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Determination of the Sub-Cellular Mechanisms Underlying Neurodegeneration in Parkinson’s Disease
by
Christopher J Yong-Kee
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Cell & Systems Biology
1.8 Models of Parkinson’s disease .......................................................................................... 36
1.8.1 In vitro models ....................................................................................................... 36
1.8.2 In vivo models ........................................................................................................ 38
1.9 Hypothesis and aims .......................................................................................................... 41
2 Mitochondrial Dysfunction Precedes other Sub-Cellular Abnormalities in an In Vitro Model Linked with Cell Death in Parkinson’s Disease ............................................. 45
3 Development and Validation of a Screening Assay for the Evaluation of Putative Neuroprotective Agents in the Treatment of Parkinson’s Disease ..................................... 73
5 Neuroprotective Actions of SIRT3 and RGM in Advanced In Vitro Models of Parkinson’s Disease .............................................................................................................. 138
5.3.1 Assessment of neuroprotective effects of RGMa and SIRT3 .............................. 149
5.3.2 Development of an in vitro α-synuclein model utilizing ventral mesencephalic primary cultures ................................................................................................... 159
5.3.3 Development of a nigro-striatal organotypic co-culture model of PD ................ 167
with Drp1 to promote self-association. Drp1 forms a spiral structure that wraps around
mitochondria during fission. Mitochondria are severed by Drp1 through a GTPase mediated
reaction. (b) Mfn1 and Mfn2 tether mitochondria together during outer mitochondrial membrane
fusion. Inner mitochondrial membrane fusion is then mediated by Opa1. GTP hydrolysis
initiates the reactions mediated by Mfn1, Mfn2 and Opa1. Abbreviations: Drp1: dynamin-
related protein 1, Mfn 1: Mitofusin 1, Mfn2: Mitofusin 2.
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Drp1
Fis1
a Fission
b Fusion
Mfn2
Mfn1
Opa1
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1.6.1 Regulation of mitochondrial dynamics
Large GTPase proteins mediate fission and fusion processes. Dynamin-related protein (Drp1)
and fission 1 (Fis1) mediate the mechanisms that orchestrate fission of mitochondria (Knott et
al., 2008). Drp1 is a cytosolic protein that contains an N-terminal GTPase domain. During
fission, Drp1 is believed to translocate from the cytosol to fission sites on mitochondria where it
self-associates to form spiral-like structures (Smirnova et al., 2001). These structures begin to
wrap around and constrict the mitochondria until the mitochondrial membrane is severed by a
GTPase dependent mechanism (Otera and Mihara, 2011). Fis1 is anchored to the outer
mitochondrial membrane with its N-terminus exposed to the cytoplasm. Studies suggest Fis1
plays a role in recruiting Drp1 to sites of fission on the mitochondria (Yoon et al., 2003). Three
large GTPases, mitofusin 1 (Mfn1), mitofusin 2 (Mfn2) and optic atrophy 1 (Opa1) mediate
mitochondrial fusion. Mfn1 and Mfn2 are anchored to the outer mitochondrial membrane with
their N-terminal GTPase and C-terminal coiled coil ends facing the cytoplasm. It is unclear how
Mfn1 and Mfn2 mediate outer membrane fusion, but they are believed to tether mitochondria
together in a GTPase dependent manner. Opa1 is situated in the inner mitochondrial membrane
with its GTPase domain facing the intermembrane space (Otera and Mihara, 2011). The process
of inner membrane fusion is unclear, but the GTPase domain of Opa1 appears to be essential
since mutations in this domain lead to mitochondrial fragmentation (Olichon et al., 2007). While
the proteins that regulate fission and fusion are controlled by GTP hydrolysis, it appears that the
activity of these proteins can also be regulated by phosphorylation by cyclin-dependent kinase 1
and cAMP-dependent protein kinase (Westermann, 2010).
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1.6.2 Function of mitochondrial dynamics
Mitochondria are constantly undergoing fission and fusion to adjust mitochondrial populations to
meet cellular demands. Processes such as axonal outgrowth and synaptic plasticity require
changes in mitochondrial content which is dependent on mitochondrial fission and fusion
(Morris and Hollenbeck, 1993; Li et al., 2004). Fission and fusion are antagonistic processes and
a delicate balance between the two must be maintained to ensure mitochondria are functional.
Indeed, damaged mitochondrial DNA is eliminated and depolarised mitochondrial membrane
potentials are restored upon completion of mitochondrial fusion. Fusion is also important for the
inheritance and maintenance of mtDNA, which in turn is key to the overall health of
mitochondrial populations (Rapaport et al., 1998). Mitochondrial fission is critical for synaptic
transmission between neurons since the proper function of synapses is reliant on functional
mitochondria at synaptic sites (Li et al., 2004; Ishihara et al., 2009). Fission is implicated in the
regulation of apoptotic processes in neurons since Drp1 is required for cytochrome c release and
caspase activation (Estaquier and Arnoult, 2007; Ishihara et al., 2009).
1.6.3 Mitochondrial dynamics and neurodegenerative disease
The functions performed by fission and fusion are critical to neuronal health which implies that
any perturbations to these processes may be linked with neurodegenerative disease. Several
neurodegenerative diseases are linked to aberrant mitochondrial dynamics. Mutations in Mfn2
cause Charcot-Marie-Tooth type 2A disease which affects sensory and motor neurons (Chen and
Chan, 2009). Dominant optic atrophy is caused by mutations in Opa1 and leads to degeneration
of retinal ganglion cells (Detmer and Chan, 2007). Interestingly, patients with Charcot-Marie-
Tooth type 2A disease display various symptoms including parkinsonism and psychiatric
disturbances (Verhoeven et al., 2006). Alterations to mitochondrial dynamics may be equally
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important in disorders such as PD where mitochondrial function is especially critical to the
health of nigral neurons.
1.7 Disease modifying agents
Presently, all treatments for PD are focused on treating symptoms of PD by using dopamine
replacement therapy, either by replacing lost dopamine in the form of the dopamine precursor L-
dopa or dopamine agonists, or by enhancing the half-life of remaining dopamine using mono
amine oxidase-B (MAO-B) inhibitors. Levodopa, rotigotine (dopamine agonist), selegiline and
rasagiline (MAO-B inhibitors) are all successful at reducing motor symptoms, however long-
term use is associated with debilitating side effects such as dyskinesias (Schapira, 2009). While
much research is focused on treating symptoms of PD, it would be much more beneficial to
patients if it were possible to stop or slow the progression of neurodegeneration by using a
neuroprotective agent. This would prevent PD progressing to more advanced stages where
patients become seriously disabled and palliative care is necessary. The Committee to Identify
Neuroprotective Agents in Parkinson’s disease (CINAPS) project was the first of its kind to
select a battery of neuroprotective agents to test in clinical trials (Heemskerk et al., 2002; Ravina
et al., 2003). Given the importance of various mechanisms such as oxidative stress and
mitochondrial dysfunction which ultimately cause apoptosis, it is conceivable that putative
neuroprotective agents should act on these pathways. Indeed, some of the various chemicals
tested by CINAPS are involved in the previously mentioned mechanisms (Table 1.2).
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Table 1.2: Agents tested by CINAPS for neuroprotective potential.Agent Mechanism of ActionCaffeine Adenosine A2A receptor antagonismCoenzyme Q10 Cofactor of electron transport chain; antioxidant propertiesCreatine Indirect antioxidant by enhancing energy transfer during periods of high energy demandEstrogen May act as an antioxidant through upregulation of Bcl-2, BDNF and GDNFGM-1 ganglioside UnknownMinocycline Inhibition of microglial activationNicotine Increases neurotrophic factorsNeuro-immunophilin A Immunophilin ligandSelegeline, Rasagiline MAO-B inhibitorRopinirole, Pramipexole Dopamine agonist with antioxidant properties
Adapted from: (Meissner et al., 2004; Hung and Schwarzschild, 2007)
Many of these agents were found to be neuroprotective in various rodent and primate models
before being considered for clinical trials, while others were tested in clinical trials without prior
testing in animal models (Meissner et al., 2004). Nicotine and caffeine were suggested to be
neuroprotective based on the low incidence rates of PD in humans that frequently consumed
them. Estrogen was selected as a putative neuroprotective agent because of the lower incidence
of PD in women, especially in women receiving estrogen replacement (Saunders-Pullman et al.,
1999). Coenzyme Q10 appears to be the most successful agent, having protective effects in
animal models and placebo-controlled clinical trials. Similarly, selegeline was found to be
beneficial when tested using the Unified Parkinson’s Disease Rating Scale (UPDRS). Indeed,
lower scores were given by patients receiving selegeline treatment, but these results were
questionable due to errors in clinical trial design (Hung and Schwarzschild, 2007). The effect of
creatine was assessed in a two year placebo-controlled clinical trial but protection was not
observed as UPDRS scores were not lowered (Bender et al., 2006). However, further study of
creatine is warranted as it was found that it could not be excluded as a possible neuroprotective
agent in a futility study (2006). It appears that compounds that are neuroprotective in animal
models are not always as successful in clinical trials. However, in cases such as with coenzyme
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Q10 and creatine, there was benefit in both animal models and clinical trials. Thus it seems there
is a disparity between the predictive capabilities of clinical trials, and improvements in clinical
trial designs are necessary to test the clinical relevance of putative neuroprotective agents.
Alternatively, improvements in cellular and animal models may be necessary so these models are
better reflective of the human disease.
1.7.1 RGMa
Repulsive guidance molecule A (RGMa) is a membrane-associated glycoprotein that was
originally isolated from chick optic tectum (Monnier et al., 2002). The C-terminal is anchored to
the plasma membrane by a glycosylphosphotidylinositol (GPI) domain and the N-terminal
contains a signal peptide sequence. A partial von Willebrand factor domain and RGD sequence
also exist to support membrane adhesion (Monnier et al., 2002). RGMa is part of a family of
RGM proteins (RGMa-c) with each having their own specific function in the central nervous
system. RGMa is localized to the soma in mature and immature neurons, while immature
neurons also contain RGMa in their axons (Brinks et al., 2004; Schwab et al., 2005). RGMa acts
as a guidance cue for retinal axons and induces the collapse of temporal growth cones during
neuronal development (Monnier et al., 2002). Other functions of RGMa include the ability to
regulate neuronal differentiation and prevent apoptosis through its interaction with neogenin, a
homologue of netrin-1 and DCC receptors. Studies reveal that neogenin is a dependence
receptor where activation of pro-apoptotic pathways normally occurs in the absence of a ligand
(Matsunaga and Chedotal, 2004; Matsunaga et al., 2004). Caspase-3 cleavage of neogenin
initiates these apoptotic signalling pathways, which are blocked in the presence of its ligand,
RGMa. Moreover, RGMa overexpression prevents pro-apoptotic signalling while RGMa siRNA
expression has no effect on cell survival (Matsunaga et al., 2004). Although RGMa has the
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ability to regulate axonal outgrowth and to modulate apoptotic pathways, surprisingly it has not
been studied in various disease models.
1.7.2 SIRT3
The novel family of protein deacetylases, sirtuins, consist of seven family members (SIRT 1-7),
each of which have shown efficacy in the treatment of a wide spectrum of diseases. Sirtuin1
(SIRT1) was the first member of the sirtuins to be extensively studied in mammals (Gan and
Mucke, 2008). Evidence suggests that SIRT1 elicits neuroprotective properties by its actions on
non-histone substrates in the nucleus (Lavu et al., 2008). Another member of the sirtuin family,
SIRT3, is less extensively studied; however, it is believed to be involved in mitochondrial
function. SIRT3 is encoded within the nucleus and then directed to the mitochondria by its N-
terminal mitochondrial localization sequence (Bao et al., 2010). Once inside the mitochondria,
SIRT3 becomes enzymatically active upon cleavage of its localization sequence by a protein
peptidase (Giralt and Villarroya, 2012). SIRT3 is primarily located in mitochondria. Studies
report SIRT3 is also localized in the nucleus and cytosol in its uncleaved form while the cleaved
form is localized in the mitochondria (Cooper and Spelbrink, 2008). Targets of SIRT3 include
acetyl-CoA, the mitochondrial permeability pore and mitochondrial complexes 1-5. Upon
regulation of the respiratory chain, SIRT3 influences energy production (Shimazu et al., 2010;
Verdin et al., 2010). The activation of these targets may be associated with SIRT3’s actions in
the cell’s stress response pathway. SIRT3 is also involved in mitochondrial biogenesis where it
mediates the effects of PGC-1α by acting as a downstream target (Kong et al., 2010). Although
SIRT3 plays many critical roles in mitochondrial function, and its family members are
efficacious in various diseases, most studies on SIRT3 have been performed in the heart, and
nothing is known about its therapeutic effect in neurons.
