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A reduction in cardiac function precedes structural adaptations in experimental spinal cord injury Fossey, Mary Pauline Mona 2019

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			A	REDUCTION	IN	CARDIAC	FUNCTION	PRECEDES	STRUCTURAL	ADAPTATIONS	IN	EXPERIMENTAL	SPINAL	CORD	INJURY	by	Mary	Pauline	Mona	Fossey	B.Sc.,	The	University	of	British	Columbia,	2017		A	THESIS	SUBMITTED	IN	PARTIAL	FULFILLMENT	OF		THE	REQUIREMENTS	FOR	THE	DEGREE	OF		MASTER	OF	SCIENCE	in	THE	FACULTY	OF	GRADUATE	AND	POSTDOCTORAL	STUDIES	(Kinesiology)		THE	UNIVERSITY	OF	BRITISH	COLUMBIA	(Vancouver)		December,	2019	©	Mary	Pauline	Mona	Fossey,	2019			 					ii	The	following	individuals	certify	that	they	have	read,	and	recommend	to	the	Faculty	of	Graduate	and	Postdoctoral	Studies	for	acceptance,	a	thesis	entitled:		A	reduction	in	cardiac	function	precedes	structural	adaptations	in	experimental	spinal	cord	injury			submitted	by	 Mary	Pauline	Mona	Fossey	 in	partial	fulfillment	of	the	requirements	for	the	degree	of		 Master	of	Science	 	in	 Kinesiology	 			Examining	Committee:	Christopher	R.	West,	Department	of	Cellular	and	Physiology	Sciences,	Faculty	of	Medicine,	UBCO	Supervisor		Matt	S.	Ramer,	Department	of	Zoology,	Faculty	of	Science,	UBC	Supervisory	Committee	Member		David	J.	Granville,	Department	of	Pathology	and	Laboratory	Medicine,	Faculty	of	Medicine,	UBC	Supervisory	Committee	Member				 					iii	Abstract	High-level	spinal	cord	injury	(SCI)	causes	the	loss	of	descending	sympathetic	control	to	the	heart	which,	in	addition	to	other	secondary	consequences	(i.e.,	changes	in	physical	activity	and	metabolism),	leads	to	premature	onset	and	increased	risk	for	cardiovascular	disease.	Our	research	team	reported	that	chronic	high-level	experimental	SCI	is	associated	with	systolic	dysfunction,	cardiomyocyte	atrophy	and	up-regulation	of	the	two	main	proteolytic	pathways	in	cardiac	tissue.	How	such	events	manifest	over	time	post-injury	is	presently	unknown.	Therefore,	the	aim	of	this	thesis	was	to	investigate	the	temporal	effects	of	high-thoracic	SCI	on	cardiac	function,	structure	and	proteolysis.	To	achieve	so,	we	used	a	pre-clinical	rodent	model	which	underwent	complete	transection	SCI	at	the	third	thoracic	spinal	level	(T3-SCI).	Rats	were	terminated	at	different	time-points	along	the	acute	timeline:	12	hours,	1	day,	3	days,	5	days	and	7	days	post-SCI.	SHAM	rats	were	used	as	controls	and	underwent	dorsal	durotomy	with	no	SCI.	Echocardiography	was	performed	on	the	7-day	SCI	and	SHAM	groups	pre-surgery	and	on	days	1,	2,	4	and	6	post-surgery	to	assess	temporal	changes	in	cardiac	volumes	and	function.	At	termination	time-points,	left-ventricle	(LV)	catheterization	was	performed	to	assess	cardiac	function	in	all	groups	except	in	the	12-hour	T3-SCI	group.	Additionally,	cardiac	tissue	was	collected	for	histological	and	gene	expression	analysis	to	quantify	cardiomyocyte	dimensions	and	the	regulation	of	proteolytic	pathways,	respectively.	We	found	a	significant	reduction	in	load-dependent	and	-independent	systolic	function	with	ventricular-arterial	uncoupling	as	early	as	1	day	post-SCI	which	persisted	into	the	chronic	setting,	but	no	changes	in	diastolic	function.	These	results	indicate	a	rapid	onset	of	cardiac	dysfunction	following	T3-SCI,	implying	that	loss	of	cardiac	sympathetic	control	and	cardiac	unloading	are	key	determinants	in	reduced	systolic	performance	post-SCI.	Furthermore,	in	T3-SCI	cardiac	tissue,	we	report	elevated	gene	expression	of	targets	involved	with	the	ubiquitin	proteasome	system,	one	of	the	two	main	proteolytic	pathways.	Although	no	significant					iv	cardiomyocyte	atrophy	was	observed,	our	results	suggest	that	the	molecular	events	ultimately	causing	chronic	cardiac	atrophy	are	initiated	acutely	post-SCI.	Together,	our	findings	imply	that	reduced	cardiac	function	precedes	structural	remodelling	following	high-thoracic	SCI.			 					v	Lay	summary	Following	high-level	spinal	cord	injury,	the	brain	signals	sent	down	the	spinal	cord	can	no	longer	reach	the	heart.	This	lack	of	signalling	negatively	impacts	heart	function,	reduces	heart	size	and	induces	a	wide	range	of	complications,	which	can	lead	to	increased	risk	for,	and	premature	onset	of,	heart	disease.	Multiple	studies	have	reported	impairments	in	heart	function	and	structure	with	associated	protein	breakdown	following	chronic	spinal	cord	injury.	However,	the	sequence	and	timeline	of	these	changes	are	unknown.	To	study	the	changes	in	the	heart	across	time	after	spinal	cord	injury,	we	collected	heart	data	at	different	time-points	using	a	rodent	model.	We	report	that,	acutely	following	spinal	cord	injury,	heart	function	was	decreased,	markers	of	protein	breakdown	were	increased	but	there	were	no	changes	in	heart	structure	within	the	first	week,	implying	that	the	reduction	in	heart	function	occurs	before	the	reduction	in	heart	size.		 					vi	Preface		 All	experimental	protocols	conducted	for	this	thesis	were	reviewed	and	approved	by	the	University	of	British	Columbia	(UBC)	Animal	Care	Committee	(A18-0344)	and	strictly	followed	the	guidelines	implemented	by	the	Canadian	Council	for	Animal	Care.	All	data	was	collected	and	analyzed	by	Mary	P.M.	Fossey	and	members	of	the	West	lab	at	International	Collaboration	on	Repair	Discoveries	(ICORD).	No	data	from	this	thesis	have	been	previously	published.		 I	was	the	lead	investigator	for	this	project.	My	responsibilities	included	concept	development,	animal	care,	surgical	assistance,	in	vivo	and	ex	vivo	data	collection,	tissue	collection,	analysis	of	physiological	and	molecular	data,	interpretation	of	results	and	writing	of	this	manuscript.	Dr.	Malihe-Sadat	Poormasjedi-Meibod	was	involved	in	concept	development,	performed	all	animal	terminal	procedures	(echocardiography	and	catheterization)	and	trained	me	on	all	molecular	techniques.	Erin	Erskine	was	involved	in	the	organization	of	this	project,	performed	all	SHAM	and	spinal	surgeries	and	further	trained	me	on	all	animal	care	procedures.	Brian	Hayes	aided	with	animal	care	and	surgical	assistance,	and	partly	trained	me	on	analysing	physiological	data.	Dr.	Matt	S.	Ramer	and	Dr.	David	J.	Granville	provided	insight	for	concept	development,	thesis	revisions	and	additional	expertise.		 Dr.	Christopher	R.	West	was	the	supervisory	author	for	this	project.	Dr.	West	was	involved	with	concept	formation	and	development,	trained	me	on	analysing	physiological	data,	aided	with	data	collection,	analysis	and	interpretation,	and	provided	thesis	revisions.			 					vii	Table	of	contents		Abstract	.............................................................................................................................................................	iii	Lay	summary	.....................................................................................................................................................	v	Preface	..............................................................................................................................................................	vi	Table	of	contents	.............................................................................................................................................	vii	List	of	tables	......................................................................................................................................................	xi	List	of	figures	....................................................................................................................................................	xii	List	of	abbreviations	.......................................................................................................................................	xiii	Acknowledgments	..........................................................................................................................................	xvi	Dedication	......................................................................................................................................................	xvii		 Literature	review	......................................................................................................................	1	1.1	 Cardiac	anatomy	...............................................................................................................................	1	1.1.1	 Gross	anatomy	of	the	cardiovascular	system	..........................................................................	1	1.1.2	 Layers	of	cardiac	tissue	............................................................................................................	2	1.1.3	 Cardiac	muscle	.........................................................................................................................	5	1.2	 The	innervation	of	the	heart,	its	conducting	system	and	the	cardiac	cycle	...................................	7	1.2.1	 Autonomic	nervous	system	.....................................................................................................	7	1.2.2	 Rhythmic	contractions	...........................................................................................................	11	1.2.3	 The	microscopic	conducting	system	......................................................................................	11	1.2.4	 Intercalated	discs	...................................................................................................................	12	1.2.5	 Generation	of	contraction	-	Summary	...................................................................................	13	1.2.6	 The	cardiac	cycle	....................................................................................................................	13	1.3	 The	renin-angiotensin-aldosterone	system	&	the	heart	...............................................................	14	1.4	 Cardiac	muscle	homeostasis	..........................................................................................................	16	1.4.1	 Cardiac	plasticity	....................................................................................................................	16	1.4.2	 Regeneration	of	cardiac	muscle	............................................................................................	16	1.4.3	 Hypertrophy	&	atrophy	..........................................................................................................	16	1.5	 Cardiac	consequences	of	SCI	.........................................................................................................	18	1.5.1	 Systolic	and	diastolic	function	following	SCI	..........................................................................	19	1.5.2	 Cardiac	and	cardiomyocyte	structure	following	SCI	.............................................................	23	1.5.3	 The	ubiquitin-proteasome	system	.........................................................................................	24	1.5.4	 Autophagy	..............................................................................................................................	28	1.5.5	 The	ubiquitin-proteasome	system	and	autophagy	following	SCI	.........................................	30	1.5.6	 Temporal	regulation	of	remodelling	pathways	in	atrophy	...................................................	31	1.6	 Closing	remarks	..............................................................................................................................	33		 Aims	and	hypotheses	.............................................................................................................	34		 Materials	and	methods	..........................................................................................................	36					viii	3.1	 Overview	.........................................................................................................................................	36	3.2	 Ethics	and	disclaimer	......................................................................................................................	37	3.3	 Animals	...........................................................................................................................................	37	3.4	 Groups	and	termination	time-points	.............................................................................................	38	3.5	 Issues	with	internal	validity	............................................................................................................	40	3.6	 Pre-surgery	animal	care	.................................................................................................................	40	3.6.1	 Housing	...................................................................................................................................	40	3.6.2	 Nutrition	and	hydration	.........................................................................................................	40	3.6.3	 Antibiotics	...............................................................................................................................	41	3.7	 Pre-surgery	data	collection:	echocardiography	(for	T3-SCI	7	days	and	SHAM	groups)	...............	41	3.8	 T3-SCI	and	SHAM	surgeries	............................................................................................................	42	3.8.1	 Anesthesia	and	preparation	...................................................................................................	42	3.8.2	 Dorsal	durotomy	(for	both	SHAM	and	T3-SCI	animals)	.........................................................	43	3.8.3	 Spinal	cord	injury	(only	for	T3-SCI	animals)	...........................................................................	43	3.8.4	 Suturing	..................................................................................................................................	44	3.9	 Post-surgery	animal	care	................................................................................................................	45	3.9.1	 Immediately	post-operation	..................................................................................................	45	3.9.2	 Continued	antibiotics	&	pain	management	...........................................................................	45	3.9.3	 Housing,	nutrition	and	hydration	..........................................................................................	45	3.9.4	 Bladder	care	...........................................................................................................................	46	3.9.5	 Health	assessments	................................................................................................................	46	3.10	 Termination	day	data	collection:	in	vivo	outcomes	.......................................................................	47	3.10.1	 Anesthesia	..............................................................................................................................	47	3.10.2	 Catheterization	.......................................................................................................................	47	3.11	 Euthanasia	......................................................................................................................................	49	3.11.1	 Surgical	endpoints	..................................................................................................................	49	3.11.2	 Non-surgical	endpoints	..........................................................................................................	50	3.12	 Collection	of	tissues	.......................................................................................................................	50	3.12.1	 Cardiac	tissue	post-mortem	...................................................................................................	50	3.12.2	 Femur	post-mortem	...............................................................................................................	51	3.13	 Genetic	analysis	..............................................................................................................................	51	3.13.1	 RNA	expression	–	mRNA	extraction,	cDNA	synthesis	and	PCR	.............................................	51	3.13.2	 UPS	and	autophagy	targets	investigated	...............................................................................	52	3.14	 Histology	.........................................................................................................................................	52	3.14.1	 Staining	for	cardiomyocyte	morphology	...............................................................................	52	3.14.2	 Imaging	...................................................................................................................................	54	3.14.3	 Analysis	...................................................................................................................................	54	3.15	 Statistics	..........................................................................................................................................	55	3.16	 Exclusion	of	data	and	standardization	...........................................................................................	56					ix		 Results	....................................................................................................................................	58	4.1	 Demographics	.................................................................................................................................	58	4.1.1	 Demographics	at	termination	................................................................................................	58	4.1.2	 Body	mass	along	the	acute	timeline	in	the	7	days	T3-SCI	and	SHAM	groups	......................	58	4.2	 In-vivo	echocardiography	data	–	Temporal	cardiac	volumetric	and	functional	indices	following	the	7	days	T3-SCI	and	SHAM	groups	..........................................................................................................	59	4.2.1	 Heart	rate	...............................................................................................................................	59	4.2.2	 Volumetric	cardiac	indices	and	systolic	function	..................................................................	59	4.3	 In-vivo	catheterization	data	–	Cardiovascular	functional	and	pressure-volume	indices	..............	62	4.3.1	 Basal	arterial	hemodynamics	.................................................................................................	62	4.3.2	 Basal	cardiac	pressure-volume	responses	.............................................................................	64	4.4	 Molecular	data	–	Gene	analysis	for	protein	degradation	pathways	.............................................	68	4.4.1	 UPS	.........................................................................................................................................	70	4.4.2	 Autophagy	..............................................................................................................................	71	4.5	 Histological	data	–	Cardiomyocyte	dimensions	.............................................................................	72	4.5.1	 Cardiomyocyte	length	and	width...........................................................................................	73	4.5.2	 Cardiomyocyte	cross-sectional	area	and	volume	..................................................................	74		 Discussion	...............................................................................................................................	76	5.1	 Hemodynamics,	cardiac	volumes	and	function	.............................................................................	76	5.1.1	 Hemodynamics	were	negatively	affected	acutely	following	high-thoracic	SCI	....................	76	5.1.2	 Cardiac	volumes	were	reduced	acutely	following	high-thoracic	SCI	....................................	77	5.1.3	 Systolic	function	and	ventricular-vascular	coupling	were	impaired	at	the	first	acute	time-point	post-SCI	and	persisted	throughout	the	acute	setting	.................................................................	78	5.1.4	 There	was	no	strong	evidence	of	diastolic	dysfunction	acutely	post-SCI	.............................	81	5.2	 Protein	degradation	.......................................................................................................................	82	5.2.1	 UPS	gene	expression	was	up-regulated	in	the	early	stages	of	acute	SCI	.............................	82	5.2.2	 No	changes	in	the	autophagy	gene	expression	were	detected	acutely	post-SCI	.................	84	5.3	 Cardiac	structure	............................................................................................................................	85	5.3.1	 Histological	data	suggested	the	commencement	of	cardiomyocyte	atrophy	acutely	post-SCI	 85		 Conclusion	..............................................................................................................................	88	6.1	 Major	findings	................................................................................................................................	88	6.2	 Relevance	.......................................................................................................................................	89	6.2.1	 Implications	............................................................................................................................	89	6.2.2	 Why	should	we	care	about	cardiac	dysfunction	and	cardiac	atrophy	post-SCI?	.................	89	6.3	 Strengths,	limitations	and	considerations	.....................................................................................	90	6.3.1	 Strengths	................................................................................................................................	90					x	6.3.2	 Limitations	..............................................................................................................................	90	6.3.3	 Considerations	.......................................................................................................................	90	6.4	 Future	directions	............................................................................................................................	91	6.4.1	 Further	molecular	analyses	following	acute	SCI	....................................................................	91	6.4.2	 Sub-acute	time-points	following	SCI	......................................................................................	91	REFERENCES	....................................................................................................................................................	92				 	 					xi	List	of	tables		Table	3.1.	List	of	investigated	dependent	variables,	organized	per	method	................................................	39	Table	3.2.	Targets	investigated	......................................................................................................................	52	Table	4.1.	Group	demographics	at	termination	............................................................................................	58	Table	4.2.	Parasternal	long-axis	volumetric	and	functional	indices	following	SHAM	and	T3-SCI	along	the	acute	timeline	.................................................................................................................................................	61	Table	4.3.	Hemodynamic	responses	to	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	....	63	Table	4.4.	Cardiac	functional	responses	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	..............................................................................................................................................................	68	Table	4.6.	Quantitative	real-time	PCR	targets,	primers	and	fold	changes	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	........................................................................................................	69	Table	4.5.	LV	myocardial	cardiomyocyte	dimensions	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	.................................................................................................................................	72		 					xii	List	of	figures	Figure	1.1.	Schematic	of	the	autonomic	innervation	of	the	heart	................................................................	10	Figure	1.2.	Representative	pressure-volume	loop	at	resting	conditions	......................................................	14	Figure	1.4.	Molecular	pathways	of	angiotensin	II	production	.......................................................................	15	Figure	1.5.	Schematic	of	the	ubiquitin	proteasome	system	..........................................................................	26	Figure	1.6.	Pathways	involved	in	protein	degradation	and	synthesis	...........................................................	