloned into a custom CAG-loxP-CAT-loxP plasmid. The linearized sequence (Pac1 digest) was injected into 1-cell B6C3F1 embryos and transplanted into pseudopregnant female mice (C57BL6N). We characterized 10 founder lines for both leakiness (basal expression void of Cre) and level of expression following Cre-mediated excision of the CAT stop site by crossing to the cardiomyocyte-specific -MHC-Cre model. Western blot analysis of cardiac lysates enabled selection of a mutant line for study. All reported experiments utilized a transgenic mouse line with 15-fold overexpression of monomeric MCUB compared to MCM control. For all experiments, littermate controls were utilized.
To conditionally overexpress MCUB in a cardiac-restricted manner MCUB-Tg mice were crossed to the α-MHC MerCreMer mouse model(Sohal et al., 2001).
Tamoxifen was injected i.p., 25 mg/kg/d for four consecutive days to induce
MCUB expression in cardiomyocytes.
Size exclusion chromatographic analysis of the high molecular weight mtCU
complex
All procedures were carried out as previously reported(Tomar et al., 2016). 2,500
grams of whole cell protein lysates or pure cardiac mitochondrial protein lysates prepared in RIPA lysis buffer were fractionated by gel filtration using fast protein liquid chromatography (AKTA Pure FPLC; GE Healthcare, using a Superdex 200
10/300 column that was equilibrated with PBS) at a flow rate of 0.5 ml/min.
Fractions were collected in 0.5 mLs of PBS and concentrated to 75 ls using the
AMICON Ultra-0.5 Centrifugal Filter Devices (with a 3,000 kD cutoff) according to
57 the manufacturers instructions. 20 ls of concentrated fractions were then loaded and run on NuPAGE NOVEX Bis-Tris Midi Gels under reducing conditions and immunoblots were performed to analyze members of the mtCU by normal western blot techniques as previously reported(Elrod et al., 2010).
qPCR mRNA analysis
All procedures were carried out as previously reported(Elrod et al., 2010; Luongo et al., 2017). mRNA was isolated using the Qiagen RNeasy Kit. qPCR analysis was conducted according to the manufacturer’s instructions (Maxima SYBR,
Thermo Scientific). We evaluated samples for mRNA expression of Mcub and
Mcu. Rps13 was used as a housekeeping gene. All experiments were analyzed in triplicate and averaged. The 2-ΔΔct was used to calculate fold change in gene expression.
Western Blot Analysis
All procedures were carried out as previously reported(Elrod et al., 2010). Briefly all protein samples were lysed by homogenization in RIPA buffer. Samples were run by electrophoresis on polyacrylamide Tris-glycine SDS gels. The only exception to this case was for all samples collected by size exclusion chromatography. These protein lysates were run on NuPAGE NOVEX Bis-Tris
Midi Gels.
58
Table 3-1. Antibody Information
Antibody Company Product Species Primary Secondary Number Ab Conc. Ab Conc. EMRE Santa Cruz sc-86337 Rabbit 1 to 250 1:12,000 FLAG Sigma F1804 Mouse 1:2,000 1:12,000 HA Roche 11867423001 Rat 1:1,000 1:12,000 MCU Cell 149975 Rabbit 1:1,000 1:12,000 Signaling MCUB Santa Cruz sc-163985 Goat 1 to 250 1:12,000 MCUR1 Custom Rabbit 1 to 500 1:12,000 MICU1 Custom Rabbit 1:1,000 1:12,000 MICU2 Abcam 101465 Rabbit 1:1,000 1:12,000 OxPhos Mitosciences MS604 Mouse 1:5,000 1:12,000 PDHE1α Abcam ab110330 Mouse 1:1,000 1:12,000 P-PHDE1α Abcam ab92696 Rabbit 1:1,000 1:12,000 Donkey anti Li-Cor 926-32214 1:12,000 Goat Goat anti Li-Cor 925-68076 1:12,000 Rat Goat anti Li-Cor 926-68071 1:12,000 Rabbit Goat anti Li-Cor 926-32210 1:12,000 Mouse
Echocardiography
Transthoracic echocardiography of the left ventricle was performed and analyzed on a Vevo 2100 imaging system (VisualSonics) as previously reported(Luongo et al., 2017). Mice were anaesthetized with 2% isoflurane in 100% oxygen during acquisition. B-mode and M-mode images were collected in both long- and short- axis views. Dimensions of M-Mode images of short axis views were analyzed at mid-ventricular sections for accurate assessment of LV structure and function.
59 Isolation of ACMs
ACMs were isolated from ventricular tissue as described previously(Zhou et al.,
2000). All cells were used within 4 h of isolation.
2+ 2+ Evaluation of mCa Uptake, Content, and Ca retention capacity experiments
All procedures were carried out as previously reported(Luongo et al., 2017). To
2+ measure mCa uptake in HeLa cells 4 x 10^6 cells were used per experiment. To
2+ measure mCa uptake in ACMs 300,000 cells were used per experiment. In brief cells were transferred to intracellular-like medium (ICM) containing: 120 mM KCl,
10 mM NaCl, 1 mM KH2PO4, 20 mM HEPES-Tris, 3 mM thapsigargin, (40 μ g ml−1 digitonin for HeLa, or 80 μ g ml−1 digitonin for ACMs), protease inhibitors
(Sigma EGTA-Free Cocktail), 10 μM succinate, at pH 7.2. Cells were gently stirred and 1 μM Fura-FF (Invitrogen) was added to monitor extra-mitochondrial
Ca2+. At 20 s JC-1 (Enzo Life Sciences) was added to monitor Δψ. Fluorescence signals were monitored at 490 nm excitation/535 nm emission for the monomer,
570 nm excitation/595 nm emission for the J-aggregate of JC-1, and 340 and 380 nm excitation/510 nm emission for Fura-FF to calculate ratiometric changes. At
350 s boluses of Ca2+ ranging from 0-10 μM Ca2+ bolus were added to HeLa cells while a 5 μM Ca2+ bolus was added in experiments with ACMs. Clearance
2+ 2+ of extra-mitochondrial Ca was representative of mCa uptake. At completion of the experiment the protonophore, FCCP, was added. All experiments were conducted at 37°C and recorded on a PTI spectrofluorometer. For Fig 3-1J the
2+ 2+ 2+ mCa uptake rate was recorded over 50s post Ca bolus, plotted versus Ca
60 bolus given, and data was fitted with one-site, specific binding with Hill slope using Prism 6 software.
2+ To evaluate mCa content, permeabilized HeLa cells in ICM were treated
2+ with RU360 and CGP-37157 to inhibit mCa flux. Fura-2 (Invitrogen) was added for ratiometric monitoring of extra-mitochondrial Ca2+ using a fluorescent spectrofluorometer. FCCP was added to uncouple the and liberate matrix
Ca2+. Fura-2 signal was calibrated to known Ca2+ standards to obtain actual
2+ 2+ matrix [Ca ]. To perform mCa retention capacity (mCRC) experiments 4 x 10^6
HeLa cells were permeabilized in ICM and underwent simultaneous ratiometric monitoring of bath Ca2+ levels (Fura-FF) and ψ (JC-1) as cells were exposed to consecutive 10 μM boluses of Ca2+ until ψ collapsed indicating membrane permeability transition. Total amount of boluses cells could take up before ψ
2+ collapsed was recorded as the mCa retention capacity (mCRC).
Mitochondria Isolation and Swelling Assay
Hearts were excised from mice and mitochondria were isolated as reported(Frezza, Cipolat, & Scorrano, 2007). For the swelling assay, mitochondria were diluted in assay buffer, and absorbance (abs) was recorded at
540 nm every 5 s. 500 M CaCl2 was injected to induce swelling(Elrod et al.,
2010).
61 2+ 2+ Adult mouse cardiomyocyte iCa and mCa transient recordings
Isolated ACMs were loaded with 1 μM Fluo-4 AM (Invitrogen) and placed in a 37
°C heated chamber on an inverted microscope stage. Cardiomyocytes were perfused with a physiological Tyrode’s buffer (150 mM NaCl, 5.4 mM KCl, 1.2 mM MgCl2, 10 mM glucose, 2 mM sodium pyruvate, and 5 mM HEPES, pH 7.4)
2+ 2+ containing 2 mM Ca . Cells were paced at 1 Hz and iCa transients were continuously recorded and analyzed using Clampex10 software (Molecular
Devices). After two minutes of baseline recording cardiomyocytes were spiked with a 100 nM isoproterenol (Sigma-Aldrich) bolus. For intracellular Ca2+ fluorescence measurements, the F0 was measured as the average fluorescence of the cell before stimulation. The maximal Fluo-4 fluorescence (F) was
2+ measured for peak amplitude. To monitor mCa transients, AAV9-mitycam (1 x
1012 particles) was injected intravenously (via retro-orbital) into mice. After 2–3
2+ weeks, ACMs were isolated and imaged the same day to monitor mCa flux during pacing (0.1 Hz). Background fluorescence was subtracted from each experiment before quantification.
Mitoplast Patch-Clamp Analysis of MCU Current
Following mitochondrial isolation, mitoplasts were prepared for patch-clamp studies. IMCU was recorded as previously described in detail(Yuriy Kirichok et al.,
2004).
62 Oxygen Consumption Assays
A Seahorse Bioscience XF96 extracellular flux analyzer was employed to measure adult cardiomyocyte oxygen consumption rates (OCR). 5,000 cardiomyocytes per well were plated in XF media pH 7.4 supplemented with 0.2 mM palmitate, 0.2 mM carnitine, and 4 mM glutamine. Basal OCR was measured, then 3 μM oligomycin was injected to inhibit ATP-linked respiration, followed by 1.5 μM FCCP to measure maximal respiration, and finally to completely inhibit all mitochondrial respiration, 2 μM rotenone/antimycin-A was injected. Detailed methodology has been previously reported(Luongo et al.,
2017).
Invasive hemodynamic measurements
Mice were anaesthetized (avertin, 25 mg kg−1) and a cervical incision was performed to isolate the right carotid artery for insertion of a 1.4-F pressure catheter (SPR-671, Millar Instruments). The pressure transducer was advanced into the left ventricle before recording. All data was analyzed using Chart 6.0 software. All details are previously described(Luongo et al., 2015).
Permanent myocardial infarction and myocardial IR Injury
For assessment of MCUB expression at the mRNA level following permanent MI and at the protein level following IR injury male C57BL/6N at 12 weeks of age were used. For evaluation of infarct size following IR injury in our transgenic mouse models male and female littermate controls were utilized. LCA ligation
63 and reperfusion was performed as previously described in(Gao et al., 2010).
Briefly, mice were anaesthetized with isoflurane and the heart was exposed via a left thoracotomy of the fifth intercostal space. For permanent myocardial infarction the left coronary artery (LCA) was permanently ligated to induce a large myocardial infarction. Alternatively, for ischemia reperfusion a slipknot was tied around the LCA to allow for reperfusion. The heart was then returned to the chest cavity and the wound was sutured revealing the slipknot. After 40 minutes of ischemia, the slipknot was released and the ischemic zone was allowed to reperfuse for 24 hours. To assess infarct size, after LCA religation, hearts were injected with 3% Evans blue dye to delineate viable tissue from area at risk, rapidly excised and cross sectioned into 1-mm heart sections were incubated with 1% triphenyl tetrazolium chloride for 5 min at 37°C to delineate infarcted tissue. Each of the first five 1-mm thick myocardial slices were weighed, had pictures taken, and the areas of infarct, at risk, and viable tissue were assessed using ImageJ to perform planimetric analysis on one side of each heart slice as previously reported(Elrod et al., 2007).
2+ 2+ cCa and mCa flux in HeLa cells
Cells were transduced with AAV6-mito-R-GECO1 for 72 hours and live cell
2+ imaging with a Zeiss microscope was performed to measure mCa exchange
2+ following histamine stimulation (100 μM). Cells were also loaded with the iCa
2+ indicator, Fluo4-AM, and live cell imaging was performed to measure cCa
64 cycling following histamine stimulation. Data were collected every 0.5 s and fluorescence was analyzed using Zen software.
Immunoprecipitations
Plasmids encoding MCU, MCUB, MICU1, and MICU2 tagged with HA or FLAG were generated. To confirm interaction 5 μgrams of plasmids were co-transfected
(using Invitrogen Lipofectamine 3000 transfection kit) into COS7 cells and allowed to express for 48 hours. Cells were lysed in 1X RIPA (with SIGMAFAST protease inhibitor (PI) cocktail tablets) by passing through a 27-gauge needle 5 times and allowed to sit on ice for 30 minutes before being centrifuged at 5,000 x g. Supernatant was collected. 50 μls was saved as input while the remaining sample was used for immunoprecipitation after preclearing samples with Santa
Cruz Protein A/G PLUS-Agarose beads for one hour at 4C. Samples were then spun at 1,000 x g and the supernatant was collected. Sigma Anti-HA High Affinity rat monoclonal antibody or Sigma Anti-FLAG monoclonal mouse antibody was then added to samples at a final concentration of 5 μgrams/mL and rotated at 4C overnight. 50 μls of Santa Cruz Protein A/G PLUS-Agarose beads was added to the immunoprecipitation samples which were rotated for four hours at 4C.
Samples were then washed 3 times in 1X RIPA plus PI followed by two washes in TBS-Tween 20. Protein was then collected in 2x SDS lysis buffer by heating at
95°C for 10 min. 20 percent of Input was loaded onto gels while 20 μls of IP samples was loaded. HA or FLAG antibody was then used to assess protein interaction. MCM and MCUB-Tg x MCM cardiac protein lysates were
65 immunoprecipitated with 50 μls of Santa Cruz Protein A/G PLUS-Agarose beads conjugated to Santa Cruz normal goat IgG (sc-2028) as controls or MCUB antibody (Santa Cruz CCDC109B (S-15) sc-163985). See exogenous immunoprecipitations for subsequent steps. MCUB was immunoblotted for to confirm pulldown of MCUB. MCU and MICU1 were immunoblotted for to evaluate protein interaction with MCUB.
Statistics
All results are presented as mean ± SEM. Statistical analysis was performed using Prism 6.0 software (GraphPad). When column analyses were performed for statistics between two groups an unpaired, two-tailed t test was used. For groups of three or more a one-way ANOVA with Bonferroni correction was used.
For grouped analyses, either multiple unpaired t-test with correction for multiple comparisons using the Holm-Sidak method or two-way ANOVA with Tukey post- hoc analysis was performed. For all echocardiographic analysis two-way repeated-measures ANOVA was used. For analysis of survival curves using the
Kaplan-Meier method the log-rank statistical method was used.
Results
2+ MCUB deletion alters mtCU formation, stoichiometry, and mCa uptake
To evaluate how MCUB contributes to mtCU function we generated a stable
MCUB knockout (MCUB-/-) HeLa cell line using CRISPR/Cas9n(Ran et al., 2013).
We targeted exon 1 by employing a double nickase strategy to avoid off-target
66 genomic editing (Fig 3-1A)(Ran et al., 2013). Following puromycin selection, clonal MCUB-/- cell lines were examined for loss of mRNA by qPCR and protein by Western blot analysis. Two clonal lines were found to be void of MCUB mRNA; line 2 was used in all subsequent experiments (Fig 3-2A). mRNA expression of mtCU subunits indicated that the ratio of MCU:MCUB in WT HeLa cells is ~3:1 (Fig 3-2B). Deletion of MCUB resulted in an increase in the expression of core mtCU subunits. Western blot analysis revealed that MCU expression increased ~2.7-fold and EMRE expression increased ~4.5-fold, as compared to wild-type (WT) controls (Fig 3-1B). Complex V (CV-S) of the ETC served as a mitochondrial loading control. We next investigated if loss of MCUB altered the composition of the mtCU complex. Protein lysates from WT and
MCUB-/- cells were fractionated by size-exclusion FPLC (Fig 3-1 C-E). FPLC fractions were calibrated to known MW standards (Fig 3-2D, 3-2E) and fractions
4-12 (corresponding to ~900-200 kD MW fractions) were collected, concentrated, and examined by Western blot under reducing conditions. MCUB deletion increased the presence of MCU in the mtCU by ~2.5-fold with significantly more expression in high-MW fractions 5-9 (Fig 3-1C). In addition, the channel regulators MICU1 and MICU2 displayed increased expression in the high-MW complex in MCUB-/- cells (Fig 3-1D, 3-1E). These results suggest that MCUB expression regulates overall mtCU formation and stoichiometry.
Given the dynamic changes in mtCU formation, we next investigated how
2+ MCUB expression impacts uniporter function. We evaluated mCa dynamics by transducing wild-type (WT) and MCUB-/- cells with adeno-associated virus (AAV)
67 encoding mito-R-GECO1, a genetic calcium reporter targeted to the mitochondrial matrix, and monitored changes in fluorescence by live-cell imaging during histamine simulation, which mediates an Inositol trisphosphate receptor
2+ 2+ (IP3R) increase in cytosolic Ca (cCa ) (Fig 3-1F). Loss of MCUB increased the
2+ -/- 2+ mCa amplitude by ~80%, suggesting MCUB cells display increased mCa
2+ 2+ uptake (Fig 3-1G). To examine if alterations in mCa uptake impacted cCa signaling, cells were loaded with the reporter, Fluo4-AM, and live-cell imaging was performed during histamine treatment (Fig 3-1H). MCUB deletion decreased
2+ the cCa transient amplitude by ~35%, suggestive of enhanced mitochondrial buffering (Fig 3-1I).