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1.8 Models of Parkinson’s disease
Various in vitro and in vivo models have been generated to replicate the mechanisms underlying
PD and the symptoms resulting from PD. Each model utilizes toxins or genetic manipulations to
induce parkinsonian features. Since each model does not fully replicate disease mechanisms or
symptoms, new models that incorporate these important aspects of PD will be invaluable for
therapeutic development. Although the present models lack certain characteristics of PD
mechanisms and symptoms, each have been useful in the development of therapeutics that are
currently available to treat PD.
1.8.1 In vitro models
In vitro models have been extremely valuable tools for determining the cellular mechanisms
underlying PD. Toxins have been applied to in vitro models to study idiopathic forms of PD,
while the insertion of exogenous DNA known to be mutated in individuals with PD into
individual cells has given insight into the mechanisms causing familial PD. Organotypic co-
cultures contain multiple cell types allowing for changes in one cell type to be directly compared
to changes in another cell type while they are in the same microenvironment. Such cultures can
be maintained for up to a few weeks in culture allowing the neurons to mature, which more
closely resembles neurons degenerating in PD. Other advantages of organotypic co-cultures
include neuronal and glial interactions, maintenance of neuronal connections and growth of
synapses while in culture (Lyng et al., 2007). More importantly, organotypic cultures express
dopamine, homovanillic acid and tyrosine hydroxylase from as little as 6 days to 17 days after
culturing (Lyng et al., 2007). Compared to other in vitro models, the production of organotypic
co-cultures is more complicated since slices of multiple brain areas must be obtained from
embryonic and post-natal rodents. Primary cultures have been employed in many studies to
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discover the biochemical pathways responsible for PD pathology (Hoglinger et al., 2003a;
Lannuzel et al., 2003). Synapses actively form between neurons in primary cultures, but the
intrinsic activity of synapses seen in the brain is not usually found in these cultures. Indeed,
these in vitro neurons were less spontaneously active and never fired in bursts, unlike in vivo
neurons (Rayport et al., 1992). While primary cultures benefit from having neurons and glia and
display the typical characteristics of mature neurons, such as TH and dopamine, generally very
few cells from the culture have these characteristics. However, the presence of dopamine
autoreceptors was identified by neuronal activity in response to quinpirole, a dopamine receptor
agonist (Cardozo, 1993). In vitro models such as cell lines have the advantage of being quickly
generated allowing reproducible effects to be shown. Even the differentiation of neuroblastoma
to neuronal-like cells can take place in a time frame as short as 72 hours, allowing for high-
throughput findings that can be inferred to actual neurons. Disadvantages of cell lines include
being dissimilar from actual neurons and having a microenvironment that is non-physiological.
Since the genes of cells in culture are easily manipulated and their protein products are rapidly
produced, the biochemical pathways and mechanisms involved in PD can be efficiently studied.
Indeed, specific molecular pathways underlying PD can be directly targeted in cellular models
(Dawson et al., 2010). In addition, specific cell types can be isolated to study changes that are
unique to that cell type. Studies show that the various toxins used to create models reliably act
upon their intended target and repeatedly produce cellular dysfunction and markers associated
with the disease state (Sherer et al., 2003b). The efficient nature of in vitro cultures enables their
use for rapid screening of therapeutic compounds. As a result, there will be a quicker “bench to
bedside” process enabling therapeutics to become available on the market more quickly.
However changes observed in cellular models do not exactly correlate with those seen in the
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disease state, thus more advanced in vivo models, which better emulate disease states, must be
utilized to further substantiate the findings obtained using in vitro models.
In vitro models lack the surrounding micro environment provided by in vivo models, but benefit
from having a more controllable environment (Alberio et al., 2012). Furthermore, most in vitro
models cannot recapitulate the complexity of multiple connections formed by cells from different
areas of the brain. This is highly disadvantageous considering PD pathology affects complex
networks organized between the SNc and striatum. Moreover, multiple cell types are affected,
which makes it difficult to model with most in vitro models, except for those models employing
organotypic cultures that are comprised of neurons from various brain areas (Snyder-Keller et
al., 2001). Given that in vitro models are isolated from organisms, they are unable to show how
changes mediated by therapeutic compounds affect disease symptoms and physiology of the
organism as a whole. Although in vitro models have many benefits, it is necessary to further
characterize and develop in vitro models which display the multitude of pathological
mechanisms found in PD, rather than modeling a single mechanism.
1.8.2 In vivo models
The use of animal models in PD research has been the main approach for developing therapeutic
compounds and studying disease symptoms. Toxin and transgenic animal models have been
used to replicate the pathology in sporadic and familial PD (Sherer et al., 2003a; Rockenstein et
al., 2007). The reserpine-treated rat model was developed in the late 1950’s, resulting in the
discovery of L-dopa as a potential treatment for parkinsonian symptoms. Reserpine application
in rodents induces muscle rigidity, postural flexion, and akinesia, closely resembling symptoms
observed in parkinsonian patients (Carlsson et al., 1957). However, administration of reserpine
produces a model that is acute, reversible and fails to replicate the pathology underlying
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idiopathic PD. A further disadvantage associated with the reserpine-treated rat is that serotonin
and noradrenaline levels are depleted to a greater extent compared to idiopathic PD. The
unilaterally-lesioned 6-OHDA rat model of PD has proved extremely useful in providing insight
into pathological, electrophysiological and pharmacological changes which occur following
degeneration of the nigrostriatal pathway, increasing our understanding of the mechanisms
underlying the generation of parkinsonian symptoms. Following stereotaxic injection of 6-
OHDA into the medial forebrain bundle, dopamine cell loss is more profound compared to that
observed following injections of 6-OHDA into the SNc (Carman et al., 1991). Metabolism of 6-
OHDA results in a phenomenal amount of oxidative stress, promoting deleterious effects of free
radicals on mitochondrial function, membrane stability and DNA integrity, which ultimately
results in cell death (Schwarting and Huston, 1996). Following unilateral administration of 6-
OHDA, rodents exhibit a spontaneous rotational behaviour ipsiversive to the side of the lesion
which represents an anti-parkinsonian action. Although exhibition of contraversive rotational
behavior in 6-OHDA lesioned animals is thought to represent an anti-parkinsonian action,
parkinsonian symptoms such as rigidity, tremor and bradykinesia are not apparent in these
animals (Ungerstedt, 1968). Furthermore adverse effects including adipsia, aphagia and high
mortality rates make this model less attractive (Ungerstedt, 1971). The MPTP mouse model is
widely used to evaluate potential therapeutic agents for PD (Cannon and Greenamyre, 2010).
This model shows progressive neurodegeneration of the nigro-striatal pathway, and sometimes
motor deficits. Thus, potential neuroprotective agents can be evaluated using post-mortem
measures of dopamine system integrity. The MPTP-lesioned mouse is easy to reproduce, and
more importantly, has proved to be one of the most useful models for assessing neuroprotective
potential of compounds in PD (Pothakos et al., 2009). One drawback of this model is that it does
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not result in the generation of protein aggregates, which are considered an important component
of PD pathology.
Upon the discovery of new genes linked with familial PD, classical toxin-induced and genetic
models became limited in their scientific value because they were unable to represent the various
forms of PD and their associated pathology. As a result, many animal models now express these
newly discovered genes that underlie PD (Li et al., 2010). This has led to the rapid identification
of the symptoms and pathology associated with certain genetic forms of PD. Models utilizing
AAV-mediated delivery and over-expression of α-synuclein in the rodent replicate the
progression of cell death and molecular pathology most closely to that observed in patients
(Koprich et al., 2010). The model also displays behavioral deficits consistent with a
parkinsonian phenotype. Following unilateral delivery of AAV-α-synuclein, dopaminergic
neurons become dysfunctional, exhibit abnormal axonal morphology and proteinase K-resistant
aggregations of α-synuclein in both terminals and nigral cell bodies, as well as increased striatal
dopamine transporter binding (DAT) (Koprich et al., 2010). These changes are accompanied by
behavioral abnormalities. At later stages, the pathology has progressed further, with marked
presence of dystrophic neurites, reductions in striatal TH, DAT and dopamine and a significant
loss of TH neurons within the nigra.
Animal models are critical for the development of neuroprotective strategies because they have
enabled the discovery of various sub-cellular deficits and various symptoms that arise from
mutations in the genes associated with PD and more importantly, reveal changes that occur in
live organisms. Moreover, these models enhance the clinical applicability of drugs since the
effect of therapeutic interventions on a live organism expressing genetic mutations closely
predicts the effect of the drug on patients. Although the various models replicate one particular
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aspect of PD, they do not emulate the multifaceted nature of the disease. This is demonstrated
by the inhibition of an individual mechanism by a particular toxin or genetic manipulation, rather
than a multitude of mechanisms as seen in PD patients. Furthermore, certain brain areas that are
not typically affected during PD appear to be vulnerable to degeneration (van der Putten et al.,
2000). Application of rotenone in animal models leads to global degeneration of neurons rather
than specific degeneration within the substantia nigra alone (Meredith et al., 2008). Markers
such as Lewy bodies that are typically found in patients exhibiting PD are not consistently
reproduced by the various toxin and genetic models (Dauer and Przedborski, 2003). Despite the
expression of these markers, many distinguishing features including nigral degeneration and loss
of tyrosine hydroxylase expression do not always occur. Although these models suffer from
disadvantages, they have been extremely useful in delineating gene specific pathology and
symptomatic treatments.
Given the advantages and disadvantages of in vitro and in vivo models with respect to
developing therapeutics to treat PD, these models must be further developed to better emulate all
the pathological mechanisms found in PD; then both models can be utilized to successfully
predict if a neuroprotective agent will be efficacious in clinical trials.
1.9 Hypothesis and aims
In PD, a variety of sub-cellular dysfunctions cause neurodegeneration of the SNc which
ultimately lead to the symptoms of PD. By understanding the function of mutations linked with
familial PD, we know that a number of separate mechanisms can trigger PD pathology.
Moreover, the utilization of environmental toxins in PD models has further highlighted that
multiple mechanisms induce PD pathology. Although a number of dysfunctional mechanisms
exist, the role of mitochondria in PD pathology is becoming more apparent. Thus, I
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hypothesized that altered mitochondrial function, due to the dysfunction of sub-cellular domains
and inhibition of mitochondrial dynamics, is a primary cause of disease pathology. Furthermore,
potential neuroprotective agents that target the mitochondria will be effective at protecting
degenerating neurons of the SNc. The initial hypothesis of this study was, that no matter what
the initial trigger, a single sub-cellular abnormality activates additional sub-cellular pathways,
such as the mitochondria, that further enhances the pathology of PD. Secondly, I hypothesized
that a correlation between the outcomes of neuroprotective agents in an in vitro model and those
of clinical trials can validate the effectiveness of an in vitro model at predicting potential
neurprotective agents to treat PD. Given the importance of mitochondrial function in the
pathology of PD, and the existence of LRRK2 on mitochondrial membranes, I hypothesized that
LRRK2 alters mitochondrial dynamics causing mitochondrial dysfunction. Moreover, a
potential neuroprotective agent such as SIRT3, which is involved in mitochondrial function, will
be effective at protecting neurons from degeneration.
The first aim of the studies described in Chapter Two was to understand how different triggers
associated with initiating the pathology of PD interact with other known components of the
neurodegenerative process associated with PD. The initial aim of these studies revealed that, no
matter what the initiating trigger, mitochondrial dysfunction occurs early in the cell death
pathway. This suggests that malfunction of mitochondria is central to all pathologies associated
with PD. The studies that were completed as part of the initial aim led to the generation of a cell
model of PD that took into account all known causes of PD. This model is quick, easy and
reliable to produce, thus, in Chapter Three my second aim of the studies validated this as a useful
model to test potential neuroprotective agents in PD using compounds that had previously passed
screening processes and were already in clinical trials. The third aim of the studies described in
Chapter Four was to further investigate how mitochondrial function is affected in PD. Since
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mitochondrial dynamics are required for physiological mitochondrial function, I investigated
changes in mitochondrial fission and fusion in cell models of PD. This aim revealed that
mitochondrial fusion is inhibited in a common genetic variant of PD. The fourth aim of the
studies outlined in Chapter Five was to evaluate the neuroprotective ability of two novel
compounds in our novel model of PD. This aim affirmed the importance of targeting the
mitochondria when trying to prevent the neurodegeneration associated with PD.