28	Figure	1.7.	A	schematic	of	macroautophagy	activation	for	purpose	to	degrade	proteins	in	bulk	...............	30	Figure	3.1.	Overview	of	methods	...................................................................................................................	37	Figure	3.2.	Representation	of	pressure-volume	data	obtained	via	left-ventricle	catheterization	and	inferior	vena	cava	occlusions	......................................................................................................................................	49	Figure	3.3.	Quadruple	immunofluorescent	stain	to	visualize	and	measure	cardiomyocyte	dimensions	....	55	Figure	4.1.	Body	mass	and	echocardiographic	indices	measured	along	the	acute	timeline	of	rats	that	have	undergone	SHAM	and	T3-SCI	surgeries	.........................................................................................................	60	Figure	4.2.	Basal	hemodynamics	following	SHAM	surgery	and	at	different	time-points	following	T3-SCI	..	63	Figure	4.3.	Averaged	baseline	pressure-volume	loops	and	representative	inferior	vena	cava	occlusions	obtained	via	LV	catheterization	......................................................................................................................	65	Figure	4.4.	Systolic	function	following	SHAM	surgery	and	at	different	times	points	following	T3-SCI	........	66	Figure	4.5.	Diastolic	function	following	SHAM	surgery	and	at	different	times	points	following	T3-SCI	......	67	Figure	4.8.	RNA	fold	changes	of	UPS	targets	following	SHAM	surgery	and	T3-SCI	at	different	time	points	along	the	acute	spectrum	(n=6)	.....................................................................................................................	70	Figure	4.9.	RNA	fold	changes	of	autophagy	targets	following	SHAM	surgery	and	T3-SCI	at	different	time	points	along	the	acute	spectrum	(n=6)	..........................................................................................................	71	Figure	4.6.	LV	myocardial	cardiomyocyte	length	and	width	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	.................................................................................................................................	73	Figure	4.7.	LV	myocardial	cardiomyocyte	cross-sectional	area	and	volume	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points	........................................................................................................	75	Figure	5.1.	Overview	of	findings	.....................................................................................................................	87		 					xiii	List	of	abbreviations	Ab	–	antibody	ACE	–	angiotensin	converting	enzyme	Ach	–	acetylcholine		Akt	–	protein	kinase	B	AMPK	–	AMP-activated	protein	kinase	ANGI	–	angiotensin	I	ANGII	–	angiotensin	II	ANOVA	–	analysis	of	variance		ANP	–	atrial	natriuretic	peptide	ANS	–	autonomic	nervous	system	Ao	–	angiotensinogen		AT1	–	angiotensin	II	receptor	1	AT2	–	angiotensin	II	receptor	2	ATG	–	autophagy	related	protein	ATG7	–	autophagy	related	7	ATG12	–	autophagy	related	12	ATG16L	–	ATG16-like	protein	AV	–	atrioventricular		BECN1	–	beclin	1	Bnip3	–	BCL2/adenovirus	E1B	19	kDa	protein-interacting	protein	3	BSA	–	bovine	serum	albumen	C#	–	cervical	segment	cDNA	–	complementary	DNA	CO	–	cardiac	output	CON	–	control		CSA	–	cross-sectional	area	CVD	–	cardiovascular	disease	DBP	–	diastolic	blood	pressure	dP/dtmax	–	maximal	rate	of	systolic	pressure	increment	-dP/dtmin	–	maximal	rate	of	diastolic	pressure	decrement	Ea	–	arterial	elastance	Ea/ESPVR	–	ventricular	vascular	coupling	ratio	EDPVR	–	end-diastolic	pressure-volume	relationship	EDV	–	end-diastolic	volume	Ees	–	end-systolic	elastance	EF	–	ejection	fraction	eIF3f	–	eukaryotic	translation	initiation	factor	3	subunit	f	ESPVR	–	end-systolic	pressure-volume	relationship	ESV	–	end-systolic	volume	FA	–	fascia	adherens	FIP200	–	FAK	family	kinase-interacting	protein	of	200	kDa	FoxO	–	forkhead	box	O						xiv	FoxO3	–	forkhead	box	O3		FS	–	fractional	shortening	HR	–	heart	rate	ICORD	–	International	Collaboration	on	Repair	Discoveries	IML	–	intermediolateral	nucleus		IVC	–	inferior	vena	cava		L#	–	lumbar	segment	LC3	–	microtubule-associated	protein	1	light	chain	3	LV	–	left-ventricle	MAFbx	–	muscle	atrophy	F-box	MAP	–	mean	arterial	pressure	MRI	–	magnetic	resonance	imaging	mRNA	–	messenger	RNA	mTOR	–	mammalian	target	of	rapamycin	MuRF1	–	muscle	RING-finger	protein	1	MyoD	–	myoblast	determination	protein	1	NDS	–	normal	donkey	serum	NE	–	norepinephrine		NF-kB	–	nuclear	factor	kappa	B	NT	–	neurotransmitter		NTS	–	nucleus	of	tractus	solitarius	PBS	–	phosphate-buffered	saline		PBS-T	–	phosphate-buffered	saline	with	Triton	Pdev	–	developed	pressure	Ped	–	end-diastolic	pressure	Pes	–	end-systolic	pressure	PI3K	–	phosphoinositide	3-kinase	PI3P	–	phosphatidylinositol	3-phosphate	Pmax	–	maximum	pressure	PRSW	–	preload	recruitable	stroke	work	PSLAX	–	parasternal	long	axis	PSNS	–	parasympathetic	nervous	system	PINK1-PRKN	–	PTEN-induced	kinase	1	&	Parkin	complex	PV	–	pressure-volume	p52	–	sporozoite	surface	protein	P36p	qPCR	-	qualitative	polymerase	chain	reaction	RAAS	–	renin-angiotensin-aldosterone	system		RBP	–	renin	binding	protein	RV	–	right	ventricle	RVLM	–	rostral	ventral	lateral	medulla	S#	–	sacral	segment	SA	–	sinoatrial	SBP	–	systolic	blood	pressure	SC	–	spinal	cord	SCI	–	spinal	cord	injury					xv	SE	–	standard	error	of	the	mean	SNS	–	sympathetic	nervous	system	SPN	–	sympathetic	preganglionic	neuron	SV	–	stroke	volume	SW	–	stroke	work	T#	–	thoracic	segment	TGF-b	–	transforming	growth	factor	beta	TNF	–	tumor	necrosis	factor	TNFa	–	tumor	necrosis	factor	a	TPR	–	total	peripheral	resistance	TRAF6	–	TNF	receptor	associated	factor	6	TRIM32	–	E3	ubiquitin-protein	ligase	TRIM32	TWEAK	–	TNF-like	weak	inducer	of	apoptosis	Ub	–	ubiquitin	UBC	–	the	University	of	British	Columbia	ULK1	–	serine/threonine-protein	kinase	ULK1		UPS	–	ubiquitin	proteasome	system	USP14	–	ubiquitin	carboxyl-terminal	hydrolase	14	USP19	–	ubiquitin	carboxyl-terminal	hydrolase	19	Vps	–	phosphatidylinositol	3-kinase	WGA	–	wheat	germ	agglutinin	6-OH-DOPA	–	6-hydroxy-dopamine			 					xvi	Acknowledgments		 First	of	all,	I	would	like	to	express	my	deepest	and	sincerest	gratitude	to	my	supervisor	Dr.	Christopher	R.	West	for	his	exceptional	mentorship,	clear	guidance	and	continuous	support.	Without	your	patience	and	expertise,	this	thesis	would	have	not	been	possible.	Thanks	to	you,	I	have	learned	so	many	skills	which	will	surely	last	for	the	entirety	of	my	scientific	career.	I	am	and	was	very	fortunate	to	have	had	you	as	a	mentor.	I	do	not	think	I	could	have	wished	for	a	better	supervisor.	Thank	you	for	this	opportunity	which	allowed	me	to	work	with	an	outstanding	team	on	a	project	which	has	further	expanded	my	passion	for	science.		Secondly,	I	would	like	to	express	special	thanks	to	Dr.	Malihe-Sadat	Poormasjedi-Meibod,	Brian	Hayes	and	Erin	Erskine	for	their	crucial	help.	Mali,	your	mentorship	and	passion	for	quality	science	has	immensely	guided	me	throughout	this	degree.	You	have	taught	me	so	much;	I	am	truly	grateful.	Brian	and	Erin,	your	friendship,	generous	help	and	support	throughout	this	project	were	invaluable	and	I	could	have	not	dreamed	of	better	teammates.		Thirdly,	I	have	profound	appreciation	for	my	committee	members	Dr.	Matt	S.	Ramer	and	Dr.	David	J.	Granville	for	their	expertise,	valuable	insight	and	contributions.	Next,	I	would	like	to	acknowledge	with	my	sincerest	thanks	all	members	of	my	lab,	the	West	lab	(particularly	Dr.	Alexandra	M.	Williams,	Dr.	Guillermo	A.	Alanis	and	Cameron	M.	Gee),	for	their	essential	assistance,	caring	encouragement	and	never-ending	positive	attitude.		Finally,	I	would	like	to	thank	ICORD	and	UBC	for	their	financial	assistance	and	high-quality	facilities.		 					xvii	Dedication	To	my	family.	 					1	 Literature	review	1.1 Cardiac	anatomy	1.1.1 	Gross	anatomy	of	the	cardiovascular	system			 The	cardiovascular	system	consists	of	three	main	components:	1)	a	fluid,	2)	a	pump	to	propel	the	fluid,	and	3)	an	extensive	network	of	vessels	to	transport	and	deliver	the	fluid	around	the	body.1,2	In	mammals,	the	heart	serves	as	the	muscular	pump	and	is	formed	of	four	chambers:	two	atria	and	two	ventricles.2,3	The	blood	functions	to	deliver	nutrients	and	oxygen,	collect	wastes	and	carbon	dioxide,	transport	communicating	molecules,	such	as	hormones	and	cytokines,	and	finally,	protect	all	tissues	with	a	myriad	of	different	immune	cells.2		1.1.1.1 Basic	structure	of	the	heart		The	heart	is	enveloped	by	the	pericardium,	which	is	a	fibrous	sheet	that	encloses	the	heart	in	a	liquid-producing	frictionless	chamber.2	The	pericardium	is	located	within	the	thoracic	cavity,	more	specifically	in	the	middle	mediastinum,	ventral	and	slightly	lateral	to	the	spinal	cord	(SC).2	The	heart	is	protected	by	the	ribs,	the	sternum,	the	vertebral	column	and	surrounding	adipose	tissue.2		The	heart	is	divided	into	two	sides	by	an	interatrial	and	interventricular	septum.2	The	right-side	of	the	heart	pumps	deoxygenated	blood	from	all	parts	of	the	body	to	the	lungs,	and	the	left-side	pumps	freshly	oxygenated	blood	from	the	lungs	to	the	rest	of	the	body.1	When	blood	enters	the	heart,	it	first	pools	into	an	atrium	then	gets	ejected	into	a	secondary	chamber	with	a	larger	lumen	and	thicker	muscular	wall,	a	ventricle.2	Both	sides	of	the	heart	contain	an	atrium	and	a	ventricle.1–3	Both	atria	and	both	ventricles	contract	in	synchrony	with	each	other,	respectively,	and	contract	to	produce	the	unidirectional	flow	of	blood.2					2	1.1.1.2 The	systemic	and	pulmonary	circuits	The	heart	pumps	blood	into	two	different	circuits:	the	systemic	and	the	pulmonary	circuits.1,2	Deoxygenated	blood	from	all	organ	systems	of	the	body	travels	back	to	the	right	side	of	the	heart	through	the	superior	vena	cava	(blood	flowing	from	the	head,	upper	limbs	and	chest	area)	and	inferior	vena	cava	(blood	flowing	from	the	abdomen,	pelvic	area	and	lower	limbs)	to	pool	into	the	right	atrium.2,3	Right	atrial	contraction	and	subsequent	ventricular	contraction	will	propel	venous	blood	to	continue	its	path	into	the	pulmonary	system.1,2	Blood	enters	the	pulmonary	trunk	which	later	divides	into	the	right	and	left	pulmonary	arteries	to	reach	each	respective	lung.2,3	In	the	capillary	beds	surrounding	lung	tissue,	the	blood	comes	into	close	contact	with	the	air	inhaled	in	the	alveoli.	As	the	blood	has	low	levels	of	oxygen	and	is	saturated	with	carbon	dioxide,	gaseous	exchange	of	both	gases	occurs	easily	by	diffusion	across	the	moist	one-cell-thick	barrier.3	Once	oxygenated,	blood	will	flow	back	to	the	left	side	of	the	heart	through	the	pulmonary	veins	and	enter	the	left	atrium.3	The	atrium	contracts	and	the	ventricle	consecutively;	this	sends	the	blood	into	the	systemic	circuit,	which	provides	oxygen	to	the	entire	body.1,3	As	the	left	side	of	the	heart	has	the	goal	to	replenish	all	tissues	from	the	body	with	freshly	oxygenated	blood,	the	muscle	wall	of	the	LV	is	thicker	than	the	right	(RV)	as	the	left-side	is	required	to	produce	more	pressure	to	propel	the	blood	at	all	extremities	of	the	body.1	Once	exchange	of	oxygen	has	occurred	in	the	systemic	capillaries,	the	blood	will	travel	back	to	the	right	side	of	the	heart	through	the	major	veins	and	exit	the	systemic	circuit.	One	important	aspect	to	take	note	of	is	that	both	sides	of	the	heart	contract	in	synchrony,	meaning	both	atria	will	contract	simultaneously,	as	will	the	ventricles	few	milliseconds	later.1	1.1.2 Layers	of	cardiac	tissue			 The	layers	of	the	heart	are	continuous	and	homologous	to	the	layers,	also	known	as	tunics,	of	blood	vessels.3	Comparable	to	the	vessel	tunics,	the	heart	has	three	layers	of	tissue:	the	endocardium,					3	homologous	to	the	tunica	intima,	the	cardiac	myocardium,	homologous	to	the	tunica	media,	and	the	epicardium,	homologous	to	the	tunica	adventitia.1–3	All	three	layers	are	present	in	the	free	walls	of	the	heart,	which	are	defined	to	be	the	walls	of	the	heart	which	are	not	in	contact	with	the	septa.2	However,	the	septa	between	the	chambers	do	not	contain	epicardium	but	rather	one	layer	of	myocardium	surrounded	by	two	layers	of	endocardium,	as	cardiac	lumen	are	found	on	each	side.2	There	exists	subtle	differences	in	layer	thickness	in	different	chambers,	for	example,	the	myocardium	located	in	the	atria	is	thinner	compared	to	the	one	in	the	ventricles,	this	difference	are	be	due	to	distinct	force	generation	requirements.2		1.1.2.1 Epicardium		 The	epicardium	is	the	most	outer	layer	of	the	heart	and	forms	part	of	the	visceral	layer	of	the	pericardium.3	The	pericardium	is	a	fibro-elastic	membranous	sac	surrounding	the	heart,	which	consists	with	two	membranous	layers:	the	parietal	and	visceral	layers,	a	pericardial	space	and	a	fibrous	outer	layer.1,3	The	serous	surfaces	secrete	a	lubricating	liquid	in	the	pericardial	cavity	to	reduce	friction	between	the	heart	with	its	surrounding	tissue	during	contraction.1–3	The	epicardium	consists	of	an	epithelial	layer	and	a	subepicardial	layer	with	loose	connective	tissue	containing	adipocytes,	serving	as	shock	absorbers,	and	infiltrating	coronary	blood	vessels,	lymph	vessels	and	nerve	supply	for	the	heart	itself.1,3		1.1.2.2 Endocardium		 The	endocardium,	the	inner	lining	of	the	heart,	has	direct	contact	with	the	blood	in	the	lumen	of	the	cardiac	chambers.1,3	The	apical	most	layer	consists	of	a	single	squamous	epithelium,	also	known	as	the	vascular	endothelium.1–3	Underneath	the	endothelium,	there	is	a	subendocardial	layer,	which	can	range	from	loose	to	dense	connective	tissue.1	The	subendocardium	mostly	contains	a	combination	of	scattered	fibroblast,	blood	and	nerve	supply,	smooth	muscle	cells	and	some	specialized	cardiac	muscle	cells,	which	form	fibers	of	the	conducting	system	of	the	heart	(i.e.,	Purkinje	fibers).1–3	As	the	tissue	grows					4	deeper	towards	and	transitions	to	myocardium,	the	connective	tissue	seems	to	accumulate	more	collagen	and	elastic	fibers.1–3	Ventricular	endocardium	has	trabeculae	which	creates	texture	to	the	lining.1	Papillary	muscles	are	expansions	of	trabeculae	into	the	lumen,	which	serve	as	attachments	for	chordae	tendineae,	important	for	valve	function.1		1.1.2.3 Myocardium	The	myocardium	is	the	middlemost	and	thickest	layer	of	the	cardiac	wall.1,3	It	mostly	consists	of	concentric	bundles	and	sheets	of	cardiac	muscle	cells,	also	known	as	cardiomyocytes,	which	surround,	in	a	circular	fashion,	all	chambers	of	the	heart.1,3	The	myocardium	is	a	complex	tissue	with	numerous	cardiomyocytes,	which	are	surrounded	by	an	extracellular	matrix	for	structural	integrity.4,5	The	matrix	around	the	muscle	cells	is	composed	of	fibroblasts,	intricate	blood	and	nerve	supply.4,5	As	the	myocardium	is	the	most	energy	demanding	tissue	in	the	entire	body,	it	is	richly	vascularized.6	It	has	been	found	that	each	cardiomyocyte	in	the	myocardium	has	direct	contact	with	at	least	one	capillary	vessel.6	Not	surprisingly,	the	thickness	of	the	myocardium	differs	between	chambers	due	to	their	function	and	their	target	circuit.1,2	While	the	ventricular	myocardium	is	thicker	than	the	atrial	myocardium,	the	left	ventricle	prevails	compared	to	the	right	ventricle	as	it	pumps	to	the	all	extremities	of	the	body.1		Cardiomyocytes	(cardiac	muscle	cells)	are	the	main	effectors	of	the	heart;	there	are	many	different	types	of	cardiomyocytes,	which	all	differ	slightly	in	their	function.3	Some	serve	to	contract	in	synchrony	to	generate	force	and	propel	the	blood	out	of	the	chamber,	others	control	the	generation	and	frequency	of	electrical	stimuli.3	The	sheets	of	cardiac	muscle	all	have	different	orientations	for	the	directionality	of	contraction	and	expulsion	of	blood.3	The	cellular	characteristics	of	cardiomyocytes	will	be	described	in	further	detail	when	discussing	the	microscopic	anatomy	of	cardiac	muscle.						5	1.1.3 Cardiac	muscle			 Cardiac	muscle	is	an	involuntary	striated	muscle,	which	serves	to	generate	mechanical	force	and	ensure	a	constant	heartbeat.1–3	The	functional	units	are	the	cardiomyocytes.1–3	Contractile	cardiomyocytes	are	branched	tubular	cells,	which	can	connect	to	multiple	neighbouring	muscle	cells	to	form	parallel	arrangements	of	cells,	called	laminae.2,3	These	laminae	have	distinct	directionality	depending	on	its	location	and	function	in	the	myocardium.2,3	As	cardiac	muscle	is	the	most	energy	demanding	tissue	in	the	entire	body,1	sheets	of	connective	tissue	lie	between	laminae	with	extensive	blood	and	nerve	supply	resulting	in	all	cardiomyocytes	to	be	in	close	proximity	to	capillaries	and	nerve	terminals.3	1.1.3.1 Cardiomyocytes	Cardiomyocytes	present	in	the	atria	are	smaller	than	their	ventricular	counterparts.3	The	different	morphologies	are	correlated	to	the	force	of	contraction	they	are	required	to	exert	to	propel	the	blood	to	its	next	destination.3	Even	within	one	lamina,	mature	cardiomyocytes	vary	in	shape	and	in	size	ranging	from	10-35	µm	in	diameter	and	80-120	µm	in	length.1,7,8	Cells	can	attach	to	multiple	different	neighbouring	cells	through	intercalated	discs;	this	branching	pattern	facilitates	the	propagation	of	electrical	impulses	throughout	the	tissue.1	Intracellularly	their	contents	can	differ	depending	on	the	species	of	interest,	and	the	location	in	the	heart.9	Despite	differences	in	organelle	quantity,	all	regular	cardiomyocytes	have	a	nucleus,	cytoskeleton	for	maintaining	cell	structure,	and	active	Golgi	and	endoplasmic	complexes.2	Although	the	common	cardiomyocyte	only	possesses	one	euchromatic	centered	nucleus,	few	can	be	doubly	nucleated.1	There	are	abundant	contractile	units,	myofibrils,	which	lie	parallel	to	the	direction	of	stimuli.2	Cardiomyocytes	contain	an	active	endoplasmic	reticulum,	specifically	called	sarcoplasmic	reticulum	and	a	plasma	membrane,	called	sarcolemma.1	Due	to	the	extensive	demand	for	ATP	for	contraction,	large	mitochondria	with	deep	cristae	(for	increased	surface					6	area)	are	present	in	large	numbers	in	cardiomyocytes	and	make	up	36%	of	the	cell,	compared	to	5%	in	skeletal	muscle	fibers.1,10	Most	of	the	mitochondria	can	be	located	densely	packed	near	the	nucleus	and	between	the	longitudinally	oriented	myofibrils.1,2,11,12	To	aid	the	high	energy	demand,	cardiomyocytes	additionally	store	large	quantities	of	glycogen	and	lipid	droplets,	which,	similar	to	mitochondria,	are	located	close	to	the	nucleus	and	myofibrils.2	These	serve	for	energy	back	up	in	case	of	low	oxygen	levels.1,2		1.1.3.2 Cardiomyocyte	contractile	apparatus,	T-tubules	and	the	sarcoplasmic	reticulum	Cardiomyocytes	have	similar	longitudinal	myofibrils	to	skeletal	muscle	cells.1,2	Cardiac	sarcomeres	are	made	up	of	the	same	proteins	as	skeletal	muscle:	alternating	thin	(~1.0	µm	long)	and	thick	filaments	(~1.6	µm	long),	predominantly	composed	of	actin	and	myosin,	respectively.2	All	filaments	are	anchored	to	the	Z-line	by	multiple	proteins,	including	a-actinin.13,14	The	contractile	proteins	make	up	more	than	55%	of	the	cytoplasm	in	cardiomyocytes.15,16	T-tubules	are	extensions	of	the	plasma	membrane	that	project	into	the	deeper	parts	of	a	muscle	cell	to	come	into	close	contact	with	the	contractile	apparatus	and	the	endoplasmic	reticulum.1,3	This	organized	arrangement	and	coming	together	of	membranes	are	located	in	close	proximity	to	Z-lines	and	serve	to	accelerate	depolarization-induced	calcium	release.1,3	As	the	electrical	stimuli	travel	down	the	t-tubule,	channels	in	the	plasma	membrane	open	to	allow	a	net	influx	of	calcium	ions	into	the	cytoplasm	of	the	cardiac	muscle	cell.3	Once	a	high	concentration	of	calcium	ions	is	detected	in	the	cytoplasm,	ryanodine	receptors	on	the	sarcoplasmic	membrane	will	open	to	generate	a	greater	influx	of	calcium	ion	into	the	cytoplasm.3	Thanks	to	both	of	these	ion	influxes,	the	contractile	apparatus	is	activated	causing	cells	to	contract,	generating	the	gross	contraction	of	the	cardiac	chamber.3							7	1.2 The	innervation	of	the	heart,	its	conducting	system	and	the	cardiac	cycle	1.2.1 Autonomic	nervous	system			 Cardiac	muscle	is	known	to	contract	rhythmically	and	spontaneously,	meaning	there	is	no	conscious	decision	or	need	for	innervation	to	stimulate	a	contraction	and	generate	a	heartbeat.1,2	It	has	been	shown	in	vitro,	that	embryonic	cardiac	muscle	cells	contract	spontaneously	with	no	direct	stimulation:	proving	that	heartbeat	is	spontaneous.2	However,	heartbeat	frequency	(chronotropism)	and	contractility	(ionotropism)	can	be	adjusted	by	the	autonomic	nervous	system	(ANS)	depending	on	the	metabolic	needs.2,17	In	addition,	the	ANS	can	change	the	conductivity	(dromotropism)	and	excitability	(bathmotropism)	of	the	cardiomyocytes,	and	the	vascular	tone	of	the	myocardial	capillaries	and	coronary	vessels.17	The	ANS	is	composed	of	complimentary	branches:	the	parasympathetic	(PSNS)	and	sympathetic	nervous	systems	(SNS),	which	act	in	opposite	ways	to	maintain	homeostasis.17,18	The	vagus	and	the	glossopharyngeal	cranial	nerves	relay	sensory	afferent	information	from	chemoreceptors,	baroreceptors,	mechanoreceptors	and	metaboreceptors	located	in	the	vasculature	and	the	sinuses	to	the	nucleus	of	tractus	solitarius	(NTS)	in	the	medulla	oblongata.19,20	1.2.1.1 Parasympathetic	nervous	system		 The	PSNS	is	activated	during	situations	of	“rest	and	digest”,	when	the	body	requires	blood	flow	directed	to	the	viscera	(splanchnic	area)	as	opposed	to	skeletal	muscle.17	Additionally,	the	PSNS	is	known	to	decrease	heart	rate	(HR)	and	decrease	contractility	of	the	heart.17	Cardiac	afferent	vagal	neurons	get	stimulated	by	stretch	receptors	in	the	carotid	sinuses	and	aortic	arch	when	blood	pressure	is	high,	to	decrease	in	HR	and	hypotensive	cardiac	reflexes.17,21,22	In	the	case	of	low	blood	pressure,	stretch	receptors	will	ultimately	cause	an	increase	in	HR	by	inhibiting	PSNS.17	Anatomical,	physiological	and	pharmacological	data	have	shown	that	the	PSNS	receives	input	from	cranial	and	sacral	nerves.17,23	Part	of	the	parasympathetic	outflow	can	occur	through	multiple	cranial					8	nerves	although	only	two	innervate	the	cardiovascular	system	(the	vagus	and	glossopharyngeal	nerve).17,19	The	rest	of	the	parasympathetic	outflow	is	located	in	the	sacral	area	(predominantly	S2-S3	compared	to	S1	and	S4)	to	innervate	the	urogenital	system	and	the	rectum.17		Parasympathetic	preganglionic	neurons,	originating	from	the	dorsal	motor	nucleus	of	the	vagus	and	the	nucleus	ambiguous	of	the	medulla	oblongata,	exit	the	vagal	nerve	and	innervate	many	organs	in	the	thorax	and	abdomen,	including	the	heart	(Figure	1.1.).17,19	Postsynaptic	nerve	terminals	from	both	branches	of	the	ANS	terminate	in	the	sinoatrial	(SA)	and	atrioventricular	(AV)	nodes	in	the	right	atrium	of	the	heart	(Figure	1.1.);	additionally,	some	branches	continue	to	run	along	the	coronary	arteries.2,19	At	the	target	organ,	for	example	the	heart,	the	parasympathetic	neurotransmitter	(NT)	released	to	decrease	HR	and	decrease	contractility	is	acetylcholine	(ACh).17	This	NT	binds	and	activates	nicotinic	(muscular	and	nervous)	and	muscarinic	receptors	(i.e.,	M1	to	M5)	on	the	surface	of	the	organ.17,24	Muscarinic	receptor	M2,	if	activated,	will	decrease	cardiac	contractility	and	conduction	velocity	mostly	in	the	atria.17	Although,	there	are	less	M2	receptors	in	the	ventricles,	if	the	stimuli	is	sufficiently	strong,	it	can	decrease	contractility	by	20%.17	M3	receptors	serve	to	mildly	dilate	the	coronary	vasculature.17			1.2.1.2 Sympathetic	nervous	system		 	The	SNS	will	be	activated	in	moments	of	“fight	or	flight”	when	cardiac	output	(CO;	volume	of	blood	pumped	out	of	the	heart	in	60	seconds)	must	be	increased,	for	example,	during	stress	or	exercise.17	To	increase	CO,	the	SNS	will	increase	the	frequency	of	depolarization	in	the	SA	node	located	in	the	right	atrium	to	increase	HR,	and	will	increase	the	frequency	of	myocardial	contraction	to	increase	stroke	volume	(SV;	volume	of	blood	pumped	out	of	the	heart	per	beat)	as	CO=SV*HR.17	Additional	excitatory	effects	of	the	SNS	include:	increased	conduction	velocity	and	decreased	refractory	period	in	the	nerves	innervating	the	heart.17						9		 Although	multiple	regions	of	the	brain	are	involved	in	sympathetic	input	(i.e.,	paraventricular	nucleus	in	the	hypothalamus,	the	rostral	ventromedial	medulla,	the	A5	region	of	the	brainstem	and	the	caudal	raphe	nuclei),	the	majority	of	the	sympathetic	premotor	neurons	which	innervate	the	heart	and	vessels	are	located	in	the	rostral	ventral	lateral	medulla	(RVLM)	of	the	medulla	oblongata.19	These	bulbospinal	premotor	sympathetic	neurons	originate	in	the	rostral	ventral	lateral	medulla	(RVLM)	in	the	brainstem	and	descend	the	SC	to	synapse	with	sympathetic	preganglionic	neuron	(SPN)	in	the	gray	commissure	around	the	central	canal,	in	the	intermediolateral	nucleus	region	of	the	grey	matter	(IML)	and	between	these	two	regions	in	segments	T1-L2	(Figure	1.1.).17,19,20	Nerves	innervating	the	cardiovascular	system	(heart	and	vessels)	and	the	heart	specifically	will	exit	at	T1-L2	and	T1-T5	(Figure	1.1.),	respectively.17	SPNs	will	enter	the	ventral	root,	follow	the	white	rami	communicantes	into	a	sympathetic	ganglion,	which	is	part	of	the	sympathetic	chain.17	SPNs	can	go	up	and	down	the	sympathetic	chain	to	reach	other	anterior	and	posterior	ganglia;	there	are	3	cervical,	11	thoracic,	4	lumbar	and	finally	4-5	sacral	ganglia.17	In	the	sympathetic	chain	ganglion,	the	SPN	will	release	ACh	to	stimulate	the	postganglionic	neuron.17,25	The	postganglionic	neuron	will	exit	the	sympathetic	chain	through	the	grey	rami	communicantes	to	then	innervate	the	target	organ,	where	it	will	release	norepinephrine	(NE)	to	activate	adrenergic	receptors.17	There	are	two	types	of	adrenergic	receptors:	a	(i.e.,	a1,	a2	and	subtypes)	and	b	(b1,	b2,	b3	and	subtypes)	adrenergic	receptors.17,26,27	In	terms	of	cardiac	functions,	a1	and	a2	mostly	constrict	coronary	vasculature,	whereas	b1	and	b2	(3:1	ratio)	induce	all	physiological	changes	previously	mentioned	in	this	paragraph	(i.e.,	increased	ionotropy,	dromotropy	and	bathmotropy).17	b2	is	found	to	be	distributed	primarily	in	all	chambers	of	the	heart,	while	the	presence	of	b3	in	the	heart	is	yet	to	be	clarified.17			 The	postganglionic	neurons	exit	the	ganglia	and	innervate	different	parts	of	the	heart	via	the	cardiac	nerves.17	There	are	eight	ganglia	on	each	side	of	the	heart:	superior,	middle	and	inferior	(stellate)					10	cervical	ganglia,	and	first	five	thoracic	ganglia	(Figure	1.1.).17–19	Sensory	nerve	endings	can	be	located	in	the	subendocardium,	and	at	conjunctions	of	different	vessels	and	chambers.17	These	will	sense	cardiac	sensory	information	which	will	be	sent	to	the	NTS	then	to	the	higher	sensory	processing	centers	located	in	the	diencephalon.17,28			Figure	1.1.	