We next evaluated if MCUB altered gating of the mtCU. WT and MCUB-/- cells were permeabilized; ER Ca2+ uptake inhibited with thapsigargin, and loaded with the ratiometric reporters Fura-FF and JC-1 to simultaneously record
2+ 2+ changes in Ca and mitochondrial membrane potential (). mCa uptake was monitored during incremental changes in bath Ca2+ [0.5 - 10 M] (Fig 3-2F). The
2+ 2+ mCa uptake rate was quantified for 200s following each bolus of bath Ca and exponentially fit to the Hill equation (Fig 3-1J). MCUB-/- cells exhibited increased
2+ 2+ mCa uptake rates across a wide-range of Ca concentration, with differences beginning at [Ca2+ > 3 µM]. Next, to examine if MCUB altered overall channel capacitance we patch-clamped whole mitoplasts, mitochondria stripped of their outer membrane, and monitored current during a voltage ramping protocol (-160 mV to 80 mV) in the presence of Ca2+ (5 mM) (Fig 3-1K). Loss of MCUB significantly increased the current density (Fig 3-1L) suggesting an increase in
68 overall functional mtCU. To see if the increase in current density and uptake-rate at high-Ca2+ impacted overall matrix content of free-Ca2+ we evaluated the
2+ release of mCa induced by FCCP (Fig 3-1M). Loss of MCUB did not significantly alter the basal matrix Ca2+ content (Fig 3-2C and Fig 3-1N). We then
2+ performed mCa retention capacity (mCRC) experiments in our permeabilized cell system during repetitive boluses of 10 M Ca2+ until collapse of (Fig 3-
1O, 3-1P). MCUB-/- cells were able to withstand ~30% less Ca2+ load before membrane potential collapse indicating a decrease in mCRC and increase in susceptibility to permeability transition (Fig 3-1Q). These data suggest that
MCUB deletion enhances overall mtCU formation and/or increases cooperative activation and uptake-rate in high-Ca2+ conditions, both of which may predispose
2+ cells to mCa overload.
MCUB incorporates into the mtCU complex after injury
2+ Aberrant iCa cycling is a key feature and contributor to the development and progression of cardiac injury(Bers, Despa, & Bossuyt, 2006). We have previously
2+ shown that mCa overload contributes to IR-injury and heart failure progression(Luongo et al., 2017; Luongo et al., 2015). To discern changes in mtCU component expression we evaluated left ventricle mRNA expression 2d and 14d after permanent ligation of the left coronary artery (LCA) which induces a large myocardial infarction (MI). We found that MCUB expression was significantly increased 48h following MI, resulting in ~3-fold increase in
MCUB/MCU expression (Fig 3-3A). Next, we evaluated the high-MW mtCU for
69 changes in MCUB expression following IR-injury. After IR mitochondria were isolated from ventricular tissue and fractionated by size-exclusion chromatography. Fractions ranging from ~200-900 kD were run under reducing conditions and immunoblotted for MCUB expression. Interestingly, MCUB was completely absent from the high-MW fractions of sham operated hearts; while
MCUB was easily detected in the ~800-600 kD fractions following IR-injury (Fig
3-3B).
Generation of a cardiac-restricted MCUB overexpression mouse model
To examine the physiological relevance of MCUB expression and incorporation into the mtCU in cardiac physiology and disease we generated a gain-of-function mouse model employing a flox-stop strategy where MCUB cDNA was cloned into a construct containing a CAG promoter followed by a floxed chloramphenicol acetyltransferase (CAT) to allow for Cre-driven deletion of CAT and expression of
MCUB (Fig 3-3C). We characterized 10 founder lines for both leakiness (basal expression void of Cre) and level of expression following Cre-mediated activation by crossing founder lines to the cardiomyocyte-specific MHC-Cre model (Fig 3-
4A). After selection of a founder with tightly regulated expression we crossbred our MCUB transgenic mouse to the MHC Mer-Cre-Mer (MCM) driver line to enable conditional expression of MCUB in the adult heart (Fig 3-3C). Tamoxifen
(tamox) was administered i.p. (25 mg/kg/d for 4d) to induce MCUB expression. 1 day following tamox treatment cardiac protein lysate was obtained to evaluate
MCUB protein expression by western blot analysis. We confirmed a significant
70 increase in MCUB overexpression, which resulted in a ~15-fold increase in the
MCUB:MCU ratio with no baseline change in the expression of other mtCU subunits (Fig 3-3D, 3-3F). Next, adult cardiomyocytes (ACMs) were isolated from
2+ MCUB-Tg x MCM and controls and mCa uptake was examined independent of intracellular calcium handling with simultaneous monitoring of mitochondrial membrane potential (Fig 3-3G). Overexpression of MCUB reduced the rate of
2+ 2+ mCa uptake by ~70% and similarly decreased the amount of Ca taken up following a 5 µM bath-Ca2+ bolus (Fig 3-3H, 3-3I). To examine Ca2+ flux in intact cardiomyocytes we injected mice with AAV encoding the mitochondrial-targeted reporter, mitycam. ACMs were isolated and live-cell imaging was performed
2+ during pacing at 0.1 Hz to resolve individual mCa transients (Fig 3-3J). MCUB-
Tg x MCM cardiomyocytes displayed a ~28% reduction in the peak amplitude of
2+ mCa transients during pacing (Fig 3-3K). The rate-of-decay, (Fig 3-4B) was not
2+ statistically different. To assess cytosolic calcium (CCa ) transients, ACMs were isolated and loaded with the Ca2+ reporter, Fluo-4, and paced at 1 Hz (Fig 3-3L,
3-3M) (+/-) 100 nM isoproterenol. At baseline there was no significant difference
2+ in the peak-amplitude of CCa transients between groups, but we did note a
2+ slight ~20% increase in the cCa transient amplitude of isoproterenol-treated
MCUB-Tg x MCM cardiomyocytes (Fig 3-3N). The rate-of-decay, (Fig 3-4C) was not statistically different.
71 2+ Overexpression of MCUB reduces mCa -overload and cardiac IR Injury
2+ To test how decreased mCa uptake resulting from MCUB gain-of-function impacted cardiac physiology function was evaluated by echocardiography 1wk and 1m after transgene induction (post-tamox treatment). 1 wk after MCUB expression fractional shortening (FS) was reduced by ~40% (Fig 3-5A) and LV end-diastolic and LV end-systolic diameters (LVEDD, LVESD) were significantly increased (Fig 3-6B, Fig 3-6C). 1m post-tamox treatment FS, LVEDD, and
LVESD of MCUB-Tg x MCM mice returned to MCM control levels indicating a transient decrease in cardiac function following MCUB overexpression. At baseline, 1w, and 1m post-tamox treatment the heart rate and LV wall thickness was not different between groups (Fig 3-6 D-G). IR results in pathogenic
2+ increases in iCa levels and we have previously reported that MCU-mediated
2+ mCa overload is a significant contributor to cardiac injury via the induction of the mitochondrial permeability transition pore. Given the transient cardiac dysfunction noted with MCUB overexpression we employed two protocols to examine how MCUB modulation of the mtCU impacted cardiac injury. LCA ligation (40m ischemia and 24h reperfusion) was performed 1wk (Protocol 1) and
1m (Protocol 2) after tamox treatment (Fig 3-5B). Surprisingly, myocardial IR performed 1wk post-tamox resulted in substantial mortality of MCUB-Tg x MCM mice. 12/13 mice overexpressing MCUB died within a few minutes of ischemia
(Fig 3-5C). In stark contrast, IR induced 1m after tamox resulted in no statistical difference in survival between MCM and MCUB-Tg x MCM (Fig 3-6J). In fact,
MCUB overexpression reduced infarct size per area-at-risk (AAR) by ~50%, as
72 compared to MCM controls (Fig 3-5D, 3-5E, Fig 3-6K). No statistical difference in AAR/LV was observed. We have previously reported that inhibition of mtCU- uptake is protective against IR-injury by lessening activation of the mitochondrial permeability transition pore (mPTP)(Luongo et al., 2015). Therefore, we examined swelling (an indicator of mPTP activity) in ventricular mitochondria isolated from MCUB-Tg x Cre and control mice. Absorbance was monitored during the delivery of 500 M Ca2+ bolus (Fig 3-5F). Increased expression of
MCUB reduced both the rate and extent of mitochondrial swelling induced by
Ca2+ overload (Fig 3-5G, 3-5H). Given the divergent post-IR phenotype of MCUB overexpression (acute expression resulted in lethality during ischemia while 1m post-gene induction was cardioprotective) we hypothesize that acute expression of MCUB impaired mitochondrial energetics such that ATP supply was insufficient during the high-stress/demand of IR-injury and that sufficient compensatory mechanisms are activated after a longer period (1m) of MCUB expression to either lessen the inhibitory effect on the mtCU or allow for the
2+ upregulation of compensatory energetic pathways such that lessening mCa overload and mPTP cell death is now more favorable.
MCUB expression impairs mitochondrial energetics and contractility during stress
Since acute MCUB expression resulted in a transient decrease in cardiac function 1w post-tamox we employed invasive hemodynamics to monitor LV function during iso infusion 1d after tamox administration (Fig 3-7A). Acute
MCUB expression slightly depressed heart rate at a single dose of iso (Fig 3-
73 7B). Overexpression of MCUB completely ablated iso-induced elevations in
2+ contractility (dP/dtmax, Fig 3-7C) and relaxation (dP/dtmin,Fig 3-7D). mCa signaling modulates energetics by modulating Ca2+-dependent dehydrogenases, as well as direct action on electron transport chain (ETC) components(Denton,
McCormack, & Edgell, 1980) (Denton, Randle, & Martin, 1972; Glancy &
Balaban, 2012). We have previously reported that mtCU-Ca2+ uptake promotes mitochondrial energetics and is necessary for increased contractility during adrenergic stimulation(Luongo et al., 2015). To examine if mitochondrial energetics were altered with MCUB expression we isolated ACMs from MCM and
MCUB-Tg x MCM mice and examined oxidative phosphorylation by monitoring oxygen consumption rates (OCR) (Fig 3-7E). MCUB expression significantly decreased maximal respiration and spare respiratory capacity, yet had no effect on basal respiration, ATP-linked respiration, or proton leak (Fig 3-7F). Next, we performed Western blots to assess phosphorylation at S293 of pyruvate dehydrogenase (p-PDH), the inhibitory phosphorylation site that is
2+ dephosphorylated by the mCa -dependent pyruvate dehydrogenase phosphatase. Densitometry revealed that MCUB gain-of-function increased baseline p-PDH ~2.3 fold (corrected to total PDH-E1), as compared to ventricular samples isolated from MCM control hearts (Fig 3-7G). There was no difference in total PDH-E1 between groups (Fig 3-8A). We also evaluated p-
PDH in cardiac lysates from hearts subjected to iso infusion and found that
MCUB expression significantly blocked dephosphorylation indicating an inability to increase PDH activity (Fig 3-7H). There was no difference in total PDH-E1
74 2+ between groups (Fig 3-8B). These results suggest that MCUB inhibition of mCa uptake is a powerful mechanism to downregulate mitochondrial bioenergetics by inactivating PDH and diminishing oxidative capacity during stress and that this mechanism may be sufficient to impair contractile reserve. Surprisingly, the effect of MCUB on LV function was even greater than what we reported with cardiomyocyte-specific deletion of Mcu(Luongo et al., 2015). We hypothesize that this may be largely dependent upon the temporal inhibition of mtCU flux as
Cre-activation of transgene expression is rapid in this model (<24h), whereas deletion of Mcu occurs over 4d due to the protein turnover rate. This thinking would also fit with other reports where a mild-phenotype was observed with germline deletion of Mcu vs. acute adult deletion and suggests alternative
2+ energetic pathways are quickly activated in response to impaired mCa uptake.
MCUB expression displaces MCU-subunits in the mtCU
(high-MW complex) to modulate MICU1/2 gating
To determine the mechanism whereby MCUB regulates mtCU function we examined the high-MW channel (~700-800 kD) in mitochondria isolated from the hearts of our mutant MCUB expression model. MCM and MCUB-Tg x MCM were injected with tamox for four consecutive days and on day five mitochondria were isolated and proteins were fractionated by size exclusion chromatography.
Fractions ranging from ~200-900 kD were run under reducing conditions and immunoblotted to evaluate the molecular composition of the mtCU (Fig 3-9 A-D).
We first examined MCUB expression and noted that MCUB was absent from the
75 mtCU in control hearts. In contrast, mitochondrial protein lysates isolated from our overexpression model (MCUB-Tg x MCM) displayed clear expression of
MCUB in the high-MW fractions (Fig 3-9A). Next, we immunoblotted for the pore- component, MCU, and found that MCUB gain-of-function decreased the overall size of the mtCU, indicated by a rightward shift in the chromatograph of MCU expression (Fig 3-9B). To ascertain if MCUB expression impacted the association of other mtCU components we also examined the mtCU regulators
MICU1 and MICU2. Incorporation of MCUB into the mtCU markedly reduced both MICU1 and MICU2 levels in the high-MW complex (Fig 3-9C, 3-9D). Given the impact of MCUB expression on MICU1/2 association with the mtCU we next sought to determine if MICUs interact with MCUB. Employing c-terminal tags we co-expressed MCUB-HA with MCU-FLAG, MICU1-FLAG, or MICU2-FLAG. 48h after transfection we immunoprecipitated (IP’d) under stringent conditions with an
HA antibody (MCUB) and immunoblotted for FLAG (Fig 3-9E). MCUB was found to interact/bind MCU, but pull-downs with MCUB resulted in no MICU1 or MICU2 immunoreactivity. MCUB expression was similar among all experimental groups
(IB:HA, Fig 3-9E). To validate this result we next performed reverse IPs and co- expressed MICU1-FLAG or MICU2-FLAG (Fig 3-9F, 3-9G) with MCU-HA or
MCUB-HA. IPs of MICU1 and MICU2 both showed clear interaction with MCU, but no binding of MCUB (Fig 3-9F, 3-9G). FLAG expression (MICU1-FLAG or
MICU2-FLAG) was similar in all groups (Fig 3-9F, 3-9G). We further confirmed this by performing endogenous immunoprecipitations for MCUB in MCUB-Tg x
MCM cardiac protein lysates (Fig 3-10A). Again we positively confirmed MCUB
76 interacted with MCU (Fig 3-10B) but did not interact with MICU1 (Fig 3-10C). In summary, these results suggest that MCUB replaces MCU in the mtCU and thereby indirectly modulates the association of MICUs with functional channel
(i.e. MICU1/2 show interaction with MCU but not MCUB and therefore are displaced when MCUB enters the high-MW complex). In totality our results suggest that the expression of MCUB is a mechanism to decrease mtCU uptake by modulating the molecular composition of the mtCU.
Discussion
Here we identified a new mechanism whereby the mtCU is regulated. We discovered that MCUB is a powerful regulator of mitochondrial calcium uptake by modulating the incorporation of channel subunits and therein modifies the stoichiometry and function of the uniporter. In addition, our study revealed that
MCUB is upregulated following acute cardiac injury and that this is likely a compensatory mechanism to lessen mitochondrial calcium overload during ischemic stress. However, we also show that increased MCUB incorporation into the mtCU does come at a cost, as MCUB-mediated downregulation of mtCU function impairs mitochondrial energetics during stress.
Raffaello et al. discovered the MCU paralog, MCUB, and in agreement
2+ with our study showed that siRNA silencing of MCUB increased mCa transients.
The Rizzuto led group also provided data that expressing MCUB alone in a lipid bilayer alone did not result in a channel with Ca2+ conductance(Raffaello et al.,
2013), in contrast to what was reported for the pore-forming subunit,
77 MCU(Chaudhuri, Sancak, Mootha, & Clapham, 2013; Diego De Stefani,
Raffaello, Teardo, Szabò, & Rizzuto, 2011). This led to the conclusion that this paralog exists as a dominant negative regulator of channel function and perhaps primarily impacted calcium permeation or pore function. While this founding discovery presented strong evidence that MCUB is a component of the mtCU many questions remained. How does MCUB regulate the mtCU? Is it a physiological regulator of mitochondrial calcium uptake? How does its association with the uniporter impact cellular function?
Employing size-exclusion chromatography we were able to examine how mtCU stoichiometry changed with loss or overexpression of MCUB. We deleted
MCUB in HeLa cells because of their relatively high level of MCUB expression
(~3:1, MCU:MCUB) and vast historical data with regards to the molecular regulation of the mtCU to allow comparisons with previously published studies.
Loss of MCUB enriched the levels of MCU, MICU1, and MICU2 in the high-MW mtCU (the physiological/functional uniporter). Conversely, cardiomyocyte-specific overexpression of MCUB in the heart decreased the overall size of the mtCU
(rightward shift in the MCU chromatogram) and decreased both MICU1 and
MICU2 association with the high-MW complex. These data suggest that MCUB may regulate the uniporter by altering the subunits that are present in the functional complex and suggests an alternative mechanism of regulation differing to, or beyond, altering the structure of the pore. Since we found that MCUB was not present in the high-MW complex at baseline but was incorporated into the mtCU after cardiac injury it suggests that this is a stress responsive regulatory
78 2+ mechanism in the heart to limit mCa overload. It is possible that this may also be a mechanism to mediate differences in uptake among differing cell types as
MCUB expression is reported to be tissue dependent(Raffaello et al., 2013).