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Chapter 2
Mitochondrial Dysfunction Precedes other Sub-Cellular Abnormalities in an In Vitro Model
Linked with Cell Death in Parkinson’s Disease
A modified version of this chapter was previously published as:
Yong-Kee CJ, Sidorova E, Hanif A, Perera G, Nash JE (2012) Mitochondrial dysfunction precedes other sub-cellular abnormalities in an in vitro model linked with cell death in
Parkinson's disease. Neurotoxicity research 21:185-194.
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2 Mitochondrial Dysfunction Precedes other Sub-Cellular Abnormalities in an In Vitro Model Linked with Cell Death in Parkinson’s Disease
2.1 Introduction
By the time patients present with symptoms of PD, approximately 50% of the nigro-striatal
pathway has degenerated (Bernheimer et al., 1973), and a substantial proportion of the remaining
nigral neurons are undergoing some form of cell stress (Murray et al., 1995). Nevertheless
symptoms are typically not severely disabling and patients still have a good quality of life. Thus,
if it was possible to prescribe a disease-modifying agent that stopped or slowed down disease
progression at this stage of the disease, patients would not experience the advanced, severely
disabling stages of the disease. Development of such disease modifying strategy depends
critically upon understanding the sub-cellular mechanisms underlying neurodegeneration in PD.
Although 90-100% of cases of PD are believed to be sporadic, clues regarding the mechanisms
underlying cell death in PD have arisen from the 5-10% of cases that have been shown to be
genetically linked. All 15 loci on the 11 genes that have been linked to familial PD encode
proteins involved in the handling or removal of misfolded proteins, either at the level of the UPS,
macro-autophagy and CMA (Kitada et al., 1998; Ardley et al., 2004; Ramirez et al., 2006; Di
Fonzo et al., 2007; Lees and Singleton, 2007; Hatano et al., 2009), the regulation of
mitochondrial function (Lannuzel et al., 2003; von Bohlen und Halbach et al., 2004; Sherer et al.,
2007) and oxidative stress (Martinat et al., 2004; Shendelman et al., 2004). Furthermore, the
genetic mutations linked with familial PD cause dysfunction in at least one, if not all, of these
regulatory mechanisms (Hardy, 2010; Magen and Chesselet, 2010). For example, loss-of-
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function mutations in the E3 ubiquitin-protein ligase Parkin, a protein most commonly associated
with early-onset autosomal recessive PD (Kitada et al., 1998) was originally shown to play a role
in tagging proteins for degradation by the proteasome (Cookson, 2003; Sakata et al., 2003).
More recently, Parkin has also been shown to play a role in the regulation of mitochondrial
dynamics (Yun et al., 2008; Whitworth and Pallanck, 2009) as well as in CMA (Olzmann and
Chin, 2008). A second example of a protein associated with familial PD, which appears to cause
multiple problems at the sub-cellular level is α-synuclein. Mutations in α-synuclein cause
impaired UPS and lysosomal function (Stefanis et al., 2001; Tanaka et al., 2001; Cuervo et al.,
2004; Chu et al., 2009), and increased susceptibility of dopaminergic neurons to mitochondrial
toxins such as MPP+, as well as oxidative stress (Kanda et al., 2000; Qian et al., 2008). Finally,
post mortem studies in parkinsonian patients suggest that similar mechanisms are responsible for
causing cell death in both familial and idiopathic cases of this disease, since all show evidence of
impaired function of mitochondrial complex 1, UPS and lysosome, as well as increased oxidative
stress in the SNc and other affected brain regions (Parker et al., 1989; Jenner, 1993; Chu et al.,
2009). If it were possible to identify a common sub-cellular dysfunction that was initiated by all
known cell mechanisms associated with neurodegeneration in PD, this would represent a
valuable target for the development of a disease modifying agent since it would allow the rescue
of the remaining 50% of neurons in the SNc.
Given that multiple sub-cellular mechanisms are involved in Parkinson’s pathology, I
hypothesized that there is a single point of convergence of these mechanisms which results in
cellular stress. The studies described in this chapter sought to identify such a point of
convergence by using a catecholaminergic neuroblastoma cell line, which has previously been
used to study the mechanisms underlying cell death in PD (Dadakhujaev et al., 2010; Xie et al.,
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2010; Xie et al., 2011; Nonaka and Hasegawa, 2009; Wu et al., 2009), in combination with
toxins that induce cell death via the three mechanisms commonly associated with
neurodegeneration in PD: inhibition of mitochondrial complex 1 (rotenone), inhibition of the
UPS using Z-Ile-Glu(OBut)-Ala-Leu-H (PSI), and disruption of the lysosomal membrane via
5,8-dihydroxy-1,4-naphthoquinone (naphthazarin). In order to determine the effect of these
toxins in early, as compared to later stages of cell stress, when cell damage is unlikely to be
reversible, sub-cellular function was measured after three and twenty four hours of exposure to
toxin.
2.2 Materials and Methods
2.2.1 Materials
SH-SY5Y cells were purchased from ATCC (USA), Dulbecco’s Modified Eagle’s Medium
(DMEM), bovine calf serum, from Wisent (Canada) and trypsin from Sigma (USA). 5,8-
dihydroxy-1,4-naphthoquinone (naphthazarin) and rotenone were purchased from Sigma (USA),
Z-Ile-Glu(OBut)-Ala-Leu-H (PSI) from BIOMOL (USA) and alamar blueTM from Biosource
(Canada). Propidum iodide was purchased from Invitrogen (USA). Lysosensor green and JC-1
were purchased from Invitrogen (USA) and proteasome sensor vector from Clontech (USA).
Ubiquitin anti-rabbit antibody was purchased from Dako (USA), β-actin anti-mouse antibody
from Sigma (USA), anti-mouse and anti-rabbit HRP-conjugated secondary antibodies from
Jackson Laboratories (USA) and nitrocellulose membrane was purchased from BIORAD (USA).
PSI was dissolved in 0.1% dimethyl sulfoxide (DMSO) plus DMEM, and rotenone and
naphthazarin in DMEM only.
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2.2.2 Cell culture
Human dopaminergic neuroblastoma SH-SY5Y cells (ATCC) (P10-P30) were grown in DMEM
(consisting of 6400 mg/L NaCl, 3700 mg/L NaHCO3, 400 mg/L KCl and 584 mg/L L-glutamine)
supplemented with 5% bovine calf serum in a sterile humidified chamber (37 oC, 5% CO2, 95%
O2) (Incubator: MCO-20AIC, Sanyo, USA) until confluent. Cells were passaged using 0.1%
trypsin for 5 min, then pelleted by centrifuging at 340xg for 5 min (Allegra 6R Centrifuge,
Beckman Coulter, USA).
2.2.3 Cell viability assays
SH-SY5Y cells were grown in 96-well plates at 1.0 x 105 cells/ml. After 24 hours, cells were
exposed to full dose response curves of naphthazarin, PSI or rotenone (0 µM–1000 µM) in the
presence of the redox sensitive dye alamar blue (0.4% of final volume). Cell viability was
assessed by measuring the change in fluorescence of alamar blue (Ex. 544 nm, Em. 590 nm)
using a plate reader (FLUOstar OPTIMA, BMG Labtech, USA) at 3 and 24 hour time points.
Data are expressed as percentage cell viability compared to control ± SEM (n = 4).
2.2.4 Cell death assays
SH-SY5Y cells were grown in 6-well plates at a density of 2.0 x 105 cells/ml. After 48 hours,
naphthazarin (2.17 µM), PSI (80.0 µM) or rotenone (40.0 µM) was added to cells. Three or 24
hours following addition of toxin, propidium iodide (2 µM) was added to cells for 5 minutes, and
the number of propidium iodide positive (dead) cells quantified. Data are presented as
percentage of propidium iodide positive cells out of 150 counted cells ± SEM (n = 4).
49
2.2.5 JC-1 and lysosensor green assays
Twenty-four hours post plating of SH-SY5Y cells, naphthazarin (2.17 µM), PSI (80 µM) or
rotenone (40 µM) were added. Following incubation with toxins for 3 or 24 hours, JC-1 (2 µM)
or lysosensor green (2 µM) was added to each well and incubated with the cells for 30 minutes,
then each well was aspirated and washed with DMEM before recording fluorescence intensity of
comparisons test post-hoc showed that toxins caused a significant decrease in cell viability.
Naphthazarin: *** P < 0.001, compared to vehicle (media). PSI: $ P < 0.05, $$ P < 0.01, $$$ P
< 0.001 compared to vehicle (DMSO). Rotenone: & P < 0.05, && P < 0.01, &&& P < 0.001
compared to vehicle (media). (c) Cell death in SH-SY5Y was assessed following 3 or 24 hours
exposure to naphthazarin (2.17 µM), PSI (80 µM) or rotenone (40 µM) using propidium iodide
(PI). Data are presented as mean number of PI positive cells out of 150 ± SEM (n = 4). Two
way ANOVA showed a significant difference in propidium iodide uptake following incubation
with toxins for 3 and 24 hours compared to control (media for naphthazarin and rotenone;
DMSO for PSI) (F4 (toxin) = 26.49, F1 (time) = 56.76; F4 (interaction toxin x time) = 16.65).
Bonferroni test post-hoc showed a significant increase in propidium iodide uptake in the cells
following incubation with naphthazarin, PSI and rotenone for 24 hours compared to control, with
no significant effect of any toxin following incubation for 3 hours (*** P < 0.001).
53
a
0 0.1 1 10 100 10000
20
40
60
80
100
120
*$
***$$$&&
***$$&
***$$$& ***
$$$&&&
concentration (τM)
cell viability (% control)
b
0 0.1 1 10 100 10000
20
40
60
80
100
120
***$$$&&&
***&& ***$
&&
***$$$&&&
concentration (τM)
cell viability (% control)
c
Napthazarin PSI RotenoneMedia DMSO
3 24 0
10
20
30
40
50
******
***
time (hr)
number of PI positive cells
3 hours
24 hours
54
2.3.2 Effect of toxin exposure on mitochondria, UPS and lysosomes
An early marker of mitochondrial impairment is depolarisation of the mitochondrial membrane
potential. To assess the impact of toxins on mitochondrial function, naphthazarin, PSI or
rotenone were added to SH-SY5Y cells for 3 or 24 hours, and JC-1 was utilised to measure
mitochondrial membrane potential. After 3 hours exposure to all toxins, the ratio of JC-1
monomer to aggregates was significantly increased compared to controls (vehicle-treated cells),
indicating a depolarisation in mitochondrial membrane potential. Post hoc analysis showed that
the monomer to aggregate ratio following naphthazarin, PSI and rotenone treatment was
increased by 2.5-fold, 2-fold and 4.6-fold respectively compared to control (media for
naphthazarin and rotenone; 0.1% DMSO for PSI) (Fig. 2.2). Longer exposure to rotenone (24
hours) enhanced mitochondrial membrane depolarisation 6.1-fold compared to control. In
contrast, following 24 hours exposure to naphthazarin and PSI, there was no significant effect
compared to control (Fig. 2.2).
55
Figure 2.2. Effect of compounds on mitochondrial membrane potential in SH-SY5Y cells.
SH-SY5Y cells were incubated with naphthazarin (2.17 µM), PSI (80 µM) or rotenone (40 µM)
for 3 hours or 24 hours. Changes in mitochondrial membrane potential were assessed using JC-
1. Data are expressed as ratio of mean fluorescence of aggregate to monomer ± SEM (N = 4).
One-way ANOVA showed a significant effect of toxins on mitochondrial membrane potential, 3
hours (F4 = 12.32) and 24 hours (F4 (toxin) = 22.85) following exposure. Tukey’s multiple
comparison test post-hoc: * P < 0.05, *** P < 0.001.
56
Media DMSO NAP PSI ROT0.0
0.5
1.0
1.5 ******
*
3 hours
24 hours
ratio
of m
onom
er/a
ggre
gate
(ave
rage
FIU
)
*
57
To determine the effect of naphthazarin, PSI and rotenone on proteasomal function, toxins were
incubated with cells previously transfected with the proteasome-sensitive fluorescent reporter
ZsProSensor-1 for 3 or 24 hours. When the proteasome is functioning normally, ZsProSensor-1
is broken down, however, when proteasomal function is compromised, ZsProSensor-1
accumulates, resulting in an increase in fluorescence intensity. Three hours of exposure to PSI
caused a significant increase (398%) in ZsProSensor-1 fluorescence compared to control,
indicating impaired proteasomal function however there was no significant effect of
naphthazarin or rotenone on proteasomal function. Following 24 hours exposure, impaired
proteasomal function was observed with all 3 toxins. Naphthazarin, PSI and rotenone increased
the fluorescence intensity of ZsProSensor-1 by 1135%, 2217% and 712% respectively (Fig. 2.3).