Schematic	of	the	autonomic	innervation	of	the	heart.	AV,	atrioventricular	node;	DMV,	dorsal	motor	nucleus	of	the	vagus;	IML,	intermediolateral	nucleus;	NA,	nucleus	ambiguous;	PSNS,	parasympathetic	nervous	system;	RVLM,	rostral	ventrolateral	medulla;	SA,	sinoatrial	node;	SNS,	sympathetic	nervous	system.17–20	1.2.1.3 The	cardiac	plexus		The	cardiac	plexus	is	a	network	of	nerves	which	serves	to	innervate	the	myocardium	of	the	interseptal	walls	(between	the	atria,	and	the	ventricles,	respectively),	the	two	nodes	of	the	conducting	system	and	some	vasculature,	with	both	the	PSNS	and	SNS.17	Due	to	its	range	of	activity,	it	is	located	at	the	base	of	the	heart,	ventral	to	the	carina	of	the	trachea	and	posterior	to	the	aortic	arch.17						11	1.2.2 Rhythmic	contractions		 A	heartbeat	consists	of	two	beats:	first,	the	atria	contract	in	synchrony	and	subsequently,	the	ventricles	do	the	same,	however	with	more	force.3	This	creates	a	rhythmical	flow	of	blood	through	the	heart.1,3	The	gross	muscular	contractions	are	generated	thanks	to	the	microscopic	contractions	of	thousands	of	cardiomyocytes	in	the	myocardium	of	these	chambers.3	Unlike	skeletal	muscle,	cardiac	muscle	requires	a	nearly	constant	release	fluctuations	of	calcium	ions	into	the	cytoplasm	and	constant	anaerobic	metabolism	to	sustain	contractions	and	relaxations.3	1.2.3 The	microscopic	conducting	system		 The	conducting	system	of	the	heart	is	made	up	of	specialized	cardiac	muscle	cells	which	have	the	ability	to	conduct	electrical	impulses	throughout	the	tissue.3	The	signals	sent	from	the	brain	descend	through	the	autonomic	nerves	transferring	the	stimuli	to	nodes,	located	in	the	right	atrial	wall.3	These	nodes	are	formed	by	the	aggregation	of	the	specialized	cardiomyocytes	abovementioned.2,3	The	SA	node,	also	known	as	the	cardiac	pacemaker	is	located	near	the	entry	of	the	major	venous	vessels.3	The	received	signals	generate	numerous	depolarization	events	within	the	cell,	which	will	result	in	a	HR	of	about	70	beats	per	minute	(bpm)	in	healthy	and	resting	human	conditions3	with	PSNS	dominance	(autonomic	input)	versus	100	bpm	without	the	latter.29	In	rats,	resting	HR	remains	at	approximately	290-370	or	350-400	bpm	depending	on	the	time	of	the	day	and	strain	(Wistar	and	Wild-type,	respectively).30	The	signal	then	flows	through	the	internodal	pathway	to	reach	the	second	atrial	node:	the	atrioventricular	(AV)	node,	located	in	the	septum	near	the	junction	of	the	right	atrium	and	the	right	ventricle.3	From	this	point	on	forward,	the	stimuli	descend	down	the	interventricular	septa	through	a	bundle	of	conducting	fibers,	consisting	of	only	specialized	cardiomyocytes,	called	the	bundle	of	His.3	Approximately	half	way	down	the	septum,	the	bundle	branches	into	right	and	left	bundles	to	surround	both	ventricles.3	The	fibers	continue					12	to	descend	and	once	they	reach	the	apex	of	the	heart,	their	direction	veers	upwards	to	envelop	the	ventricles;	these	new	branching	fibers	are	called	Purkinje	fibers.2,3	Purkinje	fibers	are	located	in	the	myocardium	of	all	chambers	but	can	be	found	in	greater	numbers	in	the	interventricular	septa.1	The	directionality	of	all	fibers	and	rhythmicity	of	stimuli	is	important	for	the	ventricles	to	contract	upwards	for	blood	to	exit	and	flow	through	the	semilunar	valves.3	Purkinje	fibers,	as	all	other	constituents	of	the	conducting	system,	are	made	of	specialized	cardiomyocytes.1	Intracellularly,	these	cardiomyocytes	are	distinct	from	others	because	of	their	disorganized	myofibrils,	and	increased	number	of	mitochondria	and	glycogen	stores,	which	serve	to	provide	these	cells	with	sufficient	energy	for	their	high	functional	demands	and	increased	ability	to	resist	hypoxia.1	In	addition	to	these	adaptations,	to	increase	their	conduction	ability,	these	cardiomyocytes	share	numerous	intercellular	channels	called	gap	junctions	which	allow	the	propagation	of	electrical	stimuli	from	cell-to-cell	via	the	flow	of	ions.1		1.2.4 Intercalated	discs		 Cardiomyocytes	are	connected	end-to-end	by	intercalated	discs,	which	have	dense	areas	of	condensed	specialized	proteins.1–3	Intercalated	discs	have	two	surfaces:	a	lateral	surface	(perpendicular	to	the	directionality	of	the	cell	and	electrical	stimuli),	and	a	transverse	surface	(parallel	to	the	cell).1–3	The	two	surfaces	have	distinct	transmembrane	proteins	with	different	functions.1–3	On	the	lateral	surfaces	of	the	intercalated	disc,	many	gap	junctions	serve	for	communication	and	conduction	of	electrical	stimuli;	thanks	to	these	channels,	depolarization	signals	are	able	to	rapidly	spread	throughout	the	tissue	to	generate	contraction.1–3	On	the	transverse	surfaces,	both	macula	adherens	(desmosomes;	also	found	on	the	lateral	surfaces)	and	fascia	adherens	(FA)	are	numerous	and	function	to	securely	anchor	cells	together.1–3	Desmosomes	and	FA	are	attached	to	intermediate	cytoskeletal	and	actin	filaments,					13	respectively	to	reduce	mechanical	stress	during	contraction.1	Cardiomyocytes	are	closely	attached	with	a	space	of	15-20	nm	between	cells.3		1.2.5 Generation	of	contraction	-	Summary		 To	generate	the	contraction	of	cardiomyocytes	and	subsequently,	the	entire	cardiac	tissue,	it	is	imperative	to	have	a	large	influx	of	calcium	ions	into	the	sarcoplasm.2,3	Once	the	depolarization	of	a	cell	is	induced,	calcium	channels	on	the	plasma	membrane	and	on	the	sarcoplasmic	membrane,	will	open	and	let	a	large	influx	of	calcium	ions.2,3	This	induces	intracellular	signal	transduction,	the	activation	of	multiple	different	pathways	and	even	further	calcium-induced	calcium	release.2,3	Calcium	ions	are	crucial	to	induce	conformational	changes	in	proteins	(i.e.,	tropomyosin	and	troponin)	blocking	myosin-binding	domains	on	thin	filaments.3	Once	myosin	heads	of	the	thick	filaments	are	able	to	attach	to	the	actin	filaments,	the	sarcomeres	are	then	able	to	shorten	in	synchrony	and	therefore	contract	the	entire	length	of	the	cell.3	T-tubules,	as	previously	described,	are	key	to	effectively	conduct	the	electrical	stimuli	and	cause	depolarization	deep	into	the	cell,	near	all	contractile	apparatus.2,3	The	specific	receptors	that	are	involved	in	the	influx	of	calcium	ions	into	the	sarcolemma	are	the	calcium-release	channels	and	the	ryanodine	receptors,	located	on	the	plasma	and	sarcoplasmic	membranes,	respectively.31	In	summary,	many	proteins	and	secondary	messengers	partake	in	cardiac	contraction.2,3	1.2.6 The	cardiac	cycle	Figure	1.2.A.	includes	pressure-volume	(PV)	data	obtained	via	placement	of	a	catheter	in	the	LV.	One	PV	loop	which	represents	one	cardiac	cycle.32	The	bottom-right	most	corner	of	the	loop	represents	the	end	of	cardiac	diastole,	the	relaxation	and	filling	phase;	at	this	stage,	the	heart	is	filling	with	blood.32	The	heart	then	enters	cardiac	systole,	the	contraction	phase.32	The	first	stage	of	systole	involves	isovolumetric	contraction	where	the	pressure	increases	with	no	changes	in	volume.32	The	increase	in					14	pressure	causes	the	aortic	valve	to	open	to	allow	for	blood	ejection	out	of	the	LV	and	therefore	causes	a	reduction	in	volume.32	Then,	diastole	commences	with	a	reduction	in	pressure.32	Once	the	pressure	is	lower	than	in	the	atrium,	the	ventricle	will	be	able	to	fill	with	blood	and	therefore	increase	in	volume	until	the	end	of	diastole.32		 		Figure	1.2.	Representative	pressure-volume	loop	at	resting	conditions.	The	different	phases	of	one	cardiac	cycle	are	explained.		1.3 The	renin-angiotensin-aldosterone	system	&	the	heart		 The	main	role	of	the	renin-angiotensin-aldosterone	system	(RAAS)	is	to	maintain	hemodynamic	homeostasis	thanks	to	endocrine	and	paracrine	secretions	from	many	organs	of	the	body.33	First,	RAAS	will	be	activated	in	response	to	low	blood	pressures	to	release	the	hormone	renin	from	the	kidney.33	Renin	converts	angiotensinogen	(Ao;	primarily	secreted	by	the	liver)	to	angiotensin	I	(ANGI).33	Angiotensin	converting	enzyme	(ACE;	secreted	by	the	pulmonary	and	renal	epithelium)	will	then	convert	ANGI	to	angiotensin	II	(ANGII)	which	will	directly	act	on	multiple	organs	(i.e.,	heart,	kidney,	adrenal	cortex	and	brain).33	ANGII	will	stimulate	the	adrenal	cortex	to	secrete	aldosterone	which	will	cause	ion	and	fluid	retention	in	the	kidneys	to	ultimately	increase	blood	pressure.33	At	the	cardiac	level,	increased	levels	of	ANGII	can	induce	changes	in	coronary	vascular	function,	muscle	metabolism,	inflammation,	structural	remodelling	(i.e.,	fibrosis	and	hypertrophy)	and	increased	cardiomyocyte	apoptosis.33	ANGII	and	atrial					15	natriuretic	peptide	(ANP;	although	primarily	produced	by	atrial	cardiomyocytes)	can	also	be	secreted	by	ventricular	cardiomyocytes	when	stretch	receptors	on	the	cells	are	activated	(Figure	1.4.).33	Additionally,	stretch	activation	of	the	same	cell	will	cause	additional	Ao	to	be	released	from	the	cardiomyocyte	(mRNA	levels	of	Ao	in	the	heart	are	less	than	0.1%	than	in	the	liver).33–37	Stretch	and	ANP	will	act	on	cardiac	fibroblasts	to	secrete	more	Ao	and	the	precursor	of	renin.33	Both	processes	will	ultimately	lead	to	further	increases	ANGII	levels	via	a	positive	feedback	loop.33	As	any	other	biological	pathway,	there	must	be	a	negative	regulator.	In	this	case,	ANGII,	itself,	will	bind	to	ANGII	plasma	membrane	receptors	(i.e.,	ANGII	receptor	1	(AT1))	on	cardiac	fibroblast	to	inhibit	renin	and	angiotensin	release.33	Interestingly,	NE,	released	from	sympathetic	neurons,	will	indirectly	increase	ANGII	levels	in	the	heart	by	promoting	Ao	production	and	release	from	cardiomyocytes,	and	Ao	and	renin’s	precursor	from	fibroblasts.33		Figure	1.3.	Molecular	pathways	of	angiotensin	II	production	in	the	heart.	ACE,	angiotensin	converting	enzyme;	ANGI,	angiotensin	I;	ANGII,	angiotensin	II;	ANP,	atrial	natriuretic	peptide;	Ao,	angiotensinogen;	AT1,	ANGII	receptor	1;	BP,	blood	pressure;	NE,	norepinephrine.33	Images	used	from	©	Servier	with	permission	(licensed	by	CC	BY	3.0).					16	1.4 Cardiac	muscle	homeostasis		1.4.1 Cardiac	plasticity	When	the	heart	is	subjected	to	different	types	of	molecular	or	physical	stimuli,	it	will	adapt	in	terms	of	its	muscle	structure	and	shape	of	its	chambers.38	In	other	words,	the	heart	is	able	to	change	in	according	to	its	physiological	needs.38	For	example,	if	there	are	changes	in	cardiac	loading,	pressure	and	volume	changes	will	be	detected	by	cardiac	mechanical	receptors,	which	will	induce	cardiac	remodelling	through	a	cascade	of	intracellular	pathways.38	This	remodelling	can	either	be	physiological	(i.e.,	exercise	and	pregnancy)	or	it	can	be	pathological	(i.e.,	denervation,	immobilization,	biochemical	stresses,	etc.).38		1.4.2 Regeneration	of	cardiac	muscle		Past	studies	have	shown	that	the	incapability	for	cardiac	myofibers	to	regenerate	was	due	to	the	absence	of	satellite	cells	(stem	cells),	otherwise	found	in	skeletal	muscle.1,2	It	is	additionally	thought	to	be	due	to	the	post-mitotic	status	of	cardiomyocytes	with	low	to	no	regeneration	ability.39	Recent	studies	investigating	heart	transplants	have	found	that	0.1%	of	cardiomyocytes	were	seen	to	have	nuclei	undergoing	mitosis	suggesting	that	cardiac	cells	could	in	fact	have	some	ability	to	regenerate	despite	its	weakness.2	Whether	these	findings	indicate	increased	cytokinesis	or	simply	karyokinesis	remains	unclear.	Regardless	of	this	evidence,	this	potential	regenerative	ability	of	these	cells	would	be	very	low.1		1.4.3 Hypertrophy	&	atrophy	Cardiac	muscle	cells	are	known	to	cease	multiplication	shortly	following	birth.39	As	the	newborn	heart	is	incapable	of	achieving	the	demands	of	an	adult	heart,	in	terms	of	function	and	mechanics,	the	organ	must	increase	in	size.39	However,	if	the	cells	are	no	longer	multiplying,	the	cells	themselves	must	compensate	for	growth.	Hypertrophy	is	the	increase	in	size	of	muscle	fibers,3	which	allows	the	growth	of	tissue	to	respond	to	new	physical	demands.39	Hypertrophy	can	be	eccentric	(change	in	mass,	dilation	of					17	the	chamber	lumens	and	sarcomeral	deposition	in	series)	or	concentric	(change	in	mass,	no	dilation	of	the	lumens	and	thickening	of	the	muscular	walls	due	to	sarcomeral	deposition	in	parallel).38	Eccentric	hypertrophy	occurs	after	volume	overload,	whereas	concentric	hypertrophy	occurs	after	pressure	overload.38	During	hypertrophy,	the	cell	will	exhibit	intracellular	changes	such	as	increased	cytoplasmic	volume,	increased	mitochondrial	density,	decreased	myofibril	size	and	increased	sarcomere	concentration.40	Researchers	hypothesize	that	these	changes	are	caused	by	the	increased	demand	in	energy	resulting	in	increased	mitochondria	fusion	and	are	due	to	the	increased	cytoplasmic	space	required	to	transport	high	energy	molecules.40	In	addition	to	an	increase	in	cardiomyocyte	size,	in	pathological	situations,	other	molecular	event	also	occur	during	hypertrophy	which	can	include	fibrosis	in	the	extracellular	matrix	and	increased	cell	death.38	It	is	important	to	note	that	pathological	hypertrophy	often	precedes	cardiac	dysfunction	and	heart	failure;	therefore,	itmust	be	taken	seriously.38		A	certain	degree	of	hypertrophic	cardiac	growth	is	crucial	to	the	development	of	the	organism	to	maintain	appropriate	cardiac	function.39	Hypertrophy	can	be	adaptive	and	maladaptive.	Following	endurance	and	strength	exercise,	the	heart	will	adapt	to	the	physical	demands.38	However,	if	hypertrophy	is	uncontrolled	or	unnecessarily	up-regulated,	it	can	become	maladaptive	and	therefore	pathogenic.39	For	example,	chronic	hypertension	will	increase	cardiac	demand	causing	the	tissue	to	respond	by	up-regulating	hypertrophic	pathways.39	The	expansion	of	cells	will	impair	myofibril	organization	subsequently	inducing	cardiomyocyte	cell	death	and	decreasing	cardiac	output,	which	all	together	will	increase	the	odds	for	heart	failure.39	Additionally,	cardiac	hypertrophy,	in	addition	to	fibrotic	remodelling,	and	cell	death,	has	been	seen	to	be	triggered	by	a	constant	over	activation	of	the	SNS.41,42	Distinguishing	physiological	versus	pathological	hypertrophy	can	be	complex	as	both	are	caused	by	cardiac	overloading	and	in	certain	cases	the	cause	(stimuli)	and	effect	are	indiscernible.43						18	Cardiac	atrophy	is	the	opposite	of	hypertrophy	where	heart	mass	is	decreased	in	size.38	It	can	occur	when	the	patient	is	immobilized	and	experiences	no-to-low	mobility	for	long	periods	of	time,	for	example,	after	high-level	SCI	and	after	decreases	in	cardiac	preload	(the	amount	of	stretch	of	the	ventricle	after	diastole;	non-invasively	estimated	with	end-diastolic	volume	(EDV)	and	pressure	(Ped)).38	During	atrophy,	cells	are	exhibiting	more	protein	degradation	and	less	protein	synthesis	events	causing	the	balance	to	shift	and	the	heart	cells	to	decrease	in	size.38	Molecular	pathways	involved	in	atrophy	will	be	discussed	in	greater	detail	in	the	following	section.	1.5 Cardiac	consequences	of	SCI	The	SC	plays	a	crucial	role	in	transporting	sensory,	motor	and	visceral	information	between	the	brain	and	the	periphery.	Unsurprisingly,	due	to	its	importance	and	omnipresent	involvement,	an	injury	to	the	SC	can	induce	various	catastrophic	consequences	on	vital	functions.	In	addition,	due	to	the	intricate	organization	of	nerves	exiting	the	SC	at	various	levels	to	innervate	different	organs,	injury	level	and	severity	will	greatly	determine	the	magnitude	and	quantity	of	secondary	consequences	on	the	body.	Following	SCI,	not	only	is	life	quality	significantly	reduced,	but	life	expectancy	is	said	to	decrease	by	30%	compared	to	able-bodied	subjects.44	In	2010,	the	Canadian	annual	incidence	for	SCI	was	4,259	with	a	total	population	of	85,556	in	Canada	44	and	2.5	million	worldwide.45	Although	SCI	only	affects	0.25%	of	the	Canadian	population,44	it	is	an	immense	burden	on	health	care	system	by	reason	of	the	gravity	and	multitude	of	complications	following	injury.46	On	average,	traumatic	SCI	(i.e.,	caused	by	an	external	physical	incident)44	costs	2.67	billion	Canadian	dollars	every	year.46	If	SCI	occurs	above	the	sympathetic	innervation	of	the	heart	and	vessels	(T1-T5	and	T1-L2,	respectively),47	the	cardiovascular	system	will	be	devoid	of	supraspinal	control	from	the	cardiovascular					19	control	center	in	the	brainstem.48	In	other	words,	the	sympathetic	signals	originating	in	the	RVLM	and	descending	through	the	SC	are	abruptly	terminated	due	to	the	injury.	This	loss	of	supraspinal	sympathetic	control	is	thought	to	explain	the	decrease	in	many	cardiac	structural	indices	seen	in	the	clinic	since	the	1980s.	For	example,	Kessler	et	al.	49	reported	decreases	in	LV	EDV,	LV	SV,	CO	and	estimated	LV	mass	in	patients	with	tetraplegia	(i.e.,	C8-T1	or	above)	compared	to	paraplegia	(i.e.,	T1	or	below).	These	cardiovascular	physiological	changes,	in	addition	to	reduced	physical	activity,	dyslipidemia	(abnormal	lipid	levels),50–53	increased	risk	for	metabolic	syndrome	(increased	obesity,54	insulin	resistance	55	and	odds	for	type	2	diabetes	56),	blood	pressure	instability	57,58	and	increased	arterial	stiffness	59	lead	to	premature	onset	and	increased	odds	for	cardiovascular	disease	(CVD)	in	this	population	by	2.67	fold	compared	to	able-bodied	subjects.60	Unsurprisingly,	CVD	is	the	leading	cause	of	death	in	this	population.60	1.5.1 Systolic	and	diastolic	function	following	SCI		1.5.1.1 Systolic	dysfunction	following	SCI		 Multiple	clinical	studies	investigating	cardiac	function	following	SCI	have	contradictory	results.	Both	Kessler	et	al.	and	Currie	et	al.	reported	decreased	SV	and	CO	in	quadriplegic	patients	compared	to	paraplegics,	indicating	worsened	function	with	higher	level	of	injury.49,61	West	et	al.	demonstrated	systolic	dysfunction,	with	decreased	LV	ejection	fraction	(EF;	%	of	blood	volume	ejected	by	the	LV	per	beat),	CO	and	SV	in	tetraplegic	athletes	compared	to	able	bodied	subjects.62	In	contrast,	Driussi	et	al.	reported	no	change	in	either	SV	or	EF	and	increased	LV	contractility	(explained	to	compensate	for	decreased	venous	return	and	inability	to	increase	HR).63	On	the	other	hand,	de	Groot	et	al.	indicated	no	systolic	dysfunction	following	SCI	with	a	decreased	trend	for	CO.64	Despite	these	inconsistent	findings,	a	meta-analysis	performed	by	our	research	team,	which	examined	echocardiographic	measures	of	cardiac	structure	and	function,	has	indicated	that	SV	is	decreased	with	no	change	in	EF	following	chronic	SCI.65					20		 EF,	though	used	widely	in	the	clinic,	is	not	an	ideal	measure	to	infer	cardiac	systolic	function	as	it	is	load-dependent	which	means	it	truly	does	not	infer	the	intrinsic	function	of	the	heart	but	the	function	of	the	entire	cardiovascular	system.	Additionally,	as	EF	is	load-dependent,	if	cardiac	volumes	are	greatly	and	proportionally	reduced,	EF	will	not	be	changed	significantly,	rendering	EF	impractical	to	infer	systolic	function	in	this	situation.	To	truly	study	intrinsic	cardiac	function	in	vivo,	it	is	required	to	remove	confounds	of	altered	loading	conditions.	A	method	has	been	developed	to	uncouple	the	heart	from	the	rest	of	the	cardiovascular	system	by	occluding	the	inferior	vena	cava	(IVC)	in	vivo.	Doing	so	will	subsequently	decrease	preload	and	create	a	series	of	PV	loops	with	different	volumes.	The	slope	of	all	connecting	end-diastolic	pressure	and	volume	points	is	used	as	an	index	of	intrinsic	LV	contractility,	the	end-systolic	pressure-volume	relationship	(ESPVR;	also	called	end-systolic	elastance	(Ees)).	As	occluding	a	vessel	is	highly	invasive,	it	is	only	suited	for	pre-clinical	studies	involving	animals.66	With	this	technique	and	using	rodent	models	of	SCI,	our	laboratory	has	demonstrated	twice	in	different	strains	of	rats	(Wistar	and	lean	Zucker)	that	systolic	dysfunction,	load-dependent	and	-independent,	is	decreased	following	high	thoracic	SCI,	as	evidenced	by	decreases	in	pressures,	the	maximal	rate	of	LV	systolic	pressure	increment	(dP/dtmax)	and	ESPVR.62,63	1.5.1.2 Diastolic	dysfunction	following	SCI	In	clinical	research,	diastolic	function	is	assessed	with	LV	filling	velocities,	obtained	via	non-invasive	techniques	(i.e.,	echocardiography).	Many	studies	disagreed	on	how	diastolic	function	changed	following	SCI	as	these	data	were	wildly	inconsistent.	63,67–71	However,	a	recent	meta-analysis	was	performed	by	our	research	group	and	it	reported	a	decrease	in	diastolic	function	in	the	human	SCI	population.65	In	pre-clinical	studies,	diastolic	dysfunction	is	assessed	more	accurately	with	more-invasive	techniques	such	as	LV	catheterization	and	IVC	occlusions.62,63	In	these	studies,	diastolic	function	can	be	inferred	with	two	load-dependent	measures	with	LV	catheterization:	tau	(LV	diastolic	time	constant)	and	-				21	dP/dtmin	(maximal	rate	of	LV	diastolic	pressure	decrement).	Intrinsic	cardiac	diastolic	function	can	be	inferred	with	the	load-independent	index	for	compliance:	the	end-diastolic	pressure-volume	relationship	(EDPVR;	the	non-linear	fit	of	all	end-diastolic	volume	and	pressure	points).	A	reduction	in	diastolic	function	following	high-level	SCI	in	pre-clinical	settings	has	not	yet	been	confirmed	as	load-independent	and	-dependent	data	are	not	in	agreement.	EDPVR	is	reported	to	not	change	significantly	in	the	sub-acute	and	chronic	settings	following	severe	high-level	SCI	(complete	transection	at	the	3rd	thoracic	segment	(T3)72	and	severe	contusion	at	the	2nd		thoracic	segment	(T2)73),	while	both	tau	and	-dP/dtmin	were	significantly	decreased	in	the	chronic	stage	following	complete	T3-SCI	but	not	in	the	sub-acute	phase	following	severe	T2	contusion.72	Due	to	this	discrepancy,	more	pre-clinical	research	on	diastolic	function	post-SCI	is	required	and	for	this,	new	contemporary	methods	to	measure	intrinsic	diastolic	function	accurately	are	currently	being	designed	in	our	laboratory.	1.5.1.3 Mechanisms	affecting	cardiac	function	following	SCI	The	function	of	the	heart	following	high-level	SCI	can	be	negatively	affected	by	multiple	mechanisms:	loss	of	supraspinal	sympathetic	74	and	sympathoadrenal	control,75,76	altered	neurohumoral	control	(i.e.,	decreased	circulating	NE	and	increased	ANGII	in	the	chronic	phase	post-SCI),72	reduced	physical	activity	77	and	cardiac	unloading	due	to	a	reduction	in	blood	volume	and	pressure.		Immediately	and	several	hours	following	traumatic	SCI,	there	is	a	sudden	loss	or	depression	in	spinal	reflexes,	defined	as	spinal	shock,	and	in	sympathetic	tone	below	the	level	of	injury,	defined	as	neurogenic	shock	which	leads	to	hypotension	and	bradyarrhythmias.78,79	High-level	injuries	with	loss	of	supraspinal	sympathetic	control	to	the	heart	and	the	vasculature	can	have	drastic	impacts	on	systemic	hemodynamic	function	due	to	reduced	sympathetic	activity	below	the	level	of	injury.74,80	Additionally,	if	the	level	of	injury	incapacitates	signals	to	reach	the	adrenal	medulla,	circulating	catecholamine	levels	will	remain	low	even	during	conditions	typically	requiring	the	activation	of	the	SNS	(i.e.,	stress	and					22	exercise).75,76	Due	to	reduced	direct	vascular	sympathetic	activity	below	the	level	of	injury,80–82	reduced	NE	circulation	48,75,76	and	reduced	motor	control	to	the	skeletal	muscle	pump,83,84	vessels	will	no	longer	constrict	effectively,	blood	pressure	will	acutely	decrease	85	and	blood	will	pool	at	the	extremities	and	the	splanchnic	area,	affecting	venous	blood	return.48,85–93	Venous	pooling,	causing	reduced	venous	pressure	and	reduced	pressure	gradient	between	the	venous	circuit	to	the	right	atrium,	and	reduced	blood	volume		will	ultimately	decrease	blood	delivery	back	to	the	heart	(which	will	decrease	preload).92,94	Acutely	after	injury,	decreased	vasoconstriction	of	the	arteries,85	due	to	the	loss	of	sympathetic	control	to	the	vasculature,80–82	and	its	subsequent	decrease	in	blood	pressure	85	should	reduce	afterload	(the	amount	of	force	the	ventricle	must	generate	to	eject	blood).	