CELLULAR AND MOLECULAR STUDIES OF
ALSIN/ALS2
Elizabeth Lilian Tudor BSc Hons
Thesis submitted in fulfilment of the degree of Doctor of Philosophy,
University of London (Kings College, Institute of Psychiatry)
September 2005
ABSTRACT
Mutations in alsin/ALS2 have been identified that cause several forms of motor neuron disease including a rare recessive form of juvenile ALS (ALS2), juvenile primary lateral sclerosis (jPLS) and infantile onset ascending hereditary spastic paraplegia (IAHSP).
Although the function of the ALS2 protein is unknown, ALS2 contains three functional domains that show sequence homology to guanine nucleotide exchange factors (GEFs) for the Ras superfamily of GTPases. The first domain shows most similarity to a Ran
GEF (RCC1-like domain), the second to Rho family GEFs (DH/PH domain) and the third to a Rab GEF (VPS9 domain).
In these studies, a polyclonal antibody to ALS2 was generated and used to investigate
ALS2 localisation in the mammalian central nervous system. ALS2 was present in several neuronal populations in the brain and in motor neurons of the spinal cord. Since the DH/PH domain of ALS2 shows close homology to Rho family GEFs, in vivo assays were performed to investigate whether ALS2 acts as a GEF for the Rho family members
RhoA, Rac1 and Cdc42. ALS2 activated Rac1 but not RhoA or Cdc42. A major downstream effector of Rac1 is p21 activated kinase 1 (PAK1) and in vitro kinase assays revealed that ALS2 also activates PAK1. Thus, ALS2 stimulates Rac1-PAK1 signalling.
Rac1 is known to regulate neurite outgrowth during development. Immunofluorescence studies revealed that ALS2 is present in growth cones of rat embryonic neurons where it co-localised with Rac1. Furthermore, overexpression of ALS2 stimulated neurite outgrowth.
2 Finally, the phosphorylation state of ALS2 was investigated. ALS2 was found to be a phosphoprotein in vivo and five serine/threonine phosphorylation sites were identified.
However, mutation of the identified phosphorylation sites did not alter the ability of
ALS2 to activate Rac1 or PAK1, or to stimulate neurite outgrowth. The findings reported in this thesis provide an insight into ALS2 function, which may contribute to our understanding of the molecular mechanisms involved in motor neuron disease.
3 ACKNOWLEDGEMENTS
I would like to thank my supervisors Chris Miller and Chris Shaw for their constant advice and support, everyone in the Miller Group for help in the lab (especially Steve
Ackerley and Kwok-Fai Lau), Steve Banner for help with the immunohistochemical studies presented in Chapter 3, Anja Schmidt for constructs and advice towards the work presented in Chapter 4, Mike Perkinton for his encouragement, advice and help with the phosphorylation studies presented in Chapter 6 and Helen Byers for the mass spectrometry studies in Chapter 6. This research was funded by a Jim Tew Memorial
Prize Studentship from the Motor Neurone Disease Association UK.
4 TABLE OF CONTENTS
ABSTRACT...... 2
ACKNOWLEDGEMENTS ...... 4
TABLE OF CONTENTS ...... 5
LIST OF FIGURES AND TABLES...... 13
PUBLICATIONS ARISING FROM THIS WORK ...... 15
ABBREVIATIONS...... 16
CHAPTER 1: INTRODUCTION...... 21
1.1 Motor Neuron Disease ...... 22
1.1.1 Clinical features and pathology of MND...... 23
1.1.2 Genetics of MND...... 25
1.1.3 Animal models of MND ...... 29
1.1.3.1 Spontaneous mutants...... 29
1.1.3.2 Targeted mutants ...... 31
1.1.4 Mechanisms of neurodegeneration in MND...... 34
1.1.4.1 Toxicity of intracellular aggregates...... 35
1.1.4.2 Oxidative stress ...... 39
1.1.4.3 Defects in axonal transport...... 42
5 1.1.4.4 Glutamatergic excitotoxicity ...... 46
1.1.4.5 Neuroinflammation and autoimmunity ...... 51
1.1.4.6 Apoptosis...... 52
1.1.4.7 Involvement of cell signalling pathways...... 54
1.2 Alsin/ALS2...... 56
1.2.1 Expression of ALS2...... 58
1.2.2 ALS2 exhibits GEF activity...... 60
1.2.3 ALS2 regulates endosomal morphology ...... 62
1.2.4 ALS2 binds to mutant SOD1 and displays neuroprotective activity...... 63
1.3 The Ras Superfamily of GTPases...... 64
1.3.1 Rho family GTPases ...... 67
1.3.2 Rab family GTPases ...... 72
1.4 Guanine nucleotide exchange factors (GEFs) ...... 74
1.4.1 DH/PH (Dbl-family) GEFs...... 74
1.4.2 Rab GEFs...... 77
CHAPTER 2: MATERIALS AND METHODS ...... 78
2.1 Materials ...... 79
2.1.1 Stock solutions...... 79
2.1.2 General molecular biology reagents ...... 81
2.1.2.1 Plasmids...... 81
2.1.2.2 Primers...... 81
2.1.2.3 Growth of E. coli for DNA purification: media ...... 82
6 2.1.2.4 Plasmid DNA preparation from E. coli ...... 82
2.1.2.5 Agarose gel electrophoresis of nucleic acids...... 83
2.1.2.6 Polymerase chain reaction (PCR) enzymes...... 83
2.1.3 Site-directed mutagenesis ...... 84
2.1.4 Purification of GST fusion proteins...... 85
2.1.5 Protein analysis...... 85
2.1.5.1 Protein sample preparation...... 85
2.1.5.2 Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE)
...... 87
2.1.5.3 Immunoblotting (Western blotting) solutions ...... 89
2.1.5.4 Protein staining...... 89
2.1.5.5 Treatment of protein samples with λ protein phosphatase...... 90
2.1.6 Antibodies...... 90
2.1.6.1 Primary antibodies...... 90
2.1.6.2 Secondary antibodies...... 90
2.1.7 Mammalian cell culture ...... 92
2.1.7.1 Chinese Hamster Ovary (CHO) cell culture...... 92
2.1.7.2 Primary neuronal culture ...... 92
2.1.8 In vitro protein kinase assays...... 93
2.1.9 Immunohistochemistry ...... 94
2.2 Methods...... 95
2.2.1 General molecular biology methods...... 95
2.2.1.1 Quantitation of nucleic acids...... 95
2.2.1.2 Restriction enzyme digestion of DNA ...... 95
2.2.1.3 Alkaline phosphatase treatment...... 96
7 2.2.1.4 Agarose gel electrophoresis of DNA...... 96
2.2.1.5 Recovery of DNA from agarose gels ...... 96
2.2.1.6 Ethanol precipitation of double-stranded DNA...... 97
2.2.1.7 Purification of nucleic acids ...... 97
2.2.1.8 Ligation of prepared vectors and DNA fragments ...... 97
2.2.1.9 Preparation of electrocompetent bacteria ...... 98
2.2.1.10 Electroporation of DH5α cells...... 99
2.2.1.11 Screening recombinant clones...... 99
2.2.1.12 Large scale preparation of plasmid DNA...... 100
2.2.1.13 DNA sequencing ...... 101
2.2.1.14 Polymerase chain reaction (PCR)...... 101
2.2.2 PCR-based site-directed mutagenesis...... 102
2.2.2.1 Digesting and polishing the PCR product ...... 104
2.2.2.2 Ligating the PCR product...... 104
2.2.2.3 Transformation into XL1-Blue supercompetent cells ...... 105
2.2.2.4 Screening for mutated plasmids ...... 105
2.2.3 Purification of GST fusion proteins...... 106
2.2.4 Protein analysis...... 108
2.2.4.1 Protein concentration determination...... 108
2.2.4.2 SDS-PAGE of protein samples ...... 109
2.2.4.3 Immunoblotting (Western blotting)...... 109
2.2.4.4 Treatment of samples with λ protein phosphatase ...... 110
2.2.5 Preparation of rabbit polyclonal antibodies...... 111
2.2.6 Mammalian cell culture and transfection...... 111
2.2.6.1 CHO cell culture...... 111
2.2.6.2 Primary embryonic rat cortical and hippocampal neuron culture ...... 112
8 2.2.6.3 Transient transfection ...... 113
2.2.7 GTPase activation assays...... 114
2.2.8 Immunoprecipitation and in vitro protein kinase assays ...... 115
2.2.8.1 Immunoprecipitation from cell lysates...... 115
2.2.8.2 Immunoprecipitation from rat brain homogenate...... 116
2.2.8.3 In vitro kinase assays...... 117
2.2.9 CHO cell fractionation...... 118
2.2.10 Immunohistochemistry ...... 118
2.2.11 Immunofluorescence...... 120
CHAPTER 3: PREPARATION OF AN ALS2 ANTIBODY
AND LOCALISATION OF ALS2 IN NEURONAL TISSUES
...... 121
3.1 Introduction...... 122
3.2 Methods...... 123
3.2.1 PCR and cloning into pGEX expression vector...... 123
3.2.2 Preparation of GST fusion protein...... 123
3.2.3 Production of a polyclonal antibody to ALS2 ...... 124
3.2.4 Affinity purification of ALS2 polyclonal antibody ...... 124
3.2.5 Cell culture and transfection...... 125
3.2.6 CHO cell fractionation...... 125
3.2.7 Antibodies...... 126
3.3 Results ...... 126
9 3.3.1 GST fusion protein preparation ...... 126
3.3.2 GST fusion protein expression and purification...... 127
3.3.3 Characterisation of ALS2 antibody ...... 128
3.3.4 Localisation of ALS2 in brain and spinal cord...... 129
3.3.5 Overexpressed ALS2 is localised in cytoplasmic and membranous fractions of
mammalian cells ...... 130
3.4 Discussion...... 130
CHAPTER 4: ALS2 ACTS AS GEF FOR RAC1 AND
ACTIVATES PAK1...... 140
4.1 Introduction...... 141
4.2 Methods...... 142
4.2.1 Plasmids...... 142
4.2.2 Cell culture and transfection...... 143
4.2.3 GTPase activation assays...... 143
4.2.4 In vitro kinase assays ...... 144
4.3 Results ...... 144
4.3.1 ALS2 acts as a GEF for Rac1 but not RhoA or Cdc42 ...... 144
4.3.2 ALS2 activates PAK1 and this is dependent on a functional DH domain ...... 145
4.4 Discussion...... 146
10 CHAPTER 5: ALS2 IS PRESENT IN NEURONAL
GROWTH CONES AND PROMOTES NEURITE
OUTGROWTH...... 153
5.1 Introduction...... 154
5.2 Materials and Methods...... 155
5.2.1 Antibodies and immunofluorescence microscopy...... 155
5.2.2 Preparation of mouse brain homogenates...... 155
5.2.3 Neurite outgrowth measurements ...... 156
5.2.4 Statistics...... 157
5.3 Results ...... 157
5.3.1 Developmental expression of ALS2...... 157
5.3.2 ALS2 is present in growth cones of hippocampal neurons, where it is co-
localised with Rac1, F-actin and α-tubulin...... 157
5.3.3 Overexpression of ALS2 (but not ALS2ΔDH) promotes neurite outgrowth in rat
embryonic cortical neurons via a Rac-dependent mechanism...... 158
5.3.4 Overexpression of ALS2 does not affect the number of neurites per cell, or the
extent of neurite branching, in cultured cortical neurons ...... 161
5.4 Discussion...... 161
CHAPTER 6: ALS2 IS A PHOSPHOPROTEIN...... 173
6.1 Introduction...... 174
11 6.2 Materials and Methods...... 175
6.2.1 Cell culture and transfection...... 175
6.2.2 Mass spectrometric sequencing of ALS2 ...... 175
6.2.3 Rac activation assay...... 176
6.2.4 PAK1 kinase assay ...... 177
6.2.5 Neurite outgrowth measurements ...... 177
6.3 Results ...... 177
6.3.1 ALS2 is a phosphoprotein ...... 177
6.3.2 Mutation of serine/threonine-proline sites to alanine does not affect the activity
of ALS2 ...... 178
6.4 Discussion...... 178
CHAPTER 7: DISCUSSION & FUTURE DIRECTIONS . 183
7.1 Summary of findings...... 184
7.2 The role of Alsin/ALS2 in motor neuron disease...... 185
7.3 Future directions ...... 187
REFERENCES...... 190
12 LIST OF FIGURES AND TABLES
Figures
Figure 1.1 Schematic of ALS2 (Long-form and Short-form) and predicted disease mutants...... 59
Figure 1.2 Schematic diagram of the small GTPase activation cycle...... 66
Figure 3.1 Schematic of ALS2 Long-form, ALS2 Short-form and GST-ALS2452-668..132
Figure 3.2 PCR amplification of sequences encoding GST-ALS2452-668...... 133
Figure 3.3 GST-ALS2452-668 and GST protein expression ...... 134
Figure 3.4 Purification of GST-ALS2452-668...... 135
Figure 3.5 Characterisation of ALS2 antibody ...... 136
Figure 3.6 Localisation of ALS2 in adult rat brain and spinal cord sections...... 137
Figure 3.7 Affinity-purified ALS2 antibody immunoprecipitates ALS2 from rat brain
...... 138
Figure 3.8 ALS2 is present in the cytosolic and membrane fractions of transfected CHO cells ...... 139
Figure 4.1 ALS2 stimulates Rac activity ...... 150
Figure 4.2 ALS2ΔDH does not activate Rac1 ...... 151
Figure 4.3 ALS2 activates PAK1...... 152
Figure 5.1 Developmental expression of ALS2 in mouse brain and cultured rat embryonic cortical neurons...... 167
Figure 5.2 Subcellular localisation of ALS2 in cultured embryonic neurons...... 168
Figure 5.3 Subcellular localisation of ALS2 in neuronal growth cones ...... 169
Figure 5.4 Transfected ALS2 and ALS2ΔDH display identical subcellular localisation to endogenous ALS2...... 170
Figure 5.5 ALS2 promotes neurite outgrowth in cultured rat cortical neurons ...... 171
13 Figure 5.6 Overexpression of ALS2 does not affect the number of neurites or the extent of neurite branching in cortical neurons...... 172
Figure 6.1 ALS2 is phosphorylated on serine/threonine-proline residues in embryonic rat cortical neurons...... 180
Figure 6.2 Phosphorylation of ALS2 ...... 181
Figure 6.3 Mutation of ALS2 phosphorylation sites does not affect ALS2 Rac GEF activity, PAK1 activation or neurite outgrowth ...... 182
Tables
Table 1.1 Classification of Motor Neuron Diseases ...... 22
Table 1.2 Loci identified in MND...... 26
Table 1.3 Loci identified in HSP...... 28
Table 1.4 Mammalian Ras GTPase Superfamily...... 65
Table 1.5 Selected effectors of the Rho GTPase family ...... 69
Table 2.1 Vectors ...... 81
Table 2.2 Mammalian expression plasmids...... 81
Table 2.3 Primary antibodies ...... 91
Table 2.4 PCR Cycling Parameters...... 102
Table 2.5 ExSite™ Mutagenesis Cycling Parameters ...... 104
Table 2.6 Transient transfection of CHO cells with Lipofectamine™ Reagent...... 113
14 PUBLICATIONS ARISING FROM THIS WORK
Peer Reviewed Research Paper
Tudor, E. L., Perkinton, M. S., Schmidt, A., Ackerley, S., Brownlees, J., Jacobsen, N. J.,
Byers, H. L., Ward, M., Hall, A., Leigh, P. N., Shaw, C. E., McLoughlin, D. M., and
C.C. Miller. 2005. ALS2/Alsin Regulates Rac-PAK Signalling and Neurite Outgrowth.
J. Biol. Chem. 280:34735-34740.
Published Abstracts
Elizabeth Tudor, Steven Ackerley, Nicholas O Jacobsen, Janet Brownlees, Steven
Banner, Anja Schmidt, Chris Shaw and Chris Miller (2003) “Alsin/ALS2 Is A Guanine
Nucleotide Exchange Factor That Is Present In Motor Neurons And Regulates Rho/Rac
Signalling” 33rd Annual Meeting of the Society for Neuroscience, New Orleans, USA.
Program No. 528.2.
Elizabeth Tudor, Steven Ackerley, Nicholas O Jacobsen, Janet Brownlees, Steven
Banner, Anja Schmidt, Chris Shaw and Chris Miller (2004) “Alsin/ALS2 Is A Guanine
Nucleotide Exchange Factor That Is Present In Motor Neurons And Regulates Rho/Rac
Signalling” 2nd European ALS Research Workshop (and European ALS Consortium
Young Investigators Meeting), Nice, France. Program No. P6.
15 ABBREVIATIONS
Amino acids: single letter code and abbreviations
Single Abbreviation Amino Acid letter code A Ala Alanine C Cys Cysteine D Asp Aspartic acid E Glu Glutamic acid F Phe Phenylalanine G Gly Glycine H His Histidine I Ile Isoleucine K Lys Lysine L Leu Leucine M Met Methionine N Asn Asparagine P Pro Proline Q Gln Glutamine R Arg Arginine S Ser Serine T Thr Threonine V Val Valine W Trp Tryptophan X Xxx Any amino acid Y Tyr Tyrosine
Other abbreviations
ABC avidin-biotinylated enzyme complex
ALS amyotrophic lateral sclerosis
16 Amp ampicillin
AMPA alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
APS ammonium persulphate
ATP adenosine 5′-triphosphate
β-gal β-galactosidase bp nucleotide base pair
BSA bovine serum albumin
Ca2+ calcium ion
CAT chloramphenicol acetyl transferase
CCD charge coupled device
C. elegans Caenorhabditis elegans
CHO Chinese hamster ovary cell line cDNA complementary DNA
CIAP calf intestinal alkaline phosphatase
CRIB Cdc42/Rac interactive binding
C-terminus carboxyl-terminus
Cu copper
DAB 3’,3’-diaminobenzidine tetrahydrochloride
Dbl diffuse B-cell lymphoma-associated protein ddH2O double distilled water
DH dbl homology
DH5α Douglas Hanahan bacterial strain 5α
DIV day(s) in vitro
DMSO dimethyl sulfoxide
DNA deoxyribonucleic acid dNTPs deoxyribonucleotide triphosphates
17 DTT dithiothreitol
E embryonic day
ECL enhanced chemiluminescence
E. coli Escherichia coli
EDTA ethylenediaminetetraacetic acid
EGTA ethyleneglycol-bis (β-aminoethylether) N,N,N′,N′-tetraacetic acid
ENU N-Ethyl-N-nitrosurea
FALS familial amyotrophic lateral sclerosis
FBS fetal bovine serum
Fgd1 faciogenital dysplasia 1-associated protein xg centrifugal force
GAP GTPase activating protein
GDI guanine nucleotide dissociation inhibitor
GDP guanosine 5’-diphosphate
GEF guanine nucleotide exchange factor
GFP green fluorescent protein
GST glutathione S-transferase
GTP guanosine 5’-triphosphate
GTPase guanosine 5’-triphosphatase
HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
HRP horse radish peroxidase
HSP hereditary spastic paraplegia
IAHSP infantile-onset ascending hereditary spastic paraplegia
IF intermediate filament
IgG immunoglobulin gamma
IP immunoprecipitation
18 IPTG isothiopropyl-1-thio-β-D-galactopyranoside kDa kiloDalton
LB Luria Bertani
LMN lower motor neuron
MBP myelin basic protein
Mg2+ magnesium ion
MND motor neuron disease
MLB magnesium lysis buffer mRNA messenger RNA
Net1 neuroepithelial cell transforming gene 1
NF neurofilament
NF-H neurofilament heavy chain
NF-L neurofilament light chain
NF-M neurofilament medium chain
NGS normal goat serum
NMDA N-methyl-D-aspartate
NP-40 Nonidet P40
N-terminus amino-terminus
OD optical density
PAK p21 associated kinase
PBD p21 binding domain
PBS phosphate buffered saline
PCR polymerase chain reaction pH inverse log of hydrogen ion concentration
PH pleckstrin homology
PI(3)K phosphatidylinositol 3-kinase
19 PLS primary lateral sclerosis
PMSF phenylmethyl sulphonyl fluoride
RBD Rho binding domain
RCC1 regulator of chromosome condensation 1 protein
RNA ribonucleic acid
ROS reactive oxygen species
SALS sporadic amyotrophic lateral sclerosis
SDS-PAGE sodium dodecyl sulphate-polyacrylamide gel electrophoresis
SMA spinal muscular atrophy
SMN survival motor neuron
SOD1 Cu/Zn superoxide dismutase 1
TBS tris-buffered saline
Tris tris(hydroxymethyl)aminomethane
Tween-20 polyoxyethylene-sorbitan monolaurate
U units
UMN upper motor neuron
UV ultraviolet
VPS9 vacuolar protein sorting 9
X-gal 5-Bromo-4-chloro-3-indoyl-β-D-galactopyranoside
Zn zinc
20
CHAPTER 1: INTRODUCTION
21 1.1 Motor Neuron Disease
The term ‘Motor Neuron Disease’ (MND) describes a group of progressive neurodegenerative disorders involving selective loss of upper motor neurons (UMN) in the cortex and/or lower motor neurons (LMN) in the brainstem and spinal cord. Clinical features include paralysis and spasticity of the limbs and bulbar muscles (UMN involvement) and muscle weakness, wasting and fasciculation (LMN involvement)
(Devon et al., 2003; Talbot, 2002). Amyotrophic Lateral Sclerosis (ALS) is the most common MND, with a prevalence of 2-3 per 100,000 people (Cleveland and Rothstein,
2001). Following progressive degeneration of both UMNs and LMNs, death usually occurs within 1-5 years. By contrast, other forms of MND involve only the upper or lower motor neurons, resulting in disorders with varying severity (Table 1.1).
Table 1.1 Classification of Motor Neuron Diseases
Combined UMN and LMN involvement • Amyotrophic lateral sclerosis (ALS)
Proximal hereditary motor neuropathy Pure LMN involvement • • Hereditary bulbar palsy • Hexosaminidase deficiency • Multifocal motor neuropathies • Post polio syndrome • Post irradiation syndrome • Monomelic, focal or segmental spinal muscular atrophy (SMA)
Pure UMN involvement • Primary lateral sclerosis (PLS) • Hereditary spastic paraplegia (HSP) • Neurolathyrism • Konzo
Mixed Motor and Sensory involvement • ALS with fronto-temporal dementia (ALS- FTD) • Charcot Marie Tooth Disease (CMT) • Distal hereditary motor neuropathy
(Donaghy, 1999; Talbot, 2002)
22 1.1.1 Clinical features and pathology of MND
Amyotrophic Lateral Sclerosis (ALS)
The ‘Classical’ form of ALS is a sporadic disease of mid- to late- life, with a mean age at onset of 58 years (Ringel et al., 1993), although rare juvenile forms have also been described with age at onset varying from 3-23 years, and with a slower progression of about 10-15 years (Ben Hamida et al., 1990). The major pathological features of classical ALS are degeneration of the corticospinal tracts and extensive loss of anterior horn cells (Ghatak et al., 1986; Hughes, 1982; Leigh and Garofolo, 1995), degeneration and loss of Betz cells and other pyramidal cells in the primary motor cortex (Hammer et al., 1979; Maekawa et al., 2004; Udaka et al., 1986), and reactive gliosis in the motor cortex and spinal cord (Ekblom et al., 1994; Kawamata et al., 1992; Murayama et al.,
1991; Schiffer et al., 1996).
An established pathological hallmark of ALS is the presence of various inclusion bodies in degenerating neurons and surrounding reactive astrocytes (Barbeito et al., 2004).
Ubiquitinated inclusions are the most common and specific type of inclusion in ALS and are found in LMNs of the spinal cord and brainstem (Matsumoto et al., 1993) and in corticospinal UMNs (Sasaki and Maruyama, 1994). They are classified as ‘Lewy body- like inclusions’ (LBIs) and ‘Skein-like inclusions’ (SLIs) (He and Hays, 2004;
Kawashima et al., 2000). The exact composition of such inclusions is not known, although proteins identified so far include (in varying amounts) ubiquitin (Leigh et al.,
1991; Murayama et al., 1989), Cu/Zn superoxide dismutase 1 (SOD1) (Shibata et al.,
1996a; Shibata et al., 1994), peripherin (He and Hays, 2004) and Dorfin (a RING-finger type E3 ubiquitin ligase) (Niwa et al., 2002). Accumulations of intermediate filament proteins (mainly hyperphosphorylated neurofilament subunits and peripherin) are found in hyaline conglomerate inclusions (HCIs) and axonal ‘spheroids’ in spinal cord motor
23 neurons (Corbo and Hays, 1992; Munoz et al., 1988; Sobue et al., 1990) and pyramidal cells of the motor cortex (Troost et al., 1992) in ALS post-mortem tissue. Additionally,
Bunina bodies (BBs), which are Cystatin C-containing inclusions, are found in the cell bodies of motor neurons in ALS (Okamoto et al., 1993; Sasaki and Maruyama, 1994), although these are now thought to be less specific for ALS than the ubiquitinated and neurofilamentous inclusions, as they are similar to structures found in neurons of aged rats and humans (Kusaka, 1999). Other neuropathological features seen in ALS include fragmentation of the Golgi apparatus (Fujita et al., 2000; Fujita et al., 2002; Gonatas et al., 1998), mitochondrial vacuolisation (Okamoto et al., 1990) and ultrastructural abnormalities of synaptic terminals (Sasaki and Iwata, 1996).
Primary lateral sclerosis (PLS)
PLS is characterised by predominantly UMN degeneration resulting in spinobulbar spasticity. LMN and extra-motor system involvement has been reported in some cases, although if present this is usually mild or only seen at a late stage in the disease (Le
Forestier et al., 2001). The majority of PLS has an average age at onset of 50 years
(Pringle et al., 1992), and rarer juvenile forms have also been described (jPLS) (Gascon et al., 1995; Grunnet et al., 1989; Yang et al., 2001). In contrast to ALS, PLS has a very slow progression of about 15 years (Pringle et al., 1992) and cortical atrophy is more extensive (Kuipers-Upmeijer et al., 2001).
Hereditary spastic paraplegia (HSP)
Hereditary spastic paraplegia or hereditary spastic paraparesis (HSP) is a group of inherited neurodegenerative disorders of lower limb spastic paralysis, caused either by failure of development or progressive degeneration of the UMNs of the corticospinal tract. HSP is described as either ‘pure’ or ‘complex’, with autosomal dominant,
24 autosomal recessive and X-linked recessive patterns of inheritance (see section 1.1.2).
Complex HSP is associated with other clinical features such as distal amyotrophy, mental retardation, pigmentary retinopathy, sensory neuropathy or ataxia (Donaghy,
1999). Degenerating neurons of the corticospinal tract exhibit a dying back axonopathy in which synaptic terminal and axonal degeneration precedes cell body degeneration, and this typically occurs in cells with the longest axons first (Donaghy, 1999).
1.1.2 Genetics of MND
Familial ALS (FALS) subgroups
Whereas approximately 90% of ALS cases are sporadic (SALS) with no known genetic linkage, 10% are inherited and known as familial ALS (FALS) (Strong et al., 1991).
