Ma_Thesis

Cardiac Troponin-C is susceptible to glycation:
Implications for diastolic dysfunction
in diabetic cardiomyopathy
BRENDAN MA
Cardiac Phenomics Laboratory
Department of Physiology
University of Melbourne
SUPERVISORS
Dr Kimberley Mellor
University of Auckland
Professor Lea M.D Delbridge
Dr David Stapleton
The Florey Institute
Submitted in partial fulfilment of the
Master of Biomedical Science (MC-BMEDSC)
November 2015

2
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3
Abstract
Context: Cardiovascular disease is the leading cause of mortality in diabetic patients. Up to 80% of diabetic
mortality is directly attributable to cardiovascular disease. Considering 1.1 million Australians are currently
suffering from diabetes, cardiovascular disease presents as a major social burden. In spite of the growing
social burden, characterisation of the diabetic heart and the molecular mechanisms linking diabetes and
heart failure are poorly understood.
Diabetic cardiomyopathy is a distinct heart disease in diabetic patients, characterised by impaired cardiac
relaxation and altered Ca2+
response. Those suffering from diabetic cardiomyopathy present with heart
failure independent of hypertension and obesity, unlike non-diabetic heart failure. This suggests that the
diabetic heart is failing under different influences to the non-diabetic heart, and a thorough molecular
understanding of the mechanisms leading to diabetic heart failure is required.
Advanced glycation end products (AGEs) have been shown to be elevated in diabetes. Studies have
demonstrated that AGES form irreversible attachments to a number of proteins, and impair their structure
and function. Experimentally, much of the focus on AGE modification and pathology has focused on
modification of extracellular matrix proteins, with the role of intracellular AGE modification only recently
coming to light. Recent studies have shown that AGE modification of Ca2+
handling proteins alters
cardiomyocyte Ca2+
response, and may be promoting diastolic dysfunction in diabetes. However no work to
date has examined the role of AGE modification of sarcomeric proteins, and linked these modifications to
altered cardiomyocyte Ca2+
response.
It is possible that AGE modification of troponin-c (TnC), the Ca2+
sensing protein of the sarcomere, may play
a key role in the altered Ca2+
response and contractility observed in diabetic cardiomyopathy. Hence AGEs
may play a crucial role in the molecular pathology of diabetic cardiomyopathy, and thus intervention of
their formation presents as a novel therapeutic target in diabetic heart cardiomyopathy.

4
Aims: To determine if cardiac troponin-C (cTnC), a key Ca2+
handling protein in cardiomyocytes is
susceptible to AGE modification, in vitro and in vivo.
Hypothesis: Cardiac troponin-c is susceptible to glycation, via hexose and AGE adduct modification, in vitro
and in vivo.
Methods: Purified human cardiac troponin-c was incubated under conditions of high glucose and high
fructose (both 2M) to identify residues which were susceptible to sugar-induced modification. Mass
spectrometric analysis of whole protein mass was employed to identify changes in protein mass due to
sugar-induced modifications. Further tandem mass spectrometric analysis of trypsin digested peptides was
utilised to identify exact sites of modification. In vivo screening of AGE adducts was then performed on
streptozotocin (STZ) induced diabetic mouse heart homogenates, and determination of AGE attachments at
in vitro identified sites on cTnC performed.
Results: In vitro incubation of purified human cTnC in 2M fructose and glucose resulted in 3.3 ± 0.2 & 9.0 ±
0.3 hexose attachments respectively. Tandem mass spectrometric identification of hexose attachment sites
revealed 10 separate attachment sites: Lys6, 17, 21, 39, 43, 92, 106, 118, 138, 157. Many of which fall within or around
key functional domains of cTnC. Analysis of in vivo STZ diabetic mouse heart homogenate found that AGE
adducts can be found on cTnC in both STZ and vehicle treated mice hearts at Lys6, 17 & 39.
Conclusions: These findings suggest that cTnC is susceptible to AGE modification, both in vitro and in vivo.
These findings complement previous findings of AGE modification of other Ca2+
handling proteins, and that
AGE formation is elevated in diabetes. However this is the first study to utilise mass spectrometric analysis
to determine all possible sites of modification. AGE modification at any/many of these sites may play a key
role in altering cardiomyocyte Ca2+
response and contractility abnormalities. These data suggest that the
diastolic dysfunction observed in diabetic cardiomyopathy may stem, in part, from AGE modification of
cTnC resulting in altered sarcomeric Ca2+
response, and impaired cardiac relaxation.

5
Declaration
This is to certify that:
- the work described in this report was performed by the author, except where otherwise
acknowledged
I declare that this thesis is a record of original work and contains no material which has been accepted for
the award of any other degree of diploma in any University. This thesis contains no material previously
published or written by another person, except where due reference is made. The work described in this
thesis was performed by the author, except where otherwise acknowledged.
This research report is not more than 20,000 words.
Professor Lea M.D. Delbridge, Co-supervisor
6th
November, 2015
Dr Kimberley Mellor, Co-supervisor
6th
November, 2015
Dr David Stapleton, Co-supervisor
6th
November, 2015
Brendan Ma
6th
November, 2015

6
Acknowledgements
I’d firstly like to express my sincere gratitude to Prof Lea Delbridge for giving me the opportunity to undertake
this research project in her laboratory group. As my supervisors, Prof. Lea Delbridge and Dr. Kimberley Mellor
have been instrumental in helping me compile this body of work. Your consistent support, mentorship and
encouragement throughout the duration of my degree has been invaluable. Your mentorship has taught me
valuable life lessons, and instilled a work ethic which I will carry through life. I truly admire your work ethic,
leadership and dedication to ensuring that your students achieve truly remarkable success. The consistently
excellent standard of work produced by this lab group is a reflection of your phenomenal supervision and
mentorship, and I am grateful to have been able to learn from you. Also many thanks to Dr. David Stapleton for
greatly assisting me with the transition to scientific life, and helping me find my feet.
I’d also like to thank the members of the Cardiac Phenomics Lab for their constant support, inspiration and
mentorship through my time here. I feel privileged to have been able to work with such generous and
enthusiastic colleagues. Their constant support and willingness to assist me with any issues during my degree
have been invaluable. Combined with my supervisors, this group of colleagues has helped me develop a set of
skills and work ethic which I can apply to all areas of my life. To Dr. Jim Bell, Dr. Claire Curl, Uspasna, Brian,
Dutchy, Lozza, Chanchal and most recently JJ, thank you.
To the departmental members and students of the student room: thank you for your comic relief and welcome
distractions during the past two years. I have enjoyed the banter and forming friendships with many of you, and
had a great time working with you every day.
To my friends: thank you for your support and encouragement through hard times. The past 2 years have
included some of the most difficult personal times of our lives, and your love and support have always lifted me
back up when I felt down in the dumps. Here’s to many more years of friendship, and here’s to Martin. To my
family, your wisdom and unwavering support throughout my life have enabled me to reach this important
milestone. I am eternally grateful for your love, and hope to someday repay your dedication and self-sacrifice. I
would not be the person I am today without your love, and am grateful to be able to come home to such a loving
household every night.

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Summary of Table of Contents
Chapter 1- Introduction 13
Chapter 2- Methodology 30
Chapter 3- Results 39
Chapter 4- Discussion 48
Chapter 5- List of references 56
Chapter 6- Appendices 64

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Table of Contents
Abstract 3
Declaration 5
Acknowledgements 6
Summary of Table of Contents 7
Table of contents 8
List of figures 10
List of abbreviations, symbols and units 11
Chapter 1: Introduction 13
1.1 Diabetes is a distinct risk factor for cardiovascular disease 14
1.2 Clinical investigation of diabetic CVD 15
1.3 Characteristics of diabetic cardiomyopathy 16
1.3.1 Cardiac Ca2+ handling is altered in diabetic cardiomyocytes 17
1.4 Troponin-C and cardiomyocyte contraction 21
1.4.1 Structure and function of cTnC in cross bridge cycling 21
1.4.2 Point mutations on cTnC impair cardiomyocyte Ca2+ handling 22
1.5 The role of AGEs in diabetes 24
1.5.1 AGE formation 24
1.5.2 AGEs in the diabetic heart 26
1.5.3 AGE Formation is accelerated with fructose 27
1.6 Summary 28
1.7 Research proposal & hypothesis 29
Chapter 2: Methodology 30
2.1 In vitro glycation of human cardiac troponin-c 31
2.2 In vivo glycation of cardiac troponin-c 31
2.2.1 Induction of diabetes, heart excision and tissue homogenisation 31
2.3 Measurement of cTnC mass shifts by Liquid Chromatographic Time-of-Flight Mass
spectrometry (LC-TOF/MS)
32
2.3.1 Preparation of in vitro cTnC samples for LC-TOF/MS 32
2.3.2 LC-TOF/MS determination of cTnC mass shifts 33
2.4 Identification of modification location on cTnC by Liquid Chromatographic
Tandem Mass Spectrometry (LC-MS/MS)
34
2.4.1 Preparation of in vitro cTnC samples for Liquid Chromatographic Tandem
Mass Spectrometry
34
2.4.2. Preparation of in vivo mouse heart samples for LC-MS/MS 35
2.4.3 LC-MS/MS sample analysis 36
2.4.4 LC-MS/MS data analysis: determination of hexose, AGE and oxidation
location on cTnC
36

9
Chapter 3: Results 39
3.1 In vitro hexose modification of purified human cardiac troponin-c 40
3.1.1 Glucose and fructose-induced cTnC hexose mass shifts 40
3.1.2 Higher levels of oxidation observed in fructose incubated cTnC relative to
glucose
40
3.2 MS/MS determination of hexose attachment sites on human cTnC following in
vitro incubation
42
3.2.1 Hexose modification of cTnC is more frequently observed in 2M glucose than
2M fructose
42
3.3 MS/MS determination of oxidative effect of in vitro incubation of cTnC 44
3.3.1 cTnC oxidation of methionine residues is more frequently following 2M
fructose incubation than 2M glucose
44
3.4 AGE adduct formation in the STZ diabetic mouse 46
Chapter 4: Discussion 48
4.1 Overview 49
4.2 Significant hexose modification of cTnC in vitro with time 49
4.3 Fructose incubation unexpectedly yields less hexose modification of cTnC than
glucose
50
4.4 AGE modification of cTnC occurs in vivo 51
4.5 Insights into AGE modification of cTnC and diastolic dysfunction 52
4.6 Summary 53
4.7 Future directions 54
Chapter 5: References 56
Chapter 6: Appendices 64
A1. Induction of diabetes and tissue excision 65
A2. Heart tissue homogenisation 66
A3. Enrichment of cTnC protein from mouse heart homogenate 67
A4. Example screen-capture of MS/MS primary data 68