In	the	chronic	phase,	vessels	will	have	undergone	remodelling	due	to	changes	in	limb	use,	blood	pressure	and	increase	in	arterial	shear	stress,85	which	will	cause	stiffening	of	the	vasculature.85	Furthermore,	both	acutely	and	chronically,	low	blood	pressure	will	increase	circulating	ANGII,33	which	has	been	observed	to	directly	cause	atrophic	95–97	and	fibrotic	remodelling	98	in	skeletal	and	cardiac	muscle,	respectively.		Collectively,	altered	neurohumoral	control	and	loss	of	supraspinal	sympathetic	control	following	SCI	lead	to	cardiac	unloading	further	impacting	cardiac	function.	Additionally	and	interestingly,	in	a	study	comparing	moderate	and	severe	high-thoracic	contusions,	our	research	team	have	shown	that	with	only	10%	of	neuronal	sparing	(descending	sympathetic	neurons)	cardiac	function	is	preserved.99	Recently,	our	laboratory	has	performed	two	experiments	to	study	cardiac	function	following	SCI:	1)	by	stimulating	adrenergic	receptors	in	the	heart	with	the	administration	of	a	b1	agonist	(dobutamine)	in	SCI	rats,73	and	2)	by	investigating	the	effects	of	intact	and	absent	supraspinal	sympathetic	control	(by	varying	the	level	of	injury).	Both	studies	show	normal	cardiac	function	in	rats	treated	with	dobutamine	73	and	in	rats	with	intact	sympathetic	control.	These	results	imply	that	the	loss	of	supraspinal	sympathetic	control	may	be	the	key	determinant	of	cardiac	dysfunction	following	SCI.					23	1.5.2 Cardiac	and	cardiomyocyte	structure	following	SCI			 Clinical	studies	have	investigated	cardiac	structure	with	echocardiography	due	to	the	inability	to	use	more	invasive	techniques	on	human	subjects.	All	report	decreases	in	estimated	LV	mass	in	quadriplegic	patients	(with	loss	of	supraspinal	sympathetic	control)	compared	to	either	able	bodied	subjects	or	paraplegic	counterparts.49,64,100	In	a	pre-clinical	study	in	our	laboratory,	Wistar	rats	showed	decreases	in	total	cardiac	mass	4	weeks	post	severe	T2	contusion	by	approximately	35%	(p<0.01)73	Squair	et	al.	was	the	first	study	to	report	significant	atrophy	of	cardiomyocytes	in	length	(by	approximately	22%;	T2	93.9	±8.54	versus	control	CON	121	±5.98	µm,	p<0.01)	and	Z-line	width	(by	approximately	26%;	T2	7.25	±0.82	versus	9.82	±1.80	µm,	p<0.05)	with	no	change	in	sarcomere	length	following	high-thoracic	sub-acute	SCI,	confirming	the	presence	of	cardiac	atrophy.73			 In	addition	to	loss	of	cardiac	mass,	the	T2	contusion	rats	demonstrated	reduced	body	mass	72,73	and	reduced	lower-limb	use	following	SCI.	Weight-loss	and	reduced	physical	activity,	by	themselves,	can	cause	cardiac	atrophy.101	As	our	current	explanation	hypothesis	for	cardiac	atrophy	following	SCI	is	due	to	the	loss	of	supraspinal	sympathetic	control,	this	could	jeopardize	our	rationale	for	why	there	might	be	atrophic	remodelling	following	SCI.	However,	outside	of	the	SCI	field	it	has	been	shown	that	ablation	of	sympathetic	neurons	to	the	heart,	alone,	caused	cardiac	atrophy	in	a	rodent	model	with	no	changes	in	body	mass	or	physical	activity.102		As	cardiomyocytes	have	been	shown	to	atrophy	following	high-thoracic	SCI	in	rodents	with	loss	of	supraspinal	sympathetic	control,	the	study	of	the	cellular	and	molecular	pathways	involved	in	this	atrophy	was	required.	In	the	literature,	there	were	no	studies	which	investigated	any	cellular	remodelling	pathways	associated	with	cardiac	atrophy	post-SCI.	Nevertheless,	there	are	many	publications	investigating	skeletal	muscle	atrophy	following	denervation	or	disuse,	and	one	publication	investigating	cardiac	atrophy	following	pharmacological	ablation	of	sympathetic	neurons	(with	6-hydroxy-dopamine;	6-				24	OH-DOPA).102	These	studies	assisted	our	research	team	to	target	important	genes	associated	with	the	two	main	proteolytic	pathways	of	the	cell:	the	ubiquitin	proteasome	system	(UPS)	and	autophagy.		In	a	healthy	cell,	protein	synthesis	and	protein	degradation	are	constantly	active	and	exist	at	equal	rates	to	maintain	homeostasis.103	Without	protein	synthesis,	the	cell	would	not	be	able	to	perform	specific	biological	and	essential	housekeeping	functions.103	Likewise,	protein	degradation,	also	known	as	proteolysis,	is	crucial	for	the	survival	of	the	cell	as	it	acts	to	recycle	proteins	which	are	old,	misfolded,	defective	or	no	longer	necessary	to	avoid	any	toxic	aggregates.103	If	proteolytic	pathways	are	up-regulated,	cells	will	atrophy	and	cause	subsequent	shrinkage	of	the	organ.104	Proteolysis	is	an	organized	process	which	can	be	accomplished	by	multiple	different	systems.103	There	are	five	intracellular	major	proteolytic	pathways	which	include:	the	UPS,	the	lysosomal	machinery	(i.e.	endocytosis	and	autophagy),	the	calpain	enzymes,	mitochondrial	proteinases	and	membrane	proteinases.103	All	processes	use	a	variety	of	enzymes	called	proteases	(>200)	which	hydrolyze	bonds	between	specific	pairs	of	amino	acids	to	break	down	the	entire	peptide.103	Hydrolysis	will	lead	to	protein	fragmentation	thanks	to	the	addition	of	water	and	a	specialized	enzyme	as	a	catalyst.103		The	mechanisms	of	calpains,	mitochondrial	and	membrane	proteases	will	not	be	discussed	in	this	thesis.	Special	focus	and	greater	detail	will	be	given	to	the	two	main	proteolytic	pathways:	UPS	and	autophagy	lysosomal	machinery.	1.5.3 The	ubiquitin-proteasome	system		1.5.3.1 Overview	&	mechanism	The	UPS	utilizes	a	series	of	specialized	enzymes	(i.e.,	E1,	E2	and	E3	ligases)	to	tag	proteins	with	ubiquitin	(Ub),	a	small	regulatory	molecule,	for	degradation.104,105	The	tagged	proteins	will	then	be	recognized	by	a	complex	of	proteases,	forming	a	proteasome,	which	will	break	down	the	peptide	bonds					25	and	annihilate	the	targets	into	amino	acids	for	recycling.104,105	Proper	function	of	UPS	is	crucial	to	degrade	and	recycle	defective	proteins	to	avoid	aggregation	and	subsequent	cell	death,	or	cell	atrophy.104,105	In	muscle,	there	exists	additional	specialized	enzymes	to	tag	sarcomeric	proteins.104	This	is	particularly	important	for	cardiac	muscle	as	it	is	the	most	energy	demanding	tissue	of	the	body	and	requires	effective	and	continuous	contraction	of	myofibrils.1					 The	UPS	mechanism	is	schematically	summarized	in	Figure	1.5.	Before	tagging	proteins	with	Ub,	the	Ub	molecules	themselves	must	be	activated	by	E1	enzymes	through	the	cleavage	of	ATP.104	The	E1	enzymes	will	then	transfer	the	activated	Ub	to	an	E2	enzyme.104	The	E2	will	bind	an	E3	ligase;	different	E2-E3	combinations	will	tag	different	proteins	for	degradation.104	More	than	650	E3	ligases	have	been	identified	in	the	human	body;106	but	not	all	will	be	transcribed	in	all	tissues.104	This	great	variety	allows	for	high	selectivity	for	degradation.104	The	rate-limiting	step	will	occur	when	the	E3	ligase	is	bound	to	both	the	E2	enzyme	and	the	targeted	protein	to	transfer	the	Ub	onto	the	target.104	Once	the	target	is	polyubiquitinated	with	several	Ub,	the	protein	ZNF216	will	recognize	and	deliver	the	tagged	target	to	the	proteasome	for	degradation.104		 Studies	investigating	UPS	have	largely	focused	on	ubiquitination	and	not	on	deubiquitination.104	Two	deubiquitinating	enzymes	have	been	identified	in	skeletal	muscle	atrophy	following	disuse	and	denervation	(USP14,	ubiquitin	carboxyl-terminal	hydrolase	14),107	and	fasting	(USP19,	ubiquitin	carboxyl-terminal	hydrolase	19).108	USP19	protein	levels	were	seen	to	be	negatively	correlated	with	muscle	mass,	which	implies	that	deubiquitination	might	recycle	Ub	from	degraded	targets	for	more	subsequent	tagging.108					26		Figure	1.4.	Schematic	of	the	ubiquitin	proteasome	system.	Reproduced	with	©	permission	from	Bonaldo	et	al.104	1.5.3.2 Muscle	atrophy	&	UPS			 In	skeletal	muscle	atrophy	following	denervation,	the	two	first	identified	up-regulated	E3	ligases	were	atrogin-1	(MAFbx;	also	called	muscle	atrophy	F-box)	and	muscle	ring	finger	1	(MuRF1).104,109	Two	important	MAFbx	targets	are	myoblast	determination	protein	1	(MyoD;	in	skeletal	muscle),	a	muscle-specific	transcription	factor,110	and	eukaryotic	translation	initiation	factor	3	subunit	f	(eIF3f),	an	important	translation	initiator	which	binds	the	small	ribosomal	unit	for	translation	activation.111	In	addition,	MAFbx	also	targets	calcineurin	which	indirectly	induces	hypertrophy.112	MuRF1	is	muscle	specific	and	targets	sarcomeric	proteins	for	degradation,	such	as	muscle	actin,	nebulin,	titin,	myosin	heavy/light	chains,	myosin	binding	protein	C	and	troponin	I.104,113–117	Two	other	lesser-investigated	E3	ligases	involved	in	atrophy	are	E3	ubiquitin-protein	ligase	TRIM32	(TRIM32)	and	tumor	necrosis	factor	(TNF)	receptor	associated	factor	6	(TRAF6).104,105	TRIM32	tags	for	degradation	thin	filament	proteins	(i.e.,	tropomyosin,	troponin	and	actin)	and	Z-line	proteins	(i.e.,	desmin	and	a-actinin),118	whereas,	TRAF6	up-regulation	is	required	for	proper	activation	of	MAFbx	and	MuRF1	through	AMP-activated	protein	kinase	(AMPK)	and	forkhead	box	O3	(FoxO3).119					27		 MAFbx,	MuRF1	and	ZNF216	expression	is	regulated	through	forkhead	box	O	(FoxO)	transcription	factors.104	FoxO	transcription	factors	need	to	undergo	posttranslational	modifications	(i.e.,	dephosphorylated,	acetylated,	etc.)	to	be	able	to	be	translocated	into	the	nucleus	and	transcribe	E3	ligases.104,120	There	are	three	isoforms:	FoxO1,	FoxO2	and	FoxO3.104		In	healthy	conditions,	insulin	and	insulin-like	growth	factor	1	(IGF-1)	bind	to	plasma	membrane	receptors	to	activate	the	PI3K-Akt	pathways	(PI3K,	phosphoinositide	3-kinase;	Akt,	protein	kinase	B)	which	leads	to	hypertrophy.121	This	causes	Akt	to	be	phosphorylated	to	promote	protein	synthesis	through	the	mammalian	target	of	rapamycin	(mTOR)	pathways	and	inhibit	protein	degradation	by	phosphorylating	FoxO.121		In	cases	of	disuse	and	fasting	where	levels	of	insulin	and	IGF-1	are	low	or	mechanical	unloading	of	the	muscle,	Akt	does	not	get	phosphorylated	and	will	cause	the	dephosphorylation	and	therefore	translocation	of	FoxO	into	the	nucleus	to	transcribe	atrophy	genes	(atrogenes;	i.e.,	E3	ligases).121,122	In	the	case	of	denervation,	loss	of	trophic	support	dephosphorylates	FoxO3	(via	β2	adrenergic	signalling),	which	causes	increased	up-regulation	of	MAFbx.102,120	However,	with	time	FoxO3	is	progressively	acetylated	and	is	tagged	for	degradation	if	catabolic	conditions	are	not	maintained.120	If	catabolic	conditions	continue,	FoxO3	will	remain	activated	and	will	cause	UPS	and	autophagy	up-regulation.120	During	high	energy	stress,	elevated	AMPK	levels,	due	to	increased	AMP:ATP	ratio,	will	up-regulate	UPS	atrogene	transcription	by	phosphorylating	FoxO3	at	Akt-independent	binding	sites.104,123–125	Other	inducers	of	UPS,	include	circulating	increased	oxidative	stress,	ANGII	(i.e.,	via	decreased	Akt	phosphorylation	or	increased	oxidative	stress	via	activation	of	AT1),	transforming	growth	factor	beta	(TGF-b),	and	tumor	necrosis	factor	a	(TNFa)		and	TNF-like	weak	inducer	of	apoptosis	(TWEAK)	which	activate	nuclear	factor	kappa	B	(NF-kB)	transcription	factors.95,104,105,126–130	A	summary	diagram	of	pathways	associated	with	muscle	atrophy	can	be	found	in	Figure	1.6.					28		Figure	1.5.	Pathways	involved	in	protein	degradation	and	synthesis.	Adapted	with	©	permission	from	Bonaldo	et	al.104	1.5.4 Autophagy	1.5.4.1 Overview	&	mechanism		 Autophagy	is	a	process	of	protein	degradation	which	utilizes	the	formation	of	vesicles	within	the	cells	around	bulks	of	less-transient	proteins	and	organelles.104,131,132	There	exist	different	types	of	autophagy	(i.e.,	macroautophagy,	microautophagy,	mitophagy,	etc.),	however,	this	section	will	solely	focus	on	macroautophagy	as	it	is	the	most	investigated	in	situations	of	muscle	atrophy.104		For	initiation	and	the	formation	of	a	phagophore	(once	called	isolation	membrane),	the	formation	of	serine/threonine-protein	kinase	ULK1	(ULK1)	complex	(composed	of:	autophagy	related	protein	(ATG)	13	,	ULK1,	ATG101,	FAK	family	kinase-interacting	protein	of	200	kDa	(FIP200))	is	required	to	activate	class	III	PI3K	(beclin	1	(BCN1),	phosphatidylinositol	3-kinase	(Vps)	Vps34,	Vps15,	ATG14)	and	subsequently	generate	phosphatidylinositol	3-phosphate	(PI3P).133,134	For	elongation	of	the	phagophore,	two	more	complexes	are	required.134	First,	ATG5	and	ATG12	must	be	covalently	bonded	by	ATG7	and	ATG10.135–137					29	The	newly	bonded	ATG5-ATG12	can	then	bond	to	ATG16-like	protein	(ATG16L)	to	form	the	first	required	complex.138,139	Second,	microtubule-associated	protein	1	light	chain	3	(LC3)	must	be	lapidated	by	the	ATG5-ATG12-ATG16L	complex,	then	conjugated	to	phosphatidylethanolamine	by	ATG7	and	ATG3	to	form	LC3II,	the	second	complex.140,141	LC3II	will	complete	elongation	of	the	phagophore	into	an	autophagosome	(as	enclosed	vesicle).142	For	mitophagy,	the	specialized	form	of	autophagy	for	mitochondria,	there	exist	specialized	proteins	involved	in	the	recruitment	of	mitochondria	to	the	phagophore	(i.e.,	PTEN-induced	kinase	1	&	Parkin	complex	(PINK1-PRKN),	sporozoite	surface	protein	P36p	(p52)	and	BCL2/adenovirus	E1B	19	kDa	protein-interacting	protein	3	(Bnip3)	factors	which	will	tag	the	surface	of	the	organelle	with	Ub).104	A	simplified	diagram	of	all	proteins	required	for	autophagy	completion	can	be	found	in	Figure	1.7.	Lysosomes	are	intracellular	vesicles	surrounded	with	a	phospholipid	bilayer,	which	enclose	an	acidic	environment	favorable	for	enzyme	activation.103	Lysosomes	will	fuse	with	endosomes	containing	extracellular	contents	(produced	via	endocytosis)	or	with	autophagosomes	to	form	an	auto-lysosome.21,103	The	proteases	located	in	the	lysosomes	will	take	action	and	digest	the	contents	of	the	fused	vesicle.103	Permeases,	located	in	the	vesicle	membrane,	will	transport	molecules	back	into	the	cytosol	for	recycling.143		Autophagy	is	up-regulated	in	periods	of	crisis	when	cells	require	rapid	recycling	of	amino	acids	as	nutrients	103.	Some	inducers	of	autophagy	include	cellular	stress,144	aging,143	fasting/starvation	103,133	and	denervation	(Figure	1.6.).104		The	activation	of	the	PI3K-Akt-mTOR	pathway	and	the	elevated	levels	of	insulin	can	inhibit	autophagy	(Figure	1.6.).133	High	AMP:ATP	ratio	will	cause	increased	in	AMPK,	which	will	inhibit	these	autophagy	inhibitors	(Figure	1.6.).133	In	addition,	previously	discussed	FoxO	transcription	factors	are	also	able	to	translocate	to	the	nucleus	and	transcribe	crucial	autophagy-related	proteins	(Figure	1.6.).104,105					30		Figure	1.6.	A	schematic	of	macroautophagy	activation	for	purpose	to	degrade	proteins	in	bulk.	Proteins	targeted	for	degradation	(i.e.,	filamin)	by	autophagy	are	tagged	with	ubiquitin	(Ub)	and	transported	to	the	site	of	phagophore	elongation	via	p52.	Adapted	with	©	permission	from	Bonaldo	et	al.104	1.5.4.2 Muscle	atrophy	&	autophagy	The	up-regulation	of	autophagy	related	proteins	(mostly	BCN1)	indicating	increased	protein	degradation	has	been	found	in	studies	investigating	both	skeletal	muscle	atrophy	following	denervation	and	fasting,104	and	cardiac	atrophy	following	denervation	and	mechanical	unloading.102,145	Cardiomyocyte	remodelling	caused	by	autophagy	following	cardiac	stress	was	seen	to	be	regulated	by	increased	ANGII	146	via	AT1.96	1.5.5 The	ubiquitin-proteasome	system	and	autophagy	following	SCI	Our	research	team	conducted	a	study	to	investigate	the	expression	of	both	UPS	and	autophagy	in	cardiac	tissue	following	high-thoracic	chronic	SCI.72	Male	lean	Zucker	rats	were	either	subjected	to	T3	complete	transections	or	uninjured	(CON;	controls).72	At	the	end	of	the	study	(12	weeks),	cardiac	tissue	was	collected	for	histology	and	gene	expression	analyses.72	Unsurprisingly,	cardiac	mass	estimated	via	magnetic	resonance	imaging	(MRI)	decreased	by	approximately	29%	in	T3-SCI	rats	compared	to	uninjured	rats.72	In	terms	of	cell	atrophy,	the	study	reports	similar	findings	to	Squair	et	al.	73,	with	decreased	cardiomyocyte	length	(by	approximately	17%;	T3	79.8	±10.7	vs.	96.0	±5.52	µm,	p<0.05)	and	Z-line	width	(by	approximately	19%;	T3	7.25	±0.39	vs.	8.92	±0.23	µm,	p<0.001)	following	high-thoracic	chronic	SCI.72					31	This	cell	atrophy	was	associated	with	the	up-regulation	of	both	UPS	and	autophagy	in	the	T3	rats	compared	to	uninjured	rats.72	Western	blots	showed	significant	up-regulation	for	MuRF1	and	BCN1	proteins	in	T3	rats	compared	to	CON.72	mRNA	expression	analyses	showed	significant	up-regulation	of	UPS	(MuRF1	and	MAFbx),	autophagy,	(ATG12	and	BCN1),	TGFb	receptors	(TGFbR1	and	R2)	and	ANGII	receptors	(AT1	and	ANGII	receptor	2	(AT2))	following	SCI	compared	to	the	controls.72	Additionally,	T3	rats	had	50%	more	circulating	ANGII	and	81%	less	circulating	NE	compared	to	CON.72	Although,	mechanical	loading	and	stretch	at	the	level	of	the	cardiomyocytes	were	not	quantified	in	this	study,	T3	rats	experienced	cardiac	unloading	compared	to	CON.72	All	results	indicate	cardiac	atrophy	to	be	associated	with	the	up-regulation	of	proteolytic	pathways	chronically	following	high-thoracic	SCI	in	rodents	which	may	be	partly	induced	by	changes	in	neurohumonal	pathways	(increased	ANGII),	changes	in	neuromechanical	loading	(cardiac	unloading)	and		the	loss	of	descending	sympathetic	control	to	the	heart	(decreased	trophic	input).72		To	decipher	a	time-sensitive	treatment	to	decrease	the	magnitude	of	cardiac	atrophy	in	the	SCI	population,	we	must	first	investigate	when	these	molecular	changes	are	occurring	and	then	characterize	in	greater	detail	the	signalling	pathways	and	molecules	involved	in	cardiomyocyte	atrophy	after	SCI.	1.5.6 Temporal	regulation	of	remodelling	pathways	in	atrophy	Once	again,	although	there	are	no	published	studies	in	the	literature	investigating	the	temporal	effects	of	SCI	on	cardiac	atrophy	at	the	molecular	level,	some	pre-clinical	studies	have	investigated	the	temporal	effects	of	denervation	in	both	cardiac	(ablation	of	sympathetic	neurons)	and	skeletal	muscle.102,147–149	The	ablation	of	sympathetic	neurons	with	a	chemical	agent	causes	complete	loss	of	sympathetic	control	to	the	heart,	while	a	high-thoracic	SCI	eliminates	the	majority	of	supraspinal	sympathetic	input	to	the	heart	the	sub-lesional	sympathetic	circuitry	remain	intact	which	may	be					32	important	for	reflexive	cardiac	control.	Although	both	ablation	of	sympathetic	neurons	and	SCI,	are	drastically	different	procedures,	the	former	gives	us	insight	on	the	molecular	events	occurring	in	the	heart	when	completely	devoid	of	any	SNS	input	and	provides	direction	for	SCI	research.	Furthermore,	the	vast	majority	of	denervation	research	investigates	skeletal	muscle.	While	skeletal	muscle	has	distinct	morphological	features	and	regenerating	capacity,1,2	its	research	is	relevant	as	all	myocyte	types	share	common	cellular	properties.		Both	Li	et	al.	(skeletal	muscle	denervation)	and	Zaglia	et	al.	(cardiac	muscle	denervation)	report	the	up-regulation	of	UPS	(i.e.,	MuRF1)	at	their	first	tested	time	point,	one	day	following	denervation	102,147	with	a	peak	at	day	3	post-denervation.147	Autophagy	activity	was	increased	following	day	30	post-denervation,	indicating	it	is	further	activated	later	than	UPS.102	UPS	proteins	remained	significantly	up-regulated	compared	to	the	control	at	the	last	time	point	in	the	study	of	Li	et	al.	(14	days	post-denervation).147	However,	contrary	to	the	results	from	Li	et	al.	and	our	preliminary	experimental	results,	Zaglia	et	al.	reported	normalized	UPS	and	autophagy	mRNA	levels,	and	normalized	UPS	protein	levels	at	eight	days	following	denervation	in	cardiac	muscle.102,147	Additionally,	Zaglia	et	al.	reported	a	decrease	in	Akt-pathways	(protein	synthesis)	at	day	one	following	denervation	which	remained	low	until	study	termination,	one	month	post-treatment.102	As	gross	cardiac	atrophy	should	be	subsequent	to	the	molecular	up-regulation	of	protein	degradation	and	down-regulation	of	protein	synthesis	pathways,	it	seems	logical	that	cardiomyocyte	atrophy	would	occur	at	a	later	time.	In	the	literature,	the	time	following	denervation	to	cause	significant	atrophy	is	less	conclusive.	Significance	for	gross	tissue	atrophy	was	seen	one,102	three	149	and	five	days	150	following	denervation.	These	inconsistencies	could	be	due	to	the	type	of	muscle	(cardiac	vs.	skeletal),	the	muscle	itself	(gastrocnemius	vs.	tibialis	anterior),	method	for	denervation	(pharmacological	ablation	of	neurons	vs.	physical	denervation)	or	the	sample	size	in	the	analysis.					33	1.6 Closing	remarks	Following	chronic	high-thoracic	SCI	in	clinically	relevant	models,	our	research	team	has	reported	the	occurrence	of	systolic	contractile	dysfunction	and	cardiomyocyte	atrophy	following	severe	contusion	SCI	and	complete	transection	at	high-thoracic	levels.72,73	Further	studies	in	our	laboratory	demonstrated	the	up-regulation	of	major	proteolytic	pathways,	UPS	and	the	autophagy	lysosomal	machinery,	in	the	heart	following	chronic	SCI,72	the	activation	of	which	has	been	shown	to	be	critical	in	induction	of	skeletal	and	cardiac	muscle	atrophy.72,102,151–153	Despite	numerous	clinical	and	pre-clinical	studies	confirming	the	physiological	aspects	of	our	findings	(i.e.,	reduced	LV	volume	and	cardiac	dysfunction),19,49,73,154	our	understanding	of	the	cellular	and	molecular	events	underlying	cardiac	atrophy	post-SCI	remains	limited.	Moreover,	there	have	been	no	studies	that	have	investigated	the	temporal	progression	of	cardiac	remodelling	from	acute-to-chronic	SCI,	which	impedes	our	ability	to	develop	a	time-sensitive	treatment	for	this	population.	The	study	is	the	first	to	define	the	temporal	changes	in	cardiac	function	and	structure,	as	well	as	the	molecular	pathways	underlying	these	responses,	across	the	acute	to	chronic	continuum	post-SCI.			 Aims	and	hypotheses		The	primary	aim	for	this	thesis	project	was	to	investigate	the	temporal	effects	of	acute	high-thoracic	SCI	on	LV	cardiac	function	(i.e.,	ESPVR),	proteolysis	(i.e.,	gene	expression	of	key	UPS	and	autophagy	targets)	and	cardiomyocyte	atrophy	(i.e.,	cardiomyocyte	length	and	width).	This	overarching	aim	was	tested	as	three	different	questions.		1) Does	cardiac	dysfunction	begin	to	occur	in	the	acute	phase	following	high-thoracic	SCI?	If	yes,	when	does	it	begin	to	occur?	2) Is	there	a	change	in	the	regulation	of	the	two-main	intracellular	proteolytic	pathways	(UPS	and	autophagy)	in	the	acute	phase	following	high-thoracic	SCI?	If	yes,	when	does	it	begin	to	occur?	3) Is	LV	cardiomyocyte	atrophy	significant	in	the	acute	phase	following	high-thoracic	SCI?	If	yes,	when	does	atrophy	begin	to	occur?	Hypothesis	1.	Systolic	dysfunction	will	occur	immediately	following	high-thoracic	SCI	due	to	immediate	loss	of	cardiac	descending	supraspinal	sympathetic	control,	while	diastolic	dysfunction	will	not	occur	in	the	acute	phase	of	SCI	due	to	there	being	no	sufficient	time	for	structural	remodeling.	Hypothesis	2.	There	will	be	an	up-regulation	of	proteolytic	pathways	occurring	during	the	acute	phase	following	high-level	SCI.	As	UPS	activity	can	be	altered	by	sympathetic	input	102,	humoral	levels72	and	mechanical	loading,105,122	and	as	there	is	an	immediate	loss	of	cardiovascular	descending	sympathetic	control	and	loss	of	lower-limb	muscle	control	following	T3-SCI,	UPS	regulation	will	increase	immediately	post-SCI	and	will	then	decrease	back	to	basal	levels	while	remaining	at	a	higher	level	than	pre-injury.	Contrarily	to	UPS,	there	is	no	clear	picture	of	the	timeline	of	autophagy	regulation	following	neural	trauma	and	it	is	therefore	difficult	to	hypothesize	an	exact	timing.	Autophagy	will	be	up-regulated	within	the	acute	timeline	post-SCI	in	cardiac	tissue	due	to	the	immediate	changes	in	blood	pressure	and	subsequent	increase	in	circulating	ANGII,	which	is	known	to	activate	the	pathway.72,98,126,155					35	Hypothesis	3.	As	atrophy	should	be	subsequent	to	the	shift	in	protein	homeostasis,	LV	cardiomyocyte	atrophy	will	occur	after	the	upregulation	of	proteolytic	pathways	and	therefore	will	be	discernible	multiple	days	following	high-thoracic	SCI.			 					36	 Materials	and	methods	3.1 Overview	A	schematic	overview	of	the	methods	can	be	found	in	Figure	3.1.	Male	Wistar	rats	(n=49)	were	assigned	to	two	groups	which	differed	in	the	type	of	surgery:	a	dorsal	durotomy	with	no	SCI	(SHAM)	or	a	complete	transection	at	the	3rd	thoracic	segment	(T3-SCI).	To	study	the	temporal	effects	of	acute	SCI,	T3-SCI	animals	were	terminated	at	different	time	points	along	the	acute	timeline:	12	hours	(n=9),	1	day	(n=8),	3	days	(n=10),	5	days	(n=7)	and	7	days	post-SCI	(n=9).	SHAM	rats	were	euthanized	at	the	last	time	point,	7	days	post-operatively	(n=6).	At	pre-intervention	and	at	different	time	points	along	the	acute	timeline	(1	day,	2	days,	4	days	and	6	days	post-surgery),	volumetric	and	functional	cardiac	indices	were	obtained	in	vivo	via	echocardiography	in	the	7-day	T3-SCI	and	SHAM	groups.	