SALS and FALS share indistinguishable clinical and pathological features (Mulder et al., 1986) which suggests the possibility of common mechanisms underlying motor neuron degeneration in both sporadic and familial ALS. FALS inheritance is mainly autosomal dominant, but autosomal recessive and X-linked modes of inheritance have also been reported (Table 1.2). SALS and the majority of FALS is adult-onset, although three loci have been described that lead to juvenile-onset disease (ALS2, ALS4 and
ALS5). Approximately 20% of FALS cases are caused by point mutations in the Cu/Zn
SOD1 gene on chromosome 21 (Deng et al., 1993; Rosen et al., 1993). Over 100 SOD1 mutations have been described so far (for an updated list see www.alsod.org) resulting in highly variable phenotypes, even within families carrying the same mutation. Since the discovery of SOD1 mutations most studies on the pathogenesis of ALS have focused on mutant SOD1 mediated motor neuron death, utilising transgenic in vivo and in vitro models.
25 Table 1.2 Loci identified in MND
DISEASE ONSET INHERITANCE LOCUS GENE REFERENCE
Combined UMN and LMN involvement ALS1 Adult AD/AR 21q22.1 SOD1 (Rosen et al., 1993) ALS2/ (Hadano et al., 2001; Yang ALS2 Juvenile AR 2q33.2 Alsin et al., 2001) ALS3 Adult AD 18q21 - (Hand et al., 2002) (Blair et al., 2000; Chen et ALS4 Juvenile AD 9q34 SETX al., 2004) ALS5 Juvenile AR 15q15-22 - (Hentati et al., 1998) (Ruddy et al., 2003; Sapp ALS6 Adult AD 16q12 - et al., 2003; Abalkhail et al., 2003) ALS7 Adult AD 20ptel - (Sapp et al., 2003) ALS8 Adult AD 20q13.33 VAPB (Nishimura et al., 2004) ALS X Adult X-linked Xp11-q12 - (Siddique et al., 1998) Pure LMN involvement (Puls et al., 2003; Puls et SMA Adult AD 2p13 DCTN1 al., 2005) (Lefebvre et al., 1995; SMA Juvenile AR 5q13 SMN1 Rodrigues et al., 1995) SBMA/ Androgen (La Spada et al., 1991; Kennedy’s Adult X-linked Xq11-q12 Receptor Biancalana et al., 1992) Disease Pure UMN involvement jPLS Juvenile AR 2q33.2 ALS2 (Yang et al., 2001) HSP See Table 1.3 Mixed sensory and motor involvement ALS- FTD/ (Siddique and Hentati, Adult AD 17q21 MAPT Parkinsonism 1995) ALS- FTD/ Adult AD 17q - (Wilhelmsen et al., 2004) Parkinsonism ALS- FTD Adult AD 9q21-22 - (Hosler et al., 2000)
Abbreviations: ALS= amyotrophic lateral sclerosis; AD= autosomal dominant; AR= autosomal recessive; SOD1= Cu/Zn superoxide dismutase 1; SETX= Senataxin; VAPB= vesicle associated membrane protein (VAMP)/ synaptobrevin-associated membrane protein B; SMA= spinal muscular atrophy; SBMA= spinobulbar muscular atrophy; jPLS= juvenile primary lateral sclerosis; HSP= hereditary spastic paraplegia; FTD= frontotemporal dementia; MAPT= microtubule associated protein tau; DCTN1= dynactin 1; SMN1= survival motor neuron 1
26 Genetic risk factors for SALS
Several genetic risk factors, or susceptibility genes, have been implicated in sporadic
ALS. These include neurofilament heavy chain (NF-H) (Al-Chalabi et al., 1999;
Figlewicz et al., 1994; Skvortsova et al., 2004; Tomkins et al., 1998), vascular endothelial growth factor (VEGF) (Greenway et al., 2004), survival motor neuron 2
(SMN2) (Veldink et al., 2001), apolipoprotein E epsilon 4 (ApoEε4) (Drory et al., 2001;
Moulard et al., 1996), the gene encoding the p150 subunit of dynactin (DCTN1) (Munch et al., 2004), the mitochondrial gene cytochrome c oxidase (Comi et al., 1998), the gene encoding DNA repair enzyme abasic endonuclease (APEX) (Hayward et al., 1999) and aberrant mRNA processing of excitatory amino acid transporter 2 (EAAT2) (Aoki et al.,
1998; Lin et al., 1998). The absence of mutations in FALS suggests that the mutations found in SALS patients are susceptibility factors rather then a direct cause of the disease
(Aoki et al., 1998; Vechio et al., 1996).
Hereditary spastic paraplegia (HSP)
21 HSP genes have been mapped so far, 9 of which have been identified as follows: cell adhesion molecule L1 (L1-CAM), the gene encoding proteolipid protein (PLP1), atlastin, spastin, the neuronal specific kinesin protein gene KIF5A, the mitochondrial chaperone heat shock protein gene HSP60 (mitochondrial chaperonin), paraplegin, non imprinted in Prader-Willi/Angelman syndrome 1 (NIPA1), spartin, Berardinelli-Seip congenital lipodystrophy (BSCL2), maspardin and the ALS2/alsin gene ALS2 (Table
1.3).
27 Table 1.3 Loci identified in HSP
GENE LOCUS DISEASE INHERITANCE REFERENCE
(De Jonghe et al., 1996; SPG4/ Spastin 2p22 Pure HSP AD Hazan et al., 1999)
SPG13/ Hsp60 2q24-34 Pure HSP AD (Hansen et al., 2002)
Pure or (Reid et al., 1999a; Reid SPG10/ KIF5A 12q13 Complicated AD et al., 2002) HSP (Dalpozzo et al., 2003; SPG3A/ Atlastin 14q11- 21 Pure HSP AD Zhao et al., 2001b)
SPG6/ NIPA1 15q11.1 Pure HSP AD (Rainier et al., 2003)
Pure or (Casari et al., 1998; SPG7/ Paraplegin 16q24.3 Complicated AR Wilkinson et al., 2004) HSP Troyer SPG20/ Spartin 13q12.3 AR (Patel et al., 2002) syndrome Complicated SPG1/ L1CAM Xq28 X-linkedR (Jouet et al., 1994) HSP Complicated SPG2/ PLP Xq21 X-linkedR (Bonneau et al., 1993) HSP (Eymard-Pierre et al., ALS2/alsin 2q33.2 IAHSP AR 2002; Gros-Louis et al., 2003b) (Patel et al., 2001; Silver SPG17/ BSCL2 11q12- 14 AD Windpassinger et al., Syndrome 2004) (Reid et al., 1999b; SPG8 8q23- 24 Pure HSP AD Rocco et al., 2000) SPG21/ Mast 15q22.31 AR (Simpson et al., 2003) Maspardin Syndrome 10q23.3- Complicated SPG9 AD (Lo Nigro et al., 2000) 24.2 HSP (Ashley-Koch et al., SPG12 19q13 Pure HSP AD 2001; Reid et al., 2000) SPG19 9q33- 34 Pure HSP AD (Valente et al., 2002) Complicated SPG14 3q27- 28 AR (Vazza et al., 2000) HSP SPG5 8q12-13 Pure HSP AR (Muglia et al., 2004) (Martinez Murillo et al., SPG11 15q13-15 ARHSP AR 1999; Shibasaki et al., 2000) Complicated SPG15 14q22-24 AR (Hughes et al., 2001) HSP Pure or SPG16 Xq11.2 Complicated X- linkedR (Steinmuller et al., 1997) HSP 12p11.1– Complicated SPG26 AR (Wilkinson et al., 2005) 12q14 HSP
Abbreviations: SPG= Spastic Gait gene locus; AD= Autosomal dominant; AR= Autosomal recessive;
28 HSP60= heat shock protein 60 (mitochondrial Chaperonin); KIF5A= kinesin heavy chain; NIPA1= non imprinted in Prader-Willi/Angelman syndrome 1; L1CAM= L1 cell adhesion molecule; PLP= proteolipid protein; IAHSP= infantile-onset ascending HSP; BSCL2= Berardinelli-Seip congenital lipodystrophy; ARHSP= autosomal recessive HSP.
1.1.3 Animal models of MND
Animal models of neurodegeneration are an invaluable tool for studying the pathogenic mechanisms involved in vivo at all stages of the disease, even before symptom onset.
Murine models of MND in particular are used to study the cellular and molecular pathways involved, as such pathways are usually highly conserved between mouse and human (Hafezparast et al., 2003a). A selection of the most widely used mouse models are described below. Other animal models of MND include mutant SOD1 rats
(Howland et al., 2002) and survival motor neuron (SMN) knockdown zebrafish
(McWhorter et al., 2003).
Transgenic mice have been created to investigate the function of proteins already known or suspected to be involved in MND, such as SOD1 and neurofilaments (NFs). This is known as a ‘genotype driven’ approach. In contrast, mice have been found
(‘natural/spontaneous mutants’) or created (by the use of irradiation or chemical mutagens) that exhibit a MND pathology, in a ‘phenotype driven’ approach. The causal gene has been identified in many of these mice and this has brought useful information regarding the proteins and pathways involved in motor neuron degeneration.
1.1.3.1 Spontaneous mutants
Wobbler mouse
The Wobbler mouse has an unsteady gait with progressive weakness, typically dying by
3 months of age (Duchen and Strich, 1968), and is often used as a model of MND.
29 However, recent studies have suggested that the disease in Wobbler mice may be a more generalised neurodegeneration than MND, as degeneration is seen in the thalamus, cerebellum and brainstem, and this precedes degeneration of the motor neurons and onset of gliosis (Rathke-Hartlieb et al., 1999). The gene mutated in this autosomal recessive disease model is not yet known but has been mapped to chromosome 11
(Kaupmann et al., 1992).
Progressive motor neuronopathy (pmn) mouse
The progressive motor neuronopathy (pmn) mutant mouse is a widely used model of
MND (most closely resembling SMA) that develops hind-limb paralysis and displays progressive degeneration of motor neurons until death occurs in the early postnatal period (6-8 weeks) (Schmalbruch et al., 1991). The autosomal recessive mutation causing this pathology was originally mapped to chromosome 13 (Brunialti et al., 1995) and has been subsequently identified as a point mutation in the tubulin-specific chaperone E (Tbce) gene (Bommel et al., 2002; Martin et al., 2002). Tbce encodes the protein cofactor E (CofE) which is a tubulin-specific chaperone and is essential in the correct assembly of microtubules (Bommel et al., 2002; Martin et al., 2002). This discovery is extremely interesting as defects of microtubule function and impairment of axonal transport have been extensively implicated in pathogenic mechanisms of MND, as discussed in section 1.1.4.3.
ENU-induced mutants
N-Ethyl-N-nitrosurea (ENU) is a chemical mutagen that produces point mutations in the genome, which enables identification of the causal gene by methods such as ‘positional candidate cloning’, and is currently being used to generate new mouse models of MND
(Brown and Balling, 2001; Chen et al., 2000; Hafezparast et al., 2003a). So far, this
30 approach has successfully created the mouse Legs at odd angles (Loa) which develops a late onset progressive motor neuron disease with neuropathological features similar to those seen in other MND mouse models, including mitochondrial swelling, Golgi fragmentation and cytoplasmic inclusions (Hafezparast et al., 2003a; Rogers et al.,
2001). The causal mutation in the Loa mouse is inherited in an autosomal dominant fashion, and has been found to occur in the cytoplasmic dynein heavy chain gene
Dnchc1, which suggests involvement of the dynein-dynactin complex in MND.
Furthermore, a different mutation in dynein heavy chain has been identified in a second
ENU mutagenesis-generated mouse, Cramping1 (Cra1). The role of the dynein-dynactin complex in the pathogenesis of MND is discussed in section 1.1.4.3.
1.1.3.2 Targeted mutants
SOD1 transgenic mice
SOD1 is a ubiquitous enzyme and is highly expressed in motor neurons (Pardo et al.,
1995). The primary function of SOD1 within the cell is to catalyse the conversion
(dismutation) of superoxide radicals (by-products of normal cellular metabolism) to hydrogen peroxide, which is then eliminated by other free radical-scavenging enzymes
(glutathione peroxidase and catalase). It also has other activities in the cell including peroxidase activity (resulting in generation of hydroxyl radicals from hydrogen peroxide or superoxide, or production of nitronium species from peroxynitrite) and protection of the enzyme calcineurin from inactivation (Hodgson and Fridovich, 1975; Wang et al.,
1996). Transgenic mice carrying the human SOD1 mutants found in FALS (the most widely-used mutants are G37R, G85R, G93A, G93R) on a wild-type mouse SOD1 background develop progressive muscle weakness and atrophy and have a pathology that highly resembles the human disease, including loss of motor neurons and interneurons, reactive astrocytosis, and inclusion bodies immunoreactive for ubiquitin,
31 NFs and SOD1 (Bruijn et al., 1997b; Cha et al., 1998; Dal Canto and Gurney, 1995;
Gonatas et al., 1998; Gurney et al., 1994; Kong and Xu, 1998; Mourelatos et al., 1996;
Tu et al., 1996; Wong and Borchelt, 1995; Zhang et al., 1997). Mice with high copy numbers (high expresser mice) show early onset of disease, whereas mice with low copy numbers (low expresser mice) are affected by a late-onset disease (Dal Canto and
Gurney, 1997), and the type of mutation seems to affect the severity/rapidity of progression (Gurney, 1997; Shibata, 2001).
SOD1 mutant mouse models have therefore been widely used to study the molecular and cellular processes occurring both before disease onset and during disease progression. It is believed that such research may lead to identification of therapeutic targets and/or elucidation of the neurodegenerative mechanisms involved in all forms of
MND. The use of mutant SOD1 mice has enabled the identification of pre-symptomatic neuropathology such as fragmentation of the Golgi apparatus and mitochondrial vacuolation (Dal Canto and Gurney, 1995; Mourelatos et al., 1996; Wong and Borchelt,
1995), and biological processes disrupted before and during disease progression such as axonal transport (Borchelt et al., 1998; Warita et al., 1998; Williamson and Cleveland,
1999; Zhang et al., 1997).
The means by which SOD1 exerts its toxicity is unknown, although it is thought to occur by a ‘gain of toxic function’ rather than loss of the dismutase activity of the protein. This is supported by several findings from the study of mutant SOD1 mice, including: SOD1 wild-type knockout mice do not develop a disease pathology (although they display subtle motor defects and their motor neurons show an increased sensitivity to axonal injury) (Flood et al., 1999; Reaume et al., 1996; Shefner et al., 1999); the toxicity of mutant SOD1 is not accelerated or reduced by loss of wild-type SOD1 in
32 transgenic mutant SOD1 mice (Bruijn et al., 1998), and is either unaffected (Bruijn et al., 1998) or enhanced (Jaarsma et al., 2000) by increasing wild-type SOD1 activity; some mutants still cause disease despite retaining their dismutase activity (Borchelt et al., 1994; Bowling et al., 1993).
Neurofilament and peripherin mouse models
Neurofilaments are type IV intermediate filament (IF) proteins that are a major component of the neuronal cytoskeleton, and are composed of 3 subunits that range in size according to the size of their tail domain; neurofilament light chain (NF-L; 68 kDa), neurofilament medium chain (NF-M; 95 kDa), neurofilament heavy chain (NF-H;
110 kDa). They are responsible for the maintenance of neuronal calibre and are particularly abundant in large myelinated neurons, such as those that are preferentially affected in MND, and are found in filamentous inclusions in spinal cord motor neurons of MND post-mortem tissue. Peripherin, a type III neuronal IF protein, is also found in the majority of motor neuron neurofilament inclusions in MND, although its expression is usually (in non-disease cases) most abundant in autonomic nerves and peripheral sensory neurons, with low levels in spinal motor neurons (Escurat et al., 1990; Parysek and Goldman, 1988; Troy et al., 1990a; Troy et al., 1990b). The discovery of intermediate filament inclusions in MND has led to the proposal that they are involved in the pathogenesis of the disease, and so transgenic mouse models have been created to investigate this possibility. Transgenic mice overexpressing human NF-H (Cote et al.,
1993), mouse NF-L (Xu et al., 1993) or mouse peripherin (Beaulieu et al., 1999) all develop an MND-like pathology with muscle atrophy, reduced axonal calibre and IF inclusions. This has led to extensive research into the role of IF proteins in the disease mechanisms that cause MND, as discussed in sections 1.1.4.1 and 1.1.4.3.
33 Mutant VEGF mice
Mice carrying a targeted deletion of the hypoxia response element (HRE) within the vascular endothelial growth factor (VEGF) promoter have a phenotype similar to that of human ALS (Oosthuyse et al., 2001). This has led to the proposal that this gene may be involved in ALS, although linkage studies have as yet failed to support this hypothesis
(Gros-Louis et al., 2003a). The mechanism by which the mouse phenotype occurs is unknown, although it has been proposed that motor neuron death could occur due to reduced perfusion under low oxygen conditions (with motor neurons being particularly vulnerable due to their large size and high energy/oxygen requirements), and/or VEGF could have neuroprotective effects on motor neurons under normal circumstances
(Oosthuyse et al., 2001). Indeed, treatment with VEGF has been found to protect motor neurons against cell death following spinal cord ischemia in mice (Lambrechts et al.,
2003) and in a mutant SOD1 rat model (Storkebaum et al., 2005).
1.1.4 Mechanisms of neurodegeneration in MND
The pathogenic mechanisms underlying the neuronal degeneration and death in MND are as yet unknown. There are currently several hypotheses concerning the pathogenic processes involved in MND, none of which are mutually exclusive. The main hypotheses include protein misfolding and aggregation, oxidative stress, disruption of axonal transport, and glutamatergic excitotoxicity; there is also evidence that inflammation, autoimmunity and apoptotic cell death pathways are involved, all of which will be discussed.
Cell types involved in MND
Activated/reactive astrocytes and microglia are often found in ALS post-mortem tissue
(Ekblom et al., 1994; Murayama et al., 1991; Kawamata et al., 1992; Schiffer et al.,
34 1996) and in mutant SOD1 transgenic mice (Bruijn et al., 1997b; Cha et al., 1998; Tu et al., 1996), and although the clinical and molecular pathology of MND indicates an obvious involvement of motor neurons, it is increasingly being recognised that glial cells also play an important role in the pathogenesis of the disease. Important insights into the interdependence of different cell types in MND pathogenesis have come from several studies in mutant SOD1 transgenic mice. The discovery that cell-specific expression of mutant SOD1 in either neurons or astrocytes did not cause disease in mice
(Gong et al., 2000; Lino et al., 2002; Pramatarova et al., 2001) indicated that both neuronal and non-neuronal cells could play a role in disease pathogenesis. A chimeric
SOD1 mouse model was subsequently created, composed of a mixture of normal cells and mutant SOD1-expressing cells in order to investigate the relationship between different cell types (Clement et al., 2003). The presence of wild-type cells delayed disease onset and extended the lifespan of the chimeric mice compared with those overexpressing mutant SOD1 ubiquitously, and more specifically, degeneration and death of mutant SOD1-expressing motor neurons was reduced when the neurons were surrounded with a sufficient number of normal non-neuronal cells, whereas normal motor neurons surrounded by mutant SOD1-expressing non-neuronal cells showed signs of degeneration such as ubiquitinated inclusions. These results indicate that the expression of mutant SOD1 in both neuronal and non-neuronal cell populations, and the interactions between them, is of fundamental importance in MND.
1.1.4.1 Toxicity of intracellular aggregates
Mutant SOD1 aggregates in FALS
The mechanism(s) by which mutant SOD1 causes FALS are not known, although a popular hypothesis is that the mutant protein forms abnormal toxic aggregates within the cell, leading to neuronal degeneration and death. It has been suggested that the
35 toxicity of these aggregates may be effected by disruption of axonal transport (Borchelt et al., 1998; Williamson and Cleveland, 1999), sequestration of heat shock proteins and other chaperones (Bruening et al., 1999; Okado-Matsumoto and Fridovich, 2002;
Shinder et al., 2001), dysfunction of the proteasome (Hoffman et al., 1996; Urushitani et al., 2002) and/or damage to mitochondria (Jaarsma et al., 2001; Kong and Xu, 1998;
Takeuchi et al., 2002a).
SOD1 aggregates are found in motor neurons and surrounding astrocytes in FALS post- mortem tissue (Kato et al., 1997; Shibata et al., 1996b); they are also an early indicator of disease, occurring before the onset of symptoms, in mutant SOD1 transgenic mice
(Bruijn et al., 1997b; Johnston et al., 2000; Stieber et al., 2000). They are not, however, a characteristic feature of sporadic ALS (Shibata et al., 1996a). Further support for a role of abnormal SOD1 aggregation in FALS pathogenesis comes from the finding that delaying the formation of such abnormal aggregates, which are high molecular weight
‘insoluble protein complexes’ (IPCs), delays the disease onset in mutant SOD1 transgenic mice. This delay in aggregate formation was achieved by the creation of double transgenic mice carrying both human SOD1G93A and chat-GluR2 (in which the
GluR2 subunit of the alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
(AMPA) receptor is overexpressed in a cholinergic neuron-specific manner, resulting in a large reduction in calcium-permeability of AMPA receptors, the significance of which will be discussed in section 1.1.4.4) (Tateno et al., 2004).
The way in which mutant SOD1 causes abnormal protein aggregation is unknown, although structural evidence suggests that it may be by means of destabilisation of normal dimers (Hough et al., 2004), and this has subsequently led to the proposal of dimer stabilisation as a potential therapeutic intervention (Ray and Lansbury, 2004).
36 Abnormal mutant SOD1 aggregates are not only present in the cytosol but have also been found in spinal cord mitochondria in both human cases and mouse models of mutant SOD1-mediated FALS (Liu et al., 2004; Pasinelli et al., 2004; Vijayvergiya et al., 2005). Together with the existence of extensive mitochondrial degeneration and vacuolation in mutant SOD1 mice at pre-clinical stages of the disease (Higgins et al.,
2002; Jaarsma et al., 2000; Wong et al., 1995), the localisation of such aggregates suggests an involvement of mitochondrial dysfunction in the pathogenesis of MND.
Intermediate filament aggregates in MND
Neurofilament subunits (NF-L, NF-M, NF-H) and peripherin are major components of the neuronal cytoskeleton and are frequently found in MND inclusion bodies (within neuronal cell bodies and axons) (Carpenter, 1968; Corbo and Hays, 1992; Hirano et al.,
1984; Migheli et al., 1993; Rouleau et al., 1996). Transgenic mice overexpressing IF proteins such as NF-L (Xu et al., 1993), human NF-H (hNF-H) (Cote et al., 1993), peripherin (Beaulieu et al., 1999) or alpha-internexin (Ching et al., 1999) all develop a motor neuron pathology with IF inclusions. In humans, mutations in NF-H have been reported in SALS patients (Al-Chalabi et al., 1999; Figlewicz et al., 1994; Tomkins et al., 1998), mutations in the NF-L gene have been found to cause the motor/sensory neuropathy Charcot Marie Tooth disease type 2 (CMT2) (De Jonghe et al., 2001;
Fabrizi et al., 2004; Georgiou et al., 2002; Jordanova et al., 2003; Mersiyanova et al.,
2000; Yoshihara et al., 2002; Zuchner et al., 2004) and two different mutations in the peripherin gene have recently been reported to cause MND (Gros-Louis et al., 2004;
Leung et al., 2004).
It has therefore been proposed that the abnormal IF accumulations found in ALS may contribute to the pathogenesis of the disease. It is unknown, however, how these
37 accumulations form and whether they play a detrimental or protective role in ALS. For example, crossing SOD1G37R mice with mice overexpressing hNF-H resulted in an extended lifespan compared with mice overexpressing SOD1G37R alone, and this correlated with the degree of perikaryal NF accumulation (Couillard-Després et al.,
1998). Similar results were found when the phenotype of peripherin overexpressing mice was rescued by overexpressing hNF-H (Beaulieu and Julien, 2003). The overexpression of hNF-H shifted the intracellular localisation of NF-H from the axonal to the perikaryal compartment, which suggests that the protective effect may be due to a reduction of axonal NF accumulation or an increase in perikaryal accumulation.
There is evidence that an increase in perikaryal accumulation is more important in protection from degeneration than a decrease in axonal accumulation, as decreasing the axonal NF content and axonal calibre in mutant SOD1 mice (by disruption in one allele of the NF-L gene), did not affect the disease severity or lifespan of the mice (Nguyen et al., 2000). It has been suggested that perikaryal NF inclusions may act as a
‘phosphorylation sink’ in which NFs are preferentially phosphorylated, thereby protecting other proteins that could be detrimental to the cell when hyperphosphorylated by kinases such as Cdk5 (Nguyen et al., 2001). In support of this hypothesis, it has been found that the perikaryal NF accumulations in the SOD1G37R/hNF-H mice are hyperphosphorylated and co-localise with Cdk5; and hyperphosphorylation of tau at
Cdk5 sites is reduced in these mice, suggesting that the NFs are preferentially phosphorylated by Cdk5 (Nguyen et al., 2001). In contrast, axonal accumulations of IF proteins are thought to have a detrimental effect, which may be caused by sequestration of essential proteins and organelles such as mitochondria, and/or by disruption of axonal transport, as discussed in Section 1.1.4.3.
38 Alternatively, sequestration of neuronal nitric oxide synthase (nNOS) in NF aggregates has been proposed to occur in neurons of mice overexpressing NF-L, leading to enhanced N-methyl-D-aspartate (NMDA)-mediated calcium influx that may cause neuronal cell death (Sanelli et al., 2004).
It is believed that NF subunit stoichiometry is a major factor in the formation of NF aggregates. This is supported by the finding that the motor neuron pathology found in hNF-H overexpressing mice was rescued by co-expressing human NF-L (hNF-L) at levels that restored the correct NF-L:NF-H stoichiometry. Additionally, reduced NF-L mRNA levels and selective alterations of NF expression have been observed in inclusion-bearing spinal cord motor neurons of ALS patients (Bergeron et al., 1994;
Menzies et al., 2002; Wong et al., 2000). Interestingly, in peripherin overexpressing mice, the disease was dramatically accelerated by a deficiency in NF-L (a phenomenon seen in ALS and in normal ageing) (Bergeron et al., 1994; Krekoski et al., 1996).
1.1.4.2 Oxidative stress
In ALS patients, biochemical changes indicative of oxidative damage, such as lipid peroxidation, free carbonyls, protein nitration and protein glycosylation, in motor neurons and spinal cord astrocytes suggests the involvement of oxidative stress in the disease (Abe et al., 1995; Beal et al., 1997; Bowling et al., 1993; Niebroj-Dobosz et al.,
2004; Shaw et al., 1995b; Shibata et al., 2001). For reviews see (Cookson and Shaw,
1999; Robberecht, 2000). Furthermore, the elevation of hydroxyl radicals in the spinal cord of pre-symptomatic SOD1 mutant mice has been reported in several studies
(Andrus et al., 1998; Bogdanov et al., 2000; Ferrante et al., 1997; Hall et al., 1998; Liu et al., 1998a), which suggests vulnerability of a particular subset of motor neurons at pre-clinical stages of the disease. The role of oxidative stress mechanisms leading to cell
39 death in MND has thus been investigated, particularly in relation to mutant SOD1- mediated FALS.
As mentioned earlier (Section 1.1.3), the toxicity of mutant SOD1 is thought to occur by means of a gain of (unknown) function rather than loss of its superoxide dismutase activity. This is suggested by the dominant inheritance pattern of FALS SOD1 mutations, the absence of any mutations causing severe truncations/loss of the protein and the finding that many of the mutants, for example D90A, retain their dismutase activity despite causing ALS. Although protein aggregation is a widely-favoured hypothesis for toxicity caused by mutant SOD1 in FALS, oxidative damage to the cell by mutant SOD1 has also been suggested as a pathogenic mechanism in FALS. It has also been postulated that the presence of such protein aggregates could be a consequence of reactive oxygen species (ROS) generation and oxidative modifications of the proteins (Gélinas et al., 2000; Rakhit et al., 2002; Valentine and Hart, 2003).