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List of Figures
Chapter 1:
Figure 1.1 Excitation contraction coupling in the cardiomyocyte 19
Figure 1.2 Diabetic mouse cardiomyocytes have increased Ca2+ response 20
Figure 1.3 Troponin C structure, conformational changes and functional domains 23
Figure 1.4 AGE formation on proteins 25
Chapter 2:
Figure 2.1 Example In vitro peptide MS/MS fragmentation data and spectrum 38
Chapter 3:
Figure 3.1 Deconvoluted LCTOF-MS spectra of time course cTnC incubations in
glucose and fructose
41
Figure 3.2 Sites of hexose modification on cTnC following incubation in glucose and
fructose
43
Figure 3.3 Sites of oxidation modification on cTnC following incubation in glucose
and fructose
45
Figure 3.4 Presence of AGE adduct on STZ mouse not seen on vehicle treated 47

11
List of abbreviations, symbols and units
2-ME 2-mercaptoethanol
µg Microgram
µL Microliter
3-DG 3-Deoxyglucosone
Å Angstrom
ACN Acetonitrile
AGEs Advanced glycation end-products
Amu Atomic mass units
CAD Coronary artery disease
CEL N-epsilon-(carboxyethyl)lysine
CHD Coronary heart disease
CID Collision Induced Dissociation
CML N(6)-Carboxymethyllysine
cTnC Cardiac Troponin-C
CVD Cardiovascular disease
Da Daltons
FFA Free fatty acid
FHC Familial Hypertrophic Cardiomyopathy
GLUT Glucose transporter
HbA1c Glycated haemoglobin
kD Kilodaltons
LC Liquid chromatography
LC-MS/MS Liquid chromatographic tandem mass spectrometry
LCTOF-MS Liquid chromatographic time-of-flight mass spectrometry
LTCC L-type calcium channel
M Molar
m/z Mass to charge ratio
MG Methylglyoxal
mM Millimolar
MS Mass spectrometry

12
NCX Sodium-calcium exchanger
ROS Reactive oxygen species
RyR Ryanodine receptor
SB Sample buffer
SDS Sodium dodecyl sulphate
SDS-PAGE SDS-polyacrylamide gel electrophoresis
SERCA Sarco/endoplasmic reticulum Ca2+ ATP-ase
STZ Streptozotocin
TCEP Tris(2-carboxyethyl)phosphine
TEAB Triethylammonium bicarbonate buffer
TFE Tetrafluoroethylene

14
1. Introduction
1.1 Diabetes is a distinct risk factor for cardiovascular disease
Cardiovascular disease (CVD) is the leading cause of mortality amongst Australians with close to 45,000
people dying in 2012 as a direct result of CVD [1, 2]. The prevalence of CVD is rising in Australia, coinciding
with increased incidence of obesity and diabetes [1, 2]. The Western diet (high fat and high sugar content)
combined with an increasingly sedentary and aging population are thought to be promoting the increased
incidence of diabetes [3]. As a result, diabetes is now Australia’s most rapidly growing chronic disease
affecting 1.1 million people, with conservative estimates predicting this number to double by 2030 [1,2].
Characterised by insulin deficiency (type 1) or insulin resistance (type 2), diabetes has been shown to
increase the risk of CVD three-fold [4]. This increased risk is evident when considering that 80% of diabetic
mortality is directly attributed to CVD [5-7]. Despite the strong epidemiological link between diabetes and
CVD, the precise mechanisms of pathophysiology remain largely unknown. There is increasing
epidemiological and clinical evidence that suggests diabetes is associated with heart failure, independent of
other risk factors such as hypertension and obesity. Although the aetiology and pathology of heart failure
has been well documented, the molecular mechanisms responsible for heart failure in diabetic patients
remain largely unknown. Historically, the Framingham Study identified a strong epidemiological link
between diabetes and heart failure [8]. The study showed that the frequency of heart failure is twice as
high in diabetic men, and five times higher in diabetic women compared with age-matched non-diabetic
patients [8]. Subsequent large scale clinical investigations showed a consistent association between
diabetes and heart failure, independent of other comorbidities [3, 5-7]. Experimentally, it has been shown
that diabetic hearts are more susceptible to failure than non-diabetic hearts [9], and that pathological
structural and molecular alterations occur in response to acute and/or chronic diabetes [9, 10]. Despite this
strong correlation, the current prognosis for diabetic patients diagnosed with early CVD remains poor, and
existing therapies fail to prolong life expectancies [11]. Thorough characterisation of the molecular
mechanisms underlying these pathological changes is vital for development of effective therapeutic targets
in the treatment of diabetic CVD.

15
The increasing prevalence of diabetes across the developed world coincides with increased consumption of
sugars. In particular, the addition of sweeteners containing fructose (in the form of sucrose and/or high
fructose corn syrup) has increased by approximately 25% over the past 3 decades [12]. Studies have shown
that excessive intake of such sweeteners confers a 24% increased risk of cardiometabolic disease [13-16].
This increased risk has been shown to occur independently of increases in blood pressure and vascular
abnormalities, suggesting a distinct fructose-related mechanism of cardiac pathology [17, 18]. Therefore
investigation of the deleterious cardiac effects of excessive sugar and fructose is warranted. Recent studies
have demonstrated contractile abnormalities in cardiomyocytes and cardiac dysfunction in response to
excess sugar [19, 20], however characterisation of precise molecular mechanisms of damage are largely
lacking.
1.2 Clinical investigation of diabetic CVD
One potential molecular mechanism which may be contributing to diabetic CVD is the role of advanced
glycation end products (AGEs, detailed in Section 1.5). AGEs are a family of permanent post-translational
modification which have been shown to be elevated in diabetes [21, 22]. Clinical studies have shown that
AGE modification of vascular epithelial cells and collagen proteins correlate strongly with the development
of hypertension in diabetic patients [23-25]. Further studies have supported this correlation in the
microvasculature, particularly in the eyes [26-28] and kidneys [29, 30]. Taken together, these studies
suggest that AGE modification of the vasculature may contribute, at least in part, to the development of
hypertension in diabetic patients.
However these clinical studies fail to recognise that heart failure in many diabetic patients occurs
independently of hypertension [31, 32]. Indeed, the majority of studies investigating the role of AGEs in
diabetic CVD have focused on AGE modification of the vasculature and extracellular matrix, and have
demonstrated AGE involvement in fibrotic infiltration and collagen crosslinking [33-35]. However despite
thorough investigation of extracellular AGEs, the role of intracellular AGEs in diabetic CVD has only recently

16
come to light in rodent models (detailed in Section 1.5.2). Given their deleterious effects in other tissues, it
is likely that intracellular AGEs, particularly in diabetic conditions of excess sugar, also play a role in
pathophysiology of diabetic CVD. There is a need to further investigate the role of AGE modification,
particularly of intracellular proteins of the myocardium. As diabetes creates an environment of high-sugar
stress, it is likely that sugar-induced AGE modification of intracellular myocardial proteins is occurring in
diabetes. Acceleration and accumulation of such modifications may play a key role in the molecular
mechanisms of diabetic cardiomyopathy.
1.3 Characteristics of diabetic cardiomyopathy
Cardiomyopathy is a disease where the myocardium becomes enlarged, thickened or stiff, leading to
significantly impaired contractility and poor heart function [32]. The prevalence of cardiomyopathy is almost
three-fold higher in diabetics than non-diabetics, and has been identified to occur independently of
hypertension, obesity and vascular abnormalities [36]. As such, diabetic cardiomyopathy is classified as a
distinct pathology, and diabetes is now recognised as an independent risk factor in the progression to heart
failure. As heart failure in diabetic cardiomyopathy occurs independently of hypertension, it is possible that
AGE modification of intracellular myocardial proteins may be contributing to the pathology.
Early stages of diabetic cardiomyopathy are characterised by impaired diastolic function, or the inability for
the heart to efficiently fill during the diastolic (relaxation) phase of the cardiac cycle [11, 37, 38]. Diastolic
dysfunction can be caused by thickness or rigidity of the ventricular walls, reducing the ability for the
ventricles to effectively relax [11, 31, 39]. This diastolic dysfunction often precedes systolic dysfunction, and
can progress to heart failure. Early clinical symptoms of ventricular dysfunction associated with
cardiomyopathy include an elevation in end diastolic blood pressure in the left ventricle, despite a normal
end diastolic volume [39]. This is often due to impaired relaxation and decreased myocardial compliance
[35].

17
Evidence suggests that increased diastolic pressure leads to mechanical wall stress in the ventricles which
results in compensatory cardiomyocyte growth to normalise wall stress [40]. Such remodelling has been
shown to result in cardiac hypertrophy and fibrotic infiltration, further reducing myocardial compliance and
ultimately leading to heart failure [41, 42]. On a molecular level, structural changes observed in the
diabetic heart have been attributed to metabolic disturbances, abnormalities in ion homeostasis and/or
alterations in structural proteins [43, 44], however precise molecular mechanisms remain largely unknown.
There is potential for AGE modification of myocardial proteins to disturb normal cardiomyocyte function,
however to date this has largely been unexplored.
1.3.1 Cardiac Ca2+ handling is altered in diabetic cardiomyocytes
In a normal cardiac cycle, Ca2+
enters the cardiomyocyte via the L-type Ca2+
channels in response to the
propagation of an action potential down the T-tubule. Upon entering the L-type channel, Ca2+
binds to the
ryanodine receptor (RyR) on the membrane of the sarcoplasmic reticulum (SR). RyR activation results in the
release of Ca2+
from the SR into the cytosol of the cell. Cytoplasmic Ca2+
then binds to the myofilaments via
troponin-c (TnC), inducing a number of conformational changes in the trimeric troponin complex (detailed
in Section 1.4). These conformational changes ultimately result in the sliding of thin and thick filaments,
leading to cardiomyocyte force development and contraction. Upon relaxation, Ca2+
is released from the
myofilament back into the cytosol, where it is sequestered by a number of key Ca2+
handling proteins. The
majority of Ca2+
is pumped back into the SR via the sarco/endoplasmic reticulum Ca2+
-ATPase (SERCA2a),
ready to be released in the next contraction cycle, while the rest is pumped out of the cell via Ca2+
ATPase
or sodium-calcium exchanger (NCX) (Figure 1.1) [45].
The role of impaired Ca2+
homeostasis in diabetic cardiomyocytes is subject to some debate. While many
studies have demonstrated disturbed Ca2+
homeostasis in diabetes [46-48], others have shown that it is
unchanged [49]. Indeed this inconsistency can be seen in studies exploring myofilament responses to Ca2+
,
with some groups showing increased cardiomyocyte Ca2+
sensitivity in diabetes [20, 50], while others

18
demonstrate decreases [51-53]. These discrepancies can be attributed to a number of factors, namely
inconsistencies with animal model, induction of diabetes and tissue preparation. Work from our lab has
shown that cardiomyocytes isolated from fructose fed mouse hearts are more sensitive to Ca2+
, requiring
less Ca2+
stimuli to contract to the same extent as non-diabetic myocytes. [20, 54]. These myocytes also
have more intracellular Ca2+
at 50% relaxation, an indicator of impaired relaxation and diastolic dysfunction
(Figure 1.2) [20]. Due to the inconsistencies in the literature, these novel findings warrant further
exploration.
The altered Ca2+
handling and impaired diastolic relaxation in these cardiomyocytes is possibly caused by
perturbations to key Ca2+
handling proteins within the cell. Indeed there has been a substantial body of
work investigating the effects of diabetes on many Ca2+
handling proteins. Some studies have shown that SR
Ca2+
release is increased in systole, shifting cytosolic Ca2+
concentrations higher and resulting in ‘hyper-
contractile’ rat cardiomyocytes [55]. Others have shown decreased activity of Ca2+
sequestering proteins,
leading to an increase in diastolic Ca2+
concentrations [56-58]. Recent studies have shown that AGE
modification of these Ca2+
handling proteins occurs in diabetic rat cardiomyocytes, and alters Ca2+
transients
(detailed in Section 1.5.2) [46-48, 59-63]. However these studies have focused on SR Ca2+
cycling, and there
has been little literature exploring the effects of diabetes on the ability for Ca2+
to directly interact with the
sarcomeres to regulate contraction and relaxation. Although the literature fails to come to an agreement
regarding Ca2+
homeostasis in diabetes, it is possible that AGE modification of sarcomeric proteins is
altering contractile responses to Ca2+
in diabetic cardiomyocytes. Through this mechanism, direct AGE
modification of the sarcomere may be contributing to diastolic dysfunction in diabetic cardiomyopathy, but
to date this has not been investigated.