At	termination,	for	all	groups	except	the	12-hour	group,	functional	load-dependent	and	-independent	cardiac	indices	were	obtained	via	LV	PV	catheterization.	Following	all	euthanasia,	cardiac	tissue	was	harvested	for	all	groups	to	assess	cardiomyocyte	morphology	via	immunofluorescence	and	to	quantify	gene	expression	of	key	targets	of	the	two	main	proteolytic	pathways.	Furthermore,	femurs	were	collected	and	measured	to	standardize	cardiomyocyte	dimensions.							37	Figure	3.1.	Overview	of	methods.	PV,	pressure-volume;	SHAM,	dorsal	durotomy	with	no	SCI;	T3-SCI,	SCI	at	3rd	thoracic	segment.	Images	used	from	©	Servier	with	permission	(licensed	by	CC	BY	3.0).	3.2 Ethics	and	disclaimer		 Ethics	approval	was	granted	by	UBC.	All	staff	in	contact	with	the	animals	were	certified	and	trained	by	the	UBC	Animal	Care	Committee.	All	protocols	involving	the	use	of	animals	and	biological	tissue	were	conducted	in	strict	accordance	with	the	Canadian	Council	for	Animal	Care.	All	techniques	and	equipment	employed	in	this	study	have	been	used	frequently	in	our	laboratory	in	the	past;	thus,	familiarity	with	data	collection	and	analysis	was	not	of	any	concern.	Additionally,	to	ensure	the	collection	of	reliable	data,	each	procedure	(i.e.,	surgery,	tissue	excision,	tissue	sectioning,	confocal	imaging,	etc.)	was	performed	by	the	same	individual	throughout	the	study.	Finally,	all	data	were	blinded	for	analysis.	3.3 Animals	Due	to	the	high	number	of	replicates	required,	rats	were	the	most	adequate	animal	model	for	such	study.	We	decided	to	use	male	Wistar	rats	as	they	have	commonly	been	studied	in	the	field	of	high-thoracic	SCI	for	over	15	years	and	animal	care	for	this	type	and	level	of	SCI,	sex	and	strain	has	been	developed	in	great	detail	at	our	facility.156	The	Wistar	strain	has	been	repeatedly	used	in	studies	investigating	the	effects	of	SCI	on	the	cardiovascular	system.73,99,154,157	Furthermore,	males	were	chosen	over	females	for	consistency	with	previous	pre-clinical	cardiovascular-SCI	research	and	because	males	constitute	over	74%	of	the	Canadian	human	population	with	traumatic	SCI.44	A	total	of	49	rats	were	ordered	from	Charles	River	Laboratories	(Wilmington,	MA,	USA)	to	be	between	the	age	of	10-11	weeks	old	and	approximately	300	to	325	g.	It	was	crucial	for	rats	to	not	vary	in	age	(>1	week	of	age)	to	minimize	differences	in	heart	size	changes	due	to	maturation.	Rats	were	received	at	the	animal	care	facility	(ICORD	Vivarium)	at	least	7	days	before	the	beginning	of	the	study	to	allow	for	acclimatisation	to	their	new	environment,	diet,	handling,	restraint	and	staff.						38	3.4 Groups	and	termination	time-points		 There	were	six	groups	of	rats	which	differed	in	surgery	type	(T3-SCI	versus	SHAM)	and	termination	time-points.	43	rats	underwent	a	complete	transection	SCI	at	the	T3	level	and	six	rats	underwent	a	dorsal	durotomy	with	no	damage	to	the	SC	(SHAM).	SHAM	rats	served	as	controls	to	reduce	the	effects	of	surgery	as	confound	variables	(i.e.	injections,	drugs,	anesthesia,	anxiety,	frequent	handling	and	restraint,	post-operative	pain,	etc.).		We	were	interested	in	the	acute	phase	of	SCI,	which	is	generally	defined	to	be	up	to	one	week	post-SCI	in	rats.158–160	The	time	of	the	T3-SCI	was	considered	to	be	time=0	and	five	endpoints	were	investigated	along	the	acute	timeline.	The	following	times	were	chosen	for	a	temporal	homogenous	spread:	12	hours,	1	day,	3	days,	5	days	and	7	days	post-SCI.		While	reviewing	chronic	studies	from	our	research	group	with	similar	procedures	and	outcomes,	a	post-hoc	G*Power	(v.	3.1.9.3)	analysis	indicated	that	for	80%	power	a	sample	size	of	two-to-four	animals	per	group	would	be	sufficient	to	detect	a	between-group	comparison	for	the	key	functional	outcome,	ESPVR,	and	a	sample	size	of	four-to-six	animals	per	group	for	the	key	structural	outcomes,	cardiomyocyte	length	and	width.	Due	to	high	mortality	following	this	SCI	model	(0-10%),156	a	sample	size	of	eight	rats	was	chosen	to	account	for	such	possibility.	For	our	controls,	we	deemed	sufficient	to	retain	a	sample	size	of	six	SHAM	rats	to	be	terminated	on	day	7	post-surgery	(the	last	endpoint).				Group	1:	Complete	T3	SCI	and	terminated	at	12	hours	post-SCI	(T3-SCI	12	hours)	Group	2:	Complete	T3	SCI	and	terminated	at	1	day	post-SCI	(T3-SCI	1	day)	Group	3:	Complete	T3	SCI	and	terminated	at	3	days	post-SCI	(T3-SCI	3	days)	Group	4:	Complete	T3	SCI	and	terminated	at	5	days	post-SCI	(T3-SCI	5	days)	Group	5:	Complete	T3	SCI	and	terminated	at	7	days	post-SCI	(T3-SCI	7	days)	Group	6:	Dorsal	durotomy	with	no	SCI	and	terminated	at	7	days	post-durotomy	(SHAM)					39	Table	3.1.	List	of	investigated	dependent	variables,	organized	per	method	Method	and	outcome	 Variable	 Notes	Echocardiography	(PSLAX)	 	 	Volumes	 EDV	(µL)	 Index	of	preload		 ESV	(µL)	 	Systolic	function	 SV	(µL)	 		 CO	(mL/min)	 		 EF	(%)	 		 FS	(%)	 	Catheterization	 	 	Arterial	hemodynamics	 SBP	(mmHg)	 		 DBP	(mmHg)	 		 MAP	(mmHg)	 Index	of	afterload		 HR	(bpm)	 	LV	systolic	function	 ESPVR	(mmHg/µL)	 Index	of	contractility		 SW	(mmHg*mL)	 		 Pmax	(mmHg)	 		 Pdev	(mmHg)	 		 dP/dtmax	(mmHg/s)	 		 TPR	(mmHg/mL)†	 Index	of	afterload		 Ea	(mmHg/µL)	 Index	of	afterload		 Ea/ESPVR	 Index	of	ventricular-arterial	coupling		 PRSW	(mmHg)	 	LV	diastolic	function	 -dP/dtmin	(mmHg/s)	 		 Ped	(mmHg)	 		 Tau	(ms)	 	Histology	 	 	Cardiomyocyte	dimensions	 Length	(µm)	 Measured	with	connexin-43		 Width	(µm)	 Measured	with	a-actinin		 CSA	(µm2)	 Measured	with	WGA		 Volume	(µm3)	 Measured	with	CSA	and	length	Gene	analysis	 	 	RNA	extraction,	cDNA	synthesis	and	qPCR	 Fold	change	each	target	 Targets	in	Table	3.2.	CO,	cardiac	output;	CSA,	cross-sectional	area;	DBP,	diastolic	blood	pressure;	dP/dtmax,	maximal	rate	of	systolic	pressure	increment;	-dP/dtmin,	maximal	rate	of	diastolic	pressure	decrement;	Ea,	arterial	elastance;	Ea/ESPVR,	ventricular	vascular	coupling	ratio;	EDV,	end-diastolic	volume;	EF,	ejection	fraction;	ESPVR,	end-systolic	pressure-volume	relationship;	ESV,	end-systolic	volume;	FS,	fractional	shortening;	HR,	heart	rate;	LV,	left-ventricle;	MAP,	mean	arterial	pressure;	Pdev,	developed	pressure;	Ped,	end-diastolic	pressure;	Pmax,	maximum	pressure;	PRSW,	preload	recruitable	stroke	work	;	PSLAX,	parasternal	long-axis	view;	SBP,	systolic	blood	pressure;	SV,	stroke	volume;	SW,	stroke	work;	Tau,	diastolic	time	constant;	TPR,	total	peripheral	resistance;	WGA,	wheat-germ	agglutinin.	†,	TPR	calculated	with	CO	obtained	via	echocardiography	and	MAP	obtained	via	catheterization	(TPR=MAP/CO).						40	3.5 Issues	with	internal	validity	Independently,	age,	weight-loss,	reduced	physical	activity	and	stress	can	cause	cardiac	atrophy.101	Therefore,	it	was	critical	to	control	for	such	confounds.	All	rats	were	ordered	to	be	the	same	age	and	were	fed	an	enriched	diet	pre-	and	post-surgery	to	avoid	excessive	weight	loss	following	the	intervention.	Due	to	the	loss	of	motor	control	in	their	hind	limbs,	rats	required	additional	help	to	move	around	their	cages	to	find	food,	water	and	shelter.	Therefore,	to	help	with	mobility	and	to	promote	physical	activity,	cages	were	equipped	with	plastic	grids	underneath	the	bedding	for	rats	to	grasp	and	facilitate	locomotion.	Proper	acclimatization	to	their	new	environment	is	crucial	to	reduce	confounds,	to	ensure	data	reliability,	to	alleviate	anxiety	and	post-intervention	complications.156	Therefore,	animals	arrived	at	the	facility	and	were	started	on	their	new	diet	a	week	prior	to	the	first	intervention.	Additionally,	animal	care	personnel	visited	the	rats	daily	with	gradual	increases	in	handling	time	and	frequency	to	ensure	human	contact	became	stress-free	before	surgery	day.156	3.6 Pre-surgery	animal	care		3.6.1 Housing	The	rats	were	located	in	rooms	with	controlled	temperature	and	12-hour	light-dark	cycles.	All	rats	were	housed	in	250	cm2	cages	containing	a	maximum	of	4	cagemates.	No	animals	were	housed	individually	as	they	require	social	enrichment.	For	the	comfort	of	the	rats,	cages	included	bedding	and	transparent	red	polycarbonate	huts.	Cages	were	changed	regularly	to	ensure	bedding	remained	dry	to	avoid	any	possible	infections.	3.6.2 Nutrition	and	hydration	All	groups	were	fed	an	enriched	diet	before	the	start	of	the	study	(i.e.	normal	rat	chow	in	addition	to	cereal,	fruit,	spinach,	nutritive	transport	gel,	etc.).	As	previously	mentioned,	an	enriched	diet	is	critical					41	to	avoid	any	cardiac	atrophy	due	to	total	weight	loss	following	SCI	for	the	T3	animals.	In	addition,	it	is	required	to	acclimatize	the	rats	to	the	diet	before	surgery	to	avoid	any	reluctance	to	ingest	new	foods	during	post-operative	care.	Water	and	food	were	provided	ad	libitum.	3.6.3 Antibiotics	Enrofloxacin	was	administered	to	the	rats	subcutaneously	once	a	day,	three	days	pre-operatively	(Baytril,	10	mg/kg).		3.7 Pre-surgery	data	collection:	echocardiography	(for	T3-SCI	7	days	and	SHAM	groups)	The	clinical	literature	indicates	preference	for	MRI	imaging	over	all	types	of	echocardiography	for	more	precise	and	accurate	measurements	of	cardiac	structure.161,162	However,	due	to	smaller	hearts	and	higher	HR	in	rodents,	MRI	has	not	yet	been	claimed	the	gold	standard	for	pre-clinical	rodent	cardiac	imaging.163	With	echocardiography,	high	frame	rates	can	be	achieved	for	greater	temporal	resolution,163	which	is	advantageous	for	our	model.	We	used	a	transthoracic	echocardiography	system	(Vevo	3100	imaging	system,	FUJIFILM	VisualSonics)	with	a	rodent	transducer	(MX250;	13-24	MHz	liner	array	transducer)	(used	in	73,99,154).	As	we	were	interested	in	the	difference	between	pre-	and	post-operative	measures	along	the	acute	timeline,	this	method	was	deemed	adequate	and	non-invasive	for	our	purposes.	A	parasternal	long-axis	(PSLAX)	view	with	B-mode	imaging	was	obtained	and	used	to	assess	systolic	function	and	volumes.	The	latter	were	obtained	by	tracing	the	epicardial	border	of	the	LV.	All	analyses	were	performed	after	data	collection	with	commercially	available	software	(Vevo	Lab).	For	all	echocardiographic	variables,	see	Table	3.1.	Due	to	the	non-invasive	nature	of	echocardiography,	the	animal	was	anaesthetized	with	isoflurane	alone	(first	with	5%	induction	with	2	L	min-1	O2	in	an	induction	box	then	1-2%	induction	with	a	nose	cone).	For	this	procedure,	the	rat	was	shaved	ventrally	(thoracic	area)	and	lightly	attached	to	the					42	ultrasound	platform	with	surgical	tape.	The	lubricated	probe	was	placed	on	the	rat’s	thorax	to	image	the	heart	to	obtain	echocardiography	images.	During	this	procedure,	the	ventilation	and	heart	rate	of	the	animal	was	monitored	at	all	times.	Once	the	procedure	was	completed,	the	rat	was	placed	individually	in	a	clean	and	comfortable	cage	to	be	monitored	closely	until	anaesthesia	fully	wore	off.	Echocardiography	images	for	these	groups	(T3-SCI	7	days	and	SHAM)	were	taken	at	pre-intervention,	12	hours,	1	day,	2	days,	4	days,	and	6	days	post-surgery	under	isoflurane.	3.8 T3-SCI	and	SHAM	surgeries	All	staff	present	in	the	surgical	room	were	certified	for	their	specific	tasks.	All	equipment	and	solutions	were	aseptic	and	sterile,	respectively.	3.8.1 Anesthesia	and	preparation	After	weighting	the	animal,	the	rat	was	anesthetized	with	inhalant	isoflurane	in	a	clean	induction	chamber	lined	with	paper	towel	(5%	induction	with	2	L	min-1	O2).	Once	the	animal	had	lost	the	righting	reflex,	the	animal	was	taken	out	of	the	induction	chamber	to	be	placed	on	its	ventral	side	and	fitted	with	a	nose	cone	(1-2%	induction	with	2	L	min-1	O2,	depending	on	depth	of	anesthesia	of	the	animal).		To	avoid	drying	of	the	eyelids	and	the	cornea,	ocular	lubricant	was	generously	placed	on	the	entire	eye	of	the	rodent.	To	prepare	the	site	for	dorsal	durotomy,	the	site	of	incision	was	shaved	and	disinfected	three	times	with	alternating	chlorhexidine	(Hibitane)	and	70%	alcohol.	Pre-surgical	care	(i.e.,	shaving	and	disinfecting)	was	performed	as	instructed	by	the	UBC	Animal	Care	Committee	to	reduce	any	risks	of	contaminating	the	surgical	wound.	To	avoid	dehydration	during	surgery,	warmed	Lactated	Ringer’s	solution	was	injected	subcutaneously	before	surgery	(10	mL/kg).	Combination	of	analgesics	and	antibiotics	was	injected	subcutaneously:	buprenorphine	(Temgesic	0.02	mg/kg)	and	enrofloxacin	(Baytril	10	mg/kg).	Once	all	pre-surgical	care	was	completed	and	the					43	palpebral	and	toe	pinch	reflexes	were	lost,	surgery	was	commenced.	At	any	point,	if	the	animal	showed	any	signs	of	a	lack	of	oxygenation	or	had	an	irregular	respiration	rate,	all	procedures	were	paused	immediately	to	attend	to	the	animal’s	needs.		At	all	times	during	pre-operation	preparation,	durotomy,	SCI	and	immediately	post-operative	care,	the	animal	remained	on	a	heat	source	(i.e.,	water-circulating	blanket)	or	in	an	incubator	to	keep	body	temperature	at	37°C	degrees.		3.8.2 Dorsal	durotomy	(for	both	SHAM	and	T3-SCI	animals)	The	animal	was	placed	in	a	prone	position	and	a	5	mL	syringe	was	positioned	underneath	its	neck	to	elevate	the	thoracic	vertebrae.	The	surgeon	palpated	the	vertebral	column	to	locate	the	T2	vertebra	thanks	to	its	uniquely	long	neural	spine.	An	incision	at	the	dorsal	midline	from	C8-T2	was	done	with	a	scalpel.	Next,	with	blunt	scissors,	the	trapezius	was	carefully	dissected.	Once	again,	the	neural	spine	of	T2	was	located	and	this	allowed	to	attain	and	cut	the	dura	between	the	T2	and	T3	vertebrae	with	micro-scissors.	For	SHAM	animals,	the	intervention	stopped	at	this	stage.	3.8.3 Spinal	cord	injury	(only	for	T3-SCI	animals)	After	the	opening	of	the	dura	was	achieved,	the	surgeon	cut	the	SC	with	micro-scissors	and	suctioned	a	segment	of	approximately	0.5	cm	at	the	T3	level.	With	a	dissecting	microscope,	the	surgeon	ensured	a	complete	injury	with	no	remaining	tissue	connecting	the	two	blunted	ends	of	the	SC.	Gelfoam,	a	hemostatic,	was	placed	between	the	rostral	and	caudal	ends	to	stop	potential	bleeding,	fill	the	now	empty	space	and	ensure	no	touching	of	the	newly	blunted	ends.	All	surgical	tools	were	autoclaved	for	each	first	use	of	the	day	and	sterilized	with	a	glass	bead	sterilizer	between	individual	surgeries.	Although,	a	complete	transection	is	not	the	most	common	type	of	injury	in	the	clinic	as	most	injuries	can	be	defined	as	partial	or	combinations	of	lacerations,	contusions	and	compressions,164	severe					44	contusions	and	complete	transections	at	high-thoracic	levels	have	shown	to	have	nearly	identical	effects	on	the	cardiovascular	system	in	a	rodent	model.73,99	A	complete	injury	was	chosen	as	this	type	of	injury,	where	the	possibility	of	differential	sparing	at	the	site	of	injury	is	minimal,	is	preferred	in	experimental	scenarios	due	to	increased	consistency	between	subjects	compared	to	a	contusion,	and	our	research	team	has	reported	preserved	cardiac	function	with	10%	of	sparing.99	Additionally,	the	majority	(55%)	of	SCI	patients	suffer	from	cervical	injuries.165	Therefore,	in	regard	to	the	level	of	injury,	it	would	be	most	clinically	valid	to	utilize	a	model	with	a	higher-level	injury	than	T3.	However,	a	higher-level	injury	could	damage	the	nerves	innervating	the	rat	forelimb	muscles	(C2-T1	166),	which	could	paralyze	the	forelimbs	and	incapacitate	the	rats	to	groom	and	move	around	the	cage	to	find	food	and	water.	In	addition,	as	the	level	and	severity	of	the	injury	is	increased,	the	risk	for	secondary	complications,	including	cardiovascular	dysfunction,	is	increased.74,167		To	avoid	tetraplegia	and	jeopardizing	the	rats’	well-being,	a	T3	injury	has	been	commonly	administered	in	the	field	of	SCI	as	a	compromise.156	Due	to	the	expansion	of	the	secondary	injury	in	the	SC,	a	T3	injury	will	damage	most	of	T2	and	part	of	T1,	essentially	causing	the	loss	of	sympathetic	control	to	the	heart	without	rendering	the	rats	tetraplegic	as	most	SPN	originate	at	the	T2	level	in	rats.168	Although,	animal	care	for	T3	rats	remains	challenging,	a	group	of	researchers	from	our	facility	have	implemented	an	animal	care	protocol	for	this	model	which	helped	to	ensure	the	best	possible	management	for	the	rats	and	to	minimize	the	burden	of	high-thoracic	SCI	compared	to	lower	level	injuries.156	3.8.4 Suturing	The	trapezius	muscle	was	sutured	with	continuous	absorbable	monocryl	(4-0).	Whereas	the	skin	was	sutured	with	interrupted	non-absorbable	prolene	(4-0).					45	3.9 Post-surgery	animal	care	3.9.1 Immediately	post-operation	All	animals,	which	underwent	surgery	(durotomy	+	T3-SCI	or	durotomy	alone),	were	placed	in	a	thermo-regulated	incubator	(37°C	degrees),	immediately	following	suturing.	The	incubator	was	lined	with	simple	laboratory	mats	to	avoid	heat	loss	and	eye	irritation	from	bedding	particles.	Staff	closely	monitored	the	rats	every	5-10	minutes	until	all	anesthesia	effects	had	worn	off.	Water	gel	and	treats	were	given	close	to	the	animals	as	mobility	was	impaired	by	analgesics	and	previous	anesthesia.	Once	rats	were	recovered	from	the	anaesthesia,	they	were	placed	in	their	initial	cages	and	monitored	every	hour	up	to	8	hours	post-operation.	3.9.2 Continued	antibiotics	&	pain	management	Following	the	surgeries,	the	same	analgesics	and	antibiotics	(buprenorphine	every	12	hours	and	enrofloxacin	every	24	hours)	were	administered	subcutaneously	for	a	duration	of	3	days	post-operatively	and	if	needed.		3.9.3 Housing,	nutrition	and	hydration	The	same	housing	arrangements	as	described	above	in	pre-surgery	animal	care	were	respected	(Section	3.6.1.).	One	difference	post-operatively	was	the	choice	of	bedding.	Oats	were	used	up	to	three	days	post-intervention	as	under	analgesics	rats	tended	to	chew	on	the	regular	bedding.	To	avoid	constipation	and	intestinal	blockages,	we	decided	to	place	the	rats	on	a	comestible	bedding	with	abundant	leaves	of	spinach.	In	addition,	to	encourage	movement	of	paraplegic	rats,	rubber	grids	were	placed	under	the	bedding	to	provide	grip	surfaces.	Water	and	the	enriched	diet	were	continued	to	be	provided	ad	libitum.	Food	was	made	to	be	easily	accessible	and	placed	at	distinct	areas	to	promote					46	physical	activity	from	the	paraplegic	animals.	Once	again	to	avoid	dehydration,	additional	warmed	lactated	Ringer’s	solution	was	injected	subcutaneously	as	needed.	3.9.4 Bladder	care	During	the	first	week	following	high-thoracic	SCI,	T3-SCI	animals	had	their	bladders	expressed	manually	by	staff	as	micturition	(i.e.,	detrusor	muscle,	urethral	sphincters	and	bladder	neck,	which	are	innervated	from	the	sympathetic	T10-L2	and	parasympathetic	S2-S4	nerves)	was	temporarily	impaired.156,169	Gentle	and	careful	bladder	care	is	crucial	to	avoid	rupture	of	the	bladder.156	The	reflexive	micturition	commonly	returns	within	one	week	to	10	days.156	However,	due	to	the	endpoints	of	this	study,	bladder	care	was	performed	for	all	T3-SCI	animals	until	euthanasia.	Additionally,	sperm	blockages	were	removed	and	bladders	were	emptied	at	least	four	times	a	day,	every	6-8	hours.		3.9.5 Health	assessments	The	rats	were	closely	monitored	four	times	daily.	Upon	the	first	morning	visit	of	each	day,	rats	were	weighted	and	assessed	for	weight	loss,	physical	appearance,	behaviour,	clinical	signs,	wounds	and	any	other	concern.	To	aid	distinction	of	pain	levels	in	rodents,	staff	utilized	the	Facial	Rat	Grimace	scale.170	Even	though	experts	have	been	skeptical	of	this	scale,171	it	was	followed,	per	the	suggestions	of	the	UBC	Animal	Care	Committee,	in	addition	to	other	behavioural	indices	(i.e.,	grooming,	arching,	flinching,	abdominal	press,	writhing,	staggering	and	twitching).	All	data	per	rat	were	manually	recorded	onto	monitoring	sheets	and	added	up	to	obtain	a	total	score.	If	the	score	equaled	or	exceeded	20	points,	the	rat	was	to	be	euthanized.	Fortunately,	this	never	occurred	in	the	present	study.					47	3.10 Termination	day	data	collection:	in	vivo	outcomes	3.10.1 Anesthesia	The	animal	was	anaesthetised	under	urethane	(1200-1500	mg/kg)	instead	of	inhalant	isoflurane	because	urethane	provides	a	more	stable	anesthetic	for	long-term	surgical	procedures,	such	as	LV	catheterization.66	Urethane	was	administered	via	intraperitoneal	injection	in	small	increments	every	15	minutes	until	the	animal	was	at	surgical	plane	(loss	of	righting,	palpebral	and	toe-pinch	reflexes).		3.10.2 Catheterization	For	LV	catheterization,	rats	were	verified	to	be	in	surgical	plane	under	urethane	anaesthesia	and	placed	in	a	supine	position.	The	following	are	widely-used	procedural	steps	for	the	insertion	of	the	admittance	PV	catheter	(Transonic,	1.9F	rat	PV	catheter)	into	the	LV	and	these	can	be	found	in	greater	and	pictorial	detail	in	Nature	Protocols.66		First,	the	rat’s	neck	was	elevated	with	a	5mL	syringe.	The	surgeon	made	an	incision	from	the	lower	jaw	to	the	sternum	through	cutaneous,	subcutaneous	tissue	and	superficial	muscles.	All	tissue	(i.e.	salivary	glands,	sternohyoid	and	sternomastoideus	muscles)	obstructing	the	right	carotid	and	trachea	was	carefully	displaced	to	the	side.	At	this	stage,	the	rat	underwent	a	tracheotomy	to	be	intubated	for	the	potential	need	of	supportive	respiratory	care.	Once	the	carotid	was	isolated	from	surrounding	tissue	and	from	the	vagal	nerve,	a	total	of	three	sutures	were	required.	Two	sutures	were	looped	and	tightly	held	around	the	most	anterior	and	posterior	ends	of	the	artery.	The	last	suture	was	loose	and	placed	towards	the	middle	of	both	previous	sutures,	which	would	later	serve	to	secure	the	catheter	to	the	carotid	once	placement	was	ideal	within	the	animal.	Thanks	to	the	sutures	in	place,	the	carotid	was	able	to	be	lightly	stretched	allowing	the	surgeon	to	incise	its	anterior	end.	The	catheter,	pre-soaked	in	warm	saline,	was	advanced	posteriorly	through	the	micro-incision	in	the	carotid.	Once	the	catheter	was	inside	the	carotid					48	artery,	the	posterior	suture	was	released	and	the	catheter	was	quickly	advanced	approximately	2-3	cm	into	the	artery,	after	which	the	middle	suture	was	lightly	fastened	to	keep	the	catheter	in	place	and	to	prevent	any	bleeding.	The	animal	was	left	to	rest	for	10	minutes	after	which	basal	arterial	data	were	collected	once	fully	stable	(i.e.,	systolic,	diastolic	and	arterial	blood	pressures,	and	heart	rate).	After	the	collection	of	arterial	data,	the	catheter	was	inserted	via	pressure	guidance	and	guided	by	echocardiography,	if	required.	The	ideal	position	of	the	admittance	PV	catheter	in	the	LV	was	checked	and	confirmed	by	echocardiography.	Following	another	10	minutes	of	rest	and	stable	vitals,	basal	PV	LV	data	were	recorded	for	10	minutes.	All	data	were	collected	via	Labchart	8.1	(AD	Instruments,	USA)	with	the	PVAN	add-on,	which	is	capable	of	generating	and	displaying	the	corresponding	PV	loops	in	real-time.		From	these	PV	loops,	collected	at	baseline,	many	cardiac	load-dependent	indices	are	measured	and	used	to	infer	systolic	and	diastolic	function	(listed	in	Table	3.1.).	Such	cardiac	indices	include	volumes	and	pressures:	end-systolic	volume	(ESV),	EDV,	SV,	end-systolic	pressure	(Pes),	end-diastolic	pressure	(Ped),	LV	developed	pressure	(Pdev)	and	minimum	and	maximum	pressures	and	volumes	(Figure	3.2.A.).	The	area	of	the	loop	can	be	measured	to	infer	stroke	work	(SW),	the	amount	of	work	done	by	the	ventricle	(Figure	3.2.A.).	Arterial	elastance	(Ea)	is	the	ratio	of	LV	Pes	to	SV	and	a	measure	of	afterload	(Figure	3.2.A.).	The	remaining	indices	such	as	tau,	dP/dtmax	and	-dP/dtmin	are	measured	via	Labchart.		 Next,	to	obtain	load-independent	measures,	inferior	vena	cava	(IVC)	occlusions	were	performed	by	isolating	the	blood	vessel	via	a	ventral	laparotomy	(closed-chest	approach).	Occluding	the	IVC	with	a	cotton-tip	applicator	allowed	for	reduced	venous	return	to	the	heart.	This	generated	multiple	PV	loops	with	changing	preload.	Load-independent	measures	are	important	to	infer	the	intrinsic	function	of	the	heart	and	not	of	the	entire	cardiovascular	system	(listed	in	Table	3.1.).	From	an	occlusion,	we	measured	two	functional	variables:	preload	recruitable	stroke	work	(PRSW;	relationship	between	SW	and	EDV)	and					49	ESPVR.	ESPVR	is	an	index	of	contractility	and	is	the	linear	slope	connecting	all	end-systolic	PV	points	(Figure	3.2.B.).	In	this	thesis,	systolic	function	from	catheterization	data	was	inferred	with	the	following	indices:	maximum	pressure	(Pmax),	Pdev,	SW,	dP/dtmax,	Ea,	ESPVR	and	PRSW.	How	well	the	heart	and	vasculature	are	working	together	was	measured	via	the	ventricular-arterial	coupling	ratio,	which	is	calculated	by	dividing	Ea	by	ESPVR	(Ees).	Concerning	diastolic	function,	it	was	inferred	with	dP/dtmin,	Ped	and	tau.	Cardiac	volumes,	CO,	EF	and	FS	were	inferred	from	echocardiography	data	due	to	higher	accuracy.			Figure	3.2.	Representation	of	pressure-volume	data	obtained	via	left-ventricle	catheterization	and	inferior	vena	cava	occlusions.	In	panel	A,	one	pressure-volume	loop	is	shown	with	its	labeled	cardiac	indices	obtained	from	the	baseline	recording.	In	panel	B,	a	series	of	loops	was	generated	via	IVC	occlusions	to	measure	ESPVR.	