LEVODOPA-INDUCED DYSKINESIA – FROM ABERRANT PRESYNAPTIC 

SIGNALING TO MALADAPTIVE POSTSYNAPTIC PLASTICITY 

 

By 

 

Rhyomi Charis Sellnow 

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

A DISSERTATION 

 

Submitted to 

Michigan State University 

in partial fulfillment of the requirements 

for the degree of 

 

Cell and Molecular Biology—Doctor of Philosophy 

 

2018 

 
 
 
 

LEVODOPA-INDUCED DYSKINESIA – FROM ABERRANT PRESYNAPTIC 

SIGNALING TO MALADAPTIVE POSTSYNAPTIC PLASTICITY 

ABSTRACT 

 

 

By 

 

Rhyomi Charis Sellnow 

 

Parkinson’s disease (PD) is a neurodegenerative disorder that arises following the 

death of dopamine (DA) neurons in the substantia nigra pars compacta (SNc). DAergic 

signaling from these neurons is required for proper signaling of the basal ganglia, a 

circuit that regulates habitual motor behaviors. As the SNc degenerates, DA signaling to 

the hub of the basal ganglia—the striatum—is drastically reduced. This progressive loss 

results in the development of parkinsonian motor symptoms, including bradykinesia, 

tremor, and gait problems. To treat these symptoms, the DA precursor L-3,4-

dihydroxyphenylalanine (L-DOPA) can be administered to reintroduce DA signaling in 

the striatum. Unfortunately, chronic treatment with L-DOPA inevitably leads to the 

development of new motor symptoms, called L-DOPA-induced dyskinesia (LID), in the 

majority of PD patients. LID development is a complex, multifaceted process. The aim 

of this dissertation is to elucidate the mechanism of LID by studying abnormal 

presynaptic signaling and the aberrant postsynaptic striatal plasticity induced in 

dyskinesia. First, we found that DA release from serotonin (5-HT) cells of the dorsal 

raphe nucleus (DRN) is a critical contributing factor to LID in a rat model of PD, and that 

regulation of DRN neurons blocks LID development. We showed this by using 

recombinant adeno-associated virus (rAAV) to express the DA autoreceptor D2Rs in 

DRN neurons, giving them ability to regulate abnormal DA release. Treatment with 

rAAV-D2Rs blocks LID development by decreasing DA efflux into the striatum. Second, 

we have characterized a novel postsynaptic molecular driver of LID, Nurr1. Nurr1 has 

been identified in genetic screens to be significantly upregulated in dyskinetic animal 

models. Therapies aimed at increasing Nurr1 are currently being investigated for PD, as 

the transcription factor is required for the health and long-term maintenance of the DA 

cells that degenerate in the disease. This dissertation provides evidence that Nurr1 

plays a direct role in LID development. Viral expression of Nurr1 in the striatum can 

induce severe LID in a rat strain that is resistant to LID. Additionally, we showed that 

LID-associated Nurr1 expression is induced by direct stimulation of the pro-movement 

pathway of the basal ganglia. Finally, we determined that Nurr1 expression causes 

changes in both the activity and morphology of striatal medium spiny neurons (MSNs). 

We have shown that, independent of L-DOPA administration, Nurr1 causes altered 

striatal activity that mimics activity changes seen in dyskinetic animals. Ectopic Nurr1 

expression also causes L-DOPA-independent decreases in dendritic spines. As 

dendritic spine plasticity is a hallmark of LID, our data suggests that Nurr1 plays a direct 

role in these maladaptive changes. Together, this dissertation provides compelling 

evidence for both presynaptic and postsynaptic mechanisms of LID development. 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

This dissertation is dedicated in memory of 

Dr. Sudhish Mishra 

A truly wonderful scientist and friend 

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

iv 

 

ACKNOWLEDGMENTS 

 
 

I first must thank my advisor, Dr. Fredric Manfredsson, for his continued guidance and 

support during my graduate career. Fredric asks some of the most insightful and 

thought-provoking scientific questions, and I have always found his drive and passion 

for the field inspiring. I am forever grateful for the way he has helped shape me on a 

scientific and personal level. I would also like to thank my committee members, Dr. 

Kathy Steece-Collier, Dr. Scott Counts, Dr. David Arnosti, and Dr. Susan Conrad for 

lending their expertise to my projects. I would like to extend particular gratitude to Kathy, 

the scientist who taught me so much about dyskinesias. The LID field is filled with 

brilliant people, and I am truly grateful to have had the chance to train with one of the 

best. Additionally, I must thank Dr. Susan Conrad, for both being a member of my 

committee as well as the director of the CMB program. She has dedicated herself to 

ensuring the success of myself and my fellow classmates, and I am very grateful for her 

support. I would also like to thank our collaborators, Dr. Christopher Bishop, Dr. 

Anthony West, and Dr. Kuei Tseng, who have all provided invaluable advice and 

assistance with experiments that have contributed to this dissertation. 

I have been lucky to have met and worked with a number of fantastic people in the 

Manfredsson lab over the last five and half years. Nathan Kuhn is the dictionary 

definition of a fantastic lab manager. He manages to keep us all afloat during even the 

most stressful of times. I have countless reasons to thank him, but mostly for his 

constant support and friendship. Dr. Ivette Sandoval is one of the best lab mates a 

graduate student could ask for. She is insightful, determined, and has always been 

there to lend a hand with surgeries or offer a story to make me laugh. I also must thank 

 

v 

 

previous lab members, Dr. Matthew Benskey and Roslyn Vashon. Matt continually 

offered his time, advice, and great book recommendations, all of which were 

instrumental in getting me through this dissertation. Roslyn was a fantastic student to 

work with, and I am grateful for the help she gave on some of my projects. 

The TSMM department is a strange and beautiful family, and there isn’t a member of 

the department who hasn’t impacted my graduate career in some way. I especially must 

thank Jennifer Stancati, who has gone above and beyond expectations by helping me 

with experiments and troubleshooting. Her friendship throughout the years is something 

I will always cherish. I would also like to thank staff from other labs, Alyson Cole-

Strauss, Christopher Kemp, and Brian Daley for all of their advice and support. 

To my fellow graduate students, I am so glad to have had the chance to get to know you 

and learn with all of you. In particular I must thank Sarah Kelly, Sandy Olenic, and 

Daniel Parell for inspiring me, driving me, and keeping me sane. 

I would finally like to thank my main cheerleaders, the Sellnows—Ellen, David, Hannah, 

Silas, and Kyrie—the Verstegens, and my other thirds—Sara Vande Walle and Brittany 

Eggert. The level of love and support you have all shown me as I’ve taken this crazy 

journey means more to me than words can express.  

My loudest and strongest support over these years has come from my partner, Joe 

Verstegen. He has been my constant rock through the most stressful and impactful 

years of my life. I cannot adequately express how much his unwavering love and 

encouragement has kept me going, and can only attempt to thank him enough for all he 

has done for me. 

 

 

vi 

 

PREFACE 

 
 

At the time of writing this dissertation, the three chapters describing my research are in 

preparation for publication. Chapter 2 has been submitted for publication, and accepted 

to Acta Neuropathologica Communications pending minor revisions. Chapters 3 and 4 

will be compiled into a single manuscript and submitted in the near future. 

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

 

vii 

 

TABLE OF CONTENTS 

 
 

LIST OF TABLES ....................................................................................................xii 
 
LIST OF FIGURES ..................................................................................................xiii 
 
KEY TO ABBREVIATIONS .....................................................................................xv 
 
Chapter 1: Introduction .........................................................................................1 
   Parkinson’s disease ...........................................................................................1 
I.  History and impact ........................................................................................1 
II.  Clinical presentation .....................................................................................2 
III.  Neuropathology of Parkinson’s disease .......................................................3 
   Treating the motor symptoms of Parkinson’s disease ...................................6 
I.  Anatomy of the basal ganglia .......................................................................6 
II.  Dopaminergic regulation of basal ganglia signaling ......................................8 
III.  L-DOPA: the beginnings of the ‘gold standard’ therapy ................................11 
IV. L-DOPA therapy and mechanism of action ..................................................13 
V.  Other dopamine-based therapies .................................................................15 
VI. Non-dopaminergic therapies.........................................................................16 
   L-DOPA-induced dyskinesia .............................................................................17 
I.  Features of LID .............................................................................................18 
II.  Clinical management strategies....................................................................19 
III.  LID risk factors ..............................................................................................20 
IV. Modeling LID for preclinical research ...........................................................21 
V.  Setting the stage for LID: dopamine depletion-induced changes .................22 
VI. Dopamine receptor signaling in LID ..............................................................23 
VII. Presynaptic mechanisms of LID: the serotonin hypothesis .........................24 
VIII. Postsynaptic LID-associated gene expression ...........................................27 
IX. Aberrant striatal plasticity in LID ...................................................................30 
   Nurr1: implications for Parkinson’s and LID....................................................34 
I.  Nurr1 biology and transcriptional regulation .................................................34 
II.  Role of Nurr1 in dopaminergic neurons and Parkinson’s disease ................36 
III.  Nurr1 in the dyskinetic striatum ....................................................................38 
   Recombinant adeno-associated virus: using gene therapy as a research tool 
 ................................................................................................................................39 
I.  rAAV biology .................................................................................................39 
II.  rAAV in neuroscience research ....................................................................40 
III.  Clinical perspective on rAAV ........................................................................41 

 

viii 

 

   Overarching significance ..................................................................................42 
   LITERATURE CITED ...........................................................................................45 
 
Chapter 2: Regulation of dopamine neurotransmission from serotonergic neurons 
by ectopic expression of the dopamine D2 autoreceptor blocks levodopa-induced 
dyskinesia ..............................................................................................................76 
   Preface ................................................................................................................76 
   Introduction ........................................................................................................76 
   Methods ..............................................................................................................81 
I.  Adeno-associated virus production ...............................................................81 
II.  Animals and surgeries ..................................................................................82 
III.  Abnormal involuntary movement (AIM) ratings and drug treatments ............84 
IV. Parkinsonian motor evaluation .....................................................................85 
V.  Tissue collection ...........................................................................................86 
VI. Immunohistochemistry ..................................................................................87 
VII. In vivo microdialysis .....................................................................................88 
VIII. High-performance liquid chromatography for monoamine tissue analysis ..89 
IX. Total enumeration of TH+ neurons to assess lesion severity .......................90 
X. Electrophysiology ..........................................................................................91 
XI. Statistical analysis ........................................................................................92 
   Results ................................................................................................................93 
I.  Validation of lesion and transgene expression .............................................93 
II.  D2Rs delivery to the dorsal raphe eliminates LID ..........................................96 
III.  D2Rs does not affect parkinsonian motor behavior .......................................98 
IV. Dopamine receptor agonists do not induce significant AIMs in L-DOPA-primed 
rAAV-D2Rs rats .............................................................................................101 

V.  D2Rs expression in the dorsal raphe reduces striatal dopamine efflux following L-

DOPA delivery ..............................................................................................104 
VI. D2Rs expression inhibits 5-HT neuron activity ..............................................106 
   Discussion ..........................................................................................................108 
   APPENDIX ...........................................................................................................115 
   LITERATURE CITED ...........................................................................................117 
 
Chapter 3: Modulating levels of Nurr1 expression in the parkinsonian rat striatum 
impacts LID severity and development ...............................................................127 
   Introduction ........................................................................................................127 
   Methods ..............................................................................................................129 
I.  Adeno-associated virus production ...............................................................129 
II.  Animals and surgeries ..................................................................................129 
III.  Abnormal involuntary movements ratings .....................................................130 

 

ix 

 

IV. Tissue collection and processing ..................................................................131 
V.  In situ hybridization .......................................................................................132 
VI. Stereology ....................................................................................................133 
VII. Statistical analysis .......................................................................................133 
   Results ................................................................................................................134 
I.  Validation of lesion and transgene expression .............................................134 
II.  Ectopic Nurr1 overexpression does not exacerbate AIMs in LID-susceptible rats 
 ...........................................................................................................................134 
III.  Ectopic Nurr1 overexpression induces severe AIMs in LID-resistant rats ....139 
IV. Nurr1 expression is induced by direct pathway activation ............................142 
V.  Viral knockdown of Nurr1 does not inhibit LID development ........................145 
   Discussion ..........................................................................................................148 
   APPENDIX ...........................................................................................................153 
   LITERATURE CITED ...........................................................................................157 
 
Chapter 4: Ectopic overexpression of Nurr1 induces a LID-like state in striatal 
medium spiny neurons .........................................................................................162 
   Preface ................................................................................................................162 
   Introduction ........................................................................................................162 
   Methods ..............................................................................................................164 
I. 
 Adeno-associated virus production ..............................................................164 
II.   Animals and surgeries .................................................................................164 
III.   Tissue collection and processing .................................................................166 
IV.  Local field potential and single cell in vivo recordings .................................167 
V.   Gogli-Cox impregnation and spine analysis ................................................168 
VI.  Stereology ...................................................................................................169 
VII.  Statistical analysis .......................................................................................169 
   Results ................................................................................................................170 
I.  Lesion validation and vector expression .......................................................170 
II.  Striatal Nurr1 expression alone induces LID-like corticostriatal signaling .....170 
III.  MSN spine density and morphology changes are induced by Nurr1 expression 
 .....................................................................................................................175 
   Discussion ..........................................................................................................177 
   LITERATURE CITED ...........................................................................................180 
 
Chapter 5: Discussion ..........................................................................................185 
   Introduction ........................................................................................................185 
   Chapter 2: Inhibiting dorsal raphe-mediated dopamine release with AAV blocks 
   LID development ................................................................................................185 
I.  Aim 1 findings, in brief ..................................................................................185 

 

x 

 

II.  Study implications and future directions .......................................................186 
   Chapter 3: Nurr1 is a molecular driver of LID development ...........................190 
I.  Aim 2 findings, in brief ..................................................................................190 
II.  Study implications and future directions .......................................................190 
   Chapter 4: Nurr1 expression induces an LID-like striatum ............................195 
I.  Aim 3 findings, in brief ..................................................................................195 
II.  Study implications and future directions .......................................................196 
   Final remarks ......................................................................................................199 
   LITERATURE CITED ...........................................................................................201 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

 

xi 

 

LIST OF TABLES 

 
 

Table 2.1 Concentrations of monoamines in lesioned versus intact hemisphere ....97 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

 

xii 

 

LIST OF FIGURES 

 
 

Figure 1.1 Schematic of basal ganglia signaling ....................................................7 
 
Figure 2.1 Experimental design and model validation ............................................94 
 
Figure 2.2 DRN D2Rs expression blocks LID development ....................................99 
 
Figure 2.3 rAAV-D2Rs does not impact L-DOPA efficacy .......................................100 
 
Figure 2.4 D2Rs-injected animals do not develop severe AIMs with DA agonist 
treatment .................................................................................................................103 
 
Figure 2.5 DRN D2Rs reduces striatal efflux of DA .................................................105 
 
Figure 2.6 Ectopic DRN D2Rs expression reduces 5-HT neuronal firing ................107 
 
Figure A.2.1 AIM scores in L-DOPA dosing paradigm ...........................................116 
 
Figure 3.1 Experimental design and model validation ............................................135 
 
Figure 3.2 Ectopic Nurr1 does not exacerbate AIMs in LID-susceptible rats ..........137 
 
Figure 3.3 Nurr1 does not impact individual AOLs in LID-susceptible rats .............138 
 
Figure 3.4 Ectopic Nurr1 expression induces severe AIMs in LID-resistant rats ....140 
 
Figure 3.5 Ectopic Nurr1 induces AIMs in LID-resistant rats that are similar to LID-
susceptible rats .......................................................................................................141 
 
Figure 3.6 Abnormal striatal Nurr1 expression is induced by direct D1 receptor 
activation .................................................................................................................143 
 
Figure 3.7 D1 stimulation induces Nurr1 expression in both indirect and direct pathway 
neurons ...................................................................................................................146 
 
Figure 3.8 Nurr1 silencing does not inhibit LID development .................................147 
 
Figure 3.9 Nurr1 shRNA does not exacerbate LID .................................................149 

 

xiii 

 

Figure A.3.1 Abnormal Nurr1 expression upregulation in dyskinetic rats ...............154 
 
Figure A.3.2 Virally-expressed Nurr1 does not induce drug-independent AIMs .....155 
 
Figure A.3.3 Nurr1 colocalization with iMSNs in quinpirole and saline treated rats 
 ................................................................................................................................156 
 
Figure 4.1 Lesion and vector confirmation ..............................................................171 
 
Figure 4.2 Striatal Nurr1 potentiates corticostriatal transmission ............................173 
 
Figure 4.3 Cortically-evoked response of striatonigral MSNs to antidromic stimulation 
 ................................................................................................................................174 
 
Figure 4.4 Nurr1-induced alterations in dendritic spine density and morphology ...176 
 
Figure 5.1 Nurr1 expression in shRNA treated animals..........................................194 
 
Figure 5.2 Proposed mechanism of LID development ............................................200 
 

 

 
 

 

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 

 

xiv 

 

 

5-HT 

6-OHDA 

α-syn 

AADC 

AIM 

AOLs 

BDNF 

BFP 

cAMP 

CNS 

COMT 

CREB 

D1 

D2 

D2Rs 

DA 

DAT 

DBS 

dMSN 

DOPAC 

KEY TO ABBREVIATIONS 

 
 

5-Hydroxytryptamine, serotonin 

6-Hydroxydopamine 

Alpha synuclein 

Aromatic amino acid decarboxylase 

Abnormal involuntary movement 

Axial, orolingual, and limb AIMs 

Brain derived neurotrophic factor 

Blue fluorescent protein 

Cyclic adenosine monophosphate 

Central nervous system 

Catechol-O-methyltransferase 

cAMP responsive element binding protein 

Dopamine receptor type 1 -- direct pathway 
specific 
  
Dopamine receptor type 2 -- indirect pathway 
specific 
  
Dopamine autoreceptor 

Dopamine 

Dopamine transporter 

Deep brain stimulation 

Medium spiny neuron of the direct pathway 

3,4-dihydroxyphenylacetic acid 

xv 

 

 

DREADD 

DRN 

Dyn 

Enk 

Designer receptor exclusively activated by 
designer drugs 
Dorsal raphe nucleus 

Dynorphin 

Enkephalin 

ERK1/2 

Extracellular signal-regulated protein kinase 

FAS 

GABA 

GFP 

GI 

GPCR 

GPe 

GPi 

HPLC 

I.P. 

IEG 

IHC 

iMSN 

ITR 

Forepaw adjusting steps 

γ-aminobutyric acid 

Green fluorescent protein 

Gastrointestinal 

G-protein coupled receptor 

Globus pallidus, external segment 

Globus pallidus, internal segment 

High-performance liquid chromatography 

Intraperitoneal 

Immediate early gene 

Immunohistochemistry 

Medium spiny neuron of the indirect pathway 

Inverted terminal repeat 

L-DOPA 

3,4-dihydroxy-L-phenylalanine, levodopa 

LFP 

LID 

LTD 

LTP 

Local field potential 

Levodopa-induced dyskinesia 

Long term depression 

Long term potentiation 

xvi 

 

 

MAO-B 

MAPK 

MFB 

MPTP 

MSK-1 

MSN 

NBRE 

NMDA 

NR4A1 

NR4A2 

NR4A3 

NurRE 

PD 

PKA 

rAAV 

RXR 

S.C. 

SERT 

SNc 

SNr 

STN 

TH 

Monoamine oxidase B 

Mitogen-activated protein kinase 

Medial forebrain bundle 

1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine  

Mitogen- and stress-activated kinase 1 

Medium spiny neuron 

NGFI-B response element 

N-methyl-D-aspartate 

Nur77 

Nurr1 

NOR-1 

Nur-responsive element 

Parkinson's disease 

Protein kinase A 

Recombinant adeno-associated virus 

Retinoid X receptor 

Subcutaneous 

Serotonin transporter 

Substantia nigra pars compacta 

Substantia nigra pars reticulata 

Subthalamic nucleus 

Tyrosine hydroxylase 

UPDRS 

Unified Parkinson's Disease Rating Scale 

xvii 

 

 

VMAT2 

Vesicular monoamine transporter 

xviii 

 

Chapter 1: Introduction 

Parkinson’s disease 

I. 

History and impact 

In 1817, an English surgeon published his characterization of a significant motor 

disorder which he had observed in six people—three of his patients, as well as three 

strangers he solely observed from a distance. The seminal work “An essay on the 

shaking palsy” by James Parkinson is the first medical characterization of the now-

termed Parkinson’s disease (PD). In his essay, Parkinson describes what are now 

recognized as the hallmark symptoms of the disease. He defined the shaking palsy as:  

“Involuntary tremulous motion, with lessened muscular power, in parts not 

in action and even when supported; with a propensity to bend the trunk 

forwards, and to pass from a walking to a running pace: the senses and 

intellects being uninjured.” (Parkinson, 2002) 

This characterization was expanded 55 years later by Jean-Martin Charcot, who 

suggested naming the disease after the English surgeon. Charcot’s characterizations 

helped separate PD from other neurological disorders and defined the basis of the 

primary motor symptoms arising from slowness rather than weakness or paralysis 

(Charcot, 1872, 1879; Gelfand, 1989). Parkinson’s recognition of a markedly distinct 

disease along with Charcot’s clinical characterization paved the way for decades of 

research to understand and treat PD. 

PD is the second most common neurodegenerative disease in the world, affecting 

approximately 1% of adults over the age of 60 (de Lau & Breteler, 2006). The 

 

1 

 

Parkinson’s Foundation Parkinson’s Prevalence Project projects that the number of PD 

patients in North America could nearly double by 2030, with over one million people 

living with the disease (Marras et al., 2018). Patient care and lost wages of both patients 

and familial caregivers placed a staggering economic burden in the United States of 

about $14 billion in 2010 (Johnson et al., 2013; Kowal et al., 2013). With the population 

shifting to a more aged demographic and projected doubling of PD patients, the 

associated economic strain demands focused research for better care of PD patients. 

II. 

Clinical presentation 

A PD diagnosis requires the presentation of cardinal motor symptoms of the disease—

termed parkinsonism. Parkinsonism describes the motor symptoms that can be caused 

by PD. These are the presence primarily of bradykinesia (slowness of movement), 

along with either a resting tremor or passive joint rigidity—both in some cases (Postuma 

et al., 2015). While the majority of PD cases are sporadic, there are familial forms of PD 

and mutations in a number of genes, such as parkin, LRRK2, and SNCA, linked to the 

disease (Klein & Westenberger, 2012; Mhyre et al., 2012). PD cases can be separated 

from other disorders that exhibit parkinsonism by evaluating further criteria, such as 

responsiveness to pharmacotherapies and non-motor symptoms like anosmia. While 

PD is traditionally considered a motor disorder, there are numerous non-motor 

symptoms that can become very debilitating, including gastrointestinal (GI) issues and 

other autonomic dysfunction, dementia, and depression (Gelb et al., 1999). 

Interestingly, some of these non-motor symptoms, such as anosmia and GI problems 

can arise years prior to the motor symptoms required for diagnosis (Mantri et al., 2018). 

 

2 

 

Though the non-motor symptoms are now deservedly being recognized, the bulk of 

research and therapeutic efforts have focused on the hallmark motor symptoms of PD. 

Motor symptoms include a resting tremor, bradykinesia, akinesia, rigidity, and postural 

instability. Many parkinsonisms are not exclusive to PD. Tremors, for instance, can be 

common with other disorders, such as dementia with Lewy bodies and multiple system 

atrophy (Wenning et al., 1995; Louis et al., 1997). Therefore, a positive diagnosis of PD 

typically requires the presence of multiple motor symptoms in combination with one 

another. Tremors are the presence of a 3-6 Hz frequency tremor of the limb while at 

rest. Tremor prevalence in patients ranges from 69%-100% (Hoehn & Yahr, 1967; 

Martin et al., 1973; Gelb et al., 1999). Bradykinesia—slow, halting movement—affects 

roughly 77% to 98% of patients (Hoehn & Yahr, 1967; Martin et al., 1973; Gelb et al., 

1999). Along with bradykinesia, many patients struggle to initiate movement due to 

akinetic ‘freezing’ as well as rigidity in the joints. Slow movements and progressive halts 

and hesitations are also seen in PD (Hoehn & Yahr, 1967). These symptoms are why 

the disease originally garnered the name paralysis agitans, which Charcot later disputed 

as these symptoms are not due to weakness or paralysis. Postural instability typically 

occurs later in disease progression, with only 37% of patients with a disease duration of 

5 years or less showing severe postural instability (Hoehn & Yahr, 1967; Gelb et al., 

1999).  

III. 

Neuropathology of Parkinson’s disease 

There are two major pathological hallmarks of PD: the presence of Lewy bodies and cell 

loss in the substantia nigra pars compacta (SNc). These pathological markers are 

evaluated post mortem and their presence confirms the PD diagnosis. 

 

3 

 

Lewy bodies are intracellular inclusions associated with PD that were first described by 

Frederich Heinrich Lewy in 1912 (Lewy, 1912; Goedert et al., 2013). Lewy bodies are 

comprised of neurofilaments, ubiquitin, and aggregated proteins found in remaining 

neurons of PD-affected brain regions (Goldman et al., 1983; Lowe et al., 1988; Pollanen 

et al., 1993; Olanow et al., 2009). One protein, alpha synuclein (α-syn), is the main 

component of these inclusions and has been studied extensively in relation to PD 

(Spillantini et al., 1997). α-Syn pathology is seen in both the brain and peripheral 

nervous system of PD patients (Braak et al., 2003; Braak et al., 2006). Multiple 

mutations in, as well as multiplications of, the α-syn encoding gene, SNCA, are 

associated with familial cases of PD, and are thought to impact the disease through 

changes in protein function and increased α-syn aggregation (Polymeropoulos et al., 

1997; Conway et al., 1998; Kruger et al., 1998; Chartier-Harlin et al., 2004; Burre et al., 

2012). Changes in α-syn expression are also observed in sporadic cases of PD, further 

suggesting its involvement in the disease process (Neystat et al., 1999; Grundemann et 

al., 2008). α-Syn pathology impacts nearly every cellular pathway that has been linked 

to PD etiology, making it one of the best validated molecular markers in the 

pathogenesis of PD (Benskey et al., 2016). While the long-standing hypothesis has 

been that α-syn aggregates are directly neurotoxic, some research has suggested that 

the toxicity may be linked to α-syn loss-of-function as the protein is sequestered into the 

aggregates (Gertz et al., 1994; Gorbatyuk et al., 2010; Kanaan & Manfredsson, 2012). 

The other key hallmark of pathology and the basis for the disease’s cardinal motor 

symptoms is loss of SNc neurons and their projections to the striatum. SNc neurons are 

dopamine (DA) producing neurons that project to the striatum, where DAergic signaling 

 

4 

 

regulates learned motor behaviors and goal-oriented behaviors (Lanciego et al., 2012). 

Anatomically, cell loss can be seen in the lack of pigment in the SNc, arising from the 

loss of neuromelanin—a byproduct of DA metabolism—as the neurons degenerate 

(Dickson; Meiser et al., 2013). It was first proposed that disruption of the basal 

ganglia—a group of interconnected nuclei which includes the striatum and regulates 

functions including voluntary motor control—may contribute to PD motor symptoms in 

the 1870s (Meynert, 1871; Engelhardt, 2013). Damage to the SNc was first reported in 

the 1920s by a number of French researchers conducting research evaluating midbrain 

pathology in PD (reviewed in (Goetz, 2011)). Post mortem evaluation of advance stage 

PD patients’ brains show severe neuronal loss in the SNc, with the caudal and 

ventrolateral tiers being most susceptible (Damier et al., 1999; Double et al., 2010; 

Jellinger, 2012). There is a corresponding loss of SNc projections throughout the 

caudate and putamen (striatum), which is correlated with dysfunctional DA metabolism 

(Kish et al., 1988; Gallagher et al., 2011). Decreased DA content due to this 

degeneration was also observed in both the striatum and SNc of PD patients (Ehringer 

& Hornykiewicz, 1960; Hornykiewicz, 1963; Hornykiewicz, 2002). It is clear that 

neurodegeneration begins long before the motor symptoms present, as nigrostriatal 

projections in the striatum are reduced roughly 50% one year after diagnosis, and that 

innervation is 70-90% depleted five years post-diagnosis (Kordower et al., 2013). With 

such dramatic levels of degeneration occurring before motor symptoms arise allowing 

for diagnosis, efforts are underway to identify biomarkers of the disease to achieve 

earlier diagnosis (Pagan, 2012; Barber et al., 2017; Frosini et al., 2017; Khodadadian et 

al., 2018; Nielsen et al., 2018). 

 

5 

 

Treating the motor symptoms of Parkinson’s disease 

While the benefit of hindsight shows the occasionally outlandish nature of early PD 

treatments—such as bloodletting and vibrating chairs—some pharmacological 

approaches taken by these pioneering clinicians hinted at the underlying cause of the 

motor symptoms (Goetz, 2011). One of the first established treatments for PD tremor 

was with belladonna alkaloids, which are now known to be anticholinergic agents and 

assist in balancing basal ganglia signaling (Price & Merritt, 1941; Weiner et al., 2013; 

Fahn, 2015). Charcot advocated for these types of treatment, along with the rye-based 

ergot fungus, which acts on DA receptors and is the basis for some current DA-agonist 

therapies such as bromocriptine (Lieberman & Goldstein, 1985; Schiff, 2006; Goetz, 

2011). Though their mechanism was not originally understood, we now know that these 

first pharmacotherapies worked by affecting the underlying cause of PD motor 

symptoms: DA dysfunction and imbalance in the basal ganglia. 

I.  Anatomy of the basal ganglia 

The basal ganglia is a series of nuclei that creates a complex circuit which regulates 

voluntary and habitual motor behavior, and goal-directed behaviors (Engelhardt, 2013). 

It is comprised of five of highly interconnected subcortical structures which, when 

properly coordinated, regulate these motor behaviors with output signals from the 

thalamus to the cortex (see Figure 1.1A for diagram of basal ganglia structures and 

signaling) (Lanciego et al., 2012; Calabresi et al., 2014). The central hub of the basal 

ganglia is the striatum. In humans and non-human primates, the striatum is 

compartmentalized into two nuclei, the caudate and putamen. In rodent models used in 

PD research, the striatum is a single nucleus. The putamen is the unit involved in basal  

 

6 

 

Figure 1.1 Schematic of basal ganglia signaling.  