MCUB gain- or loss-of-function significantly altered the biophysical
2+ properties of mCa uptake mostly at higher cytosolic calcium load. Mitoplast patch clamping studies indicated increased channel capacitance with MCUB deletion, which we attribute to either an overall increase in mtCU density (# of uniporters per mitochondrion) or alteration in mtCU gating/activation. By incrementally changing the extra-mitochondrial Ca2+ load we could resolve an
2+ increase in mCa uptake that was most apparent at concentrations >3µM. Given these changes we hypothesize that increasing MCUB expression displaces MCU subunits from the mtCU and thus is indirectly altering the association of MICU1/2 with the uniporter. This rationale is supported by our data showing that MICU1 and MICU2 can bind MCU, but do not bind MCUB. This mechanism could explain the decrease in uptake at higher calcium levels, as MICU1/2 cooperative activation of mtCU function is lost with increased MCUB expression(Csordas et al., 2013; Paillard et al., 2017; Patron et al., 2014; Payne et al., 2017).
Acute increases in MCUB had a profound effect on cardiac physiology in our transgenic mouse. We found that mitochondrial bioenergetics were severely impaired immediately following the induction of MCUB expression. As we’ve previously reported, the likely mechanism driving this impairment was an inability to activate the calcium-dependent dehydrogenases, notably pyruvate dehydrogenase (PDH). This in turn severely limited oxidative capacity decreasing
79 both maximal respiration and reserve capacity. Further, this loss of energetic signaling manifested in LV functional studies as MCUB overexpressing mice failed to increase contractility or relaxation in response to -adrenergic stimulation. This lack of energetic responsiveness was most apparent when mice with acute overexpression of MCUB (within 1w of tamox administration) were subjected to myocardial IR-injury. We found that nearly all MCUB-Tg x MCM mice died within minutes of LCA ligation (ischemic period). This is in stark contrast to MCUB overexpressing mice that were allowed to rest for 1m after tamoxifen treatment. With this protocol we found that MCUB overexpression reduced infarct size by ~50% with a noted decrease in mitochondrial swelling, indicative of reduced mPTP activation. We’ve previously reported a nearly identical phenotype with deletion of MCU, bolstering the notion that mitochondrial-calcium overload is a major contributor to IR-induced cardiomyocyte death(Luongo et al., 2015). We suspect that these divergent phenotypes are due to compensatory changes that occur following increased
MCUB expression. A few possibilities exist that may account for this difference.
Compensatory changes such as an upregulation of cardioprotective signaling, activation of alternative energetic pathways, or changes in other mtCU components to lessen the impact of MCUB on uptake could all contribute. The later notion is supported by our data that MCU expression increases overtime in the MCUB overexpression model and thereby could decrease MCUB incorporation into the mtCU (competitive effect) (Fig 3-11 A-C). This is also supported by our data showing that LV function decreases acutely after MCUB
80 expression but rebounds to wild-type levels 1m after gene induction (Fig 3-11 D-
J). It should be noted that this model recapitulates conflicting reports in MCU knockout models, which have shown very differing outcomes with regards to IR- injury. Germline deletion of MCU resulted in no change in infarct size(Pan et al.,
2013) whereas conditional deletion in the adult reduced infarct size following IR- injury(Kwong et al., 2015; Luongo et al., 2015). Here we show that compensatory mechanisms can account for a very dramatic difference in phenotype in the same mutant mouse and suggest that carefully planned experiments are needed to avoid misinterpretation of results with regards to the genetic perturbation of mitochondrial calcium exchange.
In conclusion, we report a new regulatory mechanism that greatly impacts mtCU function and mitochondrial calcium uptake. Our results suggest that the stoichiometry of the subunits associated with uniporter is critical to its function and that MCUB is a prominent regulator of uniporter composition. These findings suggest that the evaluation of subunit incorporation into the mtCU is much more important than total protein expression and suggests that post-translational regulation of the mtCU is a physiological mechanism to modulate gating of the
2+ channel and fine-tune mCa uptake to control mitochondrial bioenergetics.
81
2+ Figure 3-1. MCUB deletion alters mtCU formation, stoichiometry, and mCa uptake. A) The Cas9n double-nickase was targeted to exon 1 of MCUB. Clonal lines were evaluated after puromycin selection. B) Western blots depicting expression of MCUB, MCU, and EMRE, in wild-type (WT) vs MCUB-/- cell lines. Complex-V Sα was used as a mitochondrial loading control; n=3 for both groups. C-E) Protein lysates isolated from WT and MCUB-/- cells were fractionated by size exclusion chromatography and fractions were run on a western blot under reducing conditions. mtCU subunits were immunoblotted and densitometry was performed to generate chromatographs of relative expression for each subunit in the high-MW mtCU: C) MCU D) MICU1 E) MICU2 F) WT and MCUB-/- cells were transduced with AAV6-mito-R-GECO1 and fluorescence was recorded to monitor 2+ 2+ mCa dynamics following histamine stimulation. G) Amplitude of mCa transients -/- (peak intensity – baseline (F/F0)-F0); n=32 cells for WT, n=33 cells for MCUB . 2+ H) CCa transients (Fluo4-AM) following histamine stimulation. I) Amplitude of 2+ cCa transients (peak intensity – baseline (F/F0)-F0); n=37 cells for WT, n=41 cells for MCUB-/-. J) Ratiometric monitoring of bath Ca2+ levels (Fura-FF) and ψ (JC-1) following permeabilization. Bath Ca2+ ranged from 0.5 – 7.5 μM; n=4 runs per Ca2+ bolus. K) Mitoplasts from WT and MCUB-/- cells were patch clamped to record MCU current (IMCU). L) Current density (picoamperes/picofarad); n=8 for WT, n=7 for MCUB-/-. M) Fura-2 recording of mitochondrial free-Ca2+ content in matrix. N) Fold change in mitochondrial free-Ca2+ content in MCUB-/- cells vs WT;
82 n=7 for WT, n=6 for MCUB-/-. O) Mitochondrial calcium retention capacity (mCRC) recording in WT and P) MCUB-/- cells during repetitive additions of 10- µM Ca2+ (black arrows) until membrane potential collapse; n=4 runs per group. Q) Percent change in mCRC, MCUB-/- vs. WT; n=4 per group. Calcium traces: solid line = mean, dashed line = SEM. All data are shown as mean +/- SEM. ****p<0.0001, *** p<0.001, *p<0.05.
83
Figure 3-2. mRNA expression in HeLa cells, Fura-2 calibration, size exclusion chromatography of known protein standards for generation of a 2+ standard curve of molecular weights, and mCa uptake experiments. A) mRNA expression of MCUB in WT cells and MCUB-/- clonal lines. MCUB-/- L2 was used for all experiments. B) mRNA expression of MCU, MCUB, and the MCUB:MCU ratio in HeLa cells. C) Fura-2 was calibrated by generating a standard curve of Ca2+ (0.01 μM- 75 μM) in experimental intracellular buffer to quantify actual Ca2+ levels in Figure 1 M-N. D) Chromatograph of the protein absorbance (at 280 nms) of standards of known molecular weight size as they were fractionated by size exclusion chromatography. Thyroglobulin (670 kD), γ- globulin (158 kD), ovalbumin (44 kD), myoglobulin (17 kD), and vitamin B12 (1.3 kD). E) Molecular weight of protein standards plotted against actual fraction each known protein standard eluted at and fit non-linearly (third order polynomial cubic) to generate to a standard curve to identify fractions containing molecular weight sizes corresponding to the high-MW mtCU in all FPLC experiments. F) Depicted is the ratiometric monitoring of bath Ca2+ levels (Fura-FF) following permeabilization of WT and MCUB-/- cells as they were exposed to Ca2+ boluses 2+ 2+ ranging from 1 to 10 μM. mCa uptake rates were calculated after the Ca bolus given which were utilized to plot data in Fig 1J.
84
Figure 3-3. MCUB incorporates into the mtCU complex after injury warranting the generation of a MCUB conditional mutant mouse model. A) Mcu, Mcub, and the Mcub/Mcu mRNA ratio in cardiac tissue of mice subjected to sham, 48 hr myocardial infarction (MI), and 2w MI; n=3-4 mice per group. B) Relative expression of MCUB in the high-MW mtCU from cardiac mitochondria of mice subjected to sham procedure or 40 min ischemia 24 h reperfusion (24 h post I/R); n=4 mice per group. C) MCUB conditional gain-of-function mice (MCUB-Tg) were generated using a flox-stop strategy. D) Immunoblots of mtCU components in cardiac lysates of MCM and MCUB-Tg x MCM. Complex IV- MTCO1 was used as a loading control. E) Fold change of indicated mtCU subunit vs MCM control after correcting to CIV-MTCO1; n=4 mice per group. F) MCUB/MCU cardiac protein ratio. G-I) Ratiometric monitoring of bath Ca2+ levels (Fura-FF) in permeabilized MCM and MCUB-Tg x MCM cardiomyocytes 2+ 2+ following a 5 μM Ca bolus; n=3 mice per group. H) Percent change in the mCa 2+ 2+ uptake rate. I) Percent change of mCa uptake. J) mCa transients (Mitycam) in 2+ cardiomyocytes paced at 0.1 Hz. K) Amplitude of mCa transients (1-F/F0); n=23 2+ cells for MCM, n=30 cells for MCUB-Tg x MCM. L-M) cCa transients (Fluo4- AM) in L) MCM and M) MCUB-Tg x MCM cardiomyocytes paced at 1 Hz without 2+ vehicle (Veh) or with Isoproterenol. N) Amplitude of cCa transients (F/F0); n=51 cells for MCM, n=56 cells for MCM plus iso, n=61 cells for MCUB-Tg x MCM, n=57 cells for MCUB-Tg x MCM plus iso. Data are shown as mean +/- SEM. ****p<0.0001, **p<0.01, *p<0.05.
85
Figure 3-4. Assessing leakiness of the MCUB transgenic mouse, 2+ 2+ exponential decay rates for mCa transients and cCa transients. A) Protein cardiac lysates from -MHC-Cre, MCUB-Tg, and MCUB-Tg x Cre mice were immunoblotted for MCUB to confirm robust expression of the transgene. CV-Sα was used as the mitochondrial loading control; n=3 mice per 2+ group. B) Tau, the exponential decay rate of the mCa transients from Fig 3-3J. 2+ C) Tau, the exponential decay rate of the cCa transients from Fig 3-3L and 3- 3M. All data are shown as mean +/- SEM. ****p<0.0001.
86
2+ Figure 3-5. Overexpression of MCUB reduces mCa -overload and cardiac IR-injury. A) Fractional shortening (%FS) at baseline, 1w following tamoxifen (tamox) injections, and 1m following tamox injections; n=7 per group. B) Tamox injected mice were subjected to 40 min of ischemia and 24h of reperfusion 1w (Protocol 1) and 1m (Protocol 2) after MCUB induction and hearts were processed for Evan’s blue dye perfusion and TTC staining. C) Protocol 1: Kaplan-Meier survival curve of mice subjected to I/R injury 1w after tamox; n=13 mice per group. D-E) Protocol 2: D) Mid-ventricular cross-sections of hearts following Evan’s blue dye perfusion and TTC staining. All non-blue area indicates area-at-risk (AAR); red area, viable myocardium; white area, infarct (INF). E) Assessment of AAR per left ventricle (AAR/LV), INF/AAR, and INF/LV; n=12 mice for MCM, n=14 mice for MCUB-Tg x MCM. F) Mitochondrial swelling induced after 500 μM bath Ca2+, ventricular mitochondria isolated from α-MHC Cre and MCUB-Tg x α-MHC Cre hearts; n=5 per group. G) Rate of swelling, i.e. change in absorbance over 5 min. H) Percent change in swelling as area-over-the-curve. All data are shown as mean +/- SEM. ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05.
87
Figure 3-6. Echocardiographic assessment of parameters of cardiac function, Kaplan Meier survival curve of I/R injury 1 month post tamoxifen, and representative images of EBD/TTC staining in mouse models subjected to I/R injury 1 month post tamoxifen. A-G) MCM and MCUB-Tg x MCM mice were injected with tamoxifen for four days at 25 mg/kg/day and cardiac function was evaluated 1w and 1m later by echocardiography; n=7 mice for both groups. A) Heart rate, beats per minute. B) Left ventricular end-diastolic diameter, LVEDD. C) Left ventricular end-systolic diameter, LVESD. D) Interventricular septal end-diastolic diameter, IVS;d. E) Interventricular end-systolic diameter, IVS;s. F) Left ventricular posterior wall end-diastolic diameter, LVPW;d. G) Left ventricular posterior wall end-systolic diameter, LVPW;s. H) Left ventricular mass. I) Left ventricular mass corrected. J) Protocol 2: Kaplan-Meier survival curve of mice subjected to I/R injury 1m after tamox. K) Protocol 2: Ventricular cross-sections of hearts following Evan’s blue dye perfusion and TTC staining of mice subjected to I/R injury 1m after tamox. All data are shown as mean +/- SEM. ***p<0.001, *p<0.05.
88
Figure 3-7. MCUB expression impairs mitochondrial energetics and contractility during stress. A-D) Mice in both groups received tamoxifen (tamox; 25 mg/kg/day) for 4d and 1d later were subjected to an isoproterenol infusion (0.1-20 ng/ml); minimum n=6 per group. Invasive hemodynamic assessment of: B) heart rate (beats/m), C) dP/dt maximum, and D) dP/dt minimum. E) 1d post-tamox treatment adult cardiomyocytes were assayed for mitochondrial OxPhos function using a Seahorse bioanalyzer to measure the oxygen consumption rate (OCR). F) Maximal respiration, reserve capacity, ATP production, basal respiration, proton leak, and non-mitochondrial respiration represented as picomoles of O2/min/5k cells; n=3 mice per group. G) Immunoblots for pyruvate dehydrogenase phosphorylation at Serine293 (p-PDH), total Pyruvate Dehydrogenase E1-α (PDH- E1α), and the mitochondrial loading control Complex IV-MTCO1. Relative expression of p-PDH to PDH-E1α is depicted; n=3 per group. H) Cardiac protein lysates from post-iso stressed hearts were immunobloted for p-PDH, PDH-E1α, and Complex V-Sα; n=6 per group. Relative expression of P-PDH compared to total PDH-E1α is depicted. All data are shown as mean +/- SEM. ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05.
89
Figure 3-8. Expression levels of total pyruvate dehydrogenase E1α. A) Immunoblots in Fig 3-7G were analyzed by densitometry and the relative expression of total pyruvate dehydrogenase E1α corrected to the mitochondrial loading control CIV-MTCO1 is shown; n=3 mice per group. B) Immunoblots in Fig 3-7H were analyzed by densitometry analysis and here the relative expression of total pyruvate dehydrogenase E1α corrected to the mitochondrial loading control, CIV-MTCO1 of MCM and MCUB-Tg x MCM cardiac protein lysates of isoproterenol exposed hearts is shown; n=6 mice per group.
90
Figure 3-9. MCUB expression displaces MCU-subunits in the mtCU (high- MW complex) to modulate MICU1/2 gating. A-D) Mitochondrial cardiac protein lysates were fractionated by size exclusion chromatography and independent fractions were immunobloted for mtCU components. Densitometry analysis was performed and the chromatographs depict relative expression of the indicated mtCU subunit in the high-MW mtCU; n=3 mice per group. All fractions were fitted to the MW of known standards. A) MCUB B) MCU C) MICU1 D) MICU2 E) MCUB-HA was co-transfected with MCU-FLAG, MICU1-FLAG, or MICU2-FLAG into COS7 cells. HA- immunoprecipitation was performed and immunoblotted for FLAG and HA; n=3. F) MICU1-FLAG was co-transfected with MCU-HA or MCUB-HA into COS7 cells. FLAG immunoprecipitation was performed and Western blotted for HA and FLAG; n=3. G) MICU2-FLAG was co-transfected with MCU-HA or MCUB-HA into COS7 cells. FLAG immunoprecipitation was performed and probed for HA and FLAG immunoreactivity.
91
Figure 3-10. Endogenous immunoprecipitations confirm MCUB interacts with MCU and not MICU1. A-C) Cardiac protein lysates were obtained from MCM and MCUB-Tg x MCM mice and endogenous immunoprecipitations for MCUB were performed; n=2 for both groups. A) MCUB immunoblot. B) MCU immunoblot. C) MICU1 immunoblot.
92
Figure 3-11. Compensatory mechanisms take place 1 month following tamoxifen administration restoring cellular bioenergetics and reversing the cardiac phenotype. A-E) One month after tamoxifen injections protein was isolated from cardiac tissue of MCM and MCUB-Tg x MCM. MCUB and MCU were immunoblotted for, Complex IV-MTCO1 was used as a loading control; n=4 mice per group. B) Relative expression of MCUB. C) Relative expression of MCU. D) Immunoblots for pyruvate dehydrogenase phosphorylation at Serine293 (p-PDH), total Pyruvate Dehydrogenase E1-α (PDH-E1α), and the mitochondrial loading control Complex V-Sα. E) Relative expression of p-PDH to PDH-E1α is depicted; n=4 per group. F-G) 1mo post-tamox treatment adult cardiomyocytes were assayed for mitochondrial OxPhos function using a Seahorse bioanalyzer to measure the oxygen consumption rate (OCR). G) Maximal respiration, reserve capacity, ATP production, basal respiration, proton leak, and non-mitochondrial respiration represented as picomoles of O2/min/5k cells; n=3 mice per group. H-J) Mice in both groups received tamoxifen (tamox; 25 mg/kg/day) for 4d and 1mo later were subjected to an isoproterenol infusion (0.1-20 ng/ml); minimum n=6 per group. Invasive hemodynamic assessment of: H) heart rate (beats/m), I) dP/dt maximum, and J) dP/dt minimum. All data are shown as mean +/- SEM. **p<0.01, *p<0.05.