Ea,	arterial	elastance;	ESV,	end-diastolic	volume;	ESPVR,	end-systolic	pressure-volume	relationship;	ESV,	end-systolic	volume;	Ped,	end-diastolic	pressure;	Pes,	end-systolic	pressure;	SV,	stroke	volume;	SW,	stroke	work.		3.11 Euthanasia		3.11.1 Surgical	endpoints	At	the	end	of	the	in	vivo	outcome	procedures,	as	the	rat	remained	under	deep	urethane	anesthesia,	a	last	reflex	check	was	performed	before	euthanasia.	Euthanasia	was	completed	after	the	A	 B					50	righting,	palpebral,	toe-pinch	reflexes	were	all	confirmed	to	be	lost.	To	euthanize	the	anesthetized	animal,	a	bilateral	pneumothorax	cut	(from	the	posterior	end	of	the	thoracic	cavity)	and	an	excision	of	the	heart	were	performed.		3.11.2 Non-surgical	endpoints	If	at	any	time	an	animal	was	suffering	of	physical	injuries,	such	as	severe	self-mutilation,	unhealable	wounds	or	was	unable	to	eat	or	drink,	the	animal	was	planned	to	be	euthanized.	As	mentioned	previously,	all	abnormal	behaviours,	physical	injuries	and	weight	loss	were	recorded	as	part	of	the	regular	daily	monitoring	and	if	the	score	equaled	or	exceeded	20	points,	euthanasia	was	to	be	performed.	Fortunately,	no	abnormal	behaviours	were	noted	and	no	rats	were	euthanized	prior	to	the	experimental	endpoint.	3.12 Collection	of	tissues		3.12.1 Cardiac	tissue	post-mortem	After	completion	of	the	bilateral	pneumothorax	procedure,	the	lower	posterior	half	of	the	heart	was	excised	for	molecular	investigation.	The	apical	portion	of	the	heart	was	sectioned	for	gene	analysis	(mRNA	extraction,	cDNA	synthesis	and	qPCR).	For	tissue	destined	for	mRNA	quantification,	the	tissue	was	placed	immediately	in	RNA	later	solution	in	4°C	degrees	overnight	before	the	suctioning	of	the	solution	and	storage	at	-80°C	degrees.	Once	the	apex	was	excised,	transcardial	perfusion	was	achieved	first	with	phosphate-buffered	saline	(PBS)	then	with	4%	paraformaldehyde	for	fixation.	After	successful	fixation,	a	cross-sectional	cut	at	mid-ventricular	level	was	collected	for	histology	and	sent	to	Wax-it	Histology	Services	at	UBC	for	paraffin	embedding.						51	3.12.2 Femur	post-mortem	The	femur	was	collected	following	completion	of	fixation.	The	femur	was	then	later	measured	in	length	(cm)	for	the	standardization	of	structural	cardiac	indices.		3.13 Genetic	analysis	3.13.1 RNA	expression	–	mRNA	extraction,	cDNA	synthesis	and	PCR	To	measure	gene	expression,	the	dorsal	lateral	quadrant	of	each	apex	(previously	stored	in	-80°C	degrees)	was	thawed	and	manually	dissected	with	sterile	surgical	blades.	First,	RNA	extraction	was	performed	following	via	the	Invitrogen	TRIzol™	Reagent	procedure	(Thermo	Fischer	Scientific	15596026).	The	only	modification	to	this	protocol	was	an	increased	incubation	time	of	12	hours	in	isopropanol	at	-20°C.	Before	proceeding	to	cDNA	synthesis,	RNA	concentration,	purity	and	integrity	was	controlled	via	a	spectrophotometer	at	optical	densities	of	260	and	280	nm	(NanoDrop™	2000;	Thermo	Fisher	Scientific).	An	A260/A280	ratio	of	approximately	1.8-2	was	deemed	pure	and	acceptable	for	continuing	on.	Part	of	the	controlled	RNA	underwent	reverse	transcription	into	cDNA	via	the	SuperScript™	VILO™	MasterMix	protocol	(Thermo	Fisher	Scientific	11755-50).	To	amplify	and	quantify	the	cDNA,	samples	were	loaded	(in	duplicates,	which	were	later	averaged)	in	the	Applied	Biosystems™	ViiA	7	Real-Time	PCR	System	(Applied	Biosystems),	using	the	PowerUp™	SYBR®	Green	PCR	Master-Mix	kit	(Applied	Biosystems	A25780).	The	fold	changes	of	each	target	were	calculated	with	the	ΔΔCt	method	compared	to	SHAM	and	compared	to	the	housekeeping	gene	β-actin.						52	3.13.2 UPS	and	autophagy	targets	investigated	To	investigate	proteolytic	activity,	the	gene	expression	of	key	targets	of	UPS	and	autophagy	pathways	were	investigated.	The	chosen	targets	and	their	primer	sequences	are	reported	below	in	Table	3.2.	Table	3.2.	Targets	investigated	Pathway	 Target	 Primer	Sequence	(forward	and	reverse)	UPS	 MAFbx	 5’-ACTTCTCAGAGCGGCAGATCC-3’	5’-CTCTGGGTTGTTGGCCGT-3’		 MuRF1	 5’-GCAGGAATGCTCCAGTCGG-3’	5’-GTGAGCCCCGAACACCTT-3’	Autophagy	 ATG7	 5’-AGACCTTGAGCGTGCGTATG-3’	5’-AACTGCTACTCCATCTGTGGG-3’		 ATG12	 5’-CCCAGAAACAGCCATCCCA-3’	5’-TCACATAAATAAACAACTGCTCCGA-3’		 BECN1	 5’-CGTCGGGGCCTAAAGAATG-3’	5’-GCTCTCTCCTGGTTTCGCC-3’	Housekeeping	 b-actin	 5’-GGGAAATCGTGCGTGACT-3’																	5’-GCGGCAGTGGCCATCTC-3’		ATG7,	autophagy	related	7;	ATG12,	autophagy	related	12;	BECN1,	beclin	1;	MAFbx,	muscle	atrophy	F-box;	MuRF1,	muscle	RING-finger	protein	1.	3.14 Histology		Once	a	cross-sectional	disc	of	the	heart	at	mid-ventricular	level,	destined	to	histology,	was	excised	from	the	organism,	the	tissue	was	fixed,	dehydrated,	embedded	in	paraffin	(as	explained	in	Section	3.12.1.)	and	sectioned	at	7	µm	before	proceeding	to	staining.3	In	this	study,	we	used	immunofluorescence	to	visualize	target	proteins	and	study	LV	cardiomyocyte	morphology.		3.14.1 	Staining	for	cardiomyocyte	morphology		To	study	cardiomyocyte	morphology,	we	used	immunofluorescence	histology	on	a	cross-section	of	the	heart.	After	sectioning	our	tissue	at	a	depth	of	7	µm	per	slice	with	a	microtome,	the	deparaffinising	process	(with	different	concentrations	of	xylene	and	ethanol)	was	followed	by	rehydration,	antigen-retrieval	with	tri-sodium	citrate	buffer,	washes	with	PBS	with	Triton	(PBS-T	1%)	for	permeabilization	of					53	the	plasma	membrane	and	2-hour	blocking	of	the	tissue	(1%	bovine	serum	albumin	(BSA)	and	10%	normal	donkey	serum	(NDS);	for	the	prevention	of	non-specific	binding	of	antibodies).	The	primary	(on	for	overnight)	and	secondary	Abs	(on	for	2	hours)	solutions	(diluted	in	1%	BSA)	were	then	sequentially	added	to	the	slides.	Before	mounting	the	slides,	multiple	PBS-T	washes	were	performed	to	wash	off	any	excess	secondary	Abs	(3	x	5	min).	Finally,	DNA	was	stained	with	Hoechst’s	for	nuclear	staining	(3	x	5	min	with	PBS),	which	was	followed	by	multiple	required	washes	(3	x	5	min	with	PBS)	and	three	dips	in	distilled	water.		We	used	a	quadruple	stain	technique	to	visualize:	1)	the	cell	membranes	with	wheat	germ	agglutinin	(WGA	1:2000,	Thermo	Fisher	Scientific	1853992),	2)	the	Z-lines	with	a-actinin	(rabbit	primary	1:400,	Abcam	EP2529Y;	secondary	1:1000	donkey	anti-rabbit	AF546,	Jackson	ImmunoResearch	711-586-152),	3)	the	gap	junctions	located	at	the	intercalated	discs	with	connexin-43	(goat	primary	1:1000,	Cedarlane	NBP1-51938;	secondary	1:1000	donkey	anti-goat	AF647,	Jackson	ImmunoResearch	705-606-147),	and	4)	the	nuclei	with	Hoechst	stain	(Hoechst	33342	reagent	1:10000,	Thermo	Fischer	Scientific	1874027).	The	entire	protocol	is	a	three-day	process	and	results	in	a	quadruple	immunofluorescence	stain	which	serves	to	visualize,	under	a	confocal	microscope	(Zeiss	Axio	Observer,	equipped	with	a	Yokogawa	Spinning	Disk),	single	cardiomyocytes	longitudinally	or	in	a	cross-sectional	manner	in	the	myocardium,	both	depicted	in	Figure	3.3.	WGA	is	a	lectin,	which	binds	carbohydrates	located	on	the	plasma	membrane	of	all	mammalian	cells.172	Under	the	confocal	microscope,	the	cells	appear	to	be	delineated	by	green,	which	facilitates	width,	length	and	cross-sectional	area	(CSA)	measurements.	To	measure	cardiomyocyte	width,	we	stained	for	a-actinin,	a	protein	located	at	the	Z-lines	of	the	sarcomeres.13,14	It	functions	to	keep	the	thick	filaments	in	place	within	the	hexagonal	structure	of	a	sarcomeral	unit.13,14	The	neighboring	a-actinin	proteins	in	one	cell	form	a	linear	structure	across	the	cardiomyocyte,	allowing	us	to	measure	width.	It	is					54	important	to	ensure	the	red	line	being	measured	is	straight	and	not	bent.	If	bent,	it	could	possibly	be	two	overlapping	cardiomyocytes	and	lead	to	an	incorrect	measurement.	To	measure	cardiomyocyte	length,	connexin-43	is	stained	and	visualized.	Six	connexin-43	proteins	form	a	connexon	and	two	connexons	in	adjacent	cell	membranes	form	a	gap	junction	at	the	cardiac	intercalated	discs	(end-plates).2	As	discussed	earlier,	gap	junctions	serve	to	relay	signals	from	cell-to-cell;	in	cardiac	tissue	more	specifically,	they	communicate	electrical	signals	for	the	purpose	of	actuating	cardiac	contraction.2		3.14.2 Imaging		One	heart	section	per	rat	was	imaged.	A	minimum	of	10	images	per	section	were	taken	at	20x	magnification	with	a	confocal	microscope.	Both	longitudinally	(located	in	the	myocardium)	and	cross-sectionally	oriented	(located	in	the	sub-epicardium)	myocytes	were	imaged	to	measure	length	and	width,	and	CSA,	respectively.	All	imaging	was	performed	in	the	free	wall	of	the	LV.		3.14.3 Analysis		All	measurements	of	length,	width	and	CSA	were	performed	in	ImageJ	by	a	blinded	individual	(Fiji,	bundled	with	64-bit	Java	1.8.0	112,	version	1.52e).	All	cardiomyocytes	with	measurable	indices	and	confirmed	identity	(presence	of	a-actinin	in	the	cytoplasm)	were	quantified.	For	CSA	analysis,	to	ensure	consistent	measurement	along	the	cell,	it	must	have	had	a	red-stained	cytoplasm	and	a	centrally	located	nucleus.	We	have	used	a	method	to	automate	CSA	measurements	with	ImageJ	which	involved	an	image	conversion	to	grayscale	(8-bit)	then	the	application	of	a	B&W	threshold	and	dark-background.	This	allowed	us	to	automate	the	measurement	in	one-click.	For	accuracy	and	verification,	the	selection	of	the	cells	was	done	manually.		When	processing	the	heart,	we	could	not	control	for	which	stage	in	the	cardiac	cycle	the	cardiomyocytes	were	fixed	(i.e.,	contracted	or	relaxed)	and	how	the	cardiomyocytes	were	sectioned	(i.e.,					55	at	or	away	from	the	midline	of	the	cell)	which	could	affect	length	and	width	measurements,	respectively.	Therefore,	to	prevent	the	underestimation	of	cardiomyocyte	dimensions,	we	had	a	high	sample	size	per	group	and	analyzed	a	high	number	of	cells	per	rat.	In	total,	a	minimum	of	130	lengths,	210	widths	and	100	CSA	measurements	were	measured	per	rat.	All	cardiomyocyte	dimensions,	were	averaged	per	rat	and	then	per	group.	Cardiomyocyte	volume	(assumed	to	be	of	cylindrical	shape)	was	calculated	by	multiplying	CSA	and	length.			Figure	3.3.	Quadruple	immunofluorescent	stain	to	visualize	and	measure	cardiomyocyte	dimensions.	Wheat	germ	agglutinin,	green;	α-actinin,	red;	connexin-43,	pink;	Hoechst	nuclei	staining,	blue.	These	images	were	taken	with	a	confocal	microscope	at	a	magnification	of	x20	with	a	scale	bar	of	50µm.	Panel	A,	longitudinal	view	of	the	myocardial	free-wall	of	the	LV.	The	solid	and	dotted	lines	represent	a	cardiomyocyte	length	and	width,	respectively.	Panel	B,	cross-sectional	view	of	the	sub-epicardial/epicardial	free-wall	of	the	LV,	with	a	circled	representative	CSA	measurement.	3.15 Statistics			 All	descriptive	and	statistical	inferential	tests	were	performed	with	GraphPad	Prism	(version	6.0e)	with	set	a=0.05	(p<0.05).	Before	inputting	inferential	statistics,	we	performed	outlier	(Q=5%)	and	normality	tests.	Kurtosis	and	skewness	were	used	to	determine	if	the	data	sets	were	normal	or	not;	A	 B					56	values	between	±	1.96	were	deemed	normal.	Variables	which	showed	normality	were	analyzed	with	parametric	tests	and	the	ones	that	violated	normality	were	analyzed	with	non-parametric	tests.	For	demographic	data	(i.e.,	body	mass	at	pre-intervention	and	at	termination,	and	femur	length),	ANOVAs	were	performed	with	Tukey’s	post-hoc	test	(all	group	comparisons)	to	investigate	between-group	differences	as	body	mass	loss	can	affect	cardiac	atrophy	and	femur	length	was	used	for	standardization.	For	all	longitudinal	analyses	(i.e.,	echocardiography	pre-	and	post-intervention)	for	the	SHAM	and	7	days	T3-SCI	groups,	we	performed	two-way	repeated	measures	ANOVA	(time	x	group)	with	Dunnett’s	post-hoc	comparisons	(within-group)	and	Sidak’s	post-hoc	comparisons	(between-group).	For	histological	and	molecular	analyses,	all	replicates	per	rat	were	averaged.	All	PV	measures,	cardiomyocyte	dimensions	and	mean	target	fold	changes	were	statistically	analyzed	with	one-way	ANOVAs	for	between-subject	analyses	(group).	If	data	were	normal,	a	Dunnett’s	post-hoc	comparison	(all	groups	compared	to	SHAM)	was	performed.	If	data	were	non-normal,	a	Kruskal-Wallis	tests	with	Dunn’s	post-hoc	comparisons	was	performed	(all	groups	compared	to	SHAM).		3.16 Exclusion	of	data	and	standardization	Out	of	the	49	rats,	a	total	of	45	rats	reached	termination	day	and	four	died	prematurely	during	or	closely	after	SCI	surgery.	This	totals	the	mortality	rate	of	complete	T3-SCI	in	this	study	to	be	8.2%,	which	is	in	the	range	previously	reported	in	similar	experiments	(0-10%).156	One	rat	was	excluded	from	all	analyses	due	to	poor	health	before	and	at	termination.	This	particular	rat	showed	clinical	signs	of	a	bladder	infection	and	possible	sepsis	(i.e.,	lethargy,	pale	skin,	and	an	extensive	blood	clot	found	in	its	bladder)	which	can	confound	results.	Partial	and	complete	data	were	obtained	and	utilized	from	44	rats.	Special	exclusion	criteria	for	certain	analyses	can	be	found	in	the	following	paragraphs	and	all	sample	sizes	per	type	of	analysis	in	their	relevant	result	sections.						57	PV	data	were	not	collected	for	the	12	hours	T3-SCI	group.	This	was	due	to	some	rats	having	arrhythmic	hearts	immediately	following	the	high-level	SCI,78	which	made	it	practically	impossible	to	catheterize	the	heart	without	damaging	the	apparatus.	Furthermore,	there	are	other	potential	ethical	implications	of	conducting	multiple	surgeries	in	such	a	close	time	frame	to	each	other.	No	additional	rats	were	excluded	for	PV	analysis.		For	histology,	cardiac	tissue	was	collected	precisely	at	planned	termination	times	immediately	after	completion	of	the	in	vivo	experiments.	We	were	able	to	collect	and	fix	45	hearts	for	histology.	Out	of	these	45	samples,	five	rats	were	excluded	for	the	following	reasons.	One	rat	was	excluded	from	all	analyses	from	this	study,	as	above-mentioned,	and	four	other	rats	were	excluded	due	to	poor	perfusion	quality,	causing	late	fixation	of	the	tissue	post-mortem.	As	fresh	tissue	must	be	fixed	immediately	after	death,	we	decided	to	not	trust	the	tissue	as	lack	of	blood	supply	to	the	organ	will	cause	a	decline	in	tissue	quality.	All	cardiomyocyte	dimensional	data	were	standardized	to	femur	length,	which	is	an	approach	used	in	our	laboratory	previously.72,99	Body	mass	highly	fluctuates	following	SCI	due	to	administration	of	opioids	and	surgery,	therefore	femur	length	was	deemed	more	stable	and	accurate	as	a	method	of	standardization.	Our	method	of	standardization	followed	the	instructions	in	Hagdorn	et	al.,	which	suggested	dividing	one-dimensional	measurements	by	femur	length	(cm),	area	(two-dimensional)	measurements	by	squared	femur	length	(cm2)	and	volumetric	(three-dimensional)	measurements	by	cubed	femur	length	(cm3).173	No	outliers	for	cardiomyocyte	dimensions	were	detected.			 					58	 Results	4.1 Demographics	4.1.1 Demographics	at	termination	Demographics	obtained	at	termination	are	provided	in	Table	4.1.	Body	mass	was	similar	between	all	groups	at	pre-intervention	(p=0.175).	At	termination,	body	mass	differed	between	groups	due	to	differential	termination	time	points	along	the	acute	setting	post-SCI	(p=0.006).	As	body	mass	fluctuated	significantly	acutely	post-SCI,	all	dimensional	indices	were	standardized	to	femur	length	as	it	did	not	differ	in	any	of	the	groups	(p=0.638).173		Table	4.1.	Group	demographics	at	termination		 SHAM	 12	hours	 1	day	 3	days	 5	days	 7	days	 p	Body	mass	at	pre-intervention	(g)	328.6	(3.0)	348.9	(11.4)	361.8	(16.8)	325.3	(5.5)	359.5	(14.3)	354.6	(9.6)	0.175	Body	mass	at	termination	(g)	334.2	(3.1)	370.5	(15.8)	392.2	(19.7)	320.4	(4.9)	333.3	(14.9)	315.3	(11.9)*	0.006	Femur	length	(cm)	 3.54	(0.02)	3.54		(0.05)	3.63		(0.05)	3.53	(0.04)	3.57		(0.05)	3.58		(0.04)	0.638	Values	are	means	(standard	error	of	the	mean;	SE).	The	last	column	indicates	the	p	value	for	one-way	ANOVAs.	Post-hoc,	*p<0.05	versus	1	day.	For	body	mass	at	both	pre-intervention	and	at	termination,	sample	sizes	are	as	follows:	n=6	for	SHAM,	n=9	for	12	hours	T3-SCI,	n=7	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=7	for	5	days	T3-SCI	and	n=8	for	7	days	T3-SCI.	The	same	sample	sizes	apply	for	femur	length	data,	except	for	the	3	days	T3-SCI	group	which	was	n=7.	4.1.2 Body	mass	along	the	acute	timeline	in	the	7	days	T3-SCI	and	SHAM	groups	All	means,	SEs	and	p	values	for	body	mass	obtained	along	the	acute	timeline	in	SHAM	and	7	days	T3-SCI	rats	are	reported	and	illustrated	in	Table	4.2.	and	Figure	4.1.A.	There	was	an	interaction	effect	between	time	and	group	for	body	mass	(p<0.001),	as	well	as	a	main	effect	for	time	(p<0.001).	Post-hoc	tests	revealed	that	body	mass	was	significantly	higher	in	T3-SCI	rats	compared	to	SHAM	rats	at	2	days	post-surgery	(p=0.039)	but	did	not	differ	at	pre-injury	or	6	days	post-injury	between	the	two	groups.						59	4.2 In-vivo	echocardiography	data	–	Temporal	cardiac	volumetric	and	functional	indices	following	the	7	days	T3-SCI	and	SHAM	groups		4.2.1 Heart	rate	Changes	in	HR	from	pre-	to	post-surgery	are	reported	and	illustrated	in	Table	4.2	and	Figure	4.1.B.,	respectively.	There	was	an	interaction	effect	between	time	and	group	for	HR	(p<0.001),	as	well	as	a	main	effect	for	group	(p=0.002).	Post-hoc	analyses	revealed	that	HR	was	significantly	lower	in	the	T3-SCI	rats	compared	to	the	SHAM	rats	at	1,	2	and	4	days	post-surgery	(all	p<0.001),	and	HR	was	not	different	from	SHAM	at	day	6	post-SCI	(p=0.163).	4.2.2 Volumetric	cardiac	indices	and	systolic	function		 EDV,	ESV,	SV,	CO,	EF	and	FS	from	pre-	to	post-surgery	are	reported	and	illustrated	in	Table	4.2.	and	Figure	4.1.,	respectively.	There	was	a	significant	interaction	effect	between	time	and	group	for	EDV	and	SV	(both	p<0.001),	whereby	EDV	and	SV	were	initially	higher	in	T3-SCI	vs.	SHAM	pre-surgery	but	subsequently	became	lower	than	SHAM	by	day	6	post-SCI	(both	p<0.05).	There	was	an	interaction	effect	between	time	and	group	for	ESV	(p=0.006).	Post-hoc	tests	revealed	a	lower	ESV	in	the	SHAM	group	at	pre-surgery	versus	the	T3-SCI	group	(p=0.056),	while	ESV	did	not	differ	between	the	groups	at	any	other	time-point	post-surgery	(all	p>0.658).	In	the	T3-SCI	group,	ESV	was	reduced	acutely	post-SCI	at	all	time-points	versus	their	pre-injury	values	(all	p<0.001).	There	was	an	interaction	effect	between	time	and	group	for	CO	(p=0.001),	and	post-hoc	tests	revealed	significantly	reduced	CO	at	4	and	6	days	in	the	T3-SCI	group	compared	to	SHAM	(both	p<0.05).	There	were	no	other	changes	of	note	in	any	other	indices	of	cardiac	function	on	echocardiography.							60	P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s2 5 03 0 03 5 04 0 04 5 05 0 0T im eEDV	(µL)In te ra c t io n : 	* * * * 	< 0 .0 0 1T im e : 	0 .2 7 4S u rg e r y : 	0 .7 1 5***P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s05 01 0 01 5 02 0 0T im eESV	(µL)In te ra c t io n : 	* * 	0 .0 0 6T im e :	* * * * 	< 0 .0 0 1S u rg e r y : 	0 .8 8 6P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s1 5 02 0 02 5 03 0 03 5 04 0 0T im eSV	(µL)In te ra c t io n : 	* * * * 	< 0 .0 0 1T im e : 	* 	0 .0 4 2S u rg e r y : 	0 .7 1 3***P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s05 01 0 01 5 0T im eCO	(mL/min)In te ra c t io n : 	* * 	0 .0 0 1T im e : 	0 .1 3 0S u rg e r y : 	0 .0 8 8** *P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s5 06 07 08 09 0T im eEF	(%)In te r a c t io n : 	0 .1 3 3T im e :	* * 	0 .0 0 2S u rg e r y : 	0 .5 0 6P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s01 02 03 0T im eFS	(%)In te r a c t io n : 	0 .1 4 0T im e : 	0 .0 5 5S u rg e r y : 	0 .1 6 5P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s2 5 03 0 03 5 04 0 04 5 0T im eHR	(bpm) In te ra c t io n : 	* * * * 	< 0 .0 0 1T im e : 	0 .6 7 2S u rg e r y : 	* * 	0 .0 0 2**********P re -surge ry1 	d ay2 	d ay s4 	d ay s6 	d ay s2 5 03 0 03 5 04 0 04 5 0T im eBody	mass	(g)S H A MT 3 -S C IS u rg e r y* In te ra c t io n : 	* * * * < 	0 .0 0 1T im e :	* * * * 	< 0 .0 0 1S u rg e r y : 	0 .2 1 3	Figure	4.1.	Body	mass	and	echocardiographic	indices	measured	along	the	acute	timeline	of	rats	that	have	undergone	SHAM	and	T3-SCI	surgeries.	A)	Body	mass;	B)	HR,	heart	rate;	C)	EDV,	end-diastolic	volume;	D)	ESV,	end-systolic	volume;	E)	SV,	stroke	volume;	F)	CO,	cardiac	output;	G)	EF,	ejection	fraction;	H)	FS,	fractional	shortening.	Group	means	are	plotted	as	symbols,	while	standard	errors	of	the	mean	are	represented	with	error	bars	(mean	±	SE).	The	p	values	for	the	repeated	measures	two-way	ANOVAs	are	included	by	each	variable.	For	simplicity	we	have	shown	only	the	significant	post-hoc	comparisons	with	symbols	on	the	figures,	*p<0.05,	**p<0.01,	***p<0.001	and	****p<0.0001.	All	sample	sizes,	means	and	SEs	are	provided	in	Table	4.2.	 		A	 	B	C	 D	F	E	G	 H			Table	4.2.	Parasternal	long-axis	volumetric	and	functional	indices	following	SHAM	and	T3-SCI	along	the	acute	timeline		 SHAM	 T3-SCI	 p		 Pre-surgery	1	day	 2	days	 4	days	 6	days	Pre-surgery	1	day	 2	days	 4	days	 6	days	 Time	 Surgery	 Interaction	Body	mass	(g)	 329	(3)	 349	(3)**	 339	(3)**	 338	(4)*	 332	(3)	 355	(10)	 381	(12)**	 374	(10)**†	 339	(11)**	 320	(10)**	 <0.001	 0.213	 <0.001	HR	(bpm)	 339	(17)	 382	(7)*	 373	(15)	 397	(6)*	 364	(11)	 351	(8)	 295	(18)**††	 294	(16)**††	 300	(19)*††	 321	(9)	 0.672	 0.002	 <0.001	Volumes	 	 	 	 	 	 	 	 	 	 	 	 	EDV	(µL)	 329	(11)	 338	(37)	 400	(15)	 394	(24)	 431	(24)*	 449	(18)††	 412	(27)	 411	(28)	 324	(21)**	 335	(17)**†	 0.274	 0.715	 <0.001	ESV	(µL)	 111	(16)	 86	(15)	 107	(14)	 87	(8)	 111	(15)	 155	(7)	 96	(11)**	 85	(12)**	 74	(9)**	 101	(10)**	 <0.001	 0.886	 0.006	Systolic	function	 	 	 	 	 	 	 	 	 	 	 	SV	(µL)	 218	(10)	 252	(25)	 293	(12)*	 307	(22)*	 320	(13)**	 294	(13)†	 315	(23)	 325	(18)	 250	(17)	 233	(17)*†	 0.042	 0.713	 <0.001	CO	(mL/min)	 73.2	(2.7)	 96.5	(10.1)	 109.9	(8.2)	 120.9	(8.1)*	 116.5	(7.0)*	 103.2	(4.5)	 90.8	(4.4)	 117.4	(20.7)	 75.8	(8.0)†	 74.3	(4.2)†	 0.130	 0.088	 0.001	EF	(%)	 66.8	(4.4)	 68.7	(6.6)	 73.4	(2.8)	 77.9	(1.9)*	 74.6	(2.4)	 65.5	(1.0)	 76.6	(2.2)*	 79.8	(1.9)**	 77.4	(2.2)*	 69.5	(2.8)	 0.002	 0.506	 0.133	FS	(%)	 16.5	(1.6)	 15.3	(2.0)	 16.2	(1.4)	 19.8	(1.4)	 18.3	(1.4)	 15.2	(1.6)	 20.3	(1.4)	 22.0	(2.5)*	 22.1	(2.4)*	 17.8	(1.4)	 0.055	 0.165	 0.140	Values	are	means	(SE)	with	n=6	for	SHAM	and	n=8	for	T3-SCI.	CO,	cardiac	output;	EDV,	end-diastolic	volume;	EF,	ejection	fraction;	ESV,	end-systolic	volume;	FS,	fractional	shortening;	SV,	stroke	volume.	The	last	column	indicates	the	p	values	for	two-way	repeated	measures	ANOVAs.	Within-group	post-hoc	tests	versus	pre-injury,	*p<0.05	and	**p<0.001.	Between-group	post-hoc	tests,	†p<0.05	and	††p<0.001.						62	4.3 In-vivo	catheterization	data	–	Cardiovascular	functional	and	pressure-volume	indices	4.3.1 Basal	arterial	hemodynamics	Graphical	representations	of	all	hemodynamic	variables	can	be	found	in	Figure	4.2.	Means,	SEs	and	p	values	are	reported	in	Table	4.3.	Systolic	blood	pressure	(SBP)	was	significantly	decreased	acutely	following	T3-SCI	(p=0.001)	compared	to	SHAM.	Post-hoc	analyses	revealed	significant	reductions	at	3	days	(p=0.049),	5	days	(p<0.001)	7	days	(p=0.014),	and	a	trend	for	reduced	SBP	at	1	day	post-SCI	compared	to	SHAM	(p=0.068).	DBP	was	significantly	reduced	post-SCI	versus	SHAM	(p=0.026).	However,	post-hoc	testing	versus	SHAM	revealed	that	the	only	group	which	showed	a	significant	decrease	in	DBP	was	the	5	days	T3-SCI	group	(p=0.044),	while	all	other	time-points,	including	the	last	time-point	(7	days),	showed	no	significant	differences	in	DBP	(all	p>0.612).	Mean	arterial	pressure	(MAP)	was	significantly	decreased	acutely	following	T3-SCI	(p=0.002)	with	post-hoc	tests	versus	SHAM	revealing	significant	lower	MAPs	in	the	5	and	7	days	T3-SCI	groups	(both	p<0.05).			 		 					63	SHAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s05 01 0 01 5 0SBP	(mmHg)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s02 04 06 08 01 0 0DBP	(mmHg)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s05 01 0 01 5 0MAP	(mmHg)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s01 0 02 0 03 0 04 0 05 0 0HR	(bpm)*****0.068 ********	Figure	4.2.	Basal	hemodynamics	following	SHAM	surgery	and	at	different	time-points	following	T3-SCI.	SHAM	rats	were	terminated	at	7	days	post-surgery.	A)	SBP,	systolic	blood	pressure;	B)	DBP,	diastolic	blood	pressure;	C)	MAP,	mean	arterial	pressure;	D)	HR,	heart	rate.	Individual	rat	means	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	One-way	ANOVAs	post-hoc	tests	versus	SHAM,	*p<0.05	and	***p<0.001.	All	sample	sizes,	means	and	standard	errors	are	provided	in	Table	4.3.		Table	4.3.	Hemodynamic	responses	to	SHAM	surgery	and	T3-SCI	at	different	termination	time	points		 SHAM	 1	day	 3	days	 5	days	 7	days	 p	SBP	(mmHg)	 115.2	(3.2)	 89.3	(2.9)	 88.8	(4.5)*	 76.3	(3.5)***	 85.9	(2.2)*	 0.001	DBP	(mmHg)	 62.4	(4.2)	 61.8	(2.8)	 61.2	(4.1)	 48.3	(2.5)*	 54.1	(2.7)	 0.026	MAP	(mmHg)	 85.3	(4.0)	 75.1	(2.7)	 74.8	(4.8)	 60.1	(2.9)***	 67.2	(3.0)*	 0.002	HR	(bpm)	 374	(12)	 278	(15)*	 276	(41)*	 271	(24)*	 334	(26)	 0.030	A	 B	C	 D					64	Values	are	means	(SE).	DBP,	diastolic	blood	pressure;	HR,	heart	rate;	MAP,	mean	arterial	pressure;	SBP,	systolic	blood	pressure.	The	last	column	indicates	the	p	value	for	one-way	ANOVAs.	Post-hoc	tests	versus	SHAM,	*p<0.05	and	***p<0.001.	