(A) Basal ganglia signaling pathways achieve proper motor control through the balance of the direct (green) and indirect 
(red) pathways under normal physiological conditions. (B) In Parkinson’s disease, DAergic signaling from the SNc is lost, 
causing an imbalance in the basal ganglia and a decrease in excitatory signaling from the thalamus. (C) In L-DOPA 
induced dyskinesia, the imbalance in the basal ganglia is tipped to favor the direct pathway, promoting hyperexcitatory 
signaling from the thalamus. 

Abbreviations: DA=dopamine; GABA=γ-aminobutyric acid; L-DOPA=levodopa; GPe=globus pallidus, external segment; 
GPi=globus pallidus, internal segment; STN=subthalamic nucleus; SNr=substantia nigra pars reticulata; SNc=substantia 
nigra pars compacta 

  

 

7 

 

ganglia, with inputs from the cortex and DAergic efferents from the SNc. Striatal 

signaling regulates the rest of the basal ganglia nuclei, which include the globus pallidus 

internal and external segments (GPi, GPe, respectively) and the substantia nigra pars 

reticulata (SNr), a part of the substantia nigra separate from the SNc. GPe efferents 

project to the subthalamic nucleus (STN), which itself modulates both the SNr and the 

GPi. Basal ganglia signaling inevitably modulates thalamic signaling to the cortex, either 

directly or indirectly by affecting the signaling of basal ganglia output nuclei—GPi and 

SNr—both of which project directly to the thalamus. The regulated output signal from 

the thalamus to the cortex determines the functional movement output (Fisone et al., 

2007). 

II.  Dopaminergic regulation of basal ganglia signaling 

Proper modulation of thalamic signaling to the cortex requires an intricately fine-tuned 

series of signals throughout the basal ganglia. This is achieved through DAergic 

modulation of the two signaling pathways of the basal ganglia—termed the direct and 

indirect pathways (Fisone et al., 2007; Calabresi et al., 2014). These two pathways are 

classically thought to work in opposition with one another to balance basal ganglia 

signaling and thus balance movement. The direct pathway is canonically known as 

generating pro-movement signaling, and the indirect pathway sends anti-movement 

signals (Ghiglieri et al., 2010; Calabresi et al., 2014). These distinct pathways begin in 

the striatum and its main population of neurons, medium spiny neurons (MSNs). MSNs 

comprise ~95% of the striatum, the rest being made up by cholinergic and GABAergic 

interneurons (Kawaguchi et al., 1995; Tepper & Bolam, 2004). Their dendrites are 

studded with synaptic structures called spines, which are dynamic and altered in normal 

 

8 

 

neuronal processes as well as in disease states (Maiti et al., 2015). MSNs themselves 

utilize the inhibitory neurotransmitter γ-aminobutyric acid (GABA).  

The split between the two pathways begins with these neurons and is dependent on 

which of the two principle DA receptors each neuron expresses. Direct pathway MSNs 

(dMSNs) primarily express DA type 1 (D1) receptors, while indirect pathway neurons 

(iMSNs) express the DA type 2 (D2) isoform (Gerfen et al., 1990; Surmeier et al., 1996). 

Though D1 and D2 are the most predominant receptors, there are additional isoforms, 

being the D5 (a D1-type) and the D3 and D4 isoforms (D2-types) (Beaulieu & 

Gainetdinov, 2011). The receptor classes are defined by how they impact intracellular 

cyclic adenosine monophosphate (cAMP) levels—D1-type receptor stimulation causes 

an increase in cAMP, while D2 activation results in the opposite (Beaulieu & 

Gainetdinov, 2011). The majority of research, however, has been focused on the two 

predominant isoforms, D1 and D2. 

The final result of direct pathway signaling is increased excitatory glutamatergic 

signaling from the thalamus to the cortex, promoting movement. Beginning in the 

striatum, dMSNs send inhibitory GABAergic signals to both the GPi and the SNr. The 

efferents of both of these nuclei are also GABAergic, and thus dMSN signaling inhibits 

their inhibitory signals to the thalamus. This disinhibition allows for excitatory 

glutamatergic signals to be sent from the thalamus to the cortex, promoting movement 

(Calabresi et al., 2014). 

The indirect pathway—named such as its projections take a less direct route to the 

thalamus—also begins with GABAergic MSNs, these expressing the D2 isoform DA 

receptor. iMSNs first project to the GPe, inhibiting its signaling. The GPe projects to and 

 

9 

 

inhibits the STN with GABAergic signaling. This signal dampens STN excitatory 

glutamatergic signaling to the GPi, which outputs (as in the direct pathway) inhibit 

thalamic signaling. iMSN inhibition of the GPe lifts inhibitory signals to the STN, thus 

increasing GPi GABAergic signaling to the thalamus, leading to reduced glutamate 

release on the cortex from the thalamus. The end result is the dampening or inhibition of 

movement (Calabresi et al., 2014). 

The balance of signaling between these two pathways is paramount for successful 

motor control. The key to keeping the indirect and direct pathways working together in 

coordination is DA signaling from the SNc (Rangel-Barajas et al., 2015). Projections 

from the SNc release DA onto striatal MSNs, differentially affecting each pathway due to 

their unique receptor expression. All DA receptors are G-protein coupled receptors 

(GPCR) but D1 signaling is mediated via the coupling with excitatory Gα subunits such 

as Gαolf and Gαs, whereas D2 receptors couple with inhibitory Gαi (Corvol et al., 2004; 

Neve et al., 2004). This results in increased activity in direct pathway neurons with DA 

signaling due to an increase in cAMP signaling cascades, with the opposite effect in 

indirect pathway neurons. Tonic firing of SNc neurons during movement and goal-

directed behaviors tune the final output signal of the basal ganglia by affecting direct 

and indirect MSN activity levels (Hyland et al., 2002).  

In PD patients, the progressive loss of SNc projections in the striatum inevitably results 

in impaired DA signaling, and thus basal ganglia dysregulation (Figure 1.1B). Loss of 

DA signaling results in reduced activity of the pro-movement direct pathway, with 

simultaneous disinhibition and altered firing patterns of the indirect pathway, causing 

complications with initiating smooth and controlled movements (Raz et al., 2000). 

 

10 

 

Accordingly, decreased direct pathway signaling has been demonstrated in 

parkinsonian animal models, where dMSN firing rate has been shown to be reduced 

following DA depletion in mice (Ryan et al., 2018). This imbalance is the physiological 

root of the motor symptoms in PD. 

III.  L-DOPA: the beginnings of the ‘gold standard’ therapy 

Once the loss of DA in the striatum of PD patients was confirmed in the late 1950s 

(Ehringer & Hornykiewicz, 1960; Hornykiewicz, 1963; Hornykiewicz, 2002), therapies 

aimed at correcting basal ganglia imbalance quickly moved to the forefront. It was in the 

late 1960s that George Cotzias published impressive results of treating parkinsonisms 

by administering high-doses of the DA precursor, 3,4-Dihydroxy-L-phenylalanine, also 

called levodopa or L-DOPA (Cotzias et al., 1967; Cotzias et al., 1969). This seminal 

discovery remains the most impactful development in PD treatment history, and L-

DOPA—while not disease modifying—is still the most effective treatment for alleviating 

the motor symptoms of the disease. 

L-DOPA was first isolated from the Vicia faba bean in 1913 and chemically synthesized 

in 1921, following closely the first chemical synthesis of DA (Barger & Ewins, 1910; 

Guggenheim, 1913; Waser & Lewandowski, 1921; Fahn, 2015). L-DOPA was first 

administered to patients with PD and parkinsonisms by pharmacologist Oleh 

Hornykiewicz and his clinical partner Walther Birkmayer in 1961 (Birkmayer & 

Hornykiewicz, 1961; Fahn, 2015; Lees et al., 2015). Hornykiewicz’s previous work had 

identified that DA is lost in both the striatum and SNc of PD patients, prompting the 

movement to use L-DOPA as a therapy (Ehringer & Hornykiewicz, 1960; Hornykiewicz, 

1963; Hornykiewicz, 2002). Their study of 20 patients using L-DOPA doses from 50-

 

11 

 

150mg showed improvements in bradykinesia—but not rigidity—up to three hours 

following administration (Birkmayer & Hornykiewicz, 1961). Over the years, the two 

continued treating PD patients with weekly or biweekly administration of 25mg L-DOPA 

in over 200 cases, but results were less promising than the initial study, with nearly one 

third of patients gaining no benefit from treatment (Birkmayer & Hornykiewicz, 1964; 

Fahn, 2015). Multiple other studies using low doses of L-DOPA and D,L-DOPA (now 

known to be a less effective formulation) were similarly disappointing (Hornykiewicz, 

2002; Fahn, 2015; Lees et al., 2015). Along with lackluster motor improvement, a 

variety of GI side effects, including nausea and vomiting, occurred with L-DOPA 

treatment (for review, see (Fahn, 2015)). 

Despite growing skepticism, Cotzias moved forward with the high-dose treatment 

regimen of D,L-DOPA and published his findings 1967 (Cotzias et al., 1967). By slowly 

increasing the total daily dose to avoid the development of side effects, significant motor 

benefits in all patients began to arise with 12g daily doses. Cotzias continued to 

administer L-DOPA in conjunction with the peripheral DOPA decarboxylase inhibitor 

alpha-methyl-dopa hydrazine (carbidopa), which allowed for lower doses of L-DOPA to 

achieve motor benefits without as significant GI side effects (Cotzias et al., 1969). 

Confirmatory trials by other groups quickly began and found similar success using high-

doses of L-DOPA (Fahn, 2015). 

Intriguingly, the leap Cotzias made to significantly higher doses of L-DOPA was not 

intended to replenish lost DA in PD, but to address the loss of neuromelanin in the SNc. 

He hypothesized that the depigmentation itself was a cause of the disease, and 

proposed using L-DOPA to promote neuromelanin deposition in the brain (Cotzias et al., 

 

12 

 

1964). Without this bold approach, L-DOPA therapy may not have been pursued further 

for PD treatment for decades. 

IV. L-DOPA therapy and mechanism of action 

Exogenously supplied L-DOPA successfully ameliorates PD motor symptoms by 

replenishing lost DA content in the basal ganglia. In normal physiologic conditions, DA 

synthesis begins with the rate-limiting step of tyrosine being converted into L-DOPA by 

the highly regulated enzyme tyrosine hydroxylase (TH) (Daubner et al., 2011). L-DOPA 

is then metabolized into DA by the enzyme aromatic amino acid decarboxylase (AADC), 

the physiologic final step in DA synthesis (Blaschko, 1942; Meiser et al., 2013). When 

given exogenously, L-DOPA is taken up by L-type amino acid transporters and can be 

converted to DA indiscriminately by any neurons expressing AADC, which includes any 

remaining SNc neurons (Uchino et al., 2002; del Amo et al., 2008; Hinz et al., 2016). DA 

is then packaged into vesicles by the vesicular monoamine transporter 2 (VMAT2) (Alter 

et al., 2013) and released on striatal MSNs, reinstating the lost balance of the direct and 

indirect pathways. 

The GI side effects initially seen with L-DOPA therapy stems from DA neurons in the 

gut. There is a significant population of DA neurons in the enteric nervous system that 

regulate gut motility and are responsible for half of DA production that occurs in the 

body (Hernandez et al., 1987; Lambert et al., 1991; Eisenhofer et al., 1997; Li et al., 

2004). Thus, without intervention, large amounts of exogenously supplied L-DOPA can 

be taken up and metabolized peripherally, before it can cross the blood-brain barrier. In 

turn, this requires larger doses of L-DOPA to achieve effective levels in the brain. This 

massive increase in DA concentration and signaling in the enteric system causes the 

 

13 

 

uncomfortable GI side effects, such as nausea, vomiting, and loss of appetite. To 

counteract this, peripheral decarboxylase inhibitors such as carbidopa are used in 

combination with L-DOPA therapy to block peripheral metabolism of L-DOPA, allowing 

for higher bioavailability in the brain which reduces the required therapeutic dose 

(Barbeau et al., 1972; Nutt et al., 1985). 

In addition to carbidopa, other enzymatic inhibitors are commonly used adjunct with L-

DOPA to improve its efficacy. Catechol-O-methyltransferase (COMT) inhibitors 

(entacapone and tolcapone) and monoamine oxidase B (MAO-B) inhibitors (rasagiline, 

selegiline, safinamide) are both used to inhibit DA breakdown into inactive metabolites 

(Goetz et al., 2005; Fox et al., 2011; Meiser et al., 2013). COMT inhibitors are 

exclusively used as an adjunct therapy, while MAO-B inhibitors can be used as initial 

monotherapies for early stage and mild PD patients (Stayte & Vissel, 2014). 

Despite its therapeutic strength, L-DOPA is often not immediately prescribed, and 

instead other pharmacotherapies (discussed below) are employed with the intent to 

manage or delay the onset of L-DOPA side effects (Hubble, 2002). In fact, these side 

effects can become so debilitating that many clinicians, patients, and caregivers are 

hesitant to initiate therapy. This pervasive fear has led to a phenomenon known as “L-

DOPA phobia” which ultimately fails patients who could benefit dramatically from L-

DOPA therapy (Kurlan, 2005; Titova et al., 2018). One of these side effects include 

motor fluctuations caused during wearing-off of L-DOPA efficacy, which is observed in 

nearly half of PD patients after only two years of L-DOPA therapy (Ahlskog & Muenter, 

2001). Motor fluctuations are seen when patients alternate between periods of 

therapeutic benefit (“on” state) and a reemergence of PD symptoms as the drug wears 

 

14 

 

off and the symptoms are no longer suppressed (“off” state) (Quinn, 1998). The most 

debilitating side effect is the emergence of abnormal involuntary drug-induced 

dyskinesia (discussed below in detail). Some studies have shown that even early 

treatment with DA agonists does not prevent L-DOPA side effects once it is inevitably 

introduced, while others suggest certain agonists may at least delay their onset (Hauser 

et al., 2007; Katzenschlager et al., 2008; Fox et al., 2011; Chondrogiorgi et al., 2014). 

V.  Other dopamine-based therapies 

A number of DA receptor agonists are standardly employed to treat PD motor 

symptoms, either prior to L-DOPA therapy in early stage patients or in conjunction with 

L-DOPA as the disease progresses (Goetz et al., 2005; Fox et al., 2011; Stayte & 

Vissel, 2014). DA-agonist therapies aim to delay or prevent the onset of L-DOPA side 

effects, such as motor fluctuations seen in the wearing-off stage of the drug, and 

abnormal involuntary movements, such as dyskinesia. These therapies can be used as 

monotherapies, typically before L-DOPA therapy begins, or in combination with L-DOPA 

to better manage symptoms in advanced stages of the disease (Goetz et al., 2005). 

There are a dozen different DA agonists that have been evaluated clinically, with strong 

evidence supporting that most of these can effectively manage PD motor symptoms 

(Fox et al., 2011). Some, such as ropinirole, pramipexole, bromocriptine, and 

cabergoline have been shown to delay the onset of motor fluctuations and dyskinesia 

(Fox et al., 2011). Additionally, a subset of DA agonists (pramipexole, ropinirole, 

rotigotine, apomorphine, and pergolide) can manage motor fluctuations, but there is little 

evidence supporting that any clinical tested DA agonist treat dyskinesia (Fox et al., 

2011). 

 

15 

 

VI. Non-dopaminergic therapies 

Pharmacotherapies that do not directly affect DAergic signaling are currently being used 

and investigated clinically with the intent of increasing L-DOPA efficacy, reduce side-

effects, and/or modify disease progression (Goetz et al., 2005; Fox et al., 2011; Stayte 

& Vissel, 2014). Anticholinergics, one of the first efficacious treatment strategies 

discovered for PD, are still occasionally used today to achieve balance between 

cholinergic and dopaminergic signaling in the striatum (Katzenschlager et al., 2003). 

These drugs, however, often result in neuropsychiatric side effects such as 

hallucinations, and therefore are not used as commonly (Katzenschlager et al., 2003; 

Sprenger & Poewe, 2013). N-methyl-D-aspartate (NMDA) receptor antagonists are 

used to treat L-DOPA side effects (amantadine) and have shown improvement in 

cognition (memantine) (Verhagen Metman et al., 1998; Stayte & Vissel, 2014). 

Adenosine A2A receptor antagonists have are also in clinical trials, showing increased 

“on” L-DOPA period and decreased “off” period, but with increased time with dyskinesia 

(Stayte & Vissel, 2014). 

A number of surgical interventions are also employed by clinicians to treat their patients, 

including deep brain stimulation (DBS), targeted lesioning, cell transplantation, and 

gene therapy. DBS, the preferred surgical approach, involves the implantation of a 

stimulating electrode in basal ganglia nuclei—commonly the thalamus, STN, or GPi—to 

promote proper signaling (Goetz et al., 2005; Fox et al., 2011; Moldovan et al., 2015). 

Targeted lesions, such as thalamotomy and pallidotomy, can be useful for managing L-

DOPA motor complications, but are used much less frequently in modern therapy 

(Jankovic & Aguilar, 2008; Fox et al., 2011; Moldovan et al., 2015). Cell transplantation 

 

16 

 

has been investigated as an experimental therapy clinically since the mid-1980s, with 

varying results (Bjorklund & Kordower, 2013; Boronat-Garcia et al., 2017). While a 

subpopulation of patients experience decades of symptomatic relief from fetal ventral 

mesencephalic grafts in the striatum, another subpopulation of grafted patients see no 

benefit or loss of any benefit in a few years, with up to half experiencing major motor 

side effects (Olanow et al., 2003; Fox et al., 2011; Kefalopoulou et al., 2014; Boronat-

Garcia et al., 2017). Many clinical trials using gene therapy for Parkinson’s disease 

have been conducted, with some studies aiming to maximize the benefits of L-DOPA 

treatment by supplementing with DAergic factors, and others attempting to alter disease 

progression by delivering neuroprotective factors to diseased brain regions (Stayte & 

Vissel, 2014). These trials, while safe and well tolerated, have shown little improvement 

from current treatment options (Polinski, 2016). 

L-DOPA-induced dyskinesia 

Abnormal dyskinetic movements were first described in Cotzias’ initial reports of L-

DOPA use in PD patients (Cotzias et al., 1967; Cotzias et al., 1969). Significantly more 

debilitating than on-off fluctuations, L-DOPA-induced dyskinesia (LID) are a near 

inevitability with long term therapy, with reports ranging from 40-80% of patients 

developing them within 5 years of chronic L-DOPA treatment, and 80-90% after 10 

years (Lesser et al., 1979; Lees & Stern, 1983; de Jong et al., 1987; Ahlskog & 

Muenter, 2001; Hauser et al., 2007; Manson et al., 2012; Turcano et al., 2018). 

 

 

 

17 

 

I. 

Features of LID 

LID are a series of motor symptoms—distinct from the primary PD symptoms—that 

include excessive and abnormal hyperkinetic limb movements, abrupt jerking 

movements of the limbs, head, and neck (chorea and ballism), and sustained 

involuntary muscle contractions (dystonia). LID most commonly occur as peak-dose 

dyskinesia, where the symptoms arise when L-DOPA levels are at their highest in the 

plasma. Chorea is the most common component of LID and typically seen during peak-

dose dyskinesia. This can be significantly debilitating, in some patients growing severe 

enough to affect daily living (Quinn, 1998; Bastide et al., 2015). Dystonia is often painful 

and can occur both during peak-dose dyskinesia and during a phenomenon known as 

diphasic dyskinesias (Quinn, 1998; Bastide et al., 2015). 

LID is evaluated clinically using a number of rating scales that monitor abnormal 

involuntary movements (AIMs). Those most commonly used include the AIM scale, part 

IV of the Unified Parkinson’s Disease Rating Scale (UPDRS), the Clinical Dyskinesia 

Rating Scale, and the Rush Dyskinesia Rating Scale. The AIM scale in particular is 

noted as being one of the best tools for evaluating LID clinically (Colosimo et al., 2010). 

The AIM scale assesses severity of 10 dyskinetic movements in the face and mouth, 

limbs, and trunk. Each is given a severity score from 0-4 (Guy et al., 1976). The AIM 

scale is usefully clinically as it gives both a global AIM score as well as distinguishes 

between body areas (Colosimo et al., 2010). Researchers have adapted the AIM scale 

for use in non-human primate and rodent models of LID (Petzinger et al., 2001; 

Lundblad et al., 2002; Tan et al., 2002; Steece-Collier et al., 2003). 

 

 

18 

 

II. 

Clinical management strategies 

Once LID develop, clinicians have few effective options to manage them. The most 

straightforward approach is to reduce L-DOPA dose, but this can result in ineffective 

management of PD symptoms. It is widely thought that the pulsatile delivery method of 

oral L-DOPA and subsequent extreme peaks and valleys in DA concentration in the 

brain is a main contributor to LID (Chase, 1998; Nutt, 2007). To combat this, an 

extended release formulation can be prescribed (Fox et al., 2011). These controlled 

release formulations, while as efficacious as other formulations, may still carry the same 

risk for LID, and even increase the risk in advance stage patients (Fabbrini et al., 2010; 

Manson et al., 2012). Other approaches include constant L-DOPA/carbidopa infusion 

using an intestinal gel (Duodopa), which has been clinically shown to reduce LID 

(Antonini et al., 2016; Wirdefeldt et al., 2016). This requires surgery, and is thus more 

expensive and opens the risk for surgery related complications (Manson et al., 2012). 

Some gene therapy approaches have also been used to promote DA signaling in the 

striatum by delivering DAergic factors TH, AADC, and TH-regulating cofactor GTP-

cyclohydrolase 1 (CH1) (Palfi et al., 2018). DBS in the STN or GPi has also been shown 

to help reduce LID and lead to lower doses of L-DOPA, thus helping reduce LID 

(Anderson et al., 2005; Sankar & Lozano, 2011). While a number of pharmacotherapies 

have been tested preclinically, amantadine is the only drug that has translated well 

clinically and been approved for LID management (Wolf et al., 2010; Ory-Magne et al., 

2014; Hauser et al., 2017; Pahwa et al., 2017). 

 

 

 

19 

 

III. 

LID risk factors 

L-DOPA dose and disease duration are the two strongest predictors of LID development 

(Grandas et al., 1999; Manson et al., 2012). Longer disease duration caries a greater 

risk for LID development, which could be due to a number of factors such as increasing 

disease severity (typically requiring higher L-DOPA doses) and differing ages of onset 

(Grandas et al., 1999; Manson et al., 2012; Nicoletti et al., 2016). In fact, patients 

diagnosed before age 40 show much higher LID incidence (up to 90% in five years) 

than those diagnosed after age 60 (26% incidence with 60-69 years age of onset) 

(Kostic et al., 1991; Schrag et al., 1998; Kumar et al., 2005; Manson et al., 2012). 

Additionally, higher daily doses of L-DOPA, regardless of adjunct DA agonist therapy, 

are associated with higher rates of LID (Grandas et al., 1999; Fahn, 2005; Manson et 

al., 2012).  

Genetic risk factors for LID have also been examined, and a number of mutations in 

genes associated with familial PD, such as parkin and LRRK2, have been associated 

with higher LID risk (Dekker et al., 2003; Lesage et al., 2008; Nishioka et al., 2010). 

These mutations may not, however, be directly linked to LID as genetic forms of PD 

typically have earlier onset, which itself is a major LID risk factor. A number of studies 

have examined LID-linked polymorphisms in non-genetic (idiopathic) PD. A range of 

genes have been implicated, including members of DA metabolism and signaling 

pathways (D1, D2, D3, and D4 receptors, the dopamine transporter (DAT)) and genes 

non-DAergic genes (opioid receptors, brain derived neurotrophic factor (BDNF), MAO-

B, COMT) (Oliveri et al., 1999; Bialecka et al., 2004; Strong et al., 2006; Carta et al., 

 

20 

 

2008; Foltynie et al., 2009; Monville et al., 2009; Lee et al., 2011; Prashanth et al., 

2011; Rieck et al., 2012; Purcaro et al., 2018). 

LID etiology has been a focus of PD research since their initial observations. 

Understanding LID development is imperative to achieving more effective symptom 

management and improve the quality of life for patients. It is understood that the 

mechanisms underlying LID are multifaceted. The following sections will discuss the 

current understandings of LID etiology based on preclinical and clinical findings. 

IV.  Modeling LID for preclinical research 

Our current understanding of LID has been dramatically influenced by the use and 

characterization of animal models of LID. Non-human primates and rodents (including 

mice and rats) are the most commonly utilized models, as LID can be established, 

monitored, and modulated using a variety of techniques in these animals. The first steps 

to a model of LID begin with a model of PD, most typically generated by selectively 

lesioning the SNc with neurotoxicants (Jackson-Lewis et al., 2012). 1-Methyl-4-phenyl-

1,2,3,6-tetrahydropyridine (MPTP) is widely used to create PD-like lesions in non-

human primates and mice (rats have shown resistance to MPTP toxicity) (Jackson-

Lewis et al., 2012). The toxic properties of MPTP were inadvertently discovered when 

patients developed PD-like symptoms and pathology after taking synthetic opioids 

contaminated with MPTP (Langston et al., 1996). 6-Hydroxydopamine (6-OHDA) is also 

used regularly to induce SNc lesions in animals (Jackson-Lewis et al., 2012). An analog 

to DA, 6-OHDA is readily taken up by DAergic neurons, where its metabolism 

overwhelms the neuron with reactive oxygen species, leading to degeneration (Tieu, 

2011). Lesion techniques using either of these neurotoxicants are able to cause near 

 

21 

 

complete degeneration of the nigrostriatal system, which is useful for modeling LID as 

dyskinesia development requires at least 90% striatal DA depletion (Ulusoy et al., 2010; 

Francardo et al., 2011). Following lesioning, dyskinesia can be induced in these animals 

with acute or chronic treatment with different DA agonists or L-DOPA (Lundblad et al., 

2004; Bastide et al., 2015; Johnston & Fox, 2015). LID severity is assessed using the 

AIM scale discussed above. These models of a complex behavior are a powerful tool for 

better understanding LID etiology, as well as conducting experiments that would not be 

feasible in human patients. 

V. 

Setting the stage for LID: dopamine depletion-induced changes 

It is well established that L-DOPA “priming” is an essential step in LID development. 

Priming occurs when the DA deprived parkinsonian striatum encounters a sudden 

reintroduction of non-physiological DA signaling from either L-DOPA or DA agonist 

treatment (Jenner, 2008). DA depletion in the basal ganglia due to the disease process 

itself is required for priming. Significant changes occur in striatal MSN physiology both 

in PD patients as well as animal models. Increased D1 receptor presentation at the cell 

membrane is seen in MSNs following DA denervation, suggesting altered trafficking and 

increased sensitivity (Aubert et al., 2005; Guigoni et al., 2007; Berthet et al., 2009). DA 

depletion also increases expression of Gαolf in both post-mortem tissue and 

parkinsonian models, indicating that increases in D1 receptor coupling with Gαolf 

contribute to hypersensitization (Corvol et al., 2004; Alcacer et al., 2012). Striatopallidal 

neurons are significantly affected by alterations in glutamatergic synapses from cortical 

inputs, which drive indirect pathway signaling (Picconi et al., 2004; Day et al., 2006). 

DA-depletion does not induce similar changes in glutamatergic synapses of striatonigral 

 

22 

 

neurons of the direct pathway (Day et al., 2006). Additionally, overall dendritic spine 

density on MSNs is reduced in parkinsonian animal models (Zhang et al., 2013; 

Fieblinger et al., 2014). Changes to MSNs go beyond spine plasticity, as drastic 

dearborization of both striatopallidal and striatonigral MSNs dendrites occurs following 

DA denervation (Zaja-Milatovic et al., 2005; Deutch et al., 2007; Fieblinger & Cenci, 

2015). 

The drastic morphological and physiologic changes in the basal ganglia that occur due 

to DA depletion set the stage for LID induction. When the hypersensitized striatum is 

reintroduced to non-physiological DA signaling with L-DOPA or DA agonist therapy, the 

system becomes primed with further morphological and physiologic changes that 

inevitably lead to LID. Therapeutic efficacy of L-DOPA is lost in LID, as basal ganglia 

signaling is pushed out of balance once again, with striatal hypersensitivity driving 

aberrant pro-movement signals (Figure 1.1C). 

VI. 

Dopamine receptor signaling in LID 

Stimulating DA receptors with either exogenous L-DOPA therapy or DA direct agonists 

in the denervated striatum of PD patients and animal models results in dyskinesia-

promoting signaling. The bulk of research has focused on changes in D1 receptor 

signaling, as specific dMSN stimulation with receptor agonists can induce L-DOPA-

comparable severity dyskinesia (Rascol et al., 2001). Optogenetic stimulation of dMSNs 

also produces severe dyskinesia, further suggesting that D1 activation rather than DA 

itself plays a key role in the behavior (Ryan et al., 2018). D1 stimulation leads to 

downstream activation of the cellular signaling gene protein kinase A (PKA), which 

phosphorylates a number of downstream effectors (Neve et al., 2004). Inhibiting PKA 

 

23 

 

activity reduces LID in rodents, indicating that chronic D1 signaling and the PKA 

cascade are involved in the behavior (Lebel et al., 2010). Selectively inactivating 

downstream effectors of PKA in dMSNs is also able to reduce LID severity, further 

implicating the direct pathway signaling as being an instigator for LID (Bateup et al., 

2010). 

The research of D2 receptor signaling has been less involved than D1 signaling, but it is 

clear that these receptors also play a role in LID. This is showcased by the fact that D2 

agonism alone primes the striatum and can induce dyskinesia on its own (Gomez-

Mancilla & Bédard, 1992; Blanchet et al., 1993; Pollack & Yates, 1999; Drake et al., 

2013; Chondrogiorgi et al., 2014). D2 receptor expression is upregulated following DA-

depletion, and this is not reverted following L-DOPA therapy (Aubert et al., 2005; 

Guigoni et al., 2007). Recordings of iMSNs show that L-DOPA reduces their firing rate, 

indicating a decrease in indirect pathway signaling during LID (Ryan et al., 2018). 