Studies using the spin trapping molecule 5’,5’-dimethylpyrroline-N-oxide (DMPO)
(which reduces oxidative stress propagation in NSC-34 motor neuron-like cells overexpressing SOD1G93A) (Liu et al., 2002) revealed that mutant SOD1 can become a source of free radicals by means of enhanced peroxidase activity, and can induce oxidative damage leading to lipid peroxidation, mitochondrial dysfunction and cell death (Liu et al., 2002; Wiedau-Pazos et al., 1996; Yim et al., 1996). This peroxidase activity has not, however, been observed in all studies (Singh et al., 1998). Alternatively it has been suggested that mutant SOD1 causes oxidative stress by production of nitronium ions, which may then nitrate tyrosine residues (Beckman et al., 1994), and this may be due to the mutants’ decreased affinity for zinc (Crow et al., 1997). This is supported by the presence of free 3-nitotyrosine in both human FALS cases and in
40 mutant SOD1 mice (Beal et al., 1997; Ferrante et al., 1997). These nitrotyrosines appear not to be protein bound, however, which calls into question the significance of this finding (Bruijn et al., 1997a), although it has recently been found that free nitrotyrosine can induce apoptosis (Peluffo et al., 2004). However, both of the above oxidative damage hypotheses (enhanced peroxidase and nitration) require the presence of an active-site copper bound to SOD1, but MND still occurred in a transgenic mouse expressing a mutant in which all four copper-coordinating histidines were mutated
(Wang et al., 2003). Furthermore, the disease phenotype seen in several mutant SOD1 mouse models was unaffected when the copper chaperone for SOD1 (CCS; a protein that is essential for copper loading of SOD1) was eliminated (Subramaniam et al.,
2002).
Although the origins of oxidative stress in MND are not clear, the localisation of mutant
SOD1 in the intermembrane space (IMS) (Liu et al., 2004) and matrix (Vijayvergiya et al., 2005) of mitochondria could cause mitochondrial dysfunction (membrane depolarisation, decreased activity of respiratory complexes and Cytochrome c release), and this may contribute to oxidative stress pathways leading to neuronal death in FALS
(Beretta et al., 2003; Higgins et al., 2002; Jaarsma et al., 2001; Sturtz et al., 2001).
Mitochondrial dysfunction is also implicated in Hereditary spastic paraplegia, as two of the known mutations that cause the disease (SPG7/paraplegin and SPG13/HSP60) are found in genes encoding mitochondrial chaperone proteins, which are involved in protein folding in the IMS. Indeed, several patients with the SPG7/paraplegin mutation show signs of impaired mitochondrial oxidative phosphorylation, which is shown by the presence of abnormal mitochondria (ragged red fibers) and Cytochrome c oxidase deficient-fibers in muscle biopsies (Casari et al., 1998). Furthermore, mitochondrial abnormalities such as hypertrophy, concentric cristae, herniations and giant
41 mitochondria, found in the synaptic terminals of spinal cord motor neurons, is the earliest pathological feature seen in Paraplegin knockout mice (Gelbard, 2004).
Together these studies suggest an involvement of mitochondria in the pathogenesis of the MND.
1.1.4.3 Defects in axonal transport
The presence of IF accumulations in ALS has led to the proposal that NF transport is somehow perturbed, and subsequent studies in mouse models of MND have indicated that defects in slow axonal transport is one of the earliest pathological events in the disease (Williamson and Cleveland, 1999). In particular, axonal transport is abnormally reduced in mice with the SOD1 G93A, G37R and G85R mutations, and this precedes the onset of neuropathology (Borchelt et al., 1998; Warita et al., 1999; Williamson and
Cleveland, 1999; Zhang et al., 1997). Additionally, axonal transport defects have been observed in the Wobbler mouse (Mitsumoto et al., 1990; Mitsumoto and Gambetti,
1986), pmn mutant mouse (Sagot et al., 1998), hNF-H overexpressing mice (Collard et al., 1995) and the Paraplegin knockout mouse (Ferreirinha et al., 2004).
It has been hypothesised that NF accumulations in transport-deficient axons may impede further transport along microtubules, leading to cell death by ‘axonal strangulation’ (Williamson and Cleveland, 1999). Axonal transport defects have also been proposed to contribute to the dying back axonopathy observed in HSP due to impairment of efficient protein delivery to synaptic terminals. Furthermore, a lack of trophic support has been implicated in many forms of MND and this may also be caused by faulty transport mechanisms. Motor neurons are one of the largest cells in the nervous system (cell body diameter of 50-60 µm) with axonal processes of up to 1 metre in length and possess a high NF content due to their need for a robust cytoskeletal
42 network. Therefore, defects in axonal transport and abnormal NF accumulation could explain the selectivity of motor neuron death in MND. It is not known how the slowing of axonal transport in MND occurs, although several mechanisms have been proposed.
Role of hyperphosphorylation of NF subunits
Hyperphosphorylation of NF subunits has been proposed to play a role in deregulation of NF transport in MND. The mutations in NF-H that have been identified as a risk factor for SALS affect phosphorylation of side-arm domain Lys-Ser-Pro (KSP) repeat motifs (Al-Chalabi et al., 1999; Tomkins et al., 1998) and such KSP repeat phosphorylation is believed to be important in regulating NF transport. KSP repeats in
NF-M and NF-H are phosphorylated by multiple kinases such as glycogen synthase kinase-3α and -3β (Gsk3α/β), p42/p44 mitogen-activated protein kinases (p42/p44
MAPKs, also known as extracellular signal-related kinase (ERK2/ERK1), stress activated protein kinases (SAPKs) and the cyclin-dependent kinase-5/p35 complex
(Cdk5/p35) (Brownlees et al., 2000; Giasson and Mushynski, 1998; Guan et al., 1991;
Guidato et al., 1996; Sun et al., 1996; Veeranna et al., 1998). Indeed, studies in SOD1 mutant mice have shown that Cdk5 hyperphosphorylates NFs, leading to NF inclusions
(Nguyen et al., 2001) and SAPKs phosphorylate NF subunits in response to glutamate treatment, resulting in a slowing of NF transport, in cultured cortical neurons (Ackerley et al., 2000). However, recent studies using liquid chromatography tandem mass spectrometry (LC/MS/MS) have found that the phosphorylation of NF-H from sporadic
ALS spinal cord neurons does not differ from control samples (Strong et al., 2001), although it has been proposed that the levels of NF phosphorylation in the cell body compared with the axon may differ and this could affect the assembly and transport of
NFs (Miller et al., 2002; Strong et al., 2001).
43 Role of motor proteins
There is evidence to suggest that defects in the motor proteins which mediate axonal transport may be involved in the slowing of axonal transport in MND. Studies of green fluorescent protein (GFP)-tagged NF-M transport in cultured neurons have shown that transport is dependent on microtubules and the anterograde motor protein kinesin
(Koehnle and Brown, 1999; Yabe et al., 1999). Kinesin heavy chain mutations have been found to affect axonal transport in Drosophila (Hurd and Saxton, 1996) and more recently, mice lacking the neuronal-specific kinesin heavy chain KIF5A showed a defect in NF transport, with NF subunits accumulating in neuronal cell bodies (Xia et al., 2003). Furthermore, the transport of tubulin is impaired and axonal levels of kinesin are reduced in SOD1G85R mice months before disease onset (Warita et al., 1999; Zhang et al., 1997), levels of cDNA encoding the kinesin-like protein KIF3B are reduced in the motor neuronal cell line ‘NSC34’ stably-transfected with mutant SOD1 (Kirby et al.,
2002) and mutations in KIF1Bβ have been found to cause Charcot Marie Tooth disease type 2A (CMT2A) (Zhao et al., 2001a). Kinesin dysfunction has also been implicated in
HSP (see below; Axonal transport defects in Hereditary spastic paraplegia (HSP)).
Additionally, components of the retrograde motor protein complex (dynein/dynactin) have been found to associate with NFs and catalyse their transport in vitro (Shah et al.,
2000) and mutations in dynactin (p150 subunit) have been found to cause an autosomal dominant form of lower motor neuron disease in humans (Puls et al., 2003). As dynein knockout mutations are lethal in both mouse and Drosophila models, a transgenic mouse model with a targeted disruption of the dynein/dynactin complex was recently engineered to investigate dynein involvement in MND. This was achieved by overexpression of the dynamitin subunit of dynactin (which disassembles dynactin) in postnatal motor neurons, resulting in the development of late-onset progressive motor
44 neuron degeneration and muscle atrophy, thus confirming the critical role of axonal transport in the pathogenesis of MND (LaMonte et al., 2002). Furthermore, mouse models have been utilised to demonstrate that motor neurons are uniquely sensitive to disruption of dynein function and retrograde transport. Mice heterozygous for either of two ENU-generated mutations, Loa and Cra1, demonstrate a motor neuron degenerative phenotype similar to the dynamitin transgenic mouse, and positional cloning has revealed that both mutations are in the cytoplasmic dynein heavy chain 1 gene
(Dnchc1). Neither mutation seems to affect the localisation of dynein, or its expression levels, but its function is subtly inhibited, affecting motor neurons alone. Retrograde transport of a fluorescently-labelled fragment of Tetanus toxin was found to be significantly reduced in these mice, whereas other functions of dynein such as nuclear motility during cell division, and formation and positioning of the Golgi apparatus, were normal (Hafezparast et al., 2003b; He et al., 2005; Kieran et al., 2005).
Axonal transport defects in hereditary spastic paraplegia (HSP)
One autosomal recessive form of HSP is caused by loss-of-function mutations in the
SPG7 gene, which encodes the protein Paraplegin (a member of the ‘ATPases associated with a variety of cellular activities’ (AAA) family). Paraplegin knockout mice display an MND-like pathology and their neurons have large axonal swellings containing organelles and NFs, with impaired retrograde axonal transport (Ferreirinha et al., 2004). The mice also have abnormal mitochondria in spinal cord neurons, and this mitochondrial phenotype correlates with disease onset and neuronal degeneration, which has led to the proposal that mitochondrial dysfunction may underlie defective axonal transport. However, impairment of axonal transport in these mice is only seen after the onset of disease, which indicates that it may not be the primary cause of neurodegeneration (Ferreirinha et al., 2004). Mutations in the neuronal-specific kinesin
45 gene KIF5A have recently been found to cause a form of HSP (SPG10) in humans (Reid et al., 2002), which further implicates defective axonal transport in MND. SPG4/Spastin
(an AAA protein, but in a different subgroup to paraplegin) has been found to bind to microtubules, and has been proposed to act as a microtubule-severing enzyme (Charvin et al., 2003; Errico et al., 2002) which suggests that this form of HSP may involve cytoskeletal disassembly/disruption of transport. Interestingly, Spastin abnormally co- localises with kinesin when overexpressed in HEK cells and neurons (McDermott et al.,
2003) and recent studies using RNA interference (RNAi) in Drosophila melanogaster have revealed a role for Spastin in microtubule assembly in synaptic terminals (Trotta et al., 2004). A further microtubule-interacting protein, the novel GTPase Atlastin, is also mutated in juvenile-onset HSP (SPG3A) (Dalpozzo et al., 2003; Zhao et al., 2001b), although there is currently no evidence to suggest that this interaction has any effect on axonal transport. Atlastin shares homology with the Dynamin family of large GTPases, which are involved in molecular trafficking events such as synaptic vesicle recycling and mitochondrial dispersion (Jones et al., 1998; Nicoziani et al., 2000; Smirnova et al.,
1998).
1.1.4.4 Glutamatergic excitotoxicity
A role for glutamatergic excitotoxicity (a mechanism whereby prolonged or excessive exposure to extracellular glutamate leads to cell death) in MND was first implicated by observations of elevated glutamate levels in the cerebrospinal fluid (CSF) of SALS patients (Plaitakis and Caroscio, 1987; Rothstein et al., 1990; Shaw et al., 1995a). The level of neuronal excitation by glutamate is regulated by a number of mechanisms. Two of relevance to MND are NMDA receptor and AMPA/kainate receptor inactivation and
Na+/K+-coupled glutamate reuptake by astrocytic transporter proteins (excitatory amino acid transporters; EAAT1-5, also known as glutamate transporters; GLT1-5). There is
46 extensive evidence to suggest an involvement of both mechansims in the pathogenesis of ALS, which is interesting as it highlights the importance of both motor neurons and astrocytes in the disease, as discussed in section 1.1.4. The precise molecular mechanisms that lead to excitotoxicity-mediated cell death are not known, although several pathways have been identified that contribute, including disruption of intracellular calcium homeostasis and production of free radicals (For a recent review see (Heath and Shaw, 2002)). Additionally, the motor neurons that are preferentially affected in MND are likely to be particularly sensitive to excitotoxic insults for several reasons, including a high expression of AMPA receptors lacking the GluR2 subunit
(this makes them highly calcium-permeable) and a low level of calcium-binding proteins such as calbindin D-28k and parvalbumin, compared with the neuronal populations that are frequently spared in MND such as the oculomotor, trochlear, abducens nerve and Onuf’s nucleus motor neurons (Alexianu et al., 1994; Elliott and
Snider, 1995; Ince et al., 1993; Reiner et al., 1995). Indeed, overexpression of parvalbumin in a transgenic mouse model of ALS delayed the disease onset (Beers et al., 2001). Together, these findings support a role for excitotoxicity and disrupted calcium homeostasis in MND.
Role of glutamate transporters
Astrocytes are the principle regulators of extracellular glutamate levels (Rothstein et al.,
1996), and deficiency in glutamate uptake by astrocytes appears to play a crucial role in the pathogenesis of ALS. The exact cause of glutamate transport deficiency in ALS is not known, although it has been postulated that the reduction in transport function is caused by a loss of the astrocytic glutamate transporter EAAT2 (also known as GLT1)
(Rothstein et al., 1992). In 60-70% of SALS cases, and in SOD1 mutant mouse models, a large reduction in EAAT2 expression (which is not purely due to cell death) is seen at
47 the end-stage of disease (Bristol and Rothstein, 1996; Bruijn et al., 1997b; Rothstein et al., 1995). EAAT2 reduction has also been observed at early stages of disease (before the onset of hind-limb paralysis) in a mutant SOD1G93A rat model and this loss is specific to areas of the spinal cord that contain motor neuron cell bodies (Howland et al., 2002). Furthermore, EAAT2 knock-down, obtained by administration of antisense oligonucleotides in vitro and in vivo, resulted in progressive hind-limb paralysis and motor neuron degeneration in rats (Rothstein et al., 1996). It has been suggested that abnormal mRNA splicing of EAAT2 could underlie reduced expression and function of the protein in ALS (Lin et al., 1998), although several other groups have failed to replicate these findings (Aoki et al., 1998; Flowers et al., 2001). However, no motor neuron loss is seen in EAAT2 knockout mice (although they develop hippocampal pathology and seizures, and most die as juveniles) (Tanaka et al., 1997). This suggests that an overall loss of EAAT2 function does not lead to MND pathology. Additionally, no reduction in EAAT2 expression has been observed in the brain or spinal cord of mice overexpressing SOD1G93A (Deitch et al., 2002). Several studies suggest that the reduction in EAAT2 function may be caused by biochemical changes in transporters already present rather than differences in transporter levels. For example, studies in
Xenopus oocytes have shown that the oxidative activity of mutant SOD1 leads to a reduction in EAAT2 function, and this effect can be blocked by the antioxidant
Mn(III)TBAP (Trotti et al., 1999). This suggests that EAAT2 may be susceptible to oxidative damage, resulting in decreased glutamate uptake at the synaptic cleft, and excitotoxic damage to motor neurons. Additionally, a mutation in the EAAT2 gene which results in impaired glutamate clearance capacity has also been reported in a single SALS case (Trotti et al., 2001).
48 Role of glutamate receptors
AMPA receptors are composed of different combinations of four subunits GluR1-4 (or
GluRA-D); the GluR2 (GluRB) subunit is of particular importance in determining the calcium permeability of the assembled receptor. An mRNA-editing defect of GluR2 in spinal cord motor neurons from 5 individual SALS cases has recently been reported, which confers increased calcium-permeability on AMPA receptors, leading to cell death
(Kawahara et al., 2004). Therefore, the presence/function of the GluR2 subunit and the calcium-permeability of AMPA receptors are believed to play a significant role in excitotoxic pathways in MND. The effect of a reduction in the calcium-permeability of
AMPA receptor in motor neurons has recently been investigated in a mutant SOD1 mouse model. This was achieved by crossing SOD1G93A overexpressing mice with a mouse line overexpressing GluR2 specifically in cholinergic neurons (resulting in a large reduction in the calcium-permeability of motor neuronal AMPA receptors). These mice displayed a delay of disease onset with a correlating delay in the formation of abnormal intracellular SOD1 aggregates, as compared with mice overexpressing
SOD1G93A alone (Tateno et al., 2004), suggesting that the calcium-permeability of
AMPA receptors specifically in motor neurons affects the formation of SOD1 aggregates, leading to MND, in this model. Additionally, the amount of carbonylated proteins (a marker of oxidative stress) in the spinal cord was also reduced/delayed in these mice, and it has been proposed therefore that the protein aggregates seen in the disease may be a consequence of ROS production (Tateno et al., 2004). Complimentary to these findings was a recent study in which a mouse line was created that overexpressed a functionally-modified GluR2 subunit (GluRB(N)), in which the subunit conferred calcium-permeability on its assembled AMPA receptor but the conductivity of the receptor was unaffected. These mice displayed a phenotype of progressive MND that closely resembles human SALS, and crossing with SOD1G93A transgenic mice
49 resulted in an acceleration of disease progression and decrease in survival, which further supports a role for an increase in calcium permeability caused by defective GluR2 mRNA editing in MND (Kuner et al., 2005).
Involvement of oxidative stress in excitotoxicity
Glutamate transporters are susceptible to oxidative damage, and oxidative modifications of transporter peptides have been reported both in ALS cases and in SOD1 mutant mice
(Liu et al., 2002; Pedersen et al., 1998), indicating that the reduction of transporter function in ALS may be caused by oxidative mechanisms. The presence of signs of mitochondrial degeneration in both human ALS cases and in SOD1 mutant mouse models (in which this precedes the onset of motor defects) provides a link between oxidative stress and excitotoxic mechanisms. It has been proposed that lack of mitochondrial function, coupled with the high energy demands of motor neurons, may result in a lowering of the neuron’s membrane potential, resulting in opening of glutamate receptors and influx of calcium, and less glutamate would then be needed to have an excitotoxic effect on the cell. However, a lack of involvement of oxidative stress in downregulation of EAAT2 has been reported in astrocytes (Tortarolo et al.,
2004).
Riluzole and evidence against the excitotoxicity hypothesis
In support of the glutamatergic excitotoxicity hypothesis, Riluzole, a drug that inhibits glutamatergic transmission, has been shown to delay disease progression in several forms of ALS (Bensimon et al., 1994; Lacomblez et al., 1996). However, the failure of several other antiglutamatergic agents, for example Gabapentin, in human ALS trials
(Miller et al., 1996; Miller et al., 2001) suggests that Riluzole may have a novel mode of action. Additionally, a recent study has demonstrated a lack of glutamatergic
50 involvement in ALS, using an in vivo rat spinal cord microdialysis model, and has postulated that large and long-lasting increases of glutamate in the spinal cord do not produce motor neuron hyperexcitation or degeneration (Corona and Tapia, 2004).
1.1.4.5 Neuroinflammation and autoimmunity
Neuroinflammation has been implicated in ALS due to several findings, including upregulation of the pro-inflammatory enzyme cyclooxygenase-2 (COX-2) mRNA and protein expression, and increased COX-2 activity (measured by increased prostaglandin
E2 levels) in SALS spinal cord (Maihofner et al., 2003; Yasojima et al., 2001) and in mutant SOD1 transgenic mice (Almer et al., 2001). Furthermore, a selective COX-2 inhibitor, Celecoxib, protects spinal cord motor neurons from mutant SOD1-mediated cell death by prolonging survival and protecting against microglial activation, astrogliosis and spinal cord neuron degeneration (Drachman et al., 2002; Pompl et al.,
2003).
Inflammatory responses also recruit immune mechanisms, the main effectors of which
(astrocytes and microglia) are activated in ALS (Hall et al., 1998; Kamo et al., 1987;
Kawamata et al., 1992; Schiffer et al., 1996). Indeed, microglial activation occurs before disease onset in mutant SOD1 transgenic mice (Alexianu et al., 2001) and the anti- inflammatory compound minocycline extends survival in mouse models of ALS (Kriz et al., 2002; Van Den Bosch et al., 2002; Zhu et al., 2002), which may occur by inhibiting microglial activation (Tikka et al., 2001; Tikka et al., 2002). Additionally, infiltration of macrophages, mast cells and T cell lymphocytes have been found in
SALS spinal cord (Graves et al., 2004; Hayashi et al., 2001; Kawamata et al., 1992;
Lampson et al., 1990), which suggests an involvement of both innate and acquired immune responses in MND. Autoantibodies against various components in the CNS
51 have been reported in ALS, although it is still unclear whether this represents an involvement of autoimmunity in the pathogenesis of MND, or if such antibodies have a beneficial effect in the disease. Examples of antibodies found in SALS sera include neuronal, NF subunit and calcium channel antibodies (Couratier et al., 1998; Kimura et al., 1994; Niebroj-Dobosz et al., 1999; Smith et al., 1992). Monoclonal immunoglobulin
G (IgG) is also detectable in SALS spinal cord, motor cortex and sera (Duarte et al.,
1991; Engelhardt and Appel, 1990). However, anti-inflammatory and immunosuppressive therapies have had little beneficial effect in ALS to date (Drachman et al., 1994).
1.1.4.6 Apoptosis
There is evidence to suggest that neuronal death in SOD1-mediated FALS occurs via apoptotic signalling mechanisms, which is implicated by the presence of DNA fragmentation, decreased expression of the anti-apoptotic protein Bcl-2, and increased expression of the pro-apoptotic protein Bax, in the spinal cords of ALS patients and of transgenic mutant SOD1 mice (Fujita et al., 2002; Martin, 1999; Vukosavic et al., 1999;
Yoshiyama et al., 1994). Additionally, expression of Bcl-2 in SOD1G93A mutant mice delays disease onset and increases survival (Kostic et al., 1997) and mutant SOD1 binds to Bcl-2 in mitochondria, which suggests that mutant SOD1 may sequester Bcl-2 into abnormal aggregates and thereby prevent its ability to participate in anti-apoptotic pathways (Pasinelli et al., 2004). Caspase inhibitors have also been effective in delaying onset and prolonging life (Li et al., 2000). Indeed, sequential activation of caspase-1 and caspase-3 (which belong to the family of apoptotic effector cysteine proteases) (Villa et al., 1997) in mutant SOD1 mouse spinal cord motor neurons and astrocytes coincides with onset of motor neuron loss (Li et al., 2000; Pasinelli et al., 2000; Vukosavic et al.,
1999).
52 Three major apoptotic pathways have been the most widely-studied; the mitochondrial
(intrinsic) pathway, the death receptor (extrinsic) pathway and the endoplasmic reticulum (ER) pathway. It has been shown that Cytochrome c translocation from mitochondria to the cytosol, and Bax translocation from the cytosol to mitochondria, which are crucial steps in the mitochondrial-dependent apoptotic pathway (Kroemer and
Reed, 2000), occurs in the spinal cord of SOD1G93A transgenic mice in parallel with neurodegeneration (Guegan et al., 2001). Additionally in these mice, activation of caspase-9 followed by caspase-7, and cleavage of the X chromosome-linked inhibitor of apoptosis protein (XIAP) was reported, further supporting a role of the mitochondrial apoptotic pathway. Involvement of the death receptor pathway has also been identified in MND. Nitric oxide signalling via the death receptor Fas leads to activation of caspase-dependent specific death of embryonic spinal cord motor neurons in culture
(Raoul et al., 2002) and another death receptor p75NTR has also been implicated in cell death in mutant SOD1 mice (Kust et al., 2003; Turner et al., 2003).
Although it remains unknown how apoptosis may be triggered in MND, several hypotheses exist. Reactive astrocytes have been shown to promote apoptosis of motor neurons in culture via a nitric oxide and peroxynitrite-dependent mechanism (Cassina et al., 2002) and via nerve growth factor (NGF) in p75NTR-expressing neurons (Pehar et al., 2004). It is also widely believed that mitochondrial dysfunction plays a major role, particularly as mitochondrial abnormality, vacuolation and swelling is observed in ALS patients and FALS mouse models (Dal Canto and Gurney, 1995; Kong and Xu, 1998;
Wong et al., 1995), and mutant SOD1 localises to spinal cord and brain mitochondria
(Liu et al., 2004; Vijayvergiya et al., 2005), where it impairs Cytochrome c association with the inner mitochondrial membrane and leads to apoptosis (Kirkinezos et al., 2005).
53 1.1.4.7 Involvement of cell signalling pathways
Protein kinases
Abnormalities in the activity and/or expression of several kinases have been reported in
ALS post-mortem tissue, including protein kinase C (PKC), phosphatidylinositol 3- kinase (PI(3)-K), Cdk5 and SAPK (Bajaj et al., 1998; Krieger et al., 1996; Lanius et al.,
1995; Migheli et al., 1997; Nagao et al., 1998; Wagey et al., 1998). Furthermore a recent study, using a quantitative proteomics screening technique on human thoracic spinal cord samples from ALS patients and controls, has identified increased expression of several kinases in ALS (Hu et al., 2003b), including PKC, ERK2 and its putative downstream target ribosomal S6 kinase 1 (RSK1), phosphorylated/activated p38 MAPK
(p38), protein kinase B (PKB; also known as Akt), SAPK and Cdk5.
Mutant SOD1 transgenic mice have been useful in studying Cdk5-mediated cell death in
MND. In SOD1G37R mice, mislocalisation and hyperactivation of Cdk5 has been observed, which correlates with increased production of a truncation product (p25) of its activator p35 (Nguyen et al., 2001). The ways in which Cdk5/p25 activity may cause neurodegeneration are unknown, although aberrant phosphorylation of substrates such as NFs and tau, leading to cytoskeletal alterations/defects in axonal transport, has been suggested as a potential event leading to cell death. Additionally, Cdk5 is involved in several cellular processes, including cell adhesion and synaptic signalling (Bibb et al.,
2001; Kwon et al., 2000), alterations in both of which could contribute to neurodegeneration. Interestingly, mice overexpressing Cdk5 and p35 do not show a profound disruption of the cytoskeleton (Van den Haute et al., 2001), unlike mice overexpressing p25 (Ahlijanian et al., 2000), which suggests that p25 is the toxic mediator of events leading to neurodegeneration. Oxidative stress has also been found to promote the generation of p25 from p35 through activation of the Ca2+-dependent
54 protease calpain, resulting in increased Cdk5 activity, increased phosphorylation of NF subunits, and inhibition of axonal transport (Lee et al., 2000; Shea et al., 2004).
However, transgenic mice overexpressing SOD1G93A in a p35-null background are not phenotypically different from mice overexpressing SOD1G93A alone (Takahashi and
Kulkarni, 2004), which suggests that the production of p25 from p35 and the activation of Cdk5 by p35 is not involved in the SOD1G93A-mediated disease seen in these mice.
Therefore, the precise role of Cdk5/p25 activity in ALS is currently unclear.