93 CHAPTER 4
CONCLUSIONS AND SIGNIFICANCE
Extended Discussion and Future Directions
2+ The work herein further supports the idea that mCa is a key regulator of both cellular bioenergetics and cell death but importantly suggests MCUB is a powerful way to regulate these cellular processes (Fig 4-1). This work provides evidence that MCUB alters the stoichiometry and composition of mtCU to modulate channel function and reports the cardiac pathophysiological role of this gene. For the first time we report that MCUB is absent from the mtCU of the homeostatic heart but upon I/R injury, during which intracellular Ca2+ levels are high and ROS is generated, MCUB is upregulated and incorporates into the mtCU as early as 24 hours following reperfusion. We believe MCUB’s upregulation is an acute compensatory mechanism to provide cardioprotection
2+ during I/R injury by limiting mCa uptake thus preventing mPTP induced necrosis and cardiomyocyte loss (Fig 4-2). We also provide evidence that MCUB could be utilized as a therapeutic for the treatment of I/R injury.
Paralog Interference
2+ Various cellular processes, such as mtCU mediated mCa uptake, are carried out by molecular machines consisting of multiple proteins that assemble into a macromolecular complex that executes specific biological functions but there is little evidence indicating how these molecular machines evolved. Proteins often form multimeric complexes with identical subunits which is what we observe in
94 the case of MCU in which the determined structure of fungul MCU’s is a tetramer in a dimer of dimers configuration(Fan et al., 2018; Yoo et al., 2018). Paralogs arise from gene duplication which occurs in eukaryotic organisms at the rate of
0.01 per gene per million years(Lynch & Conery, 2000). After duplication two gene paralogs from the same ancestor gene initially still form a homomeric complex through cross reaction but as random mutations are acquired overtime the genes diverge creating the opportunity for paralog interference. It is hypothesized that there are three outcomes of paralog interference; one copy becomes silenced due to non-functional mutations (nonfunctionalization); one copy gains a new function (neofunctionalization); or paralogs generate multifunctional proteins where differential functions are distributed to each paralog (subfunctionalization)(Soria, McGary, & Rokas, 2014). While it is unclear when MCU first duplicated and how its paralog, MCUB, genetically diverted, its existence necessitates an understanding of its influence within the mtCU and an inquiry into the physiological relevance of this gene. We can possibly gain insight into its function by understanding how paralogs of other multisubunit macromolecular complexes have evolved.
The evolution of the hexameric transmembrane ring of the eukaryotic vacuolar H+ -ATPase proton pump (V-ATPase) has more recently been elucidated. The V-ATPase is a channel that uses ATP hydrolysis to transport protons across the plasma membrane or across intracellular compartments of eukaryotic cells to create a proton gradient or to acidify intercellular compartments(Forgac, 2007). The VO protein ring forms hexamers and is the
95 rotary mechanism that moves protons across the membrane in a similar fashion to the FO ATP synthase(Marshansky, Rubinstein, & Grüber, 2014). In most eukaryotes two paralogs (Vma16 and Vma3) make up the hexameric VO ring while in Fungi three paralogs (Vma16, Vma3, and Vma11) make up the hexamer(Powell, Graham, & Stevens, 2000). Between these two species the VO rings of the V-ATPase are functionally no different but their hexameric assembly is. The functional VO ring in fungi contains one Vma16 subunit, 4 Vma3 subunits, and one Vma11 subunit which are in obligate positions in the hexameric ring.
The functional VO ring in eukaryotes contains one Vma16 subunit and 5 Vma3 subunits. What has happened evolutionarily is that the fungal Vma3 and Vma11 have both diverged from the eukaryotic Vma3 and the divergent mutations have impacted macromolecular assembly leading to different compositions of the hexameric VO ring(Finnigan, Hanson-Smith, Stevens, & Thornton, 2012). These data are highly suggestive that the presence of a gene paralog in a molecular machine does not always alter actual function but may just alter its actual assembly adding further complexity to gene paralogs within macromolecular complexes.
This idea of a paralog changing macromolecular composition is also reported in our data. Again, the exact stoichiometry of a functional mtCU is still unresolved but it is the general consensus that MCU forms tetramers in a dimers of dimers configuration. With the theory of paralog interference in mind, MCUB, with ~50% a.a. similarity to MCU, likely exists and complexes directly with MCU which our data supports because we found MCU and MCUB directly interacted in
96 exogenous and endogenous immunoprecipitations. Also in silico using the computer modeling program MODELLER we modeled the structure of MCUB based off already existing models of fungal MCU and found it generates a similar protein with a structure clearly similar to MCU suggesting these two paralogs still have the ability to complex with each other (Fig 4-3). By employing size exclusion chromatography we find MCUB deletion in HeLa cells increases the presence of MCU as well as MICU1 and MICU2 in fractions containing the high-
MW uniporter. It is possible that MCUB deletion is somehow allowing an increase in uniporter density but we hypothesize that upon MCUB deletion MCU subunits are now taking the place of lost MCUB. Essentially upon MCUB deletion the stoichiometry of the pore is altered where every mtCU now has 4 monomeric units of MCU. Because MCU interacts with MICU1/2 it is logical that increasing
MCU would also increase MICU1/2 within mtCU. Further, we also show what happens to the molecular compositions of mtCU when we alter the stoichiometry of MCU:MCUB by overexpressing MCUB by about 15 fold at the protein level thus altering the MCU:MCUB protein ratio from roughly 20 MCU for every 1
MCUB to 1 MCU for every ~.6 MCUB. While it is unclear how this change in the protein ratio would reflect changes in the stoichiometry of the tetrameric pore we did assess how it changed the overall molecular composition of the mtCU by size exclusion chromatography. We found MCUB incorporated into high MW fractions and displaced MCU as well as MICU1 and MICU2 from the uniporter. This also reduced the overall molecular weight size of the uniporter.
97 Because MCUB altered the composition of the pore it led us to hypothesize that one possible difference between it and its gene paralog MCU is that it may not interact with MICU1 or MICU2. This is exactly what our endogenous and exogenous immunoprecipitations confirmed because MCU directly interacted with MICU1 and MICU2, but MCUB did not. This is quite important because the expression of MCU and MCUB is differential and also tissue independent(D. De Stefani et al., 2011). The expression level of MICU1 is also tissue dependent and its expression level has directly been shown to influence mtCU function suggesting tissue dependent expression of MICU1 reflects an ability to decode Ca2+ specific signals(Paillard et al., 2017). Whether
MCUB’s tissue dependent mRNA expression also plays a role in decoding tissue specific Ca2+ signaling warrants further investigation.
We did additionally confirm that altering the composition of mtCU by deleting MCUB actually did alter its function. Upon MCUB deletion we found increased MICU1/2 in high MW fractions containing the mtCU. MICU1/2 set a
Ca2+ threshold for entry and impart cooperativity of channel activation in the face of increasing Ca2+ concentrations(Patron et al., 2014). We did not see that
MCUB deletion altered the Ca2+ threshold for entry but we did see enhanced cooperativity further supporting the hypothesis that increased MICU1 in the uniporter enhances cooperativity. This shows the absence of MCUB is not just altering the composition of mtCU but is additionally influencing its function which further warrants investigation into what MCUB’s tissue dependent expression means physiologically. We additionally would like to point out that the mRNA
98 ratio of MCU:MCUB in the heart is actually quite high 3:1 compared to 30:1 in skeletal muscle(Raffaello et al., 2013). This does not reflect what is observed in the actual mtCU though as we are unable to observe the presence of MCUB in the mtCU of the homeostatic heart suggesting mRNA levels do not always reflect the actual composition of the high-MW mtCU which is something that is important to keep in mind as the physiological relevance of this gene is further determined.
So what are the amino acids within MCUB that do not allow interaction with MICU1/2? MCU directly and indirectly interacts with MICU1. MCU has been shown to directly act with MICU1 through a DIME interacting domain(Paillard et al., 2018). MICU1 has also been shown to interact with EMRE and it is thought this makes MCU and MICU1 to have even tighter interactions(Tsai et al., 2016).
MCUB actually does contain the same proposed DIME interacting domain but does have key amino acid differences directly around the DID suggesting these amino acids may somehow interrupt MCUB’s interaction with MICU1 although further investigation is needed to support this hypothesis. In data not shown we additionally find that MCUB interacts with EMRE and why MCUB cannot interact indirectly with MICU1 also warrants further investigation.
2+ Upon MCUB gain-of-function we see a dramatic decrease in mCa uptake
2+ rates and mCa transients in ACMs suggesting MCUB incorporation results in an inactive channel. It is unclear under what mechanism this is occurring though.
According to the studies that have determined the architecture of the pore the D within the DIME forms the narrowest portion of the pore. Mutations within D or E of DIME completely abrogate channel function. This DIME motif is preserved
99 within MCUB so it is unclear whether MCUB exerts its function on mtCU by directly acting on this pore region.
2+ So if MCUB is not directly acting on the pore how is it limiting mCa uptake and what is MCUB’s true mechanism of action. Besides the DIME motif being critical for MCU mediated Ca2+ uptake it is also suggested that the second transmembrane domain of MCU actually forms the inner pore whose hydrophilic residues are essential for Ca2+ conductance contributing to the high capacitance of the channel(Oxenoid et al., 2016). MCUB’s second transmembrane domain contains a different amino acid sequence than that of MCU and these AA differences may result in blocking Ca2+ uptake. Further investigation of the critical differences in AA between MCU and MCUB and how they affect MICU1 binding or Ca2+ uptake are warranted and of great interest.
Ischemic Cardiac Injury: Hif1-alpha
Our results show MCUB expression is increased and actually incorporated into the high-MW mtCU following both MI and I/R. It is important to consider how
MCUB expression could be induced in both of these forms of cardiac injury.
Interestingly, MCUB has been found to be highly expressed at the mRNA level and protein level in high grade glioma human brain tissue compared to low grade gliomas or normal brain tissue(Xu et al., 2017). In fact, MCUB expression in human glioblastoma multiforme brain samples was found to be highest in areas that bordered necrosis that also immunostained for Hypoxia inducible factor-1α,
HIF-1α. This study also indicated that culturing glial cells in hypoxic conditions
100 increased MCUB protein expression and silencing HIF-1α reduced MCUB mRNA expression even under hypoxic conditions. This suggests HIF-1α may regulate
MCUB activity in human glioblastomas but leads us to the question; Why would
MCUB be upregulated during hypoxia in a HIF1α dependent manner?
HIF-1α is member of the hypoxia inducible factor transcription factor family which regulate a variety of genes affecting diverse cellular processes such as survival, metabolism, and angiogenesis(Dengler, Galbraith, & Espinosa, 2014).
HIFα’s are normally hydroxylated by prolyl hydroxylase domain containing enzymes (PHDs) leading to their degradation(Weidemann & Johnson, 2008).
During hypoxia PHD activity is decreased because they depend on O2 as a substrate thus HIFα’s are no longer degraded and can activate transcription of target genes(Ivan et al., 2001).
It is well understood that HIF-1α elicits protective signaling during stroke and myocardial ischemia and aids in promoting the shift from aerobic metabolism to anaerobic glycolysis(Ratan et al., 2007; Shohet & Garcia, 2007). In fact, HIF-
1α deleted MEFs have attenuated glycolytic switching when subjected to hypoxia(Seagroves et al., 2001). A number of metabolic genes are targets of
HIF1α. HIF1α enhances glycolytic metabolism by increasing glucose transporters(Gordan, Thompson, & Simon, 2007). HIF-1α induces pyruvate dehydrogenase kinase 1 thus negatively regulating the pyruvate dehydrogenase complex preventing pyruvates entrance into the TCA cycle(J. W. Kim,
Tchernyshyov, Semenza, & Dang, 2006). HIF-1α also increases expression of
Hexokinase II which is the rate limiting step of glycolysis thus increasing
101 glycolytic metabolism(J.-w. Kim, Gao, Liu, Semenza, & Dang, 2007). Additionally in the liver it has been shown that hypoxia inhibits β-oxidation and a recent study suggests this may be due to HIF2α(Rankin et al., 2009). All of these studies suggest HIF-1α signaling which is present in ischemic cardiac injury enhances glycolysis and reduces reliance on fatty acids.
We show MCUB is upregulated at the mRNA level 48 hours following MI and it is interesting to think this may also be a HIF-1α dependent process in order
2+ to shift away from mCa dependent activation of the TCA cycle and PDH complex in order to prevent OxPhos. Indeed when a promoter blast is performed on the MCUB gene using MatInspector we find a HIF response element (HRE)
611 nucleotides upstream of the transcription start site (Fig 4-4). Further investigation is needed to establish whether this HRE is being targeted by HIF-1α in order to upregulate MCUB to aid in the glycolytic shift seen during ischemic cardiac stress. It is also worth noting that this HIF-1α upregulation of MCUB may
2+ have an either unintended or intended secondary effect of preventing mCa overload during permanent ischemia.
Ischemia Reperfusion Injury: AP-1
During IR injury a period of ischemia is followed by rapid reoxygenation of the myocardium which greatly augments oxidative stress leading to cellular damage.
Oxidative stress has long been known to activate the transcription factor activator protein 1 (AP-1) in cells(S. Y. Hong, Roze, & Linz, 2013; Lo, Wong, & Cruz,
1996). AP-1 activation has been observed during cardiac IR injury and is thought
102 to regulate apoptotic genes or cell survival genes(Alfonso-Jaume et al., 2006;
Yang et al., 2003). Promoter blasting of the MCUB gene found an AP-1 Binding
Site 956 nucleotides upstream from the transcription start site of MCUB (Fig 4-4).
2+ ROS further increases iCa levels and AP-1 dependent signaling may
2+ upregulate MCUB to protect against mCa overload during the reperfusion period of I/R injury. Reoxygenation of the ischemic myocardium causes cells to switch metabolism back to OxPhos(Bopassa, 2012). OxPhos generates mitochondrial sourced ROS and thus AP-1 may additionally be activated in this situation to
2+ upregulate MCUB in order to prevent mCa induced ROS production through stimulation of OxPhos. Further investigation is warranted.
Translational Significance
2+ We still lack specific therapeutics for cardiac I/R injury that target mCa induced mPTP activated necrotic cell death. MCU mediated Ca2+ uptake in the homeostatic heart is generally beneficial to the cardiomyocyte because it stimulates cellular bioenergetics to meet contractile responsiveness due to induction of cardiac contractility. This makes drugs that target MCU for silencing of gene expression or protein degradation not ideal because although they would
2+ prevent mCa induced mPTP they would also impair cellular bioenergetics which would have negative cellular consequences. This makes MCUB an even more attractive target for therapy because it acts directly on the uniporter to negatively regulate activity and it maybe possible to take advantage of this functionality in
2+ order to fine tune inhibition of mCa uptake during IR injury. For example, once
103 the amino acid sequence of MCUB responsible for its direct mechanism of action is determined it will allow for the development of small peptide inhibitors that mimic this amino acid sequence. These small peptides could be developed in a
2+ way in which they could be used to block mCa uptake for a short period of time but be unstable enough to become degraded rather quickly allowing mtCU to quickly become functionally again which may allow for restoration of cellular bioenergetics in a positive manner. Further research into MCUB’s translational significance is warranted and of great interest for novel drug development to treat
I/R injury.
Conclusion
Overall our work has investigated how MCUB precisely regulates the uniporter and contributes to cellular physiology using unique MCUB loss- and gain-of- function models. We show that MCUB alters the macromolecular formation of mtCU and negatively impacts mtCU function. Further, for the first time we find that MCUB is absent from mtCU in the homeostatic heart but is upregulated as a
2+ novel compensatory mechanism to limit mCa induced mPTP activated necrotic cell death during ischemic cardiac disease. Our work also identifies a novel therapeutic target for future treatment of necrotic cell death.
104
Figure 4-1. Graphical depiction of how the increase in MCUB affects 2+ cellular processes dependent on mCa signaling in the context of cardiac (patho)physiology.
105
Figure 4-2. Graphical depiction of how the increase in MCUB seen during IR 2+ injury alters the composition of mtCU to limit mCa overload.
106
Figure 4-3. Computer models of MCU and MCUB in monomeric and tetrameric form. A) Monomer of human MCU modeled from fungal MCU (pdb file, 6C5W) using the computer program MODELLER. B) Monomer of human MCUB modeled from human MCU from Fig 4-1A using the computer program MODELLER. C) Human tetrameric MCU was aligned from fungal tetrameric MCU (pdb file, 6D7W) to model MCU as a tetramer. D) Human MCUB was aligned to human MCU to model MCUB as a tetramer.
107
Figure 4-4. Promoter regions of MCUB possibly targeted by transcription factors associated with IR injury.
108
BIBLIOGRAPHY
Abel, E. D. (2004). Glucose transport in the heart. Front Biosci, 9, 201-215.
Adlakha, J., Karamichali, I., Sangwallek, J., Deiss, S., Bar, K., Coles, M., . . .
Hernandez Alvarez, B. (2019). Characterization of MCU-Binding Proteins
MCUR1 and CCDC90B - Representatives of a Protein Family Conserved
in Prokaryotes and Eukaryotic Organelles. Structure, 27(3), 464-475.e466.
doi:10.1016/j.str.2018.11.004
Akram, M. (2014). Citric Acid Cycle and Role of its Intermediates in Metabolism.