Together, the research on DA receptors in LID has shown that abnormal signaling and 

receptor dynamics in both pathways occurs during LID. 

VII.  Presynaptic mechanisms of LID: the serotonin hypothesis 

It is now understood that at the time of diagnosis and as PD patients begin DAergic 

therapy, dramatic cell loss in the SNc and DA terminals in the striatum has already 

occurred (Kordower et al., 2013). Despite this, L-DOPA therapy is still efficacious for 

symptom relief, even as the disease advances and nearly all nigrostriatal neurons are 

lost. The obvious question arises: if DAergic innervation of the striatum is lost, what 

neuronal population is processing L-DOPA and releasing DA into the striatum? While 

there are certainly compensatory mechanisms that stave off motor symptoms until 

 

24 

 

severe DA loss (Blesa et al., 2017), the leading hypothesis is that 5-hydroxytryptamine 

(5-HT, or serotonin) neurons of the dorsal raphe nucleus (DRN) become the primary 

site of DA release into the basal ganglia following L-DOPA administration. 

The DRN is a bilateral brainstem nucleus comprised of a heterogeneous population of 

neurons. It contains roughly half of the 5-HT neurons of the rat and human brain 

(Jacobs & Azmitia, 1992). It is also comprised of GABAergic and a small population of 

DAergic neurons (Belin et al., 1983; Descarries et al., 1986). Highly collateralized 5-HT 

outputs from the DRN provide a primary source of 5-HT signaling in the forebrain, with 

target structures including the amygdala, cortex, nucleus accumbens, and the striatum 

(Waselus et al., 2011). DRN target structures suggest 5-HTergic influences a wide 

variety of behaviors, from stress and anxiety to motor control. 

A key component to the 5-HT hypothesis of LID is the fact that 5-HT neurons are able to 

convert L-DOPA into DA. 5-HTergic neurons express AADC, the enzyme which 

converts L-DOPA to DA. It has been previously shown that exogenous L-DOPA is 

indeed taken up by 5-HT neurons and converted into DA (Arai et al., 1995). 

Furthermore, imaging data has suggested that the majority of striatal DA release in 

advanced PD patients comes from 5-HT neurons (Roussakis et al., 2016). Thus, the 5-

HT system is pivotal for efficacious L-DOPA therapy as the DAergic system is 

progressively lost. Further studies, however, additionally implicate DRN 5-HT neurons in 

dyskinesia development. 

5-HTergic DRN projections moderately innervate the striatum under normal physiologic 

conditions, where 5-HT signaling is thought to affect goal-directed behaviors and 

voluntary movement (Imai et al., 1986; Waselus et al., 2011; Mathur & Lovinger, 2012). 

 

25 

 

In parkinsonian animal models, 5-HTergic innervation of the striatum increases 

substantially (Maeda et al., 2003; Hiromasa Yamada et al., 2007; Rylander et al., 2010). 

This hyperinnervation occurs rapidly, with observable changes only two weeks following 

DA denervation in animal models (Maeda et al., 2003). Additional studies suggest that 

L-DOPA treatment itself also contributes to 5-HTergic axonal terminal sprouting in the 

striatum (Rylander et al., 2010). Recently, virally overexpressed BDNF was shown to 

induce serotonin fiber sprouting in the rat striatum, which correlated with more severe 

AIMs (Tronci et al., 2017). Together, this evidence suggests that 5-HT hyperinnervation 

allows for excess DA release in the striatum from DRN neurons.  

Importantly, while DRN neurons can synthesize and release DA, they cannot effectively 

regulate DA release, as they do not express DAergic regulatory factors such as DAT or 

the dopamine autoreceptor (D2Rs). These factors are essential to proper modulation of 

DA signaling from DAergic neurons (Ford, 2014). Without this regulation, it is 

hypothesized that 5-HT neurons release DA in an unregulated fashion in the striatum. 

This unregulated release contributes to the pulsatile levels of L-DOPA in the system due 

to intermittent oral dosing that has been shown to drive LID (Cenci & Konradi, 2010). 

This dyskinesia-inducing mechanism for LID is supported by data showing that ablating 

the DRN, or modulating its activity with 5-HT agonists, can eliminate or reduce LID in 

animal models (Carta et al., 2007; Eskow et al., 2009; Bezard et al., 2013; Politis et al., 

2014; Ghiglieri et al., 2016; Meadows et al., 2017).  

The rationale for this theory of dyskinesiogenesis has prompted numerous clinical 

studies utilizing 5-HT agonists to treat LID. In practice, many of these drugs have fallen 

short, either negating anti-parkinsonian efficacy of L-DOPA or causing a worsening of 

 

26 

 

PD symptoms (Kannari et al., 2002; Iravani et al., 2006; Olanow et al., 2009; Cheshire & 

Williams, 2012). Despite disappointment in clinical trials, the rationale for this 

presynaptic mechanism of LID warrants further research in order to better understand 

how 5-HT neurons are involved in LID genesis. DAergic signaling in the denervated 

striatum is widely accepted to induce multiple postsynaptic changes that lead to LID, 

and thus regulating the release of DA following L-DOPA administration is an attractive 

approach for LID management. 

VIII.  Postsynaptic LID-associated gene expression 

A variety of gene expression changes are associated with LID presentation. Changes in 

the extracellular signal-regulated protein kinase 1 and 2 (ERK1/2) signaling pathway 

and its downstream targets have been heavily researched in LID. Canonical D1 

receptor signaling induces increases in cAMP following activation of PKA, which 

promotes transcription by the cAMP responsive element binding protein (CREB) (Cole 

et al., 1994; Konradi et al., 1994). Cross talk between glutamatergic NMDA receptors 

and D1 activation also promotes ERK1/2 phosphorylation (Girault et al., 2007). 

Following DA denervation and L-DOPA treatment, however, ERK1/2 activation 

(phosphorylation) and signaling occurs in dMSNs in an NMDA-independent manner 

(Gerfen et al., 2002; Fiorentini et al., 2013). The activation of the ERK1/2 cascade 

promotes transcription of a number of genes through its downstream effectors CREB 

and mitogen- and stress-activated kinase 1 (MSK1). Chronic activation of ERK1/2 leads 

to changes in a number of transcripts that have been found to be involved in LID. A 

transcriptomic study of differential expression following L-DOPA therapy and LID 

 

27 

 

induction has confirmed this, showing increases in transcripts such as FosB, Junb, Arc, 

and Narp (Charbonnier-Beaupel et al., 2015). 

One notable category of expression changes is the increase in immediate early gene 

(IEG) expression with LID. IEGs are expressed throughout the brain and are rapidly 

induced following stimulus and can be induced by ERK1/2 cascades. Neuronal IEGs 

are heavily involved in synaptic plasticity that is required for a variety of functions, such 

as learning and memory (Okuno, 2011). The IEG FosB and its spliced isoform ΔFosB 

are one of the hallmark molecular markers of LID, and is upregulated following ERK1/2 

signaling (Feyder et al., 2016). Both isoforms are highly upregulated in MSNs of 

dyskinetic rodent and non-human primate models of LID (Andersson et al., 1999; Pavón 

et al., 2006; Cenci & Konradi, 2010) and this has been confirmed in post-mortem PD 

patient tissue (Tekumalla et al., 2001; Lindgren et al., 2011). Higher endogenous 

expression of ΔFosB protein correlates with more severe dyskinetic behavior (Feyder et 

al., 2016). While ΔFosB and other genes show elevated expression with LID, it is 

important to differentiate between gene expression that is simply a consequence of the 

behavioral phenotype and those that play an active role in the process. This can be 

achieved by modulating the levels of specific genes in LID animal models. Accordingly, 

researchers have shown that ΔFosB is actively involved in LID development, as virally-

mediated overexpression of ΔFosB exacerbates LID, and silencing the transcript can 

reduce behavior severity (Andersson et al., 1999; Engeln et al., 2016). 

FosB is not the only IEG overexpressed in LID. Other IEGs including the cytoskeleton-

associated protein Arc, transcription factor Zif268, and scaffolding protein Homer-1a 

have also been shown to become overexpressed in LID (Sgambato-Faure et al., 2005; 

 

28 

 

H. Yamada et al., 2007; Ebihara et al., 2011). Arc and Homer-1a are both associated 

with D1, but not D2, receptor activation (Sgambato-Faure et al., 2005; H. Yamada et al., 

2007). Homer-1a is an example of a molecular marker that does not appear to play a 

causative role in LID, as LID can be reduced without reducing Homer-1a expression 

(Jimenez et al., 2009). 

A handful of studies have investigated the expression signatures associated with LID in 

rat and mouse models. These have offered insights into molecular networks involved in 

LID. One study comparing dyskinetic to non-dyskinetic rats showed dysregulation of 

calcium signaling genes and synaptic plasticity (Konradi et al., 2004). Another study 

instead compared rats treated acutely with L-DOPA with chronically treated animals. 

There were 16 genes that were commonly regulated in both treatment groups (El Atifi-

Borel et al., 2009). These shared genes are involved in a number of cellular 

mechanisms, including signal transduction, phosphorylation, and gene transcription and 

translation. There were a greater number of enhanced transcripts in the animals treated 

chronically with L-DOPA, and the researchers postulated that these changes were 

involved in the induction of structural changes in MSNs leading to LID (El Atifi-Borel et 

al., 2009).  

While most of the omic-level characterizations of LID have looked in the striatum 

globally, there have been some efforts to elucidate specific changes in the dMSNs 

versus iMSNs (Heiman et al., 2014). Distinguishing between the two basal ganglia 

pathways is important, as there are differential changes in D1- and D2- dependent 

signaling in LID, as discussed above. One study by Dr. Paul Greengard and colleagues 

captured the translational profile of mouse dMSNs and iMSNs in response to chronic L-

 

29 

 

DOPA (Heiman et al., 2014). Over 4,500 genes were differentially translated in dMSNs 

of severely dyskinetic animals. Some of the primary pathways affected by the 

expression changes included the mitogen-activated protein kinase (MAPK)/ERK1/2 

pathway and actin cytoskeleton regulation. These results are indicative of the known 

increase in ERK1/2 signaling in LID, as well as morphological changes in MSNs 

(Heiman et al., 2014). Of the identified genes most highly correlated with higher L-

DOPA doses and more severe LID, only one had been previously associated with LID 

(the Trh gene) (Cantuti-Castelvetri et al., 2010). This alone shows the importance of 

large-scale expression studies in LID, as many of the other targets may not otherwise 

have been identified as being involved. Other severe LID-associated genes that were 

upregulated in dMSNs included Fosl1 (a member of the Fos family), Ire3 (an ERK1/2 

dephosphorylation inhibitor), and Nr4a2 (or Nurr1, a required transcription factor in 

DAergic neurons) (Heiman et al., 2014). Differential expression in iMSNs following 

severe LID induction was markedly less drastic, with only 415 genes showing changes 

(Heiman et al., 2014). These findings reaffirm previous work displaying heightened 

activity of dMSNs in LID with simultaneous decreases in iMSN activity (Fieblinger et al., 

2014; Ryan et al., 2018). 

IX. 

Aberrant striatal plasticity in LID 

The magnitude of gene expression changes occurring with LID are thought to induce 

maladaptive changes in the physiology and morphology of MSNs. Numerous studies 

have characterized some of these changes in both human patients with LID as well as 

animal models. 

 

30 

 

It is classically thought that LID results following a substantial decrease in neuronal 

firing from the GPi, allowing for heightened excitatory signaling from the thalamus. This 

has been supported in PD patients with recordings of the GPi taken during operations 

for DBS implantations or pallidotomies. Single cell recordings monitoring individual GPi 

neurons show a reduction in firing rate in the “on” period with dyskinesia induced by 

apomorphine (Merello et al., 1999; Levy et al., 2001; Lee et al., 2007). However, this 

mechanism is challenged by the fact that lesions of the GPi can both improve PD motor 

symptoms and attenuate LID (Calabresi et al., 2014; Krishnan & Pisharady, 2017). 

A recent study in mice recorded from individual dMSNs or iMSNs identified 

optogenetically to determine firing changes following lesion and with LID (Ryan et al., 

2018). This study showed a dramatic reduction in firing rats of dMSNs following 

denervation, with a marked increase in LID-expressing mice (Ryan et al., 2018). This 

agrees with the classical assumption of overactivity of the direct pathway in LID 

(Calabresi et al., 2014). The researchers also observed a decrease in iMSN firing with 

LID, further implicating a striking imbalance between direct and indirect pathway 

signaling as a driver of LID (Ryan et al., 2018). 

One of the most established electrophysiological hallmarks of LID is the loss of 

bidirectional plasticity in MSNs following LID development. Bidirectional plasticity allows 

for the strengthening and pruning back of synapses depending on neuronal activity, and 

is important for many processes including motor learning and memory (Graybiel, 1995). 

Synaptic changes are influenced by long term potentiation (LTP), which causes 

reinforced synaptic connections with repeated stimuli, and long-term depression (LTD), 

which decreases synaptic strength (Citri & Malenka, 2008). LTP can be induced in 

 

31 

 

MSNs with repetitive high-frequency stimulation (Calabresi et al., 1992; Charpier & 

Deniau, 1997). However, following DA denervation, MSNs lose LTP capabilities in 

animal models, resulting in unilateral plasticity only (Centonze et al., 1999; Picconi et 

al., 2003; Paillé et al., 2010). Therapeutic (non-LID-inducing) doses of L-DOPA can 

restore both LTP and LTD in lesioned animals. However, rats that develop dyskinesia 

display a reestablishment of LTP but not LTD on L-DOPA (Picconi et al., 2003). This 

suggests DA-induced strengthening of potentially maladaptive synapses that cannot be 

pruned back, promoting abnormal movements. 

The original studies identifying the loss of bidirectional plasticity did not differentiate 

between the direct and indirect pathways. Differences between the two pathways have 

been looked at in a more recent study using transgenic mice (Thiele et al., 2014). 

Indeed, the researchers found unilateral plasticity following DA denervation, with 

dMSNs expressing LTD and iMSNs expressing LTP (Thiele et al., 2014). However, they 

observed differential plasticity between the two pathways in dyskinetic animals, with 

dMSNs only expressing LTP and iMSNs capable of only LTD (Thiele et al., 2014). The 

changes with LID in each pathway are opposite to their response following denervation, 

supporting the hypothesis of basal ganglia imbalance in both PD and LID.  

As aberrant synaptic plasticity has been shown to influence LID through 

electrophysiology alterations, a number of studies have worked to identify a 

morphological basis for these changes. Specifically, studies assessing the density and 

morphology of dendritic spines on MSNs have shown that there are dramatic changes 

following DA denervation as well as LID induction (Fieblinger & Cenci, 2015). Dendritic 

spines are highly dynamic structures that are the site of synapse formation from many 

 

32 

 

input nuclei. Spines on MSNs primarily form excitatory glutamatergic synapses with 

inputs from the cortex and thalamus (Lacey et al., 2005; Ding et al., 2008). Alterations in 

spine density and their morphology can influence many neuronal processes as well as 

be indicative of some disease states (Maiti et al., 2015). Accordingly, studies have 

shown changes in synaptic architecture of MSNs in dyskinetic animals. It is established 

that striatal DA depletion leads to an overall loss of MSN dendrites and dendritic spines, 

and thus a loss of synaptic connections (Zhang et al., 2013; Suárez et al., 2014). Some 

studies suggest this loss only occurs in iMSNs (Fieblinger et al., 2014; Nishijima et al., 

2014). Drastic morphology changes are again observed following L-DOPA treatment, 

though there has been some variability presumably due to study design. In one study 

that did not differentiate between dMSNs and iMSNs, an overall increase in spine 

density, as well as an increase in mature mushroom-type spines (Zhang et al., 2013). 

Studies that have differentiated the pathways have a general consensus that LID is 

associated with an increase in iMSN spine density with a concomitant decrease in 

dMSN spines (Fieblinger et al., 2014; Nishijima et al., 2014; Suárez et al., 2014). These 

spines are the post-synaptic targets for innervating neurons from the motor cortex and 

other structures, and changes in the synaptic connectivity due to changes in spine 

number and morphology can affect the activity of the circuit. The differential pattern of 

plasticity observed following chronic L-DOPA treatment may have a role in LID genesis. 

This is further supported by a study that found unusual and irregular glutamatergic 

corticostriatal synapse formation in rats with LID, suggesting aberrant excitatory input to 

the striatum (Zhang et al., 2013). Taken together, these data implicate that maladaptive 

synaptic plasticity occurs in the denervated striatum following L-DOPA treatment and 

 

33 

 

that this contributes to the development of LID. Identifying molecular markers that are 

involved in these plasticity changes is an important effort to better understand LID. 

Nurr1: implications for Parkinson’s and LID 

Better clinical management of LID requires a stronger understanding of the molecular 

mechanisms behind the behavior. The omic-level studies discussed above have 

identified numerous important pathways and genes that may be involved in the 

maladaptive changes that lead to LID. One such gene is Nurr1, which has been 

identified as being upregulated in MSNs of dyskinetic mice (Heiman et al., 2014; 

Sodersten et al., 2014). Additional unpublished gene array data from our group 

corroborates these findings in rats, indicating that Nurr1 is a compelling target for further 

research in relation to LID. 

I. 

Nurr1 biology and transcriptional regulation 

Nurr1, or NR4A2, is a member of the NR4A nuclear receptor family and expressed in a 

number of brain regions (Law et al., 1992). Nurr1, like its family members NR4A1 

(Nur77) and NR4A3 (NOR-1), is an IEG and its expression is rapidly induced by a 

variety of cellular signals, including ERK1/2 signaling (Sacchetti et al., 2006; Campos-

Melo et al., 2013). Nurr1 mRNA is highly expressed in a range of structures in the 

developing mouse and rat brain (Zetterstrom et al., 1996; Hirokawa et al., 2008). Nurr1 

expression persists into adulthood, with transcripts localized in the olfactory bulb, 

cortex, hippocampus, and SNc, among others (Xiao et al., 1996; Zetterstrom et al., 

1996).  

 

34 

 

Nurr1 is an orphan nuclear receptor, meaning that it can regulate transcription 

independent of ligand binding (Giguere, 1999). In fact, analysis of the crystal structure 

of Nurr1 shows the ligand binding domain is filled with hydrophobic residues that take 

up the space for ligand binding seen in other nuclear receptors, further implying its 

orphan state (Wang et al., 2003). Still, Nurr1 activity can be regulated by its binding to 

fatty acids as well as introduction of synthetic compounds (Hammond et al., 2015; de 

Vera et al., 2016; de Vera et al., 2018). Nurr1 transcriptional activity has also been 

shown to change based on its expression levels (Johnson et al., 2011). For instance, 

lower levels of Nurr1 may inhibit inflammatory signaling, whereas high levels can induce 

it (Do, 2014). Nurr1-mediated gene transcription in some pathways are impacted 

similarly irrespective of Nurr1 levels, including the MAPK signaling cascade and 

DAergic synaptic proteins (Do, 2014). 

Nurr1 activity is also dependent on its dimerization state. The transcription factor can 

act as a monomer or homodimer, as well as heterodimerizing with other NR4A family 

members (Maira et al., 1999; Hawk & Abel, 2011). Nurr1 can also heterodimerize with 

non-NR4A members, such as the Retinoid X receptor (RXR) receptor, to stimulate 

differential transcription (Zetterström et al., 1996; Volakakis et al., 2015). The 

dimerization state of the NR4A family affects what DNA sequences the proteins can 

bind to. As monomers, the binding specificity of NR4A transcription factors is the NGFI-

B response element (NBRE) (Paulsen et al., 1995). As homodimers or heterodimers 

with other NR4A members, Nurr1 and its family members affect transcription by binding 

the Nur-responsive element (NurRE) (Philips et al., 1997; Maxwell & Muscat, 2006). 

Nurr1 again displays differential DNA binding when in heterodimers with RXR, where it 

 

35 

 

binds the DR5 motif to regulate retinoid signaling (Zetterström et al., 1996). Together, 

this shows that Nurr1 displays unique transcriptional activity based on its dimerization 

status and expression levels based on cellular activity. 

II. 

Role of Nurr1 in dopaminergic neurons and Parkinson’s disease 

There are a number of reasons why Nurr1 is an interesting candidate to further study in 

regards to LID. One of the most well characterized roles of Nurr1 is in the development 

and long-term health of DA neurons of the SNc (Zetterstrom et al., 1997; Jiang et al., 

2005; Kadkhodaei et al., 2009). As discussed above, the Nurr1 transcript is abundant in 

the SNc of both prenatal and postnatal rodents (Xiao et al., 1996; Zetterstrom et al., 

1996). The importance of Nurr1 in development was shown in Nurr1 null mice, which do 

not develop DAergic neurons in the SNc and do not survive long after birth (Zetterstrom 

et al., 1997). DA levels in the striatum of these mice are reduced by 98% (Le et al., 

1999). This is only observed in midbrain DA neurons, as smaller DAergic populations in 

the hypothalamus and olfactory bulbs are not susceptible in these mice (Le et al., 1999). 

This evidence shows that Nurr1 is a critical factor required for the proper development 

of the midbrain DA neurons that are susceptible in PD. The impact of Nurr1 goes 

beyond development, as heterozygous knockout Nurr1 mice begin to display DAergic 

dysfunction over time. The heterozygotes—which are viable—show motor deficits 

similar to parkinsonian models, decreased DA in the striatum, and a loss of SNc 

neurons (Jiang et al., 2005). Further, conditionally knocking down Nurr1 in the SNc of 

adult mice cause progressive SNc cell loss and motor deficits (Kadkhodaei et al., 2009). 

Nurr1 has been shown to regulate the transcription of many DA-specific genes, 

including TH, AADC, and DAT (Zetterstrom et al., 1997; Saucedo-Cardenas et al., 

 

36 

 

1998; Sakurada et al., 1999; Decressac et al., 2013). Dysregulation of these genes in 

the knockout mouse models likely contributes to DA loss and cell death. Nurr1 also is 

involved in mitochondrial gene transcription, and its reduced expression may impact 

oxidative stress that promotes cell death (Decressac et al., 2013). These pivotal 

preclinical studies indicate that Nurr1 is both required for the development of SNc DA 

neurons and important for their health and maintenance in adulthood. 

In humans, nigral Nurr1 expression decreases with aging—a major risk factor for PD 

(Chu et al., 2002). Accordingly, PD patients show a marked decrease in SNc Nurr1 

expression that correlates with TH loss (Chu et al., 2006). There have been a number of 

Nurr1 polymorphisms identified that may be associated with familial PD, but it is unclear 

if these contribute to PD risk in the general population (Le et al., 2003; Zheng et al., 

2003; Nichols et al., 2004; Liu et al., 2013).  

An additional link to PD comes from research elucidating the interaction between Nurr1 

and α-syn. Viral overexpression of α-syn in the rat SNc causes a subsequent down-

regulation of Nurr1 (Decressac et al., 2012). Forcing the overexpression of Nurr1 in 

these neurons can reverse α-syn-mediated changes in gene expression (Volakakis et 

al., 2015). This evidence shows that Nurr1 is involved in the pathology and progression 

of PD at multiple levels. Because of its clear role in DAergic neuron health, a number of 

studies have worked to identify Nurr1 agonists as potential disease-altering treatments 

for PD (Kim et al., 2015; Smith et al., 2015). 

 

 

 

37 

 

III. 

 Nurr1 in the dyskinetic striatum 

While Nur77 and NOR-1 show endogenous expression in the striatum, Nurr1 does not 

(Xiao et al., 1996; Zetterstrom et al., 1996). It is expressed in a small population of TH-

positive striatal interneurons, but not in MSNs (Cossette et al., 2004). This makes the 

upregulation of Nurr1 in the dyskinetic striatum highly unusual, and an interesting target 

for therapeutic approaches. Nothing is known about the role of Nurr1 in MSNs, as it has 

not been observed in these neurons until recently in LID-expressing animals. Thus, 

research elucidating how Nurr1 is acting in MSNs is appropriate to better understand 

LID. 

A potential role for Nurr1 in LID could be through the induction of maladaptive plasticity. 

As discussed above, aberrant spine dynamics and neuronal activity are associated with 

LID development (Bastide et al., 2015). Previous work has shown that Nurr1 plays a 

substantial role in hippocampal synaptic plasticity required for learning and memory 

(Hawk & Abel, 2011). Nurr1 transcript is upregulated in rat brains after undergoing 

spatial learning tasks (Peña de Ortiz et al., 2000). Performance in memory task is 

impaired when Nurr1 is silenced in the rat hippocampus, further suggesting its 

importance in plasticity (Colon-Cesario et al., 2006). Knowing that Nurr1 retains these 

capabilities, it is reasonable to question if the transcription factor assumes a similar role 

when abnormally expressed in MSNs during LID. 

Nurr1 expression in MSNs also opens the possibility of LID-driving transcription by 

heterodimerization with Nur77. Nur77 is endogenously expressed in the striatum, and is 

increased by L-DOPA treatment in denervated animals, primarily in dMSNs (St-Hilaire 

et al., 2003; Mahmoudi et al., 2009). In fact, it was recently shown that viral 

 

38 

 

overexpression of Nur77 in the striatum can exacerbate AIMs in rats, suggesting it plays 

a direct role in LID severity (Rouillard et al., 2018). Nur77 expression has additionally 

been shown to induce spine loss in hippocampal neurons, showing that other members 

of the NR4A family affect synaptic plasticity (Chen et al., 2014). It is possible that Nurr1 

acts in a similar fashion to—or even in conjunction with—Nur77 to alter the striatum and 

promote LID. Taken together, the Nurr1 is an intriguing target to study in LID, especially 

considering potential Nurr1 agonist therapies for PD that could affect Nurr1 in the 

striatum. 

Recombinant adeno-associated virus: using gene therapy as a research tool 

I. 

rAAV biology 

Major advancements in the current understanding of LID have been achieved by 

modulating gene expression in preclinical models using gene therapy techniques. One 

such tool used widely in neuroscience research employs recombinant adeno-associated 

virus, or rAAV. Wild type AAV is a non-enveloped parvovirus that does not cause any 

known disease or illness, with over 70% of the population having been infected at some 

time (Calcedo et al., 2009; Choudhury et al., 2017). The AAV genome was 

reengineered for research and clinical purposes as a vector delivery tool. The only 

remnants of the wild type genome in rAAV are the inverted terminal repeats (ITRs) 

which are required elements for packaging the genome into the viral capsid, as well as 

genome stabilization and transcription (Schultz & Chamberlain, 2008; Nonnenmacher & 

Weber, 2012).  

 

39 

 

The wildtype genes required for genome replication and capsid formation are removed 

in rAAV, leaving approximately 5 kilobases of space to insert genetic material of interest 

(Samulski & Muzyczka, 2014). The engineered cassette must include enhancer and 

promoter elements to either drive ubiquitous or cell-specific expression (Gray et al., 

2011). Spatial and temporal expression can also be controlled using Cre-dependent 

genomes in transgenic animals, which can allow for conditional knockouts/knock-ins or 

cell-specific expression (Schnütgen et al., 2003). There are numerous approaches to 

control gene expression with rAAV depending on the expression cassette employed. A 

single gene of interest can be inserted into the vector to drive overexpression within 

cells. Numerous approaches for dual-expression using multicistronic genomes have 

been developed to allow for expression of multiple genes from one virus (Donnelly et 

al., 2001; Ngoi et al., 2004; Fagoe et al., 2013). Conversely, gene knockdown/knockout 

approaches are widely used, employing RNA interference (shRNAs, siRNAs, miRNAs, 

etc.) and gene editing (CRISPR). The versatility of rAAV makes it a robust research tool 

for research. 

II. 

rAAV in neuroscience research 

rAAV is a strong tool to use in the central nervous system (CNS) as it is able to 

transduce non-dividing cells (Choudhury et al., 2017). A number of rAAV capsid 

varieties—called pseudotypes—are capable of efficiently transducing neurons to 

achieve long-term, stable gene expression (Burger et al., 2004; McFarland et al., 2009). 

rAAV particles bind with extracellular receptors based on their pseudotype and are 

taken into the cell and trafficked into the nucleus, where the genome is released 

(Schultz & Chamberlain, 2008; Nonnenmacher & Weber, 2012). The viral DNA does not 

 

40 

 

integrate into the host genome, but instead is held episomally (Choudhury et al., 2017). 

Episomal DNA is not lost in non-dividing cells such as neurons, allowing for long-term, 

stable expression of the viral cassette in transduced cells. Stable rAAV-mediated gene 

expression has been observed two years following delivery in rat models, and 15 years 

in non-human primates (Klein et al., 2002; Aubert et al., 2005; Sehara et al., 2017). 

rAAV has been used in many preclinical studies that have progressed our 

understanding of LID etiology. Virally-mediated expression of candidate genes of LID 

has helped elucidate genes that are involved in LID development and severity. For 

instance, rAAV-mediated overexpression of genes such as FosB, Nur77, and BDNF can 

exacerbate LID severity, indicating that these factors have direct involvement in 

dyskinesia (Berton et al., 2009; Cao et al., 2010; Cote et al., 2014; Tronci et al., 2017; 

Rouillard et al., 2018). Conversely, LID severity can be ameliorated in these models by 

virally silencing LID-driving genes, or overexpressing genes deficient in LID (Marongiu 

et al., 2016; Park et al., 2016). Thus, using rAAV is a strong tool for characterizing novel 

molecular markers in LID. 

III. 

Clinical perspective on rAAV 

rAAV is not only a powerful research tool, but useful for gene therapy in PD patients. 

rAAV vectors have been shown to be safe and well tolerated multiple clinical trials, as 

they do not generate a significant inflammatory response (Choudhury et al., 2017). Over 

the past few decades, a number of rAAV gene therapy clinical trials have been 

conducted to treat PD (Polinski, 2016). While these therapies have not shown efficacy 

above current therapeutics and research is ongoing, rAAV delivery has been found to 

 

41 

 

be clinically safe for use in these patients. This precedent for clinical rAAV use in PD 

patients paves the way for potential LID-managing gene therapies. 

