ANALYSIS OF Pcp-2/L7 GENE EXPRESSION AND FUNCTION
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Yelda Serinagaoglu, B. Sc.
* * * * *
The Ohio State University 2007
Dissertation Committee:
Dr. John Oberdick, Advisor Approved by
Dr. Heithem El-Hodiri
Dr. Anthony P. Young Advisor Dr. Mike Zhu Molecular, Cellular, and Developmental Biology Graduate Program
ABSTRACT
The mouse Purkinje cell protein-2 (Pcp-2/L7, herein called L7) gene is specifically and abundantly expressed in cerebellar Purkinje cells and in retinal bipolar neurons. Recent studies suggested that L7 functions in the tuning of P/Q type Ca2+ channels by the modulation of G-protein coupled
receptors. The goals of the present studies were to extend our
understanding on L7 function and control of L7 gene expression. To better
understand function of the L7 protein, we carried out a variety of
behavioral tests to compare wild type and L7 knock-out (L7KO) mice. We
show that L7KO mice have improved performance on accelerating rotarod
and display sexually dimorphic sensorimotor behavioral changes. Our
results support the idea that cerebellum plays a role in sensorimotor
gating, and it functions in the mediation of sensory response, which is a
non-traditional role for the cerebellum. We then expanded our studies to
investigate L7 gene expression in the mouse cerebellum and eye. Here
we report that a third isoform of L7, which we call L7C, is the only form
expressed in the eye, while two isoforms, L7A and L7B, are abundantly
found in the cerebellum. To determine the molecular and genetic ii mechanisms of the Purkinje cell-specific expression of L7, we carried out
in vivo analyses where we show that L7 structural gene does not have any
Purkinje cell-enhancement activity, and likely contains significant
repressive activity. 0.9 kb L7 proximal promoter, on the other hand, acts as an enhancer to direct expression in the cerebellar Purkinje cells. This enhancer activity requires the position-dependent action of the structural gene. In the present study, we show that retinoic acid receptor related orphan nuclear receptor-alpha (RORα) is a major activator of L7 gene expression in vivo and in vitro. Work presented here will allow us to better understand the relationship between the cellular physiology and the animal behavior. Although previous studies with RORα focused on its role during the cerebellar development, our studies show that RORα is no longer just a cerebellar developmental control molecule, but a key determinant of cerebellar physiology, as it activates L7, which in turn modulates Ca2+ channels.
iii
Dedicated to my family
iv
ACKNOWLEDGMENTS
First and foremost I would like to thank my advisor Dr. John Oberdick, for his advice, support, patience, and guidance throughout my graduate study. I would like to express my gratitude for my committee members Dr. Heithem El-Hodiri, Dr. Anthony Young, and Dr. Mike Zhu for all the help and advice. I would also like to thank Dr. David Saffen for the advice and support during my candidacy exam. I am deeply indebted to Dr. Dave Bisaro for giving me the opportunity to be a member of the MCDB program. My deepest thanks go to Jan Zinaich, who always greeted me with her warm smile and has always helped me. I would like to express special thanks to the past and current staff members of the Center for Molecular Neurobiology, Keri Bantz, Scott
Hines, Darlene Jackson, Juli Kanoski, Dave Long, Paula Monsma, and Laura Richter for taking care of everything for us in Rightmire Hall. I would like to thank the members of Rightmire Hall and Pressey Hall Animal Facilities for taking good care of mice. I am also grateful to Dr. Xin-An Pu for all she has done in our transgenic studies, and for taking time to answer my questions. I would especially like to thank past and current members of the Oberdick lab, Jamie Depelteau, Nichole Gebhart, Peijun Wu, Rui Zhang, and Xulun Zhang for sharing their knowledge and experience, and creating a fun and peaceful place to work at.
v Special thanks go to members of Dr. Tsonwin Hai’s lab, Dr. Tony Brown’s lab, Dr. Randy Nelson’s lab, and Dr. Mike Zhu’s lab for technical support, collaborations, and scientific/non-scientific conversations. I would like to send my appreciation to my friends in Turkey and in Germany, Fulya Akyar, Arzu Baysal, Aydan Işık, İpek Kayhan, and Yasemin Tosun, for cheering me up with phone calls and e-mails from overseas. I do not know how to express my deepest appreciation for Ozan Özkuşaksız, for endlessly giving me the hope for a much better future. My sincere thanks go to my friends in the USA, Sinan Doğan, Yeşim Çapa-Aydın, Tolga Aydın, Özlem Birgül, Gülsen Çolakoğlu, Suat Gümüşsoy, and İlter Sever, for being my friends and making my life much happier. I also thank Tankut Taylan for the long conversations and for patiently helping me with computer software. I am eternally grateful to my family for their endless love, support, and encouragement. My mother Ayşe, my father Mehmet, my sister Yeşim and my brother-in-law Ahmet, words are not enough to thank you. My caring fiancé David, your love and support helped make this possible. Thank you for always believing in me, and keeping everything in perspective. Last but not least, I would like to thank Prufrock, for keeping company with me during the long and late hours.
vi VITA
December 26, 1977 ...... Born – Bursa, Turkey
2001 ...... B.Sc. Molecular Biology and Genetics, Middle East Technical University, Ankara, Turkey
2001 – present...... Graduate Research Associate, The Ohio State University
PUBLICATIONS
Yelda Serinagaoglu, Rui Zhang, Yufang Zhang, Linda Zhang, Greg Hartt, Anthony P. Young, and John Oberdick, (2007). A promoter element with enhancer properties, and the orphan nuclear receptor RORalpha, are required for Purkinje cell-specific expression of a G(i/o) modulator. Mol Cell Neurosci 34, 324-342.
FIELDS OF STUDY Major Field: Molecular, Cellular, and Developmental biology
vii TABLE OF CONTENTS
Page
ABSTRACT ...... ii
ACKNOWLEDGMENTS...... v
VITA ...... vii
LIST OF FIGURES ...... xiii
LIST OF TABLES ...... xv
ABBREVIATIONS ...... xvi
CHAPTER 1
INTRODUCTION...... 1 1.1 SUMMARY ...... 1 1.2 CEREBELLUM ...... 1 1.2.1 Functions of the cerebellum ...... 2 1.2.2 Structure of the cerebellum ...... 3 1.2.3 Cerebellar circuitry ...... 5 1.2.4 Development of the cerebellum ...... 6 1.2.5 Cerebellar mutant mice: Natural mutations...... 7 1.3 L7 IS A CEREBELLAR PURKINJE CELL SPECIFIC GENE ...... 8 1.3.1 Discovery of L7 gene...... 9 1.3.2 Phenotype of L7KO mice ...... 9 1.3.3 Structure and function of L7 protein ...... 11 1.3.3.1 Heterotrimeric G proteins...... 11 1.3.3.3 Modulation of voltage-dependent Ca2+ channels by G proteins ...... 14 1.3.3.4 L7 modulates G-protein-coupled receptor-mediated inhibition of Ca2+ channels in a dose-dependent manner...... 15 1.3.4 Structure of L7 gene...... 15 1.3.5 Analysis of L7 gene expression in the mouse cerebellum ...... 17 1.3.6 Regulation of L7 gene expression...... 19 1.4 NUCLEAR RECEPTOR SUPERFAMILY...... 21 1.4.1 Orphan nuclear receptors...... 24 1.4.2 Retinoic acid receptor-related orphan nuclear receptors (RORs)...... 25 1.4.3 RORα ...... 27 1.4.3.1 Discovery of RORα ...... 27 1.4.3.2 Genomic structure of RORα isoforms...... 28 1.4.3.3 Expression patterns of RORα isoforms ...... 28 viii 1.4.3.4 Protein structure of RORα isoforms...... 29 1.4.3.5 Characterization of ROR response elements ...... 30 1.4.3.6 Staggerer phenotype and RORα-/- mice ...... 30 1.4.3.7 Genes activated by RORα ...... 33 1.4.3.8 Activation of L7 by RORα ...... 36 1.4.3.9 Transcriptional control by RORα...... 38 1.4.3.9.1 Ligand-dependent mechanism: Possible ligands for RORα...... 39 1.4.3.9.2 Ligand-independent mechanism ...... 40 1.4.3.9.3 Possible mechanism for regulation by phosphorylation ...... 41 1.4.3.9.4 Role of CaMKIV in the regulation of RORα-mediated transcription .41 1.5 OVERVIEW OF THESIS WORK ...... 42
CHAPTER 2
ROLES OF THE L7 PROXIMAL PROMOTER/ENHANCER AND THE STRUCTURAL GENE IN CONTROL OF GENE EXPRESSION IN VIVO ...... 43 2.1 ABSTRACT...... 43 2.2 INTRODUCTION ...... 45 2.3 MATERIALS AND METHODS...... 49 2.3.1 Mouse strains ...... 49 2.3.2 DNA Constructs for transgenic mice ...... 49 2.3.2.1 6k-L7 Enhancer Test construct...... 49 2.3.3 Preparation of DNA for pronuclear injection...... 50 2.3.4 Tail DNA extraction ...... 50 2.3.5 Genotyping ...... 50 2.3.6 Perfusion ...... 52 2.3.7 X-Gal staining...... 52 2.3.8 Cerebellum measurements ...... 53 2.3.9 Total RNA extraction ...... 54 2.3.10 DNase treatment ...... 54 2.3.11 Reverse Transcription (RT)...... 55 2.3.12 Polymerase chain reaction (PCR)...... 55 2.3.13 Real-Time PCR ...... 56 2.3.13.1 Determination of PCR efficiency for each primer set...... 57 2.3.13.2 Quantification of expression levels by Real-Time PCR ...... 58 2.4 RESULTS ...... 59 2.4.1 L7 proximal promoter/enhancer and the L7SGC are required in concert for Purkinje cell-specific expression ...... 59 2.4.2 Distal half of the L7 gene is required for high level expression in the cerebellum...... 65 2.5 DISCUSSION ...... 77
ix CHAPTER 3
ACTIVATION OF L7 GENE EXPRESSION BY RORα...... 84 3.1 ABSTRACT...... 84 3.2 INTRODUCTION ...... 86 3.3 MATERIALS AND METHODS...... 89 3.3.1 Mouse strains ...... 89 3.3.2 DNA Constructs...... 89 3.3.2.1 L7Prom2K ...... 89 3.3.3 Preparation of DNA for pronuclear injection...... 90 3.3.4 Tail DNA extraction ...... 90 3.3.5 Genotyping ...... 90 3.3.5.1 Promoter constructs...... 90 3.3.5.2 Staggerer mice...... 91 3.3.6 Perfusion ...... 92 3.3.7 X-Gal staining...... 92 3.3.8 Cell transfection assays ...... 93 3.3.8.1 Cell culture and transfection ...... 93 3.3.8.2 Visualization of LacZ expression ...... 94 3.3.8.3 Quantification of GFP fluorescence ...... 94 3.3.9 Electrophoretic mobility shift assay (EMSA) ...... 95 3.3.9.1 Preparation of nuclear extracts...... 95 3.3.9.2 In vitro translation...... 96 3.3.9.3 Probe preparation ...... 97 3.3.9.4 Preparation of EMSA gel ...... 98 3.3.9.5 EMSA...... 98 3.3.10 Chromatin immunoprecipitation (ChIP) and PCR ...... 99 3.3.10.1 ChIP ...... 99 3.3.10.2 PCR...... 101 3.3.11 In situ hybridization...... 102 3.3.11.1 Probe preparation ...... 102 3.3.11.2 Quantitation of radioactivity of α-35S UTP-labeled probe...... 103 3.3.11.3 Tissue and section preparation...... 103 3.3.11.4 Pretreatment of the slides...... 104 3.3.11.5 Hybridization ...... 104 3.3.11.6 Post-hybridization washing ...... 105 3.3.11.7 Dipping the slides...... 105 3.3.11.8 Developing the slides...... 106 3.3.11.9 Counterstaining...... 106 3.3.12 Western blotting ...... 106 3.4 RESULTS ...... 108 3.4.1 Conservation of transcription factor binding sites in rodent and human L7 promoters ...... 108 3.4.2 L7 promoter activation by RORα in cell culture and inhibition by L7 structural gene...... 111 3.4.3 RORα can bind to its putative binding site in the proximal promoter in vitro 115 x 3.4.4 RORα occupies the RORE in the proximal L7 promoter in vivo ...... 118 3.4.5 RORα controls both the overall level and medio-lateral pattern of L7 gene expression ...... 119 3.5 DISCUSSION ...... 127
CHAPTER 4
AN ANALYSIS OF L7 ISOFORMS EXPRESSED IN THE MOUSE CEREBELLUM AND EYE ...... 130 4.1 ABSTRACT...... 130 4.2 INTRODUCTION ...... 132 4.3 MATERIALS AND METHODS...... 138 4.3.1. Mouse strains ...... 138 4.3.2. Total RNA extraction ...... 138 4.3.3. DNase treatment ...... 138 4.3.4. Reverse Transcription (RT)...... 139 4.3.5. Polymerase chain reaction (PCR)...... 139 4.3.6. Real-Time PCR ...... 140 4.3.6.1. Determination of PCR efficiency for each primer set...... 141 4.3.6.2. Quantification of expression levels by Real-Time PCR ...... 142 4.3.7 Genotyping ...... 143 4.3.8. 5’-Rapid amplificaition of cDNA ends (5’-RACE) ...... 143 4.3.8.1. Reverse Transcription (RT)...... 144 4.3.8.2. Purification of cDNA...... 144 4.3.8.3. Recombinant terminal deoxynucleotidyl transferase (rTdT) tailing...... 144 4.3.8.4. Polymerase chain reaction (PCR) ...... 145 4.4. RESULTS ...... 146 4.4.1. L7 expression analysis in mouse cerebellum and eye...... 146 4.4.2. L7 expression analysis in human and mouse testis...... 153 4.4.3. L7 expression analysis in sg cerebellum...... 156 4.5. DISCUSSION ...... 160
CHAPTER 5
BEHAVIORAL ANALYSIS OF L7KO MICE ...... 163 5.1 ABSTRACT...... 163 5.2 INTRODUCTION ...... 165 5.3 MATERIALS AND METHODS...... 168 5.3.1 Mouse strains ...... 168 5.3.2 Tail DNA extraction ...... 168 5.3.3 Genotyping ...... 169 5.3.4 Total RNA extraction ...... 169 5.3.5 DNase treatment ...... 170 xi 5.3.6 Reverse Transcription (RT)...... 170 5.3.7 Polymerase chain reaction (PCR)...... 171 5.3.8 Real-Time PCR ...... 171 5.3.8.1 Determination of PCR efficiency for each primer set...... 172 5.3.8.2 Quantification of expression levels by Real-Time PCR ...... 173 5.3.9 Behavioral tests...... 174 5.3.9.1 Latency to Move...... 174 5.3.9.2 Open Field/Locomotor ...... 174 5.3.9.3 Grip Strength...... 175 5.3.9.4 Beam Walk...... 175 5.3.9.5 Light/Dark Preference...... 176 5.3.9.6 Hotplate...... 176 5.3.9.7 Rotating Rod ...... 177 5.3.9.8 Acoustic Startle Response (ASR)...... 177 5.3.9.9 PPI ...... 178 5.3.9.10 ASR Habituation ...... 178 5.3.9.11 Acoustic Threshold ...... 179 5.3.9.12 Elevated Plus Maze ...... 180 5.4 RESULTS ...... 181 5.4.1 Initial analysis of L7KO mice ...... 181 5.4.2 Behavioral analysis of L7KO mice ...... 182 5.4.2.1 L7KO mice have normal locomotor activity and anxiety behaviors ...... 182 5.4.2.2 L7KO mice show improved maximum performance on a motor learning test ...... 184 5.4.2.3 L7KO mice show altered sensory responsiveness...... 187 5.4.3 L7 is specifically expressed in cerebellum and retina, but not in sensory organs...... 194 5.5 DISCUSSION ...... 197
BIBLIOGRAPHY...... 201
xii LIST OF FIGURES
Figure Page
1.1 The structure of the cerebellar cortex...... 4 1.2 An overview showing the diversity of G protein-coupled receptors and signal transduction...... 13 1.3 Comparison of rodent and human L7 gene structure and gene expression...... 16 1.4 Comparison of two L7 gene constructs carrying 4 kb promoter ...... 19 1.5 Structural and functional illustration of nuclear receptors...... 22 1.6 Schematic representations of RORα gene products ...... 29 1.7 Comparison of the cerebella of a staggerer and a wild type mouse...... 32
2.1 Comparison of two L7 gene constructs carrying 4 kb promoter...... 46 2.2 Constructs used to study the transcriptional role of the structural gene and the promoter/enhancer in vivo...... 60 2.3 Analysis of L7-SGC function and L7 Enhancer Test: α6 promoter...... 71 2.4 Analysis of L7-SGC fusion and L7 Enhancer Test: hNOS1 promoter ...... 74 2.5 The distal half of the L7 structural gene is necessary for high level cerebellum expression...... 75 2.6 Comparison of Purkinje cell-specific expression patterns of different L7 constructs...... 78 2.7 Minimal promoter constructs for enhancer test in transgenic mice ...... 83
3.1 Comparison of rodent and human Pcp-2(L7) promoter/enhancers...... 109 3.2 Constructs used in the transfection assay...... 112 3.3 RORα induction of the L7 promoter/enhancer in vitro and the effect of the L7SGC...... 114 3.4 Binding of RORα to the promoter/enhancer site at -62 bp in vitro and in vivo...... 117 3.5 Regional effects of RORα in the cerebellum...... 121 3.6 Purkinje cell-specific expression in L7Prom-WT and L7Prom-ΔROR mice...... 122
xiii 4.1. Comparison of rodent and human L7 gene structure and gene expression...... 133 4.2. Comparison of L7 isoforms reported in mouse cerebellum and eye...... 134 4.3. Comparison of L7 isoforms reported in human cerebellum, testis and eye...... 135 4.4 RT-PCR analysis of L7 expression in mouse cerebellum and eye...... 146 4.5 5’-RACE analysis using RNA extracts from the cerebellum and the eye ...... 147 4.6 Sequence analysis of L7 gene in mice...... 149 4.7 Comparison of L7 isoforms expressed in mouse cerebellum and eye...... 151 4.8 Comparison of protein sequences of three L7 isoforms...... 151 4.9 RT-PCR analysis of expression of L7 isoforms in mouse cerebellum and eye...... 152 4.10 RT-PCR analysis of expression of L7 isoforms in mouse and human cerebellum, and mouse testis and liver...... 153 4.11 Sequence analysis of L7 gene in humans...... 154 4.12 RT-PCR analysis of expression of L7 in sg +/+, +/-, and -/- cerebellums...... 156 4.13 5’-RACE analysis using RNA extracts from the WT and sg cerebellums...... 159 4.14 RT-PCR analysis of expression of L7 isoforms in sg +/+, and -/- cerebellums ...... 159
5.1 Western blotting analysis of L7+/+, L7+/-, L7-/- cerebellums ...... 181 5.2 Locomotor activity and anxiety behaviors were compared...... 183 5.3 Motor coordination and motor learning tests were performed...... 185 5.4. Assessment of the sensory phenotype of L7 mutants by the acoustic startle response and hotplate tests...... 190 5.5 RT-PCR and Real-time PCR analysis of L7 expression in sensory organs of wild- types...... 196
xiv LIST OF TABLES
Table Page
2.1 Expression of α6, 0.9 kb L7 proximal promoter, and L7SGC fusion gene constructs in transgenic mice...... 67 2.2 Morphological measures of transgenic cerebella...... 72 2.3 Enhancer test using NOS1-L7SGC-LacZ base constructs...... 73
3.1 Effect of ΔROR enhancer mutation on transgene expression in the cerebellum. ...125
xv ABBREVIATIONS
α-35S-UTP Uridine 5'-[α-35S]thiotriphosphate α6 γ-aminobutyric acid type-A receptor α6 subunit γ-32P adenosine 5'-[γ-32P]triphosphate 3’UTR 3' untranslated region 5’-RACE 5’ rapid amplification of cDNA ends a6IL mα6IRES-LacZ aa amino acid AF activation function ANOVA analysis of variance AP4 activator protein 4 ATP adenosine triphosphate BGH bovine growth hormone bp base pairs BS brainstem BSA bovine serum albumin CaMKIV Ca2+/calmodulin-dependent protein kinase IV cDNA complementary deoxyribonucleic acid ChIP chromatin immunoprecipitation CMV cytomegalovirus coll/thal colliculi & thalamus COUP-TF chicken ovalbumin upstream promoter-transcription factor CRE cAMP-response element CT threshold cycle CTE C-term extension C-term carboxyl terminal CTP cytidine triphosphate dCTP deoxycytidine triphosphate DEPC-H2O diethylpyrocarbonate-water dGTP deoxyguanosine triphosphate DHR3 Drosophila hormone receptor 3 DMEM Dulbecco’s modified Eagle’s medium DMSO dimethyl sulfoxide DNA deoxyribonucleic acid DNase deoxyribonuclease dNTP deoxynucleotide ds double stranded DTT dithiothreitol E embryonic EDTA ethylenediamine tetraacetic acid EGFP enhanced green fluorescent protein EGL external granular layer EGTA ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′ tetraacetic acid EMSA electromobility shift assay FBS fetal bovine serum xvi GABA γ-aminobutyric acid GABAA γ-aminobutyric acid type-A GC granule cells GCL granule cell layer GDI guanine-nucleotide dissociation inhibitor GDP guanosine diphosphate GEF guanine nucleotide exchange factor GFP green fluorescent protein GL green lantern GPSM-4 G-protein signaling modulator-4 GTP guanosine triphosphate HEK human embryonic kidney HEPES 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid Hsp heat-shock promoter IgG immunoglobulin G IP immunoprecipitation IRES internal ribosome entry site kb kilo bases KCl potassium chloride KO knock-out L7enh L7 enhancer L7SGC L7 structural gene cassette LacZ β-galactosidase LBD ligand binding domain LBP ligand binding pocket Lc Lurcher LiCl lithium chloride MgCl2 magnesium chloride MMLV-RT Moloney murine leukemia virus-reverse transcriptase mRNA messenger ribonucleic acid NaCl sodium chloride NLS nuclear localization signal NOS1 nitric oxide synthase-1 NRSF neuron restrictive silencing factor N-term amino terminal NTG nontransgenic P postnatal PBS phosphate buffered saline PBST phosphate buffered saline-Tween-20 PC Purkinje cells Pcd Purkinje cell degeneration Pcp-2 Purkinje cell protein-2 PCR polymerase chain reaction PIPES 1,4-Piperazinediethanesulfonic acid PKA protein kinase A PKC protein kinase C PMSF phenylmethanesulphonylfluoride poly(A) poly adenosine xvii Prom promoter REST-1 RE-1 silencing transcription factor RGS regulator of G-protein signaling RNA ribonucleic acid RNase ribonuclease ROR retinoic acid receptor-related orphan nuclear receptor RORE ROR response element RORα retinoic acid receptor-related orphan nuclear receptor-α RORβ retinoic acid receptor-related orphan nuclear receptor-β RORγ retinoic acid receptor-related orphan nuclear receptor-γ RT reverse transcription/reverse transcriptase rTdT recombinant terminal deoxynucleotidyl transferase RT-PCR reverse transcription-polymerase chain reaction SD standard deviation SDS sodium dodecyl sulfate SEM standard error margin sg Staggerer SSC sodium chloride/sodium citrate SV40 Simian virus 40 T4-PK T4-polynucleotide kinase TAE triethanolamine TAE Tris/Acetate/EDTA TBE Tris/Borate/EDTA TCA trichloroacetic acid TE Tris-EDTA Tg transgenic TOR thymus orphan receptor Tris-HCl Tris-hydrochloride tRNA transfer RNA TRβ thyroid hormone receptor-β VDCC voltage-dependent Ca2+ channel WT wild type X-Gal 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside
xviii
CHAPTER 1
INTRODUCTION
1.1 SUMMARY
The work presented in this thesis focuses on the analysis of Purkinje cell protein-2 (Pcp-2/L7, herein called L7) gene expression in the mouse cerebellum, transcriptional activation of L7 gene by retinoic acid receptor-related orphan nuclear receptor-α, RORα, and the behavioral analysis of L7 knock-out mice. The introduction will start with a brief review of cerebellum function, structure, and circuitry. It will then continue with an overview of L7 protein, which will be followed by a review on control of L7 gene expression in the cerebellum. The last part of the introduction will focus on the nuclear receptors and RORα in detail.
1.2 CEREBELLUM
Cerebellum is a major part in the nervous system that constitutes only
10% of the brain but contains half of all the neurons in the brain (Rapoport et al.,
1 2000). Observations as early as 19th century revealed the fine structure of the
cerebellum, and also suggested an important role of the cerebellum in the control
of posture and voluntary movements (Middleton and Strick, 1998). Since then
cerebellum has been an ideal model for the study of the central nervous system
and neurodevelopment because of its simple yet highly organized anatomical
structure, detailed knowledge about its circuitry, well-characterized
developmental progression and the availability of viable murine mutants that affect its development and functioning (De Zeeuw et al., 1998; Goldowitz and
Hamre, 1998; Voogd and Glickstein, 1998; Zigmond, 1999).
1.2.1 Functions of the cerebellum
The major functions of the cerebellum include spatial accuracy and temporal coordination of movements, formation of balance and muscle tone, and motor learning (Doya, 2000; Ito, 2000; Ivry et al., 2002; Konarski et al., 2005;
Salmon and Butters, 1995; Thach, 1998). There is also growing evidence that cerebellum may play a role in cognition and sensory responsiveness (Barinaga,
1996; Rapoport et al., 2000). Cerebellum has been implicated in event timing
(Ivry et al., 2002), sensory data acquisition (Gao et al., 1996), and cognitive
processing (Kim et al., 1994). Since any abnormalities in the cerebellum structure and circuitry may potentially cause impairment in these functions, it is important to understand how cerebellum works.
2 1.2.2 Structure of the cerebellum
Cerebellum has a simple yet highly ordered anatomical structure
(Goldowitz and Hamre, 1998; Voogd and Glickstein, 1998). The laminar structure of the cerebellar cortex is composed of three distinct layers built from five types of neurons: Purkinje cells, granule cells, Golgi cells, basket cells and stellate cells
(Figure 1.1). The outermost molecular layer contains cell bodies of basket and stellate cells, dendrites of Purkinje cells and axons of granule cells. The axons of granule cells, whose cell bodies lie in a deeper layer, run in the medio-lateral axis and are called parallel fibers. The dendritic arborizations of Purkinje cells are oriented perpendicular to the parallel fibers in the molecular layer. Beneath this layer is the Purkinje cell layer, which is formed by a single layer of Purkinje cell bodies. Purkinje cells are the principal neurons in the cerebellum and provide the sole output of the cerebellar cortex. The innermost layer of the cerebellar cortex is called granule cell layer, which contains a large number of granule cells and a few Golgi cells. The dendrites of Golgi cells run into the molecular layer, ending near the parallel fibers. Their axons on the other hand run deeper in the granule cell layers and form cerebellar glomeruli with granule cell dendrites around mossy fiber terminals.
Cells of the cerebellar cortex are inhibitory in nature, except granule cells, which are excitatory neurons mediated by neurotransmitter glutamate.
3
Figure 1.1 The structure of the cerebellar cortex. (Adapted from Zigmond, 1999)
4 1.2.3 Cerebellar circuitry
Cerebellum receives two main types of inputs: Mossy and climbing fibers
(Kandel et al., 2000; Voogd and Glickstein, 1998; Zigmond, 1999). While both inputs form excitatory synapses with cerebellar neurons, they terminate at different layers in the cerebellar cortex and produce different firing patterns in the
Purkinje cells. Mossy fibers originate from nuclei in the spinal cord and the brain stem. They form excitatory synapses with granule cell dendrites in the granule cell layer. The parallel fibers then excite large number of Purkinje cells in the molecular layer. Each Purkinje cell receives input from many mossy fibers through a number of parallel fibers. The second input comes through climbing fibers, which originate from the inferior olivary nucleus in the lower portion of the brain stem. The climbing fibers wrap around Purkinje cell bodies and proximal dendrites and make excitatory synaptic contacts. A single Purkinje cell receives input from only one climbing fiber, but an individual climbing fiber can make synaptic contacts with 1-10 Purkinje cells. The flow of information through these inputs and its effect on the output of the cerebellar cortex require a well controlled circuitry. The information coming from the outside excite Purkinje cells through mossy, parallel and climbing fibers. Upon excitation, Purkinje cells inhibit their target neurons in the vestibular and cerebellar nuclei, which are mediated by the inhibitory neurotransmitter, γ-aminobutyric acid (GABA). On the other hand, mossy fibers also excite basket and stellate cells indirectly through parallel fibers. These cells then inhibit Purkinje cells, resulting in the release of inhibition
5 in the target nuclei. This complex yet delicate flow of information makes cerebellum an ideal model for studying the network of inputs and outputs in the
central nervous system.
1.2.4 Development of the cerebellum
Cerebellum develops from the junction of midbrain and hindbrain in the
developing embryo. Neuronal populations of the cerebellum arise from two different germinal zones. Purkinje cells and the deep cerebellar nuclear neurons
arise from the ventricular zone in the roof of the fourth ventricle. The first cells to
leave the ventricular zone are the deep cerebellar nuclear neurons, and this
migration takes place at embryonic days 10-12 (E10-12) in mice. The birth and
exit of the Purkinje cells take place at E11-13 in mice. They leave the ventricular
zone and form a temporary plate-like structure. The second germinal zone that
gives rise to cerebellar cells is the rhombic lip, the lateral boundary of the
rhombencephalon (hindbrain). Granule cells first develop in the rhombic lip and
then migrate over the Purkinje cells to form the external granular layer (EGL),
found on the exterior of the cerebellum. Granule cells are produced in this layer
at about the time when deep nuclear cells and Purkinje cells stop dividing. As
EGL is populated by granule cells, Golgi neurons are born from the diminishing ventricular zone. Postnatally, granule cells migrate from the EGL in an inward and radial manner, through the molecular layer and past the developing Purkinje cells. As migrating granule cells form the granule cell layer (GCL), basket and stellate cells that colonize the molecular layer are generated. The EGL structure
6 is completely diminished by the end of the third postnatal week in mice. During
this period, climbing fibers and mossy fibers mature and form connections with
Purkinje cells and granule cells, respectively (Goldowitz and Hamre, 1998;
Kandel et al., 2000; Zigmond, 1999).
En1, En2, Wnt1, Pax2, Pax5 and Gbx are among the early genes that are
expressed by primordial cerebellar cells and greatly affect the formation of the
cerebellum. Knockouts of En1 and Wnt1 eliminate the cerebellum (McMahon and
Bradley, 1990; Thomas and Capecchi, 1990; Wurst et al., 1994). Later, the
expression of cell-specific genes starts, for example RORα in the Purkinje cells and Math1 in the granule cells. A natural mutant of RORα and RORα knockout result in a severely defected cerebellum with a disrupted laminar structure, reduced number of Purkinje cells and loss of granule cells (Dussault et al., 1998;
Hamilton et al., 1996; Herrup and Mullen, 1979; Sidman et al., 1962; Steinmayr et al., 1998). Math1 knockout mice lack all of the granule cells due to the absence of EGL formation (Ben-Arie et al., 1997). RORα will be discussed in more detail in the following sections of this chapter.