Cell Biochemistry and Biophysics, 68(3), 475-478. Retrieved from
https://doi.org/10.1007/s12013-013-9750-1. doi:10.1007/s12013-013-
9750-1
Al-Khallaf, H. (2017). Isocitrate dehydrogenases in physiology and cancer:
biochemical and molecular insight. Cell & Bioscience, 7(1), 37. Retrieved
from https://doi.org/10.1186/s13578-017-0165-3. doi:10.1186/s13578-017-
0165-3
Alfonso-Jaume, M. A., Bergman, M. R., Mahimkar, R., Cheng, S., Jin, Z. Q.,
Karliner, J. S., & Lovett, D. H. (2006). Cardiac ischemia-reperfusion injury
induces matrix metalloproteinase-2 expression through the AP-1
components FosB and JunB. American Journal of Physiology-Heart and
Circulatory Physiology, 291(4), H1838-H1846. Retrieved from
https://doi.org/10.1152/ajpheart.00026.2006.
doi:10.1152/ajpheart.00026.2006
109 Allen, L. N. (2017). Financing national non-communicable disease responses.
Global health action, 10(1), 1326687-1326687. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/28604238
https://www.ncbi.nlm.nih.gov/pmc/PMC5496084/.
doi:10.1080/16549716.2017.1326687
Altomare, C., Terragni, B., Brioschi, C., Milanesi, R., Pagliuca, C., Viscomi, C., . .
. DiFrancesco, D. (2003). Heteromeric HCN1-HCN4 channels: a
comparison with native pacemaker channels from the rabbit sinoatrial
node. J Physiol, 549(Pt 2), 347-359. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/12702747
https://www.ncbi.nlm.nih.gov/pmc/PMC2342966/.
doi:10.1113/jphysiol.2002.027698
Argaud, L., Gateau-Roesch, O., Muntean, D., Chalabreysse, L., Loufouat, J.,
Robert, D., & Ovize, M. (2005). Specific inhibition of the mitochondrial
permeability transition prevents lethal reperfusion injury. Journal of
Molecular and Cellular Cardiology, 38(2), 367-374. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0022282804003979.
doi:https://doi.org/10.1016/j.yjmcc.2004.12.001
Asp, M. L., Sjaastad, F. V., Siddiqui, J. K., Davis, J. P., & Metzger, J. M. (2016).
Effects of Modified Parvalbumin EF-Hand Motifs on Cardiac Myocyte
Contractile Function. Biophys J, 110(9), 2094-2105.
doi:10.1016/j.bpj.2016.03.037
110 Bahaji, A., Sanchez-Lopez, A. M., De Diego, N., Munoz, F. J., Baroja-Fernandez,
E., Li, J., . . . Pozueta-Romero, J. (2015). Plastidic phosphoglucose
isomerase is an important determinant of starch accumulation in
mesophyll cells, growth, photosynthetic capacity, and biosynthesis of
plastidic cytokinins in Arabidopsis. PLoS One, 10(3), e0119641.
doi:10.1371/journal.pone.0119641
Baines, C. P., Kaiser, R. A., Purcell, N. H., Blair, N. S., Osinska, H., Hambleton,
M. A., . . . Molkentin, J. D. (2005). Loss of cyclophilin D reveals a critical
role for mitochondrial permeability transition in cell death. Nature,
434(7033), 658-662. doi:10.1038/nature03434
Barr, L. A., Lambert, J. P., Shimizu, Y., Barouch, L. A., Naqvi, N., & Calvert, J. W.
(2017). Exercise training provides cardioprotection by activating and
coupling endothelial nitric oxide synthase via a β(3)-adrenergic receptor-
AMP-activated protein kinase signaling pathway. Medical gas research,
7(1), 1-8. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/28480026
https://www.ncbi.nlm.nih.gov/pmc/PMC5402342/. doi:10.4103/2045-
9912.202904
Baughman, J. M., Perocchi, F., Girgis, H. S., Plovanich, M., Belcher-Timme, C.
A., Sancak, Y., . . . Mootha, V. K. (2011). Integrative genomics identifies
MCU as an essential component of the mitochondrial calcium uniporter.
Nature, 476(7360), 341-345. doi:10.1038/nature10234
Benjamin, E. J., Blaha, M. J., Chiuve, S. E., Cushman, M., Das, S. R., Deo, R., . .
. Muntner, P. (2017). Heart Disease and Stroke Statistics-2017 Update: A
111 Report From the American Heart Association. Circulation, 135(10), e146-
e603. doi:10.1161/cir.0000000000000485
Berg, J. M., Stryer, L., & Tymoczko, J. L. (2002). Biochemistry. 5th edition: W. H.
Freeman 2002.
Bernardi, P., Krauskopf, A., Basso, E., Petronilli, V., Blalchy-Dyson, E., Di Lisa,
F., & Forte, M. A. (2006). The mitochondrial permeability transition from in
vitro artifact to disease target. The FEBS Journal, 273(10), 2077-2099.
Retrieved from https://doi.org/10.1111/j.1742-4658.2006.05213.x.
doi:10.1111/j.1742-4658.2006.05213.x
Bernardi, P., Rasola, A., Forte, M., & Lippe, G. (2015). The Mitochondrial
Permeability Transition Pore: Channel Formation by F-ATP Synthase,
Integration in Signal Transduction, and Role in Pathophysiology. Physiol
Rev, 95(4), 1111-1155. Retrieved from
https://doi.org/10.1152/physrev.00001.2015.
doi:10.1152/physrev.00001.2015
Berridge, M. J., Lipp, P., & Bootman, M. D. (2000). The versatility and
universality of calcium signalling. Nat Rev Mol Cell Biol, 1(1), 11-21.
doi:10.1038/35036035
Bers, D. M., Despa, S., & Bossuyt, J. (2006). Regulation of Ca2+ and Na+ in
normal and failing cardiac myocytes. Ann N Y Acad Sci, 1080, 165-177.
doi:10.1196/annals.1380.015
Bick, A. G., Wakimoto, H., Kamer, K. J., Sancak, Y., Goldberger, O., Axelsson,
A., . . . Seidman, C. E. (2017). Cardiovascular homeostasis dependence
112 on MICU2, a regulatory subunit of the mitochondrial calcium uniporter.
Proc Natl Acad Sci U S A, 114(43), E9096-e9104.
doi:10.1073/pnas.1711303114
Bleier, L., & Dröse, S. (2013). Superoxide generation by complex III: From
mechanistic rationales to functional consequences. Biochimica et
Biophysica Acta (BBA) - Bioenergetics, 1827(11), 1320-1331. Retrieved
from
http://www.sciencedirect.com/science/article/pii/S0005272812011024.
doi:https://doi.org/10.1016/j.bbabio.2012.12.002
Bodi, I., Mikala, G., Koch, S. E., Akhter, S. A., & Schwartz, A. (2005). The L-type
calcium channel in the heart: the beat goes on. J Clin Invest, 115(12),
3306-3317. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/16322774
https://www.ncbi.nlm.nih.gov/pmc/PMC1297268/. doi:10.1172/JCI27167
Boerries, M., Most, P., Gledhill, J. R., Walker, J. E., Katus, H. A., Koch, W. J., . . .
Schoenenberger, C. A. (2007). Ca2+ -dependent interaction of S100A1
with F1-ATPase leads to an increased ATP content in cardiomyocytes.
Mol Cell Biol, 27(12), 4365-4373. doi:10.1128/mcb.02045-06
Boord, J. B., Fazio, S., & Linton, M. F. (2002). Cytoplasmic fatty acid-binding
proteins: emerging roles in metabolism and atherosclerosis. Curr Opin
Lipidol, 13(2), 141-147.
Bopassa, J. C. (2012). Protection of the ischemic myocardium during the
reperfusion: between hope and reality. American journal of cardiovascular
113 disease, 2(3), 223-236. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/22937492
https://www.ncbi.nlm.nih.gov/pmc/PMC3427983/.
Bowler, M. W. (2013). Conformational dynamics in phosphoglycerate kinase, an
open and shut case? FEBS Letters, 587(13), 1878-1883. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0014579313003542.
doi:https://doi.org/10.1016/j.febslet.2013.05.012
Brini, M., Cali, T., Ottolini, D., & Carafoli, E. (2014). Neuronal calcium signaling:
function and dysfunction. Cell Mol Life Sci, 71(15), 2787-2814.
doi:10.1007/s00018-013-1550-7
Bruce, J. I. E. (2018). Metabolic regulation of the PMCA: Role in cell death and
survival. Cell Calcium, 69, 28-36. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0143416017300520.
doi:https://doi.org/10.1016/j.ceca.2017.06.001
Bunik, V. I., Tylicki, A., & Lukashev, N. V. (2013). Thiamin diphosphate-
dependent enzymes: from enzymology to metabolic regulation, drug
design and disease models. The FEBS Journal, 280(24), 6412-6442.
Retrieved from https://doi.org/10.1111/febs.12512.
doi:10.1111/febs.12512
Bush, A. I. (2010). Kalzium ist nicht alles. Neuron, 65(2), 143-144.
doi:10.1016/j.neuron.2010.01.015
Bygrave, F. L., & Ash, G. R. (1977). Development of mitochondrial calcium
transport activity in rat liver. FEBS Letters, 80(2), 271-274. Retrieved from
114 http://www.sciencedirect.com/science/article/pii/0014579377804559.
doi:https://doi.org/10.1016/0014-5793(77)80455-9
Catalá, A., & Díaz, M. (2016). Editorial: Impact of Lipid Peroxidation on the
Physiology and Pathophysiology of Cell Membranes. Frontiers in
physiology, 7, 423-423. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/27713704
https://www.ncbi.nlm.nih.gov/pmc/PMC5031777/.
doi:10.3389/fphys.2016.00423
Cecchini, G. (2003). Function and Structure of Complex II of the Respiratory
Chain. Annual Review of Biochemistry, 72(1), 77-109. Retrieved from
https://doi.org/10.1146/annurev.biochem.72.121801.161700.
doi:10.1146/annurev.biochem.72.121801.161700
Chang, Y. C., Yang, Y. C., Tien, C. P., Yang, C. J., & Hsiao, M. (2018). Roles of
Aldolase Family Genes in Human Cancers and Diseases. Trends
Endocrinol Metab, 29(8), 549-559. doi:10.1016/j.tem.2018.05.003
Chaudhuri, D., Sancak, Y., Mootha, V. K., & Clapham, D. E. (2013). MCU
encodes the pore conducting mitochondrial calcium currents. Elife, 2,
e00704. doi:10.7554/eLife.00704
Chen, S., & Li, S. (2012). The Na+/Ca²+ exchanger in cardiac
ischemia/reperfusion injury. Medical science monitor : international
medical journal of experimental and clinical research, 18(11), RA161-
RA165. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/23111750
115 https://www.ncbi.nlm.nih.gov/pmc/PMC3560609/.
doi:10.12659/MSM.883533
Chen, Y. R., & Zweier, J. L. (2014). Cardiac mitochondria and reactive oxygen
species generation. Circ Res, 114(3), 524-537.
doi:10.1161/circresaha.114.300559
Clapham, D. E. (2007). Calcium Signaling. Cell, 131(6), 1047-1058. Retrieved
from
http://www.sciencedirect.com/science/article/pii/S0092867407015310.
doi:https://doi.org/10.1016/j.cell.2007.11.028
Corbalan-Garcia, S., & Gómez-Fernández, J. C. (2014). Signaling through C2
domains: More than one lipid target. Biochimica et Biophysica Acta (BBA)
- Biomembranes, 1838(6), 1536-1547. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0005273614000108.
doi:https://doi.org/10.1016/j.bbamem.2014.01.008
Crompton, M., Ellinger, H., & Costi, A. (1988). Inhibition by cyclosporin A of a
Ca2+-dependent pore in heart mitochondria activated by inorganic
phosphate and oxidative stress. Biochem J, 255(1), 357-360. Retrieved
from https://www.ncbi.nlm.nih.gov/pubmed/3196322
https://www.ncbi.nlm.nih.gov/pmc/PMC1135230/.
Csordas, G., Golenar, T., Seifert, E. L., Kamer, K. J., Sancak, Y., Perocchi, F., . .
. Hajnoczky, G. (2013). MICU1 controls both the threshold and
cooperative activation of the mitochondrial Ca(2)(+) uniporter. Cell Metab,
17(6), 976-987. doi:10.1016/j.cmet.2013.04.020
116 Cung, T. T., Morel, O., Cayla, G., Rioufol, G., Garcia-Dorado, D., Angoulvant, D.,
. . . Ovize, M. (2015). Cyclosporine before PCI in Patients with Acute
Myocardial Infarction. N Engl J Med, 373(11), 1021-1031.
doi:10.1056/NEJMoa1505489
Day, I. S., Reddy, V. S., Shad Ali, G., & Reddy, A. S. N. (2002). Analysis of EF-
hand-containing proteins in Arabidopsis. Genome biology, 3(10),
RESEARCH0056-RESEARCH0056. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/12372144
https://www.ncbi.nlm.nih.gov/pmc/PMC134623/.
De Stefani, D., Raffaello, A., Teardo, E., Szabo, I., & Rizzuto, R. (2011). A forty-
kilodalton protein of the inner membrane is the mitochondrial calcium
uniporter. Nature, 476(7360), 336-340. doi:10.1038/nature10230
De Stefani, D., Raffaello, A., Teardo, E., Szabò, I., & Rizzuto, R. (2011). A forty-
kilodalton protein of the inner membrane is the mitochondrial calcium
uniporter. Nature, 476, 336. Retrieved from
http://dx.doi.org/10.1038/nature10230. doi:10.1038/nature10230
https://www.nature.com/articles/nature10230#supplementary-information
Deluca, H. F., & Engstrom, G. W. (1961). Calcium uptake by rat kidney
mitochondria. Proc Natl Acad Sci U S A, 47, 1744-1750.
Dengler, V. L., Galbraith, M., & Espinosa, J. M. (2014). Transcriptional regulation
by hypoxia inducible factors. Critical reviews in biochemistry and
molecular biology, 49(1), 1-15. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/24099156
117 https://www.ncbi.nlm.nih.gov/pmc/PMC4342852/.
doi:10.3109/10409238.2013.838205
Denton, R. M., McCormack, J. G., & Edgell, N. J. (1980). Role of calcium ions in
the regulation of intramitochondrial metabolism. Effects of Na+, Mg2+ and
ruthenium red on the Ca2+-stimulated oxidation of oxoglutarate and on
pyruvate dehydrogenase activity in intact rat heart mitochondria. Biochem
J, 190(1), 107-117.
Denton, R. M., Randle, P. J., & Martin, B. R. (1972). Stimulation by calcium ions
of pyruvate dehydrogenase phosphate phosphatase. Biochem J, 128(1),
161-163.
Deschauer, M., Wieser, T., & Zierz, S. (2005). Muscle Carnitine
Palmitoyltransferase II Deficiency: Clinical and Molecular Genetic
Features and Diagnostic Aspects. Archives of Neurology, 62(1), 37-41.
Retrieved from https://doi.org/10.1001/archneur.62.1.37.
doi:10.1001/archneur.62.1.37
Dewenter, M., von der Lieth, A., Katus Hugo, A., & Backs, J. (2017). Calcium
Signaling and Transcriptional Regulation in Cardiomyocytes. Circ Res,
121(8), 1000-1020. Retrieved from
https://doi.org/10.1161/CIRCRESAHA.117.310355.
doi:10.1161/CIRCRESAHA.117.310355
Dhar, A., Samiotakis, A., Ebbinghaus, S., Nienhaus, L., Homouz, D., Gruebele,
M., & Cheung, M. S. (2010). Structure, function, and folding of
phosphoglycerate kinase are strongly perturbed by macromolecular
118 crowding. Proceedings of the National Academy of Sciences, 107(41),
17586. Retrieved from
http://www.pnas.org/content/107/41/17586.abstract.
doi:10.1073/pnas.1006760107
Doenst, T., Nguyen, T. D., & Abel, E. D. (2013). Cardiac metabolism in heart
failure: implications beyond ATP production. Circ Res, 113(6), 709-724.
doi:10.1161/circresaha.113.300376
Donato, R., Cannon, B. R., Sorci, G., Riuzzi, F., Hsu, K., Weber, D. J., & Geczy,
C. L. (2013). Functions of S100 proteins. Current molecular medicine,
13(1), 24-57. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/22834835
https://www.ncbi.nlm.nih.gov/pmc/PMC3707951/.
Drake, S. K., Zimmer, M. A., Kundrot, C., & Falke, J. J. (1997). Molecular tuning
of an EF-hand-like calcium binding loop. Contributions of the coordinating
side chain at loop position 3. The Journal of general physiology, 110(2),
173-184. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/9236210
https://www.ncbi.nlm.nih.gov/pmc/PMC2233790/.
Dunaway, G. A. (1983). A review of animal phosphofructokinase isozymes with
an emphasis on their physiological role. Molecular and Cellular
Biochemistry, 52(1), 75-91. Retrieved from
https://doi.org/10.1007/BF00230589. doi:10.1007/BF00230589
119 Eisner, D. A., Caldwell, J. L., Kistamás, K., & Trafford, A. W. (2017). Calcium and
Excitation-Contraction Coupling in the Heart. Circ Res, 121(2), 181-195.
Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/28684623
https://www.ncbi.nlm.nih.gov/pmc/PMC5497788/.
doi:10.1161/CIRCRESAHA.117.310230
Elrod John, W., Duranski Mark, R., Langston, W., Greer James, J. M., Tao, L.,
Dugas Tammy, R., . . . Lefer David, J. (2006). eNOS Gene Therapy
Exacerbates Hepatic Ischemia-Reperfusion Injury in Diabetes. Circ Res,
99(1), 78-85. Retrieved from
https://doi.org/10.1161/01.RES.0000231306.03510.77.
doi:10.1161/01.RES.0000231306.03510.77
Elrod, J. W., Calvert, J. W., Morrison, J., Doeller, J. E., Kraus, D. W., Tao, L., . . .