58
Figure 2.3. Effect of toxins on proteasomal function in SH-SY5Y cells. Cells were
transfected with the proteasome-sensitive fluorescent reporter ZsProSensor-1 then incubated
with naphthazarin (2.17 µM), PSI (80 µM) or rotenone (40 µM) for (a) 3 hours or (b) 24 hours,
then fluorescence intensity measured. Data are expressed as mean fluorescence ± SEM (N = 4).
One-way ANOVA showed a significant effect of toxins on proteasomal function, 3 hours (F4
(toxin) = 3.15) and 24 hours (F4 = 90.61) following exposure compared to media control
(naphthazarin and rotenone) or DMSO control (PSI). Tukey’s multiple comparison post-hoc: * P
< 0.05, *** P < 0.001.
59
3 hours
24 hours
Media DMSO NAP PSI ROT
aver
age f
luor
esce
nce (
%co
ntro
l)
050
100150200
5001000150020002500 ***
*******
60
To determine the effect of naphthazarin, PSI and rotenone on lysosomes, cells were incubated
with the 3 toxins in the presence of lysosensor green, which fluoresces when taken into the
lysosome, serving as a marker of lysosomal function. Three hours of toxin exposure led to a
significant difference in lysosensor green fluorescence compared to control (Fig. 2.4). Post hoc
analysis showed that naphthazarin decreased fluorescence by 67.24% compared to vehicle,
whereas PSI and rotenone had no effect at this time. Following 24 hours of toxin exposure, there
was a significant effect of all toxins on fluorescence of lysosensor green compared to vehicle.
Post hoc analysis showed that naphthazarin, PSI and rotenone decreased fluorescence intensity
60.93%, 40.35% and 38.05% respectively.
61
Figure 2.4. Effect of toxin exposure on lysosomal function. Lysosomal function was assessed
via measurement of fluorescence of lysosensor green 3 hours or 24 hours after the addition of
naphthazarin (2.17 µM), PSI (80 µM) or rotenone (40 µM). Data are expressed as mean
fluorescence ± SEM (N = 4). One-way ANOVA showed a significant effect of toxins on
lysosomal function, 3 hours (F4 = 3.63) and 24 hours (F4 = 15.31) following toxin exposure
compared to media control (naphthazarin and rotenone) or DMSO control (PSI). Tukey’s
multiple comparison test post-hoc * P < 0.05, ** P < 0.01, *** P < 0.001.
62
0
40
80
120
**
***
3 hours24 hours
**
*
aver
age f
luor
esce
nce (
% co
ntro
l)
Media DMSO NAP PSI ROT
63
2.3.3 Effect of toxins on ubiquitin levels
Post mortem studies have shown increased formation of ubiquitin aggregates in affected brain
regions of patients with PD (Leigh et al., 1989). However, it is not known whether such
pathologies arise due to dysfunction of the mitochondria, proteasome or lysosome, or indeed all
three. As shown in Figure 2.5a, b, a 3 hour exposure to naphthazarin, and PSI, but not to
rotenone altered ubiquitin expression. Following 3 hours of toxin exposure, naphthazarin and
PSI increased ubiquitin expression by 315.94%and 323.19% respectively compared to controls
(Fig. 2.5a,b). Following 24 hours of toxin exposure, ubiquitin expression was dramatically
increased compared to control. Post hoc analysis showed that naphthazarin, PSI and rotenone
increased ubiquitin levels by 7495%, 8954% and 10902% respectively compared to vehicle (Fig.
2.5a,c).
64
Figure 2.5. Western blots to show changes in ubiquitin levels following toxin exposure. SH-
SY5Y cells were exposed to naphthazarin (2.17 µM), PSI (80 µM) or rotenone (40 µM) for 3
hours or 24 hours, harvested, and SDS-PAGE followed by Western blot was performed to detect
levels of ubiquitin. (a) Data are expressed as mean optical density ± SEM (n = 4). One-way
ANOVA showed significant effects following 3 hours (F4 = 5.17) and 24 hours (F4 = 253.1)
following toxin exposure compared to media control (naphthazarin and rotenone) or DMSO
control (PSI). Dunnett’s multiple comparison test post-hoc * P < 0.05, ** P < 0.01, *** P <
0.001. (b) Representative Western blot following 3 hours toxin exposure. (c) Representative
Western blot following 24 hours toxin exposure.
65
a
0
200
400
6006000
8000
10000
12000
***
3 hours24 hours
**
Media DMSO NAP PSI ROT
***
***
*
aver
age o
ptic
al d
ensi
ty(%
con
trol)
bUbiquitin
ß-actin 42 kDa
200-250 kDa
MED
IA
DM
SO
NA
P
PSI
RO
T
c
DM
SO
NA
P
PSI
RO
T
Ubiquitin
ß-actin 42 kDa
200-250 kDa
3 hours
24 hours
66
2.4 Discussion
SH-SY5Y cells were exposed to toxins that cause their effects via different mechanisms that
have been linked with neurodegeneration in PD, i.e., impairment of mitochondrial, lysosomal or
UPS function. The aim of this study was to determine a common cell stress mechanism
triggered by these different sub-cellular dysfunctions during the early stages of cell stress, so that
a target for a disease modifying agent may be revealed. Thus, it was necessary to utilise
concentrations of toxins that caused a significant level of cell stress (as measured by decreased
cell viability), while causing minimal levels of cell death, as this scenario recapitulates the
situation at the time of initial diagnosis of PD, when approximately 50% of neurons in the SNc
are alive, although may be undergoing some form of sub-cellular stress (Bernheimer et al.,
1973). Whilst in this study, cells were exposed to high concentrations of toxins for a relatively
short period of time (24 hrs) to induce acute toxicity. Previous studies have shown that this acute
model does reproduce the cell death processes implicated in PD (Yong-Kee et al., 2011a).
Following 3 hours exposure of all toxins at concentrations that caused an approximate 50%
decrease in cell viability (EC50%), no cell death was observed. Following 24 hours exposure,
the approximate EC50% of all toxins caused a significant increase (approximately 2-4 fold) in
cell death. Thus, following three hours of EC50% toxin exposure, 50% of SH-SY5Y cells
exhibit decreased cell viability, and the remaining 50% are likely to be undergoing some form of
sub-cellular stress, since there is a further decrease in cell viability after 24 hours. This scenario,
in which cells are under-going moderate to high levels of cell stress, although have not yet died,
is likely to be comparable to the situation in surviving neurons of SNc in parkinsonian patients.
Specifically, following 24 hours of toxin exposure, we see a loss of cell viability and an increase
in cell death. Thus, this represents a mixed population of cells, in which some are healthy, some
67
are no longer viable, and some are dead (Bernheimer et al. 1973). Thus, exposure of cells to
toxins that mimic cell death mechanisms associated with PD for short periods (3 and 24 hours) is
a useful model for studying the early events which take place when cells are under cell stress.
Given that catecholaminergic neurons are the most susceptible to neurodegeneration in PD
(Hornykiewicz, 1972; Braak et al., 1996), the catecholaminergic neuroblastoma cell line, SH-
SY5Y, which has previously been used as a model for studying cell death mechanisms in PD
was selected for our studies (Dadakhujaev et al., 2010; Xie et al., 2010; Xie et al., 2011; Nonaka
and Hasegawa, 2009; Wu et al., 2009). Whilst primary mesencephalic cultures are a more
physiological representation of neurons of the substantia nigra pars compacta, it is difficult to
produce them in large quantities, and so it would not have been possible to perform the assays
utilised in the present studies. Previous studies have shown that lower concentrations of
rotenone (15-50 nM) were sufficient to cause cellular dysfunction (Liss & Roeper, 2001;
Shamoto-Nagai et al., 2003; Lannuzel et al., 2003; Lannuzel et al., 2006); in the present study
however, the concentration of rotenone required to decrease cell viability by 50% was 680.0 µM.
There are at least three possible explanations for these variations in the EC50% of rotenone:
firstly, cell type, secondly, the period of time in which rotenone was exposed to the cells, and
finally, the parameter utilised to measure cellular dysfunction. In one study, 15 nM of rotenone
was sufficient to inhibit mitochondrial complex 1 activity by 50% in acute mouse brain slices
(Liss & Roeper, 2001). Indeed, neurons are known to be much more sensitive to cell stress than
other cell types, such as SH-SY5Y cells that were used in the present study. Whilst SH-SY5Y
cells have a neuronal phenotype, they are not neuronal cells, and so are less sensitive to rotenone
as has been shown previously (Shamoto-Nagai et al., 2003; Lannuzel et al., 2003). Another
explanation for the difference in EC50s between the study carried out by Shamoto-Nagai and
colleagues and also Lannuzel et al., compared to the current study, is that in these two previously
68
published studies, different markers of cellular dysfunction were used to measure the impact of
rotenone on cellular dysfunction. Shamoto-Nagai and colleagues measured inhibition of
mitochondrial complex 1 activity, whereas in the current study, changes in cell viability were
assessed. Since inhibition of mitochondrial complex 1 will precede a loss of cell viability, it is
most likely that the rotenone concentration required to cause a 50% inhibition of the
mitochondrial complex 1 is less than that required to cause a 50% decrease in cell viability.
Lannuzel and co-workers utilised loss of TH immunoreactivity as a marker of cell death.
Previously it has been shown that, in dopaminergic neurons, loss of TH phenotype precedes loss
of cell viability and neuronal cell death (Paul et al., 2004), thus providing a possible explanation
as to why the concentration of rotenone required was lower than in the present study.
Interestingly, Shamoto-Nagai and co-workers also used SH-SY5Y cells, and showed that 25-50
nM of rotenone was sufficient to kill 50% of the cells. However, in this study, 50% of cells died
following incubation with toxins for 72-120 hours. Indeed, as in our study, Shamoto-Nagai et
al., observe no cell death following incubation with rotenone for 24 or 48 hours. Finally, in the
present study, we describe an acute model of cellular stress, in which the initial sub-cellular
effects of toxic insult were evaluated. This justifies the use of higher concentrations of toxins for
a shorter time course.
When mitochondria, UPS and lysosomal function was assessed following 3 and 24 hours
exposure to naphthazarin, PSI or rotenone, all toxins caused depolarisation of the mitochondrial
membrane potential at 3 hours, whereas after 24 hours, only rotenone affected mitochondrial
function. This apparent recovery of mitochondrial function may represent a compensatory
mechanism, indeed it has been shown that pre-conditioning of the mitochondria occurs with mild
forms of toxic insults (Gidday, 2006; Busija et al., 2008). Three hours following toxin exposure,
only PSI inhibited proteasomal function, whereas following 24 hours exposure, all 3 toxins
69
inhibited UPS function. Similarly, with respect to lysosomal function, after 3 hours of toxin
exposure, only naphthazarin decreased lysosomal function, whereas, following 24 hours of
exposure to toxins, all 3 toxins inhibited lysosomal function. The early effect of PSI on UPS
function and naphthazarin on lysosomal function confirms that the primary effects of these
toxins are via inhibition of the proteasome and lysosome respectively. Finally, following three
hours of toxin incubation, only naphthazarin and PSI increased levels of ubiquitination, whereas
following 24 hours treatment, all three toxins caused a significant increase in ubiquitin
expression. Both the UPS and lysosome are important for the breakdown of ubiquitinated
proteins (Kettern et al., 2010; Hochstrasser, 1992; Kubota, 2009; Todde et al., 2009), therefore,
the observed increase in ubiquitinated proteins shortly after incubation of cells with toxins that
inhibit these functions is not surprising. The finding that all 3 toxins increased ubiquitin levels
24 hours following toxin exposure is expected, since both lysosome and the proteasome-
mediated de-ubiquitination require ATP, thus extended periods of mitochondrial inhibition will
result in a lack of ATP, hindering physiological function of lysosomes and the UPS. Autophagic
processes require acidified lysosomal compartments that are maintained by ATP-dependent
proton pumps (Terman et al., 2010). Thus, reduced ATP levels may hinder lysosomal function,
by decreasing the rate of activity of the proton pump, which would slow down the ability of the
lysosome to degrade unwanted proteins. Similarly, UPS function relies heavily on a constant
supply of ATP. Access to the proteasomal core is regulated by the opening and closing of the
proteasomal cap (Betarbet et al., 2005). Therefore, since this process is ATP-dependent,
decreased ATP will also prevent protein degradation by the proteasome. Furthermore, excess
proteins in the cytosol may ultimately become deleterious to the cell because increased
misfolded proteins may form toxic aggregates which cause cell death (Mytilineou et al., 2004).
70
Indeed, reduced ATP levels were found to hamper proteasomal activity in primary cultures
(Hoglinger et al., 2003).