19
3Na+
Ca2+
Ca2+
Ca2+
RyR2
SR
Ca2+
Ca2+ ATP
SERCA
Ca2+
`
ATP
Ca2+ Ca2+
TnC
Ca2+
TnT
TnI
Myosin
Actin
Sarcomere
Ca2+
Ca2+
Sarcolemma
T-Tubule
L-type Ca2+ channel
Myofilaments
Contraction
Relaxation
NCX
Sarcolemmal Ca2+
channel
Figure 1.1. Excitation contraction-coupling in the cardiomyocyte. Propagation of membrane depolarisation
down the T-tubule opens the L-type Ca2+
channels to allow Ca2+
entry into the cell. This triggers further Ca2+
release into the cytosol via the ryanodine channels (RyR2) on the sarcoplasmic reticulum (SR). Cytosolic Ca2+
binds to cTnC on the myofilaments to initiate contraction. During relaxation, Ca2+
is pumped back into the
SR by the Sarcoplasmic/endoplasmic reticulum Ca2+
-ATPase (SERCA) and extruded from the cell by the Na+
/
Ca2+
exchanger (NCX) and the sarcolemmal Ca2+
channel. Adapted from Bell et al (2011) [54]

20
Figure 1.2. Diabetic mouse cardiomyocytes have increased Ca2+
response. (A) Representative Ca2+
shortening phase loop from control and fructose-fed mouse cardiomyocytes, descending segment (marked
with red box) indicates Ca2+
concentration during relaxation. Leftward shift in descending segment suggests
Ca2+
hypersensitivity. (B) Intracellular Ca2+
levels at 50% relaxation is decreased in diabetic cardiomyocytes
(n=17-24 cells/group). Adapted from Mellor et al (2011) [20]
A. B.
3 4 5 6
0
2
4
6
8
10
Fructose Control
Ca2+
(Fura2 F360:380)
%Shortening
(normalizedtoLo)
Control Fructose
0
1
2
3
4
5
*
Ca2+
(F360:380)
at50%relaxation

21
1.4 Troponin-C and cardiomyocyte contraction
Cardiac troponin-C (cTnC) is the Ca2+
sensing component of the sarcomeric troponin complex. Consisting of
the structural troponin-T (cTnT), inhibitory troponin-I (cTnI) and cTnC, the troponin complex is integral in
mediating contraction. The trimeric complex is anchored to actin, and lies within the grooves between actin
filaments (Figure 1.1). During relaxation, tropomyosin attachment is allosterically inhibited by cTnI,
preventing actin-myosin cross-bridges from forming. As Ca2+
binds to cTnC, cTnC’s conformation changes
which causes cTnI to also change conformation. As cTnI conformation changes, tropomyosin inhibition of
myosin is relieved, and myosin-actin cross-bridge formation can occur, leading to contraction. During
relaxation, Ca2+
is released from cTnC, conformational changes are reversed, and the sarcomere returns to
the relaxed state (detailed in Section 1.4.1).
1.4.1 Structure & function of cTnC in cross bridge cycling
Cardiac TnC is a short 161 amino acid length protein, broadly divided into 2 globular regions- the N and C-
domains. These domains are linked by a short hinge region, and contain three Ca2+
binding pockets [64].
These 3 Ca2+
binding pockets are divided into 1 x low affinity, and 2 x high affinity pockets (Figure 1.3, panel
B). [65]. The two high affinity pockets bind Ca2+
at diastolic Ca2+
concentrations, hence are always filled,
while the low affinity pocket only binds Ca2+
at systolic concentrations [66, 67]. Only when all pockets are
filled do cTnC conformation changes occur [66].
When Ca2+
is released from the SR during systole, cytosolic Ca2+
concentrations rise from 100nM to 1µM
[68]. This dramatic increase in cytosolic concentration is sufficient to cause Ca2+
binding to the low affinity
pocket on cTnC. To enable sarcomeric shortening and contraction, cTnC undergoes a conformational
change from a closed ‘apo’ conformation, to an open conformation, revealing a small hydrophobic patch
(Figure 1.3, panel A). This open conformation is unstable, and without a means to stabilise it, the protein
will inherently return to the closed conformation [44]. However exposure of this hydrophobic patch
attracts the nearby ‘switch peptide’ region of the cTnI protein [69]. cTnI’s conformation changes as the
switch peptide fills the hydrophobic patch of cTnC, stabilising the cTnC open conformation long enough for

22
a contraction to occur [69, 70]. Upon relaxation, Ca2+
in the low affinity pocket is dissociated, cTnC
conformation begins to shift to the apo position and cTnC loses the cTnI stabilisation of the hydrophobic
patch. As the hydrophobic patch closes, cTnC then returns to the closed ‘apo’ position, primed for the next
contraction. This process of contraction and relaxation is dependent on proper conformational shifts in
cTnC, and any modifications to its structure can affect this process. As cTnC is a highly dynamic protein, the
presence of mutations or AGE modifications on cTnC may have a significant impact on sarcomeric
responsiveness to Ca2+
and overall cardiomyocyte contractility.
1.4.2 Point mutations on cTnC impair cardiomyocyte Ca2+ handling
Studies have shown that alterations to cTnC structure significantly impair cardiomyocyte function [70-77].
Clinical studies have identified an array of cTnC point mutations which are present in familial hypertrophic
cardiomyopathy – the most common heritable cardiovascular disease [77]. Familial hypertrophic
cardiomyopathy is characterised by LV hypertrophy that develops into heart failure and arrhythmia.
Widespread analyses found up to 68 distinct point mutations in the cardiac troponin complex which are
linked with the disease, the first and most well characterised being the cTnC L29Q mutation [74, 77]. The
impacts of these point mutations on overall Ca2+
responsiveness in the heart is debated. Studies involving
reconstituted mutant cTnC containing the L29Q mutation have shown myofilament response to be
decreased [71, 78], increased [50, 73] or unchanged [75, 76]. These discrepancies may be due, in part, to
the different models and preparations used (i.e. animal type, myofilament preparation). Other groups have
attempted to determine the causes of the altered Ca2+
response by characterising the crystal structure of
cTnC by crystallography and NMR [65, 67]. However these efforts have been largely unfruitful due to the
highly dynamic nature of the protein, and isolation of cTnC from the troponin complex is difficult. Indeed,
only one group has accurately detailed the cTnC structure in the open, Ca2+
saturated conformation [79].
Nevertheless, when considering the small but critical conformational changes cTnC undergoes during each
contraction cycle, any alterations to its structure may impair function, and cardiomyocyte Ca2+
handling. In
addition to genetic point mutations of cTnC structure, it is also likely that AGE modification may play a role
in modifying cTnC structure and impairing function in diabetic hearts.

23
Ca2+
Ca2+
Ca2+
Ca2+
Ca2+
2x Ca2+ bound
(apo closed conformation)
3x Ca2+ bound
(open conformation)
Low affinity Ca2+ binding domain
M D D I Y K A A V E Q L T E E Q K N E F K A A F D I F
V
LGAEDGCISTKELGKVMRMLGQNPTPE
E
L
Q
E M I D E V D E D G S G T V D F D E F L V M M V R
65 76
C M
K
D
S
K
G
K
N-Domain
(1-87)
SEEELSDLFRMDK
105
NADGYIDLDE
116
L
K
I M L
D
G
DKMLEEIDDETITEDTAQ
C-Domain
(92-161)
High affinity Ca2+ binding domain
D
N-Term -
1
KC-Term -
NE NDGRIDYDEFLEFMKGV
141152161
A.
B.
Figure 1.3. Troponin C structure, conformational changes and functional domains.
(A). cTnC conformation shifts from apo closed state (Ca2+
unbound) to open conformation (Ca2+
bound) following Ca2+
binding to low affinity Ca2+
binding domain. (B). TnC amino acid sequence.
The cTnC protein is comprised of 2 globular domains (N and C), linked by a short hinge region. cTnC
contains 3 Ca2+
binding domains; 1x low affinity domain (yellow box) which is unbound at diastolic
Ca2+
concentrations and 2x high affinity domains (grey boxes) which are filled at all times.