The activity of p38 is increased in SOD1G93A mouse spinal cord (neurons, astrocytes and microglia), and this occurs before disease onset and correlates with disease progression, although no alterations in its activity have been observed in human ALS tissue compared with controls (Hu et al., 2003a; Hu et al., 2003b; Tortarolo et al., 2003). p38 has been implicated in exitotoxicity, as inhibitors of p38 have been found to rescue cells from glutamate-induced cell death (Kawasaki et al., 1997), and it is also involved in phosphorylation of cytoskeletal proteins and modulating the expression of cytokines, nitric oxide and COX-2 (Ackerley et al., 2004; Guan et al., 1998; Mielke and Herdegen,
2001; Ono and Han, 2000). Although active p38 plays a role in certain apoptotic pathways (Kummer et al., 1997) the p38 activation seen in the SOD1G93A mouse is not thought to lead to apoptosis, due to the lack of ultrastructural apoptotic features in nuclei, DNA fragmentation and activated caspase-3 immunostaining in these mice
(Bendotti et al., 2001; Migheli et al., 1999; Tortarolo et al., 2003).
L1CAM
Neuronal cell adhesion molecule L1 (L1CAM) is mutated in an X-linked form of HSP
(SPG1) and plays an important role in cell recognition and signalling (Jouet et al.,
1994). The L1CAM protein is a member of the immunoglobulin superfamily of cell
55 adhesion molecules and is found primarily in the nervous system where it is involved in cellular processes such as neurite/growth cone guidance and neuronal migration during development, and cell survival (Castellani et al., 2000; Chen et al., 1999; Dahme et al.,
1997; Fransen et al., 1998).
ALS2
Amyotrophic lateral sclerosis 2 (ALS2)/alsin was first identified in 2001 by two independent research groups as a protein that is mutated in rare autosomal recessive juvenile forms of ALS (ALS2) and PLS (jPLS) (Hadano et al., 2001; Yang et al., 2001), and has subsequently been found to cause infantile-onset HSP (IAHSP) (Devon et al.,
2003; Eymard-Pierre et al., 2002; Gros-Louis et al., 2003b). Although the exact function of ALS2 is currently unknown, it shares homology with several GTPase-regulating proteins (guanine nucleotide exchange factors; GEFs). It is predicted, therefore, to be involved in various critical cellular processes such as signal transduction, regulation of the cytoskeleton and intracellular trafficking, as discussed below.
1.2 Alsin/ALS2
Mutations in the gene ALS2, on chromosome 2q33.2, have been found that cause forms of infantile/juvenile MND with similar clinical phenotypes involving UMN degeneration: ALS (juvenile ALS; ALS2) (Hadano et al., 2001; Yang et al., 2001), PLS
(juvenile PLS; jPLS) (Yang et al., 2001) and HSP (Infantile-onset ascending HSP;
IAHSP) (Devon et al., 2003; Eymard-Pierre et al., 2002; Gros-Louis et al., 2003b).
However, there are as yet no reports of pathology in the affected individuals. Of the nine ALS2 mutations described so far, eight are small deletions and one is a nonsense mutation. All mutations discovered so far are in coding regions, are inherited in an autosomal recessive pattern, and are predicted to result in premature truncation and a
56 loss of function (Figure 1.1). There is no apparent correlation between the site of mutation and the disease phenotype (Devon et al., 2003). It has been found that at least six of the disease mutant proteins are unstable and rapidly degraded, including the mutation predicted to result in a C-terminal truncation of only 28 amino acid residues
(4844delT), therefore it may be speculated that the disease is caused by a loss of the whole protein, and any of its functional domains could potentially regulate its normal function (Yamanaka et al., 2003).
Human ALS2 comprises 34 exons (33 of which contain coding sequence) over a region of 6.5 kb and encodes the novel 184 kDa protein ALS2 (also known as Alsin). An additional transcript of 2.6 kb (predicted to be derived from alternative splicing), known as ALS2 ‘short-form’, has been detected by Northern blot (Hadano et al., 2001), although as yet no 44 kDa protein corresponding to the short form has been detected either in mouse or human tissue (Otomo et al., 2003). Furthermore, short-form ALS2 was found to be unstable and rapidly degraded when over-expressed in a human cell line (Yamanaka et al., 2003). Recently, a novel ALS2 homologous gene ALS2 carboxyl- terminal like (ALS2CL) on human chromosome 3p21.3, which encodes a 108 kDa protein, has been described (Devon et al., 2005; Hadano et al., 2004).
The function of ALS2 is unknown, although sequence analysis has revealed 3 domains that are homologous to guanine nucleotide exchange factors (GEFs), which suggests involvement in Ras superfamily signalling pathways (see section 1.3). The 3 GEF domains include an amino (N)-terminal regulator of chromatin condensation (RCC1)- like domain (putative Ran GEF), a diffuse B-cell lymphoma (Dbl) homology (DH) domain followed by a pleckstrin homology (PH) domain (which is a hallmark of Rho
GEFs), and a carboxyl (C)-terminal VPS9 domain (putative Rab GEF). There are also 8
57 copies of a sequence motif called membrane occupation and recognition nexus
(MORN), which may be involved in recruitment to membranes (Takeshima et al.,
2000). ALS2CL protein shows high amino acid sequence similarity to the C-terminal region of ALS2, but lacks certain residues corresponding to the RCC1-like domain
(Hadano et al., 2004).
1.2.1 Expression of ALS2
ALS2 mRNA distribution in various human tissues has been analysed using RT-PCR
(reverse transcription-polymerase chain reaction) (Yang et al., 2001) and Northern blot
(Hadano et al., 2001). Both long- and short-form ALS2 mRNA has been detected in heart, placenta, lung, liver, spleen, skeletal muscle, kidney and pancreas, with the highest expression seen in brain and spinal cord. Analysis of different brain regions reveals highest expression in cerebellum and cerebral cortex (Hadano et al., 2001).
Immunoblotting and immunohistochemistry shows that the pattern of ALS2 long-form protein expression is similar to that of its mRNA, with the highest-expressing tissues being brain (particularly cerebellum and cerebral cortex), spinal cord and liver, in both mouse and human (Devon et al., 2005; Otomo et al., 2003; Yamanaka et al., 2003).
Additionally, the use of neuron- and glia-specific markers (Neu-N and glial fibrillary acidic protein (GFAP) respectively) has revealed that ALS2 is expressed in various neurons, but not in glial cells (Devon et al., 2005; Otomo et al., 2003). ALS2 is believed to be expressed at generally low levels, however, representing only approximately
0.0003% of the total detergent soluble fraction of mouse brain lysate (Yamanaka et al.,
2003).
58 Figure 1.1 Schematic of ALS2 (Long-form and Short-form) and predicted disease mutants Nine ALS2 mutations have been described to date, all of which result in a premature stop codon and a predicted truncated protein.
ALS2 Long-form
RCC1 DH PH MORN VPS9
1 59 167 218 576 627 690 885 1049 1244 1551 1657 169 525 578 9011657 1007 1656
ALS2 Short-form
Unique 24 residues
1 372 396
ALS2 disease mutants (predicted proteins) Disease and Reference
261delA ALS2 (Tunisia) Unique 3 (Hadano et al., residues 2001)
1 49 1130delTA IAHSP (Italy) (Eymard-Pierre et al., 2002) 1 335
1548delAG Unique 70 ALS2/jPLS (Kuwait); residues (Hadano et al., 2001; Yang et al., 2001)
1 475 545
1594delGTTTCCCCCA IAHSP (France) (Eymard-Pierre et al., 2002) 1 493
1990delCT Unique 25 jPLS (Saudi) residues (Yang et al., 2001)
1 620 645
2660delCT IAHSP (Italy) (Eymard-Pierre et al., 2002) 1 858
59 C3115T IAHSP (Israel) (Devon et al., 2003)
1 998
3742delA IAHSP (Algeria) (Eymard-Pierre et al., 2002)
1 1206
4844delT IAHSP (Pakistan) (Gros-Louis et al., 2003b)
1 1573
1.2.2 ALS2 exhibits GEF activity
ALS2 has an amino-terminal domain that is homologous to RCC1, a known GEF for the
GTPase Ran, and ALS2 was found to weakly stimulate GDP dissociation from Ran in an in vitro assay, although preliminary experiments with the isolated RCC1-like domain were contradictory (Otomo et al., 2003). It has been suggested that it is unlikely that
ALS2 acts as a Ran GEF in vivo (Topp et al., 2004), as although there are more than 90 proteins with RCC1 domains on the available databases (Bateman et al., 2002), only
RCC1 itself exhibits Ran GEF activity (Bischoff and Ponstingl, 1991). Thus, the Ran
GEF activity of ALS2 remains unclear at this stage. Aside from a role as a GEF, the
RCC1 domain may function as a protein-protein interaction domain, since it has the potential to form a seven-bladed beta-propeller structure (Topp et al., 2004).
The presence of a DH and C-terminally adjacent PH domain is an indication that ALS2 may function as a GEF for the Rho family of GTPases, as this tandem repeat is found in the majority of Rho family GEFs (Hart et al., 1991; Yaku et al., 1994) (see section
1.4.1). In vitro binding assays have shown that a fragment of ALS2 containing only the
DH/PH domain has the ability to bind to Rac1 but not to Rac3, RhoA or Cdc42 (Topp et al., 2004), which has led to the investigation of whether ALS2 can function as a GEF
60 for Rac1 in vivo. When overexpressed in S. frugiperda (sf9) cells with Rac1, ALS2 promoted an increase in the level of GTP-bound Rac1 compared to control conditions
(Topp et al., 2004). In NIH3T3 cells endogenous ALS2 co-localised with Rac1 at leading membrane edges, and overexpressed GFP-tagged ALS2 co-localised with actin in membrane ruffles and lamellipodia, although overexpression did not seem to stimulate these events (Topp et al., 2004), which further implies a role for endogenous
ALS2 as Rac GEF. Furthermore, overexpression of ALS2 was found to increase the activity of endogenous Rac in CHO cells whereas a construct in which the DH/PH was mutated did not (These studies and Kanekura et al 2005). In contrast, ALS2 was not found to act as a Rac GEF in in vitro GDP dissociation assays (Otomo et al., 2003).
ALS2 contains a C-terminal VPS9 domain, which is homologous to Rab5 GEF domains. All VPS9 domain-containing proteins that have been studied so far exhibit
Rab5 GEF activity (Hama et al., 1999; Horiuchi et al., 1997; Saito et al., 2002; Tall et al., 2001) and this includes ALS2 (Otomo et al., 2003; Topp et al., 2004). In vitro GDP dissociation assays were used to test the Rab5 GEF activity of ALS2, and to identify the regions of the protein necessary for this activity (Otomo et al., 2003; Topp et al., 2004).
It was found that ALS2 activates Rab5a, Rab5b and Rab5c but not other Rab family members (Otomo et al., 2003), and that the minimal region necessary for this activity is the N-terminal region containing the VPS9 domain and the MORN motifs. In support of this data, in vitro binding assays using ALS2 immunoprecipitated from both CHO and
SH-SY5Y cells have shown that ALS2 binds to Rab5a (Otomo et al., 2003; Topp et al.,
2004). Furthermore, yeast 2-hybrid analysis using the VPS9 domain of ALS2 fused to the Gal4 activation domain (as ‘prey’) with various Rab5 LexA DNA binding domain fusions (‘baits’), revealed that ALS2 interacts with GDP-bound or nucleotide-free Rab5
(S34N; a mutant that cannot bind GTP) but not wild-type (GTP-bound) Rab5 (Topp et
61 al., 2004). ALS2 (endogenous and transfected) has been found to form homo-oligomers
(presumably octamers, based on data from gel exclusion assays) in COS-7 cells, and mutants which have lost the ability to form such oligomers also display a loss of Rab5
GEF activity in GDP dissociation assays (Kunita et al., 2004).
1.2.3 ALS2 regulates endosomal morphology
Endogenous ALS2 has been shown to localise to cytoplasmic punctuate membrane structures in rat embryonic hippocampal neurons (Topp et al., 2004) and biochemical analysis of rat cortex has revealed that endogenous ALS2 is found in the cytosolic and membrane fractions (co-precipitating with the recycling endosome marker transferrin receptor and the early endosome marker early endosome antigen 1 (EEA1)), where it is peripherally associated with endosomal membranes (Devon et al., 2005; Topp et al.,
2004; Yamanaka et al., 2003). Immunocytochemistry has also shown that overexpressed
ALS2 co-localises with transfected Rab5A and endogenous EEA1 in HeLa cells and cultured rat embryonic cortical neurons (Otomo et al., 2003). The localisation of ALS2 demonstrated in the above research is in agreement with the findings that ALS2 can act as a GEF for Rab5, as this GTPase is involved in protein trafficking through early endosomes (see section 1.3.2). Indeed, co-transfection of ALS2 (or the VPS9 domain of ALS2) and Rab5A in cortical neurons (or NIH3T3 cells) resulted in enlarged endosomes, which is similar to the effect seen upon overexpression of a constitutively active Rab5 (Stenmark et al., 1994), whereas constructs with mutations in the VPS9
(Rab5GEF) domain did not, implying a role for ALS2 in Rab5-mediated promotion of endosome fusion/enlargement (Otomo et al., 2003; Topp et al., 2004). Furthermore, a construct encompassing the DH/PH, MORN and VPS9 domains (ALS2660-1657) was found to act in a constitutively active manner by causing dramatic endosome enlargement and this was dependent on its ability to form homo-oligomers (Kunita et
62 al., 2004; Otomo et al., 2003).
Interestingly, Spartin, a protein that is mutated in a form of complicated HSP (Troyer syndrome), may be involved in endosome morphology and molecular trafficking events
(Patel et al., 2002; Bakowska et al., 2005), and mutations have recently been discovered in vesicle-associated membrane protein (VAMP)/synaptobrevin-associated protein B
(VAPB), which is involved in intracellular membrane trafficking, in seven families with diagnoses ranging from ALS (ALS8) to SMA (Nishimura et al., 2004).
1.2.4 ALS2 binds to mutant SOD1 and displays neuroprotective activity
The relationship between ALS2 and the first protein to be identified as causative for
MND, SOD1, has been investigated in the hope of identifying common mechanisms in disease pathogenesis. Overexpression of ALS2 long-form (but not ALS2 short-form) in the motor neuron-like cell line NSC34 delays cell death caused by SOD1A4T, SOD1G85R or SOD1G93R (as observed by trypan blue exclusion assay), and this neuroprotection is specific for mutant SOD1-mediated cell death, as ALS2 has no effect on toxicity caused by mutants of presenilin1 and 2 (PS1M146L and PS2N141I), amyloid beta precursor protein
(AβPPV642I) or α-synucleinA53T (proteins involved in familial Alzheimer’s disease and familial Parkinson’s disease) (Kanekura et al., 2004). Interestingly, ALS2 long-form has the ability to bind specifically to the aforementioned SOD1 disease mutants but not to wild-type SOD1, whereas ALS2 short-form can bind to wild-type SOD1 but not to the
SOD1 mutants (Kanekura et al., 2004). The region responsible for both the neuroprotective activity of ALS2 long-form and its binding to mutant SOD1 has been found to reside in the DH/PH (Rho family GEF) domain (Kanekura et al., 2004).
Moreover, the neuroprotective activity of ALS2 can be blocked by reduction of Rac1 expression (using small interfering RNA (siRNA) for Rac1) which further supports the
63 hypothesis that ALS2 functions as a Rac GEF in this pathway. Rac has previously been shown to promote cell survival in COS-7 cells by activation of PI(3)K and Akt (Murga et al., 2002), and it has been shown that PI(3)K and Akt family proteins may be involved in ALS2-mediated neuroprotection against mutant SOD1. Indeed, treatment with wortmannin (a PI(3)K inhibitor) and reduction of Akt3 expression (using siRNA) independently abolished ALS2 neuroprotection (Kanekura et al., 2005). Additionally, overexpression of ALS2 but not a mutant without Rac GEF activity (ALS2T701A) resulted in increased phosphorylation of Akt3. Therefore, these findings suggest that
ALS2 mediates neuroprotection specifically against mutant SOD1 by sequential activation of Rac1, PI(3)K and Akt3, although whether this function of ALS2 occurs in neuronal cells and by the endogenous protein remains to be discovered.
1.3 The Ras Superfamily of GTPases
Members of the Ras superfamily of small guanosine triphosphatases (small GTPases; also known as ‘small G proteins’) are monomeric proteins that bind and hydrolyse guanine nucleotides (Bourne et al., 1991). By cycling between guanosine triphosphate
(GTP)-bound (active) and guanosine diphosphate (GDP)-bound (inactive) conformations, Ras GTPases act as molecular switches, controlling a wide range of intracellular signalling pathways in all eukaryotes. To date, more than 80 Ras superfamily members have been identified in mammals (Scita et al., 2000), and are divided into five families: Ras, Rho, Rab, Arf and Ran (see Table 1.4), all of which possess a common structurally-conserved GTP-binding region known as the ‘G domain’
(Bourne et al., 1990; Valencia et al., 1991).
64 Table 1.4 Mammalian Ras GTPase Superfamily
RAB FAMILY RAS FAMILY RHO FAMILY ARF FAMILY RAN FAMILY Rab1A, B Ha-Ras RhoA, B, C, D Arf1- 6 Ran Rab2A, B Ki-Ras RhoE/Rnd3/Rho8 Sar1a, b Rab3A, B, C, D N-Ras RhoG Arl1- 7 Rab4A, B, C R-Ras RhoH/TTF Ard1 Rab5A, B, C M-Ras Rac1, 2, 3 Rab6A, B, C RalA, B Cdc42 Rab7 Rap1A, B Rnd1/Rho6 Rab8A, B Rap2A, B Rnd2/Rho7 Rab9A, B, C TC21 TC10 Rab10 Rit TCL Rab11A, B Rin Wrch1 Rab12-15 Rad Chp/Wrch2 Rab17, 18 Kir/ Gem Rif Rab20, 21 Rheb RhoBTB1, 2 Rab22A, B, C κB-Ras1, 2 MIRO-1, 2 Rab23 Rab25, 26 Rab27A, B Rab28A, B Rab30 Rab32A, B Rab34-39 Rab40A, B, C Rab41-43
(Burridge and Wennerberg, 2004; Stenmark and Olkkonen, 2001; Takai et al., 2001; Wherlock and Mellor, 2002)
Ras itself regulates cell growth and differentiation by activation of the MAPK cascade, whereas the other superfamily members are involved in a variety of cellular events. The
Rho (in particular Rho, Rac and Cdc42) and Rab (in particular Rab5) family GTPases, which are implicated in ALS2 function, will be discussed below.
65 The activity (nucleotide binding state) of GTPases is highly regulated. In contrast to the moderate hydrophobicity of other Ras superfamily members, Rho and Rab family
GTPases contain highly hydrophobic geranylgeranyl moieties and therefore the association of guanine nucleotide dissociation inhibitors (GDIs) is needed to stabilise
GDP-binding and sequester the inactive GTPase to the cytoplasm. Indeed, the majority of GTPase in the cell is found in the inactive GDP-bound form, associated with GDIs in the cytoplasm. An additional regulating protein known as a GDI-displacement factor
(GDF) is required for the correct membrane localisation of Rab GTPases. The active state is promoted by guanine nucleotide exchange factors (GEFs) which tether the
GTPase to a particular subcellular localisation and promote GDP release (this results in
GTP binding due to a higher intracellular level of GTP than GDP). The active state is then negatively regulated by GTPase activating proteins (GAPs), which catalyse the intrinsic ability of the GTPase to hydrolyse GTP to GDP (Figure 1.2) (Moon and
Zheng, 2003; Schmidt and Hall, 2002; Zheng, 2001)
Figure 1.2 Schematic diagram of the small GTPase activation cycle
Activation Signal
GDP GTP
GDI GEF
GDP GTP
GTPase GTPase
GDI Effector GAP GDI Cytoskeletal organisation; Apoptosis; Gene expression; Cell cycle Pi progression; Membrane traffic; Cell migration; Vesicular & nuclear transport
66 1.3.1 Rho family GTPases
Rho family GTPases are ubiquitously expressed and so far 21 genes have been identified in humans, encoding at least 23 signalling proteins (Wherlock and Mellor,
2002). More than 40 effectors, 60 GEFs and 40 GAPs have been described for the mammalian Rho family (Raftopoulou and Hall, 2004). Rho family GTPases are key regulators of multiple cellular activities including regulation of the cytoskeleton and cell adhesion, cell polarity, cell cycle progression, apoptosis, neuronal axon guidance, differentiation, oncogenesis and gene transcription.
The best characterised function of Rho family GTPases is the control of signal transduction pathways linking membrane receptors to regulation of the actin cytoskeleton (for reviews see (Van Aelst and D'Souza-Schorey, 1997; Hall, 1998)).
Such research has focused on the roles of three Rho GTPase family members, Rho (A,
B and C), Rac (1 and 2) and Cdc42, in the spatiotemporal control of cytoskeletal dynamics. Although structurally related, Rho, Rac and Cdc42 have been shown to have distinct effects on the actin cytoskeleton. In classic studies using quiescent Swiss 3T3 fibroblasts (a cell line in which serum starvation results in a very low background of filamentous (F)-actin), Rho induces stress fiber formation and clustering of integrins into focal adhesion complexes (Ridley and Hall, 1992), Rac promotes actin polymerisation at the cell periphery to form lamellipodia and membrane ruffles (Ridley et al., 1992), and Cdc42 induces the formation of microspikes or filopodia (Nobes and
Hall, 1995). Furthermore, Rho GTPases have been found to play essential roles in a variety of cellular events which involve rearrangements of the actin cytoskeleton including cell migration, phagocytosis, axonal and dendritic growth and guidance, and synaptogenesis (Allen et al., 2000; Bradke and Dotti, 1999; Jin and Strittmatter, 1997;
May and Machesky, 2001; Murphy and Montell, 1996; Santos et al., 1997; Tashiro and
67 Yuste, 2004; Thies and Davenport, 2003; Yamashita et al., 1999; Zipkin et al., 1997). It is now generally accepted that Rho exerts its effects on the actin cytoskeleton by means of actin-myosin contractility, although the precise molecular mechanisms remain to be identified, whereas Rac and Cdc42 have been shown to activate members of the
Wiskott-Aldrich syndrome protein/WASP Verprolin-homologous protein
(WASP/WAVE) family, which stimulate the actin-related protein 2/3 (Arp2/3) complex to promote actin polymerisation (Machesky and Insall, 1999; Miki et al., 2000).
Activation of Rho family GTPases can be mediated by activation of numerous receptors including growth factor receptors (Hall, 1998), cell adhesion receptors such as integrin
(DeMali et al., 2003; Price et al., 1998), cadherin (Braga, 2002) and immunoglobulin superfamily members (Thompson et al., 2002), and G-protein-coupled receptors
(Collins et al., 1996; Coso et al., 1996).
Rho family effectors
Each Rho family GTPase interacts with multiple effectors and several effectors are recognised by multiple family members. A multitude of effectors have been identified, reflecting the diversity and complexity of Rho family GTPase signalling. A selection of
Rho family effectors are shown in Table 1.5.
A motif consisting of 16 amino acids (ISXPXXXXFXHXXHVG) known as the Cdc42/
Rac Interactive Binding (CRIB) domain (also known as p21 binding domain; PBD) has been identified as a region required for binding to Rac and/or Cdc42 in their GTP- bound (active) form, and was first identified in p65PAK (also known as p21 activated kinase/PAK1-6) (Burbelo et al., 1995). Subsequently, more than 25 CRIB domain- containing proteins (from a wide variety of eukaryotic species) have been identified
68 from GenBank database searches, including the mixed lineage kinases (MLK), WAVE, the activated Cdc42-associated tyrosine kinase (ACK) family of non-receptor tyrosine kinases, and marrow stromal/endothelial cell protein (MSE55) (see Table 1.5) (Burbelo et al., 1995). PAK has been implicated in Rac/Cdc42-mediated cytoskeletal rearrangements, promoting formation of polarised filopodia and membrane ruffles, and increasing motility, in fibroblasts (Sells, 1999; Sells et al., 1997), and leading to the loss of stress fibers and focal adhesions in HeLa and Swiss 3T3 cells (Manser et al., 1997).
However, such observations are complex, and often cell-type specific (for a recent review see (Bokoch, 2003)). For example, Y40C mutants of Rac and Cdc42, which are unable to bind to CRIB domain-containing proteins, are still able to induce actin reorganisation, suggesting that Rac/Cdc42 signalling to the actin cytoskeleton may be mediated by proteins other than PAK (Westwick et al., 1997), such as Partner of Rac1
(POR1) (Ishizaki et al., 1996; Van Aelst et al., 1996).
Table 1.5 Selected effectors of the Rho GTPase family
GTPase Effectors
Rho PI(3)K; PI(4,5)K; phospholipase D; rhophilin; kinectin; rhotekin; DGKζ, PRK1/PKN, PRK2, MBS, ROK/ROCK/Rho kinase; Bni1; Bnr1; PKC; p140mDia; Fks1; Fks2 ACK; PAK; PI(3)K; WASP; N-WASP; S6-kinase; MLK2; MLK3; IQGAP; MRCK; Cdc42 MSE55; Skm1; Gic1; Gic2; Borg; Bni1; Ste20; Cla4; IRSp53 NADPH oxidase; PAK; PI(3)K; PI(4,5)K; DGK; POSH; IQGAP; MLK2; MLK3; Rac MSE55; POR1; Sra-1; S6-kinase; IRSp53 (Aspenstrom, 1999; Bourne et al., 1990)
Rho, Rac and Cdc42 regulate gene expression in mammalian cells, for example by activating the serum response factor (SRF) (Hill et al., 1995). This activation is believed to be due to Rho family member-mediated changes in actin dynamics (Gineitis and
Treisman, 2001; Mack et al., 2001; Sotiropoulos et al., 1999). Rac and Cdc42 have also been shown to activate the SAPK/Jun N-terminal kinase (JNK) and p38 pathways,
69 which modulate processes such as gene transcription, apoptosis, development, transformation, immune activation and inflammation in response to toxins, physical stress and inflammatory cytokines (Coso et al., 1995; Minden et al., 1995). For a SAPK review see (Tibbles and Woodgett, 1999). Several studies suggest that binding/activation of PAK is necessary for activation of the SAPK pathway by
Rac/Cdc42 (Bagrodia et al., 1995; Joneson et al., 1996; Lamarche et al., 1996; Zhang et al., 1995). However there is also evidence suggesting that Rac can activate the SAPK pathway independently of PAK (Teramoto et al., 1996b; Westwick et al., 1997). In addition to PAK, two targets of Rac that have been reported to contribute to SAPK activation are MLK3 (Teramoto et al., 1996a) and the scaffold protein plenty of SH3s
(POSH) (Tapon et al 1998).