1.2.5 Cerebellar mutant mice: Natural mutations
Several spontaneous mutations that lead to cerebellar degeneration have been identified. The most well known mutations are reeler, weaver, staggerer, nervous, lurcher, and Purkinje cell degeneration (Pcd) (Lalonde and Strazielle,
2006). Among these mutants, lurcher is caused by a mutation in the δ-2 glutamate receptor gene (Grid2), which is predominantly expressed in cerebellar
7 Purkinje cells (Takayama et al., 1996; Zuo et al., 1997). This mutation results in a complete absence of Purkinje cells and massive degeneration in the granule cells in the cerebellum (Zuo et al., 1997). Due to defective suckling, lurcher homozygotes cannot survive beyond the first neonatal day (Resibois et al.,
1997), but heterozygotes are available for further testing. Lurcher heterozygotes display ataxic gait and locomotor deficits (Fortier et al., 1987). Purkinje cell degeneration (Pcd) is caused by a mutation in the Nna1 gene (also known as
Agtpbp1) encoding ATP/GTP binding protein-1 (Fernandez-Gonzalez et al.,
2002), which is expressed in the Purkinje cells in cerebellum. Massive loss of
Purkinje cells is observed in the cerebellum, especially after the third post-natal week (Landis and Mullen, 1978). Pcd mice also display poor motor control and locomotor activities (Le Marec and Lalonde, 1997). Staggerer is caused by a mutation in the RORα gene (Hamilton et al., 1996; Matysiak-Scholze and Nehls,
1997), and results in a massive loss of Purkinje cells and granule cells in the cerebellum (Herrup and Mullen, 1979). Staggerer mice display staggering gait and ataxia, and will be discussed further in the following sections.
1.3 L7 IS A CEREBELLAR PURKINJE CELL SPECIFIC GENE
L7 is a GoLoco domain protein exclusively expressed in the cerebellar
Purkinje cells and retinal bipolar neurons. The following section will focus on the discovery of the L7 gene, function of the L7 protein, and control of L7 gene expression in the cerebellum.
8 1.3.1 Discovery of L7 gene
To be able to understand the function of different cells and tissues, it is important to identify cell-type specific genes. In a search to identify Purkinje cell specific genes in the cerebellum, L7, by convention also called Pcp-2 (Purkinje cell protein-2), was discovered by two separate groups in 1988 (Nordquist et al.,
1988; Oberdick et al., 1988). Oberdick et. al. identified a Purkinje cell specific clone by differential hybridization using wild type and Lurcher (Lc) cDNA probes.
Similarly, Nordquist et. al identified the clone PCD5 by subtractive hybridization between wild type and Purkinje cell degeneration (pcd) mice, and mapped this gene to mouse chromosome 8. L7 mRNA was shown to be 550-600 bp long including the polyA tail. The amino acid sequence was deduced from the cDNA sequence and was predicted to have an open reading frame of 99 amino acids and to encode a 10.7 kDa protein. Northern blot analysis suggested exclusive expression in the cerebellum when compared to brain and non-neuronal tissues, including liver, heart and kidney. In situ hybridization confirmed Purkinje cell specific expression of L7 mRNA while immunohistochemistry and western blotting showed the presence of L7 protein in the cerebellar Purkinje cells.
1.3.2 Phenotype of L7KO mice
Earlier studies on L7 focused on the expression pattern of the gene.
However, the function of the L7 protein was not known. To be able to understand the function of L7, L7/Pcp-2-knock-out (KO) mice were generated by two separate groups (Mohn et al., 1997; Vassileva et al., 1997). Absence of L7
9 mRNA and protein was determined by Northern blot and a combination of immunohistochemistry and Western blotting methods, respectively. L7KO mice were compared to their wild type (WT) littermates in neuroanatomical assays and behavioral tests. To everyone’s surprise, L7KO mice did not show any difference when compared to their WT littermates. L7KO mice developed normal pre- and postnatally, and did not show any obvious behavioral changes, for example ataxia or loss of balance. WT and L7KO mice had similar total body weights and cerebellum weights. Laminar structure of the cerebellum, gross cellular anatomy, cellular organization and the number of cells were compared between WT and
L7KO mice, and no significant difference was observed. Mohn et. al. also subjected WT and L7KO mice to a constant velocity rotarod test to check the presence of subtle behavioral changes, and did not find any significant difference. These observations suggested that L7 plays little role in the cerebellar
Purkinje cells, and its function can be compensated by other unknown factors in its absence. Nevertheless, it is possible that simple motor behavioral tests are not enough to detect any subtle behavioral differences between WT and L7KO mice. In addition, at the time of these knock-out studies, the signaling function of the L7 protein was not known. Our detailed analysis of L7KO mice on accelerating rotarod has suggested that L7KO mice have improved performance on this motor learning assay. L7KO mice also present sexually dimorphic sensorimotor changes, ranging from audition to heat sensation. These analyses will be discussed in detail in Chapter 5.
10 1.3.3 Structure and function of L7 protein
Two L7 isoforms, 99 aa-long L7A and 120 aa-long L7B, are produced by
the usage of different transcriptional start positions (Zhang et al., 2002). L7
proteins are GoLoco domain-containing proteins (Siderovski et al., 1999). L7A
and L7B contain one and two GoLoco domains, respectively. GoLoco domains
are 19 amino acid-long motifs found in binding partners of inhibitory Gα subunits
(Gαi1-3, Gαo, and Gαz) (Siderovski et al., 1999; Willard et al., 2004). Through
these GoLoco domains, L7 proteins can interact with Gαi and Gαo subunits of
heterotrimeric G proteins (Kimple et al., 2002; Kinoshita-Kawada et al., 2004; Luo
and Denker, 1999; Natochin et al., 2001). It was later suggested that L7 modulates the activity of P/Q-type Ca2+ channels (Kinoshita-Kawada et al.,
2004).
1.3.3.1 Heterotrimeric G proteins
Heterotrimeric G proteins are large proteins that mediate signal
transduction through G-protein coupled receptors. They consist of three subunits,
α (Gα), β (Gβ), and γ (Gγ). Of these three subunits, Gα is the largest, and it binds
to a guanine nucleotide (GDP or GTP). It also has the ability to hydrolyze GTP,
and it remains in its active, GTP-bound form for a short period of time. In the
inactive state, GDP-Gα forms a trimeric protein with membrane bound Gβ and
Gγ. When the G-protein-coupled receptor is activated by an agonist, Gα is
converted to its active, GTP-bound state via the guanine nucleotide exchange
factor (GEF) activity of the receptor. In its active state, GTP bound Gα
11 dissociates from the heterotrimeric G protein complex, and becomes free to
activate or inhibit several enzymes and ion channels, including adenylyl cyclase,
phospholipase-Cβ, phosphokinase C, K+ and Ca2+ channels among others,
depending on the type of the Gα subunit. For example, while Gαs stimulates
+ adenylyl cyclase, Gαi1-3, Gαo, and Gαz inhibit adenylyl cyclase and regulate K and Ca2+ channels (Figure 1.2). Depending on the cell type, Gβγ dimer can also
activate other proteins and pathways in the cell. Once Gα hydrolyzes GTP, GDP- bound Gα re-associates with Gβγ dimer, and the heterotrimeric G protein returns to its inactive state. In addition to the GTP hydrolysis ability of the Gα subunits, a group of proteins called regulators of G-protein signaling (RGS) can hasten the return of Gα to its inactive, GDP-bound state (Marinissen and Gutkind, 2001;
Morris and Malbon, 1999; Schoneberg et al., 1999).
1.3.3.2 L7 protein interacts with Gαi and Gαo
L7 can interact with Gαi2 and Gαo subunits but not the stimulating Gαs subunit (Luo and Denker, 1999). Later studies showed that L7 inhibits GDP/GTP exchange of Gαi1 and Gαo, functioning as a guanine-nucleotide dissociation
inhibitor (GDI) (Kimple et al., 2002; Natochin et al., 2001). L7B was shown to act
as a GDI and interact with Gαi1, but not Gαo or any other Gα subunits, including
Gαs (Willard et al., 2006). Because of its interaction with Gα subunits and its
function as GDI, L7 was also named as G-protein signaling modulator-4 (GPSM-
4) (Willard et al., 2006).
12
Figure 1.2 An overview showing the diversity of G protein-coupled receptors and signal transduction. (Adapted from Marinissen and Gutkind, 2001)
13 1.3.3.3 Modulation of voltage-dependent Ca2+ channels by G proteins
Voltage-dependent Ca2+ channels (VDCCs) are present in all neurons in the brain and modulate their firing patterns, thus affecting neurophysiological properties of the neurons. Moreover, Ca2+ influx into the neuron can affect physiological processes including neurotransmitter release, hormone secretion, synaptic plasticity, and so on.
P/Q-type Ca2+ channels account for more than 95% of the voltage- dependent Ca2+ channel activity in the cerebellar Purkinje cells (Llinas et al.,
1989). Although they are localized in the presynaptic terminals and participate in
neurotransmitter release in other neurons, in the Purkinje cells they reside in the
dendrites. In the cerebellar Purkinje cells, they appear to contribute the
spontaneous firing properties of these cells (Womack and Khodakhah, 2002;
Womack and Khodakhah, 2003; Womack and Khodakhah, 2004). An important
feature of P/Q-type Ca2+ channels is that they can be inhibited by
neurotransmitters via G-protein-coupled receptors (Bean, 1989; Herlitze et al.,
1996; Kaneko et al., 1999). Both Gαi/o and Gβγ subunits contribute to this effect
2+ through direct binding to the pore forming α1A subunit of the Ca channel
(Cav2.1) (Furukawa et al., 1998a; Furukawa et al., 1998b; Herlitze et al., 1996;
Ikeda, 1996; Kaneko et al., 1999; Kinoshita et al., 2001; Tosetti et al., 2002).
14 1.3.3.4 L7 modulates G-protein-coupled receptor-mediated inhibition of Ca2+ channels in a dose-dependent manner
An investigation of L7 protein function in Xenopus oocytes suggested that
L7 can modulate the inhibition of P/Q-type Ca2+ channels via G-protein-coupled receptors in a dose-dependent manner (Kinoshita-Kawada et al., 2004). It was shown in this study that low concentrations of L7 isoforms enhanced receptor- mediated inhibition of the Ca2+ channels, and this inhibitory effect was mediated through the Gβγ subunits. In contrast, high levels of L7 isoforms reduced receptor-mediated inhibition and eased the inhibitory effects of Gα subunit on
Ca2+ channels. Nevertheless, how the modulation of P/Q-type Ca2+ channels by
L7 affects Purkinje cell physiology, cerebellum functioning, and mouse cerebellum is not known. Our detailed analysis of L7KO mice, presented in
Chapter 5, would be helpful to answer this question.
1.3.4 Structure of L7 gene
Initial analysis of the L7 gene structure suggested the presence of four exons and three introns that span around 2 kb (Nordquist et al., 1988; Oberdick et al., 1988). Further analysis with 5’ rapid amplification of cDNA ends (5’-RACE) method revealed two alternative cerebellar forms of the L7 mRNA, which are produced by the usage of different transcriptional start positions (Zhang et al.,
2002). In this study, mouse, rat and human L7 gene structures were examined and found to be very similar with the most notable difference in the genomic configuration of the first exon. Two forms of L7 were named L7A and L7B.
15
Figure 1.3 Comparison of rodent and human L7 gene structure and gene expression.
As seen in Figure 1.3, in rodents, the sequence coding for the first exon of form A (exon 1A) is found upstream of that of form B (exon1B). Both forms share exons 2-4. Exon 1A has no start codon; therefore L7A mRNA initiates translation in exon 2, producing a 99 aa-long protein. Exon 1B, on the other hand, has a start codon at its 3’-end, and this is in frame following its splicing to exon 2. As a result of splicing of exon 1B to exon 2, twenty-one amino acids are added to the
N-terminus of the peptide, producing a 120 aa-long protein. These two proteins differ with respect to the number of GoLoco domains; while L7A contains one
GoLoco domain, L7B contains two (Zhang et al., 2002).
In humans, similar to rodents, two forms of L7 mRNA were identified by 5’
RACE that differ in their 5’ ends. Both forms are identical over a region showing high homology to the complete exons 2-4 in mice. Human L7B is very similar to its mouse counterpart as it includes exon 1B, which is highly homologous to mouse exon 1B, spliced to exon 2, producing a 120 aa-long protein with two
16 GoLoco domains. The human L7A, however, has only three exons. The first exon
(hExon1A) is a composite of mouse exon 2 homologous region and a continuous upstream region of 119 bp. This continuous region that forms the 5’ end of the first exon in human L7A is homologous to the downstream region of mouse intron
1B. For simplicity and to highlight the homology of mouse, the second and third exons of h1A are called exons 3 and 4. L7A in humans produces a 99 aa-long protein with one GoLoco domain. In neither rodents nor humans is there any evidence of an mRNA structure carrying both exons 1A and 1B.
In addition to these forms, a eye-specific transcript in mouse and human, and a testis-specific transcript in human have been reported (Mammalian Gene
Collection Program Team* et al., 2002). However, expression of these new isoforms in the mouse has not been examined. In addition, expression of L7A and L7B in the eye has not been analyzed. Chapter 4 of this thesis will focus on the analysis of different mRNA forms in mouse cerebellum and eye.
1.3.5 Analysis of L7 gene expression in the mouse cerebellum
L7 expression starts around E15-17 in the medial regions of mouse cerebellum (Oberdick et al., 1993; Smeyne et al., 1991). During the postnatal development of the cerebellum, L7 expression progresses towards the lateral regions. In the adult, L7 is uniformly expressed throughout the cerebellum
(Smeyne et al., 1991). To investigate Purkinje cell-specific expression of L7, an 8 kb genomic clone of L7 was isolated from mouse (Oberdick et al., 1990). This genomic DNA fragment included 2 kb L7 structural gene, 4 kb fragment upstream
17 of the transcription start site and 2 kb fragment downstream of the polyadenylation signal. This 8 kb genomic fragment was then used to drive expression of β-galactosidase in the mouse cerebellum and retina. The expression pattern of the reporter gene under the control of L7 promoter in the transgenic mice was not different than that of the endogenous gene. This observation suggested that 8 kb genomic DNA fragment included all the required elements for cell-specific expression of L7 in the cerebellum and retina.
To search for the minimal sequences required for L7 expression in the cerebellar Purkinje cells, promoter truncation constructs were made (Oberdick et al., 1993). Promoter fragments as short as 0.25 kb still drove Purkinje cell- specific expression. However, constructs with promoter fragments shorter than 1
kb resulted in weaker and non-uniform expression in the adult cerebellum. 4 kb
and 1 kb promoter constructs showed a uniform expression throughout the
cerebellum in the adult, which indicated that positively acting elements are
present in the distal regions of the promoter, driving a uniform expression in the
adult cerebellum.
Two other studies expanded initial findings about the required information
within the 4 kb promoter and the structural gene for faithful Purkinje cell-specific
expression. In the first study, 4 kb of the promoter and 2 kb structural gene are
used and the expression of this transgene recapitulated the developmental and adult expression patterns of endogenous L7 gene (Figure 1.4A) (Oberdick et al.,
1990; Smeyne et al., 1991). In another study, Orr and colleagues used a construct that included a 4 kb promoter linked to the proximal and distal portions 18 of the structural gene, with a deletion of a region spanning exons 2-4 (Figure
1.4B) (Vandaele et al., 1991). In that study, they showed strong expression of the transgene in the cerebellar Purkinje cells consistent with the first report, but they also showed strong ectopic expression in a variety of brain regions. Thus, taken together, these earlier studies hinted at a silencer role for at least part of the structural gene. In neither study, however, were the individual functions of the promoter and the structural gene investigated.
Figure 1.4 Comparison of two L7 gene constructs carrying 4 kb promoter. A) The construct carrying complete L7 structural gene, and B) the construct that included the proximal and distal portions of the structural gene, with a deletion of a region spanning exons 2-4 (tE2 stands for truncated exon 2). Both constructs drove high levels of expression in the Purkinje cells. However, the second construct also showed strong ectopic expression.
1.3.6 Regulation of L7 gene expression
To understand how L7 expression is restricted to one cell type in the
cerebellum, it is necessary to identify positive and negative regulatory
transcription factors, and cis-acting elements. Several attempts have been made
19 to this end, and a variety of potential cis-elements have been identified. These include cAMP-response element (CRE), AP-1 binding site, E-box motif, among others (Anderson et al., 1997; Anderson et al., 1998; Oberdick et al., 1993;
Sanlioglu et al., 1998; Strait et al., 1992; Vandaele et al., 1991; Zou et al., 1994).
Several transcription factors including homeodomain proteins and POU domain transcription factors have been suggested as regulators of L7 gene expression.
These studies mostly involved in vitro reporter gene assays, and lacked a
detailed in vivo analysis.
Another factor shown to activate L7 gene expression in vitro is retinoic
acid receptor-related orphan nuclear receptor-α, RORα. RORα binding site in the
proximal L7 promoter was identified, and was shown to interact with in vitro- translated RORα protein in electrophoretic mobility shift assay (EMSA) (Schrader et al., 1996). Moreover, in vitro assays showed activation of the reporter gene
under the control of L7 promoter fragment containing RORα binding element in
the presence of RORα. Even though they presented data for RORα-specific
activation of L7 gene expression, these studies lacked in vivo analyses and a more detailed set of experiments to confirm specific binding of RORα protein to
L7 promoter in vivo. In a more recent study, Gold et. al. analyzed several genes in the cerebellum and identified L7 as one of the targets of RORα (Gold et al.,
2003). However, unlike the first two studies, Gold et. al. focused on an RORα
binding element found in a region ~2 kb upstream of the transcription initiation
site. To better understand the nature of these two sites, and to further investigate
activation of L7 gene expression by RORα, we carried out a more in-depth 20 analysis. Chapter 3 of this thesis will focus on the analysis of activation of L7 gene expression by RORα in cerebellar Purkinje cells and in cell culture.
1.4 NUCLEAR RECEPTOR SUPERFAMILY
Nuclear receptors represent a large superfamily of structurally related
transcription factors which regulate several different pathways ranging from
development and cell differentiation to physiology in animals (Mangelsdorf et al.,
1995). Because of their important roles in animal physiology, and association of
the ligands with known human diseases, nuclear receptors have been useful
therapeutic agents.
The superfamily consists of receptors for steroid hormones (e.g.
glucocorticoid and estrogen), steroid derivatives (e.g. Vitamin D3), and non-
steroids (e.g. thyroid hormone and retinoic acid). In addition, there is another
group of nuclear receptors, called orphan nuclear receptors, for which no ligands have been identified (Escriva et al., 2000; Laudet and Gronmeyer, 2002).
Classification of nuclear receptors can be made by different ways. First, they can be classified into two groups by their mode of action. Type I nuclear receptors are found in the cytoplasm, interacting with the heat shock proteins. Upon ligand
binding, they dissociate from the heat shock proteins, form homodimers and translocate into the nucleus, where they bind specific response elements in DNA, and activate transcription. Type II nuclear receptors, on the other hand, are found
in the nucleus. Upon activation by ligand binding, they form heterodimers with
retinoid X receptor, and activate transcription (Novac and Heinzel, 2004). In an
21 earlier classification of nuclear receptors, they were grouped based on the type of ligand they bind. However, the identification of ligands for some of the nuclear receptors has made this classification rather superficial. They are now classified by Nuclear Receptors Nomenclature Committee into 6-7 phylogenetic subfamilies with groups containing both orphan and liganded receptors (NRNC,
1999).
Although they have different roles in the cells, all nuclear receptors share a common structure (Figure 1.5). A typical nuclear receptor contains an isoform-
Figure 1.5 Structural and functional illustration of nuclear receptors. AF, transcription activation function; NLS, nuclear localization signal. (Adapted from Escriva et. al., 2000.)
specific N-terminal domain (regions A/B), a conserved DNA binding domain
(region C), a variable hinge domain (region D), a conserved ligand binding domain (region E), and a variable C-terminal domain (region F). Two transcription activation functions (AFs) have been described within a nuclear receptor structure. AF-1 is located in the N-terminal domain, and acts as a constitutively active, ligand-independent transcriptional activator. AF-2 is located in the ligand binding domain, and is ligand inducible. Region C is important in
22 dimerization properties, and DNA binding. Nuclear localization signal (NLS) is found within the regions C and D.
Nuclear receptors can bind DNA as monomers or as homo- or heterodimers. Upon activation by ligand binding, or by unidentified means in the case of orphan nuclear receptors, nuclear receptors translocate from the cytoplasm into the nucleus. In the nucleus, they regulate transcription of many genes by directly interacting with the response elements located in the promoters of target genes, or by interacting with other transcription factors. They recognize specific response elements, which can be a single half site for monomeric receptors, and repeats of half sites for dimeric receptors. Nuclear receptors regulate transcription by several mechanisms (Chen, 2000; Glass and Rosenfeld,
2000; Laudet and Gronmeyer, 2002). They can either activate or repress transcription by direct binding to specific response elements in promoters or by binding to other transcription factors that are already bound to DNA. In the case of transcriptional activation by nuclear receptors, ligand binding is the key step.
The nuclear receptors are activated by ligand binding, and then specifically interact with response elements in the gene, and activate gene expression. They also contribute to gene expression by interacting with other transcription factor, thus acting as co-activators. A group of nuclear receptors, on the other hand, are known as orphan nuclear receptors for which no ligands have been identified.
These receptors are thought to be constitutively active. Transcriptional repression can take place in the presence or absence of the ligand binding.
Some nuclear receptors, e.g. thyroid receptor or retinoic acid receptor, can form 23 heterodimers with retinoid X receptor and actively repress transcription by binding to hormone response element in the absence of the ligand. In addition,
other nuclear receptors, e.g. glucocorticoid receptor, can indirectly repress gene
expression by binding to other transcription factors in a ligand-dependent
mechanism, and inhibiting their activity.
1.4.1 Orphan nuclear receptors
A large group of nuclear receptors are called “orphan” because no ligand
has been identified for these receptors. Like other nuclear receptors, orphan
nuclear receptors also have important functions. They are important and
essential in development and cell differentiation, regulation of metabolism,
homeostasis and circadian rhythm (Benoit et al., 2006; Laudet and Gronmeyer,
2002). For example, elimination of mouse ovalbumin upstream promoter-
transcription factor (COUP-TF) gene results in an embryonic lethal phenotype
with defects in the development of the nervous system (Cooney et al., 2001; Qiu
et al., 1997). Moreover, while orphan nuclear receptor NR2E1/Tlx plays an
important role during forebrain development (Roy et al., 2004) and retinal
development (Kobayashi et al., 1999; Miyawaki et al., 2004), retinoic acid
receptor related orphan nuclear receptor-α (RORα) is required for cerebellum
development (Hamilton et al., 1996; Sidman et al., 1962). In the adult, estrogen-
related receptor-α (ERRα) plays a role in adipogenesis and energy metabolism
(Luo et al., 2003; Sladek and Giguere, 2000) whereas liver receptor homolog-1
24 (LRH-1; NR5A2) and steroidogenic factor-1 (SF-1) are important in cholesterol metabolism (Fayard et al., 2004).
Similar to liganded nuclear receptors, orphan nuclear receptors also recognize specific response elements in the promoter of target genes. However, unlike liganded receptors, most orphan nuclear receptors recognize a half site
sequence in the promoter and bind as a monomer (Benoit et al., 2006; Giguere
et al., 1995b). In contrast to liganded nuclear receptors, orphan nuclear receptors
are considered as constitutively active receptors. Nevertheless, studies to identify
ligands for these receptors are still in progress.
1.4.2 Retinoic acid receptor-related orphan nuclear receptors (RORs)
One group of orphan nuclear receptors are called retinoic acid related orphan nuclear receptors (RORs) (Jetten et al., 2001; Laudet and Gronmeyer,
2002). This group consists of three orphan receptors, which are encoded by three separate genes: RORα, RORβ, and RORγ.
RORβ is uniquely expressed in the brain, pineal gland and retina (Andre et al., 1998b; Greiner et al., 1996). In the brain, it is detected in the cerebral cortex, and in primary sensory cortices, but no expression is detected in the cerebellum
(Becker-Andre et al., 1994; Schaeren-Wiemers et al., 1997). RORβ recognizes a
TAGGTCA half site and binds as a monomer (Carlberg et al., 1994; Greiner et al., 1996). However, transcriptional activation takes place only when two properly spaced half sites are present. Two isoforms of RORβ have been identified,
RORβ1 and RORβ2 (Andre et al., 1998b). These isoforms differ only at their N-
25 terminal regions. While RORβ1 is mainly expressed in the brain, expression of
RORβ2 is restricted to the pineal gland and the retina. RORβ-/- mice undergo normal development and do not initially present defects in the regions where
RORβ is normally expressed (Andre et al., 1998a). However, these knock-out mice display decreased muscle strength and duck-like gait later in the adulthood.
Although developing retina seems to be unaffected, degenerative cell loss is observed in retina of the adults, which results in blindness. In addition to these, regulation of the circadian rhythm was observed to be disrupted in RORβ-/- mice.
One explanation for these phenotypes would be that RORβ might regulate transcription of the genes important in regulation of circadian rhythm and retinal cell survival. However, future studies are needed to determine the exact mechanisms.
RORγ is expressed at high levels in the thymus, skeletal muscle, liver, mammary gland and kidney (He et al., 1998; Hirose et al., 1994; Medvedev et al.,
1996; Ortiz et al., 1995; Villey et al., 1999). Two isoforms of RORγ have been identified, RORγ1 and RORγ2. While RORγ1 is detected in many tissues,
RORγ2 expression is restricted to the thymus, and for this reason RORγ2 is often called thymus orphan receptor (TOR) (Ortiz et al., 1995; Villey et al., 1999).
RORγ recognizes a single core motif GGTCA preceded by a 6 bp-long AT-rich sequence (Medvedev et al., 1996; Ortiz et al., 1995). RORγ-/- mice display a normal appearance, but they lack all of the lymph nodes (Kurebayashi et al.,
2000; Sun et al., 2000), which suggests that RORγ affects lymph node development in mice. 26 The third isoform, RORα, which we show in Chapter 3 is an activator of L7
gene in the mouse cerebellum, will be examined in more detail in the following
section.
1.4.3 RORα
1.4.3.1 Discovery of RORα
Like other ROR isoforms, RORα was identified as a result of different
strategies to clone novel members of the nuclear receptor superfamily. The first
homolog of retinoic acid receptor-related orphan nuclear receptors (RORs) was
discovered in Drosophila in a screen to identify members of the steroid receptor
family (Koelle et al., 1992). The first gene identified was Drosophila hormone
receptor 3 (DHR3). Later a human homolog of DHR3 was identified and called
RORα (Becker-Andre et al., 1993; Carlberg et al., 1994; Giguere et al., 1994).
Four isoforms of RORα have been identified in humans, RORα1-4,
whereas only two isoforms RORα1 and α4 are found in mice (Becker-Andre et
al., 1993). RORα isoforms are generated by a combination of alternative
promoter usage and exon splicing. They are detected in many tissues, including
brain, heart, lungs, liver, muscle and spleen. They share the same DNA binding
domain, a hinge region and putative ligand binding domain, while each contains a different amino terminal (N-term) domain (A/B regions), which influences the receptor’s DNA binding activity.
27 1.4.3.2 Genomic structure of RORα isoforms
RORα gene was mapped to mouse chromosome 9 and human chromosome 15q21-22 (Giguere et al., 1995a). The gene spans a region of 725 kb and contains 12 exons. Isoforms, α1-4, contain long open reading frames of
1569, 1668, 1644 and 1687 nucleotides, respectively. The open reading frames are predicted to encode proteins of 523, 556, 548, and 466-468 amino acid residues (Giguere et al., 1994; Matsui et al., 1995; Matysiak-Scholze and Nehls,
1997). Different isoforms form as a result of different promoter usage and alternative splicing.
1.4.3.3 Expression patterns of RORα isoforms
RORα mRNAs have been detected in many tissues in humans, including heart, brain, skin, lungs, muscle, spleen, testis, ovary, retina, liver, thymus and peripheral blood leukocytes (Becker-Andre et al., 1993; Chauvet et al., 2002;
Giguere et al., 1994; Steinmayr et al., 1998). Although presumptive RORα2 and
α3 homologs could not be detected in mice, RORα1 and α4 are expressed in brain, cerebellum, testis and skin (Matysiak-Scholze and Nehls, 1997; Steinmayr et al., 1998). RORα1 is also expressed in developing mouse forebrain from
E12.5 to P7 (Nakagawa and O'Leary, 2003). Moreover, RORα4 is detected in adult brain including thalamus, olfactory bulbs and cerebellum, where it is present at highest levels in the Purkinje cells (Matsui et al., 1995). In addition to these, several defects observed in mice lacking RORα proteins suggests a role of RORα in muscle activity and formation, regulation of immune response and
28 bone metabolism. Therefore it would be expected to detect RORα expression in muscles cells and cells of the immune system in mice.
1.4.3.4 Protein structure of RORα isoforms
RORα proteins share a very similar structure with other members of the nuclear receptor family of transcription factors. Protein structure basically contains an amino terminal domain, DNA binding domain, a hinge domain, and a putative ligand binding domain (Becker-Andre et al., 1993; Giguere et al., 1994;
Matsui et al., 1995) (Figure 1.6).
Figure 1.6 Schematic representations of RORα gene products. (Adapted from Giguere, 1994; Matsui, 1995; Matysiak-Schulze and Nehls, 1997)
DNA binding domains, which include two zinc fingers and a carboxyl- terminal (C-term) extension, hinge region, and putative ligand binding domains are highly conserved among ROR isoforms. However, they differ in their amino- terminal (N-term) domains, and these specific domains dictate the distinct DNA- binding properties of each isoform. 29 1.4.3.5 Characterization of ROR response elements
RORs bind with high affinity to DNA elements, called ROR response
elements (RORE), which consist of the half-site core motif PuGGTCA preceded
by an AT-rich sequence (Giguere et al., 1995b; Giguere et al., 1994). They bind
along one face of DNA helix as a monomer (Giguere et al., 1995b). Two zinc
fingers in the DNA binding domain recognize the PuGGTCA core motif in the
major groove of the DNA molecule. The C-term extension (CTE) interacts with
the adjacent minor groove and makes contact with the 5’ A/T-rich sequence. The
N-term domain of each isoform makes contacts with CTE in the DNA-binding
domain and this introduces slight structural changes in the protein. These
changes regulate the interaction between CTE and the A/T rich region preceding
PuGGTCA motif. The variations in the A/T-rich sequence can greatly affect the
binding of RORα, and the interaction of N-term domain with CTE in the DNA binding domain influences the RORE binding specificity of RORα isoforms
(Giguere et al., 1995b; Giguere et al., 1994; McBroom et al., 1995). The contribution of N-term domain and CTE to recognition of ROREs indicates that zinc-finger motifs alone are not sufficient to determine the DNA-binding
properties of RORα isoforms.
1.4.3.6 Staggerer phenotype and RORα-/- mice
A natural deletion in the RORα gene results in a dramatic phenotype in mice, called staggerer. Staggerer (sg) was discovered in 1962 as a result of a spontaneous mutation in obese mice stock at Jackson Laboratory, Bar Harbor,
30 ME (Sidman et al., 1962). The mutant mice were recognized by their staggering
gait, mild tremor, hypotonia and small size. Although mutant pups were not
different than their littermates during the first week of age, symptoms became
obvious around postnatal day 8. In staggerer mice, cerebellum was severely affected and reduced to one third of a normal cerebellum. Histological analysis
revealed abnormalities in laminar structure of a cerebellum, and Purkinje cells
were found to be scattered among the granule cells, which were greatly reduced
in number.
Later studies indicated that severe cerebellar ataxia was the result of a
cell-autonomous defect in the development of Purkinje cells (Herrup and Mullen,
1979). Purkinje cells in the staggerer mouse are small and abnormal in morphology and reduced by 60-90% in numbers. Furthermore, synaptic contact between Purkinje cells and granule cells is defective, which leads to massive granule cell loss (Sotelo, 1975; Sotelo and Changeux, 1974). The effects of the
mutation show a regional variation along the mediolateral axis in the cerebellum.