Overarching significance 

The phenomenon of LID has plagued the most effective symptomatic PD treatment 

since its clinical introduction. The burden of these symptoms cannot be overstated, as 

they can dramatically affect the lives of patients and their caregivers, while physicians 

are left with few effective options for managing LID. While it is of grave importance that 

the field continues to study disease-altering treatments for PD, patients and their 

families still currently—and likely will continue to—rely on the symptomatic relief granted 

by L-DOPA. Research focused on understanding LID with the end-goal to eradicate 

these symptoms is an important aspect of the PD field. In this dissertation, I aimed to 

contribute to this field by characterizing LID etiology from both a presynaptic and 

postsynaptic perspective. Using rAAV as a research tool, I first aimed to close critical 

knowledge gaps in the presynaptic hypothesis of LID by supplementing 5-HT neurons 

with the DA autoreceptor, giving them the ability to regulate DA release. Additionally, I 

sought to understand molecular postsynaptic changes in LID by showing that the 

orphan nuclear receptor Nurr1 is a novel driver of LID. Lastly, this dissertation will 

present evidence showing that maladaptive Nurr1 expression can promote striatal 

changes that influence LID development. Together, the Specific Aims of my project add 

valuable insight to the etiology of LID, and suggest potential therapeutic strategies for 

LID management. 

 

42 

 

Aim 1: Regulation of dopamine neurotransmission from 5-HTergic neurons by 

ectopic expression of the dopamine D2 autoreceptor blocks levodopa-induced 

dyskinesia  

Addressed in Chapter 2, in Aim 1, I virally expressed the DA autoreceptor, D2Rs in 

serotonin neurons of the dorsal raphe nucleus of parkinsonian rats. I challenged these 

rats with escalating doses of L-DOPA and DA receptor agonists to determine if D2Rs 

expression allowed for regulated release of DA from dorsal raphe neurons and, in turn, 

amelioration of LID. With this Aim, I sought to confirm the serotonin hypothesis of LID by 

showing that DA is abnormally released for dorsal raphe neurons, and that regulation of 

this release can mitigate LID. 

Aim 2: Modulating levels of Nurr1 expression in the parkinsonian rat striatum 

impacts LID severity and development 

Addressed in Chapter 3, in Aim 2 my goal was to characterize Nurr1 as a novel 

molecular factor in LID. I virally modulated Nurr1 levels in LID susceptible and resistant 

rats to determine if Nurr1 expression can exacerbate LID, and if its presence in MSNs is 

required for LID development. Additionally, I utilized specific D1 or D2 receptor agonists 

to determine if abnormal Nurr1 upregulation is induced by specific receptor stimulation, 

or if it requires activation of both the direct and indirect pathways. 

Aim 3: Ectopic overexpression of Nurr1 induces an LID-like state in striatal 

medium spiny neurons 

Addressed in Chapter 4, in Aim 3, I further determine the role of Nurr1 in LID 

development by showing how its ectopic overexpression alters striatal neurons. To 

 

43 

 

achieve this, we virally overexpressed Nurr1 in L-DOPA naïve animals and measured 

the electrophysiological properties of MSNs. I also determined if Nurr1 expression 

induces changes in the density and morphology of MSNs. This Aim allowed me to 

determine how Nurr1 influences the striatum in ways that may generate LID. 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

44 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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45 

 

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Chapter 2: Regulation of dopamine neurotransmission from serotonergic neurons 
by ectopic expression of the dopamine D2 autoreceptor blocks levodopa-induced 

dyskinesia  

Preface 

In this chapter, in vivo microdialysis and HPLC experiments were carried out by Dr. 

Christopher Bishop at Binghamton University. Electrophysiology recordings were 

performed by Dr. Anthony West at Rosalind Franklin University. All other experiments 

and analysis were performed at Michigan State University by Rhyomi C. Sellnow. 

Introduction 

The hallmark motor symptoms in Parkinson’s disease (PD) arise following substantial 

dopaminergic denervation within the striatum. Denervation results from the death of 

tyrosine hydroxylase (TH) expressing DA neurons of the substantia nigra pars 

compacta (SNc) as the disease progresses (Hoehn & Yahr, 1967; Kordower et al., 

2013). The lack of proper DA signaling to the striatum creates an imbalance of the basal 

ganglia motor circuit, thus, causing bradykinesia, rigidity, tremor, and gait problems 

characteristic of PD (Goldman & Postuma, 2014). Current effective treatment strategies, 

while not affecting disease progression, are aimed at treating these primary motor 

symptoms. Since the late 1960s, L-3,4-dihydroxyphenylalanine (levodopa or L-DOPA) 

has been used as a catecholamine replacement therapy to alleviate motor symptoms 

(Cotzias et al., 1967). L-DOPA remains the gold-standard pharmacological treatment for 

PD. 

While effective initially, the therapeutic window of L-DOPA narrows with the continuous 

loss of SNc neurons as the disease progresses, and higher doses are required to 

 

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maintain the anti-akinetic effects of L-DOPA. Moreover, chronic treatment with L-DOPA 

leads to the development of L-DOPA-induced dyskinesias (LID), a series of motor 

symptoms distinct and independent from the PD motor deficits being treated (reviewed 

in (Bastide et al., 2015)). These symptoms, comprised of painful and disrupting 

movements including hyperkinesia, dystonia, and chorea, occur in a majority of PD 

patients, developing in up to 50% of patients within 5 years of beginning treatment, and 

up to 90% of patients within 10 years (Ahlskog & Muenter, 2001; Manson et al., 2012).  

Studies show that LID development is a multifaceted process. However, it is largely 

agreed upon that the intermittent oral dosing of L-DOPA results in large variations in 

extracellular DA. Ultimately, this pulsatile release of DA, together with the denervated 

state of the striatum, results in maladaptive molecular and structural changes in the DA-

responsive neurons of the striatum, specifically medium spiny neurons (MSNs), leading 

to altered basal ganglia signaling (reviewed in (Cenci & Konradi, 2010)).  Given the 

extreme nigrostriatal denervation at the time of diagnosis(Kordower et al., 2013) the 

actual source of striatal DA following L-DOPA administration has been debated over the 

past half century. The leading hypothesis is that uptake of L-DOPA and its subsequent 

dysregulated metabolism to DA, and release by 5-HTergic 5-hydroxytryptamine (5-HT) 

neurons in the dorsal raphe nucleus (DRN) may be linked to dyskinesogenesis 

(reviewed in (De Deurwaerdère et al., 2016)). These neurons express aromatic L-amino 

acid decarboxylase (AADC) and can therefore convert L-DOPA into DA. However, DRN 

neurons do not express the regulatory mechanisms to monitor and control DA synthesis 

and release into the synapse, allowing for the unregulated release of DA into a 

hypersensitized striatum (Maeda et al., 1999). Additionally, 5-HTergic innervation of the 

 

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striatum increases substantially following DA denervation, allowing the majority of L-

DOPA to be metabolized and released as DA by 5-HTergic terminals (Maeda et al., 

2003; Maeda et al., 2005; Yamada et al., 2007; Rylander et al., 2010; Roussakis et al., 

2016)}. This overwhelming exposure of the DA-depleted striatal MSNs is hypothesized 

to be a large contributor to LID. In fact, studies in rats show that specifically lesioning 

the DRN (Carta et al., 2007; Eskow et al., 2009) or co-administering L-DOPA with 5-

HT1 receptor agonists (Bezard et al., 2013; Politis et al., 2014; Ghiglieri et al., 2016; 

Meadows et al., 2017), effectively reduces or eliminates LID. 

Normal regulation of DA signaling is mediated presynaptically primarily through the DA 

active transporter (DAT) and the DA autoreceptor. DAT directly regulates the levels of 

DA in the synapse by transporting synaptic DA back into the terminal. The dopamine 

autoreceptor (D2Rs) is an isoform of the D2 DA receptor (D2RL) missing 29 amino acids 

from the third intracellular loop (Dal Toso et al., 1989). D2Rs detects synaptic DA levels 

and regulates DA signaling in three ways, 1) by downregulating DA production through 

TH regulation, 2) regulation of reuptake through DAT, and 3) by directly inhibiting DA 

release (reviewed in (Ford, 2014)). Each of these modes of action are mediated through 

the inhibitory Gi alpha protein signaling pathways following D2Rs activation. 

These canonical G-protein-coupled receptor (GPCR) signaling pathways similarly inhibit 

5-HTergic signaling in DRN neurons through 5-HT1 autoreceptor activation (Harrington 

et al., 1988; Okada et al., 1989). Previous studies using 5-HT1 agonists show promising 

reductions in LID. Unfortunately, these drugs can negate the anti-parkinsonian 

therapeutic benefits of L-DOPA animal models, and in some cases worsen PD 

 

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symptoms in clinical trials (Kannari et al., 2002; Olanow et al., 2004; Iravani et al., 2006; 

Cheshire & Williams, 2012).  

While current evidence suggests a crucial role of 5-HTergic input and activity in LID, 

direct evidence of the abnormal dopaminergic neurotransmission and dysregulated DA 

release is lacking. In the present study, we sought to provide unequivocal evidence for 

the role of 5-HTergic DA neurotransmission in dyskinesogenesis and examine a novel 

therapeutic approach of modulating this non-physiological adaptation in the 

parkinsonian brain. To do this, we provided 5-HTergic neurons with DAergic regulatory 

mechanisms by ectopically expressing the D2Rs autoreceptor in the DRN of 

parkinsonian 6-OHDA lesioned rats, and evaluated the effect of ectopic D2Rs activity on 

L-DOPA efficacy, LID formation, response to DA agonists, and striatal DA release. 

Methods 

I. 

Adeno-associated virus production 

The D2Rs and GFP coding sequences were cloned into AAV genomes under the control 

of the chicken β-actin/cytomegalovirus (CBA/CMV) promoter for ubiquitous and robust 

expression. rAAV 2/9 was produced via triple-transfection of HEK 293T cells with the 

genome and helper plasmids. Virus was recovered from cells using freeze-thaw cycles, 

purified using an iodixanol gradient (Optiprep Density Gradient, Sigma-Aldrich, St. 

Louis, MO), followed by buffer exchange and concentration using concentrator columns 

(Orbital Biosciences, Topsfield, MA) as described previously (Benskey et al., 2016). The 

viral titer was determined using digital droplet PCR (ddPCR) and normalized to 1x1013 

vector genomes (vg)/ml using Balanced Salt Solution (Sigma-Aldrich, St. Louis, MO).  

 

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II. 

Animals and surgeries. 

Studies were performed using adult male Fischer F344 rats (200-220g upon arrival; 

Charles River, Wilmington, MA) in accordance with the guidelines of Michigan State 

University (AUF MSU06/16-093-00), Binghamton University (AUF# 779-17), and 

Rosalind Franklin University (AUF# A3279-01) Institutional Animal Care & Use 

Committees. Rats were housed two per cage prior to behavioral testing, and then 

separated and individually housed with environment enrichment during behavior studies 

for the remainder of the experiments. The animals were housed in a light-controlled (12 

hours light/dark cycle) and temperature-controlled (22±1 °C) room, and had free access 

to standard lab chow and water.  

All 6-OHDA and vector surgeries were performed under 2% isoflurane. After being 

anesthetized, animals were placed in a stereotaxic frame and were injected using a 

glass capillary needle fitted to a Hamilton syringe (Hamilton, Reno, NV) (Benskey & 

Manfredsson, 2016). Three weeks following lesion surgery, animals were tested for 

spontaneous forepaw use (cylinder test) to estimate lesion efficacy. Vector treatment 

groups were normalized using forepaw deficits in order to ensure equal lesions between 

the treatment groups.  

For lesion surgeries 5mg/ml 6-OHDA hydrobromide (Sigma-Aldrich, St. Louis, MO) was 

prepared in 0.2mg/ml ascorbic acid immediately prior to the injections. A subset of 

animals (n=7) destined for electrophysiological measures did not receive a 6-OHDA 

lesion. Animals received 2µl injections of 6-OHDA into the medial forebrain bundle 

(MFB) (from bregma: Anterior Posterior (AP) – 4.3mm, Medial Lateral (ML) +1.6mm, 

Dorsal Ventral (DV) -8.4mm from skull) and the SNc (from bregma: AP -4.8mm, ML 

 

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+1.7mm, DV -8.0mm from skull), for a total of 10µg 6-OHDA per site and 20µg per 

animal. The glass needle was lowered to the site and the injection started after 30 

seconds. 6-OHDA was injected at a rate of 0.5µl/minute. The needle was removed two 

minutes after the injection was finished and cleaned between each injection.  

Vector delivery was performed three weeks following the 6-OHDA lesion via stereotaxic 

delivery (Benskey & Manfredsson, 2016). Using the same procedure as described for 

the lesion surgeries, animals received a single 2µl injection of virus (AAV2/9-DRs, 

1x1013 vg/ml; AAV2/9-GFP, 1x1013 vg/ml) to the DRN (from bregma: AP -7.8, ML -3.1, 

DV -7.5 from skull). The stereotaxic arm was positioned in a 30° lateral angle in order to 

avoid the cerebral aqueduct. 

Parkinsonian and vector-injected animals used for in vivo microdialysis were shipped to 

Binghamton University two weeks following the vector surgeries. Following quarantine, 

rats were acclimated to the colony room and habituated to handling for one week. Rats 

were then tested for baseline forepaw adjusting steps. Thereafter, microdialysis 

cannulation surgery was performed under 2-3% isoflurane in oxygen with the tooth bar 

set to 5 mm below the interaural line. Five minutes before surgery and 24 hours after 

surgery rats received an injection of Buprinex (0.03 mg/kg, i.p.). A unilateral dorsal 

striatal-directed cannula (CMA 12 Elite; Stockholm, Sweden) was implanted ipsilateral 

to lesion (from bregma AP: 1.2 mm; ML: -2.8 mm; DV: -3.7 mm). The cannula was fixed 

in place by four jeweler’s screws, jet liquid, and dental acrylic (Lang Dental, Wheeling, 

IL).  Two weeks following cannulation surgery, rats underwent behavioral testing. 

Non-lesioned rats used for electrophysiological recordings of the DRN were shipped to 

Rosalind Franklin University two weeks following the vector surgeries and housed for an 

 

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additional 4-8 weeks prior to electrophysiological recordings. Burr holes (~1 mm in 

diameter) were drilled in the skull overlying the DRN. Prior to experimentation all 

animals were anesthetized with urethane (1.5 g/kg i.p.) and placed in a stereotaxic 

apparatus. Bipolar stimulating/recording electrodes were implanted in the frontal cortex 

and DRN on the right side using a micromanipulator (coordinates from Bregma: AP: 3.2 

mm; ML: 0.8 mm lateral; DV: 4.4 mm ventral (frontal cortex) or AP: 7.8 mm; ML 3.1 mm; 

DV: 7.5 mm with the manipulator angled 30 degrees toward Bregma) as previously 

described (Chakroborty et al., 2017). 

III. 

Abnormal involuntary movement (AIM) ratings and drug treatments. 

Animals were allowed to recover for three weeks following vector injections, and to 

allow for peak expression of the viral transgene (Reimsnider et al., 2007). After this 

time, L-DOPA treatment and abnormal involuntary movement (AIM) scale ratings 

began. As has been described previously, the AIM rating scale can be used to evaluate 

the severity of LID and has been adapted for animal use (Lundblad et al., 2002; Steece-

Collier et al., 2003). Briefly, AIMs were evaluated by scoring the level of dystonia of the 

limbs and body, hyperkinesia of the forelimbs, and orolingual movements. Each AIM is 

given two numerical scores—one indicating the intensity (0=absent, 1=mild, 

2=moderate, or 3=severe) and frequency (0=absent, 1=intermittently present for <50% 

of the observation period, 2=intermittently present for >50% of the observation period, 

or 3=uninterruptable and present through the entire rating period) (Maries et al., 2006) 

Each AIM was given a severity score by multiplying the intensity and frequency, and the 

total AIM score is a sum of all the behaviors severities. An animal is considered non-

dyskinetic with a score of ≤4, as non-dyskinetic parkinsonian rats can display low level 

 

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AIMs from exhibiting normal chewing behavior and a mild parkinsonian dystonic posture 

(Maries et al., 2006). 

Animals received subcutaneous injections of L-DOPA/benserazide (Sigma-Aldrich, St. 

Louis, MO) three times per week and were rated using the AIM scale in 25-minute 

intervals post-injection until all AIM behavior had subsided. L-DOPA doses ranged 

between 2mg/kg-12mg/kg. Benserazide doses (12 mg/kg) remained constant for all L-

DOPA injections. The same injection and rating paradigm was used for AIM evaluations 

with the non-selective DA agonist apomorphine (0.1mg/kg, R&D Systems, Minneapolis, 

MN), the D2/D3 receptor agonist quinpirole (0.2mg/kg, Sigma-Aldrich, St. Louis, MO) 

and the D1 receptor agonist SKF-81297 (0.8mg/kg, Sigma-Aldrich, St. Louis, MO). Peak 

AIM scores of DA agonists were determined based off the highest average AIM scores 

of control animals during the rating period. 

IV. 

Parkinsonian motor evaluation. 

To assess whether D2Rs viral therapy affects the anti-parkinsonian properties of L-

DOPA therapy, we evaluated parkinsonian motor behavior on and off L-DOPA using the 

cylinder task and the forepaw adjusting steps (FAS) test. Rats with significant lesions 

perform poorly on both these tests, with impairment to the forepaw contralateral to the 

lesion that is alleviated with L-DOPA treatment (Chang et al., 1999; Schallert, 2006). 

The cylinder task was conducted as previously reported (Manfredsson et al., 2007). 

Animals were placed in a clear Plexiglas cylinder on top of a light box for five to seven 

minutes while being recorded. Each animal was rated by counting ~20 weight-bearing 

forepaw placements on the cylinder (contralateral to the lesion, ipsilateral to the lesion, 

both) to determine the percentage use of the forepaw contralateral to the lesion, which 

 

85 

 

is derived by dividing the sum of contralateral touches and half of both forepaw touches 

by the total forepaw touches, and multiplying this number by 100. Trials were performed 

following the initial L-DOPA treatment (AIM evaluation) period, and tested either off L-

DOPA or, on the following day, 50 minutes after receiving a 6mg/kg L-DOPA injection 

(12mg/kg benserazide). 

The FAS test was performed as described previously (Meadows et al., 2017). Briefly, 

rats were restrained by an experimenter so that only one forepaw was free to touch the 

counter. Rats were then dragged laterally along a 90 cm distance over 10 sec while a 

trained rater blind to the experimental condition counted the number of steps. Data are 

represented as forehand percent intact, which are derived by taking the number of steps 

taken by the contralateral forehand and dividing it by the ipsilateral forehand, and then 

multiplying this number by 100. The test was performed over two days either off L-

DOPA or 60 minutes following an 8 mg/kg or 12 mg/kg L-DOPA injection. 

V. 

Tissue collection 

Two hours following the final L-DOPA administration, animals from the AIM 

experimentation were sacrificed via sodium pentobarbital overdose and intracardially 

perfused with Tyrode’s solution (137mM sodium chloride, 1.8mM calcium chloride 

dihydrate, .32mM sodium phosphate monobasic dihydrate, 5.5mM glucose, 11.9mM 

sodium bicarbonate, 2.7mM potassium chloride). Brains were rapidly removed and 

coronally hemisected, with the rostral portion of the left and right striatum dissected out 

and flash frozen in liquid nitrogen for biochemical analysis. The caudal portion of the 

brain was postfixed for 72 hours in 4% paraformaldehyde (PFA) in phosphate-buffered 

saline and then cryoprotected by saturation in 30% sucrose. Brains were frozen and 

 

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sectioned coronally at 40µm thickness using a sliding microtome into free floating 

sections and stored in cryoprotectant (30% ethylene glycol, 0.8mM sucrose in 0.5X tris-

buffered saline) until further use.  

VI. 

Immunohistochemistry 

A 1:6 series of free-floating tissue was stained immunohistochemically for TH (MAB318, 

MilliporeSigma, Burlington, MA), D2R (AB5084P, MilliporeSigma, Burlington, MA), or 

GFP (AB290, Abcam, Cambridge, United Kingdom) using methods previously reported 

(Benskey et al., 2018). Sections were washed in 1x Tris-buffered saline (TBS) with 

.25% Triton x-100, incubated in 0.3% H2O2 for 30 minutes, and rinsed and blocked in 

10% normal goat serum for 2 hours. Tissue was incubated in primary antibody (TH 

1:4000, D2R 1:1000, GFP 1:20,000) overnight at room temperature. After washing, 

tissue was incubated in secondary antibody (biotinylated horse anti-mouse IgG 1:500, 

BA-2001; Vector Laboratories, Burlingame, CA; biotinylated goat anti-rabbit IgG 1:500, 

AP132B, Millipore-Sigma, Burlington, MA) followed by the Vectastain ABC kit (Vector 

Laboratories, Burlingame, CA). Tissue staining was developed with 0.5 mg/ml 3,3’ 

diaminobenzidine (DAB, Sigma-Aldrich, St. Louis, MO) and 0.03% H2O2. Sections were 

mounted on glass slides, dehydrated, and coverslipped with Cytoseal (ThermoFisher, 

Waltham, MA).  

Tissue for immunofluorescence dual labeling of D2Rs or GFP with SERT (340-004, 

Synaptic Systems, Goettingen, Germany) were washed with 1x TBS with 0.25% Triton 

x-100, blocked in 10% normal goat serum for 2 hours, and probed with primary antibody 

overnight (D2Rs 1:1000, GFP 1:20,000, SERT 1:300). Tissue was incubated with 

 

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secondary antibody (A11008 1:500, A11076 1:500; ThermoFischer, Waltham, MA) in 

the dark for two hours, and washed in TBS before being mounted and coverslipped with 

Vectashield Hardset Antifade Mounting Medium (Vector Laboratories, Burlingame, CA).  

Images were taken on a Nikon Eclipse 90i microscope with a QICAM fast 1394 camera 

(fluorescence; QImaging, Surrey, British Columbia, Canada) or a Nikon D-1 camera 

(brightfield microscopy; Nikon, Tokyo, Japan). The figures were made using Photoshop 

7.0 (Adobe, San Jose, CA) with the brightness, sharpness, and saturation adjusted only 

as needed to best represent the staining as it is viewed directly under the microscope. 

VII. 

In vivo microdialysis. 

As outlined above, a separate cohort of parkinsonian rats treated with GFP or D2Rs 

were utilized for in vivo microdialysis. The night before the procedure, striatal probes 

(CMA 12 Elite; membrane length = 3 mm; 20,000 Dalton; Stockholm, Sweden) were 

inserted into the guide cannula so that they extended from bregma DV: -3.7 to -6.7 mm 

within the dorsal striatum. Rats underwent microdialysis at least two days following the 

last L-DOPA administration. During microdialysis, rats received intrastriatal infusion of 

filtered artificial cerebrospinal fluid (aCSF) (128mM NaCl, 2.5mM KCl, 1.3mM CaCl2, 

2.1mM MgCl2, 0.9mM NaH2PO4, 2.0mM Na2HPO4, and 1.0mM glucose, pH 7.4). 

Dialysate samples were collected every 20 minutes. Briefly, rats were habituated to 

microdialysis for 1 hour. Fifty minutes into the procedure, rats received a subcutaneous 

injection of L-DOPA vehicle, which consisted of 0.9% NaCl, and 0.1 % ascorbate. Rats 

then underwent baseline testing for one hour to determine baseline levels of 

monoamines prior to L-DOPA treatment. After that a new collection tube was used and 

ten minutes later rats received an injection of L-DOPA (12 mg/kg + 12 mg/kg 

 

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Benserazide, s.c.). Samples were taken every 20 minutes for 3 hours. Following the 

procedure, rats were removed from the microdialysis bowl and striatal probes were 

replaced with a dummy probe. At least two days after microdialysis, rats were sacrificed 

via rapid decapitation, the anterior striatum was taken for verification of cannula 

placement, the posterior striatum was taken for HPLC, and the hindbrain was placed in 

4% PFA for 3 days before being placed in 30% sucrose in phosphate-buffered saline 

(PBS). Brains were shipped on ice in a 50mL conical containing 30% sucrose in 0.1 M 

PBS to MSU. 

VIII.  High-performance liquid chromatography for monoamine tissue analysis 

Striatal tissue and in vivo microdialysis samples were analyzed using HPLC. Reverse-

phase HPLC was performed on striatal tissue samples as previously described 

(Kilpatrick et al., 1986; Meadows et al., 2017). Briefly, tissue samples were 

homogenized in ice-cold perchloric acid (0.1 M) with 1% ethanol and 0.02% 

ethylenediaminetetraacetic acid (EDTA). Homogenate was spun at 4°C for 45 min at 

14,000 g. Supernatant was removed and, using an ESA solvent delivery system (Model 

542; Chelmsford, MA, USA) ESA autoinjector (Model 582),  analyzed for levels of 

norepinephrine, 3,4-dihydroxyphenylacetic acid (DOPAC), DA, 5-hydroxyindoleacetic 

acid (5-HIAA), and 5-HT. Monoamines and metabolites were detected as a generated 

current as a function of time by EZCHROM ELITE software via a Scientific Software, Inc 

(SS240x) Module. Data are displayed as peaks for monoamines and metabolites, which 

are compared to a standard curve made from monoamine and metabolite samples of 

known concentrations ranging from 1e-6 to 1e-9. Values were then normalized to tissue 

 

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weight and lesion deficits are reported as percent depletion, which is equal to 100 (1 – 

M Lesion/ M Intact).  

Dialysate samples were analyzed via reverse-phase HPLC on an Eicom HTEC-500 

System (Amuza Inc., San Diego, CA). Briefly, 10 µL of each dialysate sample was 

analyzed for NE, DA, and 5-HT using an Eicompak CAX column maintained at 35°C 

with a flow rate of 250 µL/min. Mobile phase (75mM Ammonium acetate, 9.36mM acetic 

acid, 1.33mM EDTA, 0.94mM Methanol, 50mM sodium sulfate). Samples were 

compared to known concentrations of monoamines (100, 10, 1, 0.1, and 0.05 ng/µL 

dissolved in a potassium phosphate buffer (0.1mM potassium phosphate monobasic, 

0.1mM ethylenediaminetetraacetic acid, 0.02mM phosphoric acid), resulting in a final 

value of monoamine in ng/µL. 

IX. 

Total enumeration of TH+ neurons to assess lesion severity 

Lesion severity was determined using total enumeration of TH-positive neurons in three 

representative sections within the SNc identified by the presence and proximity to the 

medial terminal nucleus (MTN) of the accessory optic tract at levels equivalent to − 5.04 

mm, − 5.28 mm and − 5.52 mm relative to bregma according to our previously validated 

method (Gombash et al., 2014) method. Briefly, the intact and lesion SNc were 

quantified for all TH immunoreactive cells using a 20X objective and MicroBrightfield 

StereoInvestigator software (MicroBrightfield Bioscience, Williston, VT). The total 

number of TH cells on the intact and lesioned hemispheres were averaged, and lesion 

efficacy was derived by dividing the lesioned hemisphere average by the intact 

hemisphere average and multiplying that value by 100. 

 

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X. 

Electrophysiology 

Recording microelectrodes were manufactured from 2.0mm OD borosilicate glass 

capillary tubing and filled with sodium chloride (2M) solution. Electrode impedance was 

5-15MΩ. The signal to noise ratio for all recordings was > 4:1. The level of urethane 

anesthesia was periodically verified via the hind limb compression reflex and maintained 

using supplemental administration as previously described (Sammut et al., 2010; 

Padovan-Neto et al., 2015). Temperature was monitored using a rectal probe and 

maintained at 37C° using a heating pad (Vl-20F, Fintronics Inc, Orange, CT). Electrical 

stimuli (duration= 500µs, intensity=1000 µA) were generated using a Grass stimulator 

and delivered in single pulses (0.5Hz) while searching for cells (Padovan-Neto et al., 

2015). Once isolated, recordings consisted of basal (pre-drug), saline vehicle, and drug-

treatment-(see below) induced changes in spike activity recorded in a series of 3-minute 

duration epochs.  

All compounds and physiological 0.9% saline were prepared daily and administered 

intravenously (i.v.) through the lateral tail vein to enable rapid examination of potential 

acute effects of vehicle or drug on DRN neuronal activity. The selective 5-HT1A agonist 

8-OH-DPAT (5µg/kg, i.v.), the selective 5-HT1A antagonist WAY100635 (100µg/kg, 

i.v.), and the D2R agonist Quinpirole (500µg/kg, i.v.) were dissolved in vehicle and 

administered systemically to either BFP or D2Rs rats. DRN 5-HT neuron activity was 

recorded prior to and immediately following drug administration as described above. 

 

 

 

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XI. 

Statistical analysis 

Statistical analysis was performed using Statview (version 5.0) or in SPSS version 23 

with α set to 0.05. All graphs were created in GraphPad Prism version 7.0 (GraphPad 

Software, La Jolla, CA) or Excel (Microsoft, Redmond, WA). Lesion status was 

evaluated using unpaired, one-tailed t-tests. AIMs differences between vector groups 

were evaluated using a non-parametric Mann-Whitney U test, with p≤0.05 being 

considered statistically significant. Bonferroni post-hoc tests were employed when 

significant main effects were detected. Cylinder and FAS data for forehand and 

backhand stepping were submitted to a mixed model ANOVA with within-subjects 

factors of treatment (2: Baseline, L-DOPA) and between-subjects factors of vector 

(GFP, D2R). Overall percent intact values for FAS were determined by taking the 

overall number of right paw steps divided by the number of left paw steps and 

multiplying the quotient by 100. Similarly, overall percent intact values were analyzed 

via a repeated-measures ANOVA with within-subjects factor of treatment and between 

subjects factor of vector. Monoamine content (as determined by HPLC) was submitted 

to a mixed-model ANOVA with within-subjects factor of treatment (2: Vehicle, L-DOPA) 

and between-subjects factor of vector. Fisher’s least significant difference (LSD) post-

hocs and planned paired-samples t-tests were employed as appropriate to clarify 

significant effects. Additionally, independent-samples t-tests were employed to reveal 

effects of vector on the timing of DA, NE, and 5-HT efflux. HPLC values for striatal 

tissue were submitted to a mixed-model ANOVA with within-subjects factor of side and 

between-subjects factor of vector. Subsequently, since DA depletion did not vary as a 

function of vector, values for each monoamine for each side were collapsed across 

 

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treatments and compared via paired-samples t-tests. For electrophysiology 

experiments, the difference between the spontaneous and evoked electrophysiological 

activity of identified DRN-5-HT neurons across groups was determined and served as 

the dependent variable for our analyses. A two-way repeated measures ANOVA (GFP 

vs. gene therapy (ectopic expression of the DA D2 AR in 5-HT DR neurons)) x 2 

(vehicle vs. drug treatment) with α set to 0.05 and all “n’s” adequately powered for 

electrophysiological studies was conducted using Sigma Stat software (San Jose, CA), 

and the potential two-way interaction effect was examined to determine how treatment 

effects differ as a function of drug treatment or gene therapy (Padovan-Neto et al., 

2015). 