Crosstalk between Rho family members
Cross-talk between Rho GTPase family members (Rho, Rac, and Cdc42) is well- documented and known to play an important role in modulating and coordinating downstream cellular responses resulting from Rho GTPase signalling. In quiescent
Swiss 3T3 fibroblasts, a hierarchical linear cascade of activation has been identified, in which Cdc42 activates Rac and Rac activates Rho, resulting in the successive formation of filopodia, membrane ruffles and stress fibers (Etienne-Manneville and Hall, 2002;
Nobes and Hall, 1995). However, in other cell systems different Rho family members have been found to antagonise each other. For example, in N1E-115 neuroblastoma cells, Rho results in neurite retraction and cell rounding whereas Rac promotes cell spreading and neurite outgrowth (Leeuwen et al., 1997), and preferential activation of either Rac or Rho inhibits the opposite phenotype. Similarly, inhibition of Rho in Swiss
3T3 or Rat1 fibroblasts induces Rac-associated phenotypes (small focal contacts, cell spreading and motility) and vice versa with inhibition of Rac (Arthur and Burridge,
70 2001; Rottner et al., 1999). The exact mechanism(s) by which Rac and Rho pathways antagonistically interact with each other are unknown, although it has been proposed to involve either cross-regulation of downstream GTPase effector pathways (Sanders et al.,
1999; van Leeuwen et al., 1999), or direct disruption of GTPase conformation (resulting in modulation of its ‘switch’ mechanism of activation) (Caron, 2003; Nimnual et al.,
2003). The downstream Rac effector PAK1, for example, has been shown to inhibit Rho signalling (actin-myosin contractility) through its phosphorylation of myosin-II heavy chain and myosin light chain kinase (Sanders et al., 1999; van Leeuwen et al., 1999), and it has recently been reported to negatively-regulate the activity of Net1 (a Rho GEF) resulting in a downregulation of Rho activity in vitro (Alberts et al., 2005). A novel mechanism has also been described recently, in which production of ROS via Rac- dependent activation of NADPH oxidase (which is already known to contribute to host defence/microbial killing, cell cycling, transformation and gene transcription) (Bokoch and Diebold, 2002; Joneson and Bar-Sagi, 1998; Kheradmand et al., 1998; Suh et al.,
1999) causes an inhibition of the low molecular weight protein tyrosine phosphatase
(LMW-PTP) leading to activation of p190RhoGAP (which inactivates Rho), and this mechanism has been shown to be both necessary and sufficient for Rho activity downregulation in fibroblasts (Nimnual et al., 2003). Interestingly, the generation of
ROS by Rac is negatively regulated by Cdc42, although the significance of this finding in relation to Rho inhibition has not yet been investigated (Diebold et al., 2004).
Another Rho family member, RhoG, has been found to activate both Rac and Cdc42 in independent mechanisms (Gauthier-Rouviere et al., 1998), and a signalling pathway involved in RhoG-mediated activation of Rac has recently been described, in which activated RhoG binds engulfment and cell motility 1 (ELMO1) and this activates the
Rac-GEF dedicator of cytokinesis 180 (DOCK180) leading to Rac activation (Katoh
71 and Negishi, 2003), although the significance of this finding is controversial (Prieto-
Sanchez and Bustelo, 2003; Wennerberg et al., 2002).
1.3.2 Rab family GTPases
Rab GTPases are key regulators of vesicular transport in eukaryotic cells, and are involved in most if not all aspects of transport including vesicle formation, targeting and docking; and membrane remodelling and fusion. Most Rab proteins are ubiquitously expressed but some are tissue specific, for example Rab3 expression is restricted to neurons. Over 60 Rab proteins have been discovered in humans (Bock et al., 2001) and several of these are known to participate in more than one intracellular transport step, including Rab1, Rab5 and Rab11. For example, Rab5 has important roles in vesicle formation at the plasma membrane (McLauchlan et al., 1998), microtubule-dependent vesicle motility (Nielsen et al., 1999), membrane remodelling (through interaction with
PI(3)K) (Christoforidis et al., 1999b) and endosome fusion (Gorvel et al., 1991;
Stenmark et al., 1994).
Rab5 effectors
A variety of effectors have been described for Rab5, which reflects its role in numerous stages of vesicular transport steps (see above). 22 proteins that specifically bind to GTP- bound Rab5 were identified from bovine brain (Christoforidis et al., 1999a) and several of these are now known to be Rab5 effectors including Rabaptin-5, EEA1, Rabenosyn,
PI(3)K, Rabkinesin-6, tail interacting protein 47 (TIP47) and yeast Vac1 (Segev, 2001).
Interestingly, Rabaptin-5 also binds to both Rab4 and the Rab3 effector Rabphilin3, which suggests crosstalk exists between endocytic and exocytic pathways.
72 Crosstalk between Rab5 and Rho family members
Several observations have suggested that there could be crosstalk between Rho family
GTPases and Rab5. Rab5 has been reported to affect Rho family GTPase-dependent cytoskeletal changes. For example, dominant negative Rab5 inhibits reconstruction of the actin cytoskeleton after phorbol myristate acetate (PMA) treatment and constitutively active Rab5 induces actin remodelling; both processes are dependent on
Rac and Rho (Imamura et al., 1998; Spaargaren and Bos, 1999). Furthermore, Rab5 is essential for a form of receptor tyrosine kinase (RTK)-induced actin remodelling called
‘circular ruffling’ (a membrane ruffling event distinct from RTK-Ras-Rac-mediated lamellipodia formation, that is important in macropinocytosis and 3-dimensional migration, and is also dependent on activation of PI(3)K and Rac), which further suggests an involvement of Rab5 in Rho family GTPase signalling pathways (Lanzetti et al., 2004). Crosstalk between Rho family GTPase-mediated signalling to the actin cytoskeleton and Rab5-mediated endocytic events is also suggested by increasing evidence of involvement of actin reorganisation in endocytosis (Qualmann et al., 2000), and the involvement of several Rho family GEFs in various endocytic trafficking events. For example, activated Rac or Rho inhibits transferrin receptor endocytosis in
HeLa cells (Lamaze et al., 1996), RhoD localises to early endosomes and alters their distribution and motility (Murphy et al., 1996), RhoB localises and activates its effector protein kinase C-related kinase 1 (PRK1; also known as protein kinase N (PKN)) on endosomes and this retards trafficking of internalised EGF receptor from endosomes to a pre-lysosomal compartment (Gampel et al., 1999; Mellor et al., 1998), Rac1 can directly interact with Synaptojanin 2 (a polyphosphoinositide phosphatase implicated in the uncoating of clathrin-coated vesicles) (Malecz et al., 2000), and Intersectin 1 (a scaffolding protein regulating the formation of clathrin-coated vesicles in endocytosis)
73 (Guipponi et al., 1998) has been recently shown to exhibit Cdc42 GEF activity in cultured cells (Hussain et al., 2001).
1.4 Guanine nucleotide exchange factors (GEFs)
Small GTPases are highly regulated proteins and guanine nucleotide exchange factors
(GEFs) are the principle mediators of their activation. They achieve this in 2 ways: by destabilising GDP-GTPase interaction leading to GDP release, and by stabilising this nucleotide-depleted transition state, enabling GTP (which is at an approximately 10-fold higher concentration in the cell than GDP) to bind to the GTPase.
1.4.1 DH/PH (Dbl-family) GEFs
Most GEFs responsible for activating Rho family GTPases share two common motifs, the diffuse b-cell lymphoma (Dbl) homology (DH) and a C-terminally adjacent pleckstrin homology (PH) domain (together known as the DH/PH domain). To date, 6
DH/PH-containing GEFs have been identified in Saccharomyces cerevisiae, 18 in
Caenorhabditis elegans, 23 in Drosophila melanogaster and over 60 in humans
(Schmidt and Hall, 2002; Venter et al., 2001). The DH domain (first identified in the oncogenic product Dbl as the minimal region exhibiting Cdc42 GEF activity) is the catalytic domain necessary for GEF activity (Hart et al., 1994) whereas the PH domain has been proposed to function in localisation to the plasma membrane, as PH domains are known to bind to both phosphorylated phosphoinositides (PIPs) and proteins
(Lemmon and Ferguson, 2000; Rebecchi and Scarlata, 1998). This function is supported by the finding that the PH domain can be substituted with a membrane-targeting signal in certain GEFs such as Lymphoid blast crisis (Lbc)’s first cousin (Lfc) (Whitehead et al., 1995). However, the role of PH domains in Dbl-family GEFs remains controversial, as they have also been reported to participate in GEF activity/GTPase binding in Dbl’s
74 big sister (Dbs) and Trio (Liu et al., 1998b; Rossman et al., 2002) and in contrast, to inhibit GEF activity by masking access to the DH domain in several GEFs including
Vav and Son of Sevenless 1 (Sos1) (Han et al., 1998; Nimnual et al., 1998).
Furthermore, some GEFs such as T-lymphoma invasion and metastasis inducing protein
1 (Tiam1) and RasGRF contain a second (N-terminal) PH domain, and it is this PH domain that is required for membrane localisation rather than the PH of the DH/PH motif (Buchsbaum et al., 1996; Michiels et al., 1997; Stam et al., 1997).
Rho family GEFs without the classic DH/PH tandem motif have also recently been described. Examples include the novel Rac-specific GEF SWAP-70 which has a PH domain that is N-terminal to the DH domain (a PH/DH domain) (Shinohara et al.,
2002), and the Rac GEF DOCK180 which does not contain a DH/PH domain
(Kiyokawa et al., 1998). DOCK180 displays Rac GEF activity which has been mapped to a region called the ‘Docker/Dedicator of cytokinesis’ domain (Brugnera et al., 2002) and it requires a cofactor protein (ELMO) for its function (Brugnera et al., 2002).
Additionally, over 10 members of this unconventional GEF family have been identified
(Cote and Vuori, 2002; Meller et al., 2002).
Specificity of Rho family GEF activity
Several GEFs show specificity towards a particular Rho family member, for example
Faciogenital dysplasia protein (Fgd1) acts as a Cdc42 GEF and p115RhoGEF is a Rho
GEF, whereas others have the ability to activate several different family members, such as Dbl, Vav, and Epithelial cell transforming sequence 2 (Ect2). Interestingly, GEF specificity often varies according to the experimental conditions, for example Tiam1 is a GEF for Rho, Rac and Cdc42 in vitro but only for Rac in vivo (Michiels et al., 1995).
Furthermore, certain GEFs contain more than one DH/PH domain, for example
75 mammalian and Drosophila Trio, its mammalian orthologue Kalirin, its C. elegans orthologue Uncoordinated locomotion (UNC-73), each contain two DH/PH domains, one preferentially activating Rac1/RhoG and the other preferentially activating Rho
(Bellanger et al., 1998; Debant et al., 1996; Penzes et al., 2000; Penzes et al., 2001;
Steven et al., 1998).
Most GEFs contain diverse functional domains in addition to the DH/PH module, which may be involved in their regulation, coupling to upstream receptors or signalling proteins, or coupling to effectors; this also suggests that many GEFs may be multifunctional by nature and participate in and/or integrate several signalling pathways. Interestingly, both alpha-PAK-interacting exchange factor (α-PIX) and Fgd1 have been shown to possess functions that are independent from their GEF activity
(Daniels et al., 1999; Nagata et al., 1998). Additional functional domains identified in
Rho family GEFs so far include Src homology 2 (SH2), Src homology 3 (SH3), Ser/Thr kinase, Tyr kinase, Ras-GEF, Rho-GAP, Ran-GEF, PSD-95/DlgA/ZO-1 (PDZ),
Regulator of G-protein signalling (RGS), and additional PH domains. ALS2 is the only
Dbl-family GEF containing an RCC1-like domain, a VPS9 (Rab5 GEF) domain and
MORN motifs, that has been described to date.
Regulation of Rho GEF activity
Like their GTPase targets, GEFs are also highly regulated proteins, and there are numerous ways in which this regulation is achieved. GEFs are sometimes negatively- regulated by intramolecular interactions between the PH domain or other regulatory domains, which mask the catalytic DH domain, preventing binding and activation of
GTPases. This is seen in several GEFs including Vav, Sos1, Dbl, Lbc and p115RhoGEF. This allosteric inhibition can be relieved by several mechanisms
76 including binding of PtdIns(3,4,5)P3 to the PH domain, or tyrosine phosphorylation
(both of which have been shown to stimulate Vav activity) (Aghazadeh et al., 2000;
Bustelo, 2000; Han et al., 1998). In contrast however, binding of either PtdIns(3,4,5)P3 or PtdIns(4,5)P2 to the PH domain of Dbl inhibits its Cdc42 GEF activity (Russo et al.,
2001). Numerous GEFs can also be activated by G protein subunits, and this binding is presumed to activate the GEF by relieving autoinhibition. For example, binding of
Gα13 to the RGS domain of p115RhoGEF activates its Rho GEF activity (Hart et al.,
1998). Binding to certain signalling or adapter proteins are also thought to have various effects on GEF activity. For example, a novel protein has recently been discovered,
TRIPalpha, which inhibits the Rho-specific GEF domain of Trio (Schmidt et al., 2002).
1.4.2 Rab GEFs
Several mammalian Rab GEFs (and one putative Rab GEF; ALS2) have been described to date (although more GEFs for the Ypt GTPases, which are yeast Rab homologues, are known, one of which is Vps9). Examples of Rab GEFs are Rabex-5 (also known as
RabGEF1; a GEF for Rab5) (Horiuchi et al., 1997), MAP kinase activating death domain (Madd; also known as Rab3 guanine nucleotide exchange protein (Rab3GEP); a
GEF for Rab3A) (Wada et al., 1997) and RIN1,2,3 (Kajiho et al., 2003; Tall et al.,
2001). The majority of Ypt/Rab GEFs are part of large protein complexes within the cell, for example Rabex-5 forms a complex with Rabaptin-5 (a Rab5 effector), although the significance of such interactions is unknown (Horiuchi et al., 1997).
77
CHAPTER 2: MATERIALS AND METHODS
78 Unless stated otherwise, all molecular biology and cell culture reagents were purchased from Invitrogen Ltd. (UK) and all other chemicals were purchased from Sigma-Aldrich
Company Ltd. (UK). Stock solutions and buffers were prepared using ultrapure H2O from an Elga Maxima purification system. When required, solutions were sterilised either by autoclaving for 15 minutes at 15 lb/square inch or by filtration through a 0.2
µm pore filter (BDH).
2.1 Materials
2.1.1 Stock solutions
Acrylamide-bis-acrylamide (37:5:1 stabilised solution; National Diagnostics, USA)
Ampicillin (Amp; 100 mg/ml, filter sterilised)
Ammonium persulphate (APS; 20% w/v)
Adenosine 5′-triphosphate (ATP; 10 mM; Pharmacia, UK)
5-Bromo-4-chloro-3-indolyl-β-D-galactoside (X-gal; 20 mg/ml in N, N-dimethyl- formamide)
Chloroform (BDH): iso-amyl alcohol (IAA, Fisons) (24:1 v/v)
4′, 6-Diamidino-2-phenylindole (DAPI; 0.5 mg/ml in Dimethyl sulphoxide (DMSO))
Dithiothreitol (DTT; 1 M) dNTPs (25mM of each, 100 mM final; Pharmacia, UK)
Ethanol (70% v/v)
Ethylenediaminetetraacetic acid (EDTA; 0.5 M)
Ethylene glycol-bis(2 aminoethylether)-N,N,N’,N’-tetraacetic acid (EGTA; 0.5 M)
Ethidium bromide (10 mg/ml)
Glucose (20% w/v, filter-sterilised)
Glycerol (50% v/v, autoclaved)
79 HEPES (pH 7.0, 0.5 M)
Isopropyl β-D-thiogalactoside (IPTG; 100 mM)
Kanamycin (Kan; 10 mg/ml, filter-sterilised)
Magnesium chloride (MgCl2; 0.5 M)
Magnesium sulphate (MgSO4; 1 M, autoclaved)
Paraformaldehyde (PFA; 4% w/v in PBS)
Phenol (Fisons): chloroform: iso-amyl alcohol (25:24:1 v/v/v)
Phenylmethysulphonyl fluoride (PMSF; 100 mM)
Phosphate-buffered saline (PBS; Potassium phosphate 1.5 mM, Sodium phosphate 8.1 mM, NaCl 140 mM, Potassium chloride 2.7 mM)
Potassium acetate (C2H3KO2; 5 M)
RNase A (20 mg/ml)
Sodium azide (2% w/v)
Sodium dodecyl sulphate (SDS; 10% w/v)
Sodium chloride (NaCl; 4 M)
Sodium fluoride (NaF; 1 M)
Sodium hydroxide (NaOH; 10 M)
Sodium orthovanadate (Na2VO4; 20 mM, activated, pH 10.0)
Tris (hydroxymethyl) amino methane (Tris-HCl, buffered with HCl or NaOH to pH range 6.8-8.8)
Tris-buffered saline (TBS, pH 7.6; Tris hydroxymethyl amino methane 100 mM, NaCl
150 mM, buffered with HCl to pH 7.6)
Tris-EDTA (TE; Tris 10 mM, EDTA 1 mM; buffered with NaOH to pH 8.0)
Trypsin-EDTA (1x liquid; 0.25% in 1 mM EDTA)
Urea (6 M in PBS)
80 2.1.2 General molecular biology reagents
2.1.2.1 Plasmids
See Tables 2.1 and 2.2 below.
Table 2.1 Vectors
Vector Vector use Manufacturer
pBluescript SK+ Cloning vector Stratagene pCIneo Mammalian expression vector Promega pGEX-5X-1 Bacterial GST fusion protein expression vector Pharmacia
Table 2.2 Mammalian expression plasmids
Protein expressed Vector backbone Reference or source of plasmid Chloramphenicol pCIneo (Irving and Miller, 1997) acetyl transferase (CAT) Enhanced green fluorescent pEGFP.C1 Clontech protein (GFP) Myc-tagged ALS2 pCIneo N. J. O. Jacobsen (IOP) ALS2ala pCIneo M. S. Perkinton (IOP) DH/PH region of ALS2 pRK5myc A. Schmidt (University College London) (ALS2(DH/PH)) Rac1 pRK5myc A. Schmidt (University College London) N17 Rac1 pRK5myc A. Schmidt (University College London) L61 Rac1 pRK5myc A. Schmidt (University College London) RhoA pRK5myc A. Schmidt (University College London) Cdc42 pRK5myc A. Schmidt (University College London) N39 Rab5 pRK5myc A. Schmidt (University College London) VavΔN pRK5myc A. Schmidt (University College London) Net1ΔN pRK5myc A. Schmidt (University College London) Fgd1(DH/PH) pRK5myc A. Schmidt (University College London) p21 associated kinase-1 pCMV6myc S. Kesavapany (NIH, Bethesda, USA) (PAK1)
2.1.2.2 Primers
Unless indicated otherwise, all primers were obtained from Oswel DNA service,
University of Southampton. Sequencing of inserts in pBluescript SK+ was carried out
81 using the T3 and T7 primers:
T3 primer: 5'-AATTAACCCTCACTAAAGGG-3'
T7 primer: 5'-CGGGATATCACTCAGCATAATG-3'
Novel primers used for mutagenesis, sequencing and generating PCR products are described in the relevant sections below.
2.1.2.3 Growth of E. coli for DNA purification: media
Luria Bertani (LB) broth (powder; Invitrogen), 20 g/L sterilised by autoclaving
LB agar (powder; Invitrogen), 32 g/L sterilised by autoclaving
LB-amp broth (LB broth; ampicillin 100 µg/ml)
LB-amp agar (LB agar; ampicillin 100 µg/ml added when agar is less than ~55°C)
LB-kan broth (LB broth; kanamycin 50 µg/ml)
LB-kan agar (LB agar; kanamycin 50 µg/ml added when agar is less than ~55°C)
Bacteria containing plasmids of interest were stored at –70°C in sterile 25% glycerol
(v/v) solution in LB-amp/kan broth.
2.1.2.4 Plasmid DNA preparation from E. coli
Alkaline lysis solutions for small-scale purification of plasmid DNA from E. coli were used as previously described (Sambrook et al., 1989).
Solution 1 (sterilised by autoclaving)
50 mM Glucose
25 mM Tris pH 8.0
10 mM EDTA
82 Solution 2 (freshly-prepared from stock solutions)
0.2 M Sodium hydroxide
1% SDS (w/v)
Solution 3
5 M Potassium acetate
Glacial acetic acid
(final solution is 3 M with respect to potassium and 5 M with respect to acetate)
2.1.2.5 Agarose gel electrophoresis of nucleic acids
Solution for loading DNA samples in agarose gels
0.25% Bromophenol blue (w/v)
40% Sucrose (w/v)
Tris-Acetate-EDTA (TAE)
40 mM Tris-acetate pH 8.0
2 mM EDTA
Nucleic acid size markers
Phage λ DNA digested with Hind III (sizes in base pairs): 23,131; 9,419; 6,434; 4,335;
2,322; 2,023; 564; 125.
ΦX174 digested with Hae III (sizes in base pairs): 1,353; 1,078; 872; 603; 310; 281;
271; 234; 194; 118; 72.
2.1.2.6 Polymerase chain reaction (PCR) enzymes
Pyrococcus furiosus (Pfu) DNA polymerase (Stratagene)
83 Thermophilus aquaticus (Taq) DNA polymerase (Boehringer Mannheim)
2.1.3 Site-directed mutagenesis
ExSite™ PCR-based site-directed mutagenesis kit was purchased from Stratagene.
XL1-Blue supercompetent cells were provided with the kit. Details of specific primers used during work towards this thesis are given in the relevant results sections.
The following materials are supplied with the ExSite™ PCR-based site-directed mutagenesis kit:
Oligonucleotide control primer #1 (5’ phosphorylated; 75 ng/µl):
5’-CGCGCTTGGCGTAATCATGGTCAT -3’
Oligonucleotide control primer #2 (75 ng/µl):
5’-AGTACTCAATTAACCCTCACTAAAGGGAAC-3’ pWhitescript™ 4.5 kb control template (1.5 µg/µl) pUC18 transformation control plasmid (0.1 ng/µl in TE buffer)
10x mutagenesis buffer
200mM Tris-HCl (pH 8.8)
100mM KCl
100mM (NH4)2SO4
20mM MgSO4
1% Triton X-100 (v/v)
1 mg/ml BSA
Nucleotide mix
25 mM dATP
84 25 mM dCTP
25 mM dGTP
25 mM dTTP
2.1.4 Purification of GST fusion proteins
Glutathione elution buffer
10 mM reduced glutathione
50 mM Tris-HCl pH 8.0
2.1.5 Protein analysis
2.1.5.1 Protein sample preparation
Cell lysis buffer
50 mM Tris-HCl pH 7.6
150 mM NaCl
1 mM EDTA pH 8.0
1% Triton® X-100
Complete protease inhibitor cocktail (Roche)
Tissue homogenisation buffer
25 mM Tris-HCl pH 7.5
50 mM NaCl
5 mM MgCl2
1% Nonidet P40 (NP-40) (v/v)
5% Sucrose (w/v)
Complete protease inhibitor cocktail (Roche)
1 mM DTT
85 20 mM NaF
0.5 mM Na orthovanadate
1 mM PMSF
Magnesium lysis buffer
25 mM HEPES pH 7.5
150 mM NaCl
1% NP-40
10 mM MgCl2
1 mM EDTA
10% glycerol
1 mM NaF
1 mM Na orthovanadate
EDTA-free Complete protease inhibitor cocktail (Roche)
Cell fractionation homogenisation buffer
10 mM Tris pH 7.4
10 mM NaCl
3 mM MgCl2
1 mM EDTA
1 mM EGTA
1 mM PMSF
Complete protease inhibitor cocktail (Roche)
Cell fractionation wash buffer
0.1% NP-40
86 10 mM Tris pH 7.4
10 mM NaCl
3 mM MgCl2
1 mM EDTA
1 mM EGTA
1 mM PMSF
Complete protease inhibitor cocktail (Roche)
2x SDS protein sample buffer
125 mM Tris-HCl pH 6.8
2% SDS (w/v)
20% Glycerol (v/v)
0.005% Bromophenol blue (w/v)
1 M DTT stock solution was added just prior to use to a final concentration of 100 mM.
5x SDS protein sample buffer
125 mM Tris-HCl pH 6.8
10% SDS (w/v)
20% Glycerol (v/v)
0.005% Bromophenol blue (w/v)
1 M DTT stock solution was added just prior to use to a final concentration of 100 mM
2.1.5.2 Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE)
Resolving gel
375 mM Tris-HCl pH 8.8
0.1% SDS (w/v)
87 8-15% Acrylamide (v/v)
0.05% TEMED (v/v)
0.075% Ammonium persulphate (w/v)
Stacking gel
50 mM Tris-HCl pH 6.8
0.1% SDS (w/v)
3.75% Acrylamide (v/v)
0.15% TEMED (v/v)
0.075% Ammonium persulphate (w/v)
Running buffer
25 mM Tris-HCl pH 8.3
192 mM Glycine
0.1% SDS (w/v)
Electrophoresis markers
Rainbow™ coloured protein molecular weight markers (Amersham).
High molecular weight, sizes in kDa: 220, 97.4, 66, 46, 30, 21.5, 14.3.
Benchmark™ pre-stained protein ladder (Invitrogen); consists of 10 pre-stained protein bands in the range of 10-200 kDa (sizes vary with batch; for exact sizes see the relevant results sections).
88 2.1.5.3 Immunoblotting (Western blotting) solutions
Blotting buffer
39 mM Glycine
48 mM Trizma® base (Tris(Hydroxymethyl)aminomethane)
0.037% SDS (electrophoresis grade) (w/v)
20% Methanol (v/v)
TBS-Tween (Tris-buffered saline; Tween-20)
TBS pH 7.6
0.1% Tween-20 (v/v)
Blocking buffer
TBS-Tween
5% Dried skimmed milk (w/v)
Antibody incubation buffer
TBS-Tween
3% Dried skimmed milk (w/v)
2.1.5.4 Protein staining
Coomassie Blue
10% Glacial acetic acid (v/v)
45% Methanol (v/v)
0.05% Coomassie Brilliant Blue R (w/v)
89 Coomassie Blue destain
5% Glacial acetic acid (v/v)
40% Methanol (v/v)
Ponceau S red solution
7% Glacial acetic acid (v/v)
0.2% Ponceau S (w/v)
2.1.5.5 Treatment of protein samples with λ protein phosphatase
10x λ protein phosphatase buffer (supplied by manufacturer, NEB, with λ protein phosphatase)
50 mM Tris-HCl
0.1 mM Na2EDTA
5 mM DTT
0.01% Brij 35
2 mM MnCl2
(pH 7.5 at 25 °C)
2.1.6 Antibodies
2.1.6.1 Primary antibodies
For a list of primary antibodies used in these studies see Table 2.3.
2.1.6.2 Secondary antibodies
Immunoblotting: Sheep anti-mouse Ig and donkey anti-rabbit Ig conjugated to horseradish peroxidase (HRP; Amersham); working concentration 1:5000.
90 Immunofluorescence: Goat anti-mouse and goat anti-rabbit Igs coupled to Alexa Fluor
350, Alexa Fluor 488 or Alexa Fluor 546 (Invitrogen); working concentration 1:400.
Immunohistochemistry: Goat biotinylated anti-rabbit IgG (Vector laboratories); working concentration 1:1000.
Table 2.3 Primary antibodies
Antibody Supplier Monoclonal/Polyclonal, Species Working Immunogen dilution Monoclonal, KLH-conjugated 9B11 synthetic peptide 1:1000(IF) New England (myc-tag) (EQKLISEEDL) corresponding Mouse 1:5000 (IB) Biolabs to residues 410-419 of human c- 1.5µg (IP) Myc c-jun (H-79) Santa Cruz Polyclonal Rabbit 1:1000 (IB) Polyclonal, Full length GST from GST Sigma Rabbit 1:5000 (IB) Schistosoma japonicum Polyclonal, synthetic peptide New England PAK1 derived from the N-terminal Rabbit 1:1000 (IB) Biolabs sequence of human Pak1 Polyclonal, KLH-conjugated synthetic peptide Rho A,B,C Upstate (SYPDTDVILMCFSIDSPDSLE Rabbit 1:1000 (IB) N-KK) corresponding to amino acids 73-94 of RhoA,B,C Monoclonal, clone 23A8, 1:2000 (IB) Rac Upstate recombinant protein containing Mouse 1:200 (IF) the full length human Rac Monoclonal, fusion protein Cdc42 Upstate corresponding to residues 1-191 Mouse 1:1000 (IB) of full-length human Cdc42 BD Monoclonal, generated from Rab5 Mouse 1:300 (IF) Biosciences human Rab5 Monoclonal, clone DM1A, DM1A 1:10,000 (IB) Sigma microtubules from chicken Mouse (α-tubulin) 1:1000 (IF) embryo Phallotoxin isolated from Amanita AlexaFluor phalloides mushroom and labelled 5-10 Units/ml 568-Phalloidin Invitrogen N/A with red-orange-fluorescent Alexa (IF) (F-actin) Fluor 568 conjugate Polyclonal, peptide corresponding presenilin 1 D. Asuni to residues 1-25 of human Rabbit 1:1000 (IB) (PS1) (IOP) presenilin 1 Monoclonal, recognises MPM-2 Upstate phosphorylated serine/threonine- Mouse 1:1000 (IB) proline residues
Abbreviations: IOP= Institute of Psychiatry; IF= immunofluorescence; IB= immunoblotting; IP= immunoprecipitation.