A sg cerebellum appears much smaller than a normal cerebellum, intermediate
regions seem to be the most affected, as they narrow to a thin bar of tissue
(Figure 1.7).
31
Figure 1.7 Comparison of the cerebella of a staggerer and a wild type mouse. (A) staggerer (B) wild type Arrows in panel A mark the divisions between medial (M), intermediate (I) and lateral (L) regions. (Adapted from Herrup and Mullen, 1979)
In addition to the cerebellar defects and abnormalities, sg mice also displayed other defects including severe atherosclerosis (hardening and loss of elasticity of medium or large arteries) when fed a high-fat diet, hypoalphalipoproteinemia (condition of abnormally low levels of alpha- lipoproteins in blood) (Mamontova et al., 1998), smooth muscle dysfunction in small resistance arteries (Besnard et al., 2002), immunodeficiencies linked to the overexpression of inflammatory cytokines (Kopmels et al., 1992), abnormalities in the bone metabolism (Meyer et al., 2000), muscular atrophy (Jarvis et al., 2002), abnormal circadian rhythm (Akachi and Takumi 2005 and Sato 2004), and allergen-induced lung inflammation (Jaradat 2006).
32 Hamilton et. al. in 1996, and Matysiak-Scholze and Nehls in 1997
independently mapped the “staggerer gene” to a 160 and 250-300 kb interval of
mouse chromosome 9, respectively, which contained the gene encoding retinoic
acid receptor-related orphan nuclear receptor-α, RORα (Hamilton et al., 1996;
Matysiak-Scholze and Nehls, 1997). Staggerer mice were found to carry a 6.5 kb
deletion which results in the removal of a 122 bp-long exon of RORα gene. This exon-skipping event removes the start of the ligand binding homology domain
and shifts the reading frame, which causes an introduction of a stop codon after
27 amino acids, therefore the full length protein cannot be translated. Two
independent studies by Steinmayr et. al and Dussault et. al. in 1998 provided
proof for staggerer phenotype as the result of RORα deletion (Dussault et al.,
1998; Steinmayr et al., 1998). Dussault et. al disrupted the function of the RORα
protein by inserting a neomycin cassette into the exon 3, which encodes the first
zinc finger of the DNA binding domain. RORα-/- homozygous mice displayed cerebellar defects similar to sg mice. Steinmayr et. al. generated mice lacking a functional RORα gene by replacing the second zinc finger of the DNA binding domain with β-galactosidase gene. RORα null mutant mice displayed similar motor deficits, anatomical and electrophysiological characteristics to sg mice.
1.4.3.7 Genes activated by RORα
RORα activates genes in several different tissues during developmental or
physiological processes.
33 One of the genes regulated by RORα is a helix-loop-helix transcription
factor, Bmal1, which plays an important role in the regulation of circadian rhythm
(Sato et al., 2004). The interaction of RORα with Bmal1 promoter was later demonstrated in vitro (Akashi and Takumi, 2005). In the murine lens, RORα
activates γF-crystallin gene, which is expressed during lens cell differentiation in
vertebrates (Tini et al., 1995). RORα also activates fibrinogen-β gene expression
in human hepatoma cells and in mouse liver (Chauvet et al., 2005). This
suggests an indirect role of RORα in the blood coagulation cascade. Several
studies examined regulation of genes involved in lipid homeostasis by RORα.
Muscle carnitine palmitoyltransferase-1 and caveolin-3 (Lau et al., 2004), and
Apolipoprotein A-I (Vu-Dac et al., 1997) are directly regulated by RORα. These
studies indicate a role of RORα in the regulation of lipid and lipoprotein
metabolism, and possibly explain the development of metabolic diseases, such
as atherosclerosis, in sg mice. Moreover, 5-lipoxygenase, cellular retinol binding
protein (CRBP), p21, bone sialo protein (Schrader et al., 1996), laminin-β-1
(Matsui, 1996), oxytocin (Chu and Zingg, 1999), N-myc (Dussault and Giguere,
1997), α-Fetoprotein (Bois-Joyeux et al., 2000), and Rev-erb-α (Delerive et al.,
2002; Raspe et al., 2002) are also targets of RORα. Nevertheless, the
identification of RORα target genes has been largely based on the presence of
ROREs in their promoters. In all cases, in vitro analyses showed that RORα
could bind promoter elements carrying RORE. In addition, in vitro reporter gene
assays suggested that these genes may be targets of RORα.
34 Regulation of gene expression by RORα has also been examined in the
cerebellum, which is greatly affected in sg mice. Two earlier studies suggested
activation of L7 expression by RORα using in vitro analyses (Matsui, 1997;
Schrader et al., 1996) (Regulation of L7 gene expression by RORα will be
discussed in detail in the following section). In an elaborate study to understand
genetic program controlled by RORα during cerebellum development, Gold et. al.
identified several target genes of RORα by a gene chip differential expression
approach followed by validation using in vivo chromatin immunoprecipitation
analysis (Gold et al., 2003). The first gene identified was Sonic hedgehog (Shh), which is required for granule cell proliferation in the developing cerebellum.
RORα was shown to regulate several genes required in Purkinje cells to process synaptic input from granule cells. One group of genes identified belongs to Ca2+-
mediated signal transduction mechanism; Pcp-4 (calmodulin inhibitor), Itpr1 (IP3 receptor and calmodulin target), Cals1 (Itpr1 binding partner), Calb1 (calcium binding protein), and L7. Another group of genes found, Slc1a6 and Spnb3, functions in glutamatergic signaling. According to the reciprocal signaling model, signaling between Purkinje cells and granule cell precursors causes activation of gene expression in the Purkinje cells, which then regulate the incoming glutamatergic signaling from the granule cells. This analysis provided insight into the mechanisms by which RORα participate in the cerebellum development.
35 1.4.3.8 Activation of L7 by RORα
Since RORα deletion greatly affects cerebellum development and
functioning, and RORα is expressed in Purkinje cells, L7 was a highly suspected
candidate as an RORα target gene due to its restricted expression in the
cerebellar Purkinje cells. Schrader et. al., Matsui and Gold et. al suggested
activation of L7 gene by RORα (Gold et al., 2003; Matsui, 1997; Schrader et al.,
1996).
Schrader et. al. systematically screened promoters of ~50 genes to
identify putative ROREs in their promoters (Schrader et al., 1996). One of the
genes that carried a putative RORE in the promoter was L7. Interaction of in
vitro-translated RORα protein and L7 promoter fragment carrying RORE was
shown by EMSA analysis. Co-transfection of L7-promoter-reporter construct and
RORα expression vector into Drosophila SL-3 cells indicated that presence of
RORα enhanced reporter gene activity by 3-4 folds. These results showed
activation of L7 gene expression by RORα in vitro.
Matsui also identified three putative ROREs in the proximal promoter
region of L7 gene (Matsui, 1997). The region -935 and +286 of L7 gene was
linked to a luciferase reporter gene. Co-transfection of RORα expression vector
and L7 promoter-luciferase reporter into P19 cells was carried out to determine if
L7 is a target of RORα. Luciferase activity was enhanced 5 folds when
cotransfected with RORα. To determine the site responsible for RORα-mediated
transcriptional activation, he constructed a series of promoter truncations and
deletions. Co-transfection was repeated with each promoter construct and RORα 36 expression vector. It was shown that truncation or deletion of -61 and -91 region
resulted in a loss of enhanced reporter gene activity. Binding of RORα to the L7
promoter element carrying this proximal region was shown in vitro with EMSA,
and this interaction was further supported by DNase I footprinting analysis of
labeled promoter fragment and in vitro-translated RORα. Matsui also applied a C-
term truncation mutant of RORα (RORα411), which had a decreased binding
affinity to the promoter fragment, and showed that it did not activate the
promoter-reporter construct. These in vitro studies revealed L7 as one of the target genes of RORα.
The analyses by Schrader et. al. and Matsui supported the idea that L7 is
one of the targets of RORα. However, these studies lacked a more detailed
analysis of activation of L7 by RORα in the cerebellar Purkinje cells. First of all,
they did not examine the L7 expression levels in sg cerebellum. Moreover, they
did not provide any in vivo data for the interaction of RORα with the RORE in the
L7 promoter.
Gold et. al. analyzed several genes in the cerebellum and identified L7 as
one of the targets of RORα (Gold et al., 2003). Microarray analysis and unpublished data suggested that L7 is not detected in sg cerebellum. Two
ROREs were identified in the L7 promoter, and chromatin immunoprecipitation
(ChIP) analysis indicated interaction of RORα with the distal RORE on L7 promoter, which was ~2 kb upstream of the transcription initiation site. Proximal
RORE, however, was not shown to interact with RORα. ChIP analysis was repeated using antibodies against known RORα coactivators. SRC-1, β-catenin, 37 and TIP-60 were identified as coactivators recruited on distal L7 promoter by
RORα. To further analyze these interactions microinjection experiments were carried out. A reporter construct containing 2 kb of the L7 promoter and a LacZ reporter gene was microinjected into CV-1 cells with or without RORα expression vector. Co-injection of RORα resulted in a dramatic increase in activation of the reporter gene. Co-infection of specific blocking antibodies to SRC-1, β-catenin, and TIP-60 in the presence of RORα dramatically decreased reporter gene activation. Blocking CBP and p/CIP did not affect reporter gene activation. Since
CBP antibodies could pul l down distal promoter region in ChIP assay, it was concluded that CBP binds to L7 promoter in an RORα-independent fashion.
Although this was a very elaborate examination of activation of L7 gene expression by RORα, it brought up a few questions. Earlier studies showed that proximal RORE in L7 promoter was important in regulation of gene expression by
RORα in vitro (Matsui, 1997). Gold et. al., on the other hand, claimed that this
site is not bound by RORα in vivo, although they do not describe the precise
position of the proximal site they examined (Gold et al., 2003). They identified a
distal promoter region, around 2 kb upstream of transcription initiation site, to be
occupied by RORα and its coactivators in vivo. Chapter 3 of this thesis will focus
on transcriptional control of L7 gene expression by RORα in more detail.
1.4.3.9 Transcriptional control by RORα
Whether RORα is regulated by a ligand-dependent or –independent
mechanism or it acts as a constitutively active receptor is still unknown. However,
38 several studies have been carried out to understand transcriptional control by
RORα.
1.4.3.9.1 Ligand-dependent mechanism: Possible ligands for RORα
RORα was called “orphan” because they did not have any known ligands.
However, several attempts have been made in order to identify ligands for
RORα. First, pineal gland hormone melatonin, and thiazolidine (CGP 52068),
which is a functional analogue of melatonin, were proposed as a ligand for RORα
(Wiesenberg et al., 1995). However, subsequent studies by few other
laboratories were not able to confirm the binding or activation of RORα by
melatonin (Jetten et al., 2001; Tini et al., 1995)
Recent studies indicated that cholesterol and/or cholesterol sulfate may be
ligands for RORα (Bitsch et al., 2003; Kallen et al., 2002). In a search to obtain
the first crystal structure of RORα, cholesterol was fortuitously discovered in a
pocket in the ligand binding domain (referred as ligand binding pocket (LBP)) by
X-ray crystallography and mass spectrometry. Point mutations introduced into the LBP caused a reduction in the transcriptional activity of RORα in a reporter gene assay. Moreover, treatment of lovastatin to deplete cholesterol levels in the
cells also caused a decreased transcriptional activity of RORα in a dose- dependent manner. RORα activity could then be reconstituted by addition of
cholesterol or a number of cholesterol analogs, demonstrating specificity of
RORα activity with changing cholesterol levels (Kallen et al., 2002). The observation that sg mice are susceptible to severe atherosclerosis when fed a
39 high-fat diet further supports these possible physiological roles of cholesterol and
cholesterol derivatives in the regulation of RORα (Boukhtouche et al., 2004;
Mamontova et al., 1998).
1.4.3.9.2 Ligand-independent mechanism
Since RORα was defined as an “orphan receptor”, and it was considered
as a constitutive activator of transcription, several attempts were made to be able
to understand and explain regulation mechanisms for control of transcriptional
activity of RORα in the absence of a ligand. The recruitment of the coactivators
by RORα, e.g. P300 (Lau et al., 1999), GR-interacting protein-1 (GRIP-1) and
VDR-interacting protein 205 (DRIP205) (Atkins et al., 1999), and CRE-binding
protein (CBP), p300, SRC-1, TIP-60, GRIP-1, and β-catenin (Gold et al., 2003)
were demonstrated in vitro and in vivo.
More recently, in an attempt to characterize the coactivator binding interface of RORα, Harris et. al. compared ligand binding domains of RORα and
thyroid hormone receptor-β (TRβ), which has an absolute requirement for a
ligand for activity (Harris et al., 2002). This molecular modeling analysis
highlighted a nonconserved amino acid and a short helical structure at the N-
term of the coactivator binding site, which enabled ligand-independent
recruitment of coactivators. Consistent with this, amino acid replacements at
proposed sites caused a reduction in the transcriptional activation and ablated
the recruitment of coactivators. These results offered an explanation for the
40 ligand-independent activity of transcriptional regulation by RORα, even though a search for a ligand has not yet been over.
1.4.3.9.3 Possible mechanism for regulation by phosphorylation
In search for identification of regulatory regions in isoform-specific N-term domains, Giguere et. al mapped a short stretch of amino acids as important in activation of RORα. This 18 aa region at the N-term domain contains three putative phosphorylation sites that may be important in regulation of RORα by phosphorylation. Ser-35 and Thr-53 are putative phosphorylation sites for protein kinase C (PKC), while Ser-49 is a putative phosphorylation site for protein kinase
A (PKA) (Giguere et al., 1995b). RORα isoforms are likely to be regulated by phosphorylation; however the importance of these sites or regulatory kinases has not yet been determined.
1.4.3.9.4 Role of CaMKIV in the regulation of RORα-mediated transcription
RORα is a Ca2+-responsive transcription factor and can be activated potently by Ca2+/calmodulin-dependent protein kinase IV (CaMKIV) (Kane and
Means, 2000). One hypothesis to explain this activation is that RORα is phosphorylated by CaMKIV at its LBD. However, point mutations at the putative phosphorylation sites do not cause a reduction of activation of RORα by
CaMKIV, which suggests an indirect mechanism for the activation of RORα by
CaMKIV.
41 1.5 OVERVIEW OF THESIS WORK
Work presented in the thesis focuses on two major areas: 1) Control of L7 gene expression, and 2) Behavioral analysis of L7KO mice. Chapter 2 describes
an analysis of regulatory properties of the L7 promoter and L7 structural genes on the control of L7 gene expression in the cerebellum. Work presented in
Chapter 3 describes the activation of L7 gene expression by RORα protein.
Chapter 4 focuses on the expression of different isoforms of L7 in mouse cerebellum and eye. Finally, Chapter 5 will describe a behavioral analysis of
L7KO mice.
42
CHAPTER 2
ROLES OF THE L7 PROXIMAL PROMOTER/ENHANCER AND THE
STRUCTURAL GENE IN CONTROL OF GENE EXPRESSION IN VIVO
2.1 ABSTRACT
In the central nervous system, L7 gene expression is restricted to
cerebellar Purkinje cells. In this study, we set out to determine the molecular and
genetic mechanisms of this restriction. The role of the structural gene was first
examined as a part of a gene engineering collaboration with Dr. Tony Young.
The 2 kb structural gene and LacZ reporter were linked to the promoter of the
human nitric oxide synthase-1 (NOS1) and γ-aminobutyric acid type-A (GABAA)
receptor α6 subunit (henceforth called α6) genes. In either case expression of these constructs in transgenic mice was excluded from Purkinje cells and was
found in brain regions and cell types expected of the promoter that was used.
These and other observations described in this chapter indicate that the L7
structural gene does not have any significant Purkinje cell promoting activity of its 43 own, and likely contains significant repressive activity. To further determine the
potential of the proximal promoter with respect to driving Purkinje cell-specific
expression, we took advantage of the above-mentioned constructs that lack
expression in Purkinje cells. Attachment of a 0.9 kb fragment of the L7 proximal promoter in an inverted orientation upstream of either construct resulted in highly reproducible and robust Purkinje cell-specific expression in transgenic mice, in addition to the characteristic expression of the respective (NOS1 and α6) promoters. Thus, the proximal promoter region appears to have the characteristics of an enhancer. However, further studies indicate that the enhancer may require cooperative action of sequences within the structural gene. Furthermore, these structural gene sequences would appear to act only when configured as part of the primary transcript, and not outside, and therefore do not appear to act as a traditional enhancer. This property could suggest that coupling of transcription and RNA processing plays a role in the specific expression of the L7 gene. Further analysis of the 0.9 kb L7 “pseudo-enhancer” and the structural gene cooperativity will be conducted in future studies by linking these sequences to a minimal promoter.
44 2.2 INTRODUCTION
Most gene expression analyses rely on in vitro model systems. For large genes (genes greater than 20 kb in length), it is often difficult to elucidate the minimal promoter and structural gene sequences that are required for proper spatial and temporal expression. However, genes are controlled by multiple negative and positive transcriptional regulators, and functioning of these regulators is mediated by the cell’s response to its environment and the input from the surrounding cells. Therefore, to better understand control of gene expression, in vivo analyses are needed.
Due to its small size the L7 gene provides a useful genetic engineering system to investigate control of gene expression in vivo. L7’s small size allows one to introduce mutations and deletions in the L7 structural gene and/or promoter sequences, and observe the effects in vivo.
Earlier in vivo studies to identify regulatory regions to direct Purkinje cell- specific expression focused on the promoter sequences. A promoter truncation analysis in vivo showed that promoter fragments as short as 0.25 kb still drove
Purkinje cell-specific expression (Oberdick et al., 1993). However, constructs with promoter fragments shorter than 1 kb resulted in weaker and non-uniform expression in the adult cerebellum. 4 kb and 1 kb promoter constructs showed a uniform expression throughout the cerebellum in the adult, which indicated that positively acting elements are present in the distal regions of the promoter, driving a uniform expression in the adult cerebellum.
45 In vivo expression analysis with 4 kb promoter plus the complete 2 kb structural gene (Figure 2.1A) showed that the transgene recapitulated the developmental and adult expression patterns restricted to the cerebellar Purkinje cells (Oberdick et al., 1990; Smeyne et al., 1991). This result indicated that all required information for faithful Purkinje cell-specific expression was contained within a DNA fragment spanning about 6 kb. Another study utilized a construct that included a 4 kb promoter fragment linked to the proximal and distal portions of the structural gene, with a deletion of a region spanning exons 2-4 (Figure
2.1B) (Vandaele et al., 1991). In that study, they showed strong expression of the transgene in the cerebellar Purkinje cells consistent with the first report, but they also showed strong ectopic expression in a variety of brain regions. Thus, taken together, these earlier studies hinted at a silencer role for at least part of the structural gene. In neither study, however, were the individual functions of the promoter and the structural gene investigated.
Figure 2.1 Comparison of two L7 gene constructs carrying 4 kb promoter. A) The construct carrying complete L7 structural gene, and B) the construct that included the proximal and distal portions of the structural gene, with a deletion of a region spanning exons 2-4 (tE2 stands for truncated exon 2). Both constructs drove high levels of expression in the Purkinje cells. However, the second construct also showed strong ectopic expression.
46 In our study, we set out to determine the molecular and genetic mechanisms of L7 gene expression restricted to cerebellar Purkinje cells using in vivo expression analysis. Based on the observations from previous studies, we decided to investigate the individual functions of L7 promoter and structural gene sequences in more detail. One way to do this is to generate expression constructs which carry L7 promoter and L7 structural gene in combination with other promoter and genes, and examine the expression of these hybrid constructs in mouse brain.
Towards this goal, we first examined the role of the structural gene as part of a gene engineering collaboration with Dr. Tony Young. The 2 kb structural gene and LacZ reporter were linked to the promoter of the human nitric oxide synthase-1 (NOS1) gene. We then expanded this analysis using the promoter of the mouse γ-aminobutyric acid type-A receptor α6 subunit (henceforth called α6) gene.
Dr. Young’s lab identified multiple and closely linked transcriptional start positions of the human NOS1 gene, and the hNOS1 promoter has been tested in cell culture (Xie et al., 1995). However an expression construct carrying 5’1-5’2- hNOS1 promoter fragment, the first exon of the NOS1 gene with and SV40 intron, LacZ coding sequence, and SV40-poly(A) did not work in vivo; no expression was observed in transgenic mice (Dr. Young, personal communication). Hypothesizing the mRNA stability and processing might be part of the problem, and the L7 structural gene may serve as a more favorable transcriptional unit for brain expression studies, two different promoter fragments, 47 5’1-5’2 and 5’3-5’4 were fused to the L7 structural gene-LacZ cassette, and the
expression pattern was examined in transgenic mice.
The mouse GABAA receptor α6 subunit gene (henceforth called α6) was previously shown to drive highly selective, granule cell-specific expression in the mouse cerebellum with little or no ectopic expression in the brain (Bahn et al.,
1997). This construct included 1 kb α6 promoter, first 8 exons of the α6 gene and a LacZ cassette under the control of an internal ribosome entry site (IRES). In our study, we linked the α6 promoter to L7 structural gene and examined expression in transgenic mice.
With either promoter, expression of these constructs was excluded from
Purkinje cells, and was found in brain regions expected of the promoter used.
The observations described in this chapter indicate that L7 structural gene does not carry any activity to promote Purkinje cell-specific expression. Attachment of a 0.9 kb fragment of the L7 proximal promoter in an inverted orientation upstream of either construct, on the other hand, resulted in highly reproducible and robust
Purkinje cell-specific expression in transgenic mice. Thus, L7 proximal promoter region appears to have the characteristics of an enhancer. However, this enhancer activity requires the cooperative action of the L7 structural gene; it works when the L7 structural gene is placed within the transcriptional unit of the construct. In this chapter we will present data for the analysis of the 0.9 kb
“pseudo-enhancer” and the L7 structural gene.
48 2.3 MATERIALS AND METHODS
2.3.1 Mouse strains
All transgenic mouse lines were produced in FVB/N strain using Standard
Pronuclear Microinjection method in Transgenic Animal Facility at the Ohio State
University.
2.3.2 DNA Constructs for transgenic mice
L7-LacZ structural gene cassette, α6 promoter constructs (a6-Prom, 3K,
6K, a6IL-L7 Enhancer, a6IL-L7SGC, and a6IL-L7SGC-L7 Enhancer Test), L7-
GFP constructs (L7full-GFP and L7Trun-GFP), and NOS1 promoter constructs
(NOS1-L7SGC-LacZ and NOS1-L7 Enhancer Test) were previously made by Dr.
Oberdick’s and Dr. Young’s lab members and described elsewhere
(Serinagaoglu et al., 2007). mα6IRES-LacZ was the courtesy of Dr. Bill Wisden
(Bahn et al., 1997).
2.3.2.1 6k-L7 Enhancer Test construct
0.9 kb L7 proximal promoter fragment was made by Polymerase chain
reaction (PCR) using EM1 vector, which carry 4 kb L7 promoter and 2 kb
structural gene, as the template. The following primer sequences were used:
Forward: AGAAGCTTAAGTCGACTCCTGAAAGGTATCTGGAGATAGG SalI
Reverse: GGCTCTTATACTCGAGTACCTATACCCCCTGTGTGTTATAG XhoI
49 The fragment was cloned into pCR/8/GW/TOPO vector using TOPO TA
cloning kit (Invitrogen, Carlsbad, CA), cut out with XhoI and SalI, and cloned into
6K vector in the XhoI site. Orientation of the fragment was determined by sequencing. The entire 6K-L7 Enhancer Test construct was excised from the vector backbone by XhoI and SphI digestion prior to pronuclear injection (total construct size = 13.1 kb.).
2.3.3 Preparation of DNA for pronuclear injection
Plasmids were extracted using QiaFilter Plasmid Maxi kit (12262, Qiagen,
Inc., Valencia, CA). 50-60 µg of DNA for each construct were digested with suitable enzymes, and sent to Transgenic Animal Facility for pronuclear injection.
2.3.4 Tail DNA extraction
A 1-2 mm piece of mouse tail was digested in 300 µl tail buffer (50 mM
Tris (pH=8.0), 100 mM EDTA and 0.5% SDS) in the presence of 1 mg/ml
Proteinase-K at 55°C overnight on a rotating shaker. After digestion, the mixture was phenol-chloroform extracted, and tail DNA was ethanol-precipitated. DNA was air-dried and dissolved in 250 µl TE buffer overnight at room temperature. 1
µl was used in PCR to identify transgenic mice carrying the construct.
2.3.5 Genotyping
To identify mice carrying expression constructs 6k-L7enh, a6IL-L7enh, a6IL-L7SGC and a6IL-L7SGC-L7enh, polymerase chain reaction (PCR) was run using primers specific to LacZ gene. In addition, a control PCR was carried out
50 using primers specific to L7 gene in order to check the efficiency of tail DNA extraction procedure. The primer sequences were as follows:
LacZ set 1:
Forward: ACGCGCGAATTGAATTATGGCCCACACCAG
Reverse: GGCGCTCAGCTGGAATTCCGCCGATACTGA
PCR produces a 261 bp-fragment.
LacZ set 2:
Forward: CCATTGTCAGACATGTATACCCCGTACGTC
Reverse: GCCACCAATCCCCATATGGAAACCGTCGAT
PCR produces a 225 bp-fragment.
LacZ set 3:
Forward: ATTGACCCTAACGCCTGGGTCGAACGCTGG
Reverse: AACATCAACGGTAATCGCGATTTGACCTCT
PCR produces a 210 bp-fragment.
L7:
Forward: CGGACCAGGAAGGCTTCTTCAACCTGC
Reverse: ATCCCAGAACCCCAGCACTCCTGCCAC
PCR produces a 194 bp-fragment.
To identify mice carrying expression constructs L7full-GFP and L7trun-
GFP, polymerase chain reaction (PCR) was run using primers specific to GFP reporter genes. In addition, a control PCR was carried out using primers specific to L7 gene in order to check the success of tail DNA extraction procedure. The primer sequences were as follows: 51 L7trun-GFP:
Forward(EGFP-F2): ACGCGCGAATTGAATTATGGCCCACACCAG
Reverse (EGFP-T7-R4): GGCGCTCAGCTGGAATTCCGCCGATACTGA
PCR produces a 391 bp-fragment.
L7full-GFP:
Forward (L7-GL-5′): TGCAGTGCTTTTCCAGATACCCAGA
Reverse (L7-GL-3′): GTGTTCTGTTGATAATGGTCGGCCA
PCR produces a 350 bp-fragment.
2.3.6 Perfusion
Transgenic mice carrying enhancer constructs were perfused with fixing
solution containing 4% Paraformaldehyde in 1X PBS and 1X Blue I (0.12 M
PIPES, 2 mM MgCl2 and 2 mM EGTA, pH=7.6). Brain and cerebellum were
dissected and further fixed in the fixing solution for one hour. Samples were
washed with PBS twice and subjected to X-Gal staining procedure immediately.
2.3.7 X-Gal staining
Dissected brain samples were incubated in freshly prepared and filter-
sterilized X-Gal (5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside) oxidation
solution containing X-Gal (1 mg/ml, in DMSO) and Blue II (4.8 mM potassium
ferricyanide, 4.8 mM potassium ferrocyanide, 2 mM MgCl2, 0.02% NP40, 0.2 mM sodium deoxycholate and 1X PBS). For one cerebellum, 5 ml of the staining solution was used. The samples were incubated at 37°C for 4-5 hours or at room temperature overnight. Samples were washed with 1X PBS twice and stored in 52 1X PBS with 50 mM EDTA at 4°C. Whole mount pictures were taken using a camera attached to the dissecting microscope.
For histochemical analysis, samples were sectioned using the cryostat.
Before sectioning, samples were kept in 22% Sucrose solution with 1 mM EDTA overnight. Using the cryostat, 40 µm-thick tissue slices were collected in 1X PBS on 24-well plates. They were further stained in X-Gal staining solution overnight to make sure cells in deeper part of the tissue could have access to the substrate. Tissue sections were post-fixed in 4% Paraformaldehyde, washed with
1X PBS twice and with water once, and mounted on slides. After air drying, sections on microscope slides were counter-stained with Nuclear Fast Red, and subjected to a series of ethanol wash to remove water (70, 80, 90, 95 and 100% ethanol). The sections were incubated in Histoclear (HS-200, National
Diagnostics, Atlanta, GA) for 10 min and coverslipped using Permount (SP15-
100, Fisher, Pittsburgh, PA). They were analyzed under the Zeiss microscope, and images were taken using the MetaVue Imaging Software.
2.3.8 Cerebellum measurements
40 µm-thick sections from 6K, 6K-L7 Enhancer test, and nontransgenic
(NTG) mice were mounted on slides, and stained with 1 % cresyl violet in 30% acetic acid solution. The slides were subjected to a series of ethanol wash (70,
80, 90, 95 and 100% ethanol). The sections were then incubated in Histoclear for
10 min and coverslipped using Permount. The sections were analyzed under the
Zeiss microscope. Cerebellar cross-sectional area and granule cell layer
53 thickness were measured on sagittal sections using the MetaVue Imaging
Software. Granule cell density was determined by counting the number of granule cells in ten areas of 2450 µm2 each, and calculating the average number of cells.
2.3.9 Total RNA extraction
Total RNA was isolated from freshly dissected mouse tissues using TRIzol reagent (15596-026, Invitrogen, Carlsbad, CA) following manufacturer’s instructions. Basically, half of the cerebellum was homogenized using a glass
Teflon homogenizer in 1 ml TRIzol reagent. Proteins and lipids were separated from the mixture by addition of chloroform. RNA was precipitated by addition of isopropanol. Precipitated RNA was air dried and dissolved in RNase-free water.
Total amount of RNA was quantitated after measuring optical density of the sample. Samples were stored at -80°C.
2.3.10 DNase treatment
DNA-Free kit (AM1906, Ambion, Austin, TX) was used to eliminate DNA contamination in RNA extracts. 10 µg total RNA extract was incubated with 2 units of DNase at 37°C for 30 min. 0.1 volume of DNase inactivation reagent was added, and the sample was incubated at room temperature, mixing occasionally.
Inactivation reagent was precipitated by centrifuging, and RNA in the supernatant was transferred to a fresh tube. Samples were stored at -80°C.
54 2.3.11 Reverse Transcription (RT)
RETROscript kit (AM1710, Ambion, Austin, TX) was used to produce cDNA from total RNA extracted from mouse brain and cerebellum tissues. ~1 µg of DNA-free RNA was incubated with Oligo(dT) primer and nuclease free water at 70°C for 3 min to heat-denature the RNA. After putting the tube on ice, remaining RT components were added: RT buffer, dNTP mix, RNase inhibitor and MMLV-RT. The mixture was incubated at 42°C for one hour, and at 92°C for
10 min to inactivate the RT. Samples were stored at -20°C.
2.3.12 Polymerase chain reaction (PCR)
GeneAmp PCR System 9600 (Waltham, MA) was used to carry out PCRs.
PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows:
L7trun-GFP:
Forward (EGFP-F2): ACGCGCGAATTGAATTATGGCCCACACCAG
Reverse (EGFP-T7-R4): GGCGCTCAGCTGGAATTCCGCCGATACTGA
RT-PCR produces a 391 bp-fragment.
L7full-GFP:
Forward (L7-GL-5′): TGCAGTGCTTTTCCAGATACCCAGA
Reverse (L7-GL-3′): GTGTTCTGTTGATAATGGTCGGCCA
55 RT-PCR produces a 350 bp-fragment.
L7:
Forward (L7-YS-1-F): AGGCTTCTTCAACCTGCAGA
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
RT-PCR produces a 234 bp fragment.