Interestingly, both proteasomal and lysosomal inhibition reduced mitochondrial membrane
potential. Mitochondrial dysfunction caused by inhibition of the proteasome has been shown in
other in vitro studies (Qui et al., 2000; Ding and Keller, 2001; Hoglinger et al., 2003). Inhibition
of the proteasome results in an increase in the number of misfolded and aggregated proteins,
which has deleterious effects on the cell, by increasing the number of reactive oxygen species.
The resulting increase in oxidative stress further augments cell stress by increasing influx of Ca2+
into the cell, leading to excitotoxic conditions (Demuro et al., 2005; Danzer et al., 2007).
Increased excitotoxicity, enhances Ca2+ influx into the mitochondria, resulting in the
accumulation of free radicals within the mitochondria, which induces depolarisation of the
mitochondrial membrane potential (Ward et al., 2000) and loss of ATP (Budd and Nicholls,
1996). Whilst, there are no previous studies to show that inhibition of lysosomal function
impacts mitochondrial function, it has been shown that following over-expression of wild-type
α-synuclein in neuronal cultures, this protein accumulates in mitochondria leading to reduced
activity of mitochondrial complex 1 and subsequent mitochondrial dysfunction (Devi et al.,
2008). Whilst the reason why α-synuclein accumulates in the mitochondria is unclear; it is
known that α-synuclein is usually degraded by the lysosome, and that mutant α-synuclein
inhibits lysosome-mediated breakdown of misfolded and toxic proteins, including α-synuclein
(Cuervo et al., 2004; Sarkar et al., 2007). Impaired lysosomal function would also prevent
lysosome-mediated uptake of free radicals, which would in turn cause mitochondrial dysfunction
via similar mechanisms to those described for the proteasome (Kubota et al., 2010). Thus, these
studies indicate that, no matter what the mechanism of insult, mitochondrial dysfunction occurs
71
early in the cell death process, indicating that targeting the mitochondria may be an effective
neuroprotective strategy against all causes of cell death linked with PD.
As mentioned above, previous in vitro studies in cell models of PD have shown a tight link
between mitochondrial and UPS function (Shamoto-Nagai et al., 2003; Sullivan et al., 2004;
Radke et al., 2008). It has also been shown previously that mitochondrial abnormalities result in
lysosomal dysfunction, suggesting that there is also a link between mitochondrial and lysosomal
integrity (Irrcher et al., 2010). The current study is the first to demonstrate the link between
UPS, lysosomal and mitochondrial abnormalities very early in the cell stress pathway. It is
known that lysosomes and the UPS work together to regulate protein degradation within the cell,
the present study also suggests that only after longer periods of impaired lysosomal function does
the UPS become impaired, and vice versa. This delayed impairment of the UPS or lysosome
following prolonged inhibition of lysosome or UPS respectively, is probably due to a build up of
aggregated proteins, as indicated by the increase in ubiquitin levels as early as 3 hours following
exposure to naphthazarin or PSI in the present study.
In conclusion, these studies show that with respect to mechanisms of cellular dysfunction linked
with PD, cells are most sensitive to mitochondrial dysfunction, since any form of cell stress,
whether caused by inhibition of mitochondria, proteasome or lysosome results in mitochondrial
dysfunction early in the cell death pathway. Finally, our findings suggest that protecting
mitochondrial function early in the disease process i.e., upon initial diagnosis could potentially
be a disease modifying strategy for preventing the cell death that characterises later stages of PD.
The assays utilised in these studies may prove useful for testing potential neuroprotective
treatments in the future.
72
Chapter 3
Development and Validation of a Screening Assay for the Evaluation of Putative
Neuroprotective Agents in the Treatment of Parkinson’s Disease
A modified version of this chapter was previously published as:
Yong-Kee CJ, Salomonczyk D, Nash JE (2011) Development and validation of a screening assay for the evaluation of putative neuroprotective agents in the treatment of
Parkinson's disease. Neurotoxicity research 19:519-526.
73
3 Development and Validation of a Screening Assay for the Evaluation of Putative Neuroprotective Agents in the Treatment of Parkinson’s Disease
3.1 Introduction
In order to prevent neurodegeneration, it is necessary to understand how cells die. Genetic and
post mortem studies suggest that inhibition of mitochondrial complex 1, aberrant protein
degradation caused by dysfunction of the UPS and lysosomes, as well as oxidative stress from
dopamine metabolism in neurons of the SNc contribute to neurodegeneration in Parkinson’s
disease (Ardely et al., 2004; Di Fonzo et al., 2007; Kitada et al., 1998; Ramirez et al., 2006).
Both the lysosome and UPS are required for the degradation of damaged proteins. Ubiquitin
motifs are attached to proteins by ubiquitin ligases, which target the protein to proteasomes,
while lysosomes employ chaperone-mediated autophagy for protein degradation. Abnormal
lysosome and UPS function leads to protein accumulation into toxic aggregates that eventually
cause cell death (Betarbet et al., 2005). Presently, there is no evidence that mutations in genes
involved in dopamine metabolism cause PD, however, in any neuron, dopamine metabolism is
inherently toxic, causing oxidative stress, due to the production of semiquinones, and other free
radicals. It is not known why dopaminergic neurons of the SNc are more susceptible to
oxidative stress than any other dopaminergic neuron however (Jenner, 2003).
It is probable that in most parkinsonian patients, a combination of some or all of the above
pathological processes contribute to cell death (Dagda et al., 2008; Canu et al., 2000; Hoglinger
et al., 2003; Pandey et al., 2007). Many animal and cell culture models of PD have been
developed to further understand how these cell death mechanisms interact (Hanrott et al., 2006;
74
Panov et al., 2005) however, it is difficult to develop a model that takes into account the
numerous cell death mechanisms that are likely to cause idiopathic PD. Furthermore, such
models take months or years to develop. Unfortunately, without such a model, it is extremely
difficult to accurately predict the true potential of a putative disease modifying agent for the
treatment of PD.
Herein I hypothesized that a direct correlation between the assay used here, and the assays
conducted by NINDS can validate the effectiveness of this assay to test potential neurprotective
agents. The studies described in this chapter outline the development of an in vitro assay, which
involved exposure of catecholaminergic neuroblastoma cells (SH-SY5Y) to toxins that mimic
what are currently believed to be the main causes of neurodegeneration in PD. To assess the
validity of this assay, we tested five compounds that were previously chosen by the NINDS
sponsored CINAPS to move forward into clinical trials (Ravina et al., 2003; Heemskerk et al.,
2002). The results of this study directly correlate with the success of putative neuroprotective
agents in clinical trials. Therefore, it appears that this cell assay is a rapid and accurate predictor
of the usefulness of potential disease modifying agents for halting the progression of PD. We
propose that this assay will be a useful predictor of potential neuroprotective agents which may
be successful in clinical trials.
3.2 Materials and Methods
3.2.1 Materials
Z-Ile-Glu(OBut)-Ala-Leu-H (PSI) was purchased from BIOMOL (USA), alamar blueTM was
purchased from Biosource (Canada) and propidium iodide from Invitrogen (USA). Dopamine
The pOCT-dsRed2 and pCMV-mitoGFP (kindly donated by Dr. Heidi McBride and Dr. Linda
Mills, respectively) constructs have CMV promoters which drive the expression of the
mitochondrially targeted fluorescent proteins. The pcDNA3.1-3xFLAG-LRRK2 and pcDNA3.1-
3xFLAG-G2019S-LRRK2 (kindly donated by Dr. Christopher Ross) constructs contain a CMV
promoter which drives the expression of the transgene. The 3xFLAG sequence is upstream of
the LRRK2/G2019S sequence. pcDNA3.1-3xFLAG-LRRK2 and pcDNA3.1-3xFLAG-G2019S-
LRRK2 have the same sequence except GGC was replaced by AGC at amino acid 2019. The
pcDNA3.1 empty vector was generated from the excision of the 3xFLAG-LRRK2 sequence at
Xho1 sites. The ptracer-3xFLAG-LRRK2-mCherry and ptracer-3xFLAG-G2019S-LRRK2-
mCherry constructs were generated from pcDNA3.1-3xFLAG-LRRK2, pcDNA3.1-3xFLAG-
G2019S-LRRK2, ptracer-CMV2, and pmCherry-C1 (Clontech, USA). The mCherry sequence
was excised, blunted and inserted into the Pml1 sites of ptracer-CMV2 so that expression of
mCherry was driven by the EF1 promoter. The LRRK2 or G2019S-LRRK2 sequence was then
ligated into the Kpn1 and Not1 sites of ptracer-CMV2-mCherry. LRRK2 and G2019S-LRRK2
was positioned upstream of the mCherry sequence and its expression was regulated by the
CMV2 promoter.
4.2.2 Cell culture
Human dopaminergic neuroblastoma SH-SY5Y cells (ATCC, USA) (P10-P25) were stored in
liquid nitrogen (-200 oC), rapidly thawed and grown in DMEM (consisting of 6400 mg/L NaCl,
3700 mg/L NaHCO3, 400 mg/L KCl and 584 mg/L L-glutamine) supplemented with 5% bovine
100
calf serum in a sterile humidified chamber (37 oC, 5% CO2, 95% O2) (Incubator: MCO-20AIC,
Sanyo, USA) until confluent. Cells were passaged using 0.1% trypsin for 5 min, then pelleted by
centrifuging at 340xg for 5 min (Allegra 6R Centrifuge, Beckman Coulter, USA).
4.2.3 SDS-PAGE followed by Western blotting
SH-SY5Y cells were transfected with pcDNA3.1-3xFLAG-LRRK2 or pcDNA3.1-3xFLAG-
G2019S-LRRK2 (5µg). After 48 hrs, cells were washed twice with ice-cold PBSx1 and scraped
from 100 mm plates with Laemmli buffer. Samples were lysed for 15 minutes in water (95 oC),
and protein concentrations determined using the Bradford method. Protein samples (10-20µg)
were loaded onto gels, and SDS-PAGE followed by Western blotting was carried out.
Nitrocellulose blots were blocked in 5% non-fat powdered milk (60 min) and incubated with
antibodies against β-actin (1:1000 µl) and FLAG (1:1000 µl) (both Sigma-Aldrich, Canada)
overnight (4oC). Following 5 washes in 0.5% Triton-X in PBSx1, mouse (1:5000 µl) HRP-
conjugated secondary antibodies (Jackson Immunoresearch, USA) were incubated at room
temperature (60 min) in a 1% non-fat powdered milk/TTBS (0.1% Tween-20) solution. Protein
levels were detected using enhanced chemiluminescence (ECL Western Blotting Substrate,
Pierce), then imaged (SRX-101A, Konica Minolta). Three replicates were produced for each
condition.
4.2.4 Expression profile of constructs
SH-SY5Y cells were transfected with pcDNA3.1, pcDNA3.1-3xFLAG-LRRK2 or pcDNA3.1-
3xFLAG-G2019S-LRRK2 (all 8.6 µg) 24 hours after plating. Cells were fixed 8, 12, 24, 48 and
72 hours after transfection using 4% PFA, permeabilized with 0.1% Trion-X plus 100 mM
glycine, and blocked non-specific proteins with 5% bovine calf serum. Mouse anti-FLAG
(Sigma-Aldrich, Canada) was incubated with cells overnight at 4oC, which was then probed with
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anti-mouse cy2 (Jackson Immunoresearch, USA). In a separate experiment, SH-SY5Y cells
were transfected with ptracer-mCherry, ptracer-3xFLAG-LRRK2-mCherry or ptracer-3xFLAG-
G2019S-LRRK2-mCherry (all 5 µg) 24 hours after plating. Images were obtained 8, 12, 24, 48
and 72 hours after transfection. Images were captured using an epifluorescent microscope with
attached Apotome grid with 40x oil immersion objective (Axiovert 200, Carl Zeiss, Canada) for
fixed samples or with a 20x long distance objective for live imaging. One hundred cells were
counted for each of four replicates that were positively stained. Data are expressed as mean
number of positive cells ± SEM.
4.2.5 Mitochondrial morphology assay
SH-SY5Y cells were plated on glass coverslips in a 6 well dish 24 hours prior to transfection.
Cells were transiently transfected with pmCherry (0.768 µg), pCMV-mtGFP (1.86 µg) and
pcDNA3.1-3xFLAG-LRRK2 or pcDNA3.1-3xFLAG-G2019S-LRRK2 (both 8.6 µg). After 48
hours, cells were fixed with 4% parformaldehyde, permeabilized with 0.1% Triton-X plus 100
mM glycine and incubated with 5% bovine calf serum for 1 hour. In order to probe for 3xFLAG,
anti-mouse FLAG primary antibody was incubated at 4 oC overnight, followed by incubation
with anti-mouse cy5 secondary antibody (Jackson Immunoresearch, USA). Cells were imaged
using an epifluorescent microscope equipped with an Apotome grid with a 63x objective. To
determine the relative amounts of mitochondria that were fragmented, elongated or clustered,
one hundred cells from four independent experiments were analyzed in a blinded fashion.