91 2.1.7 Mammalian cell culture
2.1.7.1 Chinese Hamster Ovary (CHO) cell culture
CHO cell medium
Nutrient Mixture Ham F-12 (HAM) with glutamine
10% Fetal bovine serum (FBS) (v/v)
100 U/ml Penicillin
100 µg/ml Streptomycin
1x Trypsin-EDTA solution
0.05% Trypsin (w/v)
0.53 mM EDTA
2.1.7.2 Primary neuronal culture
Supplemented Neurobasal medium
Neurobasal medium
2% B27 supplement (v/v)
100 U/ml Penicillin
100 µg/ml Streptomycin
2 mM L-Glutamine
Poly-D-lysine (PDL)
20 µg/ml poly-D-lysine (PDL) in ddH2O
Trypsin-EDTA (TE) solution
0.05% Trypsin
0.53 mM EDTA
92 HBSS without Ca2+ and Mg2+
Triturating solution
10 mg/ml Albumax
500 µg/ml Trypsin inhibitor
10 µg/ml DNAase 1
2.1.8 In vitro protein kinase assays
Lysis buffer
20 mM HEPES pH 7.4
2 mM EGTA
1% Triton X-100
10% glycerol
1 mM DTT
1 mM Na orthovanadate
50 mM β-glycerophosphate
1 mM PMSF
Complete protease inhibitor (Roche)
Wash buffer I
100 mM Tris-HCl pH 7.6
500 mM Lithium chloride
0.1% Triton X-100
1 mM DTT
93 Wash buffer II
25 mM HEPES pH 7.5
0.2% Triton X-100
1 mM EDTA
Kinase assay buffer
25 mM HEPES pH 7.5
20 mM MgCl2
20 mM β-glycerophosphate
20 mM p-nitrophenylphosphate
0.1 mM Na orthovanadate
2 mM DTT
2.1.9 Immunohistochemistry
TBSA (Tris-buffered saline; Sodium azide)
TBS pH 7.6
0.05% Sodium azide
TBS-Triton (Tris-buffered saline; Triton X-100)
TBS pH 7.6
0.3% Triton X-100 (v/v)
94 2.2 Methods
2.2.1 General molecular biology methods
2.2.1.1 Quantitation of nucleic acids
Spectrophotometric quantitation of nucleic acids was performed using an Ultrospec
3000 spectrophotometer (Pharmacia Biotech). The absorbency of samples at 260 nm and 280 nm was recorded. An optical density reading (at a wavelength of 260 nm;
OD260) of 1 corresponds to a concentration of ~50 µg/ml for double stranded DNA and
40 µg/ml for RNA. The OD260/280 ratio gives an indication of purity of the sample. Pure
DNA has a value of 1.8 and pure RNA has a value of 2.0 (Sambrook et al., 1989). In the case of low yield DNA, for example gel purified fragments, DNA concentration was also roughly quantified using ethidium bromide fluorescent quantitation. This method relies on the ability of ethidium bromide to bind to the DNA helix via intercalation. UV- induced fluorescence emitted by DNA-linked ethidium bromide is proportional to the total mass of DNA. The amount of DNA in a sample could then be quantified by visual comparison with the UV-induced fluorescence of a known quantity of a DNA standard or series of standards. This technique has a detection limit of as little as 5 ng of DNA.
2.2.1.2 Restriction enzyme digestion of DNA
Plasmid DNA was digested using the appropriate restriction enzyme and corresponding buffer according to the manufacturers’ instructions (Invitrogen and New England
Biolabs). In a typical reaction, 5-10 units of enzyme were used per microgram of DNA
(1 unit is usually the amount of enzyme required to cleave 1 µg of DNA in 1 hour at
37°C in the appropriate buffer). The volume of enzyme never exceeded 10% of final reaction volume and the incubation time with the enzyme was typically 1-2 hours at
37°C, depending on the quantity of DNA being digested.
95 2.2.1.3 Alkaline phosphatase treatment
Calf intestinal alkaline phosphatase (CIAP, Pharmacia) was used to remove 5′- phosphate groups from digested, double-stranded DNA fragments to prevent self- ligation. Typically 0.05 units of CIAP (1 µl of a 1-in-10 dilution) were used to dephosphorylate 10 µg of linearised DNA in the buffer supplied (One-Phor-All buffer,
Pharmacia). The reaction mixture was incubated for 30 minutes at 37°C. The phosphatase was then inactivated by incubating the reaction at 85°C for 15 minutes.
2.2.1.4 Agarose gel electrophoresis of DNA
Agarose (Ultra pure, electrophoresis grade) was dissolved in boiling 1x TAE buffer and cast on a gel bed with a suitable comb using a horizontal gel apparatus (Hybaid). On setting, gels were placed in an electrophoresis tank containing 1x TAE buffer to a level just above the gel surface. DNA samples containing DNA loading buffer were loaded into the sample wells and were run at 2-10 Volts/cm. Agarose concentrations of 0.6-
1.5% were used, depending on the size of the DNA fragment of interest. Gels were stained with 10 µg/ml ethidium bromide to visualise DNA. They were then placed on a
3UV™ transilluminator emitting ultra violet light (λ = 302 nm), visualised on a Sony video monitor and videographed on a Sony video graphic printer.
2.2.1.5 Recovery of DNA from agarose gels
Ethidium-illuminated DNA bands were excised from TAE/agarose gels using a clean scalpel blade, placed in Spin-X centrifuge tube filters (Costar), and centrifuged for 5 minutes at 7,000xg to recover the DNA from the gel. 50 µl ddH2O was added to the filter and the columns centrifuged again to recover residual DNA.
96 2.2.1.6 Ethanol precipitation of double-stranded DNA
DNA was precipitated from aqueous solution by the addition of NaCl to a final concentration of 150 mM. Two volumes of ethanol (100%) were added to the mixture and the solution was chilled on ice for 30 minutes. DNA was pelleted by centrifugation at 14,000xg for 15 minutes. The DNA pellet was then washed with 70% ethanol to remove excess salt. After air-drying for 5 minutes the pellet was resuspended in ddH2O.
2.2.1.7 Purification of nucleic acids
Removal of contaminants, such as proteins, from aqueous solutions of nucleic acids was achieved by extraction with phenol:chloroform:isoamyl alcohol (25:24:1 v/v/v) followed by chloroform:isoamyl alcohol (24:1 v/v). An equal volume of phenol:chloroform:isoamyl alcohol was added to the aqueous solution of DNA, and the contents of the tube vortexed for up to one minute to form an emulsion. The emulsion was centrifuged at 14,000xg for 10 minutes and the aqueous phase collected. This procedure was repeated at least one more time, or until no contaminants were visible at the interface of the organic and aqueous phases. An equal volume of chloroform: isoamyl alcohol was then added and the contents of the tube were vortexed and centrifuged as before. The nucleic acids were recovered by precipitation with ethanol as described in Section 2.2.1.6.
2.2.1.8 Ligation of prepared vectors and DNA fragments
Linearised, dephosphorylated (if necessary) vector DNA and purified insert DNA concentrations were determined by gel electrophoresis alongside standards of known concentration. Ligations were carried out as previously described (Sambrook et al.,
1989). The ratio of picomole ends for vector:insert was 1:3 and the total quantity of
DNA usually used was approximately 80 ng. The number of picomole ends was
97 calculated using the following formula:
6 (2 x 10 )/ (660 x number of base pairs) = picomole ends/µg double stranded DNA
A typical reaction contained the vector and insert DNA plus 1 unit of T4 DNA ligase in the supplied buffer and 1 mM ATP in a final reaction volume of 10 µl. Ligations were performed overnight at 12-16°C. Appropriate control ligations lacking either vector or insert DNA were carried out simultaneously.
2.2.1.9 Preparation of electrocompetent bacteria
The E. coli strain DH5α (Hanahan, 1983) was used as a host for plasmid amplification, unless otherwise specified in the text. Preparation of electrocompetent DH5α was as follows. A single bacterial colony, picked from a streak-out onto an LB-agar plate, was used to inoculate 5 ml of LB-broth and grown overnight at 37°C in a shaking incubator.
2 ml of this culture was then diluted 100 fold in fresh LB-broth and grown at 37°C with shaking until an OD600 of 0.4 was reached (approximately 2 hours); at this point the bacteria are in mid-log phase growth. The culture was chilled on ice for 10-30 minutes to halt growth and the bacteria were pelleted at 6,000xg for 15 minutes at 4°C in a
Beckman Avanti J25 refrigerated centrifuge. The bacterial pellet was washed with 40 ml ice-cold ddH2O and pelleted by centrifugation, as before. This was repeated to ensure that all traces of salts from the LB-broth were removed. The pellet was then washed in ice-cold, sterile 10% (v/v) glycerol solution. The final bacterial pellet was resuspended in ice-cold 10% (v/v) glycerol (0.3 ml 10% glycerol per 100 ml original culture) which acts as a cryoprotectant, and was then divided into 40 µl aliquots which were frozen on dry ice and stored at −70°C.
98 The transformation efficiency of electrocompetent cells was calculated by transforming an aliquot with 0.1 ng of control plasmid (pUC19, Invitrogen) and counting the resulting number of ampicillin resistant colonies. The calculated efficiency of the
7 electrocompetent bacteria was routinely 5-7 x 10 colonies/µg of plasmid DNA.
2.2.1.10 Electroporation of DH5α cells
Prior to electroporation, salt was removed from ligation reactions by microdialysis for
20 minutes on cellulose membrane filters (0.025 µm pore size, Millipore-UK Ltd). The dialysed ligation mixture was added to an aliquot of ice-thawed electrocompetent DH5α cells and mixed gently. This was quickly added to a dry, chilled, 0.2 cm electroporation cuvette (Bio-Rad) and electroporated (field strength 2.5 kV/cm, capacitance 25 µF, resistance 200 Ohms) using a Bio-Rad Gene Pulser according to the manufacturer’s instructions. Following electroporation, 1 ml of 37°C LB-broth was added to the cuvette. The solution was then transferred to a 1.5 ml Eppendorf tube and incubated at
37°C for 45 minutes to allow the antibiotic resistance gene on the plasmid construct to be sufficiently expressed. When colour selection was possible, plates were spread with
40 µl of X-gal (20 mg/ml prepared in DMSO) and 20 µl of 100 mM IPTG (prepared in ddH2O) and left for 30 minutes prior to plating. The culture was then spread onto LB- amp agar plates (200µl per plate) using a moulded, sterile Pasteur pipette, and left to allow absorption to occur. Plates were then inverted and incubated at 37°C for 14-16 hours.
2.2.1.11 Screening recombinant clones
Following overnight incubation, bacterial colonies present on the LB-amp plates were picked (when colour selection was used, only white colonies were picked) with sterile
99 inoculation loops and each was used to inoculate 5 ml LB-broth containing the appropriate antibiotic. These samples were placed overnight in a shaking incubator at
37°C. Plasmid DNA from these samples was then prepared by the method of alkaline lysis (see Section 2.1.2.4) which exploits differences in properties between plasmid and bacterial genomic DNA (Sambrook et al., 1989). Briefly, 1.5 ml of culture was pelleted in a Biofuge at 14,000xg for 2 minutes and the supernatant discarded. The pellet was resuspended in 100 µl ice-cold Solution 1 by vortexing. 200 µl freshly prepared
Solution 2 was added and mixed by inverting the tube several times, before placing the tubes on ice for no longer than 5 minutes. 150 µl Solution 3 was then added and the contents of the tube mixed by inversion. The resulting mixture was left to precipitate on ice for 5 minutes before centrifugation at 14,000xg for 10 minutes; this stage efficiently removes the precipitated genomic DNA. The supernatant, now containing plasmid DNA and bacterial RNA, was placed in a fresh tube and extracted once with phenol:chloroform:isoamyl alcohol (25:24:1 v/v/v) and once with chloroform:isoamyl alcohol (24:1 v/v). The plasmid DNA was precipitated by adding 2 volumes of 100% ethanol and leaving at room temperature for 5 minutes and the precipitate was pelleted by centrifugation at 14,000xg for 10 minutes. Pellets were washed in 70% ethanol, air- dried and then resuspended in 50 µl TE, pH 8.0 containing 20 µg/ml RNase A. An aliquot of the plasmid DNA was removed and used for restriction enzyme digestion to screen for the desired construct.
2.2.1.12 Large scale preparation of plasmid DNA
The QIAfilter Plasmid Maxi Kit (Qiagen Ltd.) was used according to manufacturer’s instructions. This kit is based on the alkaline lysis method of DNA recovery (Sambrook et al., 1989); typically 200 ml of culture was used, resulting in approximately 500 µg of pure plasmid DNA.
100 2.2.1.13 DNA sequencing
DNA was sent to MWG Biotech UK Ltd., Milton Keynes, for sequence analysis.
2.2.1.14 Polymerase chain reaction (PCR)
Two different thermostable enzymes were used for PCR. The first, Thermophilus aquaticus (Taq) DNA polymerase (Boehringer Mannheim) was used to carry out preliminary PCRs to determine optimal cycle number and annealing temperature for each primer. Taq DNA polymerase has high processivity although it does not have proofreading activity, so it is not ideal for cloning. For cloning PCRs, native Pfu DNA polymerase (Stratagene), isolated from the hyperthermophilic marine archaebacterium
Pyrococcus furiosus, was used. Pfu DNA polymerase possesses both 5' to 3' DNA polymerase and 3' to 5' exonuclease activities. The 3' to 5' proof-reading activity associated with this enzyme results in a 12-fold increase in fidelity of DNA synthesis over Taq DNA polymerase. When used for PCRs, Pfu DNA polymerase generates blunt-ended products which, once purified, can be cloned directly into blunt cloning sites such as those created by Sma1 in the pBluescript SK+ cloning vector. Reactions were performed in 0.2 ml 8-tube PCR strips (BDH), with the following additions: template DNA (100-500 ng), primers (25 pmol of each), 5 µl of 10x Taq buffer
(Boehringer Mannheim) or native Pfu buffer (Stratagene), 8 µl of 100 mM dNTPs, 2.5 units of Taq or Pfu DNA polymerase and ddH2O to 50 µl. Each reaction was mixed gently by pipetting. Reaction mixes were then placed in a T3 Thermocycler (Biometra) for the PCR reaction, as shown in Table 2.4. The conditions found to be most productive were used in subsequent PCR reactions for that particular template/primer combination. In general, the lowest number of amplification cycles was used to minimise the possibility of errors by the DNA polymerase, whilst at the same time generating enough amplified product for subsequent procedures, such as DNA
101 purification and cloning. A sample of each PCR product was analysed for size, quality and quantity by agarose gel electrophoresis alongside appropriate DNA markers
(Section 2.2.1.4).
Table 2.4 PCR Cycling Parameters
Step Cycles Temperature (°C) Time
1. Pre-incubation 1 95 5 minutes 95 1 minute 2. Melting, Primer annealing and DNA 10-30 40-70 1 minute polymerisation 1 minute (Taq) or 3 72 minutes (Pfu) 3. Extension 1 72 10 minutes
2.2.2 PCR-based site-directed mutagenesis
ExSite™ PCR-based site-directed mutagenesis kit was purchased from Stratagene. The
ExSite™ system allows site-specific mutation in double stranded plasmids, which eliminates the need for subcloning into M13-based bacteriophage vectors and for single- stranded DNA (ssDNA) rescue (Weiner et al., 1994). This system uses increased template concentration and reduced cycling number to reduce potential second-site mutations during the PCR and can be used for producing single site point-mutations, large or small deletions and/or 5’-end oligonucleotide-directed base insertions. It was used in the work presented here to delete a large sequence (237 bp) of ALS2 (see
Chapter 4). The mutagenic primers used are detailed in the relevant results sections. The protocol for the ExSite™ mutagenesis kit was followed according to the manufacturer’s instructions and is described in brief.
Control and experimental reactions were set up as shown below and subjected to the
PCR steps as shown in Table 2.5. Control reactions were carried out to monitor
102 mutation efficiency. The pWhitescript vector is the pBluescript vector with a stop codon
(TAA) within the β-galactosidase gene. Thus, even when plated onto LB-amp plates spread with IPTG and X-gal, bacteria transformed with pWhitescript appear white.
Annealing the oligonucleotide control primers to pWhitescript and following the
ExSite™ protocol converts the stop codon back to the Glu encoding codon seen in pBluescript (CAA), allowing functional β-galactosidase expression. Bacteria containing successfully mutated (β-gal+) plasmids then appear blue on LB-amp plates spread with
IPTG and X-gal.
Control reaction:
1 µl (1.5 µg; 0.5 pmol) of pWhitescript 4.5 kb control template DNA
2.5 µl of 10x mutagenesis buffer (Section 2.1.3.2)
1 µl of 25 mM dNTP mix
2 µl (15 pmol; 150 ng) of oligonucleotide control primer #1 (24 mer, phosphorylated)
2 µl (15 pmol; 150 ng) of oligonucleotide control primer #2 (30 mer)
15.5 µl ddH2O to final volume of 24 µl
1 µl (5 U/µl) ExSite DNA polymerase blend
Experimental reaction:
0.5 pmol of template DNA (0.5 pmol of template DNA = 0.33 µl/kb x size of the template, kb)
2.5 µl of 10x mutagenesis buffer
1 µl of 25 mM dNTP mix
15 pmol of each 5’-phosphorylated primer (15 pmol = 5 ng/base x size of primer, bases) ddH2O to a final volume of 24 µl
1 µl (5 U/µl) ExSite DNA polymerase blend
103 Table 2.5 ExSite™ Mutagenesis Cycling Parameters
Step Cycles Temperature (°C) Time 94 4 minutes 1 1 50 2 minutes 72 2 minutes/kb 94 1 minute 2 8 56 2 minutes 72 1 minute/kb 3 1 72 5 minutes
2.2.2.1 Digesting and polishing the PCR product
Following PCR, the reactions were placed on ice. 1 µl (10 U/µl) of the Dpn1 restriction enzyme and 0.5 µl (2.5 U/µl) of cloned Pfu DNA polymerase (both supplied with the kit) were added and reactions were incubated at 37°C for 30 minutes, followed by 72°C for an additional 30 minutes. The Dpn1 endonuclease (target sequence 5’-Gm6ATC-3’) is specific for methylated and hemimethylated DNA and is used to digest parental DNA and to select for mutation-containing amplified DNA (Nelson and McClelland, 1992).
Cloned Pfu DNA polymerase is used prior to end-to-end ligation of the linear template to remove any bases extended onto the 3’ ends of the product by ExSite DNA polymerase blend.
2.2.2.2 Ligating the PCR product
Following analysis of a small amount of the reaction samples by agarose gel electrophoresis (section 2.2.1.4), the following was added to each sample:
100 µl of ddH2O
10 µl of 10x mutagenesis buffer
5 µl of 10 mM ATP
104 10 µl of this mixture was transferred to a fresh microcentrifuge tube and incubated with
1 µl (4 U/µl) of T4 DNA ligase for 1 hour at 37°C.
2.2.2.3 Transformation into XL1-Blue supercompetent cells
2 µl of the ligation reaction was added to 80 µl thawed XL1-Blue supercompetent cells in a prechilled Falcon® 2059 polypropylene tube and incubated on ice for 30 minutes.
The cells were then subjected to a heat pulse for 45 seconds at 42°C, and then placed on ice for a further 2 minutes. Immediately, the entire volume of the cells was plated onto
LB-amp plates. The transformed control cells were plated on an LB-amp agar plate pre- spread with 40 µl of X-gal (20 mg/ml prepared in DMSO) and 20 µl of IPTG (100 mM prepared in ddH2O) for colour selection. As a further control, 0.1 ng of pUC18 transformation control plasmid was added to an aliquot of cells and incubated on ice for
30 minutes, subjected to a heat pulse for 45 seconds at 42°C, placed on ice for a further
2 minutes (as above), then 5 µl of the cells was added to 80 µl of LB broth and this was plated on LB-amp agar plates pre-spread with X-gal and IPTG as described above. The plates were incubated overnight at 37°C. The mutagenized pWhitescript control colonies should appear as blue colonies, and the expected number of colonies for the experimental reaction should be greater than 100 colonies. The transformation efficiency for the pUC18 control plasmid should be greater than 250 colonies, with more than 98% displaying the blue phenotype.
2.2.2.4 Screening for mutated plasmids
Plasmid DNA was isolated as in Section 2.2.1.11 and then screened for the desired mutation using analytical restriction enzyme digests. Clones appearing to be positive for the mutation were then sequenced to ensure that no undesirable changes were introduced during the mutagenesis protocol.
105 2.2.3 Purification of GST fusion proteins
The glutathione S-transferase (GST) gene fusion system is an integrated system for the expression, purification and detection of fusion proteins produced in E. coli. The use of the BL21 strain of E. coli is generally recommended for protein production, as this strain lacks both the lon protease and the ompT outer membrane protease, which can degrade proteins during purification (Grodberg and Dunn, 1988). The system relies on the cloning of the cDNA of interest into a pGEX plasmid vector (Pharmacia). pGEX vectors are designed for inducible, high level intracellular expression of genes/gene fragments as fusions with Schistosoma japonica GST. Fusion proteins are purified from bacterial lysates by using glutathione sepharose 4B according to manufacturer's instructions (Pharmacia). Purified proteins can then be eluted from the glutathione sepharose 4B beads. The steps involved in the purification of GST fusion proteins from
E. coli are described in detail in the manufacturer's instructions (Pharmacia). Conditions were optimised as described below:
1. Preparation of stock glutathione sepharose 4B
Glutathione sepharose 4B beads are supplied as a 75% slurry in 20% ethanol. The beads were washed three times with ice-cold PBS to remove the ethanol and re-suspended in
PBS to produce a 50% slurry.
2. Preparation of bacterial sonicates
A single colony of E. coli harbouring a recombinant pGEX plasmid was used to inoculate 5 ml of LB-amp broth. This was incubated with shaking at 37°C for 12-15 hours. The resulting culture was diluted 1:100 into fresh, pre-warmed LB-amp broth, and grown at 37°C with shaking until the OD600 reached approximately 0.6-0.8. At this stage, 100 mM IPTG was added to a final concentration of 0.4 mM, to induce protein
106 expression, and incubation was continued for a further 3 hours. The culture was then centrifuged at 6,000xg for 15 minutes at 4°C, to sediment the cells, and the supernatant was discarded. The cell pellet was re-suspended using 50 µl ice-cold PBS with complete protease inhibitor cocktail (Roche) per 1 ml of culture. Alternatively, for recovery of insoluble proteins, the pellet was resuspended in 50 µl of 6 M urea per 1 ml of culture; see step 3 below. Suspended cells were disrupted by sonication performed on ice in 30 second bursts, 4-6 times. The resulting sonicate was centrifuged at 15,000xg for 20 minutes at 4°C, and the supernatant was used in step 4, below.
3. Dialysis of salts
As described above, proteins that were insoluble in PBS were solubilised in 6 M urea with complete protease inhibitor cocktail and rocked gently at 4°C for 20 minutes. This was centrifuged at 15,000xg for 15 minutes, and the resulting supernatant was dialysed against PBS (2 l PBS per 10 ml supernatant) overnight at 4°C. Centrifugation was repeated as above, to remove any protein that had come out of solution during dialysis, and the resulting supernatant was used in subsequent steps.
4. Batch purification of fusion proteins using glutathione sepharose 4B
2 ml of the 50% glutathione sepharose 4B slurry was added to 10 ml supernatant (i.e. 1 ml bed volume of beads per 200 ml bacterial grow). The mixture was incubated with gentle agitation at room temperature for 1 hour, or 4°C for 2 hours, after which the matrix with adsorbed fusion protein was sedimented by centrifugation at 500xg for 5 minutes. The supernatant was discarded and the glutathione sepharose 4B pellet washed three times with PBS. At this stage the bound fusion protein could be quantitated by direct comparison with standard amounts of bovine serum albumin (BSA) on
Coomassie Blue stained SDS-PAGE gels. Purified bound fusion protein samples were
107 stored at 4°C (in PBS as a 50% slurry) for future use or were eluted as described in step
5.
5. Batch elution
500 µl of reduced glutathione elution buffer was added to the sedimented beads (per 1 ml bed volume of beads). The re-suspended matrix was incubated with shaking at room temperature for 10 minutes to elute the fusion protein. The matrix was re-sedimented by centrifugation, and the supernatant stored on ice. Elution and centrifugation steps were repeated a further two times before pooling the three eluates, resulting in purified GST- fusion protein dissolved in 1.5 ml glutathione elution buffer. Samples were taken at various stages throughout the above procedures, boiled in 2x SDS protein sample buffer and stored at −20°C for future analyses.
2.2.4 Protein analysis
2.2.4.1 Protein concentration determination
Determination of protein concentration was carried out using the Bio-Rad protein assay according to the manufacturer’s instructions (Bio-Rad, UK). This is based on the
Bradford dye-binding procedure and is a simple colorimetric assay for measuring total protein concentration (Bradford, 1976). The assay is based on the observation that the maximum absorbance for an acidic solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 595 nm when binding to protein occurs. Absorbance readings were recorded at a wavelength of 595 nm using an Ultrospec 3000 spectrophotometer
(Pharmacia Biotech) and values were compared to a curve drawn from a set of BSA standards, freshly prepared each time the assay was performed.
108 2.2.4.2 SDS-PAGE of protein samples
For SDS-PAGE analyses of cell lysates, cells were washed in PBS, lysed in lysis buffer
(Section 2.1.5.1) for 20 minutes on ice, then boiled for 5 minutes in appropriate amounts of 2x or 5x SDS protein sample buffer (final concentration 1% SDS). The protein samples were then separated by SDS-PAGE using the Mini Protean II gel electrophoresis system (Bio-Rad) with a discontinuous buffer system. 8-12% acrylamide gels were run at a constant voltage of 120-180 V, until the dye front reached the bottom of the gel. Protein bands could be visualised at this stage by staining in
Coomassie Blue for 1 hour and then washing for 3 successive periods of 20 minutes in
Coomassie Blue destain.
2.2.4.3 Immunoblotting (Western blotting)
Transfer of proteins to nitrocellulose
After SDS-PAGE, proteins were transferred from gels to a 0.45 µm nitrocellulose support (Schleicher and Schuell) using the following filter sandwich:
Cathode
5 filters (5 x 8 cm, grade 1, Whatman) soaked in Blotting buffer
SDS-polyacrylamide gel
Nitrocellulose membrane pre-soaked in Blotting buffer (5 x 8 cm)
5 filters (5 x 8cm, grade 1, Whatman) soaked in Blotting buffer
Anode
This sandwich was assembled on a semi-dry transfer unit (Transblotter SD, Bio-Rad) at
17 V for 45-60 minutes. Nitrocellulose membranes with protein transferred onto them were referred to as "Blots". Blots were incubated with Ponceau S for 1-2 minutes to
109 determine the efficiency of protein transfer. Blots were then washed for 5 minutes with
TBS-Tween. This method of protein detection is compatible with all subsequent methods of immunological probing.