Sample	sizes	are	as	follows:	n=6	for	SHAM,	n=7	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=6	for	5	days	T3-SCI	and	n=6	for	7	days	T3-SCI.	4.3.2 Basal	cardiac	pressure-volume	responses	All	load-dependent	and	-independent	means,	SEs	and	p	values	are	reported	in	Table	4.4.	Averaged	PV	loops	per	group	and	representative	IVC	occlusions	are	illustrated	in	Figure	4.3.	4.3.2.1 Left-ventricular	pressure-volume	derived	load-independent	function	ESPVR,	used	to	infer	systolic	function,	was	significantly	decreased	acutely	post-SCI	(p=0.005)	(Figure	4.4.A.).	Post-hoc	analyses	revealed	that	at	all	T3-SCI	time-points,	except	3	days,	ESPVR	was	significantly	lower	versus	SHAM	(all	p<0.05).	Nonetheless,	the	3	days	T3-SCI	group	tended	to	have	a	lower	ESPVR	versus	SHAM	(p=0.051).	Contrarily	to	ESPVR,	PRSW	was	not	seen	to	differ	in	any	of	the	groups	(p=0.531).	4.3.2.2 Left-ventricular	pressure-volume	derived	load-dependent	function	All	load-dependent	variables	which	infer	systolic	function	were	significantly	different	following	T3-SCI.	Pmax	(p=0.005),	Pdev	(p<0.001)	and	dP/dtmax	(p=0.006)	were	significantly	reduced	at	all	T3-SCI	time-points	versus	SHAM	(Figure	4.4.C.	and	D.).	Unstandardized	(Figure	4.4.B.)	and	standardized	(to	femur	length)	SW	were	significantly	lower	in	the	3,	5	and	7	days	T3-SCI	groups	versus	SHAM	(SW	p=0.004;	standardized	SW	p=0.013).	Although	Ea	was	not	significantly	different	in	any	of	the	groups	(p=0.145),	it	tended	to	be	lower	in	the	1	day	T3-SCI	group	versus	SHAM	(post-hoc:	p=0.059)	(Figure	4.4.E.).	Furthermore,	Ea/ESPVR,	the	ventricular	vascular	coupling	ratio,	was	significantly	increased	in	the	1,	5	and	7	days	T3-SCI	groups	versus	SHAM	(p=0.011)	(Figure	4.4.F.).	All	T3-SCI	time-points,	except	3	days	(post-hoc:	p=0.063),	showed	significant	or	strongly	suggested	reduced	-dP/dtmin	values	versus	SHAM	(p=0.006)	(Figure	4.5.A).	On	the	contrary,	other	load-				65	dependent	variables	which	infer	diastolic	function	showed	no	significant	changes	in	the	T3-SCI	groups	versus	SHAM	(Ped	p=0.766;	tau	p=0.224)	(Figure	4.5.B).	0 5 0 1 0 0 1 5 0 2 0 0 2 5 005 01 0 01 5 0SH AMV o lu m e 	(µ L )Pressure	(mmHg)0 5 0 1 0 0 1 5 0 2 0 0 2 5 005 01 0 01 5 0T3 -S C IV o lu m e 	(µ L )Pressure	(mmHg)0 5 0 1 0 0 1 5 0 2 0 0 2 5 002 55 07 51 0 01 2 5V o lu m e 	(µ L )Pressure	(mmHg)7 	d a y s 	T 3 -S C I5 	d a y s 	T 3 -S C I3 	d a y s 	T 3 -S C IS H A M1 	d a y 	T 3 -S C I	Figure	4.3.	Averaged	baseline	pressure-volume	loops	and	representative	inferior	vena	cava	occlusions	obtained	via	LV	catheterization.	A)	Averaged	baseline	PV	loops	of	all	experimental	groups.	Error	bars	represent	SEM.	B)	Representative	IVC	occlusion	following	SHAM	surgery.	C)	Representative	IVC	occlusion	following	T3-SCI	in	the	acute	phase	following	injury.	The	dotted	lines	represent	representative	ESPVR	values.	ESPVR,	end-systolic	pressure-volume	relationship;	IVC,	inferior	vena	cava;	LV,	left-ventricle;	PV,	pressure-volume;	SCI,	spinal	cord	injury.			A	B	 C					66	SHAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s01234ESPVR	(mmHg/µL)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s01 02 03 0SW	(mmHg*mL)**0 .051**SHAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s02 0 0 04 0 0 06 0 0 08 0 0 01 0 0 0 0dP/dt max(mmHg/s)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s6 08 01 0 01 2 01 4 0Pmax	(mmHg)***************SHAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s0 .00 .20 .40 .60 .8Ea	(mmHg/µL)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s0 .00 .20 .40 .60 .8Ea/ESPVR*****	Figure	4.4.	Systolic	function	following	SHAM	surgery	and	at	different	times	points	following	T3-SCI.	A)	ESPVR,	end-systolic	pressure-volume	relationship;	B)	SW,	stroke	work;	C)	Pmax,	maximum	pressure;	D)	A	 B	C	 D	E	 F					67	dP/dtmax,	maximal	rate	of	systolic	pressure	increment;	E)	Ea,	arterial	elastance;	F)	Ea/ESPVR,	ventricular	vascular	coupling	ratio.	Individual	rat	means	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	One-way	ANOVAs	post-hoc	tests	versus	SHAM,	*p<0.05	and	**p<0.01.	All	sample	sizes,	means	and	standard	errors	are	provided	in	Table	4.4.		SHAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s-6 0 0 0-4 0 0 0-2 0 0 00-dP/dt min	(mmHg/s)S HAM1 	d ay3 	d ay s5 	d ay s	7 	d ay s051 01 52 02 5Tau	(ms)***0 .063	Figure	4.5.	Diastolic	function	following	SHAM	surgery	and	at	different	times	points	following	T3-SCI.	A)									-dP/dtmin,	maximal	rate	of	diastolic	pressure	decrement;	B)	Tau,	diastolic	time	constant.	Individual	rat	means	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	One-way	ANOVAs	post-hoc	tests	versus	SHAM,	*p<0.05	and	**p<0.01.	All	sample	sizes,	means	and	standard	errors	are	provided	in	Table	4.4.		A	 B					68	Table	4.4.	Cardiac	functional	responses	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points		 SHAM	 1	day	 3	days	 5	days	 7	days	 p	Systolic	function	 	 	 	 	 	 	ESPVR	(mmHg/µL)	 2.82	(0.17)	 1.05	(0.22)**	 1.29	(0.22)	 1.10	(0.18)*	 1.10	(0.16)*	 0.005	SW†	(mmHg*mL)	 20.42	(0.86)	 18.62	(1.51)	 14.42	(1.89)*	 13.17	(1.23)**	14.37	(1.11)*	 0.004	Standardized	SW†	(mmHg*mL/cm3)		4.58	(0.16)	 3.90	(0.29)	 3.34	(0.46)	 2.92	(0.37)**	 3.12	(0.32)*	 0.013	Pmax†	(mmHg)	 118	(2)	 92	(4)*	 90	(5)*	 85	(3)**	 92	(2)*	 0.005	Pdev†	(mmHg)	 119	(2)	 91	(2)****	 91	(6)****	 88	(3)****	 91	(1)****	 <0.001	dP/dtmax	†	(mmHg/s)	8383	(230)	 5475	(320)*	 5241	(745)**	 4935	(472)**	 5765	(481)*	 0.006	TPR	(mmHg/mL)	 1.32	(0.21)	 1.69	(0.20)	 1.72	(0.34)	 1.82	(0.26)	 1.38	(0.14)	 0.464	Ea†	(mmHg/µL)	 0.43	(0.01)	 0.32	(0.02)	 0.45	(0.07)	 0.39	(0.05)	 0.36	(0.05)	 0.145	Ea/ESPVR	 0.16	(0.01)	 0.36	(0.06)**	 0.29	(0.04)	 0.36	(0.04)**	 0.36	(0.06)*	 0.011	PRSW†	(mmHg)	 122	(7)	 121	(16)	 121	(9)	 106	(10)	 141	(17)	 0.531	Diastolic	function	 	 	 	 	 	 	-dP/dtmin	(mmHg/s)	-4957	(98)	 -3094	(135)*	 -3361	(432)	 -2750	(286)**	 -3201	(83)	 0.006	Ped†	(mmHg)	 4.49	(1.46)	 5.03	(3.73)	 2.12	(1.45)	 1.47	(1.00)	 3.19	(2.63)	 0.766	Tau	(ms)	 7.58	(0.35)	 10.84	(2.01)	 12.56	(2.36)	 9.57	(0.50)	 10.85	(1.61)	 0.224	Values	are	means	(SE).	Standardized	SW	was	divided	by	femur	length3	(cm3).173	dP/dtmax,	maximal	rate	of	systolic	pressure	increment;	-dP/dtmin,	maximal	rate	of	diastolic	pressure	decrement;	Ea,	arterial	elastance;	Ea/ESPVR,	ventricular	vascular	coupling	ratio;	ESPVR,	end-systolic	pressure-volume	relationship;	Pdev,	developed	pressure;	Ped,	end-diastolic	pressure;	Pmax,	maximum	pressure;	PRSW,		preload	recruitable	stroke	work;	SW,	stroke	work;	Tau,	diastolic	time	constant;	TPR,	total	peripheral	resistance.	The	last	column	indicates	the	p	value	for	one-way	ANOVAs.	Post-hoc	tests	versus	SHAM,	*p<0.05,	**p<0.01	and	****p<0.0001.	†,	sample	sizes	are	as	follows:	n=6	for	SHAM,	n=7	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=6	for	5	days	T3-SCI	and	n=6	for	7	days	T3-SCI.For	ESPVR	and	PRSW:	n=6	for	SHAM,	n=7	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=5	for	5	days	T3-SCI	and	n=6	for	7	days	T3-SCI.	For	-dP/dtmin:	n=6	for	SHAM,	n=7	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=6	for	5	days	T3-SCI	and	n=5	for	7	days	T3-SCI.	For	tau:	n=6	for	SHAM,	n=6	for	1	day	T3-SCI,	n=6	for	3	days	T3-SCI,	n=5	for	5	days	T3-SCI	and	n=6	for	7	days	T3-SCI.		4.4 Molecular	data	–	Gene	analysis	for	protein	degradation	pathways	All	sample	sizes,	mean	fold	changes	and	SEs	per	group	for	all	UPS	and	autophagy	targets	are	reported	in	Table	4.6.		The	following	results	describe	an	increase	in	UPS	and	no	changes	in	autophagy	gene	expression	in	LV	tissue	acutely	post-SCI.					69	Table	4.5.	Quantitative	real-time	PCR	targets,	primers	and	fold	changes	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points		 SHAM	 12	hours	 1day	 3	days	 5	days	 7	days	 p	UPS	 	 	 	 	 	 	 	MAFbx	 1.06	(0.15)	 3.57	(0.48)***	 2.33	(0.51)	 2.26	(0.19)*	 1.94	(0.22)	 1.73	(0.21)	 0.005	MuRF1	 1.02	(0.10)	 1.38	(0.10)	 1.38	(0.23)	 1.49	(0.17)	 1.66	(0.28)	 1.03	(0.33)	 0.288	Autophagy	 	 	 	 	 	 	 	ATG7	 1.03	(0.12)	 1.64	(0.16)	 1.36	(0.23)	 1.30	(0.08)	 1.32	(0.21)	 1.59	(0.32)	 0.355	ATG12	 1.01	(0.09)	 1.20	(0.13)	 1.11	(0.16)	 1.07	(0.11)	 1.14	(0.20)	 1.43	(0.25)	 0.716	BECN1	 1.01	(0.06)	 1.07	(0.15)	 1.21	(0.24)	 1.05	(0.10)	 1.04	(0.14)	 1.14	(0.11)	 0.964	Values	are	mean	fold	changes	(SE)	with	n=6	for	all	groups.	ATG7,	autophagy	related	7;	ATG12,	autophagy	related	12;	BECN1,	beclin	1;	MAFbx,	muscle	atrophy	F-box;	MuRF1,	muscle	RING-finger	protein	1.	The	last	column	indicates	the	p	value	for	one-way	ANOVAs.	Post-hoc	tests	versus	SHAM,	*p<0.05	and	***p<0.001.				70	4.4.1 UPS		 The	fold	changes	of	UPS	marker,	MAFbx,	were	significantly	increased	following	T3-SCI	by	approximately	3.37-fold	versus	SHAM	(p=0.005)	(Figure	4.8.A.).	Post-hoc	analyses	revealed	that	MAFbx	levels	were	significantly	increased	compared	to	SHAM	at	12	hours	and	3	days	post-SCI	(both	p<0.05)	(Figure	4.8.A.).	MAFbx	peaked	at	12	hours	and	tended	to	gradually	decrease	back-down	with	time	(Figure	4.8.A.).	The	fold	changes	of	UPS	marker,	MurF1,	were	not	significantly	changed	in	any	of	the	T3-SCI	groups	versus	SHAM	(p=0.288)	(Figure	4.8.B.).	However,	MurF1	tended	to	be	up-regulated	in	all	but	one	animal	acutely	post-SCI	with	a	gradual	increase	starting	at	the	first	time-point,	12	hours,	and	a	peak	at	5	days	post-SCI	versus	SHAM	(Figure	4.8.B.).	MuRF1	appeared	to	decrease	back	towards	SHAM	levels	by	7	days	post-SCI	(Figure	4.8.B.).		SHAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s0246MAFbx	fold	change* * **SHAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s0 .00 .51 .01 .52 .02 .5MuRF1	fold	change	Figure	4.6.	RNA	fold	changes	of	UPS	targets	following	SHAM	surgery	and	T3-SCI	at	different	time	points	along	the	acute	spectrum	(n=6).	A)	MAFbx,	muscle	atrophy	F-box;	B)	MuRF1,	muscle	RING-finger	protein	1.	Individual	rat	mean	fold	changes	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	All	means,	standard	errors	and	p	values	are	provided	in	Table	4.6.	MAFbx,	one-way	ANOVA,	Kruskal-Wallis	test	(p=0.005)	and	Dunn’s	post-hoc,	*p<0.05	and	***p<0.001.	A	 B					71	4.4.2 Autophagy		The	fold	changes	of	autophagy	markers,	ATG7,	ATG12	and	BECN1,	were	not	significantly	changed	in	LV	cardiac	tissue	at	any	time-point	following	T3-SCI	versus	SHAM	(all	p>0.355)	(Figure	4.9.).		SHAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s0123ATG7	fold	changeS HAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s0 .00 .51 .01 .52 .02 .5ATG12	fold	changeS HAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s0 .00 .51 .01 .52 .02 .5BECN1	fold	change	Figure	4.7.	RNA	fold	changes	of	autophagy	targets	following	SHAM	surgery	and	T3-SCI	at	different	time	points	along	the	acute	spectrum	(n=6).	A)	ATG7,	autophagy	related	7;	B)	ATG12,	autophagy	related	12;	C)	BECN1,	beclin	1.	Individual	rat	mean	fold	changes	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	All	means,	standard	errors	and	p	values	are	provided	in	Table	4.6.	A	 B	 C				4.5 Histological	data	–	Cardiomyocyte	dimensions	All	final	sample	sizes	and	histological	measurements	for	all	groups	are	reported	in	Table	4.5.		All	longitudinally	oriented	cardiomyocyte	data	(i.e.,	length	and	width)	pertained	to	the	myocardial	layer	of	the	LV	free-wall,	while	cross-sectionally	oriented	data	(i.e.,	CSA)	pertained	to	the	sub-epicardial	and	epicardial	layers	of	the	LV	free-wall.	No	significant	atrophy	was	found	in	any	of	the	SCI	groups	versus	the	SHAM	group.	However,	trends	will	be	discussed	in	the	following	sections.	Table	4.6.	LV	myocardial	cardiomyocyte	dimensions	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points		 SHAM	 12	hours	 1	day	 3	days	 5	days	 7	days	 p	Standardized	 	 	 	 	 	 	 	Length	(µm/cm)	 25.64	(0.80)	24.86	(0.76)	24.82	(0.56)	23.95	(0.77)	23.93	(0.70)	23.49	(0.63)	 0.328	Width	(µm/cm)	 4.42	(0.29)	 4.28	(0.18)	 3.91	(0.24)	 4.07	(0.15)	 4.29	(0.18)	 4.05	(0.15)	 0.609	Length	to	width	ratio	5.91	(0.39)	 5.91	(0.34)	 6.46	(0.33)	 5.93	(0.33)	 5.62	(0.29)	 5.61	(0.24)	 0.503	CSA	(µm2/cm2)	 35.2	(4.8)	 29.1	(2.4)	 25.2	(1.2)	 32.2	(3.5)	 31.0	(3.7)	 27.0	(2.2)	 0.207	Volume	(µm3/cm3)	 927	(122)	 684	(69)	 623	(28)	 762	(77)	 748	(98)	 622	(49)	 0.146	Unstandardized	 	 	 	 	 	 	 	Length	(µm/cm)	 90.79	(2.48)	87.72	(1.76)	89.95	(2.15)	84.88	(3.17)	86.26	(1.70)	84.10	(2.07)	 0.264	Width	(µm/cm)	 15.66	(1.00)	15.15	(0.67)	14.22	(0.96)	14.41	(0.58)	15.49	(0.71)	14.49	(0.49)	0.763	CSA	(µm2/cm2)	 439	(57)	 375	(32)	 330	(13)	 395	(43)	 400	(37)	 344	(26)	 0.368	Volume	(µm3/cm3)	40716	(5024)	31453	(2725)	30585	(417)	32683	(3305)	34446	(3108)	28264	(1818)	 0.189	Standardized	values	were	corrected	to	femur	length.173	Values	are	means	(SE).	CSA,	cross-sectional	area.	The	last	column	indicates	the	p	value	for	one-way	ANOVAs.	Variables	obtained	via	the	longitudinal	view	(standardized	and	unstandardized	length	and	width)	had	sample	sizes	of:	n=6	for	SHAM,	n=9	for	12	hours,	n=7	for	1	day,	n=5	for	3	days,	n=5	for	5	days	and	n=8	for	7	days.	Variables	obtained	via	the	cross-sectional	view	(CSA	and	standardized	volume)	had	sample	sizes	of:	n=5	for	SHAM,	n=5	for	12	hours,	n=7	for	1	day,	n=4	for	3	days,	n=5	for	5	days	and	n=7	for	7	days.	Unstandardized	volume	had	sample	sizes	of:	n=5	for	SHAM,	n=5	for	12	hours,	n=6	for	1	day	T3-SCI,	n=4	for	3	days	T3-SCI,	n=5	for	5	days	T3-SCI	and	n=7	for	7	days	T3-SCI.					73	4.5.1 Cardiomyocyte	length	and	width		 There	was	no	significant	cardiomyocyte	atrophy	in	either	length	or	width,	standardized	to	femur	length,	following	T3-SCI	at	any	time-point	along	the	acute	spectrum	versus	SHAM	(both	p>0.328)	(Figure	4.6.C.).	Post-hoc	analyses	revealed	that	standardized	length	tended	to	gradually	decrease	along	the	acute	spectrum	with	the	lowest	value	at	7	days	post-SCI	versus	SHAM	(p=0.145).	Unstandardized	length	showed	similar	trends	to	its	standardized	counterpart	(p=0.264;	post-hoc	SHAM	versus	7	days:	p=0.135).	Standardized	width,	length-width	ratio	and	unstandardized	width	did	not	show	any	significant	cardiomyocyte	atrophy	or	trends	(all	p>0.503).	A	cardiomyocyte	with	representative	length	and	width	is	illustrated	in	Figure	4.6.A.	0 2 4 6 8 1 0051 01 52 02 5S ta n d a rd iz e d 	c a rd io m y o c y te	w id th 	(µm / cm )Relative	frequency	(%) S H A M1 2  h o u rs1  d a y3  d a y s5  d a y s7  d a y s0 1 0 2 0 3 0 4 0 5 0051 01 5S ta n d a rd iz e d 	c a rd io m y o c y te	le n g th 	(µm / cm )Relative	frequency	(%) S H A M1 2  h o u rs1  d a y3  d a y s5  d a y s7  d a y sSHAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s	7 	d ay s2 02 22 42 62 83 03 2Standardized	cardiomyocyte	length	(µm/cm)S HAM1 2	ho urs1 	d ay3 	d ay s5 	d ay s7 	d ay s23456Standardized	cardiomyocyte	width	(µm/cm)A B C	Figure	4.8.	LV	myocardial	cardiomyocyte	length	and	width	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points.	Longitudinally	oriented	cardiomyocytes	are	imaged	in	the	LV	myocardial	free-wall	and	all	cardiomyocyte	dimensions	(µm)	are	standardized	to	femur	length	(cm).	In	panel	A,	an	immunofluorescent	image	shows	a	representative	cardiomyocyte	with	a	scale	bar	measuring	20	µm	(x63).	Stained	targets	include:	connexin-43	in	far-red	(pink;	intercalated	discs),	α-actinin	in	red	(Z-lines),	wheat	germ	agglutinin	in	green	(plasma	membrane)	and	DNA	in	blue	(nuclei).	The	dotted	and	solid	lines	indicate	the	length	and	width	of	a	cardiomyocyte,	respectively.	In	panel	B,	frequency	distributions	show	skewness,	kurtosis	and	the	range	of	measurements	obtained	for	standardized	length	and	width	(Gaussian	fit).	In	panel	C,	individual	rat	means	are	plotted	as	symbols,	while	group	means	and	standard	errors	are					74	represented	with	error	bars	(mean	±	SE).	All	sample	sizes,	means,	standard	errors	and	p	values	are	provided	in	Table	4.5.	4.5.2 Cardiomyocyte	cross-sectional	area	and	volume		 Standardized	(to	femur	length)	and	unstandardized	cardiomyocyte	CSA	were	not	significantly	decreased	following	T3-SCI	at	any	time-point	in	the	acute	setting	versus	SHAM	(both	p>0.207)	(Figure	4.7.C.).	Post-hoc	testing	indicated	that	standardized	cardiomyocyte	CSA	at	1	day	post-SCI	tended	to	be	lower	versus	SHAM	(p=0.074).	An	immunofluorescent	image	with	representative	CSA	is	illustrated	in	Figure	4.7.A.	Similarly	to	CSA,	standardized	(to	femur	length)	and	unstandardized	cardiomyocyte	volume	were	not	seen	to	significantly	decrease	following	T3-SCI	at	any	time-point	along	the	acute	spectrum	versus	SHAM	(both	p>0.146)	(Figure	4.7.C.).	Furthermore,	post-hoc	tests	revealed	that	standardized	cardiomyocyte	volume	at	1	day	and	7	days	post-SCI	tended	to	be	lower	versus	SHAM	(p=0.086	and	p=0.075,	respectively).						75	0 2 0 4 0 6 0 8 0051 01 5S ta n d a rd iz e d 	c a rd io m y o c y teC S A 	(µm 2 / c m 2 )Relative	frequency	(%) S H A M1 2 	h o u r s1 	d a y3 	d a y s5 	d a y s7 	d a y sACBSHAM1 2	ho urs	1	da y3 	d ay s5 	d ay s	7 	d ay s02 04 06 0Standardized	cardiomyocyteCSA	(µm2/cm2)S HAM1 2	ho urs	1	da y3 	d ay s5 	d ay s	7 	d ay s05 0 01 0 0 01 5 0 0Standardized	cardiomyocytevolume	(µm3/cm3)	Figure	4.9.	LV	myocardial	cardiomyocyte	cross-sectional	area	and	volume	following	SHAM	surgery	and	T3-SCI	at	different	termination	time	points.	Cross-sectionally	oriented	cardiomyocytes	are	imaged	in	the	LV	epicardial	LV	free-wall.	Cross-sectional	area	(µm2)	and	volume	(µm3)	are	standardized	to	femur	length2	(cm2)	and	femur	length3	(cm3),	respectively.	Panel	A	shows	an	immunofluorescent	image	of	representative	cardiomyocytes	from	all	groups	with	a	scale	bars	measuring	50	µm	(x20).	Stained	targets	include:	α-actinin	in	red	(Z-lines	to	identify	cardiomyocyte	cytoplasm),	wheat	germ	agglutinin	in	green	(plasma	membrane)	and	DNA	in	blue	(nuclei).	In	panel	B,	frequency	distributions	show	skewness,	kurtosis	and	the	range	of	measurements	obtained	for	standardized	cross-sectional	area	(Gaussian	fit).	In	panel	C,	individual	rat	means	are	plotted	as	symbols,	while	group	means	and	standard	errors	are	represented	with	error	bars	(mean	±	SE).	All	sample	sizes,	means,	standard	errors	and	p	values	are	provided	in	Table	4.5.		 					76	 Discussion	In	this	study,	we	aimed	to	investigate	the	temporal	effects	of	acute	high-level	SCI	on	cardiac	function,	proteolysis	and	structure	in	an	experimental	model	with	LV	catheterization,	echocardiography,	histology	and	gene	expression	analysis.	This	is	the	first	study	to	demonstrate	that	a	reduction	in	cardiac	function	precedes	structural	remodelling	following	acute	experimental	high-thoracic	SCI.	We	report	reduced	load-dependent	and	-independent	systolic	function	and	ventricular-arterial	uncoupling	but	no	change	in	diastolic	function	acutely	post-SCI,	which	could	be	explained	respectively	by	the	immediate	loss	of	cardiac	descending	sympathetic	control	along	with	cardiac	unloading,	and	no	sufficient	time	for	structural	remodelling.	Furthermore,	it	appears	that	cardiomyocyte	dimensions	are	yet	to	be	affected	in	the	acute	setting	but	this	effect,	which	we	know	occurs	in	the	chronic	setting,	is	likely	driven	by	the	upregulation	of	UPS	gene	expression	that	we	observed	acutely	post-SCI.			5.1 Hemodynamics,	cardiac	volumes	and	function	5.1.1 Hemodynamics	were	negatively	affected	acutely	following	high-thoracic	SCI		We	found	reduced	SBP	and	MAP	acutely	post-SCI	and	these	findings	mirror	what	has	been	previously	described	in	sub-acute	and	chronic	studies	from	our	laboratory	with	similar	injury	models.72,73,154,157	Furthermore,	our	MAP	results	are	in	agreement	with	a	clinical	meta-analysis,	performed	by	our	research	group.65	The	observed	decreased	in	blood	pressure	in	this	thesis	and	the	literature	is	likely	explained	by	the	lack	of	vasomotor	tone	in	the	vasculature	due	to	the	loss	of	supraspinal	sympathetic	input	to	the	majority	of	the	cardiovascular	system	following	high-level	SCI.48,85,93	We	observed	an	acute	decrease	in	HR	following	high-thoracic	SCI	which	returned	to	basal	levels	at	the	end	of	the	acute	phase,	6-7	days	post-SCI.	The	decrease	in	HR	in	the	early	acute	phase	post-SCI	could	be	explained	by	the	lack	of	sympathetic	input	to	the	heart,	which	incapacitates	its	chronotropic					77	functions.78	An	increase	in	HR	in	the	later	phase	of	acute	SCI	could	be	a	compensatory	mechanism	by	the	PSNS	(decreased	input)	to	attempt	to	counteract	the	reduction	in	SV	observed	at	the	end	of	the	acute	phase.	That	HR	values	were	comparable	to	controls	at	the	end	of	the	acute	phase	is	in	agreement	with	chronic	pre-clinical	data	from	our	laboratory72	and	clinical	data.65		5.1.2 Cardiac	volumes	were	reduced	acutely	following	high-thoracic	SCI	We	report	a	significantly	lower	EDV	and	SV	in	the	T3-SCI	group	compared	to	SHAM	at	6-days	post-surgery.	Our	findings	are	in	agreement	with	both	pre-clinical	and	clinical	literature	following	sub-acute	and	chronic	SCI,	which	also	indicate	lower	EDV	and	SV	following	high-level	SCI.65,72,73,157	The	reduction	in	EDV	post-SCI	is	most	likely	explained	by	the	reduction	in	preload	post-injury	due	to	venous	pooling	in	the	splanchnic	region	and	lower	limbs,	which	is	subsequent	to	the	loss	of	sympathetic	control	to	lower-limb	skeletal	vasculature	and	impairment	of	the	skeletal	muscle	pump	(innervated	by	L2-S1)	as	well	as	a	reduction	in	blood	volume.48,83,93,94,174,85–92	This	influence	of	preload	on	LV	volume	function	was	demonstrated	in	a	previous	study	which	elicited	an	increase	in	preload	via	chronic	passive	hind-limb	exercise	and	showed	improved	EDV	and	SV	in	SCI	rats	following	severe	T2	contusions.154	Interestingly,	we	found	that	EDV	was	actually	significantly	lower	in	the	SHAM	at	group	pre-surgery	compared	to	the	T3-SCI	group	and	then	increased	with	time.	The	former	occurrence	is	perhaps	due	to	lower	but	non-significant	body	mass	in	the	SHAM	group	at	the	pre-surgery	as	these	animals	were	slightly	younger	by	4-5	days.	The	latter	could	be	explained	by	maturation	with	time	as	these	animals	were	adults	but	remained	juvenile.	There	were	no	differences	in	ESV	between	the	T3-SCI	and	SHAM	groups	along	the	acute	timeline,	however,	there	was	an	interaction	effect	for	ESV,	whereby	ESV	was	different	at	pre-surgery	in	the	two	groups.	Although	there	were	no	differences	post-surgery	in	the	SHAM	and	SCI	groups,	the	SCI	rats	had	significantly	reduced	ESV	post-SCI	at	all	time-points	compared	to	their	pre-surgery	values.	The	reduction	in	ESV	post-SCI	observed	in	this	thesis	is	in	agreement	with	pre-clinical	and	clinical	research	performed	by					78	our	group,	which	also	indicate	a	significant	reduction	in	ESV	in	the	chronic	stages	post-SCI.65,72	While	the	chronic	reduction	in	ESV	could	partly	be	explained	by	the	well-known	cardiac	atrophy	post-SCI,	this	hypothesis	is	not	suitable	to	explain	our	acute	findings	as	there	was	no	significant	atrophy	in	this	study.	Additionally,	the	acute	reduction	in	ESV	cannot	be	accounted	for	by	an	increase	in	contractility	as	ESPVR	was	reduced	at	all	time-points	post-SCI.	Instead,	the	reduction	in	ESV	is	most	likely	to	be	explained	by	the	reduction	in	afterload,	observed	as	a	non-significant	reduction	in	Ea,	which	is	subsequent	to	the	loss	of	sympathetic	control	to	the	vasculature	(innervated	by	T1-L2),48	incapacitating	vasoconstriction	and	causing	a	reduction	in	arterial	blood	pressure,	observed	as	reduced	SBP	and	MAP.	Similarly	to	EDV,	ESV	was	increased,	along	with	SBP	and	MAP,	in	SCI	rats	which	underwent	passive	hind-limb	exercise.154	It	is	important	to	note	that	the	reduction	in	cardiac	volumes	in	the	T3-SCI	group	is	most	likely	not	due	to	weight	loss	as	there	was	no	significant	main	effect	of	surgery	on	body	mass.	5.1.3 Systolic	function	and	ventricular-vascular	coupling	were	impaired	at	the	first	acute	time-point	post-SCI	and	persisted	throughout	the	acute	setting		 The	reduced	systolic	LV	function	observed	in	the	T3-SCI	group	compared	to	SHAM,	demonstrated	as	acute	reductions	in	SV,	SW,	CO,	Pmax,	Pdev,	dP/dtmax	and	ESPVR,	is	in	accordance	with	pre-clinical	sub-acute	(5	weeks	following	severe	T2	contusion	in	Sprague-Dawley	rats)73	and	chronic	studies	(12	weeks	following	T3	complete	SCI	in	lean	Zucker	rats),72	performed	by	our	research	team.	Furthermore,	SV	and	CO	are	also	significantly	reduced	in	clinical	studies	following	chronic	high-level	SCI.65	The	acute	reductions	in	SV,	SW	and	CO	could	be	explained	by	the	unloading	of	the	heart	and	subsequent	reduction	in	cardiac	volumes	post-SCI,	as	suggested	by	past	pre-clinical	research	which	reported	improved	CO	and	SV	when	increasing	preload	via	passive	hind-limb	exercise	following	severe	T2	contusions.154,157	Due	to	the	high-level	injury,	there	is	a	loss	of	descending	sympathetic	control	to	the	lower-limb	vasculature	and	impairment	to	the	skeletal	muscle	pump.48,83,85–93,174	The	decreased					79	sympathetic	input	to	the	skeletal	muscles	of	the	lower-limbs	leads	to	venous	pooling	in	the	lower-limbs	and	splanchnic	area	and	therefore	reduced	preload,48,83,85–93,174	which	was	demonstrated	as	reduced	EDV	post-SCI	in	this	thesis.	The	decreased	sympathetic	input	to	the	vasculature	leads	to	the	inability	to	vasoconstrict,48	subsequent	reduction	in	blood	pressure	and	therefore	reduced	afterload,	which	was	implied	by	reduced	MAP	and	the	non-significant	but	notable	reduction	in	Ea	acutely	post-SCI	in	the	present	study.		