24
1.5 The role of AGEs in diabetes
Advanced glycation end products (AGEs) are a heterogeneous family of post-translational modifications of
proteins, nucleic acids and lipids via non-enzymatic glycation [80]. Protein glycation is the process through
which covalent glycosidic bonds form between reducing sugar molecules (including glucose and fructose)
and basic amino acid residues (lysine and arginine). AGEs have been shown to be rapidly elevated in
diabetes, and glycated hemoglobin (HbA1c) in the blood is used as a key clinical measure of diabetic
progression [30]. AGE accumulation on cardiac proteins has been well documented in rodent models of
diabetes, on both extracellular [24, 35, 81, 82] and intracellular [46, 59, 83] proteins. These modifications
have been shown to impair protein function and contribute to cardiovascular disease. Given that glycation
of both extracellular and intracellular proteins have been shown to be accelerated in diabetic heart, it is
possible that AGE modification of cTnC may also be occurring in diabetes, and may be contributing to the
altered Ca2+
dynamics observed in diabetic cardiomyocytes.
1.5.1 AGE formation
The first step in AGE formation is the glycosidic attachment of a hexose sugar (by its aldehyde group) to the
NH2-terminal of basic amino acid residues (lysine and arginine) (See Figure 1.4). The glycosidic bond formed
between a hexose molecule and a protein is initially weak and can be enzymatically displaced [84]. This
weakly bound hexose-amino acid molecule is known as a Schiff base. Over time, the Schiff bases undergo a
number of oxidation and reduction rearrangements (collectively known as the Amadori rearrangement) to
form an Amadori product (glucose) or Heyns adduct (fructose) [85]. These AGE intermediates are strongly
bound to the protein molecule, and cannot be enzymatically displaced in vivo [86-89]. They undergo a series
of structural alterations, producing reactive dicarbonyl intermediates such as 3-deoxyglucosone (3-DG) and
glyoxal as biproducts. These reactive dicarboyl species then go on to form subsequent AGE adducts on other
proteins (as they also contain reactive aldehyde groups) [30, 90, 91]. Over a period of months, Amadori and
Heyns products eventually form a variety of stable, permanent AGE adducts on proteins [30]. As the process

25
Protein
N
H H
O H
C
C
H2O
HOH
R
Protein
N
C
H
C
HO
R
Protein Protein
NH
C
H H
C
O
R
NH
C
H
C
HO
R
H
OH
Hexose AmadoriSchiff Base AGE
Days/weeks
Reversible Irreversible Irreversible
Weeks Months
of AGE formation is a chemical reaction, it is driven largely by the concentrations of sugars and the
availability of accessible amino groups for Schiff bases to form. Hence conditions of hyperglycemia or excess
sugar are likely to create an environment conducive to increased AGE formation.
Figure 1.4. AGE formation on proteins. The aldehyde group of a reducing sugar (marked dotted red box)
interacts with the amino group of protein residues (marked dotted blue box) to form a glycosidic bond.
This attachment results in a Schiff base which undergoes the Amadori rearrangement to form an
Amadori product (glucose) or analogous Heynes product (fructose). Over the period of months, this
Amadori product stabilises to form an AGE adduct. Adapted from Brownlee (2001) [92].

26
1.5.2 AGEs in the diabetic heart
AGE modification of proteins occurs naturally in healthy individuals, due to the presence of sugars in and
around cells. However in high sugar settings (such as diabetic hyperglycemia or increased intracellular
glycogen breakdown) AGE formation has been shown to be rapidly accelerated [21, 80, 93, 94].
Paradoxically, in spite of reduced glucose transporters and glucose uptake in diabetes [42, 93, 95], recent
studies have shown that glycogen in the diabetic cardiomyocyte is two to threefold higher than non-
diabetics [19, 96]. Although the mechanisms which lead to this storage are unknown, it is possible that such
accumulation poses pathological risks to the heart. Recent studies postulate the existence of glycophagy- a
glycogen specific process that results in the bulk degradation of stored glycogen, releasing large quantities
of free glucose into the cytosol [19]. Such increases in intracellular glucose may contribute to the
accelerated AGE formation in diabetic cardiomyocytes. In particular, AGE modification of Ca2+
handling
proteins may be an underlying molecular mechanism in the pathology of diabetic diastolic dysfunction [92].
Studies have shown that AGEs can be found on short-lived intracellular Ca2+
handling proteins such as RyR
and SERCA2a in diabetic cardiomyocytes, despite short half-lives of 8 days and 2-3 days respectively [46,
59]. As the formation of AGE adducts occurs over months, this suggests that AGE-modified proteins may
become resistant to degradation. Indeed there has been evidence to suggest AGE modification interferes
with lysosomal bulk degradation of proteins in cardiomyocytes [97, 98]. These AGE modified Ca2+
proteins
have been shown to have impaired function, as compared to non-diabetic controls [59, 60, 62, 63, 99].
Despite these findings, investigation of the role of AGE modification of sarcomeric proteins is still largely
lacking. To date, only one study has investigated AGE-mediated cross-linking of myosin heavy chain in
diabetic cardiomyocytes [83], however they neglected to investigate the troponin complex, or link these
findings to changes in Ca2+
responsiveness in these cardiomyocytes. Although these findings suggest that
altered Ca2+
handling may be linked to AGE-modification of SERCA and RyR, impaired Ca2+
utilisation at the
sarcomeric level is unknown. Interestingly, recent work has demonstrated a degree of efficacy of
pharmacological AGE breakers and inhibitors in abrogating AGE-related functional deficits, however these
are restricted to studies of vascular and extracellular matrix compliance [24, 100]. Hence there is scope for

27
more in depth investigation of AGE modification of intracellular myocardial proteins in diabetes, and their
impact on cardiomyocyte function.
1.5.3 AGE formation is accelerated with fructose
In diabetic rat hearts, fructose content has been reported to be up to 60-fold higher than non-diabetic
controls [101]. As fructose has been shown to be a more reactive molecule than glucose, there is a
compelling case for fructose-mediated AGE damage in the cardiomyocyte. Studies have shown that
fructose-derived AGEs (Fru-AGEs) form and accumulate more rapidly than their glucose-derived equivalents
on vascular proteins [88, 102]. Indeed Fru-AGE derived modification of haemoglobin has been shown to be
7.5 fold higher than glucose [102, 103]. In spite of this, quantification of intracellular Fru-AGE modified
proteins is the subject of debate. Studies have found that Fru-AGEs are elevated in a number in vitro models
[80, 88, 102, 104], however existing detection assays have difficulties in differentiating Fru-AGE and Glu-
AGE adducts [105, 106]. As such the role that Fru-AGE modification of proteins in vivo has failed to reconcile
in vitro studies. In spite of this, it has been shown (as outlined in Section 1.5.2) that AGE modification of
intracellular Ca2+
proteins occurs in diabetic cardiomyocytes. It is likely that there is also AGE modification of
cTnC occurring in diabetes, and such modifications may be playing a key role in the development of Ca2+
handling abnormalities which lead to diastolic dysfunction in diabetic cardiomyopathy.

28
1.6 Summary
In overview, to understand the epidemiological link between diabetes and cardiomyopathy, thorough
molecular characterisation of the diabetic heart is critical. Although there are a number of well
characterised structural and functional deficiencies in the diabetic heart, their underlying molecular
mechanisms are largely unknown. The development of diastolic dysfunction is often the earliest symptom in
diabetic cardiomyopathy, hence characterisation and intervention at this time-point is crucial. As diastolic
dysfunction involves alterations in cardiomyocyte Ca2+
handling, perturbations in Ca2+
handling proteins is a
promising candidate for research.
The role of AGEs has been suggested to impair many Ca2+
proteins in the diabetic heart, and adduct
accumulation has been shown to significantly impair the structure and function of many proteins.
The intricate link between cTnC and Ca2+
handling in muscle (both cardiac and skeletal) has been thoroughly
explored. The importance of cTnC in healthy cardiac function has been well described, however attempts to
study the effects of cTnC function via point mutation insertion has resulted in conflicting outcomes. The
literature to date has largely failed to elucidate any differences in cTnC structure and function in the
diabetic heart. As both fructose and glucose is known to be elevated in diabetic cardiomyocytes, it is
possible that AGE modification of cTnC may be occurring in diabetes. Such modification may be an
underlying mechanism of the impaired sarcomeric Ca2+
response in diastolic dysfunction. It is vital to
understand more about cTnC in the diabetic vs non diabetic heart, to gain more insight into potential
underlying mechanisms of diastolic dysfunction and diabetic cardiomyopathy. Describing this mechanism
may prove to provide a crucial therapeutic target in the development of diabetes-specific CVD therapies to
effectively treat diabetic cardiomyopathy in the future.

29
1.7 Research proposal
In order to achieve a greater understanding of the potential mechanisms underlying diabetic
cardiomyopathy, the aim of my project was to determine if cTnC was susceptible to AGE formation in vitro
and in vivo. In vitro experiments were performed to assess if cTnC was vulnerable to hexose modification
(the first stage in AGE formation) in supra-physiological sugar concentrations. To investigate if any
differential effects of fructose existed, in vitro experiments were performed in a high glucose and high
fructose setting. By employing proteomic techniques, these in vitro experiments allow identification of
precise sites of modification on the cTnC molecule. Identification of sites of hexose modification can then be
applied to in vivo extractions of cTnC from streptozotocin (STZ) diabetic and non-diabetic hearts to screen
for AGE adducts. These outcomes aim to understand cTnC structural alterations which may be occurring in
diastolic dysfunction, and diabetic cardiomyopathy.
Hypothesis:
Cardiac TnC is susceptible to structural alterations via hexose adduct and AGE formation in conditions
of high glucose and fructose, in vitro and in vivo.

31
2. Methodology
All mass spectrometric sample preparation and experiments were performed in the Mass Spectrometry and
Proteomics Facility (MSPF) at Bio21. With the exception of induction of type 1 diabetes by streptozotocin
(STZ) injection and MS/MS sample processing, I independently optimised all protocols, performed all
sample preparation and mass spectrometry experiments.
2.1. In vitro glycation of human cardiac troponin-c
Purified human cardiac TnC (cTnC, 0.16µg/µL) (Life Diagnostics, PA, USA) was incubated in phosphate
buffered saline (PBS) containing: 2M glucose (Sigma-Aldrich, MO, USA) or 2M fructose (Sigma-Aldrich, MO,
USA) at 37o
C for 60minutes, 4 hours, 12 hours, 24 hours or 7 days. Incubations lasting longer than 12 hours
were checked every 12 hours for evaporative condensation on tube caps. A brief vortex and trituration was
performed if condensation was observed. At the end of the incubation period, cTnC samples were
immediately placed on ice prior to sample preparation for LC-TOF/MS (section 2.3.1) or LC-MS/MS (section
2.4.1)
2.2. In vivo glycation of cardiac troponin-c
Induction of type 1 diabetes in mice, heart excision and tissue storage was performed by Chanchal
Chandramouli and Melissa Reichelt. I assisted with homogenisation and processing of heart tissue for mass
spectrometry.
2.2.1. Induction of diabetes, heart excision and tissue homogenisation
Male C57Bl/6 mice were obtained from the Animal Resources Centre (WA, Australia). Animals were aged to
12-16 weeks, and housed at the research facility of the University of Melbourne, Australia under standard
conditions. All experimental procedures were performed in accordance with the Australian code of practise
for the care and use of animals for scientific purposes. This project was approved by the University of
Melbourne animal ethics committee (#1011784).