Antibody probing of membrane-bound proteins
Blots were incubated in blocking buffer (see Section 2.1.5.3) for up to 1 hour at 37°C to reduce non-specific binding. All incubations with antibodies were performed in antibody incubation buffer. Blots were incubated with an appropriate dilution of primary antibody overnight at 4°C (Table 2.3 gives the details of all primary antibodies used). The blots were then washed for 10 minutes in TBS-Tween; this was repeated a further four times, after which an appropriate dilution of secondary antibody was added for 1 hour at room temperature. Secondary antibodies used depended on the species in which the primary antibody was raised, for details see Section 2.1.6.2. Following a further four 15 minute washes in TBS-Tween, immunoreactive species were immediately visualised using enhanced chemiluminescence development reagents
(ECL; Amersham) according to the manufacturer's instructions, in conjunction with
Hyperfilm-ECL (Amersham). Film was developed using a Hyperprocesser (Amersham).
2.2.4.4 Treatment of samples with λ protein phosphatase
Immunoprecipitation of ALS2 from cultured cortical neurons was carried out with 500
µg protein as described in Section 2.2.8.1. The pelleted beads were washed 3 times with
1 ml lysis buffer without protease inhibitor, followed by 3 washes in ice-cold PBS. The beads were then resuspended in 20 µl of λ protein phosphatase buffer (Section 2.1.5.5) containing 1 µl (400 units) of λ protein phosphatase. The mixture was incubated at 30°C for 1 hour and 20 µl of 2x SDS protein sample buffer was then added before boiling for
5 minutes. An IP sample without λ protein phosphatase was also prepared as above, as a
110 negative control. The protein samples were then analysed by SDS-PAGE and immunoblotting (Sections 2.2.4.2-2.2.4.3).
2.2.5 Preparation of rabbit polyclonal antibodies
Rabbit polyclonal antibodies were prepared by immunising rabbits with GST fusion proteins. GST fusion protein production and immunisations were performed "in house".
Fusion proteins were purified and eluted as described in Section 2.2.3. Following a pre- immune test bleed, the rabbit was immunised with 100 µg of antigen (in 500 µl of glutathione elution buffer) emulsified with 500 µl of TiterMax Classic Adjuvant
(Sigma) according to the manufacturer's instructions. The rabbit was immunised with 4 x 0.2 ml antigen/adjuvant subcutaneously, at 0, 2, 4 and 6 weeks. At 8 weeks the rabbit underwent a termination bleed. Bleeds were obtained by trained staff, collected in glass bottles and left at room temperature for 1 hour to allow clotting. The clots were rimmed with a glass rod and allowed to retract overnight at 4°C. The serum was carefully removed, centrifuged at 2,000xg for 10 minutes at 4°C and then stored in aliquots at -
20°C. Once defrosted, sodium azide was added to 0.05% and aliquots were stored at
4°C for up to 6 months.
2.2.6 Mammalian cell culture and transfection
2.2.6.1 CHO cell culture
CHO cells were maintained in monolayer culture in Nutrient Mixture Ham F-12 (HAM) with glutamine, supplemented with fetal bovine serum, penicillin and streptomycin (see
Section 2.1.7.1) at 37°C under an atmosphere of 5% CO2. Cells were passaged when they formed an approximately 80% confluent layer in the culture vessel. Cell medium was aspirated and the cells washed twice with PBS. After removal of the second wash, enough trypsin-EDTA solution was added to cover the monolayer (e.g. 2 ml trypsin-
111 EDTA solution for a 175 cm2 flask). The vessel was rocked to evenly distribute the trypsin-EDTA solution before placing it in a 37°C incubator until the cells were just beginning to detach (1-2 minutes). The culture vessel was struck to dislodge remaining adherent cells. Cell medium (e.g. 8 ml for a 175 cm2 flask) was then added to inactivate the trypsin, and the cells were pipetted up and down to triturate the cells. Cell density was determined by counting a sample of the cell suspension using a haemocytometer under low power magnification before distribution of an appropriate volume into vessels for sub-culturing.
2.2.6.2 Primary embryonic rat cortical and hippocampal neuron culture
Cortical and hippocampal neurons were obtained from embryonic day 18 (E18) Sprague
Dawley rat embryos (Charles Rivers) and cultured on poly-D-lysine (PDL)-coated glass coverslips, or PDL-coated cell culture dishes in supplemented Neurobasal medium
(Section 2.1.7.2).
Dissection and preparation of neurons
A time-mated Sprague Dawley dam was sacrificed by cervical dislocation. The abdominal wall was cut through and the two horns of the uterus removed. The foetuses
(typically 10-12 per dam) were removed and dissected in HBSS without Ca2+/Mg2+. The brains were removed by cutting the skulls open with fine sprung scissors and gently scooping out using a spatula. The cortex and hippocampus were dissected under a dissecting microscope: The midbrain and cerebellum were removed using Dumont forceps and the meninges were removed to prevent contamination of cultures with fibroblasts; the cortex and hippocampus were then dissected and processed separately.
The isolated tissue was incubated with TE/HBSS solution for 20 minutes at 37°C, after which DNAse 1 solution (0.001% DNAse in HBSS) was added. The solution was
112 gently mixed; the tissue was then re-suspended in triturating solution, and triturated using glass Pasteur pipettes of decreasing bore size to produce a single-cell suspension, which was then diluted with supplemented Neurobasal medium. Cell density was determined using a haemocytometer. Various plating densities were used and cells were transfected and/or harvested at 1.5– 21 days in vitro (DIV).
2.2.6.3 Transient transfection
Lipid based transfection of CHO cell cultures
Lipofectamine™ reagent (Invitrogen) is a 3:1 (w/w) liposome formulation of the polycationic lipid 2,3-dioleyloxy-N-[2(spermine-carboxamido)ethyl]-N,N-dimethyl-1- propanaminiumtrifluoroacetate (DOPSA) and the neutral lipid dioleoyl phosphatidylethanolamine (DOPE) in membrane-filtered H2O. The positively charged and neutral lipids form liposomes that can complex with nucleic acids. When applied to cultured cells, the lipid-nucleic acid complex facilitates the uptake of nucleic acids into the cells. CHO cells were transfected essentially according to the manufacturer's instructions, using amounts of reagents as indicated in Table 2.6. The cells were plated out the day before the transfection experiment so that they reached 50-80% confluency on the day of transfection (half of this density was used for immunofluorescence), and were harvested for analysis 24 hrs post-transfection.
Table 2.6 Transient transfection of CHO cells with Lipofectamine™ Reagent
Diameter of dish Number of cells Amount of plasmid Amount of (cm) plated DNA (µg) Lipofectamine (µl) 3.5 2 x 105 1 4 6 6 x 105 3- 4 12 10 1.6 x 106 8- 10 32
113 Lipid-based transfection of primary cortical neuron cultures
Cortical neurons were transfected using Lipofectamine 2000™ reagent (Invitrogen) according to the manufacturer's instructions. Cells were cultured in 12-well plates (3.5 cm diameter) on PDL-coated glass coverslips, at a density of 3 x 105 cells per well in 1 ml of growth medium. 25 µl Optimem® reduced serum medium (Invitrogen) was mixed with 1 µl Lipofectamine 2000 and this was incubated for 5 minutes at room temperature. 1-4 µg DNA was mixed with 25 µl Optimem and this was added to the
Optimem/Lipofectamine 2000 and mixed thoroughly. After incubation at room temperature for a further 25 minutes, the mixture was added dropwise onto the cell culture medium, this was swirled gently and the cells were then returned to incubation at 37°C. Cells were harvested 4-24 hours later.
2.2.7 GTPase activation assays
Cellular Rho, Rac and Cdc42 activities were assayed using commercially available kits essentially according to the manufacturer’s instructions (Upstate). Active (GTP-bound)
Rho, Rac and Cdc42 were captured on GST-bait coupled to glutathione agarose beads.
Active Rho was captured using GST-Rhotekin Rho-binding domain (GST-RBD) and active Rac and Cdc42 captured using GST-PAK1 p21-binding domain (GST-PBD)
(Benard et al., 1999; Ren and Schwartz, 2000; Taylor and Shalloway, 1996).
CHO cells were cultured in 10 cm dishes and transfected using Lipofectamine™ reagent. 24 hours post-transfection and following 16 hours serum starvation, cells were washed twice with ice-cold PBS, harvested into 500 µl ice-cold magnesium lysis buffer
(MLB; Section 2.1.5.1), and cleared by centrifugation at 14,000xg for 5 minutes. 10 µl of each lysate sample was retained for analysis by SDS-PAGE and immunoblotting, and the remainder of each lysate was immediately incubated with 5-10 µg of either GST-
114 RBD (for Rho assay), GST-PBD (for Rac and Cdc42 assays) or GST (as a negative control) coupled to agarose beads for 45 minutes at 4°C. The beads were collected by centrifugation at 14,000xg for 5 minutes, the supernatant was discarded and the beads were resuspended in 20 µl of 2x SDS protein sample buffer. The samples were boiled at
95°C for 5 minutes and subjected to SDS-PAGE and immunoblotting.
Positive and negative controls for the assay were set up as follows: lysates of CHO cells co-transfected with either Rho, Rac or Cdc42 and pCIneo-CAT were cleared as above and EDTA was added to a final concentration of 1 mM. Each sample was divided into two tubes (one positive and one negative control): for the positive control, GTPγS was added to a final concentration of 100 µM and for the negative control GDP was added to a final concentration of 1 mM. The tubes were incubated at 30°C for 15 minutes with agitation. The reaction was terminated by placing tubes on ice and adding MgCl2 to a final concentration of 60 mM. Each sample was then incubated for a further 30 minutes with 5-10 µg of either GST-RBD (for Rho assay) or GST-PBD (for Rac and Cdc42 assays) at 4°C. The beads were collected by centrifugation at 14,000xg for 5 minutes, the supernatant was removed and the beads were resuspended in 20 µl of 2x SDS protein sample buffer. The samples were boiled at 95°C for 5 minutes and subjected to
SDS-PAGE and immunoblotting.
2.2.8 Immunoprecipitation and in vitro protein kinase assays
2.2.8.1 Immunoprecipitation from cell lysates
All steps of immunoprecipitation (IP) experiments were performed at 4°C or on ice.
Cells were washed twice in ice-cold PBS before being incubated in cell lysis buffer (e.g.
1 ml per 10 cm dish; see Section 2.1.5.1) for 20 minutes. The lysates were then scraped and pipetted into 1.5 ml tubes and centrifuged at 15,000xg for 10 minutes. The
115 supernatants were removed to fresh tubes and were then pre-cleared by incubation with
25 µl of Protein A-sepharose in cell lysis buffer (50% v/v) per 1 ml of lysate, for 1 hour at 4°C. Following centrifugation for 5 minutes at 5,000xg, lysates were transferred to fresh tubes and samples taken to check transfection and to quantitate total protein as in
Section 2.2.4.1. Protein concentration was adjusted to 1 µg/µl and equal amounts of protein (typically 500-1000 µg) for each sample were then used for IP. 100-500 µg
samples were placed into each of two fresh tubes. One of each duplicate sample was incubated with primary antibody and the other acted as a ‘no antibody’ control. Both samples were placed on a rotor for 2-16 hours after which 30 µl of protein A-sepharose beads (50% v/v in lysis buffer) was added. This was incubated for a further 1-2 hours on the rotor before centrifuging for 5 minutes at 5,000xg. The pelleted beads were retained and washed three times in 1 ml of lysis buffer. The beads were then either resuspended in the appropriate buffer or boiled in 30 µl of 2x sample buffer for 10 minutes and subjected to SDS-PAGE.
2.2.8.2 Immunoprecipitation from rat brain homogenate
Whole adult rat brain was homogenised in 5 ml of tissue homogenisation buffer, using a dounce tissue homogeniser. Homogenates were then passed through a 28-gauge needle three times and centrifuged at 15,000xg for 30 minutes. The supernatant was then precleared by incubation with 50 µl of protein A-sepharose (50% slurry in PBS) per 1 ml of homogenate, for 1 hour. Following centrifugation at 5000xg for 5 minutes, homogenates were transferred to a fresh tube and samples were taken for protein quantitation (as described in section 2.2.4.1). 2 mg of total protein in 1 ml of buffer was used per IP sample, and the IP was then carried out as for CHO cell lysates (see above).
116 2.2.8.3 In vitro kinase assays
Purification of kinases
Kinase-transfected cells were washed twice with ice-cold PBS and harvested by scraping into 1 ml kinase assay lysis buffer at 4°C (Section 2.1.8). The cell lysates were pre-cleared with 50 µl of protein A-sepharose beads (50% v/v in lysis buffer). The total protein was quantitated (section 2.2.4.1), 50 µg added to each of the three tubes and the volume made up to 1 ml with lysis buffer. The protein was then isolated by immunoprecipitation using 1.5 µg 9B11 antibody (see Section 2.2.8.1). The beads were then washed two times with 1 ml lysis buffer, then two times with 1 ml wash buffer I, and finally two times with 1 ml wash buffer II (see section 2.1.8). Negative controls were set up without immunoprecipitating antibody. “Reaction Mix” control was set up without lysate/kinase.
Kinase assays
5 µg Myelin basic protein (MBP) was used as a substrate in the PAK1 kinase assay.
Kinase reactions were set up as follows: 30 µl 20 µM ATP in kinase buffer (made from
10 mM stock of ATP), 0.259 MBq [γ-32P] ATP (0.7 ul of 370 MBq/µl; Amersham-
Pharmacia Biotech), 5 µg substrate and kinase buffer to a final volume of 35 µl. The kinase reaction mixture was then added to the immunoprecipitated kinases (or no kinase control or reaction mix only), still on beads. Samples were incubated for 20 minutes at
30°C and the reactions then terminated by the addition of 20 µl of 5x sample buffer.
The samples were analysed by performing SDS-PAGE, vacuum drying gels on a Slab
Gel Dryer vacuum pump (Savant) and exposing them to Hyperfilm-ECL (Amersham) for 2-72 hours.
117 2.2.9 CHO cell fractionation
CHO cells were cultured in 10 cm dishes and transfected with 10 µg ALS2 cDNA using
Lipofectamine™ reagent. 18 hours post-transfection cells were washed twice with ice- cold PBS and harvested into 1 ml of cell fractionation homogenisation buffer (Section
2.1.5.1). Cells were homogenised using a dounce homogeniser and centrifuged at 375xg for 5 minutes at 4°C. The pellet (nuclear fraction) was washed with cell fractionation wash buffer (Section 2.1.5.1) five times, resuspended in a final volume of 100 µl wash buffer and added to 100 µl 2x SDS protein sample buffer. The supernatant was centrifuged at 100,000xg for 30 minutes at 4°C. The resulting supernatant (cytoplasmic fraction) was added to 80 µl 2x SDS protein sample buffer and the pellet (membrane fraction) was washed with wash buffer 3 times, resuspended in 50 µl wash buffer and
50 µl 2x SDS protein sample buffer. Samples were subjected to SDS-PAGE and immunoblotting. The following amounts were used to obtain equal loading: 25 µl cytoplasmic fraction; 1.25 µl membrane fraction; 1.75 µl nuclear fraction. For tubulin immunoblots 1/10th of the above amounts were loaded on the gel.
2.2.10 Immunohistochemistry
The methods used in immunohistochemical staining are based on avidin-biotin and peroxidase methodologies (Hsu and Raine, 1981). The target protein (ALS2) was detected by a primary ALS2 antibody (described in Chapter 3) that in turn was detected with a biotinylated secondary antibody. The secondary antibody was then bound with
Avidin-Biotinylated enzyme Complex (ABC). Staining was completed by addition of the 3,3'-diaminobenzidine tetrahydrochloride (DAB) chromogen which precipitates at the antigen site, generating a brown stain.
118 Sectioning of frozen rat brain
Rat brain was fixed with 4% paraformaldehyde (PFA)/PBS (pH 7.4) for 48 hours at room temperature. Fixed brain specimens were washed once with PBS and then soaked in 50 ml of 30% sucrose in TBSA solution at 4°C (see Section 2.1.10). When the brain specimens descended to the bottom of the tube they were removed and stored at -70°C. A Leitz 1321 freezing microtome (Leitz, Japan) was used to prepare frozen brain sections. Free-floating sections in TBS were used for immunohistochemistry.
Avidin-Biotinylated enzyme Complex (ABC) staining
40 µm sections were used for staining. Endogenous peroxidase activity was blocked with 1% H2O2 in TBS for 15 minutes after which the sections were rinsed 3 times for 5 minutes in TBS. Non-specific protein binding was blocked with 15% normal goat serum (NGS; Vector Laboratories, UK) in TBS-Triton (see Section 2.1.10) for 40 minutes. Sections were then incubated overnight at 4°C, with constant gentle agitation, in primary antiserum diluted with 10% NGS/TBS-Triton. Primary antibody treated sections were rinsed 3 times for 5 minutes in TBS and incubated for 2 hours in 1:1000 diluted biotinylated secondary antiserum (Vector laboratories, USA) with 10%
NGS/TBS-Triton. Secondary antiserum treated sections were rinsed 3 times for 5 minutes in TBS and incubated for 2 hours in Vectastain® Elite® ABC solution (Vector laboratories, USA) prepared at 1:1000 dilution in TBS, 30 minutes in advance. ABC solution treated sections were rinsed 3 times for 5 minutes in TBS. Sections were then incubated in 0.05% DAB solution (preferably in darkness to reduce background staining) for 5-10 minutes. 0.05% DAB solution was prepared by crushing a 10 mg
DAB tablet (Sigma, UK) into 20 ml TBS with 6 μl of 30% H2O2 and used immediately.
Staining intensity was frequently checked during incubation. The reaction was terminated by transferring sections to ice-cold TBS. Sections were rinsed a further 3
119 times for 5 minutes in TBS. Sections were taken from the TBS, placed onto Superfrost- plus slides (BDH, UK) and air-dried overnight. Sections on slides were then left in xylene for at least 30 minutes, and mounted with a glass cover slip with a xylene-based mountant, e.g. DPX® (BDH, UK). Images were captured on a Zeiss Axioscope 2 MOT using Axiocam and Axiovision software (Zeiss, Germany).
2.2.11 Immunofluorescence
Cells on glass coverslips were washed twice in PBS at room temperature before being fixed in 4% PFA/PBS for 15 minutes, and permeabilised in 0.1% Triton X-100/PBS for
10 minutes. Cells were then incubated with 5% FBS/PBS blocking solution for 30-60 minutes to reduce non-specific binding. Incubation with primary antibodies in blocking solution was then carried out for 1 hour. The cells were washed for 10 minutes in PBS
5-6 times, before being incubated for 1 hour with 5% FBS/PBS containing goat anti- mouse and goat anti-rabbit Igs coupled to Alexa Fluor 350, Alexa Fluor 488 or Alexa
Fluor 546 (Invitrogen). Nucleic acids were stained with 50 µg 4′,6-Diamidino-2- phenylindole (DAPI; 0.5 mg/ml in DMSO) for 5 minutes, followed by a further 5-6 washes (10 minutes each wash) in PBS. Finally, coverslips were mounted onto slides using Vectashield mountant (Vector Laboratories). Conventional images were viewed using a Zeiss Axioscop microscope and captured using a charge coupled device (CCD) camera (Princeton Instruments), and confocal images captured using a Zeiss LSM 510
META confocal microscope at the appropriate excitation wavelengths.
120
CHAPTER 3: PREPARATION OF AN ALS2
ANTIBODY AND LOCALISATION OF ALS2 IN
NEURONAL TISSUES
121 3.1 Introduction
Alsin/ALS2 was first identified in 2001 by two independent research groups (Hadano et al., 2001; Yang et al., 2001) as a protein of unknown function that is mutated in rare autosomal recessive juvenile forms of MND. The identified mutations all cause a predicted loss of function of the ALS2 protein. This research project was started immediately after the publication of these findings, with the purpose of investigating the normal function of ALS2, so as to shed light on the pathological mechanisms involved in MND.
Sequence homology data suggests that ALS2 may function as a guanine nucleotide exchange factor (GEF) for the Ras superfamily of GTPases. ALS2 Long-form contains three putative GEF domains; an amino-terminal domain that displays homology to the
Ran GEF RCC1, a central region containing Dbl and pleckstrin homology domains
(DH/PH) that are found in GEFs for Rho family members, and a carboxyl-terminal vacuolar protein sorting 9 (VPS9) domain which is found in GEFs for Rab5. ALS2
Short-form has been identified at the mRNA level, although there is as yet no evidence of the presence of a short form protein. As this predicted truncated protein is not thought to contain any of the functionally homologous GEF domains an antibody was created specifically to ALS2 Long form (referred to simply as ALS2).
The study of the subcellular localisation of ALS2 could provide clues to its function within the cell. For example a cytoplasmic/intracellular or plasma membrane localisation would indicate function as a Rho or Rab GEF whereas localisation within the nucleus may suggest a function as a GEF for Ran (as Ran is activated in the nucleus)
(Hughes et al., 1998). The first step of this project was therefore the preparation and characterisation of an antibody to ALS2, in order to facilitate such studies.
122 3.2 Methods
3.2.1 PCR and cloning into pGEX expression vector
PCR reactions were carried out in a T3 Thermocycler (Biometra) in order to amplify a sequence of cDNA encoding a 651 bp fragment of ALS2 (corresponding to amino acid residues 452-668 of human ALS2). Pairs of synthetic oligonucleotide primers were designed to introduce an EcoR1 site (GAATTC) at each end of the construct (to facilitate cloning into the EcoR1 site of pGEX-5X-1), as follows:
Primer1: 5’-GCGGAATTCGAACAGGTTAAACAGGAATCAATGC-3’
Primer 2: 5’-GCGGAATTCTCCAAGCTTACTACAGGAGAGAAG -3’
Approximately 100 ng DNA was amplified in a mixture containing 25 pmol of each primer (Oswel DNA service, University of Southampton), 16 µl deoxynucleotide triphosphates (25 mM each), 1 µl Pfu DNA polymerase (2.5 units) and 10 µl 10x Pfu reaction buffer (Stratagene). The final reaction volume was 100 µl. Double-stranded
DNA was denatured at 95°C for 5 minutes, followed by a cycle of melting at 95ºC for 1 minute, primer annealing at 60°C for 1 minute, and finally DNA polymerisation at 72°C for 3 minutes. This was repeated for 12, 18, 24 or 30 cycles, and was followed by a final extension step of 72°C for 10 minutes. The PCR product (obtained from the minimum cycle number) was cloned blunt into the Sma1 site of pBluescript SK+ (Stratagene) and sequenced (MWG Biotech) to ensure no errors had occurred. The PCR product (insert) was then sub-cloned in-frame as an EcoR1 fragment into the pGEX-5X-1 GST vector
(see Section 2.2.1).
3.2.2 Preparation of GST fusion protein
The GST-ALS2452-668 construct was transformed into and expressed in BL21-
CodonPlus®-RIL (BL21-RIL; Stratagene) E. coli. BL21-RIL cells contain extra copies
123 of rarely-used E. coli tRNAs (argU, ileY and leuW) that are frequently found in A/T rich genomes. Empty pGEX-5X-1 was used to express GST alone in BL21-RIL cells as a control. A 1 ml sample was taken from each culture at 0 hrs (at OD600 0.6-0.8) and 1 hr, 2hr and 3hrs post-induction with 0.4 mM IPTG. Samples were resuspended in ice- cold PBS, lysed by sonication and separated by centrifugation (at 15,000xg for 10 minutes) into soluble (supernatant) and insoluble (pellet) fractions. 2x SDS protein sample buffer was added to each sample and the samples were boiled for 5 minutes prior to analysis by SDS-PAGE.
The GST-ALS2452-668 fusion protein was expressed predominantly in the insoluble fraction. The protein was therefore purified from the insoluble fraction by solubilising in 6 M urea followed by dialysing against PBS to a final concentration of approximately
30 mM urea. The protein was then captured using glutathione sepharose 4B beads, and eluted with reduced glutathione elution buffer.
3.2.3 Production of a polyclonal antibody to ALS2
100 µg GST-ALS2452-668 antigen (in glutathione elution buffer) was concentrated to 500
µl using Centricon® centrifugal filter units (Millipore) and was emulsified in 500 µl
TiterMax® classic adjuvant, according to the manufacturer’s instructions (Sigma-
Aldrich). An immunologically-naïve New Zealand white rabbit was injected into 4 subcutaneous sites with 200 µl antigen/adjuvant per site. This was repeated 3 more times at 10-15 day intervals until a terminal bleed was carried out (47 days after the first immunisation).
3.2.4 Affinity purification of ALS2 polyclonal antibody
Antibodies were affinity-purified from sera using immobilised antigen. Adsorbents were
124 prepared by spotting 500 µg antigen onto Immobilon PVDF membrane (Millipore). The protein was allowed to dry before blocking in 5% skimmed milk/PBS for 2 hours. 2 ml of the antiserum was incubated with the membranes and 10 ml 3% skimmed milk/PBS overnight. After incubation, the membranes were washed with PBS, and antibody was eluted in 500 µl Elution Buffer (0.5 M glycine, 0.5 M NaCl, pH 2.4) for 10 minutes.
The elution step was repeated three more times and the pooled eluate was neutralised to pH 8 before addition of sodium azide to 0.05%.
3.2.5 Cell culture and transfection
CHO cells were cultured and transiently-transfected as described in Section 2.2.6. A carboxyl-terminal myc-tagged human ALS2 cDNA was prepared by N.J.O. Jacobsen
(Institute of Psychiatry). Briefly, the 5’ end of ALS2 was amplified by PCR from a human brain cDNA library and ligated to a 5’-truncated partial ALS2 cDNA (clone
KIAA1563, obtained from Kazusa Research Institute, Japan) so as to create a full-length clone. A carboxyl-terminal myc tag was then added by PCR and the tagged full-length cDNA then cloned into pCIneo (Promega) as a Not1 fragment.
3.2.6 CHO cell fractionation
CHO cells were cultured in 10 cm dishes and transfected with 5 µg myc-tagged ALS2 cDNA using Lipofectamine™ (as described in Section 2.2.6.3). Cells were washed twice with ice-cold PBS, and incubated in 1 ml cell fractionation homogenisation buffer
(Section 2.1.5.1) on ice for 10 minutes, followed by homogenisation with a dounce homogeniser. Cell fractionation was performed as described in Section 2.2.9. Samples were boiled for 5 minutes in 2x SDS protein sample buffer, and subjected to SDS-
PAGE and immunoblotting.
125 3.2.7 Antibodies
Immunoblotting, immunohistochemistry and immunofluorescence were carried out as described in Sections 2.2.4.3, 2.2.10 and 2.2.11. The ALS2 antibody described in this chapter was used at varying concentrations as indicated in the relevant results sections.
For details and concentrations used of other antibodies in this chapter see Table 2.3.
Antibody preabsorption was carried out by incubation of ALS2 antibody with 10x molar excess of the purified GST-ALS2452-668 immunogen or with purified GST protein alone (quantified by SDS-PAGE and staining with Coomassie Blue, alongside BSA standards), prior to addition to the immunoblot.