Results 

I. 

Validation of lesion and transgene expression 

In order to assess if exogenous expression of D2Rs in the DRN could inhibit LID 

development or decrease LID severity, adult Fischer rats were rendered parkinsonian 

with 6-OHDA delivered to the SNc and MFB. Because LID is dependent on the severity 

of the lesion (Winkler et al., 2002) we validated post mortem that sufficient nigrostriatal 

denervation was achieved. Immunohistochemistry of the striatum (Figure 2.1B) and the 

SNc (Figure 2.1C) showed a near complete ablation of TH immunoreactivity with no 

difference in the number of SNc DA neurons between groups (Figure 2.1D; 

GFP=1.29%+0.29% remaining; D2Rs=1.45%+0.41% remaining; t(9)=0.31, p>0.05). 

Similarly, HPLC analysis of DA and DOPAC levels from striatal tissue from rats 

employed in the microdialysis experiment confirmed that all animals displayed a near  

 

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Figure 2.1 Experimental design and model validation 

 

 

 

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Figure 2.1 (cont’d) 

(A) Experimental timeline showing LID-inducing L-DOPA paradigm, motor behavior 
evaluations, and DA agonist treatments. AIM score ratings were taken at each injection 
where indicated. (B and C) Representative TH immunoreactivity in the striatum (B) and 
substantia nigra (C) showing complete loss of TH-positive neurons and projections 
following 6-OHDA lesions (scale bar=1mm). (D) Total enumeration of remaining TH 
neurons in the substantia nigra. (E and F) IHC for the D2 receptor (E) or GFP (F) in the 
DRN, showing successful targeting of the structure and robust expression of the 
transgene (scale bar=1mm). Cell bodies were efficiently transduced in the DRN (E and 
F, bottom insets, scale bar=50um) and could be seen filling projection fibers in the 
peduncles (E and F, top insets, scale bar=50um). (K-P) Dual labeling transgene 
expression and SERT in rAAV-D2Rs (K-M) and rAAV-GFP (N-P) animals. Transgene 
expression was visualized with D2Rs (L) or GFP (O) staining, and serotonin fiber and 
cell integrity were confirmed by staining for SERT (M and P). No adverse effects on 
SERT fibers were observed following vector transduction with either construct (K and N) 
(K-P scale bar=100um). 

 

 

 

 

 

 

 

 

 

 

95 

 

complete reduction in striatal DA levels in the lesioned hemisphere as compared to the 

intact hemisphere (DOPAC=18.11±6.68% of intact hemisphere, DA=3.48± 1.36% of 

intact hemisphere) (Table 2.1). There was no difference in striatal DA depletion between 

groups (DOPAC t(13)=0.73, p>0.05, DA t(13)=17.21, p>0.05).  

After a three-week recovery period, rAAV 2/9 expressing either D2Rs or GFP was 

delivered by stereotaxic injection into the DRN. Following sacrifice, transduction was 

confirmed with immunohistochemistry (IHC) of D2Rs or GFP (Figure 2.1E-J). Significant 

transgene expression was observed in the soma (D2Rs Figure 2.1F; GFP Figure 2.1I) of 

the DRN as well as in DRN efferent projections (D2Rs Figure 2.1E; GFP Figure 2.1H). 

The two transgenes exhibited a slightly different subcellular expression pattern where 

more GFP expression was seen in projections as compared to D2Rs expression. It is 

unclear if this is due to increased 5-HT innervation in dyskinetic (i.e. GFP treated) 

animals (Maeda et al., 2003), or due to a different distribution pattern specific to the 

transgenes. The latter is to be expected as GFP is a soluble protein and typically fills 

the entire neuron. Vector transduction and transgene expression of both constructs did 

not adversely affect SERT expression in the DRN (Figure 2.1K-P). Four animals (rAAV-

D2Rs: n=2, rAAV-GFP: n=2) that lacked sufficient vector expression in the DRN were 

removed from the analysis, leaving a total of n=15 rats included in the analysis (rAAV-

D2Rs: n=7, rAAV-GFP: n=8). 

II. 

D2Rs delivery to the dorsal raphe eliminates LID 

After a 4-week recovery period to allow for optimal transgene expression (Reimsnider et 

al., 2007), animals were treated with L-DOPA and rated for AIMs (see Figure 2.1A for  

 

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______________________________________________________________________ 

Monoamine   

Lesioned Side 

NE 

 

DOPAC 

DA 

 

5-HIAA 

5-HT   

 

 

 

 

 

14.89 ±5.77*  

311.39 ± 96.00* 

378.18 ± 144.94* 

532.68 ± 61.51 

283.07 ± 45.73* 

 

 

 

 

 

 

Intact Side 

 

% Intact 

41.93 ±2.19   

35.53 ±13.33 

2070.7± 207.05 

18.107 ± 6.68 

12027 ±1175.20 

3.48 ± 1.36 

485.18 ±64.70 

136.71 ± 26.74 

561.89 ± 105.76 

64.46 ± 12.35 

Table 2.1 Concentrations of monoamines in lesioned versus intact hemisphere 

These data represent the picograms per microliter of monoamines and metabolites for 
striatal tissue taken from animals used in microdialysis experiments. Data verify 
successful DA lesion. There were no significant differences in monoamine 
concentrations in rAAV-GFP versus rAAV-D2Rs rats. Thus, data are collapsed across 
groups. (*=p<0.05 lesioned vs. intact) 

 

 

 

 

 

 

 

 

 

 

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experimental timeline). With L-DOPA, rAAV-D2Rs treated animals did not show 

significant LID at the typical peak-dose time point (75 minutes post-L-DOPA delivery) 

LID (defined as an AIM score ≥4) (Zhang et al., 2013) at any dose level (2 mg/kg 

AIMs=0±0, 4mg/kg AIMs=0.14±0.14, 6mg/kg AIMs=0±0, 8mg/kg day 8 AIMs=0.29±0.18, 

8mg/kg day 10 AIMs=0.29±0.29, 8mg/kg day 12 AIMs=0.14±0.14, 8mg/kg day 15 

AIMs=0.29±0.29, 12mg/kg day 17 AIMs=0.14±0.14, 12mg/kg day 19 AIMs=0.36±0.18) 

(Figure 2.2A, Figure A.2.1). rAAV-GFP controls began to show mild-to-moderate peak-

dose AIMS with a moderate L-DOPA dose (6mg/kg peak dose AIMs=3±1.43), which 

increased to more significant levels of severity with higher doses of L-DOPA (8mg/kg 

peak dose AIMs: day 8=4.5±2.29, day 10=5.5±2.36, day 12=6.88±2.72, day 

15=5.25±1.76; 12mg/kg peak dose AIMs: day 17=8.69±2.06, day 19=9.44±1.93) (Figure 

2.2A, Figure A.2.1). When compared to rAAV-D2Rs subjects, rAAV-GFP animals 

showed significantly higher total peak dose AIM scores per session starting with 8mg/kg 

doses (day 12 rAAV-D2Rs (Md=0), rAAV-GFP (Md=4), U=12, p<0.05; day 15 rAAV-

D2Rs (Md=0), rAAV-GFP (Md=4.5), U=12, p<0.05; Mann-Whitney U test) (Figure 2.2B). 

This difference was maintained with the high dose of L-DOPA (12mg/kg day 17 rAAV-

D2Rs (Md=0), rAAV-GFP (Md=8.5), U=0.5, p<0.001; day 19 rAAV-D2Rs (Md=0), rAAV-

GFP (Md=9.25), U=0, p<0.001) (Figure 2.2C-F, Figure A.2.1). Taken together, these 

data show that D2Rs expression in the DRN completely blocks the development of LID 

in parkinsonian rats, even with administration of high L-DOPA doses. 

III. 

D2Rs does not affect parkinsonian motor behavior 

To assess if rAAV-D2Rs treatment alters the anti-akinetic properties of L-DOPA, we 

examined motor behavior using the cylinder task (Figure 2.3A). There were no  

 

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Figure 2.2 DRN D2Rs expression blocks LID development 

(A) rAAV-D2Rs-injected animals did not develop LID over the course of 19 days of treatment with increasing doses of L-
DOPA, where rAAV-GFP controls developed AIMs. (B) The total AIM score for each rating session was significantly 
different between groups starting on treatment day 8 with 8m/kg L-DOPA. D2Rs animals remained LID-. (C-F) AIM scores 
from days 12, 15, 17, and 19 showing LID severity in 25-minute intervals. GFP animals displayed a typical dyskinetic 
response to chronic L-DOPA treatment, with increasing AIM severity seen at higher doses. The peak-dose severity (AIM 
score at 75 minutes-post L-DOPA injection) was significantly higher in GFP animals than D2Rs animals in the last four 
days of the L-DOPA paradigm. (*=p≤0.05, **=p≤0.01, ***=p≤0.001

 

 

99 

 

 

 

Figure 2.3 rAAV-D2Rs does not impact L-DOPA efficacy 

(A) Cylinder task was performed three weeks post-lesion (pre-vector), off L-DOPA 
(post-vector, post-L-DOPA paradigm) and on L-DOPA (6mg/kg, 50 minutes post 
injection). Both vector groups showed significant impairment following lesion and vector 
delivery, which was recovered with L-DOPA treatment. There were no significant 
differences between vector groups. (B) A second cohort received the same lesions and 
vector deliveries and motor function was evaluated using the adjusting steps tests. 
While all animals in both groups showed significant impairment on the test without L-
DOPA, motor function was restored while on drug (8mg/kg and 12mg/kg). There were 
no differences between vector groups. 

 

 

 

 

 

 

 

 

100 

 

significant differences between the rAAV treatment groups without L-DOPA (F-

(1,13)=0.008, p>0.05). Pre-vector scores for both groups and post-vector scores for rAAV-

GFP showed a marked decrease from normal contralateral forepaw use, indicating 

significant impairment. rAAV-D2Rs animals post-vector showed a trend towards more 

balanced forepaw use, but the differences were not significant. Both groups showed a 

significant increase from baseline increase towards balanced contralateral forepaw use 

while on L-DOPA (6mg/kg) (F(2,26)=7.11, p<0.01). No significant differences were seen 

in impairment or improvement between vector treatment groups (F(2,26)=0.72, p>0.05). A 

second separate cohort of animals underwent the adjusting steps test, both off and on 

(8-12mg/kg) L-DOPA. Both vector treatment groups showed significantly impaired 

adjusting steps without L-DOPA (GFP baseline=3.57%±0.49% intact stepping; D2Rs 

baseline=6.33%±3.02% intact stepping; t(11)=0.98, p>0.05), however, this deficit was 

rescued with the administration of both doses of L-DOPA (Figure 2.3B; F(2,22)=9, 

p<0.01). As with the cylinder task, no differences in impairment nor improvement while 

on L-DOPA were seen between groups (F(2,22)=0.24, p>0.05). Together, this suggests 

that ectopic D2Rs expression in the DRN does not interfere with the anti-parkinsonian 

motor benefits of L-DOPA. 

IV. 

Dopamine receptor agonists do not induce significant AIMs in L-DOPA-

primed rAAV-D2Rs rats 

Next, we examined whether dopamine agonists could induce AIMs in the rAAV-D2Rs 

treated rats that had remained resistant to LID after the L-DOPA dosing paradigm. 

Animals received three repeated doses each of a non-selective DA agonist 

(apomorphine, 0.1mg/kg), a D2/3-specific receptor agonist (quinpirole, 0.2mg/kg), and a 

 

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D1-specific receptor agonist (SKF-81297, 0.8mg/kg) and were evaluated for AIM 

severity (see timeline in Figure 2.1A). These DA agonists can induce AIMs in both L-

DOPA-primed and unprimed parkinsonian animals (Boraud et al., 2001; Boyce et al., 

2001; Chondrogiorgi et al., 2014). We hypothesized that directly activating the DA 

receptors with an agonist would bypass any protective effects of the rAAV-D2Rs 

treatment in normalizing aberrant DA release, as these agonists do not require 

processing and release by DAergic or 5-HTergic terminals, and therefore would not be 

affected by presynaptic regulatory mechanisms. They also allowed us to compare DA 

receptor supersensitivity status between treatment groups. Interestingly, rAAV-D2Rs 

animals challenged with both apomorphine and quinpirole did not show significant peak 

AIMs (rAAV-D2Rs apomorphine third treatment 25-minute AIMS=1.86±1.32; quinpirole 

third treatment 25-minute AIMs=-1.57±0.66), while rAAV-GFP animals continued to 

express moderate-to-severe AIM behaviors (rAAV-GFP apomorphine third treatment 

25-minute AIMS=10.75±2.10; quinpirole third treatment 25 minute AIMs=-11.81±2.45) 

(Figure 2.4A-F). rAAV-D2Rs animals exhibited significantly lower peak-dose AIMs with 

both apomorphine and quinpirole treatment compared to rAAV-GFP animals 

(apomorphine third treatment 25-minute AIMs rAAV-D2Rs (Md=0), rAAV-GFP 

(Md=12.75), U=4.5, p<0.01; quinpirole third treatment 25-minute AIMs rAAV-D2Rs 

(Md=1.5), rAAV-GFP (Md=13), U=3.5, p<0.01). Treatment with SKF-81297 did induce 

mild-to-moderate AIM scores in rAAV-D2Rs treated animals (third treatment 50-minute 

AIMs=3.92±0.73), but these scores remained significantly less severe than their control 

counterparts (third treatment 50-minute AIMs rAAV-D2Rs (Md=3.5), rAAV-GFP (Md=13), 

U=2, p<0.001) (Figure 2.4G-I).

 

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Figure 2.4 D2Rs-injected animals do not develop severe AIMs with DA agonist treatment 

Animals were treated three times each with apomorphine (A-C) quinpirole (D-F) or SKF-81297 (G-I). (A-C) rAAV-D2Rs-
injected animals remained AIM resistant with .1m/kg pan-DA agonist apomorphine treatment, while rAAV-GFP animals 
continued to exhibit dyskinetic behaviors. (D-F) .2mg/kg quinpirole (D2 agonist) did not elicit AIMs in rAAV-D2Rs animals, 
where rAAV-GFP animals continued to exhibit moderate to severe AIMs. (G-I) rAAV-D2Rs began to show mild-to-
moderate AIMs with .8mg/kg of the D1 agonist SKF-81297 treatments, but remained significantly less severe than their 
rAAV-GFP counterparts. (*=p≤0.05, **=p≤.01, ***=p≤.001) 

 

 

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V. 

D2Rs expression in the dorsal raphe reduces striatal dopamine efflux 

following L-DOPA delivery 

In order to determine if ectopic D2Rs expression in the DRN was inhibiting LID by 

moderating DA release from 5-HTergic neurons, we generated a second cohort of 

animals in order to perform in vivo microdialysis (rAAV-D2Rs n=6, rAAV-GFP n=7). 

Animals were lesioned and received vector in an identical manner to the first cohort, 

and subsequently treated with L-DOPA to establish LID. In order to determine 

differences between vector groups in the absence of L-DOPA, striatal dialysate was 

analyzed via HPLC and data for monoamine content were examined using a 2 (vector) 

x 2 (treatment) mixed-model ANOVA. Overall, DA values were dependent upon 

treatment, F(1,11)=124.35, p<0.05, and vector, F(1,11)=7.39, p<0.05. Planned pairwise 

comparisons revealed that L-DOPA treatment increased striatal DA efflux in both 

groups. However, rats with rAAV-D2R had lower levels of DA efflux than rats with the 

GFP vector (p<0.05) (Figure 2.5A). Finally, there was a vector-by-treatment interaction, 

F(1,11)=6.66, p<0.05, such that rats with the D2R vector had lower levels of DA efflux 

than rats with the GFP vector, but only after L-DOPA treatment. There was no effect of 

vector or treatment on striatal 5-HT efflux (Figure 2.5B; F(11,121)=0.867, p>.05). DA 

values for each time point were also submitted to paired-samples t-tests in order to 

examine the effect of vector on DA efflux at each time point during microdialysis. There 

were significant differences between vector groups 60 (t(5)=3.42, p<0.05), 80 (t(5)=2.77, 

p<0.05), 100 (t(5)=4.68, p<0.01), and 120 (t(5)=2.59, p<0.05) minutes after L-DOPA 

administration, showing that rats with the GFP vector had elevated striatal DA efflux as 

compared to the rats with the D2R vector. This is the first direct evidence showing that  

 

104 

 

Figure 2.5 DRN D2Rs reduces striatal efflux of DA 

(A and B) In vivo microdialysis of rAAV-D2Rs and rAAV-GFP animals in twenty-minute 
intervals following L-DOPA injection (12 mg/kg + 12 mg/kg benserazide). (A) rAAV-
D2Rs animals showed significantly decreased DA efflux in the striatum 60-120 minutes 
following injection. (B) No changes in serotonin efflux in the striatum between vector 
groups was observed following L-DOPA injection. (*=p≤0.05, **=p≤.01, ***=p≤.001) 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

105 

 

mishandled DA by DRN neurons can be regulated exogenously, and this regulation 

reduces DA release in the striatum, thus suppressing LID. 

VI. 

D2Rs expression inhibits 5-HT neuron activity 

In order to demonstrate that the ectopically expressed D2Rs have the capacity to inhibit 

the activity of 5-HT neurons, we performed electrophysiological recordings on a 

separate cohort of non-parkinsonian, L-DOPA naïve animals. Animals received a 

stereotaxic delivery of either vector as described above, and 4-12 weeks later we 

performed in vivo single-unit extracellular recordings of DRN neurons. Putative 5-HT 

neurons were identified based initially on their firing characteristics (e.g., long-duration 

action potentials, regular firing pattern interrupted with burst activity).  Next, neurons 

were identified as 5-HTergic based on well characterized responses to systemic 

administration (i.v.) of 5-HT1A receptor agonist (8-OH-DPAT) and reversal with an 

antagonist (WAY-100635) which restored 5-HT neuron firing to that of baseline (Figure 

2.6B-D) (Hajos et al., 2007; Celada et al., 2013).  Figure 2.6A shows typical traces of 5-

HT and non-5HT DR neurons. Importantly, electrophysiologically identified 5-HT 

neurons recorded in the dorsal raphe of rats transduced with rAAV expressing BFP or 

D2Rs responded similarly to systemic administration of vehicle, 5-HT1A receptor 

agonist, and reversal of 5-HT1A inhibition by 5-HT1A antagonism.  Moreover, 5-HT cells 

recorded in AAV-D2Rs injected rats administered the D2 agonist quinpirole exhibited 

clear inhibitory effects, whereas responses to quinpirole were variable and sometimes 

excitatory in BFP controls (Figure 2.6E-F). These data show that ectopic expression of 

D2Rs in confirmed 5-HT neurons can act as a functional autoreceptor and inhibit 

impulse activity in 5-HTergic neurons. 

 

106 

 

 

Figure 2.6 Ectopic DRN D2Rs expression reduces 5-HT neuronal firing 

A) Top: Traces show typical single-unit recordings of isolated DRN 5-HT neurons (5-
HT+) (left) and non-5-HTergic (5-HT-) neurons (right). Middle/Bottom: 5-HT neurons 
often exhibit burst firing with short inter-spike intervals as well as regular spiking.  B) 
Systemic administration of the selective 5-HT1A agonist 8-OH-DPAT (1 µg/kg, i.v.), but 
not saline (0.9%) vehicle, suppressed the spontaneous firing of a DRN neuron 
exhibiting spike characteristic of a 5-HT cell. A return to baseline firing was observed 
after the local application of WAY-100635 ((100 µg/kg, i.v.), vertical blue bars). C-D) 
firing rate distributions of DRN neurons recorded in BFP and D2Rs expressing rats 
before and after the application of saline, 8-OH-DPAT and WAY-100635. Putative 5-HT 
neurons in both groups exhibited similar inhibitory responses to 5-HT1AR agonist and 
reversal of inhibition by 5-HT1AR antagonism.  E) Firing rate histograms showing the 
effects of the D2R agonist quinpirole (500 µg/kg, i.v.) on 5-HT neurons recorded in BFP 
(top) or AAV-D2AR (bottom) injected rats. Control neurons that responded to Quin 
increased their firing activity to varying degrees, whereas the majority of 5-HT cells 
recorded in AAV-D2AR injected rats were inhibited. F) Cumulative electrophysiological 
data showing the mean ± S.E.M. firing rates of 5-HT DRN cells transfected with BFP or 
D2AR prior to, and after Quin administration.  

 

 

 

107 

 

Discussion 

In this study, we used rAAV to ectopically express the dopamine autoreceptor (D2Rs) in 

order to equip DRN 5-HT neurons with a DA-mediated autoregulatory mechanism. 

Dysregulated DA release (“false neurotransmission”) by 5-HT neurons has been heavily 

implicated as a key contributor to LID development (Maeda et al., 1999; Bibbiani et al., 

2001; Maeda et al., 2003; Maeda et al., 2005; Carta et al., 2007; Eskow et al., 2009; 

Muñoz et al., 2009) . While a number of studies have supported this theory, to date, no 

direct evidence has been presented that shows that DRN neurons can release DA in 

the striatum in a way that ultimately effects LID. In the present study, our data 

demonstrate that providing DA-dependent autoregulation in 5-HT neurons can prevent 

LID formation by reducing striatal DA efflux, thus, providing unambiguous evidence that 

5-HT neurons play a central role in DA-dependent symptomology.  

A wealth of preclinical and clinical studies have shaped the serotonin hypothesis of LID, 

which suggest that DA processing and release from striatal 5-HT terminals is involved in 

AIM presentation. Studies ablating DRN neurons or dampening their activity with 

serotonin autoreceptor agonists are able to reduce or eliminate LID, the hypothesized 

reasoning being that reducing 5-HTergic neuronal activity leads to a reduction in DA 

release from DRN terminals (Iravani et al., 2006; Carta et al., 2007; Eskow et al., 2007). 

Although the mechanism by which 5-HT neurons process L-DOPA and release DA is 

not fully established, it is well established that the machinery to do so is present in 5-

HTergic neurons (Arai et al., 1995; Tanaka et al., 1999; Gantz et al., 2015).  

 

 

108 

 

Dopamine autoregulation in the dorsal raphe blocks 5-HT neuron activity and LID 

development 

In order to better delineate the role of DRN 5-HT neurons in dyskinesogenesis, we 

argued that expressing DA regulatory factors in 5-HT neurons would decrease LID 

severity. As 5-HT autoreceptors share a canonical signaling cascade with the D2-type 

DA autoreceptors—both are inhibitory GPCRs that reduce cellular cAMP to inhibit 

neuronal signaling (Harrington et al., 1988; Neve et al., 2004)—we hypothesized that 

ectopically expressing the DA autoreceptor D2Rs in DRN neurons would serve such a 

function. In support of this theory, recent work demonstrated that expression of D2Rs in 

the DRN of naïve mice results in a reduction of 5-HT-mediated currents (Gantz et al., 

2015). Indeed, in this work we found that direct recordings of single 5-HT neurons in the 

DRN expressing the autoreceptor provide evidence that ectopic D2Rs expression can 

provide an inhibitory neuromodulatory effect in 5-HT neurons, characterized by a strong 

decrease in spontaneous firing following systemic DA D2Rs agonist administration.  

Accordingly, we utilized rAAV targeted to the DRN in hemiparkinsonian rats that 

subsequently received a LID-inducing dosing regimen of L-DOPA. As hypothesized, we 

found that DRN expression of D2Rs provided complete protection against the 

development of LID, an effect that also persisted at high levels of L-DOPA. Importantly, 

there was no difference in the extent of nigrostriatal denervation between the groups, 

nor was there any demonstrable toxicity due to either treatment in the DRN. Thus, 

prevention of LID was explicitly due to expression of D2Rs in the DRN.  

 

 

 

109 

 

Dopamine efflux into the striatum is reduced with dorsal raphe D2Rs expression 

Although there is a wealth of research supporting the 5-HTergic input in LID 

development (Nicholson & Brotchie, 2002; Scholtissen et al., 2006; Bastide et al., 

2015), direct evidence showing that the contribution is due to an increase in DA release 

from these neurons is limited. Using in vivo microdialysis, we determined that L-DOPA-

mediated DA efflux into the striatum can be significantly reduced by negatively 

regulating DRN serotonin neurons. We did not see a decrease in serotonin efflux in the 

striatum in rAAV-D2Rs animals, suggesting serotonin release was not affected. 

However, one explanation for this observation may also be the lack of direct stimulation 

of the 5-HTergic system during our L-DOPA treatment, in which case one would not 

expect to see changes in 5-HT signaling. Our findings demonstrate that D2Rs can 

induce DAergic regulation in 5-HT neurons, by ‘hijacking’ endogenous signaling 

cascades and reducing neuronal activity following L-DOPA administration. Our data 

indicate that exogenously provided D2Rs can couple with Gαi subunits in DRN neurons, 

and induce the appropriate signaling cascades to reduce neuronal activity in the 

presence of DA.  In conjunction with the LID studies utilizing serotonin agonists, our 

data confirm that reducing the activity of the serotonin system can dramatically inhibit 

LID. Additionally, this is the first evidence showing that 5-HTergic neurons can control 

DA release when supplied exogenously with a single DA-regulatory factor. Our study 

delivered rAAV-D2Rs prior to beginning L-DOPA treatment. In future studies, showing 

that D2Rs delivery post-L-DOPA and post-AIM establishment would be imperative to 

determine if the addition of regulation can reduce or reverse established LID. 

 

110 

 

Ectopic D2Rs expression in the dorsal raphe blocks L-DOPA priming in the 

striatum 

We hypothesized that the protective effects of D2Rs treatment would be negated with 

DA-receptor agonist treatments which directly affect the postsynaptic target, striatal 

medium spiny neurons (MSNs). As the DA regulation of D2Rs is a presynaptic 

mechanism, treatment with postsynaptic receptor agonists—which can induce 

dyskinesia in animal models and patients (Gomez-Mancilla & Bédard, 1992; Boraud et 

al., 2001; Boyce et al., 2001; Chondrogiorgi et al., 2014)—should induce AIMs in rAAV-

D2Rs-treated animals resistant to LID. Interestingly, treatment with D1-, D2-specific, or 

pan-DA agonists did not induce severe AIMs in rAAV-D2Rs animals, where only a rather 

modest dyskinetic response was seen with the D1 agonist SKF-81297. However, it is 

well established that LID development is preceded by a “priming-period” consisting of 

discontinuous striatal DA tone, resulting in morphological and molecular changes to the 

MSNs (Morelli et al., 1989; Pinna et al., 1997; Carta et al., 2003; Simola et al., 2009; 

Steece-Collier et al., 2009; Cenci & Konradi, 2010; Zhang et al., 2013). Our data 

therefore indicates that D2Rs-treated animals were blocked from the preceding L-DOPA 

priming, suggesting that D2Rs in the DRN can counteract this discontinuous DA release 

to sufficiently block the LID-inducing priming of striatal MSNs. The fact that at the end of 

the treatment we began to observe a mild-to-moderate increase in AIM presentation in 

rAAV-D2Rs animals with DA agonist treatment as compared to L-DOPA, suggest that 

these animals were in the early stages of priming, a phenomenon that is to be expected 

as direct MSN DA receptor activation would not be mitigated by DRN D2Rs expression.   

 

111 

 

Inhibition of dorsal raphe 5-HTergic neurons does not mitigate the anti-

parkinsonian benefits of L-DOPA 

It was important to confirm that D2Rs expression in the DRN does not negatively affect 

the therapeutic efficacy of L-DOPA in our PD model, as this has been an issue with 5-

HT agonist-type therapies in clinical trials for LID (Kannari et al., 2002; Olanow et al., 

2004; Cheshire & Williams, 2012), and an imperative problem to mitigate for potential 

future therapies. We saw no changes in motor improvement between control and D2Rs 

animals of two separate cohorts using two different motor tests. Both tests 

demonstrated a significant improvement in motor function with the administration of L-

DOPA, reflecting recovery back to a pre-lesion state. This shows that D2Rs activity in 5-

HTergic terminals of the striatum (or elsewhere) does not interfere with the 

pharmacological benefits of L-DOPA, and implicates D2Rs therapy as a potential 

treatment option for LID. It is important to note that while many preclinical studies using 

5-HT agonists did not show an effect on L-DOPA-induced motor improvement, these 

results have not translated clinically. While multiple trials have used a variety of 5-HT 

agonists and seen reductions in AIM scores, many of these compounds contribute to 

worsening of parkinsonian symptoms and OFF L-DOPA periods, or have been 

abandoned due to lack of efficacy (reviewed in (Cheshire & Williams, 2012)). The 

discrepancy between our D2Rs approach and the use of agonists is unclear given that 

these two approaches conceivably evoke the same mechanism. Nevertheless, 5-HT1 

compounds may produce their own side effects (Lindenbach et al., 2015). Second, their 

effects are dependent on an exogenously administered compound and dose as 

opposed to a gene therapy approach. Nevertheless, further studies would be required to 

 

112 

 

determine if D2Rs expression in the raphe is successful in other preclinical models of 

LID. 