32
Diabetes was induced at 15 weeks of age via 5 consecutive daily intraperitoneal injections of streptozotocin
(STZ, 55mg/kg) to animals which had been fasted for 6 hours. STZ was dissolved in Na2+
citrate buffer (Tri-
sodium citrate dissolved in saline 2.94 mg/mL, pH 4.5) (Appendix A1). Vehicle treated animals were
injected with citrate buffer only. Animals were culled 8 weeks post STZ injections and hearts were excised
and sectioned for molecular analysis. To excise tissues, animals were anesthetised with a single
intraperitoneal injection of sodium pentobarbital (70mg/kg). Once animals were sufficiently anesthetised,
hearts were removed via thoracotomy, ventricles separated from atria and halved into two equally sized
sections and snap frozen in liquid nitrogen. A 5% tissue homogenate was produced by homogenising frozen
heart tissue in homogenisations buffer (NaCl 146.2mM, KCl 4.7mM, NaH2PO4H2O 0.35mM, MgSO47H2O
1.05mM, HEPES 10mM, Glucose 11mM). Hearts were homogenised with 3 x 15 second bursts of
mechanical blending (Polytron PT2500E, Thermo Scientific, MA, USA). Samples were left on ice to settle for
3 minutes before 400µL of homogenate was added to 400µL of 2x sample buffer (SB) and snap frozen in
liquid nitrogen. See Appendix A2 for homogenisation schematic and buffer information.
2.3. Measurement of cTnC mass shifts by Liquid Chromatographic Time-of-Flight Mass
Spectrometry (LC-TOF/MS)
All LC-TOF/MS experiments were performed on a modular Agilent 6220 ESI-TOF Mass Spectrometer at the
Bio21 MSPF.
2.3.1. Preparation of in vitro cTnC samples for LC-TOF/MS
At the completion of the in vitro glucose or fructose incubation period, human cTnC protein (2µg) was
diluted with 50mM triethylammonium bicarbonate buffer (TEAB) to reduce salt concentrations below
100mM. Samples were purified and further desalted using C18 ZipTip Pipette Tips (Merck & Co, New Jersey,
USA), and eluted with 40µL of 100% acetonitrile (ACN). Samples were then placed onto the Agilent 6200
Auto-Sampler injection stage for LC-TOF/MS analysis.

33
2.3.2. LC-TOF/MS determination of cTnC mass shifts
Reverse phase liquid chromatographic (LC) separation was performed via a C18 silica polymer EASY- reverse
phase column (Thermo Scientific, MA, USA). The dimensions of the LC column were 2.1x100mm, with pore
diameter of 200Å. The LC column was secured horizontal at room temperature. A pre-wash of the silica
column with 85% acetonitrile (ACN) for 5 minutes followed by 5 minutes 5% ACN was performed prior to
each sample injection to elute off any residual proteins from previous experiments. To further promote
protein elution off the column, 5µL of tetrafluoroethylene (TFE) was added during the wash period.
Following the wash protocol, 25µL of incubated cTnC protein sample was injected into the system. A 4
minute switch was included at the start of the sample protocol, where 4 minutes of 5% ACN was
maintained to send non-hydrophilic incubation compounds to waste prior to switching to the mass
spectrometer. At the conclusion of the switch protocol, cTnC was eluted off the column using a gradient of
0.1% formic acid (Solvent A) and 100% ACN (Solvent B) with a flow rate of 0.25ml/minute. Solvents were
pumped through the injection platform using the Agilent 1200 Series Binary Pump. A 20 minute gradient
was run with 5% to 55% of ACN over 10 minutes, then 55% to 85% over 3 minutes. 85% ACN was
maintained for 2 minutes, before returning to 5% for 5minutes. Ionisation of cTnC was performed via an
Agilent 6220 ESI-TOF mass spectrometer with soft ESI ioniser set to positron mode. Mass spectra from the
samples were acquired and compiled using the MassHunter Data Acquisition software (version B.06.00),
then analysed using the MassHunter Qualitative Analysis software (version B.08.01). Mass spectra were
extracted from total ion chromatograms, and presence of cTnC base peaks were confirmed via measures of
mass-to-charge (m/z). In silico deconvolution of mass spectra was also performed to calculate total protein
mass. Masses are presented as Atomic Mass Units (amu), equivalent to Daltons (Da). Determination of
hexose adduct attachment was performed via manual calculation of mass shifts on deconvoluted spectra,
with one hexose adduct known to result in +162 amu. Oxidation was also identified by a mass shift of +16
amu.

34
2.4. Identification of modification location on cTnC by Liquid Chromatographic Tandem Mass
Spectrometry (LC-MS/MS)
All LC-MS/MS experiments were performed on an OrbiTRAP Elite ETD MS/MS, Agilent 5600 QTOF, or
QExactive Triple-TOF instrument. Instrument selection was based on workload and instrument status, as
decided by MSPF staff. I performed all sample preparation, data searching and data analysis, while samples
were loaded and run by the MSPF staff.
2.4.1. Preparation of in vitro cTnC samples for Liquid Chromatographic Tandem Mass
Spectrometry (LC-MS/MS)
At the completion of the in vitro glucose or fructose incubation period, cTnC (4µg) was diluted with 50mM
TEAB to reduce salt concentrations below 100mM. The solution was then transferred from the incubation
tube to a 1.5mL Safe-Lock tube (Eppendorf, Hamburg, Germany). CTnC sample was reduced by incubation
with 5mM tris(2-carboxyethyl)phosphine (TCEP) (Life Technologies, Vic, Australia) and heated at 60o
C for 10
minutes. Following the reduction period, samples were centrifuged at 16,000g for 5 seconds and alkylated
with 100µL of 55mM proteomics grade iodoacetamide (Sigma-Aldrich, MO, USA) at room temperature, in
darkness for 45 minutes. CTnC was then digested with sequencing-grade trypsin (Sigma-Aldrich, MO, USA)
at a ratio of 1:50 (protease:sample) at 37o
C overnight to allow cleavage to occur (no more than 18 hours).
The samples were then acidified with formic acid to 1%v/v final concentration and centrifuged at 16,000g
for 10 minutes. Supernatant peptides (20µL) were transferred into an Exigen vial and placed at 4o
C pending
LC-MS/MS analysis.

35
2.4.2. Preparation of in vivo mouse heart samples for LC-MS/MS
Equal amounts diabetic and control mouse heart homogenate were loaded onto polyacrylamide gels and
separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) (4% acrylamide stacking, 15% acrylamide
resolving gel) using the Invitrogen XCell System (Life Technologies, Vic, Australia). Gels were run at 180V for
60 minutes, before being removed from cassettes and stained for 60 minutes with Coomassie Brilliant Blue
(Bio-Rad, NSW, Australia) on a rocking device. Gels were then de-stained (50% methanol, 7% formic acid) to
reduce background and enhance coomassie band visibility. Gel bands were then excised between 15 and
20kDa for each sample and diced into 1mm3
cubes (Appendix A3) and placed in 1.5mL Safe-Lock tube
(Eppendorf, Hamburg, Germany). Gel cubes were further destained overnight with 50% acetonitrile and
50mM TEAB in equal proportions on a rocking device.
Once the gel cubes were completely transparent with no visible coomassie, samples were centrifuged, de-
stain solution supernatant removed and gel cubes dehydrated with 100% ACN for 30 minutes. Dehydration
was deemed complete when gel cubes became ‘chalk-like’ (opaque, white and hard). If dehydration was
incomplete after 30 minutes, ACN was replaced with fresh 100% ACN and left for a further 10 minutes.
Following dehydration, ACN was removed and proteins were reduced using 10mM TCEP (Life Technologies,
VIC, Australia) at 55o
C for 45 minutes. Reduction solution was then removed and proteins alkylated using
50mM proteomics grade iodoacetamide (Life Technologies, Vic, Australia) for 30mins at room temperature,
in the dark. Alkylation solution was rinsed off using 50mM TEAB, with samples left to rinse for 10mins on a
rotation device. Rinsing was repeated 3 times, before gels were again dehydrated using 100% ACN. Once
dehydrated, ACN was removed and digestion solution (5µg/mL trypsin in 25mM TEAB) was added.
Digestion was allowed to occur overnight (no more than 18 hours) at 37O
C with shaking. The next day,
samples were acidified with 1% v/v formic acid, and centrifuged at 16,000g for 10minutes. 20µL of
supernatant peptides were then transferred to Exigen vials and left at 4o
C pending LC-MS/MS analysis

36
2.4.3. LC-MS/MS sample analysis
Liquid chromatographic separation of peptides for MS/MS was performed on a Nano Acquity reverse phase
LC column. Trypsin digested cTnC peptides (obtained from either the in vitro incubated purified human
cTnC, or the in vivo diabetic mouse heart samples following gel separation enrichments) were loaded onto
a 100um x 25mm ‘Magic’ C18 100Å reverse phase column for desalting and chromatographic separation
before entering gas phase. Peptides entered the gas phase through a Proxeon Nanospray source in positron
mode. Peptides were elutebhbd off the column using a gradient of 0.1% formic acid and 100% ACN with a
flow rate of 0.3µL/minute. A 60 minute gradient was run with 5% to 35% ACN for 45 minutes then 35% to
85% over 5 minutes. The 85% ACN was maintained for 5 minutes before returning to 5% over 5 minutes.
Initial peptide detection was performed using one of the three MS/MS instruments listed in Section 2.4. In
all instruments, peptide ionisation was performed via an ion trap in positron mode, before peptides were
fragmented by Collision Induced Dissociation (CID) collision cell to produce β and γ ions (Figure 2.1, Panel
A). Fragmented peptides then passed through a Transfer Quadrupole before reaching the secondary
detector. Mass spectra were collected and compiled using Bio21’s in-house MSILE platform before being
uploaded to the MASCOT Pipeline for bioinformatic trawling and analysis. All MASCOT assigned spectra
showing hexose or oxidation adduct presence were manually validated for accuracy, details outlined in
Section 2.4.4.
2.4.4. LC-MS/MS data analysis: determination of hexose, AGE and oxidation location on cTnC
MASCOT detection and assignment of cTnC is based on a number of criteria. First, tryptic peptides are
fragmented into β and γ ions, and theoretical ion masses are compiled (Figure 2.1, panel B). Then,
observed mass spectra (Figure 2.1, panel C) are compared to theoretical masses, and a confidence ‘score’ is
generated. The MASCOT software assigns peptide scores by considering the number of theoretical ions
which are detected, and checking the experimental spectra for their signals. It also accounts for the
intensity of their signals (height of peaks) relative to background noise. Ions which have many of their
theoretical ions detected, with a high intensity, are assigned a higher score. After all peptides for cTnC have

37
been assigned scores, MASCOT then determines a ‘threshold’ score, above which represents p<0.05 that
the detected peptide (and any PTM’s) are true results (Appendix A4).
MASCOT software compared detected experimental peptide spectra against theoretical in silico peptide
signatures in the UNIPROT peptide database to identify cTnC (Figure 2.1, Panel B). UNIPROT database
trawling was restricted to i) Homo sapien taxonomy for the in vitro purified cTnC samples, or Mus musculus
taxonomy for the in vivo diabetic mouse heart samples, and ii) peptides limited to those produced via
trypsin proteolysis (C-term lysine and arginine cleavage only). As trypsin cleavage is imperfect, 2 missed
cleavages per peptide were permitted by the software. Peptides generated from cTnC had MASCOT
calculated scores compiled, and unique peptides with the highest scores used to confirm the presence of
cTnC in incubated samples. Confirmation criteria were: combined cTnC score >2000, with 10 or more
unique peptides detected. Peptide confirmation was performed as specified by the PARIS guidelines.
Briefly, peptide scores greater than identity score thresholds, where the false discovery rate is <5% and
peptide sequence coverage >75%. Probing for post-translational modification attachment sites was also
performed by the MASCOT bioinformatic searching platform. Variable modifications set to be searched
were: hexose attachments on lysine or arginine and oxidation of methionine or cysteine residues. Presence
of attachments as determined by MASCOT were confirmed by manually assigning theoretical ion masses to
spectra (Figure 2.1, Panel B&C).
Identification of site specific PTM’s is performed by searching for a known mass shift, on a selected amino
acid residue. Hexose attachments only form on lysine or arginine residues, and result in a mass shift of +162
amu, so MASCOT trawling of MS/MS spectra searches for ions containing Lys or Arg residues which present
a mass shift of 162 amu. Identification of AGE adducts was performed using the same process, except
variable modifications probed also included the AGE adducts N(6)-Carboxymethyllysine (CML) and N-
epsilon-(carboxyethyl)lysine (CEL) on lysine or arginine.