Lefer, D. J. (2007). Hydrogen sulfide attenuates myocardial ischemia-
reperfusion injury by preservation of mitochondrial function. Proc Natl
Acad Sci U S A, 104(39), 15560-15565. doi:10.1073/pnas.0705891104
Elrod, J. W., Wong, R., Mishra, S., Vagnozzi, R. J., Sakthievel, B., Goonasekera,
S. A., . . . Molkentin, J. D. (2010). Cyclophilin D controls mitochondrial
pore-dependent Ca(2+) exchange, metabolic flexibility, and propensity for
heart failure in mice. J Clin Invest, 120(10), 3680-3687.
doi:10.1172/jci43171
Fan, C., Fan, M., Orlando, B. J., Fastman, N. M., Zhang, J., Xu, Y., . . . Feng, L.
(2018). X-ray and cryo-EM structures of the mitochondrial calcium
uniporter. Nature, 559(7715), 575-579. doi:10.1038/s41586-018-0330-9
120 Fearnley, C. J., Roderick, H. L., & Bootman, M. D. Calcium signaling in cardiac
myocytes. Cold Spring Harb Perspect Biol, 3(11), a004242-a004242.
Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/21875987
https://www.ncbi.nlm.nih.gov/pmc/PMC3220352/.
doi:10.1101/cshperspect.a004242
Fieni, F., Lee, S. B., Jan, Y. N., & Kirichok, Y. (2012). Activity of the mitochondrial
calcium uniporter varies greatly between tissues. Nature communications,
3, 1317-1317. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/23271651
https://www.ncbi.nlm.nih.gov/pmc/PMC3818247/.
doi:10.1038/ncomms2325
Finnigan, G. C., Hanson-Smith, V., Stevens, T. H., & Thornton, J. W. (2012).
Evolution of increased complexity in a molecular machine. Nature,
481(7381), 360-364. doi:10.1038/nature10724
Forgac, M. (2007). Vacuolar ATPases: rotary proton pumps in physiology and
pathophysiology. Nat Rev Mol Cell Biol, 8(11), 917-929.
doi:10.1038/nrm2272
Frezza, C., Cipolat, S., & Scorrano, L. (2007). Organelle isolation: functional
mitochondria from mouse liver, muscle and cultured fibroblasts. Nat
Protoc, 2(2), 287-295. doi:10.1038/nprot.2006.478
Fukui, H., Mizuguchi, H., Nemoto, H., Kitamura, Y., Kashiwada, Y., & Takeda, N.
(2017). Histamine H1 Receptor Gene Expression and Drug Action of
121 Antihistamines. Handb Exp Pharmacol, 241, 161-169.
doi:10.1007/164_2016_14
Fukuta, H., & Little, W. C. (2008). The cardiac cycle and the physiologic basis of
left ventricular contraction, ejection, relaxation, and filling. Heart Fail Clin,
4(1), 1-11. doi:10.1016/j.hfc.2007.10.004
Furuyama, K., & Sassa, S. (2000). Interaction between succinyl CoA synthetase
and the heme-biosynthetic enzyme ALAS-E is disrupted in sideroblastic
anemia. J Clin Invest, 105(6), 757-764. Retrieved from
https://doi.org/10.1172/JCI6816. doi:10.1172/JCI6816
Gao, E., Lei, Y. H., Shang, X., Huang, Z. M., Zuo, L., Boucher, M., . . . Koch, W.
J. (2010). A novel and efficient model of coronary artery ligation and
myocardial infarction in the mouse. Circ Res, 107(12), 1445-1453.
doi:10.1161/circresaha.110.223925
Gibb, A. A., & Hill, B. G. (2018). Metabolic Coordination of Physiological and
Pathological Cardiac Remodeling. Circ Res, 123(1), 107-128. Retrieved
from https://www.ncbi.nlm.nih.gov/pubmed/29929976
https://www.ncbi.nlm.nih.gov/pmc/PMC6023588/.
doi:10.1161/CIRCRESAHA.118.312017
Gibbs, C. L. (2003). Cardiac energetics: sense and nonsense. Clin Exp
Pharmacol Physiol, 30(8), 598-603.
Glancy, B., & Balaban, R. S. (2012). Role of Mitochondrial Ca2+ in the
Regulation of Cellular Energetics. Biochemistry, 51(14), 2959-2973.
Retrieved from https://doi.org/10.1021/bi2018909. doi:10.1021/bi2018909
122 Glatz, J. F. C., van Bilsen, M., & van der Vusse, G. J. (2000). Cardiac fatty acid
uptake and transport in health and disease. Cardiovascular Research,
45(2), 279-293. Retrieved from https://doi.org/10.1016/S0008-
6363(99)00263-1. doi:10.1016/S0008-6363(99)00263-1
Gordan, J. D., Thompson, C. B., & Simon, M. C. (2007). HIF and c-Myc: sibling
rivals for control of cancer cell metabolism and proliferation. Cancer Cell,
12(2), 108-113. doi:10.1016/j.ccr.2007.07.006
Grabarek, Z. (2006). Structural Basis for Diversity of the EF-hand Calcium-
binding Proteins. Journal of Molecular Biology, 359(3), 509-525. Retrieved
from
http://www.sciencedirect.com/science/article/pii/S0022283606004475.
doi:https://doi.org/10.1016/j.jmb.2006.03.066
Granger, D. N., & Kvietys, P. R. (2015). Reperfusion injury and reactive oxygen
species: The evolution of a concept. Redox Biology, 6, 524-551. Retrieved
from https://www.ncbi.nlm.nih.gov/pubmed/26484802
https://www.ncbi.nlm.nih.gov/pmc/PMC4625011/.
doi:10.1016/j.redox.2015.08.020
Griffiths, E. J., & Halestrap, A. P. (1991). Further evidence that cyclosporin A
protects mitochondria from calcium overload by inhibiting a matrix
peptidyl-prolyl cis-trans isomerase. Implications for the
immunosuppressive and toxic effects of cyclosporin. Biochem J, 274 ( Pt
2)(Pt 2), 611-614. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/1706598
123 https://www.ncbi.nlm.nih.gov/pmc/PMC1150183/.
Guerini, D., Coletto, L., & Carafoli, E. (2005). Exporting calcium from cells. Cell
Calcium, 38(3-4), 281-289. doi:10.1016/j.ceca.2005.06.032
Gunter, T. E., & Pfeiffer, D. R. (1990). Mechanisms by which mitochondria
transport calcium. American Journal of Physiology-Cell Physiology,
258(5), C755-C786. Retrieved from
https://doi.org/10.1152/ajpcell.1990.258.5.C755.
doi:10.1152/ajpcell.1990.258.5.C755
Hake, J., & Lines, G. T. (2008). Stochastic Binding of Ca2+ Ions in the Dyadic
Cleft; Continuous versus Random Walk Description of Diffusion. Biophys
J, 94(11), 4184-4201. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0006349508700766.
doi:https://doi.org/10.1529/biophysj.106.103523
Halestrap, A. P., Clarke, S. J., & Javadov, S. A. (2004). Mitochondrial
permeability transition pore opening during myocardial reperfusion--a
target for cardioprotection. Cardiovasc Res, 61(3), 372-385.
doi:10.1016/s0008-6363(03)00533-9
Halestrap, A. P., & Davidson, A. M. (1990). Inhibition of Ca2(+)-induced large-
amplitude swelling of liver and heart mitochondria by cyclosporin is
probably caused by the inhibitor binding to mitochondrial-matrix peptidyl-
prolyl cis-trans isomerase and preventing it interacting with the adenine
nucleotide translocase. Biochem J, 268(1), 153-160. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/2160810
124 https://www.ncbi.nlm.nih.gov/pmc/PMC1131405/.
Halestrap, A. P., & Pasdois, P. (2009). The role of the mitochondrial permeability
transition pore in heart disease. Biochimica et Biophysica Acta (BBA) -
Bioenergetics, 1787(11), 1402-1415. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0005272809000073.
doi:https://doi.org/10.1016/j.bbabio.2008.12.017
Hallworth, M. (2014). CHAPTER 39 - Therapeutic drug monitoring. In W. J.
Marshall, M. Lapsley, A. P. Day, & R. M. Ayling (Eds.), Clinical
Biochemistry: Metabolic and Clinical Aspects (Third Edition) (pp. 767-786):
Churchill Livingstone.
Harris, R. A., Bowker-Kinley, M. M., Huang, B., & Wu, P. (2002). Regulation of
the activity of the pyruvate dehydrogenase complex. Adv Enzyme Regul,
42, 249-259.
Hausenloy, D. J., Duchen, M. R., & Yellon, D. M. (2003). Inhibiting mitochondrial
permeability transition pore opening at reperfusion protects against
ischaemia-reperfusion injury. Cardiovasc Res, 60(3), 617-625.
Hazelton, J. L., Petrasheuskaya, M., Fiskum, G., & Kristián, T. (2009).
Cyclophilin D is expressed predominantly in mitochondria of gamma-
aminobutyric acidergic interneurons. Journal of neuroscience research,
87(5), 1250-1259. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/18951528
https://www.ncbi.nlm.nih.gov/pmc/PMC2650012/. doi:10.1002/jnr.21921
125 Hederstedt, L., & Rutberg, L. (1981). Succinate dehydrogenase--a comparative
review. Microbiological reviews, 45(4), 542-555. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/6799760
https://www.ncbi.nlm.nih.gov/pmc/PMC281527/.
Hennekens, C. H., & Gaziano, J. M. (1993). Antioxidants and heart disease:
Epidemiology and clinical evidence. Clinical Cardiology, 16(S1), 10-15.
Retrieved from https://doi.org/10.1002/clc.4960161305.
doi:10.1002/clc.4960161305
Hinkle, P. C. (2005). P/O ratios of mitochondrial oxidative phosphorylation.
Biochimica et Biophysica Acta (BBA) - Bioenergetics, 1706(1), 1-11.
Retrieved from
http://www.sciencedirect.com/science/article/pii/S0005272804002701.
doi:https://doi.org/10.1016/j.bbabio.2004.09.004
Hirst, J. (2013). Mitochondrial Complex I. Annual Review of Biochemistry, 82(1),
551-575. Retrieved from https://doi.org/10.1146/annurev-biochem-
070511-103700. doi:10.1146/annurev-biochem-070511-103700
Hogan, P. G., Chen, L., Nardone, J., & Rao, A. (2003). Transcriptional regulation
by calcium, calcineurin, and NFAT. Genes Dev, 17(18), 2205-2232.
doi:10.1101/gad.1102703
Hong, S. Y., Roze, L. V., & Linz, J. E. (2013). Oxidative stress-related
transcription factors in the regulation of secondary metabolism. Toxins
(Basel), 5(4), 683-702. doi:10.3390/toxins5040683
126 Hong, T., & Shaw, R. M. (2016). Cardiac T-Tubule Microanatomy and Function.
Physiol Rev, 97(1), 227-252. Retrieved from
https://doi.org/10.1152/physrev.00037.2015.
doi:10.1152/physrev.00037.2015
Hopper, R. K., Carroll, S., Aponte, A. M., Johnson, D. T., French, S., Shen, R. F.,
. . . Balaban, R. S. (2006). Mitochondrial matrix phosphoproteome: effect
of extra mitochondrial calcium. Biochemistry, 45(8), 2524-2536.
doi:10.1021/bi052475e
Houten, S. M., & Wanders, R. J. A. (2010). A general introduction to the
biochemistry of mitochondrial fatty acid β-oxidation. Journal of inherited
metabolic disease, 33(5), 469-477. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/20195903
https://www.ncbi.nlm.nih.gov/pmc/PMC2950079/. doi:10.1007/s10545-
010-9061-2
Huang, B., Gudi, R., Wu, P., Harris, R. A., Hamilton, J., & Popov, K. M. (1998).
Isoenzymes of pyruvate dehydrogenase phosphatase. DNA-derived
amino acid sequences, expression, and regulation. J Biol Chem, 273(28),
17680-17688.
Inserte, J., Garcia-Dorado, D., Hernando, V., & Soler-Soler, J. (2005). Calpain-
Mediated Impairment of Na+/K+–ATPase Activity During Early
Reperfusion Contributes to Cell Death After Myocardial Ischemia. Circ
Res, 97(5), 465-473. Retrieved from
127 https://doi.org/10.1161/01.RES.0000181170.87738.f3.
doi:10.1161/01.RES.0000181170.87738.f3
Israelsen, W. J., & Vander Heiden, M. G. (2015). Pyruvate kinase: Function,
regulation and role in cancer. Semin Cell Dev Biol, 43, 43-51.
doi:10.1016/j.semcdb.2015.08.004
Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., . . . Kaelin, W. G.,
Jr. (2001). HIFalpha targeted for VHL-mediated destruction by proline
hydroxylation: implications for O2 sensing. Science, 292(5516), 464-468.
doi:10.1126/science.1059817
Jeffs, G. J., Meloni, B. P., Bakker, A. J., & Knuckey, N. W. (2007). The role of the
Na+/Ca2+ exchanger (NCX) in neurons following ischaemia. Journal of
Clinical Neuroscience, 14(6), 507-514. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0967586806006187.
doi:https://doi.org/10.1016/j.jocn.2006.07.013
Ji, H., Wang, J., Guo, J., Li, Y., Lian, S., Guo, W., . . . Liu, Y. (2016). Progress in
the biological function of alpha-enolase. Animal Nutrition, 2(1), 12-17.
Retrieved from
http://www.sciencedirect.com/science/article/pii/S2405654515300755.
doi:https://doi.org/10.1016/j.aninu.2016.02.005
Jonckheere, A. I., Smeitink, J. A. M., & Rodenburg, R. J. T. (2012). Mitochondrial
ATP synthase: architecture, function and pathology. Journal of inherited
metabolic disease, 35(2), 211-225. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/21874297
128 https://www.ncbi.nlm.nih.gov/pmc/PMC3278611/. doi:10.1007/s10545-
011-9382-9
Jozef, K., Stephan, K., & Barbara, K. (2013). Calcium in the Early Evolution of
Living Systems: A Biohistorical Approach. Current Organic Chemistry,
17(16), 1738-1750. Retrieved from
http://www.eurekaselect.com/node/113713/article.
doi:http://dx.doi.org/10.2174/13852728113179990081
Kadamur, G., & Ross, E. M. (2013). Mammalian phospholipase C. Annu Rev
Physiol, 75, 127-154. doi:10.1146/annurev-physiol-030212-183750
Kalogeris, T., Baines, C. P., Krenz, M., & Korthuis, R. J. (2012). Cell biology of
ischemia/reperfusion injury. International review of cell and molecular
biology, 298, 229-317. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/22878108
https://www.ncbi.nlm.nih.gov/pmc/PMC3904795/. doi:10.1016/B978-0-12-
394309-5.00006-7
Kim, J.-w., Gao, P., Liu, Y.-C., Semenza, G. L., & Dang, C. V. (2007). Hypoxia-
Inducible Factor 1 and Dysregulated c-Myc Cooperatively Induce Vascular
Endothelial Growth Factor and Metabolic Switches Hexokinase 2 and
Pyruvate Dehydrogenase Kinase 1. Mol Cell Biol, 27(21), 7381. Retrieved
from http://mcb.asm.org/content/27/21/7381.abstract.
doi:10.1128/MCB.00440-07
Kim, J. W., Tchernyshyov, I., Semenza, G. L., & Dang, C. V. (2006). HIF-1-
mediated expression of pyruvate dehydrogenase kinase: a metabolic
129 switch required for cellular adaptation to hypoxia. Cell Metab, 3(3), 177-
185. doi:10.1016/j.cmet.2006.02.002
Kirichok, Y., Krapivinsky, G., & Clapham, D. E. (2004). The mitochondrial calcium
uniporter is a highly selective ion channel. Nature, 427(6972), 360-364.
doi:10.1038/nature02246
Kirichok, Y., Krapivinsky, G., & Clapham, D. E. (2004). The mitochondrial calcium
uniporter is a highly selective ion channel. Nature, 427, 360. Retrieved
from http://dx.doi.org/10.1038/nature02246. doi:10.1038/nature02246
https://www.nature.com/articles/nature02246#supplementary-information
Korotchkina, L. G., & Patel, M. S. (2001). Site specificity of four pyruvate
dehydrogenase kinase isoenzymes toward the three phosphorylation sites
of human pyruvate dehydrogenase. J Biol Chem, 276(40), 37223-37229.
doi:10.1074/jbc.M103069200
Kosova, A. A., Khodyreva, S. N., & Lavrik, O. I. (2017). Role of Glyceraldehyde-
3-Phosphate Dehydrogenase (GAPDH) in DNA Repair. Biochemistry
(Mosc), 82(6), 643-654. doi:10.1134/s0006297917060013
Kovács-Bogdán, E., Sancak, Y., Kamer, K. J., Plovanich, M., Jambhekar, A.,
Huber, R. J., . . . Mootha, V. K. (2014). Reconstitution of the mitochondrial
calcium uniporter in yeast. Proceedings of the National Academy of
Sciences, 111(24), 8985. Retrieved from
http://www.pnas.org/content/111/24/8985.abstract.
doi:10.1073/pnas.1400514111
130 Kroner, H. (1986). Ca2+ ions, an allosteric activator of calcium uptake in rat liver
mitochondria. Arch Biochem Biophys, 251(2), 525-535.