β-actin:
Forward (bActin-2a-F): GCATGTGCAAAGCCGGCTTC
Reverse (bActin-2-R): GGGGTGTTGAAGGTCTCAAA
RT-PCR produces a 346 bp fragment.
After PCR samples were run on a 1.5% agarose gel, and visualized by
Gel Doc 2000 gel documentation system (BioRad, Hercules, CA).
2.3.13 Real-Time PCR
The iCycler iQ Real-time PCR Detection System was used along with the iQ SYBR Green Supermix (1708882) following manufacturer’s instructions
(BioRad, Hercules, CA). The 96 well-format was used. Before the Real-time
PCR, all primers were observed to produce a PCR product that resolved as a single band with no primer–dimers on agarose gels.
The Real-time PCR cycle program was as follows:
• Automatically inserted steps to collect well factors: 30 sec at 95°C (2
cycles)
• 1st step: 15 min at 95°C
• 2nd step: 15 sec at 94°C
56 • 3rd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (60 cycles)
• 4th step: 10 sec starting at 95°C and decreasing temperature for 0.5°C at
each cycle (100 cycles: this is to draw the melt curve)
Same primers used in RT-PCR were used in Real-time PCR.
2.3.13.1 Determination of PCR efficiency for each primer set
To be able to compare the threshold cycle values from two different PCRs, it is important to use primer sets with similar efficiencies (95-100%). For this purpose, standard curve function of iCycler iQ Real-time PCR Detection System was used.
Each cDNA of interest (for example L7 and β-actin) was produced by RT-
PCR. The fragments were resolved in 1.5% gel, gel-extracted and cloned into pCR/8/GW/TOPO vector using TOPO TA cloning kit (Invitrogen, Carlsbad, CA).
Plasmids were extracted from bacterial culture, and DNA concentration was calculated. Serial dilutions of plasmid containing the cDNA of interest were prepared: 10, 1, 0.1, 0.01, and 0.001 ng/µl. Each cDNA dilution series were used in Real-time PCR with its specific primers. Real-time PCR was run using three wells for each dilution. Standard curve function was selected at the beginning of the run. At the end of the cycle, the software provided the standard curve, and automatically calculated the efficiency of the primer set used in that particular reaction. Primers with efficiencies between 95 and 100% were then used in the
Real-time PCR with experimental cDNA samples.
57 2.3.13.2 Quantification of expression levels by Real-Time PCR
For quantification, the comparative CT method was used (Table 3 of User
Bulletin #2 of the ABI Prism 7700 Sequence Detection System). Briefly, the
average threshold cycle (CT) from three reactions of each RT sample was
determined. The ΔCT was calculated by subtracting the average actin CT value of
each sample from the average L7 (or the gene being investigated) CT value of
the same sample. The ΔΔCT value was determined by subtracting the ΔCT value of the sample with higher CT value from the ΔCT value of the other sample. The level of gene of interest relative to the control gene (in our case, β-actin) was determined by the equation 2−ΔΔCT.
The standard deviation (SD) for each sample was derived from the
square-root of the summed squares of the standard deviations of the average L7
and β-actin CT values, SD-L7, and SD-actin. Data are presented as ± SEM.
58 2.4 RESULTS
2.4.1 L7 proximal promoter/enhancer and the L7SGC are required in
concert for Purkinje cell-specific expression
A construct, mα6IRES-LacZ (herein called a6-IL), carrying 1 kb of the α6 promoter, the first 8 exons of the α6 gene and a LacZ reporter gene under the control of an internal ribosome entry site (IRES) (Figure 2.2B) was previously shown to drive highly selective, granule cell-specific expression in the mouse cerebellum with little or no ectopic expression in the brain (Bahn et al., 1997). In our study, we divided the a6-IL construct minus IRES-LacZ into three subregions:
1 kb promoter, exons 1-3, and exons 4-8 (mα6IRES-LacZ was the courtesy of Dr.
Bill Wisden) (Figure 2.2B).
We then generated other constructs with α6 promoter and L7 structural
gene. Earlier unpublished studies in our lab suggested that L7 structural gene is
not capable of driving expression in Purkinje cells. We attached 1 kb α6 promoter to the promoter-less L7 structural gene cassette (L7-SGC, Figure 2.2A), and this new construct was called a6-Prom (Figure 2.2B). L7-SGC included 2 kb- L7 gene and 3 kb-LacZ reporter gene. All start codons were removed on the L7 structural gene by mutagenesis in order to obtain translation of just the LacZ protein, as previously described (Smeyne et al., 1991). This construct produced no granule cell expression, and weak expression was observed in the cortex (Table 2.1, a6-
Prom). When 3 kb of the α6 structural gene (exons 1-3) was added at the 3’-end of the α6-Prom construct (3K construct) (Figure 2.2B), two founders showed 59 Figure 2.2 Constructs used to study the transcriptional role of the structural gene and the promoter/enhancer in vivo. A) L7 structural gene cassette (L7SGC). B) Constructs used in the α6 enhancer test analysis. C) Constructs used in the NOS1 enhancer test analysis.
60 weak granule cell expression throughout the cerebellum, while a third showed
moderate expression in the caudal-most lobules (Table 2.1, 3K). These 3K
founders had no expression in the pons, and weak expression in the cortex and
midbrain. Lastly, another 3 kb of the α6 structural gene (exons 4-8) was attached
(6K construct) (Figure 2.2B), which further improved granule cell expression of the transgene (Table 2.1, 6K). We now obtained one founder with high level, and four with moderate to weak level, granule cell expression (Figure 2.3C-E). In these granule cell-expressing mice, there was no expression in any other brain regions, in particular pons, cortex or midbrain. This is in contrast to control
transgenic mice that we produced carrying the original mα6IRES-LacZ construct
(a6-IL). As previously reported, most of the founders expressing this construct
showed moderate to high level expression in cerebellar granule cells (Table 2.1,
a6-IL, and Figure 2.3A-B). In contrast to the earlier report, however, we found
that all of the mice with expression in the cerebellum also had moderate to high
level expression in the pons, and some lines expressed at weak to moderate
levels in olfactory bulb, cortex, and midbrain (Table 2.1, a6-IL, and Figure 2.3A-
B). The source of this increased ectopic expression is unknown, but could be due
to strain differences. In our studies, we used FVB/N strain whereas CBA/cba X
C57BL/6 hybrids were used in the previous report.
It would appear from these results that the entire 7 kb α6 gene fragment
carrying the promoter and the first eight exons and introns is required for
maximal granule cell expression. In addition, incorporation of the L7 structural
gene into the α6 gene represses expression of the latter, especially in non- 61 cerebellar regions. It may even repress expression in granule cells, since few
expressing lines were obtained (eight expressors out of nineteen founders
carrying 3K and 6K constructs). Those founders that do express in granule cells,
however, have an expression that is more restricted to those cells. To determine
that this reduced frequency of granule cell expressors was due to a “toxic” effect
of L7-SGC transcript expression in granule and other cells, we measured the
cerebellar cross-sectional area, granule cell layer thickness, and granule cell
density in transgenic animals with high level expression of SGC-LacZ in granule
cells. No differences in these multiple measures were observed (Table 2.2).
Only one out of thirteen carrying the 6K construct had any expression in
Purkinje cells (Table 2.1, 6K), which supports the conclusion that the L7-SGC
has no significant enhancer activity for these cells. To determine whether the
proximal L7 promoter has any enhancer activity, the 0.9 kb promoter fragment
(with a mutated TATAA box) was linked to the 6K construct (6K-L7 Enhancer
Test, Figure 2.2B). This placed the 0.9 kb promoter fragment about 1.2 kb distant
from the L7-SGC in an inverted orientation. In spite of this arrangement, eleven
founders out of seventeen had expression in Purkinje cells, and six of these had
very high level expression, comparable to what we see with the standard L7
expression vector (Table 2.1, 6K-L7 Enhancer Test, and Figure 2.3F-K). Some
founders retained granule cell expression, while some did not. Most expression
outside the cerebellum (pons, olfactory bulb, and cortex) was eliminated, but
expression was still observable in the colliculi and thalamus, similar to the 6K vector. We conclude that the proximal promoter of L7 has enhancer-like 62 properties, and is separable from the SGC, but we cannot exclude that SGC is
required for the enhancement effect.
It appears from the above-mentioned results that L7-SGC fragment has a
repressive effect on α6 promoter function. To test whether this repressive effect
could be observed in a different configuration, we linked the SGC to the a6-IL
construct (a6IL-L7SGC; Figure 2.2B). However, the pattern of expression was
identical to that observed with the base a6-IL construct (Table 2.1, a6IL-L7SGC
and a6-IL). Therefore, if the SGC does carry a repressor, it would not appear to
act when placed outside of the transcriptional unit. Similarly, 0.9 kb L7 promoter
fragment, either alone (a6IL-L7 Enhancer Test; Figure 2.2B) or in combination
with the L7-SGC (a6IL-L7SGC-L7 Enhancer Test; Figure 2.2B), had no effect on
expression of the a6-IL transgene (Table 2.1, a6IL-L7 Enhancer Test and a6IL-
L7SGC-L7 Enhancer Test).
In another set of experiments1, two different promoter fragments of
hNOS1, 5’1-5’2 and 5’3-5’4, were fused to the L7 structural gene-LacZ cassette
(NOS1-L7SGC-LacZ construct, Figure 2.2C). Nine transgenic lines were obtained, four carrying 5’1-5’2 and five carrying 5’3-5’4 promoters. While some
line-to-line variations were observed, each displayed expression in multiple brain
regions known to express Nos1 (Figure 2.4A-F). In the cerebellum, four out of
nine lines had high to moderate levels of transgene expression in the granule cells, while two showed weak granule cell expression, three showed expression in the granule cell layer but probably in Golgi cells or glia, and one showed no
1 These experiments were carried out by Dr. Greg Hartt in Dr. Tony Young’ s lab. 63 expression in the cerebellum (Table 2.3). Sporadic or restricted expression was
observed in the Purkinje cell layer of a few lines, but in two of these the
expression is likely in Bergmann glia and not Purkinje cells. Some expression in
inhibitory interneurons was also observed. Transgene expression in the granule
cells with this construct is consistent with the earlier reports that showed Nos1 immunostaining in the cerebellar granule cells (Baader et al., 1997; Bredt et al.,
1990).
One characteristic of the NOS1-L7SGC-LacZ construct was its lack of
expression in cerebellar Purkinje cells. To determine whether the 0.9 kb L7
proximal promoter fragment could act to enhance expression of this construct in
Purkinje cells, we attached this fragment, again in an inverted orientation, at the
5’-end of the NOS1 promoter of the NOS1-L7SGC-LacZ construct. Nine founders
were obtained, and moderate to high levels of Purkinje cell expression was
observed in most of the lines. In spite of the high levels of Purkinje cell
expression in the cerebellum, most founders retained hallmark features of the
NOS1-L7SGC-LacZ expression pattern outside the cerebellum (Figure 2.4G-L).
Thus the L7 proximal promoter/enhancer and the hNOS1 promoter seem to act
in a modular and additive fashion for the most part. Within the cerebellum,
however, only Purkinje cell expression was observed. Therefore, as with the α6
gene, the L7 promoter and SGC can cooperate to “re-program” expression to
Purkinje cells.
The α6 and NOS1 data support the view that the L7 proximal
promoter/enhancer and the L7SGC are required in concert for Purkinje cell- 64 specific expression. Furthermore, this Purkinje cell enhancement is observed when the SGC was configured to drive the reporter gene, rather than placed outside the transcriptional unit.
2.4.2 Distal half of the L7 gene is required for high level expression in the
cerebellum
Purkinje cell enhancement was observed when both L7 proximal
promoter/enhancer and the L7SGC are present, and when L7SGC was placed
within the transcriptional unit of the construct. To identify the regulatory regions in
the L7SGC, we tested a truncated version of the L7 gene that includes the proximal promoter, and the structural gene missing the second half of the exon 2, exon 3 and 4, and introns 2 and 3, called L7trun-GFP (Figure 2.5A). The missing part in the gene was replaced with green fluorescent protein (GFP) gene, and
SV40 3’UTR fragment. We then compared expression of this construct to another
L7-GFP construct carrying the 0.9 kb proximal promoter, L7 structural gene and
GFP, called L7full-GFP (Figure 2.5A). We obtained seven founders carrying
L7trun-GFP. All brains were analyzed by in situ hybridization for detection of
mRNA, and confocal imaging and western blotting for detection of GFP expression, and no expression was observed by these methods. However, five of these founders were also examined by reverse transcription-PCR (RT-PCR) and
Real-time RT-PCR. RT-PCR revealed weak transgene expression in the cerebellum that ranged from 150-fold (Tg5) to 3000-fold (Tg18) less than that of endogenous mRNA (Figure 2.5B-D). In addition, expression was also found in
65 the brain minus cerebellum, roughly equal to that in the cerebellum in two cases
(Tg5 and Tg18). Tg 17 and Tg 3388 showed very weak expression of the transgene in the cerebellum, whereas Tg 3368 did not express at all. In contrast, the endogenous L7 mRNA, and L7full-GFP transgene mRNA expressed from the previously reported full-length transgene showed expression in the cerebellum that was 60,000-fold and 5000-fold greater, respectively, than in the rest of the brain. Therefore, even though these data do not allow us to distinguish cell type- specific expression in the cerebellum, it seems clear that the 0.9 kb promoter plus the proximal half of the structural gene is insufficient to drive high level cerebellum expression. The distal part of the gene is necessary. Moreover, this distal part is also required to suppress ectopic expression in the brain, consistent with the previous results.
66
Table 2.1 Expression of α6, 0.9 kb L7 proximal promoter, and L7SGC fusion gene constructs in transgenic mice
67 Mouse ID# GC PC Pons Olf. bulb Cortex Coll./ B.S. thal. a6-Prom
3 + + +
5 + + +
16 + + + 23 + + 24 + ++
27 + + +
33 + + 2 + + + 3K
3 + + 4 ++(post) + + 6 + + +
13 + +
14 + + + 17 + + 6K 2 + + 3 + 9 + + 10 ++ + 12 +(post) + + 13 ++ + + 17
20 + + 22 +++ 23 +(post) + 31 30 33 ++ +
6K-L7 Enhancer test inj. 1 1 + +++ inj. 2 8 + 15 + inj. 3 1 +++ ++ + ++ 3 + +++ + 8 + + 9 +++ 13(4) +++ + + 15 +++ + + 17 + ++ 21 + 22(5) inj. 4 1 + (post) + + + 11 + + 12 +++ + 20 + (post) ++ 21 + (post) Table 2.1. Continued
68 Table 2.1. Continued
Mouse ID # GC PC Pons Olf. Cortex Coll./ B.S. bulb Thal. a6-IL inj. 1 42 + (ant) + 47 43 57 +++ + 58 ++ + 54 + + ++ ++ 45 ++ + 59 +++ ++ + + + 53 ++ ++ ++ + ++ 41 ++ +++ + 49 ++ + inj. 2 3 + + + 8 + 9 ++ + + 10 + 11 + + + 14 +++ ++ + ++ 15 +++ ++ + 19 + 20 22 + + ++ 26 ++ (ant) ++ + 27 + +++ + 28 +++(ant),++(post) ++ a6IL-L7 Enhancer Test inj. 1 11 12 + 13 + 5 + 18 + 19 +++ ++ inj. 2 5 ++ + + 8 +++ + 11(a) 12(b) ++ + ++ +++ 15 + 17(c) ++ +++++ ++ 21 ++ 22 + + 23 +++ +++ Table 2.1. Continued
69 Table 2.1. Continued
Mouse ID # GC PC Pons Olf. Cortex Coll./ B.S. bulb Thal. a6IL-L7SGC 4 +++ +++ +++ +++ +++ +++ 5 +++ +++ 12 + +++ 14 ++ ++ + 15 ++ (ant), + (post) +++ 18 + +++ 20 +++ + 21 +++ +++ 24 + ++
27 + 35 ++ 37 + 40 +++ +++ 41 +++(post), +(ant) ++ a6IL-L7SG C- L7 Enhancer Test 6 + ++ 16 38 + +++ 41 +++(ant), +(post) +++ 73
post = expression in poste rior cerebellum only; ant = exp ression in anterio r cerebel- lum only; GC = granule cell s; PC = Purkinje cells; coll/thal = colliculi & thalamus; BS = brainstem
70
Figure 2.3 Analysis of L7-SGC function and L7 Enhancer Test: α6 promoter. A-E) Test of α6 gene fragments attached to the L7SGC. A-D) Whole-mount LacZ stains of hemi-brains (cut through mid-line) viewed in sagittal lane. A) Line 53, and B) line 59 carrying the mα6IRES-LacZ construct. C) Line 22 and D) line 33 carrying the 6K construct. E) Sagittal section of brain in C, showing granule cell-specific LacZ expression in the cerebellum. F-K) L7 Enhancer Test assay. F, G, I, J) Whole-mount stains of hemi- brains. F) Founder 15, and I) founder 1 (from injection 3) carrying the 6K-L7 Enhancer Test construct. G-J) Same as F and I, respectively, but close-up view of the cerebellum. H) Sagittal section of cerebellum in F and G showing LacZ expression restricted to Purkinje cells. K) Sagittal section of cerebellum in I and J, showing LacZ expression in both Purkinje cells and granule cells. Scale bars: D, 1 mm; F, 0.75 mm; J, 500 µm; E, K, 20 µm. ML=Molecular layer; GC=Granule cell layer, PC=Purkinje cell layer.
71
Construct name- L7-lacZ Cbm area IGL thickness GC density founder # expression (mm2 +SD) (mm +SD) (cells/2450 domain µm2 +SD)a 6K-22 high in GC 9.39+0.70 0.231+0.048 76.0+5.9 6K-33 low in GC 9.11+0.94 0.224+0.033 75.7+3.9 6K-17 none 9.43+0.54 0.225+0.024 74.7+2.3 6K-L7 enh-1 high in GC & PC 10.03+0.49 0.256+0.050 77.4+3.1 6K-L7 enh-15 high in PC 9.09+0.26 0.272+0.050 76.3+6.3 NTG (wild-type) none 9.10+0.37 0.249+0.057 75.3+5.4
Table 2.2 Morphological measures of transgenic cerebella. NTG = non- transgenic; PC = Purkinje cell; GC = granule cell; aGC number in a 70X35 µm box
72
Construct-Line # Granule cell Purkinje cell layer Basket/Stellate layer Cells NOS1 Line 2 + (sporadic) - - 3 +++ - ++ 18 ++ + (~30% of PCs) - 19 + + (B.Glia?) - 73 +++ - ++ 81 +(Gol,glia?) + (B.Glia?) - 87 ++ - - 88 +(Gol,glia?) ++ (PC,vermis + only) 99 NE NE NE
L7-NOS1 F0# 4 - ++ - 6 - ++ - 7 - +++ - **11 - ++ - 12 NE NE NE 16 NE NE NE **18 - +++ - 19 - +++ - 22 - +(sporadic) -
Table 2.3 Enhancer test using NOS1-L7SGC-LacZ base constructs. NE=no expression; PC=Purkinje cells; B. Glia=Bergmann glia NOS1 construct = 5’1+5’2 Lines 3, 18, 73, 99 & 5’3+5’4 Lines 2, 19, 81, 87, 88 **Indicates two Enhancer Test mice in which expression was restricted to cerebellum
73
Figure 2.4 Analysis of L7-SGC fusio n and L7 Enhancer Test: h NOS1 promoter. A-F ) Test of the hNOS1 promoter linked to the L7SGC. A, B, D, E) Whole-mount LacZ stains o f hemi-brains (cut through the mid-line) v iewed in sagittal plane. A) Line 73 and D) line 3 carrying the hNOS1-5’1-5’2 promoter li nked to L7SGC. B and E) Same brains as A and D, respectively, but close-up view of the cerebellum. C) Sagittal section of the cerebellum shown in A and B showing very high level expression in granule cells and weaker expression in molecular layer interneurons. F) Sagittal section of the cerebellum shown in D and E showing expression in mainly in basket and stellate cells, in some granule cells and/or Golgi interneurons, and possibly weak expression in Bergmann glia or Purkinje cells. G-M) L7 Enhancer Test assay. G, H, J, K, L, M) Whole-mount LacZ stains of hemi-brains. G) Line 7, J) line 18, and L) line 19 carrying the NOS1-L7 Enhancer Test vector. H, K, M) Same as G, J, and L, respectively, but close-up view of the cerebellum. I) Sagittal section of the cerebellum in G and H showing LacZ expression in Purkinje cells. The section was counterstained for calbindin antigen with HRP revelation. Scale bars: D, 1 mm; C, F, 40 µm; E, M, 500 µm; I 25 µm. GC=Granule cell layer, ML=Molecular layer.
74
Figure 2.5 The distal half of the L7 structural gene is necessary for high level cerebellum expression. A) Schematic diagrams of the L7full-GFP and L7trun-GFP constructs. B) RT-PCR used to determine whether L7trun-GFP was expressed in the cerebellum and cerebellum minus brain. C and D) Actin-normalized relative expression levels of L7trun-GFP, L7full-GFP and endogenous L7 mRNAs determined by Real-Time PCR. Panel C represents the data in logarithmic form, and panel D in linear form.
75 76 2.5 DISCUSSION
In this chapter, we presented data showing that the L7 0.9 kb proximal promoter/enhancer in conjunction with a 2 kb structural gene fragment can behave like a classic position-independent enhancer that is specific for cerebellar
Purkinje cells. This enhancer activity can re-program or silence other strong neuronal promoters in favor of Purkinje cell-specific expression. Neither fragment alone has significant Purkinje cell enhancement activity; and the distal half of the structural gene is necessary for the enhancement. In addition, L7 structural gene carries repressive signals that limit the expression in the brain.
From the data presented, it is clear that 0.9 kb proximal promoter can act as a Purkinje cell-specific enhancer when the L7 structural gene fragment is within the transcriptional unit. As shown in α6 promoter studies, attachment of proximal promoter fragment has no enhancer effect when the L7 structural gene is placed outside the transcriptional unit. Nevertheless, placement of the proximal part of the L7 structural gene within the transcriptional unit is not sufficient to drive Purkinje cell-specific expression when attached to the L7 promoter (L7Trun-
GFP); the distal part of the gene is also required. One interpretation of this is that
L7 promoter/enhancer function may be coupled to mRNA processing signals.
Recent studies in yeast suggested that the initiation of transcription and mRNA
3’-end processing are coupled by interaction of transcriptional co-activators and mRNA 3’-end processing factors before or during the formation of transcription initiation complex (Calvo and Manley, 2001; Hirose and Manley, 2000; Proudfoot,
2000; Proudfoot, 2004). It may be possible that certain end processing factors 77 that recognize specific sequences at the 3’-end of the L7 gene may contribute
this coupling mechanism, thus enhancing L7 expression in Purkinje cells.
Figure 2.6 Comparison of Purkinje cell-specific expression patterns of different L7 constructs.
Several observations support this coupling hypothesis. First of all, L7 structural gene has no Purkinje cell enhancement activity (Figure 2.6A); and promoter and the structural gene are required for Purkinje cell-specific expression (Figure 2.6B). Next, in spite of the fact that the original truncated construct with a deletion in the structural gene (Figure 2.1 and 2.6E) was expressed ectopically outside the cerebellum, it was nevertheless highly
78 expressed in Purkinje cells (Vandaele et al., 1991). On the other hand, our
L7Trun-GFP construct, which carried SV40-3’UTR and 3’-end processing signal
instead of the L7 counterparts (Figure 2.6D), was not expressed at detectable levels in Purkinje cells. However, trace levels detected by Real-Time PCR showed equivalent levels in cerebellum and brain minus cerebellum. Based on these results, we concluded that the 3’ region including the 3’UTR and 3-end processing signal is required for strong Purkinje cell-specific expression. We also conclude that structural gene region between exons 2 and 4 causes repression of expression in the brain. We can rule out that L73’UTR plays a role in Purkinje cell-specific expression, since when this region alone is replaced with SV40-
3’UTR (Figure 2.6C), there is still strong expression in Purkinje cells (Zhang,
2007). The only difference is that the mRNA is not localized in the dendrites with this construct, which suggests that L73’UTR is required for mRNA localization within the Purkinje cells. Nevertheless, this indicates the function of 3’-end processing region as the primary co-enhancer. Thus, this region may influence transcription in a different way from classical enhancement, and it may be an active component in the coupling between L7 transcription and 3’-end mRNA processing.
Studies with α6 promoter suggest that L7 structural gene has a repressive action on the expression in non-cerebellar regions in the brain. However, hNOS1 promoter construct cannot tell us anything specific about the repressor function of the structural gene. At the same time, it suggests that the repressor function is of moderate strength, and can be overridden by strong promoters. It is possible 79 that the moderate repressive capability of the L7 structural gene can be used to
tailor gene expression patterns by dampening down or eliminating weak sites of
transgene expression, as we observed using the α6 promoter.
One possible mechanism that explains repression of L7 gene in non- cerebellar regions may be the interaction of repressor proteins with binding sequences in the structural gene and the proximal promoter. Neuron restrictive silencing factor (NRSF)/RE-1 silencing transcription factor (REST-1) may be the first candidate for this repression mechanism. NRSF is a Zn-finger “silencer” protein extensively expressed in non-neuronal tissues. It recognizes a 21 bp-long
DNA element and suppresses expression of neuronal genes in the non-neuronal tissues, thus restricting expression to neurons (Chong et al., 1995; Schoenherr and Anderson, 1995a; Schoenherr and Anderson, 1995b). Some of the genes that are known to be targets of NRSF are the voltage-gated sodium type II channel (NaV1.2), superior cervical ganglion 10 (SCG10), and synapsin (Bruce et al., 2004; Quinn, 1996; Schoenherr et al., 1996). While NRSF would be the first candidate for repression of L7 expression in non-neuronal cells, we have not identified a binding site for NRSF in the human or rodent genes. A recently
identified repressor complex is composed of activator protein 4 (AP4) and the
corepressor geminin (Gem), which form a functional complex to repress target
gene expression in nonneuronal cells (Kim et al., 2006). AP4 is a ubiquitously
expressed helix-loop-helix transcription factor that binds to the DNA consensus
sequence of CAGCTG (Hu et al., 1990; Kim et al., 2006). Geminin, on the other
hand, is a DNA replication inhibitor protein (Wohlschlegel et al., 2000). A 80 repressor complex of AP4 and geminin was shown to repress expression of
phytanoyl-CoA α-hydroxylase-associated protein-1 (PAHX-AP1) in the non-
neuronal cells (Kim et al., 2006). This complex may be another candidate to
suppress L7 gene expression in the non-neuronal cells. However, we could not
identify AP4 binding sites in the human and mouse L7 genes. Nevertheless,
other factors with similar repressor functions may explain the lack of L7 expression outside the cerebellum.
Other repressors include the fruitfly HLH protein, Hairy, and its mammalian HES counterparts (Akazawa et al., 1992; Parkhurst, 1998). These factors bind to the so-called N-box sequence (CACNAG), and in Drosophila act to recruit co-factors such as Groucho to repress transcription of target genes. In fact, three N-boxes are located in roughly the same positions and orientation in the rodent and human L7 genes. Two of these are found in the structural gene portion. In mouse one is found in intron 2 and the other in intron 3 while in human two are located only 18 bases apart in intron 3. Loss of this part of the structural gene results in weak ectopic brain expression and loss of Purkinje cell expression, as we have shown using the L7trun-GFP construct. In both rodent and human the third N-box is found in the proximal portion of the promoter: in mouse it lies just upstream of the putative SRY/SOX-5 element (see Figure 3.1) while in human it lies proximal to the RORα binding site. All three N-boxes are inverted relative to the canonical CACNAG. Further ChIP and promoter mutagenesis studies will have to be performed to resolve the relevance of these sites. 81 In our study we showed that the cooperative action of the 0.9 kb L7
“pseudo-enhancer” and the structural gene are required for driving Purkinje cell- specific expression of the transgene. To further analyze this, we will conduct another expression analysis in vivo. To this end, we will prepare a set of constructs using a minimal promoter, heat-shock promoter (Hsp) (Figure 2.7). A single 0.9 kb L7 “pseudo-enhancer” and its concatamers will be attached to the
Hsp minimal promoter- LacZ cassette in two separate constructs (Figure 2.7A-B).
These constructs will contain the bovine growth hormone (BGH) poly(A) and 3’- end processing signals. Our expectation is that BGH 3’-end signals will not support Purkinje cell-specific expression, much as the SV40 sequences did not
(Figure 2.6D). The construct will express in the cerebellar Purkinje cells only when the L7 3’UTR and 3’-end signals replace BGH-3’ (Figure 2.7C), but will not work when these signals are added downstream of the BGH-3’ (Figure 2.7D).
Exons 1 up to the first half of exon 2 were included in previous construct where high levels of Purkinje cell-specific expression were obtained (Figure 2.6B and
E). Therefore, it is possible that some sequences in this region are also required for Purkinje cell-specific expression. If this is the case, a construct which include proximal half of the gene in the transcriptional unit (Figure 2.7E) will result in a high level Purkinje cell-specific expression in vivo.
82
Figure 2.7 Minimal promoter constructs for enhancer test in transgenic mice.
In summary, our results indicate different but cooperative actions of 0.9 kb
proximal promoter/enhancer and the structural gene in the control of gene
expression in vivo. Different constructs utilizing these sequences would be useful in vivo genetic engineering tools to better understand molecular and genetic
mechanisms of Purkinje cell-specific gene expression.
83
CHAPTER 3
ACTIVATION OF L7 GENE EXPRESSION BY RORα
3.1 ABSTRACT
L7 is a GoLoco domain protein which we hypothesize to function in the
“tuning” of P/Q type Ca2+ channels by the modulation of G protein-coupled receptors. L7 knock out (L7KO) mice have improved performance on accelerating rotarod, and they also display sensorimotor changes in the behavioral assays. L7 is specifically and abundantly expressed in Purkinje cells in the cerebellum. In this study we show that retinoic acid receptor related orphan nuclear receptor-α (RORα) activates L7 gene expression. The L7 promoter is activated by RORα in HEK293 cells, and RORα can bind to its putative binding site in the L7 promoter in vitro. Furthermore, RORα occupies in vivo the RORα response element in the L7 promoter, as shown by chromatin immunoprecipitation assay. L7 expression is greatly eliminated in staggerer 84 mutants, which lack RORα. Additional studies in vivo show that while L7 expression is dependent on RORα throughout the cerebellum, this dependence is greatest in the intermediate region between the vermis and far lateral hemispheres. Results presented here provide a combination of in vitro and in vivo analyses of L7 gene expression, and show activation of L7 by RORα. Our studies will also provide insight into the recently indicated role of RORα in Ca2+-
mediated signaling mechanisms in the cerebellum, and help us understand the
connection between the physiology of Purkinje cells and animal behavior.
85 3.2 INTRODUCTION
L7 is a GoLoco domain protein. Through the GoLoco domains, L7 can interact with inhibitory Gα subunits, Gαi and Gαo. With this interaction, L7 can
“tune” the activity of P/Q type Ca2+ channels by modulating G protein-coupled receptor-mediated inhibition of these channels (Kinoshita-Kawada et al., 2004).
L7KO mice have improved performance on accelerating rotarod, and they also present sexually dimorphic sensorimotor changes, ranging from audition to heat sensation (See Chapter 5). To better understand the physiological relevance of this phenotype, it is necessary to investigate control of L7 gene expression in the cerebellar Purkinje cells.
The L7 gene is exclusively expressed in Purkinje cells in the cerebellum.
Initial analyses on L7 gene expression showed that it is regulated spatially and temporally during cerebellum development (Smeyne et al., 1991). L7 expression starts at around E17 in the medial cerebellum and progresses in the medial- lateral direction as parasagittal bands as the cerebellum develops, until it is expressed uniformly throughout the cerebellum in adult. Promoter truncation analyses showed that truncation of the promoter from 4 kb to 1 kb still causes high levels of expression in the cerebellar Purkinje cells (Oberdick et al., 1993).