38
7A A8 V9 E10 Q11 L12 T13 E14 E15 Q16 K17(+Hex)
γ10
β1 β2
γ9
β3
γ8
β4
γ7
β5
γ6
β6
γ5
β7
γ4
β8
γ3
β9
γ2
β10
γ1
Example TnC MS/MS peptide fragmentation
(Peptide amino acids 7-17)
B.
A.
C.
200 400 600 800 1000 1200
Intensity
Mass (amu)
(γ7)KQEETLQ
(γ8)KQEETLQE
(γ9)KQEETLQEV
(γ2)KQ
(γ6)KQEETL
γ ions
β ions
(β3)AAV
(β4)AAVE
(β5)AAVEQ
(β6)AAVEQL
(β7)AAVEQLT
(β8)AAVEQLTE
(β9)AAVEQLTEE
(β10)AAVEQLTEEQ
(γ1)K(+Hex)
(γ3)KQE
(γ5)KQEET
(γ4)KQEE
β-ions
Peptide
Residue
γ-ions
`Ion sequence # Theoretical
Mass (amu)
Theoretical
Mass (amu)
# Ion sequence
A 1 72.04 A7 11
AA 2 143.082 A8 1336.648 10 KQEETLQEVA
AAV 3 242.150 V9 1265.611 9 KQEETLQEV
AAVE 4 371.193 E10 1166.542 8 KQEETLQE
AAVEQ 5 499.251 Q11 1037.500 7 KQEETLQ
AAVEQL 6 612.335 L12 909.441 6 KQEETL
AAVEQLT 7 713.382 T13 796.357 5 KQEET
AAVEQLTE 8 842.425 E14 695.309 4 KQEE
AAVEQLTEE 9 971.468 E15 566.267 3 KQE
AAVEQLTEEQ 10 1099.527 Q16 437.224 2 KQ
K17 (+Hex) 309.167 1 K(+Hex)
Figure 2.1: Example of in vitro peptide MS/MS fragmentation data and spectrum. (A) Collision
induced disassociation (CID) fragmentation pattern to produce β and γ ions from peptide 7-17. (B)
Theoretical sequences and masses (amu) of fragmented β and γ ions along with residues included in
each ion. (C) Mass spectrum of peptide 7-17 produced by MS/MS instrument. Ions in mass spectra
(Panel C) were automatically assigned by MASCOT software by comparing observed masses with in
silico masses. Manual validation of MASCOT assignment of peaks was performed. MASCOT assigned
peaks were manually compared to theoretical masses (Panel B) to ensure accuracy. Amino acid
residues A7 and A8 not detected in this instance (grey text).

40
3. Results
3.1. In vitro hexose modification of purified human cardiac troponin C
3.1.1. Glucose and fructose-induced cTnC hexose mass shifts
To determine whether cardiac TnC (cTnC) is vulnerable to hexose modification, in vitro incubations of
purified human cTnC with 2M glucose and 2M fructose was performed. Results from LC-TOF/MS analysis of
cTnC mass show that both glucose and fructose incubations resulted in mass shifts corresponding to hexose
attachments (+162 Da) on cTnC. 2M glucose time course incubations resulted in +1 hexose peak appearing
following a 4 hour incubation, increasing to 2 hexose at 24 hours and 8 adducts at 7 days. 2M fructose
incubations also showed +1 hexose at 4 hours and +2 at 24 hours, however by 7 days there were only 3
hexose adducts detected.
Mean data from 63 experimental replicates showed that 2M glucose induced hexose modification is
significantly increased at 24 hours and 7 days vs PBS control (3.3 ± 0.2 & 9.0 ± 0.3 hexose peaks
respectively, p < 0.05). This significant increase in hexose modification was also seen in fructose incubated
cTnC (2.4 ± 0.2 & 2.8 ± 0.2 hexose attachments at 24 hours and 7 days respectively, p < 0.05 vs PBS control),
although less hexose adducts were observed compared to glucose at the 7 day time point.
3.1.2. Higher levels of oxidation observed in fructose incubated cTnC relative to glucose
To determine whether oxidation modification of cTnC was more evident in fructose or glucose incubations,
2M fructose time course cTnC incubations were compared with 2M glucose. Results from LC-TOF/MS
analysis of fructose incubated cTnC showed multiple +16 amu oxidation peaks in the LC-TOF/MS trace
which were absent in glucose incubated cTnC (Figure 3.1 panel A vs C). These findings suggest that in vitro
incubation of cTnC in fructose results in more oxidation modification compared to glucose.

41
PBS
1 hour
4 hours
24 hours
7 days
Deconvoluted mass (amu)
Relativeintensity(Counts)
TnC
+1H
+1H
+2H
+2H
+1H
+3H +4H
+5H
+6H
+7H +8H
A.
Ctrl 1hr 4hrs 24hrs 7d
B. Troponin C: Glucose
PBS High Glucose
Numberhexoseattachments
P B S 1 4 24 168
0
4
8
12
4
8
12
*
*
Troponin C: Glucose
x105
3
1
2
3
1
2
3
1
2
3
1
2
3
1
2
Deconvoluted mass (amu)
Relativeintensity(Counts)
PBS
1 hour
4 hours
24 hours
7 days
TnC
+1H
+1H
+2H
+1H +2H
+3H
C.
Numberhexoseattachments
0
4
8
12
Troponin C: Fructose
D.
Ctrl 1hr 4hrs 24hrs 7d
PBS High Fructose
4
8
12
* *
x105
3
1
2
3
1
2
3
1
2
3
1
2
3
1
2
1860018500 18700 18800 18900 19000 19100 19200 19300 19400 19500 19600 19700 19800
1860018500 18700 18800 18900 19000 19100 19200 19300 19400 19500
Figure 3.1: Deconvoluted LCTOF spectra of time course TnC incubations in glucose and fructose.
(A) Representative deconvoluted mass spectrum of TnC following time course incubation in 2M glucose.
‘TnC’ peak represents unmodified TnC (18,442 amu), +1H represents a mass shift of +162 amu,
corresponding to a hexose adduct to TnC. (B) Mean data showing number of hexose additions at each
time point under PBS control or high glucose conditions. (C) Deconvoluted spectrum of time course
incubated TnC in 2M fructose, annotations are identical to Panel A. (D) Mean data showing number of
hexose additions at each time point. Data presented mean ± SEM; n(PBS) = 28, n(Glucose) = 8-11
samples per group, n(Fructose) = 3-7 samples per group. 1 way ANOVA, Bonferroni post-hoc test.
*p<0.05 vs PBS.

42
3.2. MS/MS determination of hexose attachment sites on cTnC following in vitro incubation
3.2.1. Hexose modification of cTnC is more frequently observed in 2M glucose than 2M fructose
To determine the amino acid location of the hexose modifications on the cTnC molecule, MS/MS analysis of
cTnC incubated in 2M glucose or 2M fructose was performed. MS/MS analysis of glucose incubated cTnC
identified hexose adducts at 9 out of a total of 13 lysine residues in the cTnC sequence, but none of the 4
arginine residues were modified. Data from 49 glucose incubated cTnC experimental replicates (2M
glucose, heat-block incubated, and trypsin proteolysed) revealed that Lys21 was the most frequently
modified residue (observed in 70% of the 49 samples analysed, Figure 3.2, panels A&D). Consistent with
the LC-TOF/MS data, incubation in 2M fructose resulted in a different hexose modification attachment site
frequency. Lys138 was the most frequently hexose modified residue (evident in 58% of 22 fructose
incubated cTnC samples), while Lys21 was the 4th
most frequently modified (27%). All identified hexose
adduct attachment sites lie within or near key functional domains of cTnC (Figure 3.2, panel F).

43
FructoseIncubated TnC
Amino AcidResidue
K17 K21 K39 K43 K92 K106 K118 K138 K158
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Legend
Not modified
Belowthresholdscore
Above thresholdscore
Glucose IncubatedTnC
AminoAcidResidue
K17 K21 K39 K43 K92 K106K118K138K158
ExperimentalReplicates
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
A. B.
P B S in c u b a te d s a m p le s
% H e x o s e m o d ific a tio n o c c u rre n c e
n = 3 4
R es id u e # (L y sin e)
%ofsamplesanalysed
0
20
40
60
80
100
< T hreshold
> T hreshold
6 17 21 39 43 92 106 118 138 158
34 34
26
32
33 34 34 34 34 348
Troponin C: PBS ControlC.
1 .8 1 -2 M G lu c o s e s a m p le s
% H e x o s e m o d ific a tio n o c c u rre n c e
n = 5 6
R es id u e # (L y sin e)
%ofsamplesanalysed
0
20
40
60
80
100
< T h re s h o ld
> T h re s h o ld
6 17 21 39 43 92 106 118 138 158
U n m o d ifie d
47
21
10
21
34
18
27
34
25
38
7
21
13
21
20
8
10
8
11
6
1220141930143314
Troponin C: GlucoseD.
%Hexoseoccurrence%Hexoseoccurrence
Residue# (Lysine)
Residue# (Lysine)
E.
1 .8 1 -2 M F ru c to s e s a m p le s
% H ex o s e m o d ific a tio n o c c u rre n ce
n = 22
R esid u e s # (L ysin e)
%ofsamplesanalysed
0
2 0
4 0
6 0
8 0
1 0 0
< T hreshold
> T hreshold
6 17 21 39 43 92 106 118 138 158
8
9 10
18
13
15
17
9
12
21
8
5
3
4
5
6
91244986
Residue# (Lysine)
E.
V
E
L
Q
65 76
C M
K
D
S
K
G
K
N-Domain
(1-87)
SEEELSDLFRMDK
105
NADGYIDLDE
116
L
K
I M L
D
G D
K
MLEEIDDETITEDTAQ
C-Domain
(92-161)
D
N-Term -
1
KC-Term -
141152161
6 17 21
3943
106
138
158
F.
Figure 3.2: Sites of hexose modification on TnC following incubation in glucose and fructose.
(A) Heatmap showing sites of hexose modification in glucose incubated cTnC experimental replicates (n=49).
Modification of a residue shown as columns. ‘Threshold score’ refers to MASCOT confidence score (p<0.05) that
the presence of hexose modification is true result, and not false positive (see Section 2.3.3 for more information).
(B) Heatmap of fructose incubated cTnC experimental replicates (n=22). (C). Mean data of cTnC hexose
modification occurrence in PBS incubated samples n=34. (D). Mean data of cTnC hexose modification occurrence
in all glucose incubated samples (1.81M and Glu-C proteolysed included; n=56). (E). Mean data of cTnC hexose
modification occurrence in fructose samples (n=22). (F). Visualisation of hexose modification sites in relation to
functional domains of cTnC. Key: Dark red = above MASCOT significance threshold, orange = below threshold, grey
= no modification detected.