Kruman, II, & Mattson, M. P. (1999). Pivotal role of mitochondrial calcium uptake
in neural cell apoptosis and necrosis. J Neurochem, 72(2), 529-540.
Kubasiak, L. A., Hernandez, O. M., Bishopric, N. H., & Webster, K. A. (2002).
Hypoxia and acidosis activate cardiac myocyte death through the Bcl-2
family protein BNIP3. Proc Natl Acad Sci U S A, 99(20), 12825-12830.
doi:10.1073/pnas.202474099
Kwong, J. Q., Lu, X., Correll, R. N., Schwanekamp, J. A., Vagnozzi, R. J.,
Sargent, M. A., . . . Molkentin, J. D. (2015). The Mitochondrial Calcium
Uniporter Selectively Matches Metabolic Output to Acute Contractile
Stress in the Heart. Cell Rep, 12(1), 15-22.
doi:10.1016/j.celrep.2015.06.002
Lambert, J. P., Nicholson, C. K., Amin, H., Amin, S., & Calvert, J. W. (2014).
Hydrogen sulfide provides cardioprotection against myocardial/ischemia
reperfusion injury in the diabetic state through the activation of the RISK
pathway. Medical gas research, 4(1), 20-20. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/25525500
https://www.ncbi.nlm.nih.gov/pmc/PMC4269946/. doi:10.1186/s13618-
014-0020-0
Lander, N., Chiurillo, M. A., Bertolini, M. S., Storey, M., Vercesi, A. E., &
Docampo, R. (2018). Calcium-sensitive pyruvate dehydrogenase
phosphatase is required for energy metabolism, growth, differentiation,
131 and infectivity of Trypanosoma cruzi. J Biol Chem, 293(45), 17402-17417.
doi:10.1074/jbc.RA118.004498
Lanner, J. T., Georgiou, D. K., Joshi, A. D., & Hamilton, S. L. (2010). Ryanodine
receptors: structure, expression, molecular details, and function in calcium
release. Cold Spring Harb Perspect Biol, 2(11), a003996-a003996.
Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/20961976
https://www.ncbi.nlm.nih.gov/pmc/PMC2964179/.
doi:10.1101/cshperspect.a003996
Lawlis, V. B., & Roche, T. E. (1981). Regulation of bovine kidney .alpha.-
ketoglutarate dehydrogenase complex by calcium ion and adenine
nucleotides. Effects on S0.5 for .alpha.-ketoglutarate. Biochemistry, 20(9),
2512-2518. Retrieved from https://doi.org/10.1021/bi00512a023.
doi:10.1021/bi00512a023
Li, X.-B., Gu, J.-D., & Zhou, Q.-H. (2015). Review of aerobic glycolysis and its
key enzymes - new targets for lung cancer therapy. Thoracic cancer, 6(1),
17-24. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/26273330
https://www.ncbi.nlm.nih.gov/pmc/PMC4448463/. doi:10.1111/1759-
7714.12148
Li, Y., Park, J.-S., Deng, J.-H., & Bai, Y. (2006). Cytochrome c oxidase subunit IV
is essential for assembly and respiratory function of the enzyme complex.
Journal of bioenergetics and biomembranes, 38(5-6), 283-291. Retrieved
from https://www.ncbi.nlm.nih.gov/pubmed/17091399
132 https://www.ncbi.nlm.nih.gov/pmc/PMC1885940/. doi:10.1007/s10863-
006-9052-z
Liu, J. C., Liu, J., Holmstrom, K. M., Menazza, S., Parks, R. J., Fergusson, M. M.,
. . . Finkel, T. (2016). MICU1 Serves as a Molecular Gatekeeper to
Prevent In Vivo Mitochondrial Calcium Overload. Cell Rep, 16(6), 1561-
1573. doi:10.1016/j.celrep.2016.07.011
Lo, Y. Y., Wong, J. M., & Cruz, T. F. (1996). Reactive oxygen species mediate
cytokine activation of c-Jun NH2-terminal kinases. J Biol Chem, 271(26),
15703-15707.
Logan, C. V., Szabadkai, G., Sharpe, J. A., Parry, D. A., Torelli, S., Childs, A.-M.,
. . . Sheridan, E. (2013). Loss-of-function mutations in MICU1 cause a
brain and muscle disorder linked to primary alterations in mitochondrial
calcium signaling. Nature Genetics, 46, 188. Retrieved from
https://doi.org/10.1038/ng.2851. doi:10.1038/ng.2851
https://www.nature.com/articles/ng.2851#supplementary-information
Longo, N., Amat di San Filippo, C., & Pasquali, M. (2006). Disorders of carnitine
transport and the carnitine cycle. Am J Med Genet C Semin Med Genet,
142c(2), 77-85. doi:10.1002/ajmg.c.30087
Luongo, T. S., Lambert, J. P., Gross, P., Nwokedi, M., Lombardi, A. A.,
Shanmughapriya, S., . . . Elrod, J. W. (2017). The mitochondrial
Na(+)/Ca(2+) exchanger is essential for Ca(2+) homeostasis and viability.
Nature, 545(7652), 93-97. doi:10.1038/nature22082
133 Luongo, T. S., Lambert, J. P., Yuan, A., Zhang, X., Gross, P., Song, J., . . . Elrod,
J. W. (2015). The Mitochondrial Calcium Uniporter Matches Energetic
Supply with Cardiac Workload during Stress and Modulates Permeability
Transition. Cell Rep, 12(1), 23-34. doi:10.1016/j.celrep.2015.06.017
Lushchak, O. V., Piroddi, M., Galli, F., & Lushchak, V. I. (2014). Aconitase post-
translational modification as a key in linkage between Krebs cycle, iron
homeostasis, redox signaling, and metabolism of reactive oxygen species.
Redox Report, 19(1), 8-15. Retrieved from
https://doi.org/10.1179/1351000213Y.0000000073.
doi:10.1179/1351000213Y.0000000073
Lynch, M., & Conery, J. S. (2000). The Evolutionary Fate and Consequences of
Duplicate Genes. Science, 290(5494), 1151. Retrieved from
http://science.sciencemag.org/content/290/5494/1151.abstract.
doi:10.1126/science.290.5494.1151
Mailloux, R. J., Beriault, R., Lemire, J., Singh, R., Chenier, D. R., Hamel, R. D., &
Appanna, V. D. (2007). The tricarboxylic acid cycle, an ancient metabolic
network with a novel twist. PLoS One, 2(8), e690.
doi:10.1371/journal.pone.0000690
Maklashina, E., Sher, Y., Zhou, H.-Z., Gray, M. O., Karliner, J. S., & Cecchini, G.
(2002). Effect of anoxia/reperfusion on the reversible active/de-active
transition of NADH–ubiquinone oxidoreductase (complex I) in rat heart.
Biochimica et Biophysica Acta (BBA) - Bioenergetics, 1556(1), 6-12.
Retrieved from
134 http://www.sciencedirect.com/science/article/pii/S0005272802002803.
doi:https://doi.org/10.1016/S0005-2728(02)00280-3
Mallilankaraman, K., Cardenas, C., Doonan, P. J., Chandramoorthy, H. C., Irrinki,
K. M., Golenar, T., . . . Madesh, M. (2015). MCUR1 is an essential
component of mitochondrial Ca(2+) uptake that regulates cellular
metabolism. Nat Cell Biol, 17(7), 953. doi:10.1038/ncb3202
Mallilankaraman, K., Doonan, P., Cardenas, C., Chandramoorthy, H. C., Muller,
M., Miller, R., . . . Madesh, M. (2012). MICU1 is an essential gatekeeper
for MCU-mediated mitochondrial Ca(2+) uptake that regulates cell
survival. Cell, 151(3), 630-644. doi:10.1016/j.cell.2012.10.011
Marks, A. R. (2003). Calcium and the heart: a question of life and death. J Clin
Invest, 111(5), 597-600. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/12618512
https://www.ncbi.nlm.nih.gov/pmc/PMC151912/. doi:10.1172/JCI18067
Marshansky, V., Rubinstein, J. L., & Grüber, G. (2014). Eukaryotic V-ATPase:
Novel structural findings and functional insights. Biochimica et Biophysica
Acta (BBA) - Bioenergetics, 1837(6), 857-879. Retrieved from
http://www.sciencedirect.com/science/article/pii/S0005272814000280.
doi:https://doi.org/10.1016/j.bbabio.2014.01.018
McCormack, J. G., Daniel, R. L., Osbaldeston, N. J., Rutter, G. A., & Denton, R.
M. (1992). Mitochondrial Ca<sup>2+</sup> transport and the
role of matrix Ca<sup>2+</sup> in mammalian tissues.
Biochemical Society Transactions, 20(1), 153. Retrieved from
135 http://www.biochemsoctrans.org/content/20/1/153.abstract.
doi:10.1042/bst0200153
McCormack, J. G., & Denton, R. M. (1981). A comparative study of the regulation
of Ca2+ of the activities of the 2-oxoglutarate dehydrogenase complex and
NAD+-isocitrate dehydrogenase from a variety of sources. Biochem J,
196(2), 619-624.
Minarik, P., Tomaskova, N., Kollarova, M., & Antalik, M. (2002). Malate
dehydrogenases--structure and function. Gen Physiol Biophys, 21(3), 257-
265.
Mitchell, P. (1966). Chemiosmotic coupling in oxidative and photosynthetic
phosphorylation. Biol Rev Camb Philos Soc, 41(3), 445-502.
Monassier, J. P. (2008). Reperfusion injury in acute myocardial infarction. From
bench to cath lab. Part I: Basic considerations. Arch Cardiovasc Dis,
101(7), 491-500. Retrieved from
http://www.sciencedirect.com/science/article/pii/S1875213608001058.
doi:https://doi.org/10.1016/j.acvd.2008.06.014
Mourier, A., & Larsson, N. G. (2011). Tracing the trail of protons through complex
I of the mitochondrial respiratory chain. PLoS Biol, 9(8), e1001129.
doi:10.1371/journal.pbio.1001129
Nakagawa, T., Shimizu, S., Watanabe, T., Yamaguchi, O., Otsu, K., Yamagata,
H., . . . Tsujimoto, Y. (2005). Cyclophilin D-dependent mitochondrial
permeability transition regulates some necrotic but not apoptotic cell
136 death. Nature, 434(7033), 652-658. Retrieved from
https://doi.org/10.1038/nature03317. doi:10.1038/nature03317
Nakayama, S., & Kretsinger, R. H. (1994). Evolution of the EF-hand family of
proteins. Annu Rev Biophys Biomol Struct, 23, 473-507.
doi:10.1146/annurev.bb.23.060194.002353
Nalefski, E. A., & Falke, J. J. (1996). The C2 domain calcium-binding motif:
structural and functional diversity. Protein Sci, 5(12), 2375-2390.
doi:10.1002/pro.5560051201
Nishizuka, Y. (1988). The molecular heterogeneity of protein kinase C and its
implications for cellular regulation. Nature, 334(6184), 661-665. Retrieved
from https://doi.org/10.1038/334661a0. doi:10.1038/334661a0
Øie, E., Bjørnerheim, R., Clausen, O. P. F., & Attramadal, H. (2000). Cyclosporin
A inhibits cardiac hypertrophy and enhances cardiac dysfunction during
postinfarction failure in rats. American Journal of Physiology-Heart and
Circulatory Physiology, 278(6), H2115-H2123. Retrieved from
https://doi.org/10.1152/ajpheart.2000.278.6.H2115.
doi:10.1152/ajpheart.2000.278.6.H2115
Oxenoid, K., Dong, Y., Cao, C., Cui, T., Sancak, Y., Markhard, A. L., . . . Chou, J.
J. (2016). Architecture of the mitochondrial calcium uniporter. Nature,
533(7602), 269-273. doi:10.1038/nature17656
Paillard, M., Csordas, G., Huang, K. T., Varnai, P., Joseph, S. K., & Hajnoczky,
G. (2018). MICU1 Interacts with the D-Ring of the MCU Pore to Control Its
137 Ca(2+) Flux and Sensitivity to Ru360. Mol Cell, 72(4), 778-785.e773.
doi:10.1016/j.molcel.2018.09.008
Paillard, M., Csordas, G., Szanda, G., Golenar, T., Debattisti, V., Bartok, A., . . .
Hajnoczky, G. (2017). Tissue-Specific Mitochondrial Decoding of
Cytoplasmic Ca(2+) Signals Is Controlled by the Stoichiometry of MICU1/2
and MCU. Cell Rep, 18(10), 2291-2300. doi:10.1016/j.celrep.2017.02.032
Pan, X., Liu, J., Nguyen, T., Liu, C., Sun, J., Teng, Y., . . . Finkel, T. (2013). The
physiological role of mitochondrial calcium revealed by mice lacking the
mitochondrial calcium uniporter. Nat Cell Biol, 15(12), 1464-1472.
doi:10.1038/ncb2868
Parmacek, M. S., & Solaro, R. J. (2004). Biology of the troponin complex in
cardiac myocytes. Progress in Cardiovascular Diseases, 47(3), 159-176.
Retrieved from
http://www.sciencedirect.com/science/article/pii/S0033062004000556.
doi:https://doi.org/10.1016/j.pcad.2004.07.003
Patel, M. S., Nemeria, N. S., Furey, W., & Jordan, F. (2014). The pyruvate
dehydrogenase complexes: structure-based function and regulation. J Biol
Chem, 289(24), 16615-16623. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/24798336
https://www.ncbi.nlm.nih.gov/pmc/PMC4059105/.
doi:10.1074/jbc.R114.563148
Patel, M. S., & Roche, T. E. (1990). Molecular biology and biochemistry of
pyruvate dehydrogenase complexes. Faseb j, 4(14), 3224-3233.
138 Patron, M., Checchetto, V., Raffaello, A., Teardo, E., Vecellio Reane, D.,
Mantoan, M., . . . Rizzuto, R. (2014). MICU1 and MICU2 finely tune the
mitochondrial Ca2+ uniporter by exerting opposite effects on MCU activity.
Mol Cell, 53(5), 726-737. doi:10.1016/j.molcel.2014.01.013
Patron, M., Granatiero, V., Espino, J., Rizzuto, R., & De Stefani, D. (2019).
MICU3 is a tissue-specific enhancer of mitochondrial calcium uptake. Cell
Death Differ, 26(1), 179-195. doi:10.1038/s41418-018-0113-8
Payne, R., Hoff, H., Roskowski, A., & Foskett, J. K. (2017). MICU2 Restricts
Spatial Crosstalk between InsP3R and MCU Channels by Regulating
Threshold and Gain of MICU1-Mediated Inhibition and Activation of MCU.
Cell Rep, 21(11), 3141-3154. doi:10.1016/j.celrep.2017.11.064
Perez-Reyes, E. (2003). Molecular Physiology of Low-Voltage-Activated T-type
Calcium Channels. Physiol Rev, 83(1), 117-161. Retrieved from
https://doi.org/10.1152/physrev.00018.2002.
doi:10.1152/physrev.00018.2002
Perez-Reyes, E., & Bers, D. M. (1999). Ca channels in cardiac myocytes:
structure and function in Ca influx and intracellular Ca release.
Cardiovascular Research, 42(2), 339-360. Retrieved from
https://doi.org/10.1016/S0008-6363(99)00038-3. doi:10.1016/S0008-
6363(99)00038-3
Perocchi, F., Gohil, V. M., Girgis, H. S., Bao, X. R., McCombs, J. E., Palmer, A.
E., & Mootha, V. K. (2010). MICU1 encodes a mitochondrial EF hand
139 protein required for Ca(2+) uptake. Nature, 467(7313), 291-296.
doi:10.1038/nature09358
Philipson, K. D., Nicoll, D. A., Ottolia, M., Quednau, B. D., Reuter, H., John, S., &
Qiu, Z. (2002). The Na+/Ca2+ exchange molecule: an overview. Ann N Y
Acad Sci, 976, 1-10.
Pinnell, J., Turner, S., & Howell, S. (2007). Cardiac muscle physiology.
Continuing Education in Anaesthesia Critical Care & Pain, 7(3), 85-88.
Retrieved from https://doi.org/10.1093/bjaceaccp/mkm013.
doi:10.1093/bjaceaccp/mkm013
Piot, C., Croisille, P., Staat, P., Thibault, H., Rioufol, G., Mewton, N., . . . Ovize,
M. (2008). Effect of Cyclosporine on Reperfusion Injury in Acute
Myocardial Infarction. New England Journal of Medicine, 359(5), 473-481.
Retrieved from https://doi.org/10.1056/NEJMoa071142.
doi:10.1056/NEJMoa071142
Plattner, H., & Verkhratsky, A. (2015). Evolution of calcium signalling. Cell
Calcium, 57(3), 121-122. doi:10.1016/j.ceca.2015.02.007
Plovanich, M., Bogorad, R. L., Sancak, Y., Kamer, K. J., Strittmatter, L., Li, A. A.,
. . . Mootha, V. K. (2013). MICU2, a paralog of MICU1, resides within the
mitochondrial uniporter complex to regulate calcium handling. PLoS One,
8(2), e55785. doi:10.1371/journal.pone.0055785
Powell, B., Graham, L. A., & Stevens, T. H. (2000). Molecular characterization of
the yeast vacuolar H+-ATPase proton pore. J Biol Chem, 275(31), 23654-
23660. doi:10.1074/jbc.M004440200
140 Raffaello, A., De Stefani, D., Sabbadin, D., Teardo, E., Merli, G., Picard, A., . . .
Rizzuto, R. (2013). The mitochondrial calcium uniporter is a multimer that
can include a dominant-negative pore-forming subunit. Embo j, 32(17),
2362-2376. doi:10.1038/emboj.2013.157
Rafieian-Kopaei, M., Setorki, M., Doudi, M., Baradaran, A., & Nasri, H. (2014).