3.3 Results
3.3.1 GST fusion protein preparation
A GST-ALS2 expression plasmid (GST-ALS2452-668) was created in order to purify a fragment of ALS2 to use as an antigen for the production of ALS2 antibodies. This involved isolating a 651 bp fragment of ALS2 corresponding to amino acid residues
452-668 of human ALS2 (Figure 3.1), which was generated by PCR as described above.
A sample of each PCR product was checked for size, quality and quantity by agarose gel electrophoresis, in order to identify the most productive conditions (Figure 3.2).
PCR reactions were initially carried out with Taq DNA polymerase in order to optimise the primer annealing temperature. Reactions were carried out with annealing temperatures of 55°C, 60°C, 65°C, and 70°C and samples from these reactions were analysed by agarose gel electrophoresis. 60°C was found to be the optimum temperature
(Figure 3.2 A). PCR reactions were then carried out with Pfu DNA polymerase at 60°C with different cycle numbers (12, 18, 24 and 30). 18 cycles was used as this was the lowest number of cycles (minimising the possibility of errors by the DNA polymerase)
126 that generated enough amplified product for the subsequent procedures (Data not shown). The PCR product was then purified from the TAE gel (see section 2.2.1.5) and cloned into the Sma1 cloning site of pBluescript SK+. Following colour selection and sequencing (MWG Biotech), a clone with the correct insert was identified, and the insert was then sub-cloned in-frame into pGEX-5X-1 as an EcoR1 fragment. Clones were screened using an analytical digest with EcoR1 and BamH1 and a clone containing the GST-ALS2452-668 insert was identified (referred to as GST-ALS2452-668).
3.3.2 GST fusion protein expression and purification
The bacterial expression plasmid GST-ALS2452-668 was transformed into BL21-RIL E. coli and the protein was expressed as described (Section 2.2.3). Samples taken during expression were analysed by SDS-PAGE followed by staining with Coomassie Blue and immunoblotting (Figure 3.3). The GST-ALS2452-668 fusion protein was identified as a band migrating at approximately 55 kDa (which is the predicted size) and the presence of this protein was confirmed by immunoblotting with a polyclonal antibody to GST
(Figure 3.3 A, B). The majority of this protein was found in the insoluble fraction.
Empty pGEX-5X-1 was expressed under the same conditions, as a control; the increase in expression with time of a band migrating at approximately 29 kDa, mainly in the soluble fraction, confirmed that GST was expressed in abundance under these conditions (Figure 3.3 C and D). The conditions were optimised for maximal expression of both the GST control and GST-ALS2452-668 by varying the amount of IPTG (0.1 mM,
0.25 mM and 0.4 mM) and the temperature during expression (30°C and 37°C).
However, the GST-ALS2452-668 fusion protein was expressed in the insoluble fraction in all of the above conditions, and the maximal expression was with a temperature of 37°C and induction with 0.4 mM IPTG. After capture with glutathione sepharose beads,
127 elution with reduced glutathione buffer and concentration, the GST-ALS2452-668 fusion protein was analysed and quantified by SDS-PAGE (Figure 3.4).
3.3.3 Characterisation of ALS2 antibody
Following production of the polyclonal ALS2 antibody, the antibody was characterised by immunoblotting and immunofluorescence. Myc-tagged ALS2 was expressed in CHO cells and 16 hours later cells were harvested in lysis buffer, boiled in 2x SDS protein sample buffer and subjected to SDS-PAGE and immunoblotting. Equal amounts of
CHO lysate were probed with varying amounts of ALS2 antibody in order to determine the specificity of the antibody and the optimal concentration for detection of ALS2 protein (Figure 3.5 A). A single species was detected in ALS2 transfected but not non- transfected CHO cells, with both the ALS2 and myc (9B11) antibodies (Figure 3.5 A).
This reactive species migrated at approximately 185 kDa, which is the predicted size for the myc-tagged ALS2 construct, and the optimal concentration of the ALS2 antibody was determined to be 1:1000. As negative controls, preimmune serum from the rabbit that produced the ALS2 antibody was used in place of a primary antibody on one blot, and primary antibody was omitted on another.
Immunoblotting of rat or mouse brain homogenates with the polyclonal ALS2 antiserum was initially unsuccessful. ALS2 is now known to be a low abundance protein, and is thought to represent only approximately 0.0003% of the total detergent- soluble fraction of mouse brain lysate (Yamanaka et al., 2003). The antibody was therefore affinity-purified with GST-ALS2452-668 antigen. The affinity-purified antibody resulted in lower non-specific background, allowing blots to be exposed for a longer period of time, which is more efficient for detecting low-abundance proteins. The affinity-purified ALS2 antibody was able to detect a band of the predicted molecular
128 mass of ALS2 in rat brain homogenate, co-migrating with the 185 kDa band in transfected CHO cell lysate (Figure 3.5 B). To further determine the specificity of the antibody, it was incubated with 10x molar excess of the antigen prior to incubation with the blot. This preabsorption with the antigen resulted in the loss of the 185 kDa band, whereas incubation with GST alone did not affect the intensity of the band. Therefore, the ALS2 sequence of antigen (rather than the GST sequence) was effectively
“competing out” the band, which indicates that the band detected is ALS2.
CHO cells transfected with myc-tagged ALS2 were also subjected to immunofluorescence; cells were incubated with the ALS2 antibody at a concentration of
1:100 and anti-myc (9B11) antibody. The pattern of staining obtained with ALS2 antibody was identical to that of 9B11 (Figure 3.5 C). The overexpressed ALS2 protein appeared to be localised to the cytoplasmic (particularly perinuclear) region of the cell, but excluded from the nucleus.
3.3.4 Localisation of ALS2 in brain and spinal cord
The ALS2 antibody was used to detect endogenous ALS2 in rat brain and spinal cord sections by immunohistochemistry (Figure 3.6). Frozen brain and spinal cord sections were incubated with ALS antibody at a concentration of 1:100 and staining was observed in a variety of neuronal populations. This staining was found to be specific for
ALS2 as preincubation of the antibody with GST-ALS2452-668 resulted in a lack of immunoreactivity (Figure 3.6). High magnification of the sections revealed that ALS2 staining within neurons appears to be in both the cell body and neuronal processes.
Interestingly, ALS2 was detected in large neurons in the spinal cord that are located in a motor neuron-rich area of the spinal cord. Therefore, ALS2 seems to be present in
129 motor neurons. Particularly intense staining was seen in neurons of the cerebellum, and
ALS2 staining was also observed in neurons of the cortex, hippocampus and brainstem.
The affinity-purified ALS2 antibody was then used to immunoprecipitate endogenous
ALS2 from rat brain. A single band was identified, co-migrating with ALS2 immunoprecipitated from transfected CHO cells using anti-myc antibody 9B11, at approximately 185 kDa (Figure 3.7). Thus, the ALS2 antibody produced in these studies successfully detects ALS2 by immunofluorescence, immunoblotting, and immunohistochemistry, and can immunoprecipitate ALS2 from rat brain.
3.3.5 Overexpressed ALS2 is localised in cytoplasmic and membranous fractions of mammalian cells
Differential centrifugation of ALS2-transfected CHO cell homogenate was used to determine the subcellular localisation of overexpressed myc-tagged ALS2. ALS2 was found to be localised to the cytoplasmic and membrane fractions but not the nuclear fraction (Figure 3.8). To check the purity of the prepared fractions, the samples were probed for endogenous proteins that are known to be specific to the cytoplasmic, membrane and nuclear fractions (α-tubulin, presenilin1 and c-jun respectively). This confirmed that the different fractions had been separated successfully.
3.4 Discussion
In these studies, a polyclonal antibody that is specific to endogenous ALS2 was prepared and used successfully for immunoblotting, immunofluorescence, immunohistochemistry and immunoprecipitation. The antibody detected a major species of 185 kDa, which is the predicted size of the long form of ALS2. Specificity of the antibody has been demonstrated by pre-absorption with the antigen. One further test for specificity would be to use on ALS2 mouse knockout tissue, or cells expressing an
130 ALS2 RNAi construct, although as yet neither of these have as yet been published.
Recently, the ALS2 C-terminal like (ALS2CL) gene has been identified as an ALS2 homolog, which encodes a 108 kDa protein ‘ALS2CL’ (Devon et al., 2005; Hadano et al., 2004). Although the antigen used to prepare the antibody in these studies shows approximately 92% sequence similarity to ALS2CL, no band migrating at 108 kDa was seen in immunoblots of rat brain homogenate or cultured rat embryonic cortical neurons, and therefore this antibody might not detect ALS2CL.
To investigate the subcellular localisation of ALS2, a myc-tagged ALS2 construct was overexpressed in CHO cells, and immunofluorescence and cellular fractionation (by differential centrifugation) studies were carried out. ALS2 was found to be mainly localised to the cytoplasm and membrane fractions and excluded from the nucleus, which is in agreement with studies of endogenous ALS2 in rodent brain and spinal cord
(Devon et al., 2005; Topp et al., 2004; Yamanaka et al., 2003). However, the cell fractionation studies reported here show a higher level of ALS2 in the cytoplasmic fraction compared with the membrane fraction. This is most likely due to these experiments being carried out in transfected CHO cells and this overexpression of ALS2 would result in higher levels in the cytoplasm. Immunohistochemical staining revealed a high abundance of ALS2 in cerebellar neurons, which is consistent with the finding that the cerebellum is the brain region with the highest ALS2 expression in immunoblots of mouse brain homogenates (Yamanaka et al., 2003). Most interestingly, ALS2 was detected in large cells in spinal cord which closely resemble motor neurons. Thus it appears that ALS2 is present in motor neurons, the cell type that is predominantly affected in MND.
131 Figure 3.1 Schematic of ALS2 Long-form, ALS2 Short-form and GST-ALS2452-668
Schematics of ALS2 Long-form, ALS2 Short-form and the predicted GST-ALS2452-668 fusion protein are shown as indicated. The ALS2 fragment corresponds to amino acids 452-668 of human ALS2, which encompasses sequences of the RCC1-like domain.
ALS2 Long-form RCC1 DH PH MORN VPS9
1 59 167 218 525 576 627 690 885 1049 1244 1551 1657 169 578 9011657 1007 1656
ALS2 Short-form Unique 24 residues
1 372 396
GST-ALS2452-668
452 668
132 Figure 3.2 PCR amplification of sequences encoding GST-ALS2452-668
GST-ALS2452-668 fusion protein was created by PCR and the products were subjected to agarose gel electrophoresis for analysis. PCR products resulting from reactions carried out with Taq DNA polymerase at different annealing temperatures (55°C, 60°C, 65°C and 70°C) were as shown. The PCR product ran at the predicted size of approximately 670 base pairs (bp). L is DNA ladder ΦX174HaeIII. Subsequent PCR reactions were carried out with Pfu DNA polymerase with an annealing temperature of 60°C. PCR products were cloned into pBluescript SK+, clones were identified by colour selection and restriction digest with EcoR1, and positive clones were sequenced (MWG Biotech). The insert from a chosen clone was then subcloned into pGEX-5X-1 as an EcoR1 fragment, and positive clones were identified by an analytical digest with EcoR1 and BamH1 followed by agarose gel electrophoresis.
L 55°C 60°C 65°C 70°C bp 1353 1078 872 603 310
133 Figure 3.3 GST-ALS2452-668 and GST protein expression
GST-ALS2452-668 fusion protein and GST were expressed in BL21-RIL E. coli and samples were taken when uninduced (0 hrs) and at hourly time points post-induction with 0.4 mM IPTG (1 hr, 2 hrs and 3 hrs). Samples were separated into total (T), soluble (S) and insoluble (I) fractions; proteins were separated by SDS-PAGE and either stained with Coomassie Blue (A, C) or immunoblotted and probed for GST-ALS2452-668 or GST using a rabbit polyclonal GST antibody (Sigma) (B, D). GST control was found to be expressed mainly in the soluble fraction (migrating at approximately 29 kDa) whereas
GST-ALS2452-668 was found mainly in the insoluble fraction. GST-ALS2452-668 was found to migrate at its predicted size of ~55 kDa and the presence of the fusion protein in the samples was confirmed by immunoblotting with anti-GST.
A 0 hrs 1 hr 2 hrs 3 hrs kDa T S I T S I T S I T S I 220 97
66 GST-ALS2452-668 45
30
B GST-ALS2452-668
C 0 hrs 1 hr 2 hrs 3 hrs
T S I T S I T S I T S I kDa
66 45
30 GST D GST
134 Figure 3.4 Purification of GST-ALS2452-668
GST-ALS2452-668 was expressed in BL21-RIL cells for 3 hrs, solubilised in 6 M urea, dialysed against PBS and then captured with glutathione sepharose 4B beads. The GST-
ALS2452-668 antigen was eluted by addition of reduced glutathione buffer, concentrated by centrifugal filtration and analysed by SDS-PAGE alongside BSA standards (stained with Coomassie Blue). The sample shown represents 1/60th of the yield from 200 ml of E. coli. The total yield was therefore approximately 150 µg per 100 ml of E. coli culture.
BSA standards (μg) GST- kDa 0.5 1 2 5 ALS2452-668 97
66
45
135 Figure 3.5 Characterisation of ALS2 antibody (A) non-transfected CHO cells (NT) and CHO cells transfected with myc-tagged ALS2 (ALS2) were subjected to SDS-PAGE and immunoblotting with varying concentrations of rabbit polyclonal ALS2 antisera (1:100, 1:500, 1:1000, 1:2500; 1:25,000) or with myc antibody (9B11). A single band was observed at the predicted size of approximately 185 kDa. Blots were incubated with pre-immune serum (PI) or without a primary antibody (No primary) as controls. (B) CHO lysates (NT and ALS2- transfected) and adult rat brain homogenate (RB) were subjected to SDS-PAGE and immunoblotting with affinity-purified ALS2 antibody at a concentration of 1:1000, either without or with (Preabsorbed GST-ALS2452-668) incubation with 10x molar excess of GST-ALS2452-668 antigen. Incubation of affinity-purified ALS2 antibody with GST protein was carried out as a control (Preabsorbed GST). Samples were also immunoblotted without primary antibody as a negative control (No antibody). The 185 kDa band was not detected upon preabsorption specifically with the GST-ALS2452-668 antigen. (C) immunofluorescence was carried out in CHO cells transfected with myc- tagged ALS2, co-staining with both rabbit polyclonal ALS2 antisera and mouse monoclonal myc (9B11) antibodies. Scale bar is 50 µm.
A PI 1:100 1:500 1:1000 1:2500 1:25,000 No primary 9B11 kDa NT ALS2 NT ALS2 NT ALS2 NT ALS2 NT ALS2 NT ALS2 NT ALS2 NT ALS2 220
97
66 45
B Preabsorbed Preabsorbed ALS2 antibody GST-ALS2452-668 GST No antibody NT ALS2 RB ALS2 RB ALS2 RB ALS2 RB kDa
220
97
C ALS2 Myc (9B11)
136 Figure 3.6 Localisation of ALS2 in adult rat brain and spinal cord sections ALS2 immunoreactivity is seen in various brain and spinal cord regions. (1) is Cortex incubated with the ALS2 antibody preabsorbed with the GST-ALS2452-668 immunogen. (2) is Cortex (without preincubation with immunogen). (3) is Brainstem, (4) is Cerebellum and (5) is Hippocampus. (6) is cervical spinal cord and (7) is high magnification of the area outlined in (6). Scale bars are 200 µm (1-5) and 100 µm (6,7).
137 Figure 3.7 Affinity-purified ALS2 antibody immunoprecipitates ALS2 from rat brain CHO cells were transfected with myc-tagged ALS2 and immunoprecipitation was carried out with (+) or without (-) 9B11 antibody to the myc-tag. Adult rat brain (RB) was homogenised and immunoprecipitation was carried out with (+) or without (-) affinity-purified ALS2 antibody. The samples were then probed on immunoblots with affinity-purified ALS2 antibody. An approximately 185 kDa species was immunoprecipitated, both from transfected CHO cells with 9B11 antibody and from rat brain with affinity-purified ALS2 antibody.
CHO RB kDa - + - +
182
120
88
138 Figure 3.8 ALS2 is present in the cytosolic and membrane fractions of transfected CHO cells CHO cells were transiently transfected with myc-tagged ALS2 (ALS2) and homogenised in buffer containing 1% Triton (see section 3.2.6). Homogenates were separated into cytosolic (C), membrane (M) and nuclear (N) fractions by differential centrifugation. Samples were then subjected to SDS-PAGE and immunoblotting using ALS2 antibody. NT and ALS2 are samples of non-transfected and ALS2-transfected CHO cell lysates. Samples were also probed for c-jun (nuclear-specific protein), presenilin 1 (PS1; membrane-specific protein) and α-tubulin (cytosol-specific protein) as fractionation controls.
NT ALS2 C M N
ALS2
c-jun
PS1
α-tubulin
139
CHAPTER 4: ALS2 ACTS AS GEF FOR RAC1 AND
ACTIVATES PAK1
140 4.1 Introduction
Although the function of ALS2 is as yet unclear, its amino acid sequence predicts that it functions as a GEF. GEFs positively regulate the activity of members of the Ras superfamily of GTPases. These GTPases cycle between inactive (GDP-bound) and active (GTP-bound) conformational states and GEFs stimulate GTP-binding so as to promote activation of the GTPase (For reviews see Etienne-Manneville and Hall, 2002;
Schmidt and Hall, 2002). ALS2 contains three putative GEF domains; an amino- terminal domain that displays homology to the Ran GEF RCC1, a central region containing Dbl and pleckstrin homology (DH/PH) domains that are found in GEFs for members of the Rho family of GTPases (for example Rho, Rac and Cdc42), and a carboxyl-terminal vacuolar protein sorting 9 (VPS9) domain which is found in GEFs for
Rab5 (Hadano et al., 2001; Yang et al., 2001). Indeed, there is now evidence that ALS2 functions as a GEF for Rab5, and this is via its VPS9 domain (Otomo et al., 2003; Topp et al., 2004).
To investigate the hypothesis that ALS2 functions as a GEF for the Rho family of
GTPases via its DH/PH domain, I have studied the level of activation of the three most characterised Rho family GTPases (Rho, Rac and Cdc42) in the presence of ALS2.
These studies were carried out in mammalian (CHO) cells. I have demonstrated that
ALS2 causes an increase in activation of Rac but not Rho or Cdc42. The catalytic activity of Dbl-family (DH/PH) GEFs is predicted to be mediated by the Dbl homology
(DH) domain, based on studies of the Dbl protein (Hart et al., 1994), and this has subsequently been confirmed for a number of GEFs (Aghazadeh et al., 1998; Kubiseski et al., 2003; Matsuo et al., 2002; Movilla and Bustelo, 1999; Whitehead et al., 1995).
Therefore, I have disrupted the DH domain of ALS2 by deletion of its key residues, and have found that an intact DH domain is necessary for its function as a Rac GEF.
141 Active (GTP-bound) Rac binds to a variety of effectors including members of the p21- activated kinase (PAK) family. In order to further elucidate the downstream components of ALS2-Rac1 signalling, I have investigated the activation of PAK1 by ALS2, using in vitro protein kinase assays. These studies have revealed that ALS2 overexpression results in activation of PAK1 and that this activation is dependent on ALS2 Rac GEF activity (as shown by the use of the ALS2 mutant described above). Thus, ALS2 has the ability to function as a Rac GEF and activate PAK1 in mammalian cells.
4.2 Methods
4.2.1 Plasmids
Plasmids used were as follows: RhoA, Rac1, Cdc42, L61Rac1, Net1ΔN, VavΔN and
Fgd1(DH/PH) (all in pRK5myc expression vector), myc-tagged ALS2 (in pCIneo), pCIneo-CAT and PAK1 (in pCMV6myc) were as described (Table 2.2). ALS2(DH/PH) was prepared by Anja Schmidt (University College London): Briefly, the DH/PH GEF domain (amino acids 681-1010) of ALS2 was generated by PCR and cloned as a
BamH1-EcoR1 fragment into pRK5myc.
A mutant ALS2 clone (ALS2ΔDH) in which the DH/PH GEF domain was disrupted was created, by deletion of sequences encoding the DH domain (residues 747-826), which has been identified as the region necessary for the catalytic GEF activity of the
Dbl-family GEFs. The mutagenesis was performed using an ExSite™ PCR-based site- directed mutagenesis kit as described in Section 2.2.2. The following 5’-phosphorylated primers were used (Oswel DNA service, University of Southampton):
Primer 1: 5’-ACTCAGTTGATGGAAATACTGAATAC-3’
Primer 2: 5’-ACACAGCTTGCTGAATCGGCTAGC-3’
142 PCR products were analysed by restriction enzyme digest and sent for sequencing to confirm that the mutagenesis had been successful (MWG Biotech).
4.2.2 Cell culture and transfection
Transfections were carried out with Lipofectamine™ reagent as described in Section
2.2.6.3. For GTPase activation assays, CHO cells were cultured in 10 cm dishes and transiently co-transfected with 2 µg of either myc-tagged RhoA, Rac1 or Cdc42 and 8
µg of ALS2 or pCIneo-CAT cDNA. As positive controls, CHO cells were co- transfected with RhoA, Rac1 or Cdc42 and constitutively active GEFs for Rho, Rac or
Cdc42 (Net1ΔN, VavΔN and Fgd1(DH/PH) respectively). As a negative control for the
Rac activation assay (and to investigate the ability of ALS2ΔDH, an ALS2 mutant in which the DH domain is disrupted, to act as a GEF for Rac) cells were co-transfected with 2 µg Rac and 8 µg ALS2ΔDH. Additionally, the ability of the isolated DH/PH domain to stimulate activation of Rac was investigated and so CHO cells were transfected with Rac1 and myc-tagged ALS2(DH/PH). For PAK1 kinase assays, CHO cells were cultured as above and co-transfected with 2 µg myc-tagged PAK1, 2 µg Rac1 and 6 µg of either pCIneo-CAT as a negative control, ALS2, ALS2ΔDH or constitutively active Rac1 (L61Rac1) as a positive control.
4.2.3 GTPase activation assays
CHO cells were cultured in 10 cm dishes and co-transfected as described above. Cells were harvested 24 hours after transfection, following 16 hours of serum-starvation (in order to obtain a low basal level of Rho family GTPase activation (Jalink et al., 1993;
Jalink et al., 1994; Tigyi and Miledi, 1992)). Cellular Rho, Rac and Cdc42 activities were assayed using commercially available kits essentially according to the manufacturer’s instructions (Upstate) as described in Section 2.2.7. Captured Rho, Rac
143 and Cdc42 were detected on immunoblots using anti-myc antibody (9B11) and the relative amounts quantified by pixel densitometry using a Bio-Rad GS710 imaging densitometer and Quantity 1 software.
4.2.4 In vitro kinase assays
CHO cells were cultured in 10 cm dishes and co-transfected as described above. Cells were harvested 24 hours after transfection, following 16 hours of serum-starvation.
PAK1 was immunoprecipitated from the lysate using 1.5 µg of anti-myc epitope tag
(9B11) antibody, and in vitro kinase assays were carried out in kinase assay buffer
(Section 2.1.8) with 0.26 MBq γ[32P]-ATP, 20 μM ATP and 5 µg myelin basic protein
(MBP) as substrate, as described in Section 2.2.8.3. Samples were boiled in 2x SDS protein sample buffer and subjected to SDS-PAGE followed by staining with
Coomassie Blue and autoradiography, or immunoblotting.
4.3 Results
4.3.1 ALS2 acts as a GEF for Rac1 but not RhoA or Cdc42
ALS2 contains a central DH/PH domain which shows homology to GEFs that regulate the Rho family of GTPases. To determine whether ALS2 activates the 3 most widely- studied Rho family GTPases RhoA, Rac1 and Cdc42, in vivo pull-down assays were utilised to monitor the activities of these GTPases in transfected CHO cells. Active
(GTP-bound) Rho binds to the Rhotekin “Rho binding domain” (RBD; corresponding to residues 7-89 of mouse Rhotekin) whereas active Rac and Cdc42 both bind the PAK1
“p21 binding domain” (PBD; corresponding to residues 67-150 of human PAK1). GST-
RBD and GST-PBD “baits” can thus be used to isolate GTP-bound RhoA, Rac1 and
Cdc42 from experimentally manipulated cells; the amounts of these GTPases detected on immunoblots correlates with their activities.
144 Co-transfection of cells with ALS2 induced a significant increase in the amount of Rac1 but not Cdc42 pulled-down by GST-PBD (Figure 4.1B, C). ALS2 did not activate
RhoA (Figure 4.1A). Control samples, in which GST-PBD or GST-RBD was substituted with GST alone, did not result in activation of Rac, Rho or Cdc42. Similarly, pre-loading with GDP resulted in no active (GTP-bound) GTPase in the cells.
Transfection of dominant active GEF constructs, or pre-loading with non-hydrolysable
GTPγS, resulted in significant stimulation of GTPase activity in these assays (Figure
4.1). The isolated DH/PH GEF domain (residues 681-1010) did not activate Rac1 in these assays, which suggests that other regions of ALS2 may be necessary for control of its Rac1 GEF function (Figure 4.1 D). Thus, ALS2 stimulation of Rac1 activity requires the full-length ALS2 protein.
To confirm that the stimulatory effect on Rac activity was indeed mediated by the
DH/PH domain, an ALS2 mutant (ALS2ΔDH) was created in which the majority of the catalytic DH domain (residues 747-826) was deleted, using the ExSite™ PCR-based site-directed mutagenesis system (Figure 4.2 A, B, C). Rac activation assays were carried out as described above. ALS2ΔDH did not activate Rac1 (Figure 4.2 D), which shows that the ALS2 Rac1 GEF activity occurs via its catalytic DH domain.
4.3.2 ALS2 activates PAK1 and this is dependent on a functional DH domain
Immediate downstream targets for Rac include members of the p21-activated kinase
(PAK) family of serine/threonine kinases. PAK1 is a major neuronal member of the
PAK family. The ability of ALS2 to stimulate PAK1 activity was therefore investigated.
In vitro PAK1 kinase assays were performed from CHO cells co-transfected with PAK1
+/- ALS2 as described above, using MBP as a substrate. These experiments revealed that ALS2 stimulated PAK1 activity by approximately 1.5-fold, as measured by the
145 extent of phosphorylation of MBP. This stimulation was not seen in cells co-transfected with the non-functional mutant ALS2ΔDH, which indicates that this activity is dependent on a functional DH/PH domain and implicates Rac activity in ALS2 activation of PAK1 (Figure 4.3). Thus, ALS2 functions as a GEF to regulate Rac1-
PAK1 signalling.
4.4 Discussion
In these studies I have demonstrated that ALS2 functions as a Rac GEF via its DH domain, and have identified PAK1 as a downstream target of this activation. During the course of these studies (as discussed in Section 1.2.2), two research groups also reported that ALS2 functions as a Rac GEF. This was demonstrated by the use of similar pull- down assays utilising GST-PBD, in insect sf9 cells (Topp et al., 2004) and in CHO cells
(Kanekura et al., 2005). The level of Rac1 activation observed upon overexpression of
ALS2 was similar to that reported here. However, Kanekura et al. (2005) studied levels of endogenous Rac1 whereas in the studies reported here Rac1 was transfected into the
CHO cells. In agreement with the findings reported here, no activation of endogenous
Cdc42 or RhoA (pulled down with GST-RBD) was observed (Kanekura et al., 2005).
Furthermore, these authors used a mutant ALS2, in which a key threonine residue of the
DH domain had been mutated to alanine (residue 701) to demonstrate that a non- mutated