Furthermore, our enhancer test analysis suggested that 0.9 kb proximal promoter has enhancer-like properties (See Chapter 2, (Serinagaoglu et al., 2007). Based on these observations, we decided to examine L7 proximal promoter in more
86 detail to identify transcription factors and cis-acting elements that they interact with to control L7 gene expression.
Up to now several attempts have been made to identify positive and negative regulators of L7 expression in the Purkinje cells. Most of these studies included the identification of putative binding elements for transcription factors by sequence analysis, and the analysis of promoter activation in cell culture assays.
A few of the putative binding sites identified in the L7 promoter are cAMP- response element (CRE), AP-1 binding site, and an E-box motif (Anderson et al.,
1997; Anderson et al., 1998; Oberdick et al., 1993; Sanlioglu et al., 1998; Strait et al., 1992; Vandaele et al., 1991; Zou et al., 1994). Several transcription factors including homeodomain proteins and POU domain transcription factors have been shown to activate L7 gene expression in vitro. These studies mostly involved in vitro reporter gene assays, and lacked a detailed in vivo analysis.
Another transcription factor shown to activate L7 expression in vitro was
RORα. An RORα response element was found in the proximal promoter of L7
(Matsui, 1997; Schrader et al., 1996), and was shown to bind to in vitro translated
RORα protein in binding assays. Moreover, L7 promoter carrying the RORα response element was shown to be activated by RORα in vitro. Even though these two studies presented data for RORα-specific activation of L7 gene, these studies lacked in vivo analyses and a more sophisticated set of experiments to confirm specific binding of RORα protein to L7 promoter in vivo. In a more recent study, Gold et. al. analyzed several genes in the cerebellum and identified L7 as one of the targets of RORα (Gold et al., 2003). However, unlike the first two 87 studies, Gold et. al. focused on an RORα binding element found in a region ~2
kb upstream of the transcription initiation site. Based on these observations, we
decided to examine L7 proximal promoter and activation of L7 gene by RORα in
more detail. Here we report that RORα binds to and activates expression through
the L7 proximal promoter in vitro. We examined this interaction in vivo, and
showed that RORα occupies the RORα binding element in the proximal promoter
in vivo. L7 expression is greatly eliminated in staggerer mutants, which lack
RORα proteins. Additional studies in vivo show that while L7 expression is
dependent on RORα throughout the cerebellum, this dependence is greatest in
the intermediate region between the vermis and far lateral hemispheres. Our
results presented here will provide a better understanding of role of RORα in
Ca2+-mediated signaling mechanisms in the cerebellum, and help us understand the connection between the physiology of Purkinje cells and animal behavior.
88 3.3 MATERIALS AND METHODS
3.3.1 Mouse strains
All transgenic mouse lines were produced in FVB/N strain using Standard
Pronuclear Microinjection method in Transgenic Animal Facility at the Ohio State
University. The staggerer mice were in C57BL/6 background, and were originally obtained from the Jackson Laboratory (Bar Harbor, ME).
3.3.2 DNA Constructs
L7BG1, L7BG3, L7PromWT, L7PromΔROR, L7full-GFP and L7trun-GFP constructs were previously made in the lab. The pCMX-hRORα1 plasmid expressing RORα1 was kindly provided by Dr. Vincent Giguere, McGill University
Health Centre, Montreal, Canada (Giguere et al., 1994). pcDNA3.1 was obtained from Invitrogen, Carlsbad, CA. pEGFP-C3 was obtained from BD Biosciences
(Clontech), Mountain View, CA.
3.3.2.1 L7Prom2K
2 kb L7 promoter fragment was made by Polymerase chain reaction
(PCR) using EM1 vector, which carry 4 kb L7 promoter and 2 kb structural gene, as the template. The following primer sequences were used:
Forward: TCTTTAACGGATCCCAGTCCTTAACCTGCAAGGC BamHI
Reverse: AACTTAAAGGATCCATCTGATCTAAGAAACAGGA BamHI
89 The fragment was cut with BamHI, and cloned into TY-L7-LacZ vector in the BglII site. Orientation of the fragment was determined by sequencing. The entire L7Prom2K construct was excised from the vector backbone by XhoI and
SalI digestion prior to pronuclear injection (total construct size = 7 kb.).
3.3.3 Preparation of DNA for pronuclear injection
L7PromWT and L7PromΔROR plasmids were extracted using QiaFilter
Plasmid Maxi kit (12262, Qiagen, Inc., Valencia, CA). 50-60 µg of DNA for each construct were digested with suitable enzymes, and sent to Transgenic Animal
Facility for pronuclear injection.
3.3.4 Tail DNA extraction
A 1-2 mm piece of mouse tail was digested in 300 µl tail buffer (50 mM
Tris (pH=8.0), 100 mM EDTA and 0.5% SDS) in the presence of 1 mg/ml
Proteinase-K at 55°C overnight on a rotating shaker. After digestion, the mixture was phenol-chloroform extracted, and tail DNA was ethanol-precipitated. DNA was air-dried and dissolved in 250 µl TE buffer overnight at room temperature. 1
µl was used in PCR to identify transgenic mice carrying the construct.
3.3.5 Genotyping
3.3.5.1 Promoter constructs
To identify mice carrying expression constructs L7PromWT,
L7PromΔROR, and L7BG3, polymerase chain reaction (PCR) was run using primers specific to LacZ gene. In addition, a control PCR was carried out using 90 primers specific to L7 gene in order to check the efficiency of tail DNA extraction
procedure. The primer sequences were as follows:
LacZ set 1:
Forward: ACGCGCGAATTGAATTATGGCCCACACCAG
Reverse: GGCGCTCAGCTGGAATTCCGCCGATACTGA
PCR produces a 261 bp-fragment.
LacZ set 2:
Forward: CCATTGTCAGACATGTATACCCCGTACGTC
Reverse: GCCACCAATCCCCATATGGAAACCGTCGAT
PCR produces a 225 bp-fragment.
LacZ set 3:
Forward: ATTGACCCTAACGCCTGGGTCGAACGCTGG
Reverse: AACATCAACGGTAATCGCGATTTGACCTCT
PCR produces a 210 bp-fragment.
L7:
Forward: CGGACCAGGAAGGCTTCTTCAACCTGC
Reverse: ATCCCAGAACCCCAGCACTCCTGCCAC
PCR produces a 194 bp-fragment.
3.3.5.2 Staggerer mice
To identify genotypes of staggerer (sg) litters, polymerase chain reaction
(PCR) was run using primers specific to the WT and mutant alleles. PCR cycle
program was as follows:
91 • 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 54°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows: sg WT allele:
Forward (oIMR1233): TCTCCCTTCTCAGTCCTGACA
Reverse (oIMR1234): TATATTCCACCACACGGCAA
PCR produces a 318 bp-fragment. sg mutant allele:
Forward (oIMR1235): GATTGAAAGCTGACTCGTTCC
Reverse (oIMR1236): CGTTTGGCAAACTCCACC
PCR produces a 450 bp-fragment.
3.3.6 Perfusion
Transgenic mice carrying L7PromWT, L7PromΔROR, and L7BG3Xsg constructs were perfused with fixing solution containing 4% Paraformaldehyde in
1X PBS and 1X Blue I (0.12 M PIPES, 2 mM MgCl2 and 2 mM EGTA, pH=7.6).
Brain and cerebellum were dissected and further fixed in the fixing solution for one hour. Samples were washed with PBS twice and subjected to X-Gal staining procedure immediately.
3.3.7 X-Gal staining
Dissected brain samples were incubated in freshly prepared and filter- sterilized X-Gal (5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside) oxidation 92 solution containing X-Gal (1 mg/ml, in DMSO) and Blue II (4.8 mM potassium ferricyanide, 4.8 mM potassium ferrocyanide, 2 mM MgCl2, 0.02% NP40, 0.2 mM sodium deoxycholate and 1X PBS). For one cerebellum, 5 ml of the staining solutio n was used. The samples were incubated at 37°C for 4-5 hours or at room temperature overnight. Samples were washed with 1X PBS twice and stored in
1X PBS with 50 mM EDTA at 4°C. Whole mount pictures were ta ken using a camera attached to the dissecting microscope.
3.3.8 Cell transfection assays
3.3.8.1 Cell culture and transfection
HEK-293 cells were grown in 10 cm culture plates (Fisher, Pittsburgh, PA) at 37°C in a humidified incubator (5% CO2) in Dulbecco’s modified Eagle’s
medium (DMEM) (Sigma, St. Louis, MO) supplemented with 10% fetal bovine
serum (FBS). Twenty-four hours prior to transfection, cells were plated on 6-well plates with 2 ml DMEM (5×105 cells/well). Transient transfection was performed
using Lipofectamine and Plus reagents (Invitrogen, Carlsbad, CA) following the
manufacturer’s protocol. HEK293 cell stocks were kindly provided by Dr. Mike
Xhu, The Ohio State University, Center for Molecular Neurobiology, Columbus,
OH, 43210.
Each reporter construct was used to transfect HEK 293 cells. In one set,
they were co-transfected with the plasmid carrying RORα coding sequence, while
the other set received a balancer plasmid, pcDNA3.1, and two wells were
93 transfected in each set. In experiments with LacZ reporters, the pEGFP-C3 vector (CMV promoter) was used to normalize the transfection efficiency.
3.3.8.2 Visualization of LacZ expression
Cells were washed with 1X PBS 36 h after transfection and fixed with 2%
formaldehyde in 1X PBS solution by incubation at 37°C for 30 min. After washing
with 1X PBS, they were stained with freshly prepared and filter-sterilized X-Gal
(5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside) oxidation solution containing X-Gal (1 mg/ml, in DMSO) and Blue II (4.8 mM potassium ferricyanide, 4.8 mM potassium ferrocyanide, 2 mM MgCl2, 0.02% NP40, 0.2 mM sodium deoxycholate and 1X PBS). 0.5 ml staining solution was used for each well. Plates were incubated at 37°C for 4 h/overnight. Ten random areas in two culture wells per condition were examined under the microscope with 10X objective and LacZ-positive cells, which appeared blue, were counted. The average number of positive cells per field was calculated for each cotransfection set. The entire experiment was repeated twice and the two data sets were pooled.
3.3.8.3 Quantification of GFP fluorescence
Cells were plated on glass bottom dishes (Willcowells, D3522P, Warner
Inst, Hamden, CT). Medium was removed before imaging, and the glass-bottom well was filled with 1X PBS, and a circular coverslip (12-545-102 25cir-1, Fisher,
Pittsburgh, PA) placed over it. 8-bit color fluorescence images were collected on a Zeiss Axiophot upright microscope (20X objective) with an Optronics analog 94 camera (Optronics Engineering, Goleta, CA), converted to grey-scale in
Photoshop, and then imported as TIFFs into Image J (NIH). With each set of images the +RORα condition was consistently the brightest and a non-saturating exposure time was selected using these cells. Once in Image J, all visible cells were highlighted by segmenting each image with the image/adjust/threshold tool.
To measure the area occupied by the highlighted cells the analyze/measure tool was used. This relative measure should be analogous to cell counts, and this was confirmed by manually counting cells in some images. The fold-differences observed by manual cell counting was no different from that obtained using
Image J tools. The integrated pixel density was also determined in order to measure actual differences in intensities within the segmented areas using different constructs. Fold differences calculated using these measures were identical to those from area measures. The area data from 10 random fields were imported into Excel for processing and statistical analysis. The entire experiment was repeated four times.
3.3.9 Electrophoretic mobility shift assay (EMSA)
3.3.9.1 Preparation of nuclear extracts
One cerebellum is homogenized using a motor-driven homogenizer in 1ml of homogenization buffer (2 M sucrose, 10 mM HEPES (pH=7.6), 25 mM KCl,
0.15 mM spermine, 0.5 mM spermidine, 1 mM EDTA, and 10% Glycerol). For sg extracts, 2-3 sg cerebella were combined. The homogenate was layered on a cushion of homogenization buffer in ultracentrifuge tubes, and centrifuged at 95 24,000 rpm for 30 min at 4°C. The supernatant was removed. The pellet was resuspend in 80 μl of lysis buffer (10 mM HEPES (pH=7.6), 100 mM KCl, 3 mM
MgCl2, 0.1 mM EDTA, 10% Glycerol, and freshly added protease inhibitors: 1
mM DTT, and 0.1 mM PMSF). The nuclei were allowed to extract for 30 min with
continuous gentle mixing at 4°C. Extracted nuclei was centrifuged at 14000 X g
(13000 rpm) for 15 min at 4°C. The supernatant was transferred to a new tube
and dialyzed against dialysis buffer (10 mM Tris-HCl (pH=7.9), 1 mM EDTA, 5
mM MgCl2, 10 mM KCl, 10 % Glycerol, and freshly added protease inhibitors 1
mM DTT and 0.1 mM PMSF). For dialysis, Slide-A-Lyzer mini-dialysis tubes
(69552, Pierce/Fisher, Pittsburgh, PA) are used. Nuclear extract was removed
from the dialysis tube and centrifuged at 14000 X g (13000 rpm) for 10 min at
4°C. Protein concentration was determined using Lowry assay. The extracts were stored at -80°C.
3.3.9.2 In vitro translation
TNT® T7 Coupled Reticulocyte Lysate System (L-4610, Promega,
Madison, WI) was used to make RORα proteins from the pCMX-hRORα1 expression vector. Basically, 1 μg vector was mixed with TNT rabbit reticulocyte lysate, TNT reaction buffer, amino acid mixes, ribonuclease inhibitor, and nuclease-free water. T7 RNA polymerase was added, and the mixture was incubated at 30°C for 90 min. To check the efficiency of the reaction, Western blotting was carried out using anti-RORα antibodies (sc-6062, Santa Cruz
Biotechnology, Inc, Santa Cruz, CA). 96 3.3.9.3 Probe preparation
Double stranded (ds) probes are radioactively end-labeled using [γ-32P]
ATP by T4-polynucleotide kinase (T4-PK). 50 ng dsDNA was mixed with 5 μl [γ-
32P] ATP (specific activity: 3000 Ci/mmole, 10 μCi/μl), water, 10X T4-PK reaction buffer (final concentration 1X), and T4-PK (10 units). The mixture was incubated at 37°C for 30 min. Samples were run on 15% Polyacrylamide gel (40% acrylamide (Acrylamide:Bisacrylamide, 19:1)) in 1X TBE. One of the glass plates holding the gel was detached, and the gel was wrapped tightly using a plastic wrap. It was exposed onto the Classic Blue Autoradiography Film B-Plus
(EBA45, MidSci, St. Louis, MO). The gel and the film were aligned, and the DNA fragment of interest was located on the gel. The gel was excised, cut into smaller pieces, transferred to a 1.5 ml tube, and DNA probe was eluted. For the elution,
5 μg tRNA (carrier) and 300-400 μl elution buffer (0.5 M ammonium acetate, 10
mM magnesium acetate, 1 mM EDTA (pH 8.0), and 0.1% SDS) was added to the
gel pieces. The samples were incubated at 42°C for at least 4 hours. DNA was
extracted by phenol-chloroform, and precipitated by ethanol. The pellet was
dissolved in 50 μl TE, and radioactivity was measured using the scintillation
counter. Probe sequences are as follows:
97 L7Prom-WT probe:
GATCAAGTCTCTGGAGTCCCCTGACCCAGTTACTATAACACAC WT-RORE
L7Prom-ΔROR probe
GATCCAAGTCTCTGGAGTCCCCTCCAGTTACTATAACACACAG Δ-RORE
3.3.9.4 Preparation of EMSA gel
5 % native polyacrylamide gel was prepared using 40% acrylamide mix
(Acrylamide:Bisacrylamide, 79:1), and let polymerized horizontally for at least
half an hour at room temperature.
3.3.9.5 EMSA
In vitro translated RORα protein or cerebellar nuclear extract is incubated
with 1 μg poly(dIdC) in binding buffer (10 mM Tris (pH=7.9), 5 mM MgCl2, 50 mM
NaCl, 5% glycerol, 5% sucrose, 1mM EDTA, 1mM DTT and 0.1 mM PMSF) at
37°C for 15 min. 20,000 cpm probe is added into each reaction. Samples are incubated at RT for 15 min, and run on 5% native polyacrylamide gel in 0.5X TBE buffer for 4-5 h at 4°C. The gel is dried, and exposed to X-ray film at -80°C for
14-16 h. For supershift assay, 0.4 μg of RORα antibody (sc-6062, Santa Cruz
Biotechnology, Inc, Santa Cruz, CA) is added to each reaction and incubated at room temperature for 15 min.
98 3.3.10 Chromatin immunoprecipitation (ChIP) and PCR
3.3.10.1 ChIP
This protocol was adapted from Farnham Lab’s protocol
(http://genomics.ucdavis.edu/farnham/protocols/chips.html).
Freshly dissected cerebellum is chopped into small pieces with razor blade. Tissue pieces are transferred into a 1.5 ml tube and 1% formaldehyde in
1X PBS is added. After 15 min incubation at room temperature, crosslinking reaction is stopped by adding glycine (2.5 M stock, final concentration of 0.125
M). Samples are centrifuged at low speed (2000 rpm), and supernatant is removed. Pellet is washed with cold 1X PBS once. 1 ml homogenization buffer
(2M sucrose, 10 mM HEPES (pH=7.6), 25 mM KCl, 0.15 mM spermine, 1.5 mM spermidine, 1 mM EDTA, and 10% glycerol) is added to the pellet. Tissue pieces are homogenized by using a motor-driven homogenizer at 4°C. Homogenate is layered on 1 ml homogenization buffer cushion in 2 ml ultracentrifuge tube, and centrifuged at 24,000 rpm for 30 min at 4°C to pellet the nuclei. Supernatant is removed. Nuclei are resuspended in 300 μl nuclei lysis buffer (50 mM Tris-HCl
(pH=8.1), 10 mM EDTA, 1% NP40, 0.1 μM PMSF and protease inhibitor cocktail
(P8340, Sigma, Saint Louis, MO)). Sample is incubated on ice for 20 min.
Chromatin is sonicated to an average length of about 500 bp while keeping samples on ice (10 sec sonications with 30 sec intervals at power setting 4.0).
Sample is centrifuged at 14,000 rpm for 10 min at 4°C. Supernatant is transferred to a new tube. Samples are stored at -80°C overnight.
99 Next day, chromatin is pre-cleared by adding 50 μl blocked protein G-
sepharose beads. Samples are incubated on a rotating platform at 4°C for 1h,
and centrifuged at 14,000 for 10 min. Supernatant is transferred to a new tube,
and divided into two. A mock sample is also included which contains 1X dialysis
buffer (2 mM Tris-HCl (pH=8.0), 2 mM EDTA, 0.2% Sarkosyl) instead of
chromatin. Volume is adjusted to 300 μl by adding IP dilution buffer (16.7 mM
Tris-HCl (pH=8.1), 1.2 mM EDTA, 1% NP40, and 167 mM NaCl). 2 μg of RORα antibody (sc-6062, Santa Cruz Biotechnology, Inc, Santa Cruz, CA) is added to the first half of the sample. Samples are incubated on the rotating platform at room temperature for 2 hours and centrifuged at 14,000 rpm for 10 min at 4°C.
Supernatant is transferred to a new tube (this centrifugation step is carried out twice). 50 μl protein G-sepharose beads are added to each sample. Samples are incubated on the rotating platform for 2 h at room temperature. They are centrifuged at 2,000 rpm for 3 min. Supernatant from the “no antibody” sample is saved as “total input chromatin”. Pellets are washed 2X with 1.4 ml 1X dialysis buffer and 4X with wash buffer (100 mM Tris-HCl (pH=9.0), 500 mM LiCl, 1%
NP40 and 1% deoxycholic acid). For each wash, pellet is dissolved in buffer, incubated on the rotating platform for 3 min at room temperature, and centrifuged at 2,000 rpm for 3 min. After the last wash, samples are centrifuged again and last traces of buffer are removed. DNA/protein/antibody complexes are eluted by
150 μl freshly made elution buffer (50 mM sodium bicarbonate and 0.2% SDS).
For elution, samples are incubated on the rotating platform for 15 min at room temperature, and centrifuged at 14,000 rpm for 3 min. Supernatant is transferred 100 to a clean tube. Elution is repeated and both elutions are combined in the same
tube. Sample is centrifuged at 14,000 rpm for 5 min to remove any traces of the
pellet. Supernatants are transferred to clean tubes. 0.33 mg/ml RNase and 0.3 M
NaCl are added. Samples are incubated in the 67°C water bath for 4 h to reverse
formaldehyde crosslinking. 80 μl 5X Proteinase K buffer (50 mM Tris-HCl
(pH=7.5), 25 mM EDTA and 1.25% SDS) is added and samples are incubated at
45°C for 1 h. DNA is extracted once with 300 μl phenol:chloroform:isoamyl alcohol (25:24:1) and once with 300 μl chloroform:isoamyl alcohol (24:1). 30 μl
NaCl (5M), 1 μg tRNA, and 750 μl 100% Ethanol are added to each sample.
DNA is precipitated in -20°C freezer overnight.
In the morning, samples are centrifuged at 14,000 rpm for 20 min at 4°C.
Pellets are air-dried, and resuspended in 30 μl TE and DNA was purified with
Qiagen PCR purification kit (28104, Qiagen, Inc., Valencia, CA). 3-5 μl DNA is used as template in PCR.
3.3.10.2 PCR
GeneAmp PCR System 9600 (Waltham, MA) was used to carry out PCRs.
PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows:
101 PCR1 for proximal RORa binding site:
Forward: TCAGACCTTCTAGACAAGGT
Reverse: ACCTACTATCTCTTGGGGGA
PCR produces a 240 bp fragment.
PCR2 for distal RORa binding site
Forward: CAG TCC TTA ACC TGC AAG GC
Reverse: CCT GGA ACT CCT GCT GTC AT
PCR produces a 203 bp fragment.
PCR3 Control region around 1 kb of promoter:
Forward: GAAGCTACAGATGGTTGTGAGCTGTTATGT
Reverse: GAAACTACCGGATGCTCCTGTTTATCTGCC
PCR produces a 300 bp fragment.
3.3.11 In situ hybridization
3.3.11.1 Probe preparation
10 µg DNA templates for L7 (pL7hom1) and calbindin were linearized by
HindIII and PstI, respectively. DNA was purified using Qiagen PCR purification kit
(28104, Qiagen, Inc., Valencia, CA). Linearized templates were used in in vitro transcription. DNA templates were mixed with transcription buffer, DTT (final concentration 100 mM), RNase inhibitor (15518-012, Invitrogen, Carlsbad, CA),
ATP, GTP, CTP, SP6/T7 grade α-35S UTP (SJ603, Amersham, Piscataway, NJ), nuclease-free water and T7 RNA polymerase (13229-72, Invitrogen, Carlsbad,
CA). The mixture was incubated at 37 ºC for 1 h. 1 µl RNase-free DNase was 102 added to digest the template, and the mixture was further incubated at 37 ºC for
15 min. The volume was increased to 300 µl by DEPC-H2O, and RNA was
extracted by phenol-chloroform method. 25 µg tRNA and 8 M ammonium acetate
(final concentration 1 M) were added to the mixture, and RNA was precipitated by addition of 850 µl 100% Ethanol. The mixture was stored at -80ºC overnight.
Next morning, it was spun at highest speed for 15 min. The pellet was washed with 70% Ethanol, and it was air-dried. It was dissolved in 300 µl DEPC-H2O. 25
µg tRNA and 8 M ammonium acetate (final concentration 1 M) were added to the mixture, and RNA was re-precipitated by addition of 850 µl 100% Ethanol. The mixture was stored at -80 ºC for 30 min. Then, it was spun at highest speed for
15 min. The pellet was washed with 70% Ethanol, and it was air-dried. It was
dissolv ed in 30 µl DEPC-H2O.
3.3.11.2 Quantitation of radioactivity of α-35S UTP-labeled probe
1 µl of probe was diluted to 200 µl with DEPC-H2O. 1 µl of this dilution was precipitated in a separate tube by adding 500 µl, 2% BSA, and 1 ml 15% TCA, and storing on ice for 10 min. The precipitate was filtered with glass-fiber filter paper (1822-024, Fisher, Pittsburgh, PA) using a vacuum pump. The filter was washed twice with 5% TCA, and dried in a vacuum oven for 1 min. The radioactivity of the filtered paper was measured using a scintillation counter.
3.3.11.3 Tissue and section preparation
After the mouse is sacrificed, the head was placed immediately on ice for
5 min, and the brain was dissected. It was placed on dry ice for fast-freezing. The 103 frozen tissue was sectioned immediately, or stored at -80ºC for later use. For sectioning, cryostat was used, and the brain was sliced into 12 µm-thick sections.
Sections were placed on microscope slides as they were being sectioned (Two sections per slide). Slides were kept at -80ºC for storage.
3.3.11.4 Pretreatment of the slides
Immediately before the experiment, slides were taken out of the freezer, and kept at room temperature for 20 min. They were immersed into 4%
Paraformaldehyde in 1X PBS (pH=7.2) for 10 min. After rinsing twice in 1X PBS, they were immersed into 0.25% acetic anhydride in 0.1 M triethanolamine (TAE, pH=8.0) for 10 min. They were then subjected to a series of ethanol and chloroform washing: in 70% ethanol for 1 min, 80% ethanol for 1 min, 95% ethanol for 2 min, 100% ethanol for 2 min, 100% chloroform for 5 min, 100% ethanol for 1 min, and 95% ethanol for 1 min. The slides were air-dried at room temperature. All solutions were prepared using DEPC-H2O.
3.3.11.5 Hybridization
The probe was diluted in hybridization cocktail with 50% formamide (0973-
50 ml, Amresco, Solon, OH). The mixture was heated at 95ºC for 2 min to unfold secondary structure, and placed in ice-water. 5 M DTT was added to a final concentration of 0.1 M. 50 µl hybridization mixture was added on each slide.
Coverslip was placed, and the slides were sealed with Gurr (361254D, VWR Int.,
Poole, England). They were put in a closed plastic container, and sealed with
104 plastic wrap. The slides were then incubated at 55ºC overnight to allow the probe
to hybridize to targets.
3.3.11.6 Post-hybridization washing
The Gurr sealing was peeled off, and the slides were placed in 2X SSC at
room temperature. After the removal of the coverslip, they were washed twice
with 2X SSC plus 5 mM DTT at 45ºC for 15 min each. To remove the unbound
probe, they were then washed in RNase buffer with 20 µg/µl RNase at 37ºC for
30 min. After this, sections were washed with a series of SSC with decreasing
concentrations at increasing temperatures to further remove unbound probes,
and to obtain high stringency. For this purpose, the sections were washed five
times in 2X SSC at 37ºC for 15 min each, twice in 0.5X SSC at 37ºC for 15 min
each, and four times in 0.1X SSC at 75ºC for 15 min each. After the final wash,
the sections were dried in a series of ethanol washes: in 70% ethanol with 0.3 M
ammonium acetate (pH=7.4) for 1 min, 80% ethanol with 0.3 M ammonium
acetate (pH=7.4) for 1 min, 95% ethanol for 1 min, 100% ethanol for 5 min. The
sections were air dried, and exposed to Classic Blue Autoradiography Film B-
Plus (EBA45, MidSci, St. Louis, MO) overnight at room temperature without
using the intensifying screens.
3.3.11.7 Dipping the slides
The LM-1 emulsion for light microscopy (RPN40, Amersham, Piscataway,
NJ) was thawed at 45ºC for at least 30 min. The vertical glass container was
filled with the emulsion, and the slides were dipped. The emulsion on the back of 105 the slide was wiped away by tissue paper. The slides were leaned on a wall to remove the excess emulsion. They were put in a closed box, wrapped with aluminum foil, and stored at 4ºC for 1-2 weeks, depending on the strength of the signal. This protocol was carried out in the dark room with the red light on.
3.3.11.8 Developing the slides
The slides were developed in the dark room in D-19 developer for 4 min, rinsed with water for 30 sec, and fixed in fixer for 5 min. They were rinsed in the running distilled water for 30 min in the lab.
3.3.11.9 Counterstaining
The sections were stained with cresyl violet for 5 min at room temperature.
They were washed twice with water, and once for 5 min with 95% ethanol with a few drops of glacial acetic acid. They were rinsed with water, and air dried. They were then kept in Histoclear (HS-200, National Diagnostics, Atlanta, GA) for 10 min, coverslipped using Permount (SP15-100, Fisher, Pittsburgh, PA). Dark-field and bright field images were taken using the Zeiss microscope and MetaVue
Imaging Software.
3.3.12 Western blotting
Equal aliquots of protein extracts (10-15 μg/sample) were analyzed by
Western blotting using 12% SDS-polyacrylamide gel, and then blotted to
Hybond-C extra nitrocellulose membrane (Amersham, Piscataway, NJ) in the cold room (at 100 V for 1 h). The blots were blocked in 5% non-fat dried milk
106 powder in 1X PBS with 0.1% Tween-20 (PBST). Primary antibody was then added, and the blots were incubated at 4°C overnight. For L7, rabbit polyclonal anti-L7 antibody (1/500), for calbindin, mouse monoclonal anti-Calbindin-D-28K antibody (1/600) (c-9848, Sigma, St. Louis, MO), and for RORα goat polyclonal anti-RORα antibody (1/500) (sc-6062, Santa Cruz Biotechnology, Inc, Santa
Cruz, CA) were used. The blots were washed several times with 1X PBST, and incubated for 1 h at room temperature with 1/5000 dilution of horserarish peroxidase-conjugated IgG secondary antibodies in 5% milk solution described above. For L7, goat anti-rabbit IgG (A0545, Sigma, St. Louis, MO), for calbindin, goat anti-mouse IgG (A9917, Sigma, St. Louis, MO), and for RORα, donkey anti- goat IgG (705-035-003, Jackson ImmunoResearch Laboratories, West Grove,
PA) were used. After several washing steps in 1X PBST, the SuperSignal West
Pico Chemiluminescence Substrate (ECL) (34077, Pierce/Fisher, Pittsburgh, PA) was used to detect the protein signals with exposure to Classic Blue
Autoradiography Film B-Plus (EBA45, MidSci, St. Louis, MO).
107 3.4 RESULTS
3.4.1 Conservation of transcription factor binding sites in rodent and human L7 promoters
To identify candidate transcription factors which interact with the 0.9 kb proximal promoter/enhancer fragment, we compared mouse, rat and human L7 proximal promoter sequences. Direct Blast sequence comparison yielded no significant homologies. Then the promoter sequences from each species were individually scanned using the MotifFinder program (GenomeNet, Kyoto
University, Japan), which identified the positions of putative transcription factor binding sites. The rodent and human results were then compared and binding sites were identified that occupied the same relative position on the linear sequence. RORα response element TGACCC (inverted relative to the core sequence published previously; (Giguere et al., 1994)), POU/homeodomain consensus sequence RTAATNA (Verrijzer et al., 1992), Pbx/Meis1 consensus
core sequence TGACA(A/G) (Bertolino et al., 1995; Rave-Harel et al., 2004),
SRY/SOX5 consensus core sequence (A/T)(A/T)CAA(A/T) (Mertin et al., 1999),
and several SP1-like sites were identified in the proximal promoter (Figure 3.1).
Once we oriented the promoters from each species to have each consensus site to be aligned, we noticed that the precise relative spacing between the proposed sites was conserved. For example, RORE is 33 bases downstream of the putative homeodomain protein binding site in all species. Similarly, RORE was
108 Figure 3.1 Comparison of rodent and human Pcp-2(L7) promoter/enhancers. A number of core consensus transcription factor binding sites, including one for RORα, were identified with the same relative positions and orientations in the human and mouse promoters.