Atherosclerosis: process, indicators, risk factors and new hopes.
International journal of preventive medicine, 5(8), 927-946. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/25489440
https://www.ncbi.nlm.nih.gov/pmc/PMC4258672/.
Rahman, F. A., Abdullah, S. S., Manan, W. Z. W. A., Tan, L. T.-H., Neoh, C.-F.,
Ming, L. C., . . . Khan, T. M. (2018). Efficacy and Safety of Cyclosporine in
Acute Myocardial Infarction: A Systematic Review and Meta-Analysis.
Frontiers in pharmacology, 9, 238-238. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/29970999
https://www.ncbi.nlm.nih.gov/pmc/PMC6018391/.
doi:10.3389/fphar.2018.00238
Ran, F. A., Hsu, P. D., Wright, J., Agarwala, V., Scott, D. A., & Zhang, F. (2013).
Genome engineering using the CRISPR-Cas9 system. Nat Protoc, 8(11),
2281-2308. doi:10.1038/nprot.2013.143
Rankin, E. B., Rha, J., Selak, M. A., Unger, T. L., Keith, B., Liu, Q., & Haase, V.
H. (2009). Hypoxia-inducible factor 2 regulates hepatic lipid metabolism.
Mol Cell Biol, 29(16), 4527-4538. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/19528226
141 https://www.ncbi.nlm.nih.gov/pmc/PMC2725738/.
doi:10.1128/MCB.00200-09
Ratan, R. R., Siddiq, A., Smirnova, N., Karpisheva, K., Haskew-Layton, R.,
McConoughey, S., . . . LaManna, J. (2007). Harnessing hypoxic
adaptation to prevent, treat, and repair stroke. J Mol Med (Berl), 85(12),
1331-1338. doi:10.1007/s00109-007-0283-1
Reed, L. J. (2001). A trail of research from lipoic acid to alpha-keto acid
dehydrogenase complexes. J Biol Chem, 276(42), 38329-38336.
doi:10.1074/jbc.R100026200
Rizzuto, R., & Pozzan, T. (2006). Microdomains of Intracellular Ca2+: Molecular
Determinants and Functional Consequences. Physiol Rev, 86(1), 369-408.
Retrieved from https://doi.org/10.1152/physrev.00004.2005.
doi:10.1152/physrev.00004.2005
Rottenberg, H., & Scarpa, A. (1974). Calcium uptake and membrane potential in
mitochondria. Biochemistry, 13(23), 4811-4817. Retrieved from
https://doi.org/10.1021/bi00720a020. doi:10.1021/bi00720a020
Rutter, G. A., & Denton, R. M. (1988). Regulation of NAD+-linked isocitrate
dehydrogenase and 2-oxoglutarate dehydrogenase by Ca2+ ions within
toluene-permeabilized rat heart mitochondria. Interactions with regulation
by adenine nucleotides and NADH/NAD+ ratios. Biochem J, 252(1), 181-
189.
142 Samtleben, S., Jaepel, J., Fecher, C., Andreska, T., Rehberg, M., & Blum, R.
(2013). Direct imaging of ER calcium with targeted-esterase induced dye
loading (TED). J Vis Exp(75), e50317. doi:10.3791/50317
Sancak, Y., Markhard, A. L., Kitami, T., Kovacs-Bogdan, E., Kamer, K. J.,
Udeshi, N. D., . . . Mootha, V. K. (2013). EMRE is an essential component
of the mitochondrial calcium uniporter complex. Science, 342(6164), 1379-
1382. doi:10.1126/science.1242993
Schönfeld, P., & Reiser, G. (2013). Why does brain metabolism not favor burning
of fatty acids to provide energy? Reflections on disadvantages of the use
of free fatty acids as fuel for brain. Journal of cerebral blood flow and
metabolism : official journal of the International Society of Cerebral Blood
Flow and Metabolism, 33(10), 1493-1499. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/23921897
https://www.ncbi.nlm.nih.gov/pmc/PMC3790936/.
doi:10.1038/jcbfm.2013.128
Seagroves, T. N., Ryan, H. E., Lu, H., Wouters, B. G., Knapp, M., Thibault, P., . .
. Johnson, R. S. (2001). Transcription factor HIF-1 is a necessary
mediator of the pasteur effect in mammalian cells. Mol Cell Biol, 21(10),
3436-3444. doi:10.1128/mcb.21.10.3436-3444.2001
Sedaghat, F., & Notopoulos, A. (2008). S100 protein family and its application in
clinical practice. Hippokratia, 12(4), 198-204. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/19158963
https://www.ncbi.nlm.nih.gov/pmc/PMC2580040/.
143 Shamseldin, H. E., Alasmari, A., Salih, M. A., Samman, M. M., Mian, S. A.,
Alshidi, T., . . . Alkuraya, F. S. (2017). A null mutation in MICU2 causes
abnormal mitochondrial calcium homeostasis and a severe
neurodevelopmental disorder. Brain, 140(11), 2806-2813.
doi:10.1093/brain/awx237
Sharov, V. S., Dremina, E. S., Galeva, N. A., Williams, T. D., & Schoneich, C.
(2006). Quantitative mapping of oxidation-sensitive cysteine residues in
SERCA in vivo and in vitro by HPLC-electrospray-tandem MS: selective
protein oxidation during biological aging. Biochem J, 394(Pt 3), 605-615.
doi:10.1042/bj20051214
Shin, O. H., Han, W., Wang, Y., & Sudhof, T. C. (2005). Evolutionarily conserved
multiple C2 domain proteins with two transmembrane regions (MCTPs)
and unusual Ca2+ binding properties. J Biol Chem, 280(2), 1641-1651.
doi:10.1074/jbc.M407305200
Shohet, R. V., & Garcia, J. A. (2007). Keeping the engine primed: HIF factors as
key regulators of cardiac metabolism and angiogenesis during ischemia.
Journal of Molecular Medicine, 85(12), 1309-1315. Retrieved from
https://doi.org/10.1007/s00109-007-0279-x. doi:10.1007/s00109-007-
0279-x
Sohal, D. S., Nghiem, M., Crackower, M. A., Witt, S. A., Kimball, T. R., Tymitz, K.
M., . . . Molkentin, J. D. (2001). Temporally regulated and tissue-specific
gene manipulations in the adult and embryonic heart using a tamoxifen-
inducible Cre protein. Circ Res, 89(1), 20-25.
144 Song, L., Alcalai, R., Arad, M., Wolf, C. M., Toka, O., Conner, D. A., . . .
Seidman, J. G. (2007). Calsequestrin 2 (CASQ2) mutations increase
expression of calreticulin and ryanodine receptors, causing
catecholaminergic polymorphic ventricular tachycardia. J Clin Invest,
117(7), 1814-1823. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/17607358
https://www.ncbi.nlm.nih.gov/pmc/PMC1904315/. doi:10.1172/JCI31080
Soria, P. S., McGary, K. L., & Rokas, A. (2014). Functional divergence for every
paralog. Mol Biol Evol, 31(4), 984-992. doi:10.1093/molbev/msu050
Stammers, A. N., Susser, S. E., Hamm, N. C., Hlynsky, M. W., Kimber, D. E.,
Kehler, D. S., & Duhamel, T. A. (2015). The regulation of
sarco(endo)plasmic reticulum calcium-ATPases (SERCA). Can J Physiol
Pharmacol, 93(10), 843-854. doi:10.1139/cjpp-2014-0463
Sutton Martin, G. S. J., & Sharpe, N. (2000). Left Ventricular Remodeling After
Myocardial Infarction. Circulation, 101(25), 2981-2988. Retrieved from
https://doi.org/10.1161/01.CIR.101.25.2981.
doi:10.1161/01.CIR.101.25.2981
Tabarean, I. V. (2016). Histamine receptor signaling in energy homeostasis.
Neuropharmacology, 106, 13-19. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/26107117
https://www.ncbi.nlm.nih.gov/pmc/PMC4686377/.
doi:10.1016/j.neuropharm.2015.04.011
145 Taegtmeyer, H., Young, M. E., Lopaschuk, G. D., Abel, E. D., Brunengraber, H.,
Darley-Usmar, V., . . . Wang, T. J. (2016). Assessing Cardiac Metabolism:
A Scientific Statement From the American Heart Association. Circ Res,
118(10), 1659-1701. doi:10.1161/res.0000000000000097
Tang, W., Blair, C. A., Walton, S. D., Málnási-Csizmadia, A., Campbell, K. S., &
Yengo, C. M. (2017). Modulating Beta-Cardiac Myosin Function at the
Molecular and Tissue Levels. Frontiers in physiology, 7, 659. Retrieved
from https://www.frontiersin.org/article/10.3389/fphys.2016.00659.
Tarasov, A. I., Griffiths, E. J., & Rutter, G. A. (2012). Regulation of ATP
production by mitochondrial Ca(2+). Cell Calcium, 52(1), 28-35. Retrieved
from https://www.ncbi.nlm.nih.gov/pubmed/22502861
https://www.ncbi.nlm.nih.gov/pmc/PMC3396849/.
doi:10.1016/j.ceca.2012.03.003
Temple, I. P., Inada, S., Dobrzynski, H., & Boyett, M. R. (2013). Connexins and
the atrioventricular node. Heart rhythm, 10(2), 297-304. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/23085482
https://www.ncbi.nlm.nih.gov/pmc/PMC3572393/.
doi:10.1016/j.hrthm.2012.10.020
Territo, P. R., Mootha, V. K., French, S. A., & Balaban, R. S. (2000). Ca(2+)
activation of heart mitochondrial oxidative phosphorylation: role of the
F(0)/F(1)-ATPase. Am J Physiol Cell Physiol, 278(2), C423-435.
doi:10.1152/ajpcell.2000.278.2.C423
146 Tomar, D., Dong, Z., Shanmughapriya, S., Koch, D. A., Thomas, T., Hoffman, N.
E., . . . Madesh, M. (2016). MCUR1 Is a Scaffold Factor for the MCU
Complex Function and Promotes Mitochondrial Bioenergetics. Cell Rep,
15(8), 1673-1685. doi:10.1016/j.celrep.2016.04.050
Tsai, M. F., Phillips, C. B., Ranaghan, M., Tsai, C. W., Wu, Y., Willliams, C., &
Miller, C. (2016). Dual functions of a small regulatory subunit in the
mitochondrial calcium uniporter complex. Elife, 5. doi:10.7554/eLife.15545
Vais, H., Mallilankaraman, K., Mak, D. D., Hoff, H., Payne, R., Tanis, J. E., &
Foskett, J. K. (2016). EMRE Is a Matrix Ca(2+) Sensor that Governs
Gatekeeping of the Mitochondrial Ca(2+) Uniporter. Cell Rep, 14(3), 403-
410. doi:10.1016/j.celrep.2015.12.054
Valderrábano, M. (2007). Influence of anisotropic conduction properties in the
propagation of the cardiac action potential. Progress in biophysics and
molecular biology, 94(1-2), 144-168. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/17482242
https://www.ncbi.nlm.nih.gov/pmc/PMC1995420/.
doi:10.1016/j.pbiomolbio.2007.03.014
Vasington, F. D., Gazzotti, P., Tiozzo, R., & Carafoli, E. (1972). The effect of
ruthenium red on Ca2+ transport and respiration in rat liver mitochondria.
Biochimica et Biophysica Acta (BBA) - Bioenergetics, 256(1), 43-54.
Retrieved from
http://www.sciencedirect.com/science/article/pii/0005272872901612.
doi:https://doi.org/10.1016/0005-2728(72)90161-2
147 Vatrinet, R., Leone, G., De Luise, M., Girolimetti, G., Vidone, M., Gasparre, G., &
Porcelli, A. M. (2017). The α-ketoglutarate dehydrogenase complex in
cancer metabolic plasticity. Cancer & Metabolism, 5(1), 3. Retrieved from
https://doi.org/10.1186/s40170-017-0165-0. doi:10.1186/s40170-017-
0165-0
Vecellio Reane, D., Vallese, F., Checchetto, V., Acquasaliente, L., Butera, G., De
Filippis, V., . . . Raffaello, A. (2016). A MICU1 Splice Variant Confers High
Sensitivity to the Mitochondrial Ca2+ Uptake Machinery of Skeletal
Muscle. Mol Cell, 64(4), 760-773. Retrieved from
http://www.sciencedirect.com/science/article/pii/S1097276516306232.
doi:https://doi.org/10.1016/j.molcel.2016.10.001
Vos, T., Abajobir, A. A., Abate, K. H., Abbafati, C., Abbas, K. M., Abd-Allah, F., . .
. Murray, C. J. L. (2017). Global, regional, and national incidence,
prevalence, and years lived with disability for 328 diseases and injuries for
195 countries, 1990–2016: a systematic analysis for the Global
Burden of Disease Study 2016. The Lancet, 390(10100), 1211-1259.
Retrieved from https://doi.org/10.1016/S0140-6736(17)32154-2.
doi:10.1016/S0140-6736(17)32154-2
Wang, L., Yang, X., Li, S., Wang, Z., Liu, Y., Feng, J., . . . Shen, Y. (2014).
Structural and mechanistic insights into MICU1 regulation of mitochondrial
calcium uptake. Embo j, 33(6), 594-604. doi:10.1002/embj.201386523
Wang, T., Pentyala, S., Elliott, J. T., Dowal, L., Gupta, E., Rebecchi, M. J., &
Scarlata, S. (1999). Selective interaction of the C2 domains of
148 phospholipase C-beta1 and -beta2 with activated Galphaq subunits: an
alternative function for C2-signaling modules. Proc Natl Acad Sci U S A,
96(14), 7843-7846. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/10393909
https://www.ncbi.nlm.nih.gov/pmc/PMC22149/.
Weidemann, A., & Johnson, R. S. (2008). Biology of HIF-1α. Cell Death Differ,
15, 621. Retrieved from https://doi.org/10.1038/cdd.2008.12.
doi:10.1038/cdd.2008.12
Whitaker, M. (2006). Calcium at fertilization and in early development. Physiol
Rev, 86(1), 25-88. doi:10.1152/physrev.00023.2005
Wiegand, G., & Remington, S. J. (1986). Citrate synthase: structure, control, and
mechanism. Annu Rev Biophys Biophys Chem, 15, 97-117.
doi:10.1146/annurev.bb.15.060186.000525
Wierenga, R. K., Kapetaniou, E. G., & Venkatesan, R. (2010). Triosephosphate
isomerase: a highly evolved biocatalyst. Cell Mol Life Sci, 67(23), 3961-
3982. doi:10.1007/s00018-010-0473-9
Williams, R. J. P. (2006). The evolution of calcium biochemistry. Biochimica et
Biophysica Acta (BBA) - Molecular Cell Research, 1763(11), 1139-1146.
Retrieved from
http://www.sciencedirect.com/science/article/pii/S0167488906002540.
doi:https://doi.org/10.1016/j.bbamcr.2006.08.042
Xu, R., Han, M., Xu, Y., Zhang, X., Zhang, C., Zhang, D., . . . Wang, J. (2017).
Coiled-coil domain containing 109B is a HIF1alpha-regulated gene critical
149 for progression of human gliomas. J Transl Med, 15(1), 165.
doi:10.1186/s12967-017-1266-9
Yáñez, M., Gil-Longo, J., & Campos-Toimil, M. (2012). Calcium Binding Proteins.
In M. S. Islam (Ed.), Calcium Signaling (pp. 461-482). Dordrecht: Springer
Netherlands.
Yang, J., Marden, J. J., Fan, C., Sanlioglu, S., Weiss, R. M., Ritchie, T. C., . . .
Engelhardt, J. F. (2003). Genetic redox preconditioning differentially
modulates AP-1 and NF kappa B responses following cardiac
ischemia/reperfusion injury and protects against necrosis and apoptosis.
Mol Ther, 7(3), 341-353.
Yogev, O., Yogev, O., Singer, E., Shaulian, E., Goldberg, M., Fox, T. D., & Pines,
O. (2010). Fumarase: a mitochondrial metabolic enzyme and a
cytosolic/nuclear component of the DNA damage response. PLoS Biol,
8(3), e1000328. doi:10.1371/journal.pbio.1000328
Yoo, J., Wu, M., Yin, Y., Herzik, M. A., Jr., Lander, G. C., & Lee, S. Y. (2018).
Cryo-EM structure of a mitochondrial calcium uniporter. Science,
361(6401), 506-511. doi:10.1126/science.aar4056
Yu, H. (1999). Extending the size limit of protein nuclear magnetic resonance.
Proc Natl Acad Sci U S A, 96(2), 332-334. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/9892632
https://www.ncbi.nlm.nih.gov/pmc/PMC33545/.
Zhao, Q., Wang, S., Li, Y., Wang, P., Li, S., Guo, Y., & Yao, R. (2013). The role
of the mitochondrial calcium uniporter in cerebral ischemia/reperfusion
150 injury in rats involves regulation of mitochondrial energy metabolism. Mol
Med Rep, 7(4), 1073-1080. doi:10.3892/mmr.2013.1321
Zhou, Y. Y., Wang, S. Q., Zhu, W. Z., Chruscinski, A., Kobilka, B. K., Ziman, B., .
. . Xiao, R. P. (2000). Culture and adenoviral infection of adult mouse
cardiac myocytes: methods for cellular genetic physiology. Am J Physiol
Heart Circ Physiol, 279(1), H429-436.
doi:10.1152/ajpheart.2000.279.1.H429
151