Pharmacological manipulations of 5-HT neurons in the treatment of LID, although 

successful pre-clinically, have not been fully translated.  The transient nature of the anti-

dyskinetic effect of currently available 5-HT approaches may be due to pharmacologic 

limitations of these drugs, including lack of specificity and potency for the specific 

receptor. Moreover, timing and comparative pharmacodynamics with L-DOPA delivery 

may be preventative (Mazzucchi et al., 2015). Because of this, a genetic approach in 

the form of continuous 5-HT inhibition should bypass such pharmacological limitations 

and provide meaningful and lasting protection against LID. Moreover, the finding that 

D2Rs gene therapy does not interfere with L-DOPA efficacy in our rat model provides 

promise for such an approach. Of course, the DR innervates a large part of the brain, 

providing many crucial functions, and the D2Rs therapy undertaken here does not 

distinguish between various projections. Thus, understanding any off-target effects from 

DA-mediated regulation of 5-HT neurons remains one important caveat that requires 

further research. Further studies to better understand potential side effects and the 

effect on the 5-HTergic system may be warranted. 

Our findings shed new light on previous work demonstrating changes in 5-HT 

innervation occurring concomitant with nigrostriatal denervation and PD. Both 5-HT 

hyperinnervation (Politis et al., 2010; Rylander et al., 2010; Bedard et al., 2011) as well 

as a decrease in 5-HT terminals (Scatton et al., 1983; Kim et al., 2003; Guttman et al., 

2007; Kish et al., 2008) has been documented in human disease. Although the cause of 

these divergent findings is unknown, it is highly likely that 5-HT neurons play an 

 

113 

 

important role in PD symptomology and, as our findings would suggest, in LID. As 

nigrostriatal denervation in human PD is near complete at the time of diagnosis 

(Kordower et al., 2013) it is conceivable to speculate that changes in 5-HT innervation 

and function—and the capacity of these neurons to release DA—is a crucial component 

to dyskinesogenesis. To that end, understanding both the mechanisms of how 5-HT 

neurons process and release DA, and the underlying etiology of presynaptic 5-HT 

changes are important components as we begin to understand LID etiology and PD 

nonmotor symptoms, and represents a new therapeutic modality.  

In conclusion, the current study shows that DA release from DRN 5-HT neurons can be 

regulated with ectopic expression of D2Rs, altering the release activity of these neurons. 

These data add important evidence to the current understanding of LID and serve as to 

confirm the serotonin hypothesis in LID, showing that directly regulating serotonin 

neurons activity can inhibit LID development.  

 

 

 

 

 

 

 

 

 

114 

 

 

 

 

 

 

 

 

 

 

 

APPENDIX 

 

 

 

 

 

 

115 

 

Figure A.2.1 AIM scores in L-DOPA dosing paradigm 

AIM scores of days 1-10 in the L-DOPA dosing regimen, ranging from 2mg/kg-8mg/kg. Significantly more severe AIMs 
were observed in rAAV-GFP animals starting on day 10 with 8mg/kg. Peak-dose severity scores taken at 75 minutes post 
L-DOPA. (*=p≤0.05, **=p≤.01, ***=p≤.001) 

 

 

 

 

 

 

 

 

 

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Chapter 3: Modulating levels of Nurr1 expression in the parkinsonian rat striatum 

impacts LID severity and development 

Introduction 

Levodopa (L-DOPA) is considered the gold-standard pharmacologic therapy for the 

motor symptoms in Parkinson’s disease (PD). The precursor to dopamine (DA), L-

DOPA therapy alleviates motor symptoms by restoring balance to basal ganglia 

signaling following a loss in DA signaling from degenerated substantia nigra pars 

compacta (SNc) neurons (Cotzias et al., 1967; Fox et al., 2011). Unfortunately, chronic 

treatment with L-DOPA leads to the inevitable development of drug-induced abnormal 

involuntary movements in the majority of patients (Ahlskog & Muenter, 2001; Manson et 

al., 2012). These symptoms, termed levodopa-induced dyskinesia (LID), consist of 

disruptive hyperkinetic movements and dystonic muscle tone associated with L-DOPA 

plasma levels.  

Studies have focused on gene expression changes associated with LID to better 

understand their development and etiology. A previous study looking at gene 

expression differences between the direct and indirect pathway of the basal ganglia in 

dyskinetic rats showed a marked increase in expression of the transcription factor Nurr1 

(Heiman et al., 2014). This has been corroborated by a study that observed Nurr1 

transcript increase globally in the striatum of parkinsonian mice treated chronically with 

L-DOPA (Sodersten et al., 2014). Additionally, non-peer reviewed data from our group 

has shown LID-associated Nurr1 mRNA colocalized with markers of both the direct and 

indirect pathway, with a noted higher abundance in the direct pathway. Nurr1 is an 

orphan nuclear transcription factor and a member of the NR4A family (Giguere, 1999). It 

 

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has long been studied in PD as Nurr1 expression is crucial for DAergic neuronal 

development and long-term maintenance. Nurr1 knockout mice are not viable, and 

Nurr1+/- heterozygotes show DA dysfunction and loss of SNc neurons (Zetterstrom et 

al., 1997; Jiang et al., 2005; Kadkhodaei et al., 2009). Nurr1 expression in the SNc has 

been shown to decrease with age in humans, and in fact multiple Nurr1 isoforms have 

been found to be associated with some familial forms of PD (Dekker et al., 2003; Le et 

al., 2003). Because of this, Nurr1-based therapeutics are being researched as potential 

disease-modifying treatments for PD. The association of Nurr1 with LID, thus, begs for 

further characterization (Kim et al., 2015; Smith et al., 2015; Dong et al., 2016). 

The upregulation of Nurr1 in the striatum of dyskinetic animals is especially of note as 

Nurr1 is not normally expressed in striatal medium spiny neurons (MSNs) (Xiao et al., 

1996). This abnormal expression pattern associated with abnormal behavior suggests 

that Nurr1 plays a direct role in LID. However, this has not been shown directly. Nurr1 

has been shown to be involved in synaptic plasticity required for learning and memory in 

the hippocampus (Peña de Ortiz et al., 2000; Colon-Cesario et al., 2006; Hawk & Abel, 

2011; Hawk et al., 2012). As synaptic plasticity and spine dynamics have been 

previously implicated in LID formation (Fieblinger & Cenci, 2015), it is reasonable to 

question if Nurr1 plays a role in these changes. In the present study, we aim to show 

that Nurr1 is a key player in LID development and that it’s overexpression can induce 

LID. 

 

 

 

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Methods 

I. 

Adeno-associated virus production 

The human Nurr1 and GFP gene were cloned into the rAAV genome under control of 

the chicken β-actin/cytomegalovirus (CBA/CMV) promotor to allow for robust and 

ubiquitous expression in neurons. For shRNA experiments, a cassette expressing GFP 

as a transduction marker with either an shRNA targeted for the Nurr1 transcript 

(shRNA-Nurr1) or a scrambled control with no gene specificity (shRNA-SCR) was 

cloned under the same promoter element. rAAV pseudotype rAAV2/5 was produced via 

double transfection of HEK293 cells with rAAV genome and helper plasmids as 

previously described in this dissertation (Benskey et al., 2016). Virus was recovered 

using an iodixanol gradient and concentrated in concentration columns. Viral titer was 

ascertained by dot blot, and adjusted to a working titer of 1.0x1013 vg/mL. 

II. 

Animals and surgeries 

Adult male Fischer F344 and Lewis rats (200-220g on arrival, Charles River, 

Wilmington, MA) were used in the studies. Studies were conducted in accordance with 

the guidelines of Michigan State University (AUF MSU06/16-093-00). Rats were housed 

two per cage up until LID behavior testing began, when they were separated and singly 

housed. Animals were housed in a light-controlled (12 hours light/dark cycle) and 

temperature-controlled (22±1 °C) room, and had free access to standard lab chow and 

water. 

All surgeries were performed as described previously in this dissertation. Animals were 

anesthetized with isoflurane and placed into a stereotaxic frame. All materials were 

 

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injected with a Hamilton syringe fitted with a glass capillary needle. Parkinsonian lesions 

were induced using 5mg/ml 6-hydroxydopamine (6-OHDA) hydrobromide mixed in 

0.2mg/ml ascorbic acid immediately before injections. Animals received two 2µl 

injections of 6-OHDA (10µg per injection), one in the medial forebrain bundle (MFB, 

from bregma: Anterior Posterior (AP) – 4.3mm, Medial Lateral (ML) + 1.6mm, Dorsal 

Ventral (DV) - 8.4mm from skull) and one in the SNc (from bregma: AP - 4.8mm, ML + 

1.7mm, DV - 8.0mm from skull). The needle was lowered to the site and the injection 

began after 30 seconds. The needle was removed two minutes after the injection was 

finished and cleaned between each injection. Lesion efficacy was estimated two and a 

half weeks following 6-OHDA injection with the cylinder task as previously described 

(Schallert, 2006; Manfredsson et al., 2007).  

Viral delivery surgeries were performed similarly three weeks following 6-OHDA lesions. 

Animals in the overexpression studies received a single 2µl injection of either rAAV-

Nurr1 or rAAV-GFP targeting lateral striatum (from bregma: AP + 0.5mm, ML + 3.7mm, 

DV – 5.3mm from skull). Animals in the knockdown studies received two 2µl injections 

of either rAAV-shRNA-Nurr1 or rAAV-shRNA-SCR. Injections targeted the striatum (1st 

injection from bregma: AP + 0.0mm, ML + 3.0mm, DV – 5.2mm from dura. 2nd injection 

from bregma: AP + 1.6mm, ML + 2.7mm, DV – 4.9mm from dura). 

III. 

Abnormal involuntary movements ratings 

Drug-induced dyskinesia severity was evaluated using the abnormal involuntary 

movement (AIM) scale described earlier in this dissertation. Animals received 

subcutaneous injections of either increasing doses of L-DOPA (2-8mg/kg) with 

benserazide (12mg/kg), or the DA receptor agonists SKF-81297 (0.8mg/kg) and 

 

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quinpirole (0.2mg/kg). Dosing occurred three days a week for three weeks for L-DOPA 

studies and one week for DA agonist studies. The AIM scale was used to rate drug-

induced AIMs, as has been previously described (Steece-Collier et al., 2003; Maries et 

al., 2006). Briefly, AIM severity is evaluated by scoring the level of dystonia of the limbs 

and body, hyperkinesia of the forelimbs, and hyperoral movements. Each AIM is given 

two scores—one indicating the intensity (0=absent, 1=mild, 2=moderate, 3=severe) and 

frequency (0=absent, 1=intermittently present for <50% of the observation period, 

2=intermittently present for >50% of the observation period, 3=uninterruptable and 

present through the entire rating period). Each AIM is given a severity score by 

multiplying the intensity and frequency, and the overall AIM score for each timepoint is a 

sum of all the behaviors severities. The sum of all AIM scores from each timepoint 

makes up the total AIM score. Peak-dose dyskinesia is considered to be 75 minutes 

post drug administration. An animal is considered non-dyskinetic with a score of ≤4 

(Maries et al., 2006). Animals were observed in 25-minute increments following drug 

delivery until AIMs subsided. 

IV. 

Tissue collection and processing 

Animals received a final injection of either L-DOPA or DA agonists two hours prior to 

sacrifice. Rats were anesthetized deeply with a lethal dose of sodium pentobarbital, and 

intracardiacally perfused with Tyrode’s solution (137mM sodium chloride, 1.8mM 

calcium chloride dihydrate, .32mM sodium phosphate monobasic dihydrate, 5.5mM 

glucose, 11.9mM sodium bicarbonate, 2.7mM potassium chloride) followed by 4% 

paraformaldehyde (PFA). Brains were removed rapidly and post-fixed for 72 hours in 

4% PFA before being transferred into 30% sucrose. Brains were sectioned on a 

 

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freezing sliding microtome at 40µm and stored at -20°C in cryoprotectant (30% ethelyne 

glycol, 0.8mM sucrose in 0.5X tris-buffered saline). 

Immunohistochemistry (IHC) was performed as previously reported (Benskey et al., 

2018). A 1:6 series of free-floating tissue was stained for TH (MAB318, MilliporeSigma, 

Burlington, MA) Nurr1 (AF2156, R&D Systems, Minneapolis, MN) or GFP (AB290, 

Abcam, Cambridge, United Kingdom). Briefly, sections were washed in 1x TBS with 

.25% Triton x-100, incubated in 0.3% H2O2 for 30 minutes, and rinsed and blocked in 

10% normal goat or donkey serum for 2 hours. Tissue was incubated in primary 

antibody (TH 1:4000, Nurr1 1.5ug/ml, GFP 1:20,000) overnight at room temperature. 

After washing, tissue was incubated in secondary antibody (Biotinylated horse anti-

mouse IgG 1:500, BA-2001, Vector Laboratories, Burlingame, CA; biotinylated donkey 

anti-goat IgG 1:500, AP180B, Millipore-Sigma, Burlington, MA; biotinylated goat anti-

rabbit IgG 1:500, AP132B, Millipore-Sigma, Burlington, MA) followed by the Vectastain 

ABC kit (Vector Laboratories, Burlingame, CA). Tissue staining was developed with 0.5 

mg/ml 3,3’ diaminobenzidine (DAB, Sigma-Aldrich, St. Louis, MO) and 0.03% H2O2. 

Sections were mounted on slides, dehydrated, and coverslipped with Cytoseal 

(ThermoFisher, Waltham, MA). 

V. 

In situ hybridization 

In situ hybridization with IHC was performed using the RNAscope® 2.5 HD Duplex 

Assay according to the manufacturers’ protocol (Advanced Cell Diagnostics, Newark, 

CA). 40µm striatal sections were treated with the hydrogen peroxide solution for at least 

10 minutes, or until active bubbling from the tissue subsided. Sections were washed in 

1x TBS, mounted onto slides, and allowed to dry for at least 48 hours. Slides were then 

 

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boiled for 10 minutes in the Target Retrieval solution, followed by Protease Plus 

treatment. Tissue was hybridized with target probe for direct and indirect pathway 

markers (dynorphin (Dyn) or enkephalin (Enk), respectively) (Lu et al., 1998) for 2 hours 

at 40°C. Slides were rinsed in 1x RNAscope® Wash Buffer. Sequential amplification 

steps were then applied to the slides, with 1x Wash Buffer rinses between each 

amplification. After the sixth amplification, the probe was visualized using the Detect 

Red Signal solution for 10 minutes. 

Immediately following in situ hybridization, the tissue was stained 

immunohistochemically for Nurr1. IHC was performed as described above, and Nurr1 

protein was visualized using Vector SG Peroxidase kit (Vector Laboratories, 

Burlingame, CA). 

VI. 

Stereology 

Unbiased stereology was used to determine lesion status via TH loss as previously 

described (Polinski et al., 2015). Using Stereo Investigator software with the optical 

fractionator probe (MicroBrightfield Bioscience, Williston, VT), TH-positive neurons in 

every sixth section of the whole SNc were counted on the intact and lesioned 

hemisphere, giving an estimate of total TH-positive cells in the SNc. 

VII.  Statistical analysis 

Statistical analysis was performed using Statview (version 5.0) or GraphPad Prism 

version 7.0 (GraphPad Software, La Jolla, CA. All graphs were created in GraphPad. 

Lesion status was evaluated using unpaired, one-tailed t-tests. AIMs were evaluated 

using a Mann-Whitney U test or the Kruskal-Wallis test. Differences between vector 

 

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groups were compared with p≤0.05 being considered statistically significant. Bonferroni 

post-hoc tests were employed when significant main effects were detected. 

Results 

I. 

Validation of lesion and transgene expression 

Adult male Fisher and Lewis rats were lesioned with 6-OHDA injected into the medial 

forebrain bundle and substantia nigra. Lesion efficacy was validated post-mortem with 

TH staining and stereology. All animals included in analysis were sufficiently lesioned 

with ≥94% TH loss (Figure 3.1B-D). rAAV 2/5 expressing either Nurr1 (Fischer n=7, 

Lewis n=5) or GFP (Fischer n=7, Lewis n=5) was injected into the striatum of both 

strains of rats. Vector transduction and expression was confirmed with post-mortem IHC 

(Figure 3.1E-H). Vector delivery results in Nurr1 expression markedly higher than what 

is seen in rAAV-naïve, LID+ Fischer animals, where Nurr1 is abnormally upregulated 

(Figure A.3.1A). Lewis animals, however, express much lower level AIMs than Fischer 

rats, and do not express Nurr1 in the striatum (Figure A.3.1B). 

II. 

Ectopic Nurr1 overexpression does not exacerbate AIMs in LID-susceptible 

rats 

Three weeks were allowed after the virus delivery to ensure maximal transgene 

expression before animals were placed on a L-DOPA treatment paradigm (see Figure 

3.1A for experimental timeline). AIMs were evaluated in 25-minute intervals post-L-

DOPA delivery. Animals were first evaluated with vehicle (0mg/kg L-DOPA, 12mg/kg 

benserazide) to ensure that Nurr1 overexpression does not cause drug-independent 

AIMs. Indeed, both groups showed no AIMs with the vehicle administration (Figure  

 

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Figure 3.1 Experimental design and model validation 

(A) Experimental timeline showing surgeries and LID induction timeline. AIM scores 
were evaluated at each injection where indicated. (B-D) 6-OHDA injections caused 
near-complete loss of TH fibers in the striatum (B) as well as TH-positive cells in the 
nigra (C). Stereology confirmed that all animals had greater than 94% TH cell loss in the 
nigra (D). (E-H) Nurr1 (E and F) and GFP (G and H) vector driven expression were 
confirmed in the striatum with IHC.  
B, C, E, H scale bar=1mm 
F, H scale bar=100µm  
 

 

135 

 

A.3.2). When treated with L-DOPA, there was no significant difference in total AIM 

scores at any rating timepoint in Fischer rats treated with rAAV-Nurr1 or GFP (Figure 

3.2) (total AIM sum: D1 2mg/kg rAAV-Nurr1 (Md=0.5) rAAV-GFP (Md=1), U=17, p>0.05; 

D3 4mg/kg rAAV-Nurr1 (Md=8) rAAV-GFP (Md=6.5), U=10, p>0.05; D5 6mg/kg rAAV-

Nurr1 (Md=39.5) rAAV-GFP (Md=29), U=14, p>0.05; D8 8mg/kg rAAV-Nurr1 (Md=55) 

rAAV-GFP (Md=42), U=12, p>0.05; D10 8mg/kg rAAV-Nurr1 (Md=61.5) rAAV-GFP 

(Md=52.5), U=11, p>0.05; D12 8mg/kg rAAV-Nurr1 (Md=57.5) rAAV-GFP (Md=48), 

U=14, p>0.05; D15 8mg/kg rAAV-Nurr1 (Md=55) rAAV-GFP (Md=63), U=15, p>0.05; 

D17 8mg/kg rAAV-Nurr1 (Md=62.5) rAAV-GFP (Md=54), U=13, p>0.05). In order to 

determine if Nurr1 affected specific AIM behaviors that may not be apparent in the 

overall AIM score, we separated the overall AIM score into individual axial, orolingual, 

and limb scores (AOL scores). Again, Nurr1 did not drive a difference in AIM severity in 

any individual LID behavior criteria (Figure 3.3). There was a difference observed on the 

final treatment (day 17) in forelimb AIM severity at 150 minutes post L-DOPA (rAAV-

Nurr1 (Md=2) rAAV-GFP (Md=0), U=7, p<0.05). While statistically significant, this is 

likely not biologically relevant, as by this time point the AIMs of both groups were below 

our threshold for what is considered a dyskinetic score (AIM score≥4 to be LID+) 

(Maries et al., 2006). This data shows that Nurr1 did not have a specific effect on one 

particular set of AIM behaviors. We observed moderate and severe LID in both groups 

of Fischer rats, suggesting that ectopic Nurr1 expression does not exacerbate the AIMs 

of the LID-prone strain. 

 

 

 

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Figure 3.2 Ectopic Nurr1 does not exacerbate AIMs in LID-susceptible rats 

(A) Peak-dose LID severity (75 minutes post injection) over each rating period. No differences in peak-dose severity 
between vector groups was observed in Fischer rats. (B) The total AIM score sum of each rating session showed no 
differences between rAAV-Nurr1 and rAAV-GFP animals. Both groups developed severe AIMs similarly over the 
treatment regimen. (C-F) Animals were treated with 8mg/kg L-DOPA on days 10-17. Both groups showed similar AIM 
expression over time (left panels) and peak-dose AIMs (right panels).  

 

 

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Figure 3.3 Nurr1 does not impact individual AOLs in LID-susceptible rats 

(A) Axial AIMs—comprised of trunk and neck dystonia—are not different between rAAV-Nurr1 and rAAV-GFP Fischer 
rats. (B) Orolingual AIMs—comprised of chewing and tongue protrusions—are not different between vector groups. (C) 
Forelimb AIMs—comprised of forelimb hyperkinesia and dystonia—are not different at most time points between vector 
treatment groups. There was a significant difference between the groups at 150 minutes post injection. Rating period 
shown at day 17 with 8mg/kg L-DOPA dosing.

 

 

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III. 

Ectopic Nurr1 overexpression severe induces AIMs in LID-resistant rats 

Notably, when treated chronically with L-DOPA, rAAV-Nurr1 Lewis rats developed 

severe LID, where their rAAV-GFP control counterparts expressed low-level AIMs 

(Figure 3.4). This difference was first observed at the 6mg/kg treatment, in both total 

AIM score sum and peak-dose LID (total AIM sum 6mg/kg rAAV-Nurr1 (Md=34) rAAV-

GFP (Md=3), U=3, p<0.05; peak-dose AIM 6mg/kg rAAV-Nurr1 (Md=13.5) rAAV-GFP 

(Md=4), U=0, p<0.01). rAAV-Nurr1 treated Lewis rats also displayed more severe AIMs 

than their rAAV-GFP counterparts on days 8, 10, and 19 with 8mg/kg L-DOPA (peak 

dose AIMs: day 8 rAAV-Nurr1 (Md=16) rAAV-GFP (Md=3), U=2, p<0.05; day 10 rAAV-

Nurr1 (Md=16) rAAV-GFP (Md=6), U=0, p<0.01; day 17 rAAV-Nurr1 (Md=13.5) rAAV-

GFP (Md=4), U=0, p<0.01; day 19 rAAV-Nurr1 (Md=16) rAAV-GFP (Md=3), U=2, 

p<0.05). The induction of severe AIMs with ectopic Nurr1 expression in an otherwise 

LID resistant strain shows that Nurr1 is directly involved in LID development. 

When compared, rAAV-Nurr1 Lewis rats AIM scores were indistinguishable from rAAV-

GFP Fischer rats (Figure 3.5). On day 17 of treatment, rAAV-GFP Lewis rats were 

significantly less dyskinetic than both rAAV-Nurr1 Lewis and Fischer treated animals 

(peak dose AIMs: rAAV-Nurr1 Lewis (Md=13.5) rAAV-GFP Lewis (Md=4), U=0, p<0.01; 

rAAV-Nurr1 Fischer (Md=16) rAAV-GFP Lewis (Md=4), U=2, p<0.01). Interestingly, 

rAAV-GFP Fischer animals did not express significantly more severe AIMs than rAAV-

GFP Lewis rats (peak dose AIMs: rAAV-GFP Fischer (Md=17) rAAV-GFP Lewis 

(Md=4), U=3, p>0.05).  Together, this data shows that the presence of Nurr1 is sufficient 

to overcome resistance to severe LID seen in Lewis animals. 

 

 

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Figure 3.4 Ectopic Nurr1 expression induces severe AIMs in LID-resistant rats 

(A) Peak-dose LID severity (75 minutes post injection) show rAAV-Nurr1 treated Lewis rats developing significantly more 
sever AIMs than rAAV-GFP animals at 6 and 8mg/kg doses. (B) Total AIM score sum showed exacerbated AIMs on days 
5, 8, 10, 12, and 19 in rAAV-Nurr1 treated Lewis rats. (C-D) Individual rating periods on days 10-19 with 8mg/kg 
treatment. rAAV-Nurr1 animals developed AIMs more severe than rAAV-GFP animals at multiple timepoints during the 
observation period. (*=p≤0.05, **=p≤.01, ***=p≤.001) 

 

 

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Figure 3.5 Ectopic Nurr1 induces AIMs in LID-resistant rats that are similar to LID-susceptible rats 

(A) Day 17 AIM rating time course comparing rAAV treated LID-resistant Lewis rats to LID-susceptible Fischer rats. rAAV-
Nurr1 Lewis animals showed indistinguishable AIMs compared to both groups of Fischer rats. (B) Peak dose (75 minutes 
post injection) AIM severity on day 17 of treatment. rAAV-GFP Lewis animals showed significantly lower LID severity than 
both rAAV-Nurr1 treated Lewis and Fischer rats. rAAV-GFP treated Fischer rats did not show significantly higher AIMs 
than rAAV-GFP treated Lewis rats. (*=p≤0.05, **=p≤.01, ***=p≤.001) 

 

 

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IV. 

Nurr1 expression is induced by direct pathway activation 

We were next interested in characterizing Nurr1 induction in LID and if activation of 

either the direct or indirect pathway is sufficient for the abnormal expression to occur. 

Previous reports have found L-DOPA-induced Nurr1 expression in both direct and 

indirect pathway MSNs (dMSNs and iMSNs, respectively). To test this, we induced 

AIMs in parkinsonian Fischer rats with D1 agonist SKF-81297 or D2 agonist quinpirole 

(saline injection as a control). Animals were treated for one week and sacrificed 2 hours 

following the final dosing. Moderate and severe AIMs developed in animal treated with 

SKF-81297 (Figure 3.6). Quinpirole-treated rats expressed mild and moderate AIMs 

(Figure 3.6). No saline-treated animals developed AIMs. A Kruskal-Wallis test showed 

that drug-induced AIMs were significantly different between treatment groups (peak-

dose AIMs: Kruskal-Wallis statistic=6.78, p<0.05). IHC showed Nurr1 expression in the 

lesioned striatum of SKF-81297 rats with AIMs, but not in those that did not develop 

AIMs, suggesting that Nurr1 is associated with AIM behavior and can be upregulated 

with direct pathway activation alone (Figure 3.6C-F). Notably, no staining for Nurr1 

protein was observed in AIMs-expressing animals treated with quinpirole, despite 

moderate AIM scores (Figure 3.6G-J). This shows that indirect pathway activation alone 

does not induce maladaptive Nurr1 expression, and that direct pathway activation is 

required for this event. 

In order to determine if direct pathway activation lead to Nurr1 expression only in D1 

MSNs, we performed in situ hybridization with IHC to localize Nurr1 protein with mRNA 

of direct and indirect pathway markers (dynorphin (Dyn) and enkephalin (Enk), 

respectively) (Lu et al., 1998). We observed colocalization of Nurr1 protein and both  

 

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Figure 3.6 Abnormal striatal Nurr1 expression is induced by direct D1 receptor activation 

 

 

143 

 

Figure 3.6 (cont’d) 

(A) AIM scores from the final treatment with either D1 agonist (SKF-81297), D2 agonist 
(quinpirole), or saline. D1-agonist treated rats displayed severe AIMs, while D2 agonist 
treated animals expressed moderate AIMs. No AIMs were observed in saline treated 
animals. (B) Peak-dose AIM severity from the final drug treatment.  (C-L) Nurr1 IHC in 
animals treated with SKF-81297 (C-F), quinpirole (G-J) or saline (K-L). Nurr1 protein 
was only observed in the lesioned striatum of dyskinetic rats treated with SKF-81297 
(C), but not in LID- animals with the same treatment (E) nor in the intact hemisphere (D 
and F). Nurr1 was not observed in the lesioned or intact striatum of LID+ rats treated 
with quinpirole (G and H), nor in LID- quinpirole treated animals (I-J). No Nurr1 was 
seen in saline treated animals (K and L). Scale bar=100µm 

 

 

 

 

 

 

 

 

 

 

 

 

 

144 

 

Dyn and Enk in SKF-81297 AIM-expressing animals (Figure 3.7). These findings 

suggest that Nurr1 expression in the indirect pathway is dependent on direct pathway 

activation. We also tested this in rats treated with quinpirole and saline, but no 

colocalization was observed as no Nurr1 protein was expressed in these animals 

(Figure A.3.3). 

V. 

Viral knockdown of Nurr1 does not inhibit LID development 

Our next objective was to determine if inhibiting the abnormal LID-associated Nurr1 

expression in MSNs would block LID development. To achieve this, we used a vector 

cassette expressing an shRNA for Nurr1 (rAAV-shRNA-Nurr1) or a scrambled 

sequence with no gene specificity as a control (rAAV-shRNA-SCR). Animals were 

lesioned with 6-OHDA followed by stereotaxic vector delivery as described above. 

When challenged with L-DOPA, no significant reduction in AIMs was observed in rAAV-

shRNA-Nurr1 animals (Figure 3.8). In fact, rAAV-shRNA-SCR animals expressed less 

severe AIMs than the rAAV-shRNA-Nurr1 animals (Total AIM sum, day 19: rAAV-

shRNA-Nurr1 (Md=13.5) rAAV-shRNA-SCR (Md=2), U=31, p<0.05).  This suggests that 

silencing Nurr1 prior to L-DOPA delivery is not sufficient to block LID in susceptible 

animals, indicating that there are likely redundant pathways to LID development that do 

not require Nurr1. 

We noted that the animals treated with the rAAV-shRNA-SCR expressed notably less 

severe AIMs than expected for the strain. This could be due to inconsistent 6-OHDA 

lesions, or shRNA toxicity. To show that rAAV-shRNA-Nurr1 treatment did not 

exacerbate LID severity beyond what is expected for parkinsonian Fischer rats, we 

compared the rAAV-shRNA-Nurr1 treated animals to the control rAAV-GFP Fischer rats  

 

145 

 

 

Figure 3.7 D1 stimulation induces Nurr1 expression in both indirect and direct 
pathway MSNs 

(A) In situ hybridization for dynorphin (red) and IHC for Nurr1 protein (blue) in striatum 
of LID+ animal treated with D1 agonist SKF-81297. Dyn transcript and Nurr1 protein are 
colocalized (white arrows) in some neurons, but not others (black arrows). (B) In situ 
hybridization for enkephalin (red) and IHC for Nurr1 protein (blue) in the striatum of an 
LID+ animal treated with D1 agonist SKF-81297. The enk transcript is seen to colocalize 
with Nurr1 protein (white arrows). Some cells show Nurr1 expression with no enk 
transcript (black arrows) Scale bar=100µm 

 

 

 

 

 

 

 

 

 

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Figure 3.8 Nurr1 silencing does not inhibit LID development 

(A) Peak dose (75 minutes post injection) AIM severity over the study duration. rAAV-shRNA-Nurr1 treated animals 
developed severe peak-dose AIMs over the study, while rAAV-shRNA-SCR animals developed moderate AIMs. (B) The 
sum AIM score from each rating period. shRNA-Nurr1 animals developed severe AIMs over time, more significant than 
shRNA-SCR animals. (C-H) Individual rating timepoints at 8mg/kg dosing. shRNA-Nurr1 did not inhibit sever AIMs from 
developing. rAAV-shRNA-SCR animals displayed moderate AIMs throughout the study.   