109
110 348 and 343 bases downstream of the putative SRY/SOX5 binding site in human
and in rodents, respectively. The conservation of these sites and similar relative
spacing between the sites among species strongly suggested that these sites are
conserved in evolution and are important in the transcriptional regulation of L7 gene expression.
3.4.2 L7 promoter activation by RORα in cell culture and inhibition by L7 structural gene
Next we focused on the activation of L7 promoter by RORα in vitro. To determine this, we prepared two different sets of promoter constructs. The first set carried varying lengths of L7 promoter and the L7-LacZ cassette (Figure
3.2A), and the second set included constructs with 0.9 kb proximal promoter and full and truncated versions of L7 structural gene with a GFP reporter (Figure
3.2B). HEK293 cells were then co-transfected with each construct and either
RORα expression vector or an empty vector. We then analyzed the number of cells expressing the reporter gene.
The first set included four promoter constructs: L7BG1, with a 4 kb L7 promoter fragment, L7Prom-2k with a 2 kb L7 promoter fragment, L7PromWT, with a 0.9 kb L7 promoter fragment with an intact RORE, and L7PromΔROR with the same 0.9 kb L7 promoter fragment as L7PromWT but with a 3 bp-deletion in the RORE (Figure 3.2A). Each construct was co-transfected into HEK293 cells with either an empty expression vector, pcDNA3.1 (Invitrogen, Carlsbad, CA) or
RORα expression vector, hCMX-RORα1 (Giguere et al., 1994). pEGFP-3C (BD
111
Figure 3.2 Constructs used in the transfection assay. A) All constructs have the L7 structural gene–LacZ cassette. Then, in descending order, promoters include 4 kb, 2 kb, 0.9 kb, 0.9 kb plus ΔROR mutation, and 0.5 kb. The last construct is not used in the transfection assay, but used in Figure 3.4F-H. B) These two constructs have the EGFP coding sequence in the full structural gene (L7Full-GFP) or in a truncated structural gene (L7trun-GFP). Both constructs have 0.9 kb promoter.
112 Biosciences/Clontech, Mountain View, CA) was also co-transfected in each case to normalize the transfection efficiency and to determine cell survival. To determine if presence of RORα protein affected cell survival, we analyzed and found the number of EGFP-expressing cells in +RORα and –RORα wells, indicating that presence or absence of RORα did not affect cell survival (Figure
3.3C). Then we focused our attention on the expression of the LacZ gene in plates with different combination of promoter constructs and RORα expression vector or pcDNA3.1. There was a 3-4 fold increase in the number of LacZ+ cells with the L7BG1, L7Prom-2k and L7PromWT constructs in the presence of RORα, compared to the –RORα wells (Figure 3.3A). In parallel, we tested L7PromΔROR construct in the presence or absence of RORα. L7PromΔROR construct had a 3 bp-deletion in the RORE in the proximal promoter and this deletion completely disrupted the interaction of RORα and promoter fragment, as we will explain in the following section. This mutated version did not show any changes in the number of LacZ+ cells in the presence or absence of RORα expression vector
(Figure 3.3A). As a result, we showed that the number of expressing cells decreased as the length of the promoter was decreased. Moreover, disruption of
RORE in the proximal L7 promoter caused unresponsiveness of the promoter construct to the presence of RORα protein. As a result we concluded that RORα activates L7 promoter in vitro.
The second set of the constructs were the same GFP constructs that we explained in Chapter 2. L7Full-GFP included 0.9 kb proximal promoter and L7-
GFP reporter gene (Figure 3.2B). L7Trun-GFP contained a truncated version of 113
Figure 3.3 RORα induction of the L7 promoter/enhancer in vitro and the effect of the L7SGC. A) Analysis of the LacZ constructs in HEK293 cells. B) Analysis of the GFP constructs (L7full-GFP and L7trun-GFP) in HEK293 cells. C) Analysis of cell survival in the presence or absence of RORα. D-L) Images depicting what is summarized in B. D-F) Low magnification (10X) images comparing expression of L7trun-GFP construct with (D) and without (E) RORα to expression of pEGFP-C3 (CMV-promoter) (F). The L7 gene promoter is very inactive in these cells compared to the CMV promoter. Nonetheless the increased expression in the presence of RORα is clear. G-H) High magnification (40X) view of similar cultures to those shown in D and E transfected with L7trun-GFP. Again, a substantial increase is observed in the presence of RORα. I and J) Thresholded versions of images in G and H showing relative areas occupied by visible cells (used to calculate area data in B). K and L) Thresholded images of cultures transfected with the L7full-GFP
114 the L7 gene that includes the 0.9 kb proximal promoter, and the structural gene missing the second half of the exon 2, exon 3 and 4, and introns 2 and 3. The
missing part in the gene was replaced with a reporter gene, green fluorescent
protein (GFP), and SV40 3’UTR fragment and this construct was called L7trun-
GFP (Figure 3.2B). The two constructs were transfected into HEK293 cells with
or without RORα expression vector. L7Full-GFP construct revealed 2-fold
increase in the GFP expression area in the presence of RORα, similar to what
was observed with LacZ construct (Figure 3.3B). In contrast, L7trun-GFP
construct was induced by more than 4-fold in the presence of RORα.
Furthermore, the basal expression, i.e. in the absence of RORα, of the latter was
nearly 2-fold greater than that of the former. These data supported a weak
repressor function for the L7 structural gene as we concluded in the in vivo
studies described in Chapter 2. They also showed that the repressor acts to
modulate the inducing effect of RORα. While this effect has so far been tested in
HEK293 cells, it may partially explain the absence of L7 expression in other non-
cerebellar tissues known to express RORα.
3.4.3 RORα can bind to its putative binding site in the proximal promoter in
vitro
We next investigated the interaction of RORα with RORE in the proximal
L7 promoter. We carried out electrophoretic mobility shift assay (EMSA) to
determine whether in vitro translated RORα protein or endogenous RORα protein
in the cerebellar nuclear extracts would interact with an L7 promoter fragment
115 containing RORE. In this assay, we used two 32P-labeled oligonucleotides
corresponding to the proximal RORE. The first probe, WT, contained intact
RORα binding site whereas the second one, Δ, had a 3 bp-deletion in the RORE.
This was the same mutation that we tested in the co-transfection assays with
L7PromΔROR construct in vitro (see section 3.4.2 above), and in a LacZ reporter
assay in vivo (see the following section). In EMSA, an intense shifted band was
observed (arrow) when we incubated in vitro translated RORα with WT probe, but this band was not present when the Δ probe was used (Figure 3.4A, Lanes 5 and 6). This major band was supershifted when we incubated the probe-protein mix with antibodies against RORα (Figure 3.4A, Lanes 7 and 8, doublet arrow).
Similarly, a major band shifted when we incubated WT probe with cerebellar nuclear extract (prepared from P10 mice) (Figure 3.4A, Lane 9), and this band was supershifted using antibodies against RORα (Figure 3.4A, Lane 11). This interaction was missing when Δ probe was used (Figure 3.4A, Lanes 10 and 12).
In all cases we observed multiple bands, but RORα antibodies caused a supershift of only one band, and this band was missing in the lanes where we
used Δ probe. The supershifted bands in each case ran as doublets (doublet arrow), presumably as a result of the multiple antibody species since RORα
antibody was polyclonal. In addition to these, we also tested nuclear extracts prepared from staggerer (sg) mutant cerebella (prepared from P10 mice). The
intensity of the band corresponding to RORα in the sg +/- cerebellum was less
than that corresponding to RORα in the sg +/+ cerebellum (Figure 3.4B, Lanes 3 and 5). This band was completely missing when nuclear extracts prepared from 116
Figure 3.4 Binding of RORα to the promoter/enhancer site at -62 bp in vitro and in vivo. A) EMSA and supershift analysis using in vitro translated RORα and cerebellar nuclear extracts. B) EMSA analysis using sg nuclear extracts. C) Chromatin immunoprecipitation assay. PCR1 is to detect proximal binding site, and PCR is to detect the distal binding site. PCR3 is to exclude the possibility that PCR1 and PCR2 regions were pulled down as one ling DNA fragment.
117 sg -/- cerebellum, which is devoid of RORα, was used (Figure 3.4B, Lane 7).
Thus, we concluded that RORα protein can bind to RORE in vitro.
3.4.4 RORα occupies the RORE in the proximal L7 promoter in vivo
EMSA described above showed that RORα protein can interact with
RORE in the proximal L7 promoter in vitro. To show that this interaction actually takes place in vivo and RORE in the proximal promoter is occupied by RORα protein in the cerebellum, we carried out chromatin immunoprecipitation (ChIP) assay. ChIP performed by another group indicated that another RORE in the distal promoter, around 2 kb upstream of the transcription start site, was occupied by RORα in P0 cerebella (Gold et al., 2003). Our own analysis of sequence in this region revealed an inverted RORE in mouse that lies 1782 bp upstream of the proximal one. In humans, there are two sites in this approximate position, one at 1727 bp, and another at 1868 upstream of the proximal one. The more distal site is inverted like the one in mouse. Therefore, although two distal sites are present in human and only one in mouse, the general position and spacing of all sites are conserved in rodents and human. To extend the observations of the other group, we performed ChIP assay using P10 cerebellum as opposed to P0. We chose this time because it represents the period of greatest increase in L7 gene expression in the cerebellum (Nordquist et al.,
1988; Oberdick et al., 1998), and because most of the in vivo reporter gene expression analysis we describe in this study and in our earlier work presented in
Chapter 2 involves post-natal animals. In this assay, we pulled down DNA-RORα
118 complexes in the cerebellum nuclear extracts using antibodies against RORα.
After the purification of the DNA fragments, we ran PCR to determine if both proximal and distal ROREs were pulled down. Both proximal and distal ROREs were bound by RORα protein in vivo (Figure 3.4C, PCR1 and PCR2). To determine if these sites were being pulled down separately instead of a long DNA fragment which contains both sites, we ran a control PCR corresponding to a fragment that lies mid-point between the proximal and distal ROREs. We did not expect to see the amplification of this fragment because we sheered DNA to 500 bp-long fragments by sonication prior to the pull-down step. As expected, the control primers did not amplify this middle region (Figure 3.4C, PCR3). These results showed that RORα interacts with the ROREs in the proximal and the distal L7 promoters in vivo at P10.
3.4.5 RORα controls both the overall level and medio-lateral pattern of L7 gene expression
If L7 is one of the targets of RORα, then we would expect L7 expression to be missing or greatly reduced in the surviving cells of sg mutant mice. To examine this, we performed in situ hybridization and western blotting to detect presence of L7 mRNA and protein, respectively in sg cerebella. While all or most
Purkinje cells in sg expressed relatively high levels of calbindin mRNA, a marker for Purkinje cells in the cerebellum (Figure 3.5B), very little L7 mRNA was detectable (Figure 3.5A). L7 mRNA was only detectable around the midline and the far lateral regions of the sg cerebellum (Figure 3.5B, arrowheads).
119 Expression of L7 mRNA in adult sg is more similar to expression at P0 in wild-
type (Figure 3.5C) than it is to adult wild-type (Figure 3.5D).
Western blotting revealed that L7 protein was reduced in sg +/- and
undetectable in sg -/- cerebellum (Figure 3.5E). In contrast, calbindin protein was clearly detectable in sg -/- cerebellum, but reduced relative to the +/+ due to extensive Purkinje cell loss. We concluded that L7 is very much reduced in sg cerebellum, but some mRNA was still detectable in restricted regions. The latter suggests a role for RORα in the medio-lateral pattern of L7 expression.
One way to examine the effect of RORα on cerebellar medio-lateral patterning is to test the expression of sagittal band markers in sg mutants. To do this, we crossed L7BG3 reporter transgenic mouse line (Figure 3.2A) (Oberdick et al., 1993) with sg mutants. This reporter line carries an L7-LacZ transgene with a 0.5 kb L7 proximal promoter, which includes the RORE, and it expresses in a pattern of sagittal stripes from E15 to adult. Unfortunately no expression was detected in the sg mutants (Figure 3.5H), consistent with the western blotting
results for the endogenous L7 protein. However, L7BG3 was detectable but
reduced in sg +/- cerebella (Figure 3.5G). In this case there was a selective loss
of expression of discrete stripes in the paravermal and intermediate zones, which
are the severely affected regions in the sg mutants.
Next we tested whether the RORα binding site in the proximal promoter could be shown to play a related role in L7 expression. We produced transgenic mice carrying L7PromWT and L7PromΔROR constructs described above (Figure
120
Figure 3.5 Regional effects of RORα in the cerebellum. A–D) In situ hybridization analysis of sg and wild-type cerebellum, L7 (A) and calbindin (B). Arrowheads in A shows residual L7 expression. Expression of L7 mRNA in adult sg is more similar to expression at P0 in wild-type (C) than it is to adult wild-type (D). E) Western blotting analysis of sg and wild-type cerebellum. F–H) Expression of L7BG3 reporter in sg cerebellum. I–L) Whole-mount staining for lacZ expression in the cerebella of adult transgenic mice expressing the L7Prom-WT construct (I) and the L7Prom-ΔROR construct.
121
Figure 3.6 Purkinje cell-specific expression in L7Prom-WT and L7Prom-ΔROR mice. A) Purkinje cell-specific expression of LacZ in L7Prom-WT mice (founder #39, injection #2). B) Spatially restricted and Purkinje cell-specific expression of LacZ in L7Prom-ΔROR mice (Line 32A). C) Magnified view of far lateral expression in L7Prom-ΔROR mice showing LacZ in Purkinje cells.
122 3.2A). Twenty-six founders were obtained carrying the L7PromWT constructs, and sixteen founders were obtained carrying L7PromΔROR construct (Table
3.1). Three lines were established for L7PromWT and five lines for
L7PromΔROR. The rest of the founders were sacrificed and analyzed directly for
LacZ expression in the brain. Thirteen out of twenty-six lines or founders carrying
L7PromWT construct had high levels of LacZ expression that was restricted to the cerebellar Purkinje cells (Table 3.1, Figure 3.6A). The staining pattern of whole-mount cerebella appeared uniform (Figure 3.5I). Seven other lines or founders had weak but uniform expression in the cerebellum. All other lines and founders carrying this construct had no expression in any brain region. Most lines or founders (twelve out of sixteen) carrying the L7PromΔROR construct had no detectable brain expression (Table 3.1, Figure 3.5J). Only one line (Line 32) had strong expression specific to the cerebellar Purkinje cells (Figure 3.6B-C). This line was resolved into two sub-lines, 32A and 32B. Both of these sub-lines displayed heterogeneous expression in the cerebellum defined by sharp boundaries along the medio-lateral axis. Sub-line 32A revealed a unique whole- mount staining pattern in the cerebellum, in which LacZ staining was restricted to the midline and far lateral regions (Figure 3.5K). Intermediate zones on either side of the midline showed no expression. Sub-line 32B showed a similar pattern that was observed previously with L7 promoter constructs with 0.5 kb or less promoter fragment (Figure 3.5L). In spite of the differences between these two patterns, one common observation was the near complete absence of expression in the intermediate zone that lies between the vermis (midline region) 123 and far lateral hemispheres. We concluded that expression of the reporter gene is mostly eliminated by mutation of the proximal RORE, and in rare transgenic lines that express L7PromΔROR construct (presumably due to the effects of favorable integration site), there is a tendency for this effect to be more severe in the intermediate zone than elsewhere. This matches the pattern of residual expression of endogenous L7 mRNA in sg mutants. Similarly, intermediate zones are the most severely affected regions in the sg mutants. Therefore, our studies support a heterogeneous function of RORα with respect to cerebellar development in the formation of the medio-lateral axis of the cerebellum.
124
Table 3.1 Effect of ΔROR enhancer mutation on transgene expression in the cerebellum.
125
Construct & Line or Founder # Level and pattern of expression in whole mount cerebellum L7Prom-WT inj. 1 Line 5 +; uniform 3 +; uniform Line 23 +++; uniform (See Figure 3.5I) 25 +++; uniform 35 +++; uniform Line 9 - 25-2 - 20 - 16 - inj. 2 4 - 10 +; uniform 16 ++; uniform 18 ++; uniform 23 +++; uniform 32 +; uniform 35 +++; uniform 39 +++; uniform inj. 3 6 +; uniform inj. 4 1 - 7 +; (cortex and olf. bulb stained very intensely) 9 +++ caudal; + rostral 11 ++; uniform 18 +++; uniform 22 ++; uniform 24 ++; uniform 25 +; uniform
L7Prom-ΔROR inj. 1 Line 1 - 2- Line 5a +; not stripy, but heterogeneous 5b - 8- 9- Line 10 +; lobule VI/VII only, but looks like GCs or pontine mossy fibers, not Purkinje cells(?) Line 11 - Line 32a ++; midline + far lateral only (See Figure 3.5K) Line 32b ++; stripy, excluded from intermediate zone (See Figure 3.5L) 41 - inj. 2 2 - 8- 11 - 12 - 15 -
126 3.5 DISCUSSION
In this chapter, we presented data demonstrating activation of the L7 gene by RORα, and interaction of RORα with its binding element in the L7 promoter in
vitro and in vivo. Following the results reported in Chapter 2, we focused our
attention on the 0.9 kb proximal promoter of L7, which behaved like a classic
position-independent enhancer for Purkinje cells in conjunction with the 2 kb
structural gene fragment. We identified an RORα binding site in this promoter region, -62 bp upstream of the transcription start site (relative to the cerebellum- specific transcription start +1 bp of Exon 1A (Zhang et al., 2002)). We showed that the L7 promoter is activated by RORα in vitro, and deletion of the RORα
binding element in the promoter results in reduction or complete elimination of
transgene expression in the cerebellum. We also provided in vitro and in vivo
evidence to show the interaction of RORα protein with its binding element in the
L7 promoter. In addition, expressi on of L7 was examined in the staggerer
mutants, which are natural mous e mutants with a deletion in the RORα gene.
Only traces of L7 mRNA were detected in the staggerer cerebellum, while L7 protein was completely absent.
Our co-transfection analysis is consistent with a previous study, in which 4 kb and 1 kb promoter sequences were attached to the L7 structural gene-LacZ cassette (Oberdick et al., 1993). R oughly 3-fold higher levels of Purkinje cell-
specific L7 expression were obtained in mice carrying the 4 kb vs. 1 kb promoter.
In our in vitro analysis, 4 kb, 2 kb and 1 kb promoters drove progressively lower
127 basal levels of LacZ expression in HEK293 cells in the absence of RORα protein.
In addition, they drove progressively lower absolute levels of RORα-inducible
expression. It is possible that at least part of this increased expression mediated
by sequences upstream of the -1 kb is due to the activity of the distal RORα
binding element around -1.8 kb. This site was shown to be occupied in vivo by
RORα in our study as well as in an earlier study from another group (Gold et al.,
2003). It is also possible that other transcription factor binding sites may affect
the increased transcription obtained with the longer promoter. Nevertheless, our
studies suggest that the specificity of expression in cerebellar Purkinje cells, in
conjunction with the structural gene, is directed by the 0.9 kb proximal promoter.
In addition, our studies with transgenic animals carrying 0.9 kb promoter with a deletion in the RORα binding element indicate that the proximal RORα site is the critical one for Purkinje cell-specific expression.
In this study, we further analyzed the repressor function of the L7 structural gene. In Chapter 2 we suggested that the L7 structural gene carries repressive signals that work to limit expression within the brain. This result is consistent with the in vitro studies using full length and truncated versions of L7-
GFP constructs, as reported in this chapter. At least in HEK 293 cells, the distal part of the structural gene had a dampening effect on the promoter induction by
RORα. This may provide a clue to explain why L7 gene expression is not observed in the many other tissues known to express RORα within and outside the brain.
128 RORα is Ca2+-responsive and can be activated by CaMKIV (Kane and
Means, 2000). In addition to its response to changing Ca2+ concentrations, RORα
further participates in Ca2+-signaling by activating expression of genes that
function in the the Ca2+-mediated signal transduction mechanisms (Gold et al.,
2003). This group of genes includes Pcp-4, a calmodulin inhibitor, Itpr1, IP3 receptor and calmodulin target, Cals1, and Hrp1 binding partner, Calb1, calcium binding protein, and L7. L7 was shown to be indirect modulator of the P/Q-type voltage-dependent Ca2+ channels (Kinoshita-Kawada et al., 2004). Our detailed
analysis of activation of L7 by RORα provides further support for the functioning of RORα in Ca2+-signaling.
129
CHAPTER 4
AN ANALYSIS OF L7 ISOFORMS EXPRESSED IN THE MOUSE
CEREBELLUM AND EYE
4.1 ABSTRACT
The mouse L7 gene is specifically expressed in cerebellar Purkinje cells and retinal bipolar neurons. Two isoforms of L7, L7A and L7B, are present in the rodent and human cerebellums, but it is not known which isoform is expressed in the eye. In addition, a third form has been reported to be found in the mouse eye in GenBank, although the size of this transcript differs significantly from the size of previously known L7 transcripts. Moreover, three previously unknown L7 isoforms are reported to be expressed in human testis and one isoform in human eye in GenBank. It is not known if L7 is expressed in mouse testis.
As we show in Chapter 3, only a trace amount of L7 mRNA is detected in staggerer (sg) cerebellums. L7 isoform found in trace levels in sg mice has not 130 yet been identified; thus it is not known how the absence of RORα in the cerebellum affects the choice of first exon.
To address these issues, we attempted to identify L7 isoforms expressed in wild-type mouse cerebellum and eye, human and mouse testis, and in sg cerebellum. In this chapter, we will present data to show that while L7A and L7B are the most abundant L7 isoforms in the cerebellum, a third form, which we call
L7C, is expressed at much lower levels. In the eye, however, only L7C is expressed. We also present the sequence of L7C cDNA, which was found to differ from that reported in GenBank. We also show that in human and mouse testis, no L7 transcripts are found. In addition, we present data showing that L7B is the only isoform expressed in sg cerebellum at much lower levels when compared to a wild-type cerebellum.
131 4.2 INTRODUCTION
L7 is a GoLoco domain protein expressed in cerebellar Purkinje cells and retinal bipolar neurons (Nordquist et al., 1988; Oberdick et al., 1998; Oberdick et al., 1990; Siderovski et al., 1999; Zhang et al., 2002). Initial analyses reported that L7 gene contains four exons, and spans a region of 2 kb on mouse chromosome 8 (Nordquist et al., 1988; Oberdick et al., 1998). Later, another isoform of L7 was identified in rodent cerebellum (Zhang et al., 2002). The first form was called L7A, while the second form was called L7B (Figure 4.1). In addition, human L7 gene structure was found to be very similar to the mouse gene, except the positioning of the first exon (Figure 4.1). In rodents, the sequence coding for the first exon of form A (exon 1A) is found upstream of that of form B (exon 1B). Both forms share exons 2-4. Exon 1A has no start codon; therefore L7A mRNA initiates translation in exon 2, producing a 99 aa-long protein. Exon 1B, on the other hand, has a start codon at its 3’ end, and this is in frame following its splicing to exon 2. As a result of splicing of exon 1B to exon 2, twenty-one amino acids are added to the N-terminus of the peptide, producing a
120 aa-long protein. These two proteins differ with respect to the number of
GoLoco domains; L7A carries one GoLoco domain, while L7B has two.
In humans, similar to rodents, two forms of L7 mRNA were identified by 5’
RACE that differ at their 5’ ends. Both forms are identical over a region showing high homology to the complete exons 2-4 in mice. Human L7B is very similar to its mouse counterpart as it includes exon 1B, which is highly homologous to
132 mouse exon 1B, spliced to exon 2, producing a 120 aa-long protein with two
GoLoco domains. The human L7A, however, has only three exons. The first exon
(hExon1A) is a composite of mouse exon 2 homologous region and a continuous
upstream region of 119 bp. This continuous region that forms the 5’ end of the
first exon in human L7A is homologous to the downstream region of mouse intron
1B. For simplicity and to highlight the homology of mouse, the second and third
exons of h1A are called exons 3 and 4. L7A in humans produces a 99 aa-long
protein with one GoLoco domain. In neither rodents nor humans, there is no
evidence of an mRNA structure carrying both exons 1A and 1B.
Figure 4.1. Comparison of rodent and human L7 gene structure and gene expression.
Although the two isoforms expressed in the cerebellum are well known,
the L7 isoform expressed in the retina has not been characterized. Recently, an
elaborate and multi-institutional study was carried out by the National Institutes of
Health Mammalian Gene Collection (MGC) Program to identify and sequence
cDNA clones containing complete open reading frames (ORFs) for each human
and mouse gene. It was reported in this analysis that a third form of L7 transcript 133 was identified in the mouse eye (Mammalian Gene Collection Program Team* et al., 2002). This eye-specific transcript is about 1500 bp-long, and includes a 56 bp-long first exon, which we call exon 1C, whose sequence is different than previously defined exon 1A and exon 1B. In addition, unlike forms A and B, it also contains the last intron, between exon 3 and exon 4, which results in a transcript with much larger size (Figure 4.2). This is in contrast to what was reported previously on L7A and L7B. Initial studies by Northern blotting showed that L7 mRNA had an estimated size of 550-600 bp (Nordquist et al., 1988).
Moreover, in our analyses, the L7 cDNA detected by reverse transcription-PCR
(RT-PCR) never had a size of ~1500 bp. For this reason, we decided to examine
L7 expression in the mouse eye, and find out the exact nature of this eye-specific transcript.
Figure 4.2. Comparison of L7 isoforms reported in mouse cerebellum and eye. Note that Exon 1C in this eye-specific transcript is located upstream of Exons 1A and 1B. Moreover, this reported cDNA sequence also includes the last intron as a part of the mature transcript.
134 In addition to L7A and L7B expressed in the cerebellum, two other isoforms were reported in GenBank to be expressed in the human testis, and a third isoform in human testis and eye. In both tissues, the first exons reported in the database were located upstream of exon 1B (Figure 4.3). It is not known if this testis-specific isoform is expressed in mouse testis. The isoform expressed in human eye was found at the 3’ end of the two testis-specific first exons (Figure
4.3), and this isoform was also found in human testis. The sequence of the eye- specific first exon in humans is not similar to the eye-specific exon sequence reported in mice.
Figure 4.3. Comparison of L7 isoforms reported in human cerebellum, testis and eye. Note that Exon 1C in the testis- and Exon 1E in the eye-specific transcripts are located upstream of Exons 1B.
135 As shown in Chapter 3, retinoic acid receptor-related orphan nuclear
receptor-α (RORα) binds to L7 promoter and activates L7 gene (Serinagaoglu et
al., 2007). A natural mutation of RORα gene in mice causes a phenotype known as staggerer (sg) (Hamilton et al., 1996). In sg mice, no L7 protein was produced, whereas only minute amounts of L7 mRNA can be detected by in situ
hybridization. The L7 isoform that is found in trace levels in sg mice has not yet
been characterized; thus how the absence of RORα in the cerebellum affects the
choice of first exon is not known. Moreover, the level of L7 mRNA in sg
cerebellum has not yet been measured.
Based on these observations, we decided to analyze L7 isoforms expressed in the wild-type mouse cerebellum, testis, and eye, in the human testis, and in the sg cerebellum. To this end, we first extracted total mRNA from the above-mentioned tissues, and performed reverse transcription-polymerase chain reaction (RT-PCR) to identify expression of alternative first exons. We found that while L7A and L7B are the most abundant isoforms in the cerebellum, the third form, L7C, is expressed in much lower levels. We also found that the
only isoform expressed in the eye is L7C. In human or mouse testis, we could not detect any L7 mRNA. Next we performed 5’-Rapid Amplification of cDNA Ends
(5’-RACE) to identify the sequences of alternative 5’ ends of L7 mRNA using total
mRNA isolated from mouse cerebellum and eye. We obtained the sequence of
the eye-specific isoform, L7C, which was found to differ than that presented in an
earlier study (Mammalian Gene Collection Program Team* et al., 2002). This isoform generates another form of L7 protein, which is 137 aa-long, and carries 136 only one GoLoco domain. In addition, we showed that L7B is the only isoform
expressed in sg cerebellum in much lower levels when compared to a wild-type cerebellum.
137 4.3 MATERIALS AND METHODS
4.3.1. Mouse strains
C57BL/6 strain was used to obtain mouse tissues. The staggerer mice
were in C57BL/6 background, and were originally obtained from the Jackson
Laboratory (Bar Harbor, ME).
4.3.2. Total RNA extraction
Total RNA was isolated from freshly dissected mouse tissues using TRIzol reagent (Invitrogen, Carlsbad, CA) following manufacturer’s instructions.
Basically, half of the cerebellum was homogenized using a glass Teflon homogenizer in 1 ml TRIzol reagent. Proteins and lipids were separated from the mixture by addition of chloroform, and RNA was precipitated using isopropanol.
RNA pellet was air dried and dissolved in RNase-free water. Total amount of
RNA was quantitated after measuring optical density of the sample. Samples were stored at -80°C.
4.3.3. DNase treatment
DNA-Free kit (Ambion, Austin, TX) was used to eliminate DNA contamination in RNA extracts. 10 µg total RNA extract was incubated with 2 units of DNase at 37°C for 30 min. 0.1 volume of DNase inactivation reagent was added, and the sample was incubated at room temperature, mixing occasionally.
Inactivation reagent was precipitated by centrifuging, and RNA in the supernatant
was transferred to a fresh tube. Samples were stored at -80°C. 138 4.3.4. Reverse Transcription (RT)
RETROscript kit (AM1710, Ambion, Austin, TX) was used to produce cDNA from total RNA extracted from cerebellum, eye and testis tissues. ~1 µg of
DNA-free RNA was incubated with Oligo(dT) primer and nuclease free water at
70°C for 3 min to heat-denature the RNA. After chilling the samples on ice, remaining RT components were added: RT buffer, dNTP mix, RNase inhibitor and MMLV-RT. Samples were incubated at 42°C for one hour, and at 92°C for 10 min to inactivate the RT. They were stored at -20°C.
4.3.5. Polymerase chain reaction (PCR)
GeneAmp PCR System 9600 (Waltham, MA) was used to carry out PCRs.
PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows:
L7:
Forward (L7-YS-1-F): AGGCTTCTTCAACCTGCAGA
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
RT-PCR produces a 234 bp fragment.
L7A:
Forward (L7-Exon1A-F): ATTCTTAGTACTGTCCCCCA
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
139 RT-PCR produces a 328 bp fragment.
L7B:
Forward (L7-Exon1B-F): CAGGTCAGGGAAGGAGACTC
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
RT-PCR produces a 319 bp fragment.
L7C:
Forward (L7-Exon1C-F): AGAGGGAGGCTCAGACCTTC
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
RT-PCR produces a 287 bp fragment.
β-actin:
Forward (bActin-2a-F): GCATGTGCAAAGCCGGCTTC
Reverse (bActin-2-R): GGGGTGTTGAAGGTCTCAAA
RT-PCR produces a 346 bp fragment.
Human testis:
Forward (L7human-tst-F): GAGAAGACGAGGAAGGCTCA
Reverse (hL7-R3): ATCAGTTTTTGGGGGCCGAG
Fragments w ere r esolved in 1.5% agarose gel in 1X Tris-acetate buffer (TAE).
4.3.6. Real-Time PCR
The iCycler iQ Real-time PCR Detection System was used along with the iQ SYBR Green Supermix following manufacturer’s instructions (1708882,
BioRad , Hercules, CA). The 96 well-format was used, and three reactions in three wells were run for each sample. Before the Real-time PCR, all primers
140 were confirmed to produce a PCR product that resolved as a single band with no primer–dimers on agarose gels.