Mitochondria were categorized as fragmented, elongated or clustered if 50% of the total
mitochondrial population in a cell displayed one morphology type. Data are expressed as mean
number of cells having a particular mitochondrial morphology ± SEM.
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4.2.6 Tetramethylrhodamine ethyl ester perchlorate (TMRE) assay SH-SY5Y cells were cultured on glass coverslips in a 6 well dish 24 hours prior to transfection
with pTag-BFP-C (0.78 µg) (Evrogen, USA), pcmv-mtGFP (1.86 µg) and pcDNA3.1,
pCDNA3.1-3xFLAG-LRRK2 or pCDNA3.1-3xFLAG-G2019S-LRRK2 (all 8.6 µg). On the
second day after transfection, cells were labelled with 20 nM TMRE (Invitrogen, USA) for 20
min in DMEM. TMRE is a lipophillic cationic dye that binds to the inner and outer portions of
the mitochondrial inner membrane upon proportion of membrane potential (Scaduto and
Grotyohann, 1999). Fluorescence intensity is reduced as a consequence of depolarisation in
membrane potential. Coverslips were placed in a live imaging chamber with imaging buffer
(116 mM NaCl, 5.4 mM KCl, 0.4 mM MgSO4, 20 mM HEPES, 0.9 mM Na2HPO4, 1.2 mM
CaCl2, 10 mM glucose, 5 mM pyruvate, pH 7.4) and imaged on an epifluorescent microscope
with Apotome with a 40x oil immersion objective. The fluorescence intensity of 30-50
randomly chosen cells from each of four independent experiments was measured using ZEN
2009 (Carl Zeiss, Canada). Then the average fluorescence intensity of all four independent
experiments was determined. Data are expressed as mean TMRE intensity ± SEM.
Figure 5.12. Effect of toxin exposure on TH expression in nigro-striatal organotypic co-
cultures. Co-cultures (D24) were incubated with toxins 24 hours prior to being fixed and
immunolabelled with antibodies against GIRK2 and TH. Images show the effect of naphthazarin
(80 µM), PSI (100 µM) and rotenone (70 µM) on TH expression in GIRK2 expressing cells.
Scale bars = 20 µm.
176
DMSO
DAPIGIRK2 TH MergeNAP
PSI
ROT
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5.4 Discussion
Familial and sporadic PD are likely to have similar mechanisms underlying their pathology,
however a treatment that halts, slows or reverses PD has yet to be developed. To this end, I
developed and characterized cell models of PD that enable more rapid screening of potential
neuroprotective compounds in a relatively high throughput and cost effective manner. Whilst
these models are less physiological than in vivo models, I have shown that they are likely to have
clinical relevance.
Previously I validated an SH-SY5Y cell model of PD which proved to be effective in testing
potential neuroprotective agents (Yong-Kee et al., 2011b). Here, I utilized this same model to
test the neuroprotective effect of RGMa. Interestingly, RGMa was protective during dopamine
and rotenone toxicity, both highly relevant in PD pathology. Given that only rotenone and
dopamine toxicity was blocked, it is likely that RGMa protects cells by preventing ROS
generation. Here, the greatest protective effect of RGMa was during rotenone toxicity. Since
mitochondria are highly active organelles that generate large amounts of ROS, inhibition of
mitochondrial complex 1 would lead to a large rapid increase in ROS (Lin and Beal, 2006).
Preventing this increase may explain the higher protective effect seen during rotenone toxicity
than in dopamine toxicity. Studies revealed an interaction between RGMa and neogenin
receptors prevents the activation of proapoptotic pathways (Matsunaga and Chedotal, 2004). It
could be possible that RGMa blocks apoptosis given that toxic effects of dopamine and rotenone
are mediated by apoptotic processes (Hanrott et al., 2006; Imamura et al., 2006). However, a
study using neural precursor cells revealed the activation of apoptosis in response to lysosome
inhibition (Walls et al., 2010). Thus, if RGMa blocked apoptotic processes, it would be expected
that it would be effective during naphthazarin toxicity as well.
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The role of SIRT3 in mitochondria and cell survival made it a potential candidate for a
neuroprotective agent in PD. The effectiveness of SIRT3 as a neuroprotective agent was tested
in various in vitro models of PD. SIRT3 protected neuroblastoma and neurons from rotenone
toxicity. A study by Ahn et al. (2008) revealed the deacetylase activity of SIRT3 can enhance
the activity of mitochondrial complex 1. This in turn regulates the production of ATP levels in
the cell. Since mitochondrial activity at complex 1 is reduced by rotenone, it is feasible that the
actions of SIRT3 can reverse these effects on mitochondria. This explains how effective SIRT3
is at negating the effects of rotenone and not any of the other toxins tested. An alternative
function of SIRT3 can warrant the protective effect of SIRT3. A study in myotubes revealed the
SIRT3 dependent up-regulation of PGC1α leads to mitochondria biogenesis (Kong et al., 2010).
An up-regulation of mitochondria may compensate for damaged mitochondria which ultimately
prevents cell loss. These results prompted the examination of the effectiveness of SIRT3 during
LRRK2 induced toxicity. Given the effects of LRRK2 on mitochondria, SIRT3 would be a
prime candidate for neuroprotection in PD (Wang et al., 2012b). Indeed, SIRT3 prevented
LRRK2 induced cell death in differentiated SH-SY5Y cells. This necessitates further study of
SIRT3 to protect cells from dysfunction associated with mutant LRRK2. SIRT3 is effective at
protecting mitochondria which infers that it may be a valuable tool for other mitochondrial
disorders.
In vitro models utilizing cell lines are useful for high throughput experiments where multiple
compounds can be tested, but they lack physiological elements that are essential for more
rigorous testing of compounds. Thus, the more complex in vitro models, such as the organotypic
and primary cultures were characterized for various neuronal markers. Both the organotypic and
primary cultures used here express TH, which is a key component in the production of
dopamine. Other studies in ventral mesencephalic and organotypic cultures have shown the
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expression of TH and homovanillic acid, proteins involved in dopamine production (Lyng et al.,
2007; Zhang et al., 2007). Furthermore, these cultures are composed of neurons from the SNc,
as shown by prominent GIRK2 expression. Neurons that degenerate in the early stages of PD
are specific to the SNc and have a dopamine producing phenotype (Hornykiewicz, 1966; Obeso
et al., 2000). These findings make them an appropriate model, from a physiological standpoint,
to study PD. In vivo models benefit from the retention of synaptic networks when compared to
in vitro models. Indeed, the cultures used here form synaptic connections and have key
components of synapse formation. This is demonstrated by the expression of the synaptic
proteins PSD95 and synaptophysin. This model is in agreement with other studies that found
PSD95 and synaptophysin expression in organotypic and primary cultures (Buckby et al., 2004;
Wakita et al., 2010). The models studied here display characteristics similar to those of in vivo
models and benefit from relatively faster production. This in turn enables testing of various
compounds in an environment that is more physiological than those provided by cell lines, while
retaining easier production than animal models.
Toxin based in vitro and in vivo models have been used as tools to mimic the pathology seen in
sporadic PD. Most in vitro models lack the physiological environment that resembles PD, while
in vivo models are inconsistent at obtaining a complete loss of dopaminergic neurons (Betarbet et
al., 2000; Hoglinger et al., 2003b). It is apparent here that a variety of toxins produced a loss of
cells at specific concentrations in organotypic cultures. Cell death induced by naphthazarin, PSI
and rotenone is associated with a loss of TH specifically in SNc neurons. A similar study in
ventral mesencephalic cultures shows a loss of TH after MPTP exposure (Jakobsen et al., 2005).
Interestingly, rat models which fail to produce behavioural changes similar to those found in PD,
do not show a reduction in TH because these behavioural changes are associated with losses of
dopamine (Lapointe et al., 2004). A retraction of SNc axons and terminals occurs in patients, but
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the toxins used here do not lead to similar morphological changes. This could be a consequence
of the time cultures were exposed to the toxins, thus extended exposure times may allow for a
loss of axons. Indeed, multiple studies have shown axon degeneration in organotypic cultures
using rotenone incubation periods from three days to more than a week (Testa et al., 2005;
Ullrich and Humpel, 2009). This organotypic co-culture model is advantageous for studying
mechanisms related to PD because it employs toxins linked with PD pathology to consistently
produce a loss of dopaminergic neurons.
Current transgenic models suffer from the inability to consistently produce dopaminergic
degeneration (Meredith et al., 2008). Furthermore the only transgenic animals expressing α-
synuclein are mice; which makes it extremely difficult to dissect the nigra. Thus, I generated an
in vitro primary culture model that expresses α-synuclein. The α-synuclein primary culture
model used here repeatedly shows degeneration of GIRK2 expressing cells. This specificity is
important because it replicates the early stages of disease pathology, whereby nigral cells begin
to degenerate (Hornykiewicz, 1966; Obeso et al., 2000). Lewy bodies, which are composed of
α-synuclein and ubiquitin, do not form in these neurons. Lewy bodies are often associated with
PD linked with α-synuclein (Crowther et al., 2000; Saito et al., 2004). It is likely that increased
culture periods would allow for the formation of Lewy bodies because α-synuclein would have
more time to accumulate and aggregate. Since this model does not require transgenic animals, it
is a relatively cost-effective way to study pathology linked with PD.
These studies reveal that in vitro models are a valuable tool for studying PD because they are
easily produced and are similar to cells affected in PD. Furthermore, they display typical losses
of function and viability when presented with toxin and gene induced cell stressors. Granted
there are disadvantages to these models, their advantages are highly beneficial when used in a
181
high throughput environment. This is demonstrated by the elucidation of the protective effect of
RGMa and SIRT3. The rescue of cells from mitochondrial dysfunction caused by toxins and
genetic over-expression implies that PD could be a mitochondrial disorder. Furthermore, RGMa
and SIRT3 may be important agents for treating other mitochondrial disorders.
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Chapter 6
Summary and Future Directions
183
6 Summary and Future Directions
To conclude, following exposure to toxins that mimic PD pathology, UPS and lysosome function
are inhibited, consequently reducing mitochondrial function. Following genetic over-expression
of LRRK2 and G2019S-LRRK2, mitochondrial fusion becomes inhibited, resulting in
fragmentation of mitochondria. Compounds that primarily target mitochondrial function in
models that both mimic PD pathology and contain neurons that are similar to those which
degenerate in PD are essential for developing therapeutics to treat PD. The findings described
here suggest that alterations to mitochondria may be a central factor contributing to PD onset,
and that enhancing the function of mitochondria is a potential avenue for preventing
neurodegeneration causing PD.
Recently it has been shown that ROS production, as well as malfunctions to the UPS and
lysosome are causative factors of the disease (Cuervo et al., 2004; Betarbet et al., 2005). In
Chapter Two, studies reveal that malfunction of multiple sub-cellular systems inhibit
mitochondrial function early on during cell stress. More specifically, these experiments reveal a
tight link between the UPS, lysosomes and mitochondria, and that convergence of these
dysfunctional processes can be disastrous for cellular health. These experiments not only reveal
the convergence of multiple mechanisms, but they also present evidence that mitochondria are
sensitive to abnormalities in other sub-cellular systems, such as the UPS and lysosome. These
studies are the first to show a link between the mitochondria, UPS and lysosome. Moreover, the
link between the UPS and lysosome is further established here since prolonged inhibition of the
UPS or lysosome can impair the lysosome or UPS, respectively (Sherer et al., 2002). In future,
studies should examine the pathways leading to the interactions between the mitochondria, UPS
184
and lysosome. The regulation of mitochondrial protein quality control by the UPS and the
removal of dysfunctional mitochondria by lysosomes are possible pathways interconnecting
these sub-cellular domains. These pathways can be studied by measuring the accumulation of
ubiquitnated proteins in mitochondrial fractions from cell lysates during proteasomal inhibition
or by measuring the number of mitochondria encapsulated by autophagosomes during lysosomal
inhibition. However, it is still unclear why mitochondria are specifically affected and why
mitochondria are affected so early during cellular stress when processes such as fission and
fusion are in place to rescue mitochondria. Since mitochondria provide energy to the cell, it is
possible that slight perturbations to mitochondrial function can have profound effects on neurons
that have high energy demands. Moreover, nigral cells are highly susceptible to changes in
homeostasis because they are constantly metabolizing dopamine, which produces large quantities
of ROS, thus, further production of ROS by mitochondrial damage may bring imbalance to
nigral cell homeostasis. This would also account for nigral neurons being unable to quickly
compensate for losses of mitochondrial function and ROS production. Thus, future studies
should compare the function of mitochondria and their effects on ROS in nigral and non-nigral
neurons to determine if nigral cells are more susceptible to these changes. It is clear from these
studies that mitochondria play an important role in disease pathogenesis.