 

 

147 

 

 

from the overexpression study, as these animals developed severe AIMs more typical of 

the strain. rAAV-shRNA-Nurr1 animals developed AIMs similarly to rAAV-GFP animals 

(Figure 3.9) (rAAV-shRNA-Nurr1 day 17, peak dose AIMs=12.21±1.97, rAAV-GFP day 

17, peak dose AIMs=14±3.15). This shows that the shRNA-Nurr1 delivery did not 

exacerbate LID, but also showed no benefit in reducing AIM severity. 

Discussion 

We aimed to test if Nurr1 impacts LID by inducing AIMs through altering the physiology 

of MSNs. Nurr1 has been previously associated with a dyskinetic state in rodent 

models, and its involvement is of note for multiple reasons (Heiman et al., 2014; 

Sodersten et al., 2014). Firstly, Nurr1 is known to be required for not only the 

development but also the long-term survival and health of DA neurons (Zetterstrom et 

al., 1997; Jiang et al., 2005; Kadkhodaei et al., 2009). This has prompted multiple 

groups to begin pursuing Nurr1-based therapies as PD-modifying treatments, including 

Nurr1 agonist compounds (Kim et al., 2015; Smith et al., 2015). How Nurr1 is involved 

in in LID must thus be well characterized and understood, as increasing its expression 

or activity could exacerbate LID. Second, Nurr1 is involved in learning-based plasticity 

in the hippocampus. This implies that Nurr1 may be influencing LID in the striatum by 

promoting spine plasticity that has been shown to be associated with LID. Finally, while 

other NR4A transcription factors are expressed endogenously in the striatum, Nurr1 is 

not (Xiao et al., 1996; Zetterstrom et al., 1996). This abnormal expression seen only 

with these adverse motor symptoms suggests the sole role of Nurr1 in MSNs is to 

promote these maladaptive changes. 

 

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Figure 3.9 Nurr1 shRNA does not exacerbate LID 

(A) Individual rating period comparing AIMs between rAAV-shRNA-Nurr1 and rAAV-
GFP treated Fischer rats. Both groups show similar AIM expression patterns. (B) Peak 
dose (75 minutes post injection) AIMs show similar AIM severity in rAAV-shRNA-Nurr1 
animals compared to rAAV-GFP animals. 

 

 

 

 

 

 

 

 

 

 

 

 

149 

 

Utilizing LID-prone Fischer rats and LID-resistant Lewis rats, we have shown that 

ectopic Nurr1 expression can induce severe LID in the Lewis animals. GFP-treated 

control Lewis rats displaying mild AIM scores do not show an endogenous upregulation 

of Nurr1 (see Figure A.3.1B), suggesting that the lack of Nurr1 contributes to the strain’s 

resistance to severe LID. Interestingly, delivering rAAV-Nurr1 to Fischer rats did not 

induce more severe AIMs than control Fischer rats. It is likely that in strains that are 

prone to develop LID, Nurr1 cannot exacerbate the severe AIMs that would happen 

without intervention. This ‘ceiling effect’ suggests that Nurr1 is important for LID 

development and the appearance of severe AIMs, but increasing expression does not 

correlate with worsening AIMs. While vector delivery results in much higher Nurr1 

expression than what is seen in an LID+ rAAV-naïve animal (see Figure 3.1E-F and 

Figure A.3.1A), our data suggests that the hyperexpression from viral delivery does not 

further LID severity. Likely, there is a critical threshold level of endogenous (or ectopic) 

Nurr1 expression must be reached to allow for LID induction, but higher expression 

levels do not exacerbate the behavior. This suggests that Nurr1 plays a role primarily in 

severe LID development. Additionally, it has been shown that Nurr1 displays differential 

transcriptional activity based on its expression level (Johnson et al., 2011; Do, 2014). It 

is possible that the higher expression levels achieved with viral expression promote 

gene transcription that is not related to LID escalation. However, the fact that LID can 

be induced in LID-resistant Lewis rats with viral Nurr1 expression suggests that this 

level of expression still allows for Nurr1-mediated transcription that influences LID. Our 

study did not evaluate the minimum level of Nurr1 expression or number of MSNs 

 

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expressing Nurr1 for LID to present, so further characterization of Nurr1 expression in 

LID is warranted. 

Severe AIM expression in rAAV-Nurr1 treated Lewis animals suggests that Nurr1 is a 

key factor in LID development, as this strain of animals does not develop severe LID 

without Nurr1 ectopically being expressed. This additionally shows that Nurr1 is not a 

downstream byproduct of LID expression, and rather does play a role in their 

development. The ectopic expression of Nurr1 was sufficient to overcome LID 

resistance in these rats and induce AIMs as severe as those seen in LID-prone Fischer 

rats, suggesting that Nurr1 expression is a key difference in the two strains 

susceptibility. Identifying genetic or epigenetic differences between the strains that may 

explain their differential upregulation of Nurr1 in response to L-DOPA would be a 

valuable study to identify molecular traits involved in LID susceptibility. 

Nurr1 has previously been observed in striatal MSNs in primarily the direct pathway 

(Heiman et al., 2014). We were curious as to whether direct DA receptor activation 

could induce Nurr1 expression in either or both the direct or indirect pathway. We 

observed Nurr1 induction in MSNs of AIM-expressing parkinsonian rats treated with a 

D1 receptor agonist, and confirmed that the expression was occurring in both MSNs of 

the direct and indirect pathway. This is notable because it indicates that Nurr1 induction 

is dependent on direct pathway activation, and additionally that D1 receptor activation is 

sufficient to induce Nurr1 expression in indirect pathway neurons. These data align with 

studies of other immediate early genes in LID, such as Homer-1a and FosB, which 

show that D1 priming is required for expression in D2 neurons (Pollack & Yates, 1999; 

Yamada et al., 2007; Pollack & Thomas, 2010). 

 

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Interestingly, no Nurr1 protein was observed in AIM-expressing animals treated with the 

D2 agonist quinpirole. This suggests that there are D2-dependent pathways to LID 

development that do not require Nurr1 expression, and further confirms that Nurr1 

induction is reliant on direct pathway signaling. This could suggest a potential 

mechanism for low level LID development in Lewis rats that do not endogenously 

upregulate Nurr1 with AIMs. It is possible that differences in the DA receptors of Lewis 

rats promote more signaling through D2 receptors than D1, causing indirect pathway 

LID with no Nurr1 expression. 

It is important to note that both Lewis rats treated with L-DOPA and Fischer rats treated 

with the D2 agonist quinpirole develop mild and moderate AIMs, much less severe than 

Fischer rats treated with L-DOPA or a D1 agonist. As both of these groups with 

moderate AIMs do not show abnormal Nurr1 upregulation, it is reasonable to 

hypothesize that Nurr1 is required for severe AIM development. Nurr1 may act as a 

‘gate keeper’ of sorts that, when expressed, can induce striatal changes that lead to 

more severe dyskinesia.  

In conclusion, this study has shown that Nurr1 is not only a marker of LID, but is directly 

involved in LID development and severity. This characterization of Nurr1 highlights its 

important role in LID, and suggests that Nurr1 agonists-based therapies for PD could be 

detrimental to patients by reducing the efficacious period of L-DOPA by promoting LID 

development.  

 

 

 

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APPENDIX 

 

 

 

 

 

 

 

 

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Figure A.3.1 Abnormal Nurr1 upregulation in dyskinetic rats 

(A) Nurr1 IHC in the lesioned striatum of an LID+ Fischer rat. Abnormal Nurr1 
expression can be seen throughout the lesioned striatum. (B) Nurr1 IHC in the lesioned 
striatum of an LID+ Lewis rat. No Nurr1 expression is seen in these animals. Scale 
bar=100µm. 

 

 

 

 

 

 

 

 

 

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Figure A.3.2 Virally-expressed Nurr1 does not induce drug-independent AIMs 

Fischer rats treated with rAAV-Nurr1 or rAAV-GFP do not express AIMs when treated 
with vehicle (0mg/kg L-DOPA, 12mg/kg benserazide) showing that Nurr1 
overexpression does not promote AIMs without drug treatment. 

 

 

 

 

 

 

 

 

 

 

 

 

 

155 

 

 

Figure A.3.3 Nurr1 colocalization with iMSNs in quinpirole and saline treated rats 

(A) In situ hybridization for enkephalin (Enk) and with IHC for Nurr1 in a dyskinetic rat 
treated with quinpirole. No Nurr1 protein was observed in these animals. (B) In situ 
hybridization for Enk with IHC for Nurr1 in a saline treated animal show no abnormal 
Nurr1 induction in saline treated rats. 

 

 

 

 

 

 

 

 

 

 

 

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Hansen, K. (2014). Dopamine signaling leads to loss of Polycomb repression and 
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Movement Disorder Society, 18(12), 1442-1454. doi:10.1002/mds.10588 

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in situ hybridization. Neuroscience, 75(1), 221-230.  

Yamada, H., Kuroki, T., Nakahara, T., Hashimoto, K., Tsutsumi, T., Hirano, M., & 

Maeda, H. (2007). The dopamine D1 receptor agonist, but not the D2 receptor 
agonist, induces gene expression of Homer 1a in rat striatum and nucleus 
accumbens. Brain Res, 1131(1), 88-96. doi:10.1016/j.brainres.2006.11.011 

Zetterstrom, R. H., Solomin, L., Jansson, L., Hoffer, B. J., Olson, L., & Perlmann, T. 

(1997). Dopamine neuron agenesis in Nurr1-deficient mice. Science, 276(5310), 
248-250.  

Zetterstrom, R. H., Williams, R., Perlmann, T., & Olson, L. (1996). Cellular expression of 

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system. Brain Res Mol Brain Res, 41(1-2), 111-120.  

 

 

 

 

 

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Chapter 4: Ectopic overexpression of Nurr1 induces a LID-like state in striatal 

medium spiny neurons 

Preface 

In this chapter, electrophysiology experiments were performed by Dr. Anthony West and 

Dr. Kuei Tseng at Rosalind Franklin University. All other experiments and analysis were 

performed at Michigan State University by Rhyomi C. Sellnow. 

Introduction 

Levodopa (L-DOPA) induced dyskinesias (LID) are a debilitating side effect that occur in 

the majority of Parkinson’s disease (PD) patients due to long term treatment with L-

DOPA (Cotzias et al., 1967; Ahlskog & Muenter, 2001; Manson et al., 2012). LIDs are 

involuntary motor behaviors that include chorea, dystonia, and limb hyperkinesia that 

are distinct from parkinsonian motor behaviors. Though these symptoms can become 

debilitating and affect the daily living of patients, L-DOPA remains the gold-standard 

treatment for the motor symptoms in PD. 

Much research has focused on LID-associated changes in the electrophysiology of the 

basal ganglia and its connecting structures. In vivo recordings of PD patients with deep 

brain stimulation (DBS) therapy have revealed impaired depotentiation in basal ganglia 

output structures associated with severe LID (Prescott et al., 2014). Indeed, this data 

reflects findings from animal studies which established a loss of bidirectional plasticity in 

LID (Picconi et al., 2003). Following DAergic denervation, both long-term potentiation 

(LTP) and long-term depression (LTD) are lost in the striatum. While L-DOPA treatment 

restores this corticostriatal plasticity, LTD is not present when LID develop (Picconi et 

al., 2003). Recent studies using optogenetically labeled neurons have shown dramatic 

 

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increase in activity in a specific population of direct pathway neurons, with simultaneous 

decreases in indirect pathway neuron activity (Ryan et al., 2018).This abnormal 

corticostriatal response and striatal neuron activity is a key factor in LID expression. 

Some of these LID-associated changes in plasticity may be accounted for by changes in 

the dendritic spines on the majority population of neurons of the striatum—medium 

spiny neurons (MSNs). Multiple studies have shown that spine density and morphology 

changes dramatically in animal models following LID induction (Zhang et al., 2013; 

Fieblinger et al., 2014; Nishijima et al., 2014; Suárez et al., 2014). Dendritic spines 

cover MSN dendrites. These structures act as synaptic ‘hot spots’, forming connections 

with presynaptic terminals on the head and neck of the spines (Maiti et al., 2015). 

Spines are dynamic structures, and their strengthening and pruning back can occur due 

to normal neuronal processes such as memory and learning, or be indicative of 

diseased states (Bagetta et al., 2010; Spiga et al., 2014; Maiti et al., 2015). In animal 

models, dramatic spine loss is observed following DA depletion, followed by a 

reestablishment of spines in LID (Zhang et al., 2013; Fieblinger et al., 2014; Nishijima et 

al., 2014; Suárez et al., 2014). Some reports show differential changes in direct versus 

indirect pathway neurons, suggesting opposing mechanisms between pathways 

(Fieblinger et al., 2014; Suarez et al., 2016). Changes in spine morphology have also 

been observed, with increase in mature mushroom type spines being associated with 

LID (Zhang et al., 2013). While total spine density appears to return to a more basal 

level in LID, evidence suggests that the synapses being formed on these spines are 

maladaptive. There is a dramatic increase in corticostriatal synapses in dyskinetic 

animals, with increased multi-synaptic connections and synapses on dendrites (Zhang 

 

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et al., 2013). Together this research has shown that changes in spine density and 

morphology are an important event in LID development, but further research is needed 

to understand what factors influence these changes. 

Nurr1 has been shown to be involved in synaptic plasticity required for learning and 

memory in the hippocampus (Peña de Ortiz et al., 2000; Colon-Cesario et al., 2006; 

Hawk & Abel, 2011; Hawk et al., 2012). As synaptic plasticity and spine dynamics have 

been previously implicated in LID formation (Fieblinger & Cenci, 2015), it is reasonable 

to question if Nurr1 plays a role in these changes. In the present study, we aim to show 

that Nurr1 influences LID by altering the physiology and morphology of MSNs. 

Methods 

I. 

Adeno-associated virus production 

The same vector design and production methods were used as described in Chapter 3 

of this dissertation. Briefly, the human Nurr1 and GFP gene were cloned into the AAV 

genome under control of a ubiquitous promoter. The genome was packaged into 

rAAV2/5 via double transfection of HEK293 cells with AAV genome and helper plasmids 

as previously described. Virus was recovered using an iodixanol gradient and 

concentrated in concentration columns. Viral titer was ascertained by dot blot, and 

adjusted to a working titer of 1.0x1013 vg/mL. 

II. 

Animals and surgeries 

Adult male Sprague Dawley or Fischer rats (200-220g on arrival, Charles River, 

Wilmington, MA) were used in the studies. Studies were conducted in accordance with 

the guidelines of Michigan State University (AUF MSU06/16-093-00) and Rosalind 

 

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Franklin University (AUF# A3279-01). Rats were housed two per cage up until LID 

behavior testing began, when they were separated and singly housed. Animals were 

housed in a light-controlled (12 hours light/dark cycle) and temperature-controlled (22±1 

°C) room, and had free access to standard lab chow and water. 

Surgeries were performed under similar conditions as described in Chapter 3 of this 

dissertation. Lesions were performed using 5mg/ml 6-hydroxydopamine (6-OHDA) 

hydrobromide mixed in 0.2mg/ml ascorbic acid immediately before injections. Sprague 

Dawley rats used for local field potential recordings and spine analysis were lesioned 

with two 2µL injections of 6-OHDA (10µg per injection) in the left striatum (1st injection 

from bregma: Anterior Posterior (AP) + 1.6mm, Medial Lateral (ML) 2.4mm, Dorsal 

Ventral (DV) – 4.2mm from skull; 2nd injection from 1st injection site: AP – 1.4mm, ML 

+0.2mm, DV – 2.8mm). Fischer rats used in antidromic cell recordings received two 2µl 

injections of 6-OHDA (10µg per injection), one in the medial forebrain bundle (MFB, 

from bregma: Anterior Posterior (AP) – 4.3mm, Medial Lateral (ML) + 1.6mm, Dorsal 

Ventral (DV) - 8.4mm from skull) and one in the SNc (from bregma: AP - 4.8mm, ML + 

1.7mm, DV - 8.0mm from skull). The needle was lowered to the site and the injection 

began after 30 seconds. The needle was removed two minutes after the injection was 

finished and cleaned between each injection. Lesion efficacy was estimated two and a 

half weeks following 6-OHDA injection with the cylinder task as previously described 

(Schallert, 2006; Manfredsson et al., 2007).  

Viral delivery surgeries were performed similarly three weeks following 6-OHDA lesions, 

as described previously (Benskey & Manfredsson, 2016). Animals received two 2µl 

injection of either rAAV-Nurr1 or rAAV-GFP targeting lateral striatum (1st injection from 

 

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bregma: AP + 1.0mm, ML + 3.0mm, DV – 4.0mm from dura; 2nd injection from bregma: 

AP – 1.6mm, ML + 3.8mm, DV – 5mm from dura).  

III. 

Tissue collection and processing 

Animals received a final injection of either L-DOPA or DA agonists two hours prior to 

sacrifice. Rats were anesthetized deeply with a lethal dose of sodium pentobarbital, and 

intracardiacally perfused with Tyrode’s solution (137mM sodium chloride, 1.8mM 

calcium chloride dihydrate, .32mM sodium phosphate monobasic dihydrate, 5.5mM 

glucose, 11.9mM sodium bicarbonate, 2.7mM potassium chloride) followed by 4% 

paraformaldehyde (PFA). Brains were rapidly removed and post-fixed for 72 hours in 

4% PFA before being transferred into 30% sucrose. Brains were sectioned on a 

freezing sliding microtome at 40µm and stored at -20°C in cryoprotectant (30% ethelyne 

glycol, 0.8mM sucrose in 0.5X tris-buffered saline). 

Immunohistochemistry (IHC) was performed as previously reported. A 1:6 series of free-

floating tissue was stained for TH (MAB318, MilliporeSigma, Burlington, MA) Nurr1 

(AF2156, R&D Systems, Minneapolis, MN) or GFP (AB290, Abcam, Cambridge, United 

Kingdom). Briefly, sections were washed in 1x TBS with .25% Triton x-100, incubated in 

0.3% H2O2 for 30 minutes, and rinsed and blocked in 10% normal goat or donkey serum 

for 2 hours. Tissue was incubated in primary antibody (TH 1:4000, Nurr1 1.5ug/ml, GFP 

1:20,000) overnight at room temperature. After washing, tissue was incubated in 

secondary antibody (Biotinylated horse anti-mouse IgG 1:500, BA-2001, Vector 

Laboratories, Burlingame, CA; biotinylated donkey anti-goat IgG 1:500, AP180B, 

Millipore-Sigma, Burlington, MA; biotinylated goat anti-rabbit IgG 1:500, AP132B, 

Millipore-Sigma, Burlington, MA) followed by the Vectastain ABC kit (Vector 

 

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Laboratories, Burlingame, CA). Tissue staining was developed with 0.5 mg/ml 3,3’ 

diaminobenzidine (DAB, Sigma-Aldrich, St. Louis, MO) and 0.03% H2O2. Sections were 

mounted on slides, dehydrated, and coverslipped with Cytoseal (ThermoFisher, 

Waltham, MA). 

IV. 

Local field potential and single cell in vivo recordings 

Striatal local field potential (LFP) response evoked by cortically stimulation were 

recorded as previously described (Thomases et al., 2013; Jayasinghe et al., 2015). 

Animals in both electrophysiology studies were shipped to Rosalind Franklin University 

three weeks following vector delivery. Animals were deeply anesthetized with 8% 

chloral hydrate. Electrodes were placed in the cortex for stimulation (from bregma: AP + 

3.5mm, ML + 2.0mm, DV – 2.0mm from dura) and the striatum for recording (from 

bregma: AP + 0.7, ML + 3.4mm, DV – 4.5mm from dura). The cortex was stimulated 

with pulses delivered every 15 seconds (Thomases et al., 2013). LFP recordings were 

taken 30-120 minutes following injection of 5mg/kg L-DOPA with 12mg/kg benserazide. 

In vivo single cell recordings of striatonigral projection neurons were measured in vector 

treated animals without L-DOPA treatment, or in established dyskinetic animals. 

Electrical stimulation and antidromic activation of striatonigral neurons was performed 

as previously described (Threlfell et al., 2009). Spikes evoked by stimulating the 

substantia nigra pars reticulata (SNr) were determined to be antidromically activated 

based on spike collision with orthodromic spikes occurring consistently over 10 trials 

(Threlfell et al., 2009).  

 

 

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V. 

Golgi-Cox impregnation and spine analysis 

Animals for spine analysis were perfused as described above with Tyrode’s solution 

followed by 4% PFA. Brains were removed and hemisected. The caudal portion of the 

brain was then postfixed in 4% PFA and used for lesion evaluation. The rostral portion 

was postfixed for 1 hour and then transferred to 0.2M phosphate buffer until further 

processing. The rostral section was sectioned on a vibratome at 100µm. Sections were 

then processed for Golgi-Cox impregnation as described previously (Levine et al., 

2013). Briefly, sections were sandwiched gently between two glass slides and placed 

into the Golgi-Cox solution (1% mercury chloride, 1% potassium chromate, 1% 

potassium dichromate) in the dark for 14 days. Sections were transferred into a 1% 

potassium dichromate solution for 24 hours. Sections were mounted on 4% gelatin-

coated slides and the stain was developed with 28% ammonium hydroxide followed by 

15% Kodak fixer. Slides were dehydrated in alcohol and xylene and coverslipped. 

Spine density and morphology were quantified using Neurolucida (MicroBrightfield 

Bioscience, Williston, VT) as has been previously described (Zhang et al., 2013). 

Neurons of the dorsal striatum were selected for analysis. To be selected for 

quantification, a neuron needed at least four primary dendrites that projected radially, 

not bidirectionally, from the cell body. One dendrite per neuron was traced and the 

spines quantified and typified. Each spine was typified into one of four classes: thin, 

mushroom, stubby and branched (Maiti et al., 2015). Ten individual dendrites per 

hemisphere per animal were quantified. Spine quantity and phenotype were evaluated 

based on total dendrite length, or proximal (dendrite branch orders 1-2) or distal (branch 

orders 3-n) dendritic regions. 

 

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VI. 

Stereology 

Unbiased stereology was used to determine lesion status via TH loss as previously 

described (Polinski et al., 2015). Using Stereo Investigator software with the optical 

fractionator probe, TH-positive neurons in every sixth section of the whole SNc were 

counted on the intact and lesioned hemisphere, giving an estimate of total TH-positive 

cells in the SNc. 

VII.  Statistical analysis 

Statistical analysis was performed using Statview (version 5.0) or GraphPad Prism 

version 7.0 (GraphPad Software, La Jolla, CA). All graphs were created in GraphPad. 

Lesion status was evaluated using unpaired, one-tailed t-tests. Differences in spine 

quantity and morphology were evaluated using unpaired t-tests. Differences between 

vector groups at each recording timepoint within a single treatment were compared with 

t-tests, with p≤0.05 being considered statistically significant. Bonferroni post-hoc tests 

were employed when significant main effects were detected. A two-way repeated 

measures ANOVA (GFP vs. gene therapy (ectopic expression of the DA D2 AR in 5-HT 

DR neurons)) x 2 (vehicle vs. drug treatment) with α set to 0.05 and all “n’s” adequately 

powered for electrophysiological studies was conducted using Sigma Stat software (San 

Jose, CA), and the potential two-way interaction effect was examined to determine how 

treatment effects differ as a function of drug treatment or gene therapy (Padovan-Neto 

et al., 2015). 

 

 

 

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Results 

I. 

Lesion validation and vector expression 

Animals for all studies were rendered parkinsonian with 6-OHDA injections as described 

previously. TH immunoreactivity was markedly reduced in the injected hemisphere 

(Figure 4.1A-B). Animals included in the analysis showed ~75-80% cell loss in the 

injected hemisphere and there was no difference in lesion severity between vector 

groups (rAAV-Nurr1 % TH neurons remaining=21.48±3.12; rAAV-GFP % TH neurons 

remaining=27.87±5.26; t13)=1.08, p>0.05). These lesions would be sufficient to induce 

striatal changes that occur following DA depletion. Transgene expression was 

confirmed with IHC, and all included animals showed robust transgene expression in 

the striatum (Figure 4.1.D-E). Animals were then used for electrophysiology (rAAV-

Nurr1 n=5, rAAV-GFP n=5) or spine analysis (rAAV-Nurr1 n=4, rAAV-GFP n=3). 

II. 

Striatal Nurr1 expression alone induces LID-like corticostriatal signaling 

We were first interested in how Nurr1 could impact striatal medium spiny neurons on a 

physiological level. Rats were rendered parkinsonian and received striatal injections of 

either rAAV-Nurr1 (n=5) or rAAV-GFP (n=5). These animals were not treated chronically 

with L-DOPA—and thus non-dyskinetic—in order to evaluate the effect of Nurr1 on 

striatal MSN activity independent of L-DOPA. Local field potential (LFP) was measured 

in the striatum following cortical stimulation in 30 minutes intervals after receiving a 

therapeutic dose of L-DOPA (5mg/kg, 12mg/kg benserazide). Over the recording 

period, rAAV-GFP animals showed depotentiation in corticostriatal response compared 

to baseline. This response has been shown previously in parkinsonian rats (Mallet et al.,  

 

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Figure 4.1 Lesion and vector confirmation 

(A-C) Lesion status was confirmed with IHC for TH. TH immunoreactivity was 
dramatically reduced in the interjected hemisphere of the striatum (A) and substantia 
nigra (B). Stereological estimates of remaining TH-positive neurons show significant cell 
loss in both vector groups (C). (D-E) Transgene expression from viral vector delivery 
was confirmed in the striatum with IHC for Nurr1 (D) or GFP (E). 
A and B scale bar=1mm 
D and E scale bar=1µm 
 
 
 
 

 

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2006). Animals treated with rAAV-Nurr1, however, showed a potentiated response 

(Figure 4.2). This suggests that Nurr1 promotes increased neuronal activity and 

potentiation in DA-depleted MSNs. Importantly, the LFP profile of rAAV-Nurr1 non-

dyskinetic animals was indistinguishable from that of rAAV-naïve animals with 

established LID. This indicates that Nurr1 expression induces LID-like activity in MSNs 

independent of L-DOPA.  

To further understand how Nurr1 expression can impact MSN activity and signaling, we 

performed in vivo single cell recordings of direct pathway MSNs (dMSNs) of L-DOPA 

naïve parkinsonian rats injected with rAAV-Nurr1 or rAAV-GFP. Striatonigral dMSNs 

were identified by antidromically stimulating the SNr and recording from single neurons 

in the striatum. After a dMSN was identified, the cortex was stimulated to record 

cortically-evoked responses in the neurons (Figure 4.3A). Vector treated animals were 

compared with rAAV-naïve rats with established LID. dMSNs of rAAV-Nurr1 injected 

animals showed an increase in spike probability and average number of spikes 

compared to rAAV-GFP animals (Figure 4.3B-C). Importantly, rAAV-Nurr1 cortically-

evoked spike response mimicked the spike profile of rAAV-naïve dyskinetic rats. This is 

similar to what we observed with LFP recordings. Together, this data suggests that 

dMSNs acquire LID-like activity with ectopic Nurr1 expression, and that these changes 

in firing rates are not dependent on DA signaling. 

 

 

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Figure 4.2 Striatal Nurr1 potentiates corticostriatal transmission 

Local field potential (LFP) of the striatum in response to a therapeutic 5mg/kg dose of L-
DOPA following cortical stimulation in animals injected with either rAAV-GFP (blue) or 
rAAV-Nurr1 (orange). A subset of animals did not receive a vector injection, but were 
instead rendered dyskinetic with L-DOPA priming (red). rAAV- GFP injected control 
subjects exhibited a suppression of the corticostriatal response, while rAAV-Nurr1 
treated subjects exhibited a potentiation of corticostriatal transmission which was 
virtually identical to that of LID+ subjects. As expected, saline treatment (Vehicle) did 
not induce a response in rAAV-Nurr1 treated animals. 

 

 

 

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Figure 4.3 Cortically-evoked response of striatonigral MSNs to antidromic 
stimulation 

Comparisons between cortically-evoked spike characteristics of antidromically-activated 
striatonigral projection neurons recorded from DA-depleted rAAV-naïve rats treated with 
chronic L-DOPA (LID+), rAAV-GFP injected L-DOPA naïve animals, or rAAV-Nurr1 
injected L-DOPA naïve animals. (A) Traces of typical cortically-evoked responses from 
isolated striatonigral projection neurons. Ten consecutive overlaid responses are 
shown. (B-D) graphs compare the spike probability (B), average number of total spikes 
evoked during a stimulus trial at each current intensity tested (C), and onset latency of 
evoked spikes (D) during cortical stimulation. Stimulus intensity-dependent effects on 
cortically-evoked spike probability of striatonigral projection neurons were observed in 
both LID+ and Nurr-1 overexpressing rats as compared to vehicle treated DA-depleted 
GFP-expressing controls (p< 0.001).  Post-hoc comparisons revealed a significant 
increase in the probability and number of evoked responses to cortical stimulation at the 
400-600 µA current intensities in both LID+ and Nurr-1 overexpressing rats as 
compared to vehicle treated DA-depleted, GFP-expressing control rats (*p< 0.05).  No 
significant differences in onset latency (p> 0.05) or SD of latency (data not shown) of 
cortically-evoked responses were observed.  