The Real-time PCR cycle program was as follows:
• Automatically inserted steps to collect well factors: 30 sec at 95°C (2
cycles)
• 1st step: 15 min at 95°C
• 2nd step: 15 sec at 94°C
• 3rd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (60 cycles)
• 4th step: 10 sec starting at 95°C and decreasing temperature for 0.5°C at
each cycle (100 cycles: this is to draw the melt curve)
Same primers used in RT-PCR were used in Real-time PCR.
4.3.6.1. Determination of PCR efficiency for each primer set
To be able to compare the threshold cycle values from two different PCRs, it is important to use primer sets with similar efficiencies (95-100%). For this purpose, standard curve function of iCycler iQ Real-time PCR Detection System was used.
Each cDNA of interest (for example L7 and β-actin) was produced by RT-
PCR. The fragments were resolved in 1.5% gel, gel-extracted and cloned into pCR/8/GW/TOPO vector using TOPO TA cloning kit (Invitrogen, Carlsbad, CA).
Plasmids were extracted from bacterial culture, and DNA concentration was calculated. Serial dilutions of plasmid containing the cDNA of interest were prepared: 10, 1, 0.1, 0.01, and 0.001 ng/µl. Each cDNA dilution series were used
141 in Real-time PCR with its specific primers. Real-time PCR was run using three
wells for each dilution. Standard curve function was selected at the beginning of
the run. At the end of the cycle, the software provided the standard curve, and
automatically calculated the efficiency of the primer set used in that particular
reaction. Primers with efficiencies between 95 and 100% were then used in the
Real-time PCR with experimental cDNA samples.
4.3.6.2. Quantification of expression levels by Real-Time PCR
For quantification, the comparative CT method was used (Table 3 of User
Bulletin #2 of the ABI Prism 7700 Sequence Detection System). Briefly, the
average threshold cycle (CT) from three reactions of each RT sample was
determined. The ΔCT was calculated by subtracting the average actin CT value of
each sample from the average L7 (or the gene being investigated) CT value of
the same sample. The ΔΔCT value was determined by subtracting the ΔCT value of the sample with higher CT value from the ΔCT value of the other sample. The
level of gene of interest relative to the control gene (in our case, β-actin) was
determined by the equation 2−ΔΔCT.
The standard deviation (SD) for each sample was derived from the
square-root of the summed squares of the standard deviations of the average L7
and β-actin CT values, SD-L7, and SD-actin. Data are presented as ± SEM.
142 4.3.7 Genotyping
To identify genotypes of staggerer (sg) litters, polymerase chain reaction
(PCR) was run using primers specific to the WT and mutant alleles. PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 54°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows: sg WT allele:
Forward (oIMR1233): TCTCCCTTCTCAGTCCTGACA
Reverse (oIMR1234): TATATTCCACCACACGGCAA
PCR produces a 318 bp-fragment. sg mutant allele:
Forward (oIMR1235): GATTGAAAGCTGACTCGTTCC
Reverse (oIMR1236): CGTTTGGCAAACTCCACC
PCR produces a 450 bp-fragment.
4.3.8. 5’-Rapid amplificaition of cDNA ends (5’-RACE)
5’-RACE reaction is composed of four steps: Reverse transcription (RT), purification of cDNA, Recombinant terminal deoxynucleotidyl transferase (rTdT) tailing, and PCR.
143 4.3.8.1. Reverse Transcription (RT)
Reverse transcription was carried out using the same method described above, except one change. In the RT reaction, no RNase inhibitor was added.
After the RT, 1 µl RNase A (20 mg/ml) is added to the reaction to remove mRNA templates (12091-021, Invitrogen, Carlsbad, CA), and samples were incubated at
37°C for 30 min. After RT, samples were stored at -20°C.
4.3.8.2. Purification of cDNA
Because RT buffer and rTdT buffer compositions are different, cDNA
should be purified and eluted in water before the tailing reaction. For this
purpose, Qiagen PCR purification kit (28104) was used (Qiagen, Inc., Valencia,
CA), and samples were eluted in 50 µl water that was pre-warmed to 65°C. After
the purification, samples were stored at -20°C.
4.3.8.3. Recombinant terminal deoxynucleotidyl transferase (rTdT) tailing
rTdT tailing was carried out using 10 µl purified cDNA for each sample.
First, cDNA was mixed with 1X tailing buffer and 40 µM dCTP in a 24 µl reaction, and this mixture was incubated at 94°C for 2 min. Samples were chilled on ice for
1 min, and 1 µl rTdT was added (10533-065) (Invitrogen, Carlsbad, CA).
Samples then were incubated at 37°C for 1 h, and were stored at -20°C.
144 4.3.8.4. Polymerase chain reaction (PCR)
After rTdT tailing, 2 µl of each sample was subsequently used in PCR with the following primers:
Anchored abridged primer-5’:
GGCCACGCGTCGACTAGTACGGGIIGGGIIGGGII
L7 primer, reverse (L7-YS-1-R):
CGTTTCTGCATTCCATCCTT
PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
After the first round of PCR with 30 cycles, nested PCR was carried out. 1
µl PCR sample from the first reaction was diluted 100 times with TE, and 1 µl from this dilution was used as a template in the second round of PCR using the same primers. The fragments were resolved in 2% agarose gel in TAE buffer.
Each band was cut out and gel-extracted using Qiagen gel extraction kit (28704)
(Qiagen, Inc., Valencia, CA). Fragments extracted were cloned into pCR 4-TOPO vector using TOPO TA cloning kit (Invitrogen, Carlsbad, CA). Several colonies were picked; plasmids were extracted, and sent to sequencing. L7 reverse primer
(L7-YS-1-R) was used in sequencing reaction. Because the templates had a G- rich stretch, dGTP kit was used in the sequencing reaction.
145 4.4. RESULTS
4.4.1. L7 expression analysis in mouse cerebellum and eye
Before analyzing L7 isoforms expressed in mouse cerebellum and eye, we first performed reverse transcription-PCR (RT-PCR) using RNA extracted from each tissue from wild-type mice (Postnatal day 21, P21). RT-PCR showed that
L7 was expressed in both cerebellum and eye in the mouse (Figure 4.4A). β-
Actin was also detected in both tissues (Figure 4.4-B). Further analysis by Real-
Time PCR revealed that the level of expression of total L7 was 8-fold higher in the cerebellum than in the eye (Figure 4.4C).
Figure 4.4 RT-PCR analysis of L7 expression in mouse cerebellum and eye. A) L7 primers for the common region of isoforms were used to detect total L7 mRNA. L7 was detected in both cerebellum and eye. B) β-Actin was detected in both samples. C) Actin-normalized relative expression levels of L7 mRNAs in the cerebellum and eye, determined using Real-time PCR and the comparative CT method.
146 In a search to identify the mRNA forms transcribed in mouse cerebellum
and eye, first, total RNA was extracted from each tissue from wild-type mice
(Postnatal day 21, P21), and 5’-RACE was performed. An Oligo-dT primer was used in the RT reaction to produce cDNA from mRNA molecules. Then poly(C) tails were added to cDNA molecules by rTdT. L7 cDNA was amplified using a G- rich anchored abridged forward primer and an L7-specific reverse primer, which corresponds to 20 bases in exon 4. After one round of nested PCR using the same primer pair, two bands in the cerebellum and one band in the eye were resolved on agarose gel (Figure 4.5).
Figure 4.5 5’-RACE analysis using RNA extracts from the cerebellum and the eye. In the cerebellum two products were observed, whereas in the eye only one product was detected. Each band was cut out, extracted from the gel, cloned into a vector, and sequenced.
Each band was purified, cloned into a vector, and sequenced. In the
cerebellum, two out of seven clones were found to carry L7A cDNA, while five contained a shorter L7B. The latter group carried last 20-35 bases of the previously published sequence of the exon 1B (Figure 4.6) (Zhang et al., 2002).
Nevertheless, because the start codon is found at the very 3’-end of the exon 1B, which was present in all of the fragments sequenced, protein sequence predicted
147 for this isoform was not affected. None of the clones sequenced contained form-
C sequence.
In the eye, only the third isoform, which we call L7C, was found in all three
clones sequenced. The exon sequence we identified is 19 bp longer than that reported in GenBank (Mammalian Gene Collection Program Team* et al., 2002).
In addition, this isoform we identified did not include the intron sequence between exons 3-4 reported in the database (Figure 4.7). Our result supports the previously published Northern blotting analysis where L7 cDNA was shown to be
550-600 bp-long (Nordquist et al., 1988). Exon 1C carries a start codon, which
results in the production of a 136 aa-long protein, which we call L7C. L7C
includes 17 aa at its N-terminal which are not found in forms A and B (Figure
4.8). This 17 aa-long peptide does not have any homology to any other known protein domains. Nevertheless, L7C carries two GoLoco domains, like L7B.
The sequence analysis of three L7 isoforms can be seen in Figure 4.6.
After obtaining the sequences, we performed RT-PCR for each form to
identify expression of alternative first exons in the cerebellum and eye. L7 mRNA
was detected in both cerebellum and the eye (Figure 4.9A). L7A and L7B forms
were detected in the cerebellum only (Figure 4.9A-B). L7C was detected in the
cerebellum and eye, although the expression level in the cerebellum was much
lower than that in the eye (Figure 4.9C). β-Actin was detected in all tissues at
similar levels (Figure 4.9D). Expression of L7C in the eye was 37-fold more than
the expression of L7C in the cerebellum (Figure 4.4E).
148 Figure 4.6 Sequence analysis of L7 gene in mice. Sequence analysis reveals three transcriptional start site positions (indicated by arrows). Exon sequences are in bold and underlined. Start codons are indicated in red boxes. Alternative transcriptional start sites for Exon 1B revealed by 5’-RACE are indicated by blue- dotted arrows.
149
150
Figure 4.7 Comparison of L7 isoforms expressed in mouse cerebellum and eye. Note that Exon 1C we obtained in this eye-specific transcript is also located upstream of Exons 1A and 1B. However, it is 81 bases longer than that reported in GenBank. Moreover, as opposed to the cDNA sequence reported in GenBank, our cDNA sequence did not include the last intron as a part of the mature transcript.
Figure 4.8 Comparison of protein sequences of three L7 isoforms. The GoLoco motifs are indicated by boxes.
151
Figure 4.9 RT-PCR analysis of expression of L7 isoforms in mouse cerebellum and eye. A) L7 primers for the common region of isoforms were used to detect total L7 mRNA. L7 was detected in both cerebellum and eye. B) Forward primers used were in Exon 1A and Exon 1B in two separate reactions. Forms A and B were detected only in the cerebellum, but not in the eye. C) Forward primer was in Exon 1C. Form C was detected both in the cerebellum and the eye, although the level of expression in the cerebellum was lower than that in the eye. D) β-Actin was detected in both samples. E) Actin-normalized relative expression levels of L7C mRNAs in the cerebellum and eye, determined using Real-time PCR and the comparative CT method.
152 4.4.2. L7 expression analysis in human and mouse testis
As mentioned previously, in addition to L7A and L7B expressed in the cerebellum, three other isoforms were reported to be expressed in the human testis and eye (Figure 4.11). In both testis and eye, all three exons reported in
GenBank were located upstream of exon 1B. One form is reported in both human testis and eye (GenBank accession numbers AK130855 and BX647746, respectively). The sequence of the eye-specific first exon in humans does not carry any homology to the mouse eye-specific exon. The other two forms are reported only in the testis (GenBank accession numbers BC038715 and
BC025387).
In order to confirm that L7 is expressed in human testis, and to determine whether it is also expressed in mouse testis, we carried our RT-PCR with both tissues. Although we detected β-Actin in mouse and human testis (Figure 4.10B), we did not detect L7 using primers that would amplify the common region of L7 isoforms (Figure 4.10A).
Figure 4.10 RT-PCR analysis of expression of L7 isoforms in mouse and human cerebellum, and mouse testis and liver. A) L7 primers for the common region of isoforms were used to detect total L7 mRNA. L7 was detected in only cerebellum. B) β-Actin was detected in all samples.
153
Figure 4.11 Sequence analysis of L7 gene in humans. Sequence analysis of cDNA forms in human cerebellum reveals two transcriptional start site positions (indicated by arrows). Alternative transcriptional start sites were reported in GenBank (testis-specific forms are indicated by orange-dotted arrows, and eye-specific form is indicated by pink- dotted arrow). Exon sequences are in bold and underlined. Start codons are indicated in red boxes.
154
155 4.4.3. L7 expression analysis in sg cerebellum
Next, we repeated RT-PCR and 5’-RACE analysis using total RNA
samples extracted from WT and sg cerebellums (P14). RT-PCR revealed
expression of L7 in sg cerebellum at lower levels (Figure 4.12A). β-Actin was
detected in all tissues at similar levels (Figure 4.12B). Further analysis by Real-
Time PCR showed that the level of expression of total L7 was 104-fold higher in
the WT cerebellum than in the sg cerebellum (Figure 4.12C). This confirms the in
situ hybridization data we presented in Chapter 3, in which only residual levels of
L7 mRNA was shown to be detected in sg cerebellum (Serinagaoglu et al.,
2007).
Figure 4.12 RT-PCR analysis of expression of L7 in sg +/+, +/-, and -/- cerebellums A) L7 primers for the common region of isoforms were used to detect total L7 mRNA. L7 was detected in all three genotypes, although the level of L7 was lower in sg -/-. B) β-Actin was detected in all samples. C) Actin-normalized relative expression levels of L7 in WT and sg cerebellums, determined using Real-time PCR and the comparative CT method.
156 To determine the exact sequences of the isoform(s) expressed in the sg
cerebellum, we then performed 5’-RACE analysis using total RNA isolated from
P14 mice. An Oligo-dT primer was used in the RT reaction to produce cDNA
from mRNA molecules. Then poly(C) tails were added to cDNA molecules by
rTdT. L7 cDNA was amplified using a G-rich anchored abridged forward primer
and an L7 specific reverse primer, which corresponds to 20 bases in exon 4.
After one round of nested PCR using the same primer pair, two bands in the WT
cerebellum and one band in the sg cerebellum were resolved on agarose gel
(Figure 4.13). Each band was purified, cloned into a vector, and sequenced. In
the WT cerebellum, one out of eleven clones carried cDNA for the form A, while
seven out of eleven contained a shorter form B. Similar to the previous results with P21 cerebellum, the latter group carried last 31-35 bases of the previously published sequence of the exon 1B (Zhang et al., 2002). Furthermore, three out of eleven clones contained the intron between exon 1B and exon 2 as a part of the cDNA, which was never observed with P21 cerebellum. While we do not know the reason for the presence of these intron sequences in L7 cDNA, it is
possible that these are splicing artifacts.
In the sg cerebellum, we identified the sequences of form A (one out of seven) and form B (six out of seven). Similar to what we observed with P21 and
P14 WT cerebellums, exon 1B detected in the sg cerebellum contains the last
31-35 bases in the exon 1B reported previously.
We then performed RT-PCR for each form in the WT and sg cerebellums.
L7 mRNA was detected in both samples, but the level of expression was lower in 157 the sg cerebellum (Figure 4.14A). L7A and L7B were detected in both samples, although the level was lower in the sg cerebellum, especially for L7A (Figure
4.14B and C). L7C was not detected in the sg cerebellum (Figure 4.14D). β-Actin was detected in all tissues at similar levels (Figure 4.14E).
158
Figure 4.13 5’-RACE analysis using RNA extracts from the WT and sg cerebellums. In the WT cerebellum, even though they were not well-separated, two products were observed, whereas in the sg cerebellum only one product was detected. Each band was cut out, extracted from the gel, cloned into a vector, and sequenced.
Figure 4.14 RT-PCR analysis of expression of L7 isoforms in sg +/+, and -/- cerebellums A) L7 primers for the common region of isoforms were used to detect total L7 mRNA. L7 was detected in both genotypes, although the level of L7 was lower in sg -/-. B) L71A is not detected in sg -/-, nor is L71C (D). C) L71B is detected in both sg +/+ and -/-. E) β-Actin was detected in all samples.
159 4.5. DISCUSSION
Two cerebellum-specific L7 isoforms in mice have been identified so far.
Here we report the expression of a third L7 isoform in mouse eye. This new form differs from L7A and L7B at its 5’-end and is produced by usage of an alternative
first exon, called exon 1C. A translation initiation codon is found in exon 1C. It is
predicted that a 136 aa-long protein can be transcribed from the L7C mRNA, and this protein contains two GoLoco domains, like L7B. Seventeen amino acids found at the N-term of L7C are not present in forms A or B, and do not match with any known domain sequences when scanned using ScanProsite by the
ExPASy (Expert Protein Analysis System) proteomics server of the Swiss
Institute of Bioinformatics (http://www.expasy.ch/tools/scanprosite) and
InterProScan by European Bioinformatics Institute
(http://www.ebi.ac.uk/InterProScan). It is possible that this 5’ region may be
important for mRNA localization or translation in the eye.
L7C is the only isoform expressed in the eye, but it is found at very low
levels in the cerebellum. In fact, expression of L7C is 37-fold less in the
cerebellum than in the eye. One reason for this may be the presence of eye-
specific transcription factors directing the start of transcription at exon 1C in the
eye, but not in the cerebellum. Another reason may be that the distal RORα
binding site found around 2 kb upstream of the transcription initiation site of L7
takes a more significant role in the eye-specific expression.
160 L7C cDNA sequence we identified is different than that reported in
GenBank. The exon sequence identified in our study contains 19 more bases at
its 5’-end. However, addition of these bases does not change the protein
sequence, since they are found at the 5’ untranslated region of the mRNA
molecule. Furthermore, none of the cDNA sequences include the intron between
exon 3 and 4. This is also supported by our RT-PCR study, where we never
detected an L7 cDNA fragment with a size of 1500 bp. The cDNA sequence in
GenBank may be a result of incomplete splicing.
Although L7 was reported to be expressed in human testis, we could not
detect it by RT-PCR. We could not detect L7 in mouse testis, either. Therefore, we cannot confirm the previous results reported in GenBank. However, the human testis sample that we used to extract RNA was not fresh. It is possible
that mRNA was degraded in this tissue, so we did not obtain high quality cDNA
to begin with. We will repeat this analysis using a fresh tissue. We will also obtain
an eye sample and try to identify human eye-specific transcript reported in
GenBank.
In sg cerebellum, we detected only trace levels of L7 mRNA by in situ
hybridization (Serinagaoglu et al., 2007). Here we confirm this result by Real-
Time PCR analysis, where expression of L7 in sg cerebellum was found to be
104-fold less than that in WT cerebellum. 5’-RACE analysis revealed the
sequences of the isoform expressed in sg cerebellum, which are L7A and L7B.
However, the level of L7A seems to be much lower than that of L7B as indicated
by RT-PCR analysis. In fact, we could obtain L7A sequence by 5’RACE only in 161 one clone, which further supports our expression analysis by RT-PCR. Since the
level of L7 expression is much lower in the sg cerebellum, we conclude that
RORα is required for expression of both isoforms in the cerebellum, and usage of exon 1A is affected more by the absence of RORα.
162
CHAPTER 5
BEHAVIORAL ANALYSIS OF L7KO MICE
5.1 ABSTRACT
L7 is a GoLoco domain protein specifically expressed in cerebellar
Purkinje cells. L7 interacts with Gαi and Gαo subunits of heterotrimeric G
proteins, and regulates the degree of Gαi/o-mediated inhibition of Cav2.1 (P/Q-
type) voltage-dependent Ca2+ channels in a dose dependent manner. Initial behavioral and neuroanatomical analysis of L7 knock-out (KO) mice indicated that L7KO mice displayed no difference when compared to wild type (WT) mice.
We have expanded the behavioral analysis of these mutant mice to include a variety of sensory and motor tests. Our detailed analysis of WT and L7KO mice on accelerating rotarod shows gests that mutant mice have improved maximal performance on this motor learning assay. We also show sensorimotor changes in L7KO mice ranging from audition to heat sensation, and that these defects are 163 sexually dimorphic; the mutant males are more sensitive than all other
combinations of genotype and sex. These changes are due to a loss of L7
protein in the cerebellum, and not due to sensory organ defects, supported by electrophysiological analysis of Purkinje cell firing patterns, behavioral assays
that indicate sensory organs have normal function, and sensitive expression
analysis to show mRNA is not detectable in any other brain region or in sensory
tissues of WTs. We conclude that the cerebellum plays a role in sensorimotor
gating and functions in the mediation of sensory response, which is a non- traditional role for cerebellum.
164 5.2 INTRODUCTION
L7 is a GoLoco domain protein specifically expressed in the Purkinje cells in the cerebellum and localized in the Purkinje cell bodies and dendrites
(Oberdick et al., 1988; Siderovski et al., 1999; Zhang et al., 2002). The GoLoco domain was discovered in 1999 as one of the conserved domains found in several proteins of the Regulator of G-protein Signaling (RGS) family, and in other proteins which are not in the RGS family, including L7 (Siderovski et al.,
1999). The GoLoco domain typically acts as a binding domain and guanine nucleotide dissociation inhibitor (GDI) for Gαi and Gαo subunits (Kimple et al.,
2001; Kimple et al., 2002; Willard et al., 2004). L7 directly interacts with these subunits (Kimple et al., 2002; Luo and Denker, 1999; Natochin et al., 2001). It was later suggested that L7 regulates the degree of Gi/o-mediated inhibition of
Cav2.1, or P/Q-type voltage-dependent Ca2+ channels, in a dose dependent manner (Kinoshita-Kawada et al., 2004). However, how L7 protein affects cerebellum functioning has not been widely examined or fully understood.
Long before the analysis of L7 protein, L7/Pcp-2-knock-out (KO) mice were generated by two separate groups to be able to understand the effects of
L7 protein loss on cerebellar functioning (Mohn et al., 1997; Vassileva et al.,
1997). Initial analyses of L7KO animals focused on neuroanatomical and basic behavioral differences and concluded that there is no significant difference between WT and L7KO mice (Mohn et al., 1997; Vassileva et al., 1997). L7KO mice developed normally both pre- and postnatally, and they had similar total
165 body and cerebellum weights. The laminar structure of the cerebellum, gross
cellular anatomy, cellular organization and the number of cells were compared between WT and L7KO mice, and no significant difference was observed. L7KO mice did not show any obvious behavioral changes, for example ataxia or loss of balance. Behavioral analysis by constant velocity rotarod test also did not reveal any differences between WT and L7KO animals. These observations suggested that either L7 plays a little role in the cerebellar Purkinje cells, or its function can be compensated by other unknown factors in its absence. Nevertheless, when these neuroanatomical and simple behavioral analyses were carried out, the signaling function of the L7 protein was not known. Thus, it is possible that a broader range of tests are necessary to detect any subtle behavioral changes in
L7KO mice.
Based on the above observations and discussions, we decided to re-
examine the phenotype of L7KO mice using behavioral tests to analyze anxiety
behaviors, motor learning, and sensory responsiveness. Here we report that
L7KO mice show improved performance on a motor learning assay, and altered
sensory responsiveness to stimuli ranging from sound to heat. We also show that
these changes are sexually dimorphic, meaning in particular that the mutant
males are more sensitive to stimuli than all other sex-genotype combinations. We
show that these changes are due solely to cerebellar response to the lack of L7
protein and not to sensory organ deficits. We support this by behavioral assays
showing normal functioning, by electrophysiological analysis of Purkinje cell firing
patterns, and by sensitive expression analysis which confirmed that L7 mRNA 166 was not detectable in other brain regions or sensory tissues of WT mice. Our data support a non-traditional role of cerebellum in sensory responsiveness.
167 5.3 MATERIALS AND METHODS
All experiments were carried out by the author unless otherwise stated.
5.3.1 Mouse strains
L7KO homozygous breeders were obtained from St. Jude Children’s
Research Hospital (Vassileva et al., 1997). L7KO heterozygotes (L7+/-) were crossed in each generation to commercially obtained stock C57BL/6NTac mice
(Taconic), and the L7+/- pups identified by PCR of tail biopsy. All animals in the behavioral and physiological studies were obtained from crosses between L7+/- parents that were 8-10 generations in C57BL/6. All animals were 4-6 months old at the time of behavioral testing, except for the accelerating rotarod studies in which the animals were 5-6 weeks old. Five different animal sets were used for behavioral studies.
All experiments on animals were conducted in compliance with the guidelines for animal research described in “PHS Policy On Humane Care and Use Of
Laboratory Animals” and the PHS “Guide for the Care and Use of Laboratory
Animals” from the U.S. Dept. of Health and Human Services.
5.3.2 Tail DNA extraction
A 1-2 mm piece of mouse tail was digested in 300 µl tail buffer (50 mM
Tris (pH=8.0), 100 mM EDTA and 0.5% SDS) in the presence of 1 mg/ml
Proteinase-K at 55°C overnight on a rotating shaker. After digestion, the mixture was phenol-chloroform extracted, and tail DNA was ethanol-precipitated. DNA
168 was air-dried and dissolved in 250 µl TE buffer overnight at room temperature. 1
µl was used in PCR to identify transgenic mice carrying the construct.
5.3.3 Genotyping
To identify mice carrying expression constructs L7PromWT,
L7PromΔROR, and L7BG3, polymerase chain reaction (PCR) was run using primers specific to LacZ gene. In addition, a control PCR was carried out using primers specific to L7 gene in order to check the efficiency of tail DNA extraction procedure. The primer sequences were as follows:
L7-WT allele:
Forward (L7KO-5'): CGGACCAGGAAGGCTTCTTCAACCTGC
Reverse (L7KO-3'): ATCCCAGAACCCCAGCACTCCTGCCAC
PCR produces a 194 bp-fragment.
L7-KO allele:
Forward (L7Forward-3): GCGCGAATTC-GATTCTTAGTACTGTCCCCC
(L7 sequence to right of hyphen)
Reverse (L7Neo-5'B): ATCCCAGAACCCCAGCACTCCTGCCAC
PCR produces a 600 bp-fragment.
5.3.4 Total RNA extraction
Dorsal root ganglion, olfactory epithelia, and cochlea were obtained from
20 rats at 1 months of age along with two cerebella and two eyeballs (Pelfreeze,
Rogers, AR). Total RNA was isolated from freshly dissected tissues using TRIzol reagent (15596-026, Invitrogen, Carlsbad, CA) following manufacturer’s 169 instructions. Basically, half of the cerebellum was homogenized using a glass
Teflon homogenizer in 1 ml TRIzol reagent. Proteins and lipids were separated from the mixture by addition of chloroform. RNA was precipitated by addition of isopropanol. Precipitated RNA was air dried and dissolved in RNase-free water.
Total amount of RNA was quantitated after measuring optical density of the sample. Samples were stored at -80°C.
5.3.5 DNase treatment
DNA-Free kit (AM1906, Ambion, Austin, TX) was used to eliminate DNA contamination in RN A extracts. 10 µg total RNA extract was incubated with 2 units of DNase at 37°C for 30 min. 0.1 volume of DNase inactivation reagent was added, and the sample was incubated at room temperature, mixing occasionally.
Inactivation reagent was precipitated by centrifuging, and RNA in the supernatant was transferred to a fresh tube. Samples were stored at -80°C.
5.3.6 Reverse Transcription (RT)
RETROscript kit (AM1710, Ambion, Austin, TX) was used to produce cDNA from total RNA extracted from mouse brain and cerebellum tissues. ~1 µg of DNA-free RNA was incubated with Oligo(dT) primer and nuclease free water at 70°C for 3 min to heat-denature the RNA. After putting the tube on ice, remaining RT components were added: RT buffer, dNTP mix, RNase inhibitor and MMLV-RT. The mixture was incubated at 42°C for one hour, and at 92°C for
10 min to inactivate the RT. Samples were stored at -20°C.
170 5.3.7 Polymerase chain reaction (PCR)
GeneAmp PCR System 9600 (Waltham, MA) was used to carry out PCRs.
PCR cycle program was as follows:
• 1st step: 2 min at 94°C
• 2nd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (30 cycles)
• 3rd step: 10 min at 72°C
Primer sequences were as follows:
L7:
Forward (L7-YS-1-F): AGGCTTCTTCAACCTGCAGA
Reverse (L7-YS-1-R): CGTTTCTGCATTCCATCCTT
RT-PCR produces a 234 bp fragment.
β-actin:
Forward (bActin-2a-F): GCATGTGCAAAGCCGGCTTC
Reverse (bActin-2-R): GGGGTGTTGAAGGTCTCAAA
RT-PCR produces a 346 bp fragment.
After PCR samples were run on a 1.5% agarose gel, and visualized by
Gel Doc 2000 gel documentation system (BioRad, Hercules, CA).
5.3.8 Real-Time PCR
The iCycler iQ Real-time PCR Detection System was used along with the iQ SYBR Green Supermix (1708882) following manufacturer’s instructions
(BioRad, Hercules, CA). The 96 well-format was used. Before the Real-time
171 PCR, all primers were observed to produce a PCR product that resolved as a single band with no primer–dimers on agarose gels.
The Real-time PCR cycle program was as follows:
• Automatically inserted steps to collect well factors: 30 sec at 95°C (2
cycles)
• 1st step: 15 min at 95°C
• 2nd step: 15 sec at 94°C
• 3rd step: 30 sec at 94°C, 30 sec at 55°C, and 30 sec at 72°C (60 cycles)
• 4th step: 10 sec starting at 95°C and decreasing temperature for 0.5°C at
each cycle (100 cycles: this is to draw the melt curve)
Same primers used in RT-PCR were used in Real-time PCR.
5.3.8.1 Determination of PCR efficiency for each primer set
To be able to compare the threshold cycle values from two different PCRs, it is important to use primer sets with similar efficiencies (95-100%). For this purpose, standard curve function of iCycler iQ Real-time PCR Detection System was used.
Vectors carrying each cDNA of interest (for example L7 and β-actin) were obtained. For L7, pL7hom1 was used. β-actin cDNA was obtained by RT-PCR, resolved in 1.5% gel, gel-extracted and cloned into pCR/8/GW/TOPO vector using TOPO TA cloning kit (Invitrogen, Carlsbad, CA). Plasmid was extracted from bacterial culture, and DNA concentration was calculated. Serial dilutions of plasmid containing the cDNA of interest were prepared: 10, 1, 0.1, 0.01, and
172 0.001 ng/µl. Each cDNA dilution series were used in Real-time PCR with its specific primers. Real-time PCR was run using three wells for each dilution.
Standard curve function was selected at the beginning of the run. At the end of the cycle, the software provided the standard curve, and automatically calculated the efficiency of the primer set used in that particular reaction. Primers with efficiencies between 95 and 100% were then used in the Real-time PCR with experimental cDNA samples.
5.3.8.2 Quantification of expression levels by Real-Time PCR
For quantification, the comparative CT method was used (Table 3 of User
Bulletin #2 of the ABI Prism 7700 Sequence Detection System). Briefly, the
average threshold cycle (CT) from three reactions of each RT sample was determined. The ΔCT was calculated by subtracting the average actin CT value of
each sample from the average L7 (or the gene being investigated) CT value of
the same sample. The ΔΔCT value was determined by subtracting the ΔCT value of the sample with higher CT value from the ΔCT value of the other sample. The
level of gene of interest relative to the control gene (in our case, β-actin) was
determined by the equation 2−ΔΔCT.
The standard deviation (SD) for each sample was derived from the
square-root of the summed squares of the standard deviations of the average L7
and β-actin CT values, SD-L7, and SD-actin. Data are presented as ± SEM.
173 5.3.9 Behavioral tests
All behavioral tests, except the accelerating rotarod test, were done by
Stephanie Bowers-Kidder in Dr. Randy Nelson’s Lab.
5.3.9.1 Latency to Move
[1-3 hr. after lights-out, 30 min. acclimation period] A mouse was placed in the middle of a drawn circle with a diameter of 10 cm. (the average body length of the group). Latency for the animal to place all four paws outside the circle (or move one body length) was recorded (60 sec. maximum). The test was performed twice, with an inter-trial interval of 10-20 sec.