Since multiple sub-cellular mechanisms are responsible for disease pathogenesis, it is necessary
for a model to incorporate all of these mechanisms. Moreover, such a model can be deemed
representative of the condition, if when tested against a battery of potential neuroprotective
agents, has a high degree of similarity when correlated to the outcomes of clinical trials. Indeed,
the outcomes of the neuroprotective assay used in Chapter Three are very similar to the results of
clinical trials where various neuroprotective agents were tested. In particular, coenzyme Q10
was highly protective during stress induced by multiple mechanisms, such as mitochondrial, UPS
185
and lysosomal inhibition. A correlation between this study and clinical trials can be drawn since
coenzyme Q10 showed promise in clinical trials, thus validating the in vitro assay used here.
Substantial evidence for a causative role of mitochondria in PD pathology is provided by the
neuroprotective effect of coenzyme Q10. Since mitochondrial dysfunction results from UPS and
lysosomal inhibition, it is expected that a potential therapeutic which acts directly on
mitochondria would be highly protective, and indeed, coenzyme Q10 did protect cells from the
toxic effects of UPS and lysosomal inhibition. Coenzyme Q10 is an integral part of the electron
transport chain, and the protective effects seen here could be attributed to an increase in ATP
production. However, increased ROS production associated with inhibited respiratory activity
could be prevented when coenzyme Q10 restores oxidative phosphorylation. Thus, future
studies can examine if the aforementioned pathways are affected by blocking key molecules
essential for each pathway in the presence of coenzyme Q10 and then determine if cell viability
is enhanced. Alternatively, ATP and ROS levels can be measured when coenzyme Q10 is
present. These studies are the first to describe an in vitro model that is easy to produce and can
effectively test potential neuroprotective agents to treat PD. Moreover, it will enable rapid and
efficient testing of a wide range of compounds and make available more compounds for testing
in clinical trials. These studies suggest that for a therapeutic to have any potential to treat PD
patients it must enhance mitochondrial function or induce the biogenesis of mitochondria to
compensate for mitochondria that have lost function due to cell stress.
Recently research has redirected its focus back to the field of mitochondria by suggesting
mitochondrial dynamics is an important mechanism in neurodegenerative disease (Chen and
Chan, 2009). Thus, given the findings of my initial study, subsequent studies in my thesis
focused on elucidating the role of mitochondrial dynamics in PD. Mitochondrial dysfunction in
PD may arise from alterations to fission and fusion since these processes are imperative for
186
proper mitochondrial function (Otera and Mihara, 2011). This hypothesis is confirmed in
Chapter Four by finding that mitochondrial fusion becomes inhibited by LRRK2 and G2019S-
LRRK2 expression which ultimately fragments mitochondria. Furthermore, it appears that
mitochondrial fragmentation occurs before cell stress occurs. Although mitochondrial
dysfunction was not shown during LRRK2 and G2019S-LRRK2 expression, it may be that
altered fusion is the first step leading to reduced membrane potential, and hence, dysfunctional
mitochondria. Indeed, mitochondrial fragmentation is a prerequisite for mitochondrial
dysfunction and eventual cell death (Frank et al., 2001; Breckenridge et al., 2003). While
increased mitochondrial fission could lead to fragmented mitochondria, it was not observed in
this study. This result however, may be specific to the time point used. At later time points
fission may indeed play a role in the fragmentation caused by LRRK2 and G2019S-LRRK2.
Given the discovery of altered fusion, future studies should focus on elucidating the changes that
occur in fusion proteins. It is possible that changes in the activity of Mfn1, Mfn2 and Opa1 are
regulated by the kinase function of LRRK2. Moreover, the GTPase activity of LRRK2 could
limit the available GTP required for fusion protein function since LRRK2, which is located on
the outer mitochondrial membrane, would be situated close to Mfn1 and Mfn2. Future studies
can delineate if the kinase or GTPase domain of LRRK2 is responsible for the inhibition of
fusion protein activity by knocking down these specific domains of LRRK2 and subsequently
measuring fusion rates. While studies suggest mitochondrial fission is key to abnormal
mitochondrial function, this study is the first to suggest that inhibition of mitochondrial fusion by
LRRK2 could affect mitochondrial function. It is possible that patients having a LRRK2 genetic
background may suffer neurodegeneration caused by abnormal mitochondrial fusion.
It is imperative for in vitro models to replicate the pathology found in PD as precisely as possible
so as to foster the efficient production of therapeutics. Various studies, including the ones
187
described here, suggest compounds which target the mitochondria are highly effective at treating
PD. In Chapter Five, upon characterization of various in vitro models, neuronal cultures were
found to be very similar to the neurons that degenerate in the SNc of parkinsonian patients.
Moreover, the novel mitochondrial protein, SIRT3, protected neurons from degeneration
associated with cell stress induced by LRRK2 over-expression. The high therapeutic value of
SIRT3 can be concluded for two reasons. Firstly, the feasibility of SIRT3 as a neuroprotective
agent is confirmed since it was able to protect mature neurons having a dopaminergic phenotype
from degeneration. Secondly, since SIRT3 protected neurons from the toxic effects of LRRK2, a
genetic mutation that is common among PD patients, it has the potential to treat a broad
spectrum of patients with PD. The next logical study is to test SIRT3 in various animal models
that closely mimic the pathological mechanisms underlying PD. In addition to mitochondrial
dysfunction, these models should display ROS production and protein aggregation as these are
classical markers of PD pathology. Further studies should be conducted to examine the
mechanisms by which SIRT3 provides neuroprotection. Indeed two main roles of SIRT3 are
likely responsible for its therapeutic effect, one being the regulation of mitochondrial complex 1
function, and two being its role in mitochondrial biogenesis (Ahn et al., 2008; Cimen et al.,
2010; Kong et al., 2010). These pathways can be studied by measuring changes in ATP levels or
by quantifying levels of mitochondrial DNA during SIRT3 expression. These studies are the
first to show the protective effect of SIRT3 in a neuronal model that has a high degree of
similarity to the neurons that degenerate in the SNc. Furthermore, SIRT3 could have a broad use
in the clinic since it could be used to treat patients having either familial PD caused by LRRK2,
or idiopathic forms caused by environmental toxins.
188
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8 Appendices
A modified version of this appendix was previously published as:
Yong-Kee CJ, Warre R, Monnier PP, Lozano AM, Nash JE (2012) Evidence for synergism between cell death mechanisms in a cellular model of neurodegeneration in
Parkinson's disease. Neurotoxicity research 22(4):355-364.
My contributions to the following appendix include data analyses and design of the
figures listed in the text.
Abstract
Delineation of how cell death mechanisms associated with PD interact and whether they
converge would help identify targets for neuroprotective therapies. The purpose of this study
was to use a cellular model to address these issues. Catecholaminergic SH-SY5Y neuroblastoma
cells were exposed to a range of compounds (dopamine, rotenone, naphthazarin, and PSI) that
are neurotoxic when applied to these cells for extended periods of times at specific
concentrations. At the concentrations used, these compounds cause cellular stress via
mechanisms that mimic those associated with causing neurodegeneration in PD, namely
Characterisation of interactions between cell death mechanisms
Concentration response curves were performed to assess cell viability in response to a dose
response curve of each compound (dopamine, rotenone, PSI, naphthazarin), either alone or in the
presence of an EC10–20% concentration of a second compound. The decrease in the percentage of
viable cells from 100% in the presence of a compound alone (compound 1) was compared with
the decrease in the percentage of viable cells from 100% in the presence of compound 1 plus an
approximate EC10–20% concentration of compound 2. As described above, the EC10-20%
concentrations used were 10 µM dopamine (EC20%), 100 µM rotenone (EC10%), 50 µM PSI
(EC10%), and 3 µM naphthazarin (EC15%).
As an EC20% of compound 2 should produce an additional decrease in cell viability of 20% if the
relationship is simply additive, synergism between the compounds was identified as an
additional decrease in the percentage of viable cells of greater than 25% in the presence of
compounds 1 and 2 in comparison with compound 1 alone for at least two points within the first
five on the concentration response curve. Similarly, where an EC10% or EC15% of compound 2
was added in combination with compound 1, an additive effect would be expected to decrease
cell viability by 10% and 15%, respectively. In these cases, synergism was defined as an
additional decrease in cell viability of 12.5% and 18.75%, respectively, for at least two points on
the concentration response curve for compounds 1 and 2 when compared with compound 1
alone.
Synergism is indicated in Figures 8.2 to 8.5 by diagonal shading. The occurrence of synergism
suggests that the two cell death mechanisms activated by the compounds are part of the same
pathway. An additive effect was defined when the decrease in the percentage of viable cells in
222
the presence of compound 1 and 2 was 20%, 15%, 10% or less depending on the identity of
compound 2, indicating that the compounds cause cell death by independent pathways. In some
cases the combination of compound 1 and 2 produced no additional decrease in cell viability
over compound 1 indicating that the effects of compound 1 occlude the effects of compound 2.
Effect of rotenone, PSI, or naphthazarin on dopamine-induced toxicity
SH-SY5Y cells were exposed to a range of dopamine concentrations (0.01 µM–600 µM) either
alone, or in the presence of an EC10% of rotenone, an EC10% of PSI, or an EC15% of naphthazarin.
Incubation with dopamine alone caused a concentration-dependent decrease in cell viability (Fig.
8.2). The combination of rotenone (100 µM) with dopamine caused a significant additional
decrease in the percentage of viable cells of 25.7%, 27.1%, and 26.2% for 0.1, 1, and 10 µM
dopamine (P < 0.01), respectively, indicating that dopamine and rotenone work synergistically
(Fig 8.2a). Similarly, the addition of PSI (50 µM) to the cells in the presence of dopamine also
caused a synergistic decrease in cell viability, which ranged from 19.0% to 32.6%, for all
concentrations of dopamine between 0.1 µM and 300 µM (P < 0.001) (Fig. 8.2b). In contrast, the
addition of naphthazarin (3 µM) to the dopamine dose response curve only caused an additive
effect with a significant additional decrease in the cell viability of 24.6% only observed for 1 µM
dopamine. For the remaining concentrations of dopamine in the concentration-response curve
the addition of naphthazarin caused additional decreases in the percentage of viable cells of
11.5% to 18.5% (Fig. 8.2c).
223
Figure 8.2. Effect of rotenone, PSI, or naphthazarin on dopamine-induced toxicity. SH-
SY5Y cells were incubated with dopamine (DA, 0.01 µM–600 µM), either alone, or in the
presence of approximate EC10–20% concentrations of (a) rotenone (ROT, 100 µM ≈ EC10%), (b)
PSI (50 µM ≈ EC10%), or (c) naphthazarin (NAP, 3 µM ≈ EC15%) for 24 hours. Cell viability (%
control) was assayed using alamar blue and concentration-response curves were plotted (mean ±
SEM). Diagonal shading indicates a difference of greater than 25% between the two curves for
dopamine and rotenone (a) and a difference of greater than 12.5% between the two curves for
dopamine and PSI (b), which indicates the presence of synergy between the pairs of compounds.
The effect of naphthazarin in combination with dopamine was additive (c). Differences between
the curves were analyzed using two-way ANOVA with Bonferroni’s multiple comparison test
post-hoc. * indicates P < 0.05, ** P < 0.01, and *** P < 0.001. Overall effect of rotenone
treatment: F1,70 = 28.11, P < 0.0001, PSI treatment: F1,70 = 123.9, P < 0.0001, and napthazarin
treatment: F1,70 = 18.20, P < 0.0001.