Pressure	generation	and	contractility	were	reduced	immediately	following	high-level	severe	SCI,	as	suggested	by	reductions	in	pressures	(Pmax	and	Pdev),	dP/dtmax	and	ESPVR	starting	at	the	first	investigated	PV	time-point,	one	day	post-SCI.	Pressure	reductions	are	most	likely	explained	by	the	direct	loss	of	descending	sympathetic	control	to	the	vasculature	and	heart	due	to	the	injury	itself,	incapacitating	its	ionotropic	functions,	and	cardiac	unloading.	This	potential	explanation	is	supported	by	past	studies	from	our	research	group	which	have	shown	that	sympathetic	stimulation	of	the	heart	via	dobutamine	(a	β1	agonist	with	some	β2	and	α1	agonistic	abilities)	improved	LV	pressure	generation	capacities	in	rats	with	severe	high-level	contusions,73	while	increasing	preload	via	passive	hind-limb	exercise	attenuated	reductions	in	the	rate	of	contraction	+dP/dt	and	the	rate	of	relaxation	-dP/dt	post-SCI.154,157	Au	contraire,	no	improvements	in	ESPVR	post-SCI	were	observed	in	rats	which	underwent	passive	hind-limb	exercise,157	indicating	that	decreased	contractility	post-SCI	must	predominantly	be	due	to	the	loss	of	cardiac	sympathetic	input.	Importantly,	reduced	ESPVR	in	T3-SCI	groups	highly	implies	that	the	intrinsic	systolic	function	of	the	heart	is	impaired	acutely	post-SCI.	To	decipher	the	effects	of	descending	sympathetic	control	on	intrinsic	systolic	function,	ESPVR	should	be	assessed	following	different	levels	of	complete	SCI,	above	and	below	the	innervation	of	the	cardiovascular	system.	An	unpublished	study	from	our	research	group	investigated	complete	T3	(reduced	cardiovascular	sympathetic	control)	versus	L2	(intact	cardiovascular	sympathetic	control)	transections	in	Wistar	rats.	The	results	showed	lower	systolic					80	function	(i.e.,	ESPVR,	dP/dtmax	and	pressures)	in	the	T3	rats	versus	L2	rats	8	weeks	post-SCI.	Furthermore,	we	reported	reduced	systolic	function	(i.e.,	ESPVR,	dP/dtmax	and	pressures)	following	complete	T3	transection	(12	weeks	post-SCI)72	and	severe	T2	contusion	(5	weeks	post-SCI)73,	while	Lujan	et	al.	reported	no	reduction	in	dP/dtmax	following	complete	T5	transection	21	days	post-SCI	(intact	cardiac	sympathetic	control).175	In	addition,	Currie	et	al.	reported	lower	LV	function	in	tetraplegic	athletes	with	cervical	injuries	compared	to	their	paraplegic	counterparts	with	mid-thoracic	to	lumbar	injuries.61	Altogether,	our	findings	and	the	literature	suggest	that	descending	sympathetic	control	to	the	cardiovascular	system	is	crucial	for	both	load-dependent	and	-independent	LV	function.		Pre-clinical	research	has	reported	increased	Ea/ESPVR	ratios,	indicating	ventricular-arterial	uncoupling	sub-acutely	and	chronically	following	high-level	SCI.72,73	In	the	present	study,	the	increase	in	Ea/ESPVR,	driven	by	the	significant	decrease	in	contractility,	implies	ventricular-arterial	uncoupling	occurs	acutely	post-SCI	and	is	largely	due	to	cardiac	dysfunction.	Such	ventricular-arterial	uncoupling	has	been	associated	with	reduced	LV	mechanical	efficiency,	reduced	exercise	capacity	and	LV	remodelling,	increasing	the	risk	for	heart	failure.176,177	In	pre-clinical	and	clinical	research,	EF	is	not	altered	after	chronic	SCI	despite	clear	changes	in	both	EDV	and	ESV.65,72	This	is	in	agreement	with	our	acute	findings:	no	changes	in	EF	and	FS	at	the	end	of	the	acute	phase	post-SCI	compared	to	SHAM.	EF	and	FS,	more	specifically	EF,	are	often	used	in	research	for	inferring	systolic	function	as	they	can	be	non-invasively	measured.	However,	these	are	not	adequate	measures	when	volumes	are	highly	reduced	(i.e.	in	the	SCI	field)	as	these	changes	will	be	concealed	by	the	ratios.	The	latter	might	explain	why	we	report	altered	volumes	but	no	changes	in	EF	and	FS	six	days	and	chronically	post-SCI.		Altogether,	we	report	for	the	first	time	a	reduction	in	systolic	function	and	the	occurrence	of	cardiovascular	uncoupling	acutely	post-SCI,	and	further	demonstrate	these	occurrences	following	high-				81	thoracic	SCI	in	an	additional	strain	of	rats	(Wistar),	which	are	most	likely	due	to	the	combined	action	of	decreased	cardiac	sympathetic	control	and	cardiac	unloading.	How	such	events	post-SCI	could	be	disadvantageous	are	discussed	in	Section	6.2.2.	5.1.4 There	was	no	strong	evidence	of	diastolic	dysfunction	acutely	post-SCI	-dP/dtmin	was	the	only	diastolic	functional	index	which	significantly	changed	acutely	post-SCI	versus	SHAM.	An	increased	-dP/dtmin	post-SCI	indicated	an	impaired	rate	of	pressure	decrement.	This	impairment	is	likely	to	be	explained	by	the	decrease	in	pressure	gradient	between	the	LV	and	the	arterial	system,	and	the	decrease	in	preload	post-SCI.	-dP/dtmin	is	known	to	be	affected	by	MAP	and	EDV,178	which	were	both	decreased	in	this	study	post-SCI	and	follow	the	latter	hypothesis.	The	increase	in	-dP/dtmin	is	in	accordance	with	chronic	SCI	pre-clinical	studies	from	our	laboratory,	which	have	showed	significantly	72	and	non-significantly	73,154	impaired	-dP/dtmin	post-SCI.		As	all	other	diastolic	indices	(Ped	and	tau)	showed	no	changes	following	T3-SCI	compared	to	SHAM,	we	cannot	confidently	assert	an	occurrence	of	diastolic	dysfunction	acutely	following	high-thoracic	SCI	in	our	experimental	model.	The	lack	of	distinct	changes	in	diastolic	function	acutely	post-SCI	could	be	explained	by	the	insufficient	time	for	cardiac	unloading	to	cause	structural	remodelling,	which	has	been	detected	chronically	post-SCI	(i.e.	concentric	remodelling	and	suggested	collagen	deposition).63,154,179	A	reduction	in	diastolic	function	at	a	more	chronic	time	is	expected	as	such	is	observed	in	pathologies	associated	with	development	of	cardiac	fibrosis	and	LV	remodelling	(i.e.,	aortic	stenosis,	myocardial	infarction	and	hypertension).180–182	As	mentioned	in	diastolic	dysfunction	following	SCI	(Section	1.5.1.2.),	pre-clinical	and	clinical	SCI	studies	have	not	reached	a	consensus	on	the	occurrence	of	intrinsic	cardiac	diastolic	function	chronically	post-SCI	as	data	collection	techniques	and	injuries	are	not	consistent.63,67–71	Here,	our	results	further	perpetuate	the	question	of	diastolic	function	post-SCI.					82	5.2 Protein	degradation		5.2.1 UPS	gene	expression	was	up-regulated	in	the	early	stages	of	acute	SCI	The	RNA	fold	change	of	UPS	target,	MAFbx,	was	significantly	increased	in	the	acute	phase	following	high-thoracic	SCI	in	LV	tissue.	MAFbx	peaked	at	the	first	experimental	T3-SCI	time-point	by	3.37-fold	(12	hours	post-SCI)	and	tended	to	gradually	decrease	back	to	SHAM	levels.	These	results	closely	mirror	what	was	found	by	Zaglia	et	al.	when	investigating	cardiac	atrophy	and	protein	regulation	following	denervation	via	chemical	ablation	of	sympathetic	neurons.102	They	have	reported	a	two-fold	increase	in	MAFbx	gene	expression	via	RT-qPCR	at	their	first	time-point,	1	day	post-denervation.102	Although	our	MAFbx	fold	change	was	not	significantly	higher	in	the	1	day	T3-SCI	group	compared	to	SHAM,	the	mean	fold	change	at	1	day	showed	a	two-fold	increase	comparable	to	Zaglia	et	al.	values.102	Contrarily	to	Zaglia	et	al.,	we	did	not	observe	MAFbx	gene	expression	levels	to	return	to	SHAM	levels	at	approximately	a	week	post-intervention.102	At	the	last	time-point	of	this	thesis,	7	days	post-SCI,	MAFbx	tended	to	be	increased	versus	SHAM	at	a	non-significant	value	of	1.73	±0.21.	This	value	is	similar	to	the	value	our	research	team	has	previously	reported	in	the	chronic	phase	post-SCI	using	the	same	injury	model	in	a	different	strain	of	rats	(Zucker	lean),	where	we	reported	a	significant	increase	of	MAFbx	gene	expression	at	a	significant	value	of	1.80	±0.79	compared	to	SHAM	at	12	weeks	post-SCI.72	These	results	suggest	that,	following	high-thoracic	SCI,	MAFbx	gene	expression	is	immediately	increased,	peaks	at	12	hours,	then	gradually	decreases	within	the	acute	phase	but	sustains	an	elevated	expression	into	the	chronic	phase.	Elevated	transcription	of	MAFbx	could	imply	an	increase	in	its	E3	ligase	activity	and	therefore	an	increase	in	degradation	of	its	target	proteins,	such	as	eIF3f,111	which	could	explain	the	atrophy	seen	in	in	the	chronic	setting	post-SCI.	The	RNA	fold	change	of	UPS	target,	MuRF1,	was	not	significantly	affected	by	high-thoracic	SCI	in	the	acute	phase.	However,	MuRF1	tended	to	increase	immediately	post-SCI	with	a	peak	at	5	days	post-SCI					83	and	seemed	to	decrease	back	to	SHAM	levels	at	the	end	of	the	acute	phase.	Li	et	al.	and	Zaglia	et	al.	reported	a	significant	increase	in	the	regulation	of	MuRF1	at	one	day	following	denervation	in	skeletal	and	cardiac	muscle,	respectively,102,147	with	a	peak	at	day	3	in	skeletal	muscle.147	MuRF1	remained	significantly	up-regulated	compared	to	the	control	at	14	days	post-denervation	in	the	study	of	Li	et	al.,	while	in	the	study	performed	by	Zaglia	et	al.	levels	had	returned	to	control	levels	by	day	8.102,147	Although,	the	reason	why	our	results	for	this	target	have	not	reached	significance	is	likely	due	to	low	expression	in	one	animal	in	the	5	days	T3-SCI	group,	in	general,	our	MuRF1	results	are	in	accordance	with	the	literature.	The	slight	discrepancy	in	MuRF1	timing	between	studies	could	be	explained	by	different	injury	type	(physical	and	chemical	denervation	versus	SCI),	different	type	of	muscle	(skeletal	versus	cardiac	muscle),	and	different	rodent	species	and	strains	(Sprague-Dawley	rats	and	mice	versus	Wistar	rats).	An	increase	in	MuRF1	gene	expression	could	indicate	an	increase	in	its	E3	ligase	activity	and	therefore	an	increase	in	the	degradation	of	sarcomeric	proteins,104,113–116	which	could	explain,	along	with	the	MAFbx	results,	the	trending	decrease	in	cardiomyocyte	length	seen	in	this	thesis	(Section	5.3.1).	In	our	laboratory,	we	previously	reported	an	increase	in	MuRF1	RNA	and	protein	levels	12-weeks	following	a	complete	T3	transection.72	More	investigation	is	required	in	the	sub-acute	and	chronic	phases	post-SCI	between	the	first	and	12th	week	to	map	the	temporal	changes	of	MuRF1	activity.		Although	there	were	no	significant	changes	in	the	fold	change	of	MuRF1,	our	results	indicate	that	UPS,	one	of	the	main	proteolytic	pathways,	was	up-regulated	immediately	following	high-thoracic	SCI	and	may	be	sustained	at	higher	levels	into	the	chronic	phase.	Our	results	suggest	an	increase	in	protein	degradation	activity	in	LV	tissue	acutely	following	high-thoracic	SCI	in	a	complete	T3	transection	rodent	model.	However,	to	confidently	conclude	the	latter,	protein	levels	of	MAFbx	and	MuRF1	and	their	own	targets	would	require	quantification	as	RNA	fold	changes	do	not	specifically	correlate	with	changes	in	protein	levels.					84	In	skeletal	muscle	wasting	research,	decreased	mechanical	loading	105,122	and	elevated	ANGII	levels	95,128–130	are	associated	with	increased	UPS	activity	in	the	myocytes.	As	acute	SCI	leads	to	cardiac	unloading	and	reduced	MAP,	followed	by	increased	circulating	ANGII	levels,	neuromechanical	and	neurohumoral	changes	could	potentially	explain	the	observed	UPS	up-regulation	in	cardiac	tissue	post-SCI.	Nonetheless,	prior	studies	investigating	cardiac	muscle	atrophy	strongly	suggest	that	sympathetic	input	is	the	main	factor	regulating	cardiomyocyte	morphology.183	Firstly,	it	has	been	shown	that	following	chemical	ablation	of	sympathetic	neurons,	up-regulation	of	UPS	activity	and	cardiac	atrophy	occurred	quickly	post-denervation	via	decreased	b2	adrenergic	receptors	activation	and	subsequent	FoxO	activity.102	Secondly,	more	recent	studies	from	the	same	research	team	reported	major	changes	in	cardiomyocyte	dimensions	when	the	heart	was	denervated	physically	and	chemically,	and	reported	a	correlation	between	cardiomyocyte	cross-sectional	area,	protein	degradation	and	the	density	of	cardiac	innervation.183	The	literature	and	our	data	further	strengthen	the	hypothesis	that	the	acute	up-regulation	of	UPS	observed	post-SCI	is	due	to	the	loss	of	sympathetic	control	to	the	heart.	This	decrease	in	trophic	input	would	activate	a	signalling	cascade	to	dephosphorylate	FoxO3,	which	would	allow	it	to	translocate	to	the	nucleus	and	start	transcribing	UPS-related	genes.	This	increase	in	UPS	activity	would	likely	cause	increased	protein	degradation,	cardiomyocyte	atrophy,	and	further	decreases	in	volumetric	indices	and	cardiac	contractile	function	with	time.		5.2.2 No	changes	in	the	autophagy	gene	expression	were	detected	acutely	post-SCI	The	RNA	fold	changes	of	all	investigated	autophagy	markers,	ATG7,	ATG12	and	BECN1,	were	not	significantly	affected	by	acute	high-thoracic	SCI.	This	suggests	that	none	of	these	genes	underwent	increased	transcription.	Our	results	are	in	accordance	with	Zaglia	et	al.,	which	reported	no	changes	of	RNA	levels	for	all	of	their	chosen	autophagy	markers	(i.e.,	P62,	Bnip3,	BECN1	and	LC3)	at	8	days	following	denervation	of	mice	cardiac	tissue.102						85	The	autophagy	results	from	this	thesis	refute	my	hypothesis	which	stated	an	expected	increase	in	gene	expression	of	key	targets	of	the	pathway	as	ANGII,	an	up-regulator	of	autophagy,	is	predicted	to	increase	acutely	post-SCI	due	to	reduction	in	blood	pressure	post-SCI.	For	future	directions,	temporal	ANGII	levels	must	be	quantified	to	determine	if	there	is	an	association	with	increased	autophagy	post-SCI.	Furthermore,	protein	levels	of	these	markers	and	their	targets	should	be	quantified	to	observe	if	their	translation	is	increased	and	if	protein	degradation	via	autophagy	is	increased,	respectively.	Our	research	team	has	previously	published	results	involving	the	increase	of	ATG12	and	BECN1	RNA	fold	changes	in	cardiac	tissue	in	the	chronic	stage	following	the	same	injury	model	in	a	different	strain	of	rat	(Zucker	lean	rats	terminated	at	12	weeks	post-SCI).72	With	those	results,	we	knew	that	autophagy	was	affected	in	the	chronic	phase	post-SCI.	From	this	thesis,	our	results	indicate	that	autophagy,	one	of	the	main	proteolytic	pathways,	is	perhaps	not	yet	up-regulated	in	the	acute	phase	following	T3-SCI	in	our	experimental	model.	Interestingly,	Zaglia	et	al.	reported	an	increase	in	autophagy	activity	subsequent	to	UPS	up-regulation	30	days	post-denervation	(LC3II	in	cardiac	muscle).102	Therefore,	it	is	plausible	that	autophagy	is	up-regulated	later	than	UPS	in	cardiac	muscle	atrophy		following	such	neural	interventions.102,121		5.3 Cardiac	structure	5.3.1 Histological	data	suggested	the	commencement	of	cardiomyocyte	atrophy	acutely	post-SCI	None	of	the	cardiomyocyte	dimensions	(standardized	to	femur	length	and	unstandardized),	length,	width,	CSA	or	volume,	were	significantly	different	in	the	T3-SCI	groups	at	any	time-point	versus	the	SHAM	group.	Nonetheless,	standardized	length	tended	to	gradually	decrease	along	the	acute	setting	with	the	lowest	value	in	the	7	days	T3-SCI	group	versus	SHAM.	These	results	suggest	that	cardiomyocyte	atrophy	might	be	initiated	acutely	without	reaching	significance.	We	hypothesize	that	cardiomyocyte	atrophy	will	be	significant	early	in	the	sub-acute	phase	following	high-level	SCI	as	length	and	width	have					86	been	reported	to	be	significantly	decreased	at	5-weeks	post-contusion	and,	in	the	chronic	phase,	12-weeks	post-transection,72,73	and	CSA	has	been	observed	to	decrease	8-weeks	post-transection	in	an	unpublished	study	from	our	laboratory.	The	initiation	of	cardiomyocyte	atrophy	could	be	explained	by	the	increase	in	UPS	explained	in	Section	5.2.1.	A	potential	reason	why	cardiomyocyte	length	would	decrease	prior	to	width	could	be	the	degradation	of	actin	and	myosin	filaments	by	E3	ligase,	MuRF1,	causing	either	a	decrease	in	the	number	of	sarcomeres	or	shortening	of	the	sarcomeres.113,116,117	A	previous	study	from	our	group,	which	also	reported	cardiomyocyte	atrophy,	did	not	report	significant	differences	in	sarcomere	number	or	length	in	SCI	animals	compared	to	controls.73	However,	sarcomeric	number	or	length	were	not	quantified	in	this	thesis	as	IF	was	not	deemed	accurate	for	this	type	of	measurement.	Such	quantification	should	be	investigated	next	with	higher-resolution	techniques,	such	as	electron	microscopy	to	measure	length	of	thick	filaments	(A-bands)	and	estimate	the	number	of	sarcomeres	in	parallel	per	cell.		Cardiac	atrophy	can	be	caused	by	a	reduction	in	body	mass.	However,	in	this	thesis,	the	decreasing	trend	in	cardiomyocyte	length	post-SCI	was	most	likely	not	due	to	weight	loss	in	our	paralyzed	animals,	as	there	were	no	differences	in	body	mass	between	the	SHAM	and	T3-SCI	groups	when	we	collected	the	samples	at	termination.	Furthermore,	our	dimensions	were	standardized	to	femur	length,	which	also	did	not	differ	between	groups.						87		Figure	5.1.	Overview	of	findings.	Underlined	events	are	new	findings	acutely	post-SCI.	In	blue	are	variables	measured	in	this	thesis.	In	grey	are	hypothesized	events	which	require	more	investigation.	ANGII,	angiotensin	II;	CO,	cardiac	output;	dP/dtmax,	maximal	rate	of	systolic	pressure	increment	;	EDV,	end-diastolic	volume;	EF,	ejection	fraction;	ESPVR,	end-systolic	pressure-volume	relationship	(index	of	contractility);	ESV,	end-systolic	volume;	FS,	fractional	shortening;	LV,	left-ventricle;	MAFbx,	muscle	atrophy	F-box;	MAP,	mean	arterial	blood	pressure;	MuRF1,	muscle	RING-finger	protein	1;	Pmax,	maximum	pressure;	SCI,	spinal	cord	injury;	SBP,	systolic	blood	pressure;	SV,	stroke	volume;	SW,	stroke	work;	UPS,	ubiquitin	proteasome	system.		 					88	 Conclusion		 The	literature	and	previous	findings	from	our	laboratory	agreed	on	the	occurrence	of	systolic	dysfunction	and	cardiomyocyte	atrophy	with	an	associated	increase	in	proteolytic	activity	in	the	chronic	phase	following	severe	high-level	SCI.	However,	previous	to	this	study,	there	was	no	research	investigating	the	temporal	progression	of	cardiac	dysfunction,	cardiac	remodelling	and	the	underlying	molecular	events	acutely	following	high-thoracic	SCI.	Therefore,	there	existed	a	knowledge	gap	regarding	when	these	events	began	to	occur	and	in	which	order	they	took	place.	To	investigate	the	temporal	effects	of	acute	high-thoracic	SCI	on	LV	cardiac	function,	proteolysis	and	cardiomyocyte	morphology,	I	conducted	a	pre-clinical	experiment	using	a	T3	complete	transection	rodent	model	along	with	state	of	the	art	in	vivo,	histological,	and	molecular	techniques	to	demonstrate	a	reduction	in	cardiac	function	precedes	structural	remodelling	post-SCI.		6.1 Major	findings	Our	main	functional	findings	were:	1)	systolic	function	was	reduced	and	ventricular-arterial	uncoupling	occurred	acutely	post-SCI;	2)	there	was	no	clear	evidence	of	changes	in	diastolic	function	acutely	post-SCI.	We,	therefore,	report	that	cardiac	dysfunction	and	vascular-arterial	uncoupling	took	place	early	in	the	acute	phase,	persisted	throughout	the	acute	phase	and	was,	therefore,	likely	due	to	the	immediate	loss	of	descending	sympathetic	control	to	the	heart.	Our	molecular	findings	are:	1)	UPS	gene	expression	was	increased	in	LV	tissue	at	the	earliest	investigated	time-point	post-SCI;	2)	there	was	no	evidence	of	changes	in	autophagy	regulation	in	LV	tissue	acutely	post-SCI.	We,	therefore,	report	the	upregulation	of	proteolytic	pathway	activity	in	cardiac	tissue	occurred	early	in	the	acute	setting.	Our	structural	findings	were:	1)	there	was	no	significant	atrophy	in	any	cardiomyocyte	dimensions	acutely	post-SCI;	2)	cardiomyocyte	length	tended	to	be	lower	at	the	end	of	the	acute	setting	following	SCI.	We,					89	therefore,	report	that	cardiomyocyte	atrophy	was	initiated	but	did	not	reach	significance	acutely	following	high-thoracic	SCI.	Thus,	together	the	major	finding	of	this	thesis	was	that	the	reduction	in	cardiac	function	preceded	cardiac	structural	remodelling	following	experimental	high-thoracic	SCI.	6.2 Relevance	6.2.1 Implications	All	the	information	collected	throughout	this	thesis	contributes	to	the	fields	of	SCI	and	cardiovascular	health,	by	demonstrating	that	the	acute	phase	is	a	crucial	and	decisive	time	for	the	adaptive	changes	in	the	heart’s	function	and	structure	following	high-level	SCI	and	should	be	more	thoroughly	investigated	in	future	research	for	potential	time-sensitive	treatments.		6.2.2 Why	should	we	care	about	cardiac	dysfunction	and	cardiac	atrophy	post-SCI?	Firstly,	the	general	population	tends	to	neglect	some	of	the	enumerable	secondary	consequences	following	SCI	and	most	often	do	not	consider	the	effects	of	SCI	on	the	heart.	The	level	and	severity	of	the	injury	itself	and	the	quantity	of	secondary	complications	all	have	a	role	in	recovery	speed,	individual	well-being	and	life	expectancy.	Therefore,	all	side	effects	of	SCI,	direct	or	indirect,	acute	or	chronic,	should	be	studied	as	all	bodily	systems	interplay	together.	Secondly,	the	acute	and	chronic	changes	in	heart	function	and	structure	observed	in	this	thesis	and	in	the	literature	could	be	of	adaptive	nature	instead	of	maladaptive,	contrary	to	what	some	might	think.	As	SCI	leads	to	many	physiological	and	postural	changes,	the	heart	might	simply	be	adjusting	to	its	new	conditions.	However,	this	does	not	negate	the	fact	that	these	changes	could	be	deleterious	to	the	individual	in	certain	conditions.	For	example,	a	smaller	heart,	at	first,	does	not	seem	inconvenient	when	considering	blood	volume	or	lower-limb	muscle	perfusion	as	both	are	decreased	following	SCI	and	lead	to	lesser	cardiac	demands	at	resting	conditions.	However,	a	smaller	heart,	vascular-arterial	uncoupling	and	decreased	cardiac	function	can	lead	to	decreased	performance					90	capacity	and	can	be	disadvantageous	under	strenuous	conditions	such	as	exercise	and	other	stresses	(i.e.,	autonomic	dysreflexia,	orthostatic	stress,	stress,	etc.).	Finally,	in	addition	to	decreased	physical	activity,	dyslipidemia,50–53	blood	pressure	instability,57,58	arterial	stiffness	59	and	hormonal	changes,55	cardiac	dysfunction	and	cardiac	atrophy	lead	to	premature	onset	and	increased	risk	for	CVD	observed	in	individuals	with	SCI,	which	is	the	leading	cause	of	death	in	this	population.60			6.3 Strengths,	limitations	and	considerations	6.3.1 Strengths	In	this	thesis,	working	with	rodents	allowed	us	to	perform	invasive	techniques	which	would	have	been	impossible	to	perform	on	human	subjects	for	ethical	reasons	(i.e.,	catheterizations,	tissue	collection,	etc.).	In	addition,	we	were	capable	of	collecting	robust	and	accurate	cardiac	functional	and	volumetric	data	thanks	to	contemporary	techniques	and	equipment	such	as	the	use	of	an	admittance	catheter,	which	measures	both	resistive	and	conductive	capacity,	and	echocardiography.		6.3.2 Limitations	 	Due	to	logistics,	we	unfortunately	were	not	able	to	perform	an	excision	of	the	entire	organ	in	a	timely	manner	without	affecting	tissue	quality	for	ex-vivo	experiments.	To	determine	net	gross	cardiac	atrophy,	total	cardiac	mass	should	be	obtained	via	a	clean	excision	of	the	heart	and	subsequent	weighing	of	the	tissue	after	careful	removal	of	all	of	the	associated	major	vessels.		6.3.3 Considerations	Pre-surgery	EDV	measures	were	found	to	significantly	different	in	our	SHAM	and	T3-SCI	groups.	This	unexpected	finding	could	be	explained	by	a	non-significant	but	slight	weight	difference	between	the	two	groups	due	to	age.	Although	the	rats	were	ordered	to	be	adults	at	the	time	of	the	study,	the	rats					91	remained	juvenile	enough	to	undergo	growth.	The	T3-SCI	rats	were	4-5	days	older	at	the	time	of	surgery	compared	to	the	SHAM	rats	and	this	was	solely	due	to	logistics.	Standardization	of	the	measurements	with	femur	length	or	body	mass	did	not	alter	the	significance	of	the	results.	If	this	study	were	to	be	repeated,	we	would	ensure	day-to-day	age	matching	between	groups	and	order	more	mature	rats	with	no	potential	for	further	growth.	Although	this	discrepancy	between	pre-surgery	EDV	is	an	important	consideration,	the	deleterious	effects	of	severe	and	high-level	SCI	on	EDV	were	clear	and	indisputable.	6.4 Future	directions	6.4.1 Further	molecular	analyses	following	acute	SCI	To	confirm	the	increase	in	protein	degradation	in	the	acute	phase	post-SCI,	the	quantification	of	protein	levels	of	our	UPS	targets	and	their	own	targets	is	required	via	western	blots.	Furthermore,	enzyme-linked	immunosorbent	assays	(ELISA)	could	be	performed	to	detect	circulating	NE	and	ANGII	levels	in	the	serum	throughout	the	acute	timeline	post-SCI.	These	experiments,	informed	by	what	we	know	in	the	chronic	setting	already,	would	aid	to	understand	the	connection	between	the	occurrence	of	SCI	and	the	regulation	of	proteolysis	in	the	cardiac	tissue	post-SCI.	Other	interesting	molecular	analyses,	which	have	not	yet	been	investigated	following	SCI	in	cardiac	tissue,	include	the	density	of	sympathetic	neurons	and	pathways	such	as	apoptosis	and	angiogenesis.	6.4.2 Sub-acute	time-points	following	SCI	Additional	sub-acute	time-points	should	be	investigated	to	pinpoint	the	time	for	cardiomyocyte	atrophy	and	detect	the	changes	in	the	regulation	of	autophagy.	Next,	cardiac	atrophy	could	be	measured	via	measurement	of	total	cardiac	mass	along	the	acute	and	sub-acute	timelines.			 					92	REFERENCES	1.		 Ovalle	WK,	Nahirney	PC.	Netter’s	Essential	Histology.	2nd	ed.	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