5.3.9.2 Open Field/Locomotor
[1-4 hrs. after lights-out, 30 min. acclimation period] Locomotor activity was assessed using the Flex-Field Photobeam Activity System (San Diego
Instruments, Inc., San Diego, CA). The apparatus consists of a 36 x 36 cm.
Plexiglas arena inside a ventilated, darkened cabinet. A frame with 2 rows of infrared photocell detector beams surrounds the arena; interruptions in the infrared beams made by the animals’ horizontal and vertical movements are tallied via computer. Activity (beam breaks) was recorded in a 30 min. session; data were analyzed for total activity, time spent in the periphery vs. the center of the arena, and number of rears. The arena was wiped with 70% ethanol and lined with fresh corncob bedding between each animal.
174 5.3.9.3 Grip Strength
[1-3 hrs. after lights-out, 30 min. acclimation] Balance and grip strength
were measured using a standard wire cage lid. Masking tape was placed around
the perimeter of the lid to keep the mouse on the bottom and upside down. The
test is performed by placing the mouse on top of the wire cage lid, gently shaking
the lid to cause the mouse to grip the wires, and slowly turning it upside down.
The lid was held approximately 20 cm. above the home cage to keep the animal
from easily jumping, but not causing harm should the animal fall. Latency-to-fall
off the cage lid was quantified using a stopwatch (60 sec. maximum); the test
was performed twice, with an inter-trial interval of 10-20 sec.
5.3.9.4 Beam Walk
[1-2 hrs. after lights-out; 30 min. acclimation] A mouse with significant
motor deficits will typically fall or move more slowly than a normal mouse while
beam walking. A wooden dowel rod (2 cm. diameter, 60 cm. above the tabletop)
is divided into 7 cm. sections (lines drawn on the underside and side of the rod,
not visible to the mouse). The mouse is placed in the center of the beam, and the number of beams (lines) crossed, latency-to-reach platform, and, when necessary, the latency-to-fall was recorded (120 sec. maximum). The test was performed twice, with an inter-trial interval of 60 sec., and the beam was wiped with 70% ethanol between animals.
175 5.3.9.5 Light/Dark Preference
[30 min. acclimation; 10 A.M.] The testing apparatus consisted of a transparent box, 1.35 m. long, 60 cm. wide, and 15 cm. high. A foam board divider separated the box into two chambers, and had a 10 x 10 cm. square cut in the center to allow passage from one side to the other. One chamber was completely sealed and lined with black foam board, and the other was unlined, uncovered and illuminated by a 60 W desk lamp. For the 5 min. test, the mouse was placed in the center of the lit chamber, facing the passage to the other chamber. Latency-to-move, latency-to-enter the dark chamber, total transitions and total time spent in the dark was recorded and computed. Surfaces were wiped with 70% ethanol between animals.
5.3.9.6 Hotplate
[30-60 min. after lights-out; 30 min. room acclimation] Animals were tested on a hotplate (Model 39, IITC Life Science, Inc, Woodland Hills, CA) heated to
53°C. An acrylic cylinder was placed on the surface, and mice were placed in the cylinder to keep them contained. Latency for the mouse to lift, lick, or shake either front paw was recorded. The surface of the apparatus was wiped with 70% ethanol between animals. Mice were tested as described every 7-9 days for 5
(22406 animal set) or 4 (1105 animal set) consecutive weeks both to increase trial numbers and to look for evidence of habituation.
176 5.3.9.7 Rotating Rod
[1-4 hrs. after lights-out; 30 min. room acclimation] Ability of an animal to
remain on a constantly rotating rod (ENV-576M, Med Associates, St. Albans,
Vermont) was tested. Three different trials were given at each of three increasing speeds (20, 28, and 32 RPM). There was a 60 sec. interval between trials, and
45-60 min. between speed changes. The apparatus was wiped with 70% ethanol
between animals, and all speeds were tested on the same day. For the
accelerating rod we used a Rotamex-4/8 apparatus (Columbus Instruments,
Columbus, OH). The rod was accelerated from 4-50 rpm in 5 min., the maximum
duration of the test. Analysis was performed over 7 consecutive days with three
trials per day. All animals were 5-6 wks. old at the time of testing.
5.3.9.8 Acoustic Startle Response (ASR)
The chamber (SR-LAB, San Diego Instruments, San Diego, CA) consists
of a nonrestrictive Plexiglas cylinder resting on a platform inside a ventilated
cabinet. A high-frequency loudspeaker inside the cabinet produces both a
continuous background noise of 65dB and the various acoustic stimuli. Vibrations of the Plexiglas cylinder caused by the whole-body startle response of the animal are transduced into analog signals by a piezoelectric unit attached to the bottom of the platform. The signals are digitized and stored by computer. The inter-trial intervals for each test were randomly selected prior to testing, so that the order of
interval presentation was the same for each animal. Other than assigned “no
pulse” entries at the beginning and end of the testing session, trial presentation
177 was randomized prior to testing and then programmed so that the animals would, again, be tested similarly.
5.3.9.9 PPI
[1-5hrs. after lights-out; 10 min. acclimation inside chamber] Sixty-five
readings are taken at 1 msec. intervals, starting at stimulus onset, and the
maximum velocity of the response is used to determine the acoustic startle
response. All pre-pulse inhibition test sessions consist of startle trials (pulse
only), pre-pulse trials (pre-pulse + pulse), and no-stimulus (no stim). The pulse
only trials consist of a 40 msec., 120dB pulse of broadband noise. Pre-pulse
inhibition is measured by pre-pulse + pulse trials that consist of a 20 msec. noise
pre-pulse, a 100 msec. delay, then a 40 msec., 120dB pulse. The acoustic pre- pulse intensities are 69, 73, and 81 dB. The no-stim trials consist of background noise only. The test session begins and ends with five presentations of the pulse only trial; in between, each acoustic or no-stim trial type is presented 10 times in pseudorandom order as described above. The pulse only trials are used to determine the average Vmax, or the startle amplitude. The percent PPI is calculated relative to the Vmax.
5.3.9.10 ASR Habituation
[1-6hrs. after lights-out; 3 min. acclimation in chamber] The testing paradigm consisted of 60 trials of a 120 dB pulse and 10 trials of no pulse. The trials were separated by pseudorandom inter-trial intervals that varied from 15s to 25s.
178 5.3.9.11 Acoustic Threshold
[1-5hrs. after lights-out; 5 min. acclimation in chamber] Testing consisted
of 10 trials each, presented in pseudorandom order, of no pulse, 70, 80, 90, 100,
110 and 120 dB pulses. By curve fitting (P < 0.015, R > 0.99) we could determine the Rmax (maximal response, or Vmaxmax) as well as the dB50 (sound level at which 1/2 the maximal response was obtained). Comparison of the 95% confidence limits of the dB50 values shows that these values are indeed not significantly different with respect to genotype or sex (L7+/+ males, dB50 =
100.86 - 104.51; L7+/- males, 100.07 – 107.24; L7-/- males, = 100.79 - 106.36;
L7+/+ females, dB50 = 104.89 - 107.89; L7+/- females, = 100.98 - 109.38; L7-/- females, = 103.58 - 110.91). However, there is a clear sex difference in the maximal response as the males overall reached a higher Rmax compared to females. Furthermore, the L7-/- males reached a higher Rmax value than males
of other genotypes, and analysis of 95% confidence limits suggests that this is
significant (tested by the GT2 Method cf Sokal and Rohlf, 1994, p. 501) (L7+/+ males, Rmax = 220.03 - 255.60; L7+/- males, Rmax = 196.53 - 254.09; L7-/- males, Rmax = 233.45 - 296.82). The greater Rmax value of L7-/- males is also
consistent with the observations on acoustic startle amplitude (Vmax) and
habituation where a genotype*sex effect was also observed. In spite of this
difference in Rmax, dB50 values were not significantly different in any
genotype*sex combination. Therefore hearing is not impaired in the mutant.
179 5.3.9.12 Elevated Plus Maze
[1-4hrs. after lights-out; 30 min. room acclimation] The elevated plus maze
consists of two open and two closed 5-cm. wide arms in a plus-sign configuration
1 m. off the floor. The closed arms are enclosed by 15 cm. tall black Plexiglas on
either side. Each mouse was placed in the center of the apparatus, facing an open arm, and allowed to move freely on the maze for a 5-min. videotaped testing period. Animals were considered to have entered an arm only when all
four paws crossed onto the arm. All surfaces were wiped with 70% alcohol
between animals. Videotapes were later scored with The Observer software
(Version 5, Exeter Software, Setauket, NY) for the latency-to-enter any arm,
duration and percentage of time spent in open arms, and duration and
percentage of time spent in closed arms.
180 5.4 RESULTS
All behavioral tests, except the accelerating rotarod test, were done by
Stephanie Bowers-Kidder in Dr. Randy Nelson’s Lab.
5.4.1 Initial analysis of L7KO mice
We obtained the first of the previously described lines of L7KO mice
(Vassileva et al., 1997), which were produced in C57BL/6 X 129 background.
Since strain of mice can greatly affect the behavioral phenotype (Crawley, 1999), we decided to breed L7KO mice in a pure background. To this end, we backcrossed them more than eight generations in C57BL/6 strain. RT-PCR and
Western blotting were carried out to confirm the absence of L7 mRNA and protein, respectively, in L7-/- animals (Figure 5.1). The genotypes of the mice
from each litter were identified by PCR of tail biopsy. Before the behavioral
analysis, all animals were given a general physical assessment which included
body weight (before and after the testing period), body length, touch reflex, eye
appearance, muscle tone, and cage behaviors. By these criteria, L7-/- mice appeared normal when compared to their WT littermates.
Figure 5.1 Western blotting analysis of L7+/+, L7+/-, L7-/- cerebellums. Reduced expression of L7 protein is observed in L7+/-, and no expression in L7-/-. Calbindin (CaBP) expression does not change.
181 5.4.2 Behavioral analysis of L7KO mice
After the preliminary observations, mice were subjected to several
behavioral tests covering the following modalities: Locomotor activity, anxiety,
motor coordination/balance and sensory responsiveness.
5.4.2.1 L7KO mice have normal locomotor activity and anxiety
behaviors
Locomotor activity was tested using the open-field and latency-to-move
tests. 10 L7+/+, 18 L7+/-, and 6 L7-/- males; 8 L7+/+, 15 L7+/-, and 12 L7-/- females were used in each test. Both locomotor tests showed that L7-/- mice were as
active as wild-types (Figure 5.2A-B). The only significant difference was for sex;
females overall engaged in more exploratory activity and had shorter latency to
move.
Anxiety activity was tested using elevated plus maze and light-dark
preference test. In the former test, 6 L7+/+, 10 L7+/-, and 3 L7-/- males; 3 L7+/+, 11
L7+/-, and 2 L7-/- females were used, while 10 L7+/+, 18 L7+/-, and 6 L7-/- males; 8
L7+/+, 15 L7+/-, and 12 L7-/- females were used in the latter. Similar to the
locomotor tests, there was no significant genotype effect on the anxiety tests.
However, females of all genotypes had a shorter latency to move into the dark in the light-dark preference tests (Figure 5.2C).
These results suggest that L7-/- mice have normal locomotor activity and
anxiety behaviors.
182
Figure 5.2 Locomotor activity and anxiety behaviors were compared. A) Open field locomotor test shows a sex effect in both total and peripheral activity, but no significant effect of genotype. Females overall spend more time in exploration. B) Latency-to-move test shows a clear-cut sex effect, but no significant genotype effect. Females move more rapidly out of the circle. C) L-D preference test shows a sex effect in latency-to-enter dark, i.e., females move more rapidly into the dark chamber. However, there is no significant genotype effect.
183 5.4.2.2 L7KO mice show improved maximum performance on a motor
learning test
Since cerebellum is important in motor learning and coordination of
movements, we next tested animals in motor learning tests. A previous study
detected no motor deficits in L7-/- mice using a constant velocity rotarod test
(Mohn et al., 1997). This previous study was performed using a different mutant
line and also in a hybrid background strain. Moreover, this study did not address
the issue of genotype*sex interaction. Here we tested mice using constant
velocity rotarod test and accelerating rotarod test.
The constant velocity rotarod test was carried out at three different speeds
(20, 28 and 32 rpm) and for three trials at each speed. 11 L7+/+, 16 L7+/-, and 10
L7-/- males; 12 L7+/+, 12 L7+/-, and 16 L7-/- females were used in this test. Our
data concur with the earlier study that there is no detectable overall effect of
genotype on latency to fall (Figure 5.3B; 3-way ANOVA, genotype,
F(2,675)=1.043, p=0.353). However, there are significant sex (F(1,675)=42.793, p<0.001) and rotarod speed (F(2,675)=7.751, p<0.001) effects, and sex*genotype interaction revealed by ANOVA (F(2,675)=8.062, p<0.001). L7-/- males show an increase in performance compared to the WTs while this increase depends on the rotarod speed. L7-/- females, on the other hand, show a modest decrease compared to the WTs at all speeds. This subtle but oppositely-oriented effect between the sexes would explain why no genotype interaction is observed
in our study and possibly in the previous study.
184
Figure 5.3 Motor coordination and motor learning tests were performed. L7 mutant mice of both sexes show a significant increase in maximal performance on a motor learning test. A) Grip strength was measured using the wire hang test. No significant genotypic differences were observed. B) Constant velocity rotarod shows that there is no significant overall genotype effect on motor coordination. However, there is a significant sex effect, and there is also a significant genotype*sex interaction. Male L7-/- mice tend to stay on the rod longer than wild-type males, and female L7-/- mice tend to fall off faster than female wild-types. C) An increased maximal level of performance is observed in L7-/- mice of both sexes after repeated daily training on an accelerating rotarod. Data are plotted for the late generations animal set only.
185 The accelerating rotarod test, which test both motor coordination and
motor learning, was carried out on a rod with acceleration from 4 to 50 rpm in 5
minutes, for seven days with three trials at each day. We obtained two data sets
in this test. In the first set, we used mice backcrossed 4-6 obtained two data sets
in this test. In the first set, we used mice backcrossed 4-6 generations in
C57BL/6 (early generations). 6 L7+/+, 14 L7+/-, and 5 L7-/- males; 14 L7+/+, 14 L7+/-
, and 13 L7-/- females were used in this data test. The second data set included
mice backcrossed 8-10 times in C57BL/6 strain (late generations). 17 L7+/+, 19
L7+/-, and 15 L7-/- males; 13 L7+/+, 23 L7+/-, and 8 L7-/- females were used in this
data test. In late generations, both sexes improved over time, and there was a
strong genotypic difference for both males and females (Figure 5.3C). L7-/- mice of both sexes reached a maximum level of performance that was about 30% higher than that of L7+/+ and L7+/- mice. A similar effect was observed in the
“early generations” data set (data not shown). Three-way ANOVA for the late
generations animal set indicates that there are significant effects of genotype and
day, but not of sex (for genotype, F(2,1695)=45.93, p<0.001; for day,
F(6,1695)=91.06, p<0.001; for sex, F(1,1695)=0.037, p=0.849. There are also significant interactions for genotype*sex (F(2,1695)=5.141, p=0.006) and genotype*day (F(12,1695)=2.245, p=0.008) but no three-way interactions.
Comparison of the curves for L7-/- males and females reveals that the shapes are slightly different. The mutant males appear to approach the maximum time on rod more gradually than the mutant females and ultimately reach a
higher value. To show that the improvements observed in this test was not due to 186 increased strength of the L7-/- mice, mice were subjected to grip strength test,
and no significant differences were observed (Figure 5.3A). Moreover, the
improvement is probably not due to increased motor coordination because L7-/- mice were not significantly different on the constant velocity rotarod test.
Nevertheless, both rotarod paradigms revealed a significant genotype*sex interaction, and in both cases L7-/- males performed better relative to the same-
sex wild types than did L7-/- females.
5.4.2.3 L7KO mice show altered sensory responsiveness
To show if L7-/- mice have any changes in sensory responsiveness, we
carried out several tests including acoustic startle response, pre-pulse inhibition,
acoustic startle habituation, and hot plate test.
The acoustic startle response (ASR) test was carried out to determine
whether L7-/- mice were more or less sensitive to loud noises compared to wild-
-/- types. L7 males showed slightly increased startle amplitude (Vmax) relative to
other genotypes, while L7-/- females show the opposite trend (Figure 5.4A). In
addition, time required to reach the Vmax was calculated and compared (Figure
5.4B). However, ANOVA revealed no significant genotypic effect on Vmax
(F(2,764)=0.342, p=0.710). Nevertheless, sex did have a significant influence
(F(1,764)=39.373, p<0.001), and there was a significant genotype*sex interaction
(F(2,764)=3.719, p=0.025). Post hoc individual comparison reveals that the L7-/- males have a significantly greater startle amplitude than L7+/- males (P<0.05),
and comparison with L7+/+ males reveals a similar trend (p=0.168). Consistent
187 with the ANOVA results for sex, male wild-types have a significantly greater Vmax
(1.4-fold) than female wild-types (p<0.001) and the sex difference increases (to
-/- 1.8-fold) in L7 (p<0.001). An inverse relationship between the startle amplitude
(Vmax) and time (in msec) required to reach Vmax (Tmax) was observed as expected, although this genotypic differences in Tmax did not reach significance
(F(2,764)=0.640, p=0.527), while there was a significant sex effect
(F(1,764)=6.379, p=0.012). There also was no significant genotype*sex
interaction for Tmax (F(2,764)=1.643, p=0.194), although the data trend is
consistent with increased sensitivity in male mutants and decreased sensitivity in
female mutants, as observed for Vmax.
Next, pre-pulse inhibition was examined. In this test a pre-pulse is applied,
which generally causes a reduced startle response in mice. Pre-pulse inhibition,
PPI, % inhibition of the startle amplitude in the presence of a pre-pulse, is widely
considered to be a CNS mechanism of sensorimotor gating (Crawley, 1999;
Fendt et al., 2001; Koch, 1999). 11 L7+/+, 16 L7 +/-, and 10 L7-/- males; 12 L7+/+, 12
L7+/-, and 16 L7-/- females were used. In this test, there is a small increase of
%PPI in L7-/- males relative to wild-types at all pre-pulse levels, but little change
in the females (Figure 5.4C). ANOVA (with genotype, sex, and loudness of the
pre-pulse as independent variables) revealed a significant effect of the genotype
(F(2,1952)=3.746, p=0.024), no effect of sex (F(1,1952)=0.683, p=0.409), and a
significant effect of the pre-pulse loudness (F(2,1952)=39.509, p<0.001). There
was a significant interaction between genotype and sex (F(2,1952)=8.132,
p<0.001), which reflects the fact that in males, L7 gene inactivation tends to 188 increase PPI at all prepulse levels, whereas in females, it has no such effect.
There were no other 2- or 3-way interactions.
We also examined the effect of the pre-pulse on Tmax. In parallel with the decrease in Vmax in response to a pre-pulse, there should be an increase in Tmax.
ANOVA revealed that the effect of genotype alone was not significant
(F(2,1952)=1.704, p=0.182). There was a significant effect of sex
(F(1,1952)=13.748, p<0.001), but the effect of pre-pulse loudness was not
significant (F(2,1952)=2.521, p=0.081). There was a significant interaction
between genotype and sex (F(2,1952)=15.164, p<0.001), which reflects the fact
that, in males, L7 gene inactivation tends to increase the change of Tmax,
whereas in females, it decreases this change (Figure 5.4D). In addition, there
was interaction between genotype and pre-pulse loudness (F(2,1952)=2.544,
p=0.038), but not between sex and pre-pulse loudness (F(2,1952)=2.243,
p=0.106).
189
Figure 5.4. Assessment of the sensory phenotype of L7 mutants by the acoustic startle response and hotplate tests. Sexually dimorphic effects were observed on sensory responses in L7 mutants. A) The pulse-only trials from the PPI testing paradigm were used to measure the startle amplitude, or Vmax, of the startle response. B) The time to Vmax, or Tmax, was determined from the same pulse-only trials from the PPI testing paradigm. C) PPI of the ASR. D) %-change in Tmax during PPI. E) Short-term habituation of the ASR. F) Short-term habituation of ASR viewed by linear regression of all trials. G) Acoustic threshold test. H) Hotplate test. Results were explained in the text. (The fill key in panel A pertains to all panels.)
190
191 We also examined the habituation of the ASR. Previous studies in rats
have indicated a role of the cerebellum in the habituation of the ASR (Koch,
1999; Leaton and Supple, 1991; Maschke et al., 2000). In this test, sixty trials
were presented (120 dB sound pulse), and the extend to which there was a
decreased response in the last thirty trials compared to the first thirty was
determined. 7 L7+/+, 8 L7+/-, and 7 L7-/- males; 7 L7+/+, 8 L7+/-, and 6 L7-/- females were used. All females showed habituation, while only L7-/- males showed
habituation for the males (Figure 5.4E). By ANOVA, genotype had a significant
effect on Vmax (F(2,2568)=3.835, p=0.022), as did sex (F(1,2568)=15.864,
p<0.001) and the trial set (1-30 versus 31-60) (F(1,2568)=33.130, p<0.001).
However, the largest effect in this assay is not on habituation per se, but on the
amplitude of the startle response in L7-/- males. This result confirms the
increased startle amplitude of L7-/- males that was observed in the ASR test
(Figure 5.4A). In summary, L7-/- males are more sensitive to loud noise than all
other genotype-sex combination, and they are more sensitive to the inhibitory
effects of the pre-pulse on startle amplitude than are wild-type males.
In order to exclude the possibility that any of the observed differences is
due to impairment of hearing, we performed an acoustical threshold test. 7 L7+/+,
8 L7+/-, and 7 L7-/- males; 7 L7+/+, 8 L7+/-, and 6 L7-/- females were used in this
test. As it can be seen in (Figure 5.4G), the curve upswing begins at the same
sound level for all genotype-sex combinations. In addition, L7-/- males reach a
higher maximal response (Rmax) than males of other genotypes and females. In
spite of this difference in Rmax, the sound level at which ½ the maximal response 192 was obtained was not significantly different in any genotype-sex combination.
Therefore hearing is not impaired in the L7-/- mice.
Finally, mice were subjected to hot plate test to measure their sensitivity to heat. In this test, 11 L7+/+, 12 L7+/-, and 16 L7-/- males; 12 L7+/+, 12 L7+/-, and 16
L7-/- females were used. We found that both sexes are sensitized by the loss of
the L7 gene product, but male L7-/- mice were significantly more sensitive to heat
than any other genotype sex combination (Figure 5.4H). ANOVA showed
significant differences for both sex (F(1,342)=20.296, p<0.001) and genotype
(F(2,342)=6.236, p=0.002). The genotype*sex interaction was not significant
(F(2,342)=2.596, p=0.078). Using multiple post hoc comparisons male L7-/- mice
were significantly different from male L7+/+ (P<0.05) and L7+/- (P<0.01). Female
L7-/- and L7+/- were significantly different from female L7+/+ (p<0.01 and p<0.05,
respectively). Male versus female L7+/+ were significantly different (males more
sensitive, p<0.001) and male versus female L7-/- were significantly different
(males more sensitive, p<0.01). L7+/- males and females were not significantly
different. In summary, these results suggest an altered sensory responsiveness
in L7-/- mice, especially in males.
To show that these phenotypes are due to disturbed functioning of the
cerebellum and Purkinje cells, where L7 is specifically expressed,
electrophysiological tests carried out in Dr. Mike Zhu’s Lab by Emilia Iscru and
Jinbin Tian suggest sexually dimorphic changes in the intrinsic spontaneous
firing patterns of Purkinje cells. This supports the conclusion that sexually
193 dimorphic behavioral changes in L7KO mice are actually a result of changes in the Purkinje cells, thus the cerebellum.
5.4.3 L7 is specifically expressed in cerebellum and retina, but not in sensory organs
Based on the previous observations, L7 expression is restricted to the retina and cerebellum. This restricted expression pattern suggests that the sensory changes that have been observed in L7-/- mice are truly part of a cerebellar phenotype. In order to rule out expression of L7 in sensory organs that might contribute to the phenotype of the global null mutants, we have performed
RT-PCR and real-time RT-PCR using RNA extracted from wild-type rat dorsal root ganglia, cochlea, and olfactory epithelium. In addition, we extracted RNA from mouse cerebellum, eye, brain and liver as positive and negative controls.
Among the sensory tissues used, dorsal root ganglia and cochlea were specifically chosen based on the behavioral phenotypes observed using the hot plate and acoustic startle tests. Expression of L7 was clearly detectable in both cerebellum and retina, and expression in the retina was 6-fold less than that in cerebellum (Figure 5.5). However, no L7 mRNA was detected in any sensory tissues that were tested. Actin mRNA, in contrast, was clearly detected and at comparable levels in all tissues. By this assay we can conclude that any low-level
L7 expression in these other tissues would have a maximum value at least 200- fold less than that in the retina and more than 10,000-fold less than that in the cerebellum. In combination with the experiments described above, this suggests
194 that it is unlikely that the phenotypes we observed are due to loss of L7 expression not previously observed in sensory tissues.
195
Figure 5.5 RT-PCR and Real-time PCR analysis of L7 expression in sensory organs of wild-types. A) RT-PCR was performed (PCR, 35 cycles) and the reaction products analyzed by agarose gel electrophoresis. Primers for L7 revealed product only in retina and cerebellum. β-actin was detected in all samples. B) RealTime PCR was performed on the same RNA sample as in A. Again, product was only detected in cerebellum and retina. M = mouse; R = rat; cbm = cerebellum; Br = brain; DRG = dorsal root ganglion; Olf Epi = olfactory epithelium; Coch = cochlea; Liv = liver; +RT and –RT = with and without reverse transcriptase.
196 5.5 DISCUSSION
Previous analyses of L7KO mice reported no obvious motor deficits, gait, or cage behaviors indicative of gross cerebellar dysfunction in L7 null mutants
(Mohn et al., 1997; Vassileva et al., 1997). Our analysis, on the other hand, indicates an improved motor performance of the mutants compared to their wild- type littermates. While both sexes show this improvement in motor learning, the increase relative to wild-types is greatest in males. We suspect that this is not due to improved cerebellar function, but rather a secondary effect of defective sensory processing: the null mutants show a significant change in sensorimotor responsiveness that is sexually dimorphic, with the male mutants showing a general heightened sensitivity to heat and sound. These sensory defects are cerebellum-dependent, since RT-PCR assays show no traces of L7 mRNA in wild-type sensory organs, including dorsal root ganglion and cochlea, which are the sensory organs involved in each of the behavioral tests, or in any other brain regions. In order to show physiological relevance of the observed behavioral changes to the Purkinje cells functioning, we examined the firing properties of
Purkinje cells in these mutants using extracellular recordings in slices. We found a decreased burst firing rate in mutant males, and genotypic effects in other parameters that were not sex-specific, or alternatively were enhanced in females.
As no gross motor deficits were observed in these mice, we conclude that the contribution of L7 to Purkinje cell physiology affects mainly sensory integration and timing of motor output. We suggest that L7 protein is not essential for basic
197 cerebellum function, and its function is not essential for Purkinje cell survival.
However, our results suggest subtle behavioral changes in L7 null mutants, and
they indicate that the function of L7 protein is more sensory than it is motor.
Cerebellum has long been believed to primarily function in the generation,
spatial accuracy and temporal coordination of movements, formation of balance
and muscle tone, motor learning, and fine-tuning the motor patterns. However, in
recent years, several different studies have provided evidence for the
contributions of the cerebellum to cognitive and sensory processing (Fendt et al.,
2001; Gao et al., 1996; Geyer et al., 1990; Kim et al., 1994; Koch, 1999;
Rapoport et al., 2000). Moreover, a number of studies have linked cerebellum to
sensorimotor gating, which is a mechanism the central nervous system uses to
filter out excessive sensory information that would otherwise tend to disrupt the
cognitive and integrative function of the brain. Pre-pulse inhibition (PPI) of the acoustic startle response (ASR) is one operational measure of sensorimotor gating (Fendt et al., 2001; Geyer et al., 1990; Koch, 1999). Cerebellum is not classically considered to be part of the ASR brain circuitry, which involves cochlear nuclei, caudal pontine reticular nucleus, and inter- and motorneurons.
Nevertheless, long-term habituation of the ASR is suggested to be mediated by the cerebellum in mice and rats (Leaton and Supple, 1991; Maschke et al.,
2000). A study similar to ours showed that genetic inactivation of a Purkinje cell- specific isoform of NMDA receptor, GluRδ2 in mice resulted in enhanced PPI
(Takeuchi et al., 2001). Moreover, imaging studies also suggested that the
cerebellum plays a role in heat-induced pain and hyperglasia (Helmchen et al., 198 2003; Zambreanu et al., 2005). Studies of olfactory and tactile functions of the
cerebellum also suggest that the cerebellum modulates motor systems to
optimize sampling of sensory information (Mainland et al., 2005). The results
presented here are consistent with this view that the cerebellum likely performs
functions that are primarily sensory, or different in some other way from the
traditional roles typically attributed to the cerebellum.
It was previously suggested that L7 regulates the degree of Gi/o-mediated
inhibition of Cav2.1 (P/Q-type) voltage-dependent Ca2+ channels in a dose
dependent manner (Kinoshita-Kawada et al., 2004). In this report we analyzed
changes in the spontaneous firing properties in the Purkinje cells. It is suggested
that spontaneous bursting is dependent on the dendritic Ca2+ spikes that are
eliminated by blockade of P/Q type Ca2+ channels (Womack and Khodakhah,
2004). Thus our data is consistent with a role for L7 in modulating the intrinsic firing properties of Purkinje cells via P/Q channel effector function.
We observed sexual dimorphism in our analysis, which supports increasing evidence of cerebellum-related sex differences. For example, as we have provided evidence in Chapter 3, a key transcriptional modulator of L7 gene
expression is RORα (Serinagaoglu et al., 2007). A natural mouse mutant of
RORα is known as staggerer (Gold et al., 2003; Hamilton et al., 1996). In
staggerer heterozygous mice, cerebellar Purkinje cell loss during aging appears
early in males when compared to females (Doulazmi et al., 1999). While we do
not know whether there is a sex-specific effect on L7 gene transcription, it is clear
from our current work that loss of L7 protein has a sexually dimorphic influence 199 on behavior and cerebellar physiology, which could help us explain some of the
dimorphic effects observed in heterozygous staggerer mice.
Recently, we have observed that the human PCP2 and CACNA1A genes
are separated by only 5 Mb and are both localized on human 19p13.2
(unpublished observations with in silico analysis). CACNA1A gene encodes the
α1A pore-forming subunit of the P/Q-type Ca2+ channel that we hypothesize is a
downstream effector of L7. Human 19p13.2 has been identified in a number of
studies as a susceptibility locus for autism (Buxbaum et al., 2004; Liu et al.,
2001; McCauley et al., 2005; Philippe et al., 1999; Shao et al., 2002). While the cerebellum has been implicated in many studies of the neurological origins of autism, little is known about its specific role in the disorder (Allen et al., 2004;
Pierce and Courchesne, 2001; Weber et al., 2000). Increased startle amplitude
and heightened sensitivity to a variety of sensory stimuli are characteristics of
disorder (Frankland et al., 2004; McAlonan et al., 2002). In addition, the disorder
shows a roughly 3:1 prevalence in males (Baron-Cohen et al., 2005). While the
phenotype of L7 mutant mice does not prove any relationship to autism, certain
features of the phenotype are at least consistent with its candidacy as a
susceptibility gene. These features would include the increased sensitivity of
mutants to loud sound and to heat, which in turn are stimuli to which wild-type
males are naturally more sensitive than females, as we have shown.
In conclusion, we provide evidence consistent with a sexually dimorphic
sensorimotor gating function of the cerebellum and L7.
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