44
3.3. MS/MS determination of oxidative effect of in vitro incubation of cTnC
3.3.1. cTnC oxidation of methionine residues is more frequent following fructose incubation than
glucose
To further investigate the differential oxidation effects of 2M glucose and 2M fructose incubation, MS/MS
experiments were performed to identify sites of oxidation modification on cTnC. Analysis of oxidation sites
showed more consistent methionine oxidation in fructose incubated cTnC compared to glucose at all
methionine residues, with the exception of Met47. Met137 and Met157 were oxidised in 100% of fructose
incubated samples compared to ~70% of glucose incubated samples, suggesting a greater oxidative effect
of fructose at these sites in particular. These experiments also revealed oxidation in the absence of sugar,
with oxidative modification of methionine residues observed in PBS incubated samples (Figure 3.3, panel
C).

45
Glucose incubated TnC Fructose incubated TnC
AminoAcidResidue AminoAcidResidue
M80 M81 M103M120M137M157 M80 M81 M103M120M137M157
ExperimentalReplicates
1 1
2 2
3 3
4 4
5 5
6 6
7 7
8 8
9 9
10 10
11 11
12 12
13 13
14 14
15 15
16 16
17 17
18 18
19 19
20 20
21 21
22 22
23
24 Legend
25 No modification
26 Belowthreshold
27 Above Threshold
28
29
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
P B S in c u b a te d s a m p le s
% O x id atio n m o d ifica tio n o cc u rre n ce
n = 34
R e s id u e # (M e th io n in e )
%ofsamplesanalysed
0
20
40
60
80
100
47 80 81 103 120 137 158
> T h re s h o ld
< T h re s h o ld
U n m o d ifie d
34 2 5 17 13 16 16
32
27
17
21
18 18
2
Troponin C: PBS control
1 .8 1 -2 M G lu c o s e s a m p le s
n = 56
R e s id u e # (M e th io n in e )
%ofsamplesanalysed
0
20
40
60
80
100
47 80 81 103 120 137 158
>Threshold
<Threshold
5
45
11
50
42
6 11 33 39 42 46
12
13 13
5
5
4
5
Troponin C: Glucose
1 .8 1 -2 M F ru c to s e s a m p le s
n = 22
R e s id u e s # (M e th io n in e )
%ofsamplesanalysed
0
20
40
60
80
100
47 80 81 103 120 137 158
> T hreshold
< T hreshold
21
8
5
6
6
8 11 20 21 22 22
A. B. C.
D.
E.
%Oxidationoccurrence%Oxidationoccurrence%Oxidationoccurrence
V
E
L
Q
65 76
C M
K
D
S
K
G
K
N-Domain
(1-87)
SEEELSDLFMRDK
105
NADGYIDLDE
116
L
K
I M L
D
G M
K
DLEEIDDETITEDTAQ
C-Domain
(92-161)
D
N-Term -
1
KC-Term -
141152161
6 17 21
3943
106
138
158
47
80 81
103
137120
157
F.
Figure 3.3: Sites of oxidation modification on TnC following incubation in glucose and fructose.
(A) Heatmap showing sites of oxidation modification in 2M glucose (n=49) incubated samples. Frequency of
modification at a particular residue shown as columns. ‘Threshold score’ refers to MASCOT confidence score
(p<0.05) that the presence of oxidation modification is true result, and not false positive (see Section 2.3.3 for
more information). Glucose heatmap excludes varied samples (1.81M instead of 2M, Glu-C protease digestion
instead of trypsin). (B) Heatmap showing sites of oxidation modification in 2M fructose (n=22) incubated samples.
(C). Mean data of oxidation modification occurrence in PBS incubated samples; n=34. (D). Mean data of hexose
modification occurrence in all glucose incubated samples; n=56. (E). Mean data of hexose modification occurrence
in all fructose samples; n=22. (F). Visualisation of hexose modification sites in relation to functional domains of
cTnC. Key: Dark blue = above MASCOT significance threshold, light blue = below threshold, grey = no modification
detected.

46
3.4. AGE adduct formation in the STZ diabetic mouse
To determine if in vitro identified hexose modification sites corresponded to AGE adduct sites in vivo,
MS/MS analysis of cTnC isolated from STZ-induced diabetic and vehicle mouse heart homogenate were
performed. Analysis of SDS-PAGE excised cTnC from STZ and vehicle treated animals showed that AGE-
modified cTnC is present in both STZ diabetic mice and vehicle treated mouse hearts. Although in vitro
incubations demonstrated hexose attachments at 10 lysine residues, only 3 lysine residues were found to
be AGE modified in vivo, Lys6, 17 & 39. MASCOT searching for AGE adducts was restricted to CML and CEL. Of
the three modification sites, two sites presented CML and CEL adducts in both STZ and vehicle treated
animals (Lys6 & 17). However fragmentation of the peptide Ala22-Lys39 revealed that the presence of CEL
adduct formation on Lys39 was specific to STZ, and not seen in vehicle treated mouse hearts (Figure 3.4,
Panel C). The CEL adduct on Lys39 resulted in a mass shift of 44.81 amu of the residue (from 174.32 amu to
219.13 amu), and all subsequent fragmentation γ ions containing the Lys39 + CEL adduct (Figure 3.4, Panel
B).

47
30G A31 E32 D33 G34 C35 I36 S37 T38 K39 (±CEL)
γ9
β1 β2
γ8
β3
γ7
β4
γ6
β5
γ5
β6
γ4
β7
γ3
β8
γ2
β9
γ1
TnC MS/MS peptide fragmentation
Peptide amino acids 22-39 (only 30-39 shown)
β-ions
Peptide
Residue
γ-ions
Ion sequence #
Theoretical mass
(amu)
Theoretical Mass
(amu) # Ion sequence
STZ Vehicle Vehicle STZ
Not shown A22 - L30 Not shown
AAFDIFVLG 1 934.50 934.50 G30 1038.44 1052.46 10 KTSICGDEAG
AAFDIFVLGA 2 1005.54 1005.54 A31 981.42 995.44 9 KTSICGDEA
AAFDIFVLGAE 3 1134.58 1134.58 E32 910.38 924.40 8 KTSICGDE
AAFDIFVLGAED 4 1249.61 1249.61 D33 781.34 795.36 7 KTSICGD
AAFDIFVLGAEDG 5 1306.63 1306.63 G34 666.31 680.33 6 KTSICG
AAFDIFVLGAEDGC 6 1409.64 1409.64 C35 609.29 623.31 5 KTSIC
AAFDIFVLGAEDGCI 7 1522.72 1522.72 I36 506.28 520.98 4 KTSI
AAFDIFVLGAEDGCIS 8 1609.76 1609.76 S37 393.20 407.21 3 KTS
AAFDIFVLGAEDGCIST 9 1710.80 1710.80 T38 306.17 320.18 2 KT
10 K39 174.32 219.13 1 K (±CEL)
Figure 3.4: AGE modification on cTnC isolated from STZ mouse heart not seen on vehicle treated.
(A) Fragmentation pattern of peptide 22-39 and production of ions. Amino acids 22-30 not shown. (B)
Theoretical masses and sequences of ions produced in STZ and Vehicle treated animals. Note the mass
shifts in all γ ions in the STZ animal due to the presence of CEL on K39 (underlined/bold). (C) MS/MS
spectra demonstrating mass shift of unmodified K39 due to CEL adduct- not present in vehicle treated
animal. Remaining spectra (220-1800amu) not shown. Other small peaks correspond to background
noise and non-cTnC ion signatures reaching the detector.
K39 +CEL=219.13 amu
K39 = 174.32 amu
Vehicle mouse
STZ mouse
160 170 180 190 200 210 220
160 170 180 190 200 210 220
Mass (amu)
Relativeintensity
Vehicle treated mouse
STZ treated mouse
A.
B.
C.

49
4. Discussion
4.1. Overview
This study is the first to demonstrate that cTnC is susceptible to glycation, both in vitro and in vivo. These
findings provide the first evidence that human cTnC contains 10 lysine residues which are susceptible to
hexose attachment, all of which are located in or in close proximity to key functional domains. In vitro
experiments revealed that 2M fructose has a greater oxidative effect on cTnC than 2M glucose. In vivo
screening for AGE adducts showed, for the first time, direct AGE modification of cTnC in the diabetic mouse
heart. Both CML and CEL AGE adducts were identified to modify cTnC in vivo.
4.2. Significant hexose modification of cTnC in vitro with time
LC-TOF/MS experiments revealed that following a 7 day 2M glucose incubation, 9.0 ± 0.3 hexose adducts
can accumulate on a single cTnC molecule. These adducts were detected as a 1,458 amu mass shift, or 8%
increase in total TnC mass. Identification of modification sites via LC-MS/MS showed modification of 10 out
of total 13 lysine residues in cTnC. These 10 residues represent 6.2% of cTnC’s total 161 amino acids and
thus are a substantial modification of the protein. Many of the identified hexose modification sites occur
within or near functional domains. Lys6, 17, 21, 39, 43 all lie within the dynamic N-domain of cTnC. This is the
region where cTnC conformation changes occur following Ca2+
binding to allow transition from closed ‘apo’
conformation to open conformation. The most well characterised point mutation in cTnC in familial
hypertrophic cardiomyopathy also lies in this region (Leu29), and has been shown to result in significant
impairment of cardiomyocyte contraction [51] and relaxation [77]. Lys92 lies within the short hinge region
between the N and C-domains. This flexible hinge region allows N-domain conformation changes to occur.
Lys106 lies within a high affinity Ca2+
binding pocket, and Lys118 and Lys138 are also located in close proximity
to high affinity Ca2+
binding domains. Such modifications may play a role in altering Ca2+
response by
impairing Ca2+
binding and/or unbinding, however association/dissociation kinetics were not explored in
this study. Studies have shown that modification of a single cTnC residue can result in significant
impairment myofilament Ca2+
response in familial hypertrophic cardiomyopathy, despite the L29Q mutation