 

 

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III.  MSN spine density and morphology changes are induced by Nurr1 

expression 

Dramatic changes in spine density and morphology in LID models has been previously 

shown (Fieblinger & Cenci, 2015). As Nurr1 has been shown to be involved in learning 

and memory-associated plasticity (Peña de Ortiz et al., 2000; Colon-Cesario et al., 

2006), we sought to determine if Nurr1 expression can affect the spines of MSNs 

without exposure to L-DOPA. Parkinsonian rats were generated as in the 

electrophysiology experiment, and sacrificed three weeks following striatal vector 

delivery of rAAV-Nurr1 (n=4) or rAAV-GFP (n=3). These animals did not received L-

DOPA at any time, allowing us to determine the effect of Nurr1 on DA-depleted MSNs 

independent of drug treatment. The striatum was processed using Golgi-Cox 

impregnation to visualize the spines. Spine analysis showed there was a significant 

decrease total spine density in rAAV-Nurr1 treated animals (rAAV-Nurr1 total 

spines/10µm=4.85±0.10, rAAV-GFP total spines/10µm=5.98±0.31; t(5)=3.68, p<0.05) 

(Figure 4.4A). This difference was not specific to either the proximal or distal portions of 

the dendrite (rAAV-Nurr1 proximal spines/10µm=3.54±0.31, distal 

spines/10µm=5.58±0.41; rAAV-GFP proximal spines/10µm=4.24±1.45, distal 

spines/10µm=6.89±0.74). This indicates that Nurr1 can exert spine changes in MSNs 

and suggests its role in plasticity changes accompanying LID. Additionally, this effect is 

L-DOPA independent, showing that the capability of Nurr1 to induce spine changes 

does not require DA signaling. 

We additionally compared spine morphology between rAAV-Nurr1 and rAAV-GFP 

animals. Spine morphology can be an indicator of synaptic strength and spine dynamics  

 

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Figure 4.4 Nurr1-induced alterations in dendritic spine density and morphology 

(A) Dendritic spine density of MSNs in the lesioned hemisphere of rats treated with 
rAAV-Nurr1 or rAAV-GFP. Nurr1 expression caused a decrease in total number of 
spines. (B) rAAV-Nurr1 expression induced a decrease in both thin and branched type 
spines compared to rAAV-GFP controls. (C) Representative image of dendritic spines 
on Golgi-Cox stained MSNs. Scale bar=50µm. 

 

 

 

 

 

 

 

 

176 

 

(Maiti et al., 2015). We found that ectopic Nurr1 expression lead to a change in two 

spine classes. rAAV-Nurr1 animals displayed significantly fewer thin type spines than 

controls (rAAV-Nurr1 thin spines/10µm=3.38±0.09, rAAV-GFP thin 

spines/10µm=4.48±0.43; t(5)=2.98, p<0.05). Thin spines are typically transient and more 

likely to be formed and eliminated over time, unlike thicker spines that are more stable 

(Holtmaat et al., 2005). This suggests that Nurr1 could be influencing spine dynamics by 

altering turnover rates. We also observed fewer branched (also known as cupped or 

bifurcated) spines in rAAV-Nurr1 (branched spines/10µm=0.15±0.01) than in rAAV-GFP 

animals (branched spines/10µm=0.23±0.02; t(5)=3.35, p<0.05) (Figure 4.4B). Branched 

spines are mature and much less dynamic than thin spines (Hering & Sheng, 2001). No 

difference between groups was observed with stubby or mushroom spines. The 

decrease in thin and branched spines with Nurr1 expression suggests that the 

transcription factor can independently alter spine dynamics and turnover. This 

implicates Nurr1 as a molecular regulator of the maladaptive spine plasticity that has 

been shown in animal models of LID. 

Discussion 

We studied the electrophysiology of the striatum with ectopic Nurr1 expression in order 

to understand how Nurr1 may be influencing LID. Our study shows that Nurr1 

expression alone—independent of L-DOPA—can induce an LID-like signaling in the 

striatum as shown with LFP and single cell recordings. These animals were not 

chronically treated with L-DOPA, and thus the low dose given at the time of recording 

did not induce AIMs. This allowed us to determine that Nurr1 changes the activity of the 

striatum overall and in individual neurons independently, and its effect on LID 

 

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development is dependent on expression levels, not L-DOPA administration. The fact 

that rAAV-Nurr1 injected animals showed a near identical activity profile to rAAV-naïve, 

LID+ animals indicates Nurr1 has an active role in striatal changes associated with LID.  

Additionally, we have shown that Nurr1 can independently induce changes in MSN 

architecture by affecting spine dynamics. Ectopic Nurr1 expression caused alterations in 

spine density and morphology, which have been shown previously to be linked with LID 

(Zhang et al., 2013; Fieblinger et al., 2014; Nishijima et al., 2014; Suárez et al., 2014). 

Interestingly, our study showed a decrease in total spine density in neurons ectopically 

overexpressing Nurr1. This is contrary to much of the literature investigating spine 

dynamics in LID, which have shown an overall increase in spine density associated with 

the behavior. However, some studies have shown that LID is associated with a 

decrease in spine density exclusively on dMSNs (Fieblinger et al., 2014; Nishijima et al., 

2014; Suárez et al., 2014). As our study design did not allow for differentiation between 

dMSNs and indirect pathway MSNs (iMSNs), it is possible that through random 

sampling we selected primarily dMSNs. This would suggest that Nurr1 has an impact on 

the LID-associated spine loss in dMSNs. Alternatively, Nurr1 expression could lead to a 

differential effect on spine density between the two pathways that requires L-DOPA 

administration.  

Our data implicate Nurr1 as a molecular factor involved in promoting these maladaptive 

changes. Nurr1 has previously been shown to be heavily involved in learning and 

memory-associated plasticity in the hippocampus (Peña de Ortiz et al., 2000; Colon-

Cesario et al., 2006; Hawk et al., 2012). We have shown that Nurr1 can similarly affect 

plasticity when abnormally expressed in MSNs. Another NR4A family member Nur77 

 

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has been shown to induce spine loss in pyramidal neurons (Chen et al., 2014). 

Members of the NR4A family can form heterodimers to promote transcription (Maira et 

al., 1999). Since Nur77 is endogenously expressed in MSNs, it is possible that during 

abnormal Nurr1 induction in LID, an interaction between Nurr1 and Nur77 could 

promote transcriptional changes that induce the spine loss we observed in our 

experiments.  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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LITERATURE CITED 

 

 

 

 

 

 

 

 

 

 

 

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Bagetta, V., Ghiglieri, V., Sgobio, C., Calabresi, P., & Picconi, B. (2010). Synaptic 

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Chapter 5: Discussion 

Introduction 

In this dissertation, we have addressed the hypothesis that LID is caused both by 

abnormal 5-HTergic DA release as well as maladaptive expression of Nurr1 in the 

striatum. First, I demonstrated the role of the DRN in LID development, and answered 

previously unexplored topics that provide compelling support for the 5-HT hypothesis of 

LID. I achieved this by virally expressing the DA autoreceptor in the DRN and observing 

changes in AIM behavior and DA signaling in the striatum. My second objective was to 

characterize postsynaptic molecular changes in MSNs that contribute to LID, specifically 

by determining that aberrant Nurr1 expression can induce LID. Finally, I demonstrated 

that Nurr1 may be able to influence LID by maladaptively altering striatal physiology 

through its activity alone, without L-DOPA. Together, the Aims of the present 

dissertation intend to close multiple knowledge gaps in the etiology of LID. There is an 

unmet clinical need for better management strategies for LID. Our hope with this 

dissertation is to shed light on integral aspects of LID development to broaden our 

understanding and lead to new therapeutic strategies. 

Chapter 2: Inhibiting dorsal raphe-mediated dopamine release with AAV blocks 

LID development 

I. 

Aim 1 findings, in brief 

In Chapter 2, we showed LID development can be blocked by virally expressing the D2 

autoreceptor, D2Rs, in 5-HTergic neurons of the DRN. Animals treated with rAAV-D2Rs 

showed a complete inhibition of LID development without losing the anti-akinetic effects 

 

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of L-DOPA. D2Rs expression was able to block the priming event required for LID 

development, as shown by the lack of AIMs expressed in these animals when treated 

with DA receptor specific agonists. Using in vivo microdialysis, we showed a marked 

decrease in striatal DA efflux in animals treated with rAAV-D2Rs. This is the first direct 

evidence showing that inhibition of DA release from 5-HT neurons is directly related to 

LID amelioration. We additionally showed that D2Rs is functionally active in DRN 

neurons, as D2 agonist treatment caused reduced firing rates in transduced neurons. 

Together, this chapter confirms the 5-HT hypothesis of LID and shows that viral 

mediation of DRN neurons effectively blocks LID development. 

II. 

Study implications and future directions 

The 5-HT hypothesis of ‘false neurotransmission’ from DRN neurons has been a long-

standing idea in the LID research community (Cheshire & Williams, 2012; De 

Deurwaerdère et al., 2016). There have been a number of strong studies that have 

supported this hypothesis and rationalized clinical trials modulating the 5-HT system in 

LID. The most compelling evidence supporting this hypothesis is that ablation of the 

DRN can eliminate LIDs completely (Carta et al., 2007; Eskow et al., 2009). Many 

studies using 5-HT receptor agonists in animal models have shown that 

pharmacological modulation of these neurons is also effective at reducing LID (Bezard 

et al., 2013; Politis et al., 2014; Ghiglieri et al., 2016; Meadows et al., 2017). However, 

until our study, no direct evidence has been offered that shows unequivocally 

modulating 5-HT neurons reduces abnormal DA release and effects LID severity. Our 

study has closed this knowledge gap by directly measuring a reduction in DA release 

with 5-HT modulation. Additionally, this is the first study to our knowledge utilizing 

 

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DAergic modulatory factors in 5-HT neurons in order to regulate their activity. While 

there have been a number of clinical trials aimed at modulating 5-HT activity using 5-HT 

receptor agonists, these studies have not shown dramatic improvements, and in some 

cases actually caused worsening PD symptoms (Kannari et al., 2002; Olanow et al., 

2004; Iravani et al., 2006; Cheshire & Williams, 2012). The successful blockade of LID 

development in our model implicates D2Rs gene therapy as a potential therapeutic 

strategy for modulating 5-HT neurons in a non-pharmacological way. Our study is the 

first of its kind, however, and further characterization would be necessary to determine if 

this type of therapy would be a viable clinical option, as many 5-HT agonists also 

showed highly promising preclinical results. Still, the precedent of AAV-based clinical 

trials in PD as well as the rationale and use of 5-HT agonists for LID treatment make 

D2Rs an interesting candidate for therapeutic exploration. 

There are a number of things that deservedly should be researched further with this 

approach. Firstly, a critical understand on how D2Rs expression impacts 5-HTergic 

signaling will be essential moving forward. It has been shown previously that L-DOPA-

derived DA displaces 5-HT in vesicles, suggesting that L-DOPA therapy itself 

downregulates global 5-HT signaling from DRN neurons (Waldmeier, 1985). This 

contributes further to LID as there is less activation of the 5-HT autoreceptors, causing 

over activity of these neurons and thus continual DA release. However, our treatment 

with D2Rs reduces the activity and DA release from these neurons when treated with L-

DOPA. While we did not observe changes in 5-HT efflux in the striatum, we were not 

directly activating 5-HTergic signaling during the recording period. Further analysis 

 

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looking at how D2Rs expression impacts 5-HT release in different brain regions would 

be necessary to determine if D2Rs negatively impacts the 5-HT system. 

Our viral injections broadly targeted the entire DRN. However, not all DRN 5-HTergic 

neurons project to the striatum, the anatomical hub of LID. While detailed mapping of 

DRN projections is limited, it is clear that there are discreet cell populations that project 

to certain nuclei, including the striatum (Imai et al., 1986). A logical next step would 

therefore be to specifically target these neurons that project to the striatum, rather than 

the DRN as a whole. As DRN neurons release DA on other nuclei such as the 

substantia nigra, hippocampus, and prefrontal cortex (Navailles et al., 2010) it is 

important to determine if all DRN neurons or only those projecting to the striatum are 

involved in LID. There are a few ways this could be achieved. One approach would be 

to utilize a split rAAV vector and two-site delivery (Hirsch et al., 2016). Split vectors 

have been developed to circumvent the small packaging capacity of AAV. A transgene 

can be split between two vectors and packaged separately. When a cell is transduced 

with both vectors, the genomes naturally join via homologous recombination (Hirsch et 

al., 2016). For this application, one vector would be delivered to the DRN and the other 

to the striatum. If the striatal virus delivered was an rAAV pseudotype designed for 

efficient retrograde transport, this approach would result in the full transgene being 

expressed only in DRN cells that project to the striatum. The D2Rs gene could be 

reengineered into a split vector and used in this way to determine if the strength of its 

anti-LID effect remains when expressed specifically in these neurons. This technique 

could also be used with a split designer receptor exclusively activated by designer drugs 

(DREADD). Further, this design would allow for specific ablation of striatal-projecting 

 

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DRN neurons by introducing vectors expressing split forms of apoptotic-promoting 

proteins, such as Caspase. As full ablation of the DRN has been shown to block LID 

(Carta et al., 2007), this specific lesion would definitively determine if only striatal-

projecting DRN neurons are involved in LID. 

We showed that D2Rs DRN expression can effectively block L-DOPA priming by 

promoting more physiologic DA release on MSNs. This is evident in the fact that L-

DOPA-primed rAAV-D2Rs treated with direct DA agonists do not develop AIMs. As the 

protective mechanism of D2Rs expression is a presynaptic mechanism, future 

experiments should challenge D2Rs expressing animals that have not been treated with 

L-DOPA with DA agonists. I would hypothesize that these rats would develop severe 

AIMs similar to control animals, as the DRN signaling is not involved in DA receptor 

activation mediated by direct agonists. Additionally, as DA receptor agonists are known 

to cause dyskinesia in parkinsonian models and patients (Kuno, 1997; Rascol et al., 

2001), it is likely that if we had extended DA agonist treatments in our behavioral cohort, 

severe AIMs would have eventually developed in the D2Rs—expressing animals as they 

became primed. 

In this study, we delivered rAAV prior to treating with L-DOPA and establishing LID and 

successfully blocked LID development. This brings out the natural question: can 

delivering rAAV-D2Rs to an animal with established dyskinesia revert the phenotype? 

Answering this question would have obvious clinical implications, as a patient receiving 

therapy would have been exposed to L-DOPA and developed LID. Thus, this reversal 

experiment would be very informative to determine if this approach could translate 

clinically. This study could be designed nearly identically to the study we performed, 

 

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only changing the order of treatments. Lesioned animals would be treated with L-DOPA 

to establish LID. Only after this would we delivery rAAV. If D2Rs is capable of reversing 

LID, we would expect AIM scores to drop when these animals were reintroduced to L-

DOPA. 

Chapter 3: Nurr1 is a molecular driver of LID development 

I. 

Aim 2 findings, in brief 

In Chapter 3, we tested the hypothesis that Nurr1 plays an active role in LID 

development. We showed this by inducing severe AIMs in rats that are typically 

resistant to LID with viral overexpression of Nurr1 in the striatum. While Nurr1 

overexpression did not further exacerbate LID in Fischer rats that are susceptible to 

sever LID development, the overexpression did induce severe AIMs in LID-resistant 

Lewis rats. These rats exhibited LID as severe as the Fischer rats. We next utilized 

direct DA receptor agonists to determine how pathway-specific activation affected Nurr1 

expression. We found that Nurr1 expression is dependent on direct pathway signaling, 

and that D1 activation is sufficient to induce abnormal Nurr1 expression in both dMSNs 

and iMSNs. Finally, we found that an shRNA against Nurr1 was not sufficient to inhibit 

LID development in susceptible rats. Overall, our study provides evidence that Nurr1 is 

a novel molecular driver of LID. 

II. 

Study implications and future directions 

LID development is a complex, multifaceted process involving thousands of molecular 

changes. Gene expression studies have shown drastic expression changes in 

thousands of genes in dyskinetic models, highlighting a range of cellular pathways that 

 

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influence LID (Konradi et al., 2004; Heiman et al., 2014; Sodersten et al., 2014; 

Charbonnier-Beaupel et al., 2015). Nurr1 has been previously identified as being 

upregulated in MSNs of dyskinetic mice (Heiman et al., 2014; Sodersten et al., 2014). 

Our study has showed that Nurr1 is a critical factor in LID development, and not a 

molecular byproduct with no active role in the behavior. This identification and 

characterization of a novel molecular driver of LID is an important addition to the current 

understanding of how LID develop. Identification of key genes involved in LID etiology 

allows for better understanding of their development, which is necessary for developing 

novel treatment strategies. Additionally, we believe that our study demands a critical 

reappraisal of new Nurr1-based therapeutics intended to deliver neuroprotection PD. 

Pharmacological strategies to promote Nurr1 activity in the SNc could concurrently 

promote maladaptive activity in the striatum. It is possible that this could lead to earlier 

development or more severe LID in patients. Thus, caution moving forward with these 

strategies is warranted. Studies interrogating the effect of Nurr1 agonists on LID 

development in animal models would describe the risk associated with these therapies. 

There is convincing evidence in the literature that can explain how this abnormal Nurr1 

expression occurs in LID. The MAPK signaling pathway through and activation of 

ERK1/2 is well established as being hyperactive in LID (Fiorentini et al., 2013). In vitro 

studies have found that ERK1/2 signaling is required for the transcription of NR4A 

receptors including Nurr1 (Darragh et al., 2005). Nurr1 expression is regulated by a 

number of MAPK downstream effectors, including CREB and MSK-1 (Saucedo-

Cardenas et al., 1997; McEvoy et al., 2002). In fact, inhibition of the MAPK cascade 

completely blocks activity-induced Nurr1 expression (Darragh et al., 2005). The 

 

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persistent MAPK signaling and ERK1/2 activation that occurs in LID is thus sufficient to 

promote Nurr1 expression in MSNs. 

The MAPK cascade is, of course, not specific to a dyskinetic state and activated under 

normal physiological conditions in MSNs (Girault, 2012; Hutton et al., 2017). This 

physiological signaling does not, however, lead to Nurr1 expression in MSNs. However, 

the understanding of molecular changes in LID and Nurr1 regulation suggest that 

histone modifications may be involved in this abnormal allowance of Nurr1 expression. 

MSK-1 induces histone phosphorylation—specifically phosphorylation of histone H3—

following its activation by extracellular stimuli (Kim et al., 2008). Notably, one study 

found dramatic increases in H3 phosphorylation mice treated with L-DOPA (Sodersten 

et al., 2014). Importantly, the researchers found that Nurr1 gene regulation was 

associated with specific H3 phosphorylation, and that Nurr1 transcript drastically 

increased in the striatum of dyskinetic mice (Sodersten et al., 2014). Together, this 

evidence suggests that histone modification by MSK-1 is essential in LID-associated 

gene expression, including Nurr1. 

Our study using shRNA to block Nurr1 upregulation failed to inhibit LID development. 

This would suggest that Nurr1 is not required for LID development and that there are 

functionally redundant pathways. This is contradictory to our study showing that Nurr1 is 

required to induce LID in resistant Lewis rats. A possible explanation is that limitations 

with our shRNA approach did not achieve sufficient knockdown of Nurr1. While we 

achieved strong transduction with our vectors, it is near impossible to transduce 100% 

of neurons. Indeed, while our shRNA successfully blocked Nurr1 expression in 

transduced cells, we observed LID-associated Nurr1 upregulation in non-transduced 

 

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neurons (Figure 5.1). Because of this, it is likely that the critical threshold of Nurr1 

expression to induce LID was achieved in these animals. This would reflect in the 

behavior data, as we saw severe LID development even with shRNA treatment. 

Currently, we do not know what the minimum amount of Nurr1 required for LID 

development is. Until this is characterized further, we can assume that the number of 

non-transduced neurons able to upregulate Nurr1 was sufficient to induce LID in our 

study. An shRNA approach may not be the strongest way to address the hypothesis 

that Nurr1 silencing can attenuate LID, and thus further studies using more robust 

knockdown approaches would be useful. Full genetic silencing of Nurr1 in the striatum 

using a CRISPR approach would ensure complete knockdown of the gene. However, 

this approach would require viral delivery of the CRISPR cassette, and would result in 

the same limitation as the shRNA if not enough cells are transduced. A conditional 

knockout model may be most viable, which would allow specific ablation of Nurr1 by 

using a tamoxifen-inducible Cre driven by MSN-specific regulatory elements. A similar 

approach has previously been used to selectively knockout Nurr1 in DAergic neurons 

(Kadkhodaei et al., 2013). This approach would ensure Nurr1 silencing in the entire 

striatum and allow us to determine if LID can be blocked in susceptible animals by 

disallowing Nurr1 expression. This model additionally would allow us to determine if 

silencing Nurr1 in established dyskinetic animals can revert AIM behavior. 

Our study found that Nurr1 expression in both the direct and indirect pathway is 

dependent on D1 signaling. While dyskinesia-induced Nurr1 expression occurs in both 

classes of MSNs, we did not characterize the role of Nurr1 in dMSNs versus iMSNs. In 

order to determine if Nurr1 expression has a differential effect on LID in dMSNs or  

 

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Figure 5.1 Nurr1 expression in shRNA treated animals 

(A-D) IHC for Nurr1 (red, B), transduction marker GFP (green, C) and dapi (blue, D) in the striatum of an animal treated 
with rAAV-shRNA-Nurr1. No Nurr1 was observed to colocalize with GFP, and Nurr1 was only observed in neurons that 
were not transduced (arrows). (E-H) IHC for Nurr1 (red, E), transduction marker GFP (green, G), or dapi (blue, H) in the 
striatum of animals treated with the control vector rAAV-shRNA-SCR. Nurr1 can be seen colocalizing with GFP (arrows). 
Scale bar=50µm.

 

 

194 

 

iMSNs, a Cre-dependent genetic vector could be used to drive expression specifically in 

D1 or D2 expressing neurons. Cre-dependent FLEx vectors can be used to achieve 

expression of the viral transgene only in cells expressing Cre (Schnütgen et al., 2003). 

These vectors could be packaged into rAAV and delivered to the striatum of transgenic 

rats with Cre expression driven by the promoter of either the D1 receptor or D2 

receptor. Animals would then be treated with L-DOPA to monitor changes in AIMs due 

to specific Nurr1 expression. This would address the hypothesis that Nurr1 expression 

primarily effects LID through the direct pathway. 

Chapter 4: Nurr1 expression induces an LID-like striatum 

I. 

Aim 3 findings, in brief 

In Chapter 3, we tested the hypothesis that Nurr1 expression induces abnormal 

changes in the firing patterns and ultrastructure of MSNs. We characterized the effect of 

Nurr1 on MSNs independent of L-DOPA. Using rAAV-mediated gene modulation, we 

showed altered corticostriatal activity in the striatum transduced with rAAV-Nurr1. These 

animals exhibited a potentiation in response to continuous stimuli, where control 

animals showed a depotentiation over time. Additionally, we found that the firing of rate 

individual striatonigral neurons was increased in rAAV-Nurr1 treated animals. In both of 

these studies, the activity profile of L-DOPA naïve rAAV-Nurr1 treated rats mimicked the 

behavior of neurons from a dyskinetic rat. Finally, we found that ectopic Nurr1 

expression reduced dendritic spine density and lowered the number of thin and 

branched type spines. Taken together, this aim shows that Nurr1 can induce activity 

and morphology changes in MSNs that promote LID. 

 

195 

 

II. 

Study implications and future directions 

A number of previous studies have found a strong link between altered MSN activity 

and morphology and LID (Picconi et al., 2003; Bastide et al., 2015; Fieblinger & Cenci, 

2015). Our study provides evidence that Nurr1 is a molecular instigator of these 

alterations. Importantly, Nurr1 is able to induce these changes independent of L-DOPA, 

suggesting that Nurr1-regulated activity and transcription in MSNs does not require 

DAergic signaling. This evidence shows that once Nurr1 expression is induced in 

MSNs, it can promote these changes through its transcriptional activity. 

We showed that individual dMSNs display increased firing rates with ectopic Nurr1 

overexpression. Antidromic labeling cannot reliably identify iMSNs, as stimulus of the 

GPe would lead to inadvertent activation of dMSN fibers passing through the structure 

(Threlfell et al., 2009). As the antidromic labeling technique could only identify dMSNs, 

future studies should evaluate activity changes in iMSNs as well. This has been done 

recently in parkinsonian and dyskinetic mice using optogenetic identification of D1 or D2 

expressing neurons in the striatum (Ryan et al., 2018). A similar approach could be 

employed in animals virally overexpressing Nurr1. An alternative approach would be to 

drive DREADD expression in dMSNs or iMSNs. Neurons could then be recorded from 

and identified by applying a designer drug to determine the neuron class. This or 

optogenetic identification would allow us to determine how Nurr1 alters the activity of 

both pathways, with or without L-DOPA. iMSNs show a decrease in firing potential in 

dyskinetic mice (Ryan et al., 2018), thus we hypothesize that iMSNs transduced with 

Nurr1 would show reduced firing rates independent of L-DOPA. 

 

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We saw a marked decrease in total spine density as well as thin and branched class 

spines in animals with rAAV-Nurr1 delivered to the striatum. These results indicate that 

Nurr1 can impact dendritic spine dynamics in MSNs, independent of L-DOPA. This is 

the first characterization of the impact of Nurr1 on dendritic spines of MSNs. Previous 

work has found that NR4A family member Nur77 can reduce spine density in 

hippocampal neurons, a similar effect we observed in MSNs with Nurr1 overexpression 

(Chen et al., 2014). As it is known that NR4A family members can heterodimerize to 

regulate transcription (Maira et al., 1999) Nurr1 could be impacting spine dynamics 

through its interaction with Nur77 in these neurons. Nur77 expression additionally has 

been shown to impact LID development in parkinsonian rats (Rouillard et al., 2018). As 

Nur77 and Nurr1 are not endogenously coexpressed in MSNs, their presence together 

in LID suggests that they coregulate maladaptive gene expression that drives LID. 

Further studies to determine if a Nurr1/Nur77 interaction occurs in MSNs and its effect 

on LID-associated gene transcription would add compelling evidence to support that 

NR4A nuclear orphan receptors are molecular drivers of LID. 

LID-associated alterations in MSNs, from gene expression to spine dynamics to activity, 

are not consistent between the direct and indirect pathway. The use of Golgi-Cox 

impregnation of neurons does not allow for differentiation between the two neuron 

classes, and while we did observe Nurr1-induced changes in spine density and 

morphology, our study does not inform how these changes are reflected in each 

pathway. Previous studies have used retrograde labeling using neuron-filling dye 

injected into the SNr to identify dMSNs or the GPe to identify iMSNs (Nishijima et al., 

2014). Other groups have used transgenic mouse models with fluorescent proteins 

 

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expressed under the D1 or D2 promoter to differentiate between pathways (Fieblinger et 

al., 2014; Suárez et al., 2014). Fluorescent neurons can be identified in slices and 

injected with dyes to fill the neuron and spines. Using the same experimental design 

from our study, employing one of these approaches would allow us to determine the 

effect of Nurr1 expression on each specific class of MSNs. We hypothesize that Nurr1 

induces LID-associated spine loss in dMSNs and spine increases on iMSNs, and that 

this effect is independent of L-DOPA. In the present study, we saw global decreases in 

spine numbers when not differentiation between neurons. Assuming that we sampled 

dMSNs and iMSNs equally, our current data would suggest that Nurr1 reduces spines in 

both neuron classes. This could be confirmed with these approaches for differentiating 

dMSNs and iMSNs. 

In the present dissertation, we studied the downstream effects on behavior and striatal 

physiology that are caused by Nurr1 overexpression. However, as Nurr1 is a 

transcription factor, it is likely not the direct effector of these changes, rather a regulator 

of genes that have direct impact. While our study characterized behavioral and 

physiological changes associated with Nurr1, we did not explore the downstream 

effectors regulated by Nurr1 and their effect on LID. Further studies identifying the 

Nurr1-regulated molecular pathway affecting LID are warranted. The data from this 

dissertation would suggest that the effector genes modulate synaptic plasticity through 

spine dynamics and signaling activity. Indeed, multiple genes regulated by Nurr1, 

including BDNF, the glutamate receptor GluR2, and kinesin family member KIF5C 

(Volpicelli et al., 2007; Do, 2014) can effect dendritic spines and synaptic transmission 

(Passafaro et al., 2003; Kellner et al., 2014; Willemsen et al., 2014). Pathway analysis 

 

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has shown that Nurr1 affects a wide array of cellular pathways, including MAPK 

signaling, cellular response to stimuli and stress, and cell growth and architecture (Do, 

2014). Further characterization of genes regulated by Nurr1 in LID is necessary to 

better understand how its expression promotes LID. 

Final remarks 

This dissertation offers new evidence exhibiting the critical role of both the 5-HT system 

and Nurr1 in LID development. Dysregulated DA release from 5-HT neurons of the DRN 

promote a host of changes in MSNs, including the expression of the newly identified 

driver of LID, Nurr1. There are many networks and pathways involved in LID, and this 

dissertation sheds new light on how both presynaptic input and abnormal induction of 

one gene can influence the behavior. This dissertation supports a multifaceted 

mechanism for LID where abnormal DA release from 5-HT neurons induces Nurr1 

expression in MSNs, which changes striatal physiology in a manner that promotes LID 

(Figure 5.2). Additionally, our studies provide compelling rational for the clinical 

development of novel LID management strategies targeting DRN neurons. Further 

characterization of LID-associated pathways, including aberrant Nurr1 expression, will 

be important as the field moves closer towards effective therapies for both PD and LID. 

The results presented in this dissertation have closed multiple gaps in knowledge in LID 

etiology, highlighting the impact of 5-HT regulation in LID and the role of Nurr1 

expression in LID-associated plasticity. 

 

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Figure 5.2 Proposed mechanism of LID development 

(A) This schematic shows the proposed mechanism of LID development supported by 
the findings of this dissertation. Serotonin neurons of the dorsal raphe nucleus (DRN) 
convert L-DOPA to DA and indiscriminately release the neurotransmitter onto striatal 
MSNs. Hypersensitive MSNs respond to DAergic stimulation by maladaptively 
upregulating the transcription factor Nurr1, which induces changes in dendritic spines 
and MSN signaling activity. These aberrant changes promote LID. (B) LID development 
can be blocked by regulating DA release from DRN neurons. Ectopic expression of the 
DA autoreceptor D2Rs in the DRN regulates striatal DA efflux, disallowing priming and 
maladaptive expression and synaptic plasticity required for LID development. 

 

 

 

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