224
0.01 0.1 1 10 100 10000
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Effect of dopamine, PSI, or naphthazarin on rotenone-induced
mitochondrial dysfunction
SH-SY5Y cells were exposed to a range of rotenone concentrations (0.01 µM–1000µM) either
alone, or in the presence of an approximate EC20% of dopamine, an EC10% of PSI, or an EC15% of
naphthazarin. Incubation with rotenone alone caused a concentration-dependent decrease in cell
viability (Fig. 8.3). The addition of dopamine (10 µM) to the rotenone concentration-response
curve caused a significant additional decrease in cell viability at all concentrations of rotenone
between 0.01 µM and 100 µM (P < 0.01). The additional decrease in the percentage of viable
cells ranged from 24.8% to 34.0% for the concentrations of rotenone between 0.01 µM and 100
µM indicating that the effects of dopamine and rotenone were synergistic (Fig. 8.3a). The
addition of PSI (50 µM) (Fig. 8.3b) or naphthazarin (3 µM) (Fig. 8.3c) to the rotenone
concentration-response curve both produced a synergistic effect at 1 µM and 10 µM rotenone
producing a decrease in cell viability of 18.5% and 15.8%, respectively with PSI, and 20.3% and
22.6%, respectively with naphthazarin (P < 0.01) (Fig. 8.3 b,c).
226
Figure 8.3. Effect of dopamine, PSI, or naphthazarin on rotenone-induced mitochondrial
dysfunction. SH-SY5Y cells were incubated with rotenone (ROT, 0.01 µM–1000 µM), either
alone, or in the presence of approximate EC10–20% concentrations of (a) dopamine (DA, 10 µM ≈
EC20%), (b) PSI (50 µM ≈ EC10%), or (c) naphthazarin (NAP, 3 µM ≈ EC15%) for 24 hours. Cell
viability (% control) was assayed using alamar blue and concentration-response curves were
plotted (mean ± SEM). Diagonal shading indicates a difference of greater than 25% between the
two curves for rotenone and dopamine (a), a difference of greater than 12.5% between the two
curves for rotenone and PSI (b), and a difference of greater than 18.75% between the two curves
for rotenone and naphthazarin (c), which indicates the presence of synergy between the pairs of
compounds. Differences between the curves were analyzed using two-way ANOVA with
Bonferroni’s multiple comparison test post-hoc. * indicates P < 0.05, ** P < 0.01, and *** P <
0.001. Overall effect of dopamine treatment: F1,70 = 84.01, P < 0.0001, PSI treatment: F1,70 =
16.41, P = 0.0001, and naphthazarin treatment: F1,70 = 37.89, P < 0.0001.
227
ba
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Effect of dopamine, rotenone, or naphthazarin on PSI-induced proteasome
dysfunction
SH-SY5Y cells were exposed to a range of PSI concentrations (0.1 µM–500 µM) in combination
with an approximate EC20% of dopamine, an EC10% of rotenone, or an EC15% of naphthazarin.
Incubation with PSI alone caused a concentration-dependent decrease in cell viability (Fig. 8.4).
Incubation of PSI with dopamine (10 µM) caused a significant additional decrease in cell
viability compared to PSI alone at all concentrations of PSI between 0.1 µM and 500 µM (P <
0.01 for 0.1 µM–10 µM, P < 0.05 for 50 µM and 150 µM, P > 0.05 for 500 µM). The additional
decrease in the percentage of viable cells ranged from 15.5% to 29.8% indicating that PSI and
dopamine act synergistically (Fig. 8.4a). The combination of PSI and rotenone (100 µM) also
produced a synergistic decrease in the percentage of viable cells that ranged from 19.1% to
25.0% for the concentrations of PSI between 0.1 µM and 150 µM (Fig. 8.4b). The addition of
naphthazarin (3 µM) to the PSI concentration-response curve produced additional decreases in
the percentage of viable cells of 16.7% to 19.6% between 0.1 µM and 10 µM PSI indicating that
the effects of napthazarin and PSI were additive, rather than synergistic as the additional
decrease in cell viability was not sustained at 18.75% or greater (Fig. 8.4c).
229
Figure 8.4. Effect of dopamine, rotenone, or naphthazarin on PSI-induced proteasomal
dysfunction. SH-SY5Y cells were incubated with PSI (PSI, 0.1 µM–500 µM), either alone, or
in the presence of approximate EC10–20% concentrations of (a) dopamine (DA, 10 µM ≈ EC20%),
(b) rotenone (ROT, 100 µM ≈ EC10%), or (c) naphthazarin (NAP, 3 µM ≈ EC15%) for 24 hours.
Cell viability (% control) was assayed using alamar blue and concentration-response curves were
plotted (mean ± SEM). Diagonal shading indicates a difference of greater than 25% between the
two curves for PSI and dopamine (a) and a difference of greater than 12.5% between the two
curves for PSI and rotenone (b), which indicates the presence of synergy between the pairs of
compounds. The effect of naphthazarin in combination with PSI was additive (c). Differences
between the curves were analyzed using two-way ANOVA with Bonferroni’s multiple
comparison test post-hoc. * indicates P < 0.05, ** P < 0.01, and *** P < 0.001. Overall effect of
dopamine treatment: F1,70 = 71.06, P < 0.0001, rotenone treatment: F1,70 = 31.73, P < 0.0001, and
naphthazarin treatment: F1,70 = 32.62, P < 0.0001.
230
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Effect of dopamine, rotenone, or PSI on naphthazarin-induced lysosome
dysfunction
SH-SY5Y cells were incubated with a range of naphthazarin concentrations (0.01 µM–1000
µM), either alone, or in combination with an approximate EC20% of dopamine, an EC10% of
rotenone, or an EC10% of PSI. Incubation of the cells with naphthazarin alone caused a
concentration-dependent decrease in cell viability (Fig. 8.5). The addition of dopamine (10 µM)
to the naphthazarin concentration-response curve caused a significant additional decrease in cell
viability at all concentrations of naphthazarin between 0.01 µM and 30 µM (P < 0.001). For 0.01
µM and 0.1 µM naphthazarin the addition of dopamine (10 µM) caused an additional decrease in
the percentage of viable cells of 49.8% and 25.6%, respectively, indicating that the two
compounds were acting synergistically. Although the difference between the concentrations
curves for 1 µM naphthazarin with and without dopamine (10 µM) was only 18.5%, an
additional decrease in the percentage of viable cells of 23.4% was observed for dopamine (10
µM) combined with 10 µM naphthazarin, which suggests that there is synergism between
naphthazarin and dopamine (Fig. 8.5a). The addition of either rotenone (100 µM) or PSI (50
µM) to the naphthazarin concentration-response curve had little effect on the naphthazarin-
induced decrease in cell viability indicating the effects of rotenone and PSI are occluded by the
presence of naphthazarin (Fig. 8.5b,c).
232
Figure 8.5. Effect of dopamine, rotenone, or PSI on naphthazarin-induced lysosomal
dysfunction. SH-SY5Y cells were incubated with naphthazarin (NAP, 0.01 µM–1000 µM),
either alone, or in the presence of approximate EC10–20% concentrations of (a) dopamine (DA, 10
µM ≈ EC20%), (b) rotenone (ROT, 100 µM ≈ EC10%), or (c) PSI (PSI, 50 µM ≈ EC10%) for 24
hours. Cell viability (% control) was assayed using alamar blue and concentration-response
curves were plotted (mean ± SEM). Diagonal shading indicates a difference of greater than 25%
between the two curves and the presence of synergy between naphthazarin and dopamine (a).
The effect of rotenone or PSI was occluded by the presence of naphthazarin, (b) and (c).
Differences between the curves were analyzed using two-way ANOVA with Bonferroni’s
multiple comparison test post-hoc. * indicates P < 0.05, ** P < 0.01, and *** P < 0.001. Overall
effect of dopamine treatment: F1,70 = 247.4, P < 0.0001, rotenone treatment: F1,70 = 2.377, P =
0.1276, and PSI treatment: F1,70 = 1.121, P = 0.2934.
233
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Discussion
Many studies have shown that dysfunction of the mitochondria, proteasome, and lysosome, as
well as increased oxidative stress and elevated protein aggregate formation contribute to cell
death in PD (Hoglinger et al. 2003; Rideout et al. 2001; Sherer et al. 2003). Furthermore, these
studies showed that some of these sub-cellular processes interact to accelerate
neurodegeneration. While previous studies have enabled a better understanding of the
mechanisms underlying neurodegeneration in PD, all the models used require lengthy
preparation times. In order to address this issue we developed a cell model that is simple and
quick to produce with characteristics that appear to mimic those in more sophisticated chronic
models of PD (Yong-Kee et al. 2011). However, due to the cellular nature of this model, caution
should be used in the extrapolation of results to processes underlying neurodegeneration in
patients with PD. Nonetheless, we predict that this simplified system is a useful method for
understanding interactions between intracellular mechanisms that are difficult to study in the
whole animal.
In this study we used our cell model to further characterise the interactions between the various
cell death mechanisms that destroy catecholaminergic neurons. In particular we were interested
in any potential synergistic relationships between different compounds that activate cell death
mechanisms associated with PD. As an EC10–20% of a second compound was added to a full
concentration response curve of the first compound, we defined an additive effect as an
additional decrease in cell viability of 10% to 20% or less, and synergism as an additional
decrease in cell viability of 12.5% to 25% or more for at least two points on the concentration
response curve, depending on the EC value of the second compound. According to this
definition, when combined the two compounds needed to produce an effect that was at least 25%
235
greater than a simple addition of the effects of the two compounds to be identified as synergistic.
The definition also required that the synergistic effect be present for at least two points within the
first five of the concentration curve. Thus, we were able to identify pairs of compounds where
there was synergism, and also whether this synergism was reciprocally mediated.
The results of this study indicate that oxidative stress has synergistic effects on all other cell
death mechanisms, causing a greater decrease in cell viability than would be expected from
combining the individual effects of activating each pathway. For mitochondrial and proteasomal
dysfunction the synergism with oxidative stress was reciprocal but the relationship between
lysosomal dysfunction and oxidative stress was only unidirectional (Fig. 8.6), with an EC20%
concentration of dopamine enhancing cellular impairment caused by naphthazarin, but not vice
versa. A possible explanation for this is that, as well as being involved in protein degradation,
the lysosome also accumulates free radicals, to reduce oxidative stress. Thus, when the
lysosome is inhibited, addition of even low levels of dopamine may have a more dramatic effect
because the cells have less ability to remove the quinones and reactive oxygen species generated
through dopamine metabolism, enhancing the sensitivity of the cell to dopamine. This finding
also suggests that oxidative stress may occur as a consequence of lysosomal dysfunction. The
uni-directionality of the effect indicates that lysosomal-independent oxidative stress through
addition of dopamine does not affect lysosome function. This lack of reciprocality is probably
due to the ‘all or nothing’ effect of naphthazarin on cell death, meaning that once the lysosome is
significantly damaged, the cell has probably reached a point of no return, with regards to loss of
cell viability, thus inhibition of proteasomal or mitochondrial function will have relatively little
impact. Thus, whilst there appears to be close relationships between mitochondrial and
proteasomal dysfunction and oxidative stress, decreased cell viability as a consequence of
lysosome dysfunction is a relatively independent cell death pathway.
236
Figure 8.6. Schematic diagram to show the interactions between the different mechanisms
in relation to changes in cell viability. Proteasomal, lysosomal, and mitochondrial dysfunction
form independent cell death pathways. Furthermore, proteasomal, lysosomal and mitochondria
dysfunction are exacerbated by oxidative stress. In addition there are reciprocal links between
oxidative stress, proteasomal dysfunction, and mitochondrial dysfunction.
237
Mitochondriadysfunction(rotenone)
Proteasome dysfunction
(PSI)
Oxidative stress
(dopamine)
Lysosomedysfunction
(naphthazarin)
238
Previous studies have shown that cell death caused by mitochondrial and proteasomal
dysfunction, as well as oxidative stress synergise to further enhance cell death (Hoglinger et al.
2003; Rideout et al. 2001; Sherer et al. 2003). The current studies take our understanding of cell
death mechanisms in PD further to show the relationship between these cell death mechanisms
and lysosomal dysfunction. Based on these studies, it appears that oxidative stress enhances
mitochondrial, lysosomal and proteasomal dysfunction in a synergistic manner. Furthermore, as
we have previously described, whether the site of the primary insult is the lysosome, proteasome,
mitochondria or increased oxidative stress, the mitochondria always become impaired (Yong-
Kee et al., 2012). Thus, all cell death mechanisms appear to converge on mitochondrial
dysfunction (Greenamyre and Hastings, 2004; Yong-Kee et al., 2012).
These studies suggest that an effective neuroprotective therapy in PD would need to target
multiple cell death pathways. Given that oxidative stress has a universal synergistic effect, and
that mitochondria are affected by all cell death mechanisms, perhaps decreasing oxidative stress
combined with enhancing mitochondrial health may prove to be the two most effective targets
for neuroprotective agents. Exposure of SH-SY5Y cells to a battery of compounds that
recapitulate the cell death mechanisms in PD, as was done here, in combination with potential
neuroprotective therapies may be a rapid and reliable method for screening such compounds.
239
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