50
not being in a Ca2+
binding domain. Therefore the 9 distinct hexose modifications seen during in vitro sugar
incubation in the present study is likely to represent a severe impairment.
The process of AGE modification is reliant on 2 primary factors: concentration of sugars and time. Diabetic
concentrations of fructose in the rat heart are ~2mM [101], while intracellular glucose measurements have
been difficult to accurately perform due to the number of pathways in which it is utilised. The 2M used in
this study was chosen as a supra-physiological concentration to saturate all potential hexose attachment
sites on cTnC. However our incubations only extended for 7 days maximum, while AGE modification in vivo
take months to form after the initial hexose adduct attachment [92]. Hence, although the concentrations of
sugars used are supra-physiological, the incubation times are much shorter than the time required for
hexose attachment in vivo. Therefore hexose modification to the extent seen in this in vitro study can
possibly be replicated in vivo, given enough time. As such, a long term sugar incubation, with a lower
concentration of sugars is a logical next step.
4.3. Fructose incubation unexpectedly yields less hexose modification of cTnC than glucose
Previous studies directly comparing the glycation effects of fructose and glucose in vitro have reported that
fructose exposure results in more extensive glycation. Suarez et al [103] showed increased AGE
modification of BSA with fructose vs glucose after a 32 day incubation, with AGE-modified BSA proteins
being approximately 2-fold more abundant in fructose incubations. Given that fructose is well established
to be more reactive than glucose [16, 102, 103, 107], we expected to see that hexose modification was
more abundant and occurred earlier in fructose vs glucose incubations, but our data suggest this was not
the case.
Although fructose is more reactive than glucose, the process of hexose formation is dependent upon
factors other than sugar reactivity. The initial step in the formation of hexose attachments requires amino
acid residues to have positively charged side chains (for the aldehyde group of the sugar to attach to the
amino group of the amino acid) [108]. It is possible that oxidation of methionine residues occurs prior to

51
the formation of hexose attachments on lysine, and such oxidation may be impairing hexose formation by
altering the charge status of nearby lysine and arginine residues. In fact studies have shown that pH and
oxidation-reduction potential of environments are intricately linked in vivo, whereby rates of oxidation of
NADP+
reductase are altered when pH is manipulated [109]. Importantly, the reverse has also been shown,
where unregulated oxidation via reactive oxygen species can alter the intracellular pH of cardiomyocytes
[110, 111]. Although this is an in vitro setting, and some factors (such as altered H+
transporter activity in
cardiomyocytes) may not be applicable in these experiments, the chemical oxidation of methionine may
still be playing a role in explaining the reduced hexose formation. Indeed in the experiments performed in
this study, the formation of hexose adducts in fructose incubated cTnC was associated with increased
frequency of methionine oxidation. Extending the fructose incubation periods beyond the 7 days used in
the present study may result in our findings matching those of the Suarez 32 day incubation.
4.4. AGE modification of cTnC occurs in vivo
This study was the first to show the presence of AGE modification of cTnC in mouse hearts. Screening for
AGE adducts across the cTnC amino acid sequence revealed AGE adducts at Lys6, 17 & 39. Importantly, Lys39
was only AGE modified in the STZ mouse, and not the vehicle. However these results are preliminary, and
are n=1 of each treatment and require further repeats to gain further insights. But as a proof of concept,
these data are the first to show that AGE modification of cTnC occurs in vivo. Previous literature has shown
that AGE modification of intracellular Ca2+
handling proteins RyR and SERCA2a to be increased following STZ
diabetes, and we would expect AGE-modified cTnC to be in line with this. However in this study AGE
abundance was not quantified, and this is rational direction for future research.
It is well established that AGE formation is a long-term process (ie. Up to 4 months [92]), and thus most
previous studies have focused on AGE-modification of long-lived proteins such as collagen [10, 86, 112,
113]. The findings from the present study revealed that AGE formation on the short-lived cTnC protein also
exists. The half-life of cTnC in normal conditions is 5.3 days [114], and previous studies have also found AGE
modification of RyR and SERCA2a, two other proteins with relatively short half-lives of 8 and 2-3 days

52
respectively [46, 59]. The presence of AGEs on such short lived proteins has led to the hypothesis that AGE
formation impairs protein degradation and turnover [115, 116], and the AGE-modification of cTnC in this
study is consistent with this notion. Although the relative proportions of AGE-modified vs unmodified
proteins has not been explored, it is likely that the impaired turnover of AGE-modified proteins results in an
accumulation of modified, potentially defunct, proteins in diabetic cardiomyocytes.
Although functional implications of such modifications were not explored in this study, many studies have
shown that even a single modification of a cTnC residue (via point mutations) can have significant
implications on cardiomyocyte Ca2+
response [51, 71-75, 77], even if the modified residue does not
explicitly lie within a Ca2+
binding domain (L29Q mutation in FHC). Given that diabetes creates a high sugar
intracellular environment, it is likely that AGE modification of cTnC is occurring at an accelerated rate in
diabetes. As studies have shown cTnC modification at a single residue affect cardiomyocyte Ca2+
response
[77], it is likely that AGE modification at a single or multiple sites on cTnC will also alter cardiomyocyte Ca2+
response and function. Such impaired cardiomyocyte function may play a crucial role in the development of
diastolic dysfunction in diabetic heart disease. Ultimately, direct measurement of Ca2+
binding properties of
cTnC in the presence of hexose/AGE modification is an important next step.
4.5. Insights into AGE modification of cTnC and diastolic dysfunction
Diastolic dysfunction is often characterised by impaired relaxation in cardiomyocytes. Previous studies by
Mellor et al have established fundamental disturbances in Ca2+
response from mice fed a high fructose diet
[20]. These cardiomyocytes are shown to be hyper-sensitive to Ca2+
, requiring less Ca2+
to relax, an indicator
of impaired relaxation and early diastolic dysfunction.In healthy cardiomyocytes, cTnC exists
predominantly in the closed ‘apo’ conformation (the conformation in which the sarcomere is relaxed), as
Ca2+
dissociation takes longer than association [78, 117]. However, recent work speculates that cTnC may
also exist in an intermediate state, known as the ‘primed’ or ‘partially active’ conformation [118, 119]. This
is a stage where the angle between the N and C-domains is slightly larger than the ‘apo’ conformation. This
conformation ‘primes’ the cTnC molecule for Ca2+
stimulus, so that the transition to the open conformation

53
may occur more rapidly. Evidence for cTnC existing in this stage comes in the form of NMR spectroscopic
data suggesting that some cTnC molecules change from the ‘apo’ to the ‘open’ conformations faster than
expected, hence the authors suggest that there may be some cTnC molecules existing in an intermediate
stage [118, 119]. It may be possible that the presence of hexose and AGE adducts along the cTnC molecule
may be impairing the ability for cTnC to return to the ‘apo’ and ‘open’ conformations appropriately, and
hence more cTnC molecules may be found in an intermediate conformational stage such as the ‘primed’
state. It is also possible that AGE modification of cTnC may be extending the Ca2+
dissociation time, resulting
in a reduction of cTnC molecules in the closed, relaxed conformation. A higher proportion of cTnC
molecules in the ‘primed’ or ‘open’ states may be the underlying molecular mechanism behind the diabetic
cardiomyocyte Ca2+
hypersensitivity seen by Mellor et al.
4.6. Summary
The work presented here is the first to show AGE modification of cTnC, both in vitro and in vivo.
Interestingly there was a differential hexose modification occurrence seen under high glucose and high
fructose conditions. In vitro incubations of human cTnC in glucose resulted in multiple hexose modifications
while fructose incubation promoted oxidation and hexose modification, although the extent of hexose
modification was less than glucose incubation. This reduced potency in promoting hexose modification is
surprising when considering fructose is a more reactive molecule, however more long term in vitro
incubations may increase hexose modification, as has been previously shown [16, 103]. Understanding the
mechanisms of fructose damage is becoming increasingly important as dietary fructose intake increases
across the Western world.
Direct AGE modification of Ca2+
handling proteins has been shown to modulate diabetic cardiomyocyte Ca2+
handling [46, 62]. Hence AGE modification of cTnC as shown in this study is also likely to affect myofilament
responsiveness to Ca2+
. As a key regulator of the contractile machinery of the cardiomyocyte, this
modification of is likely to have a significant impact on cardiomyocyte contraction/relaxation in response to
Ca2+
stimuli.

54
The data presented in this study provides a comprehensive examination of all potential AGE adduct
attachment sites along the cTnC molecule. The identification of 10 hexose attachment sites is vital in
progressing the understanding of functional impacts AGE modification of cTnC may have in diabetes.
As diabetic cardiomyocytes have been shown to have increased intracellular fructose, and speculated to
have increased glucose, there is potential for increased AGE modification of cTnC to play a crucial role in
the development of diastolic dysfunction in diabetic patients. As such, there may be scope for targeting
AGE modification of cTnC as a future therapeutic intervention for the treatment of diabetic
cardiomyopathy.
4.7. Future directions
The experiments detailed here indicate that AGE-modification of cTnC occurs under diabetic conditions,
however the functional deficits resulting from such modifications are yet to be investigated. Although the
experiments in this study have provided qualitative evidence of AGE-modification of cTnC, quantification of
the abundance of AGE-modified cTnC is required to determine the extent of glycation in vivo.
To progress this understanding, quantification of AGE modified cTnC via Western immunoblots should be
undertaken in diabetic hearts and non-diabetic hearts to determine if AGE modified cTnC is accumulating in
the diabetic setting. It will be also interesting to quantify the relative proportions of modified vs unmodified
cTnC molecules in diabetic and non-diabetic animals, to gain an understanding of the extent of cTnC AGE-
modification in diabetic hearts. Correlating the abundance/accumulation of cTnC AGE modification and
cellular Ca2+
handling abnormalities may provide further insight into the mechanisms responsible for the
altered Ca2+
response seen in diabetic cardiomyocytes.
Previous work on AGE-modified Ca2+
handling proteins detailed changes in cardiomyocyte Ca2+
response,
however few studies have correlated these functional deficiencies with AGE modification of sarcomeric
proteins. In fact, studies associating AGE-modified cTnC and cardiomyocyte Ca2+
response are entirely
lacking. Measurements of cardiomyocyte contraction and relaxation in response to Ca2+
stimulation may
provide a valuable insight into the functional effects AGE-modification of cTnC may have on the single cell

55
level. Previous studies have also shown the efficacy in pharmacological AGE-cleaving agents in restoring
vascular compliance [25, 35]. Hence the role of similar agents in abrogating Ca2+
abnormalities in
cardiomyocytes should be explored. Ultimately, an important experimental outcome will be to pursue the
efficacy of cardiac-specific AGE-inhibitors or breakers which may provide a viable therapeutic option for the
treatment of diabetic cardiomyopathy.

56
CHAPTER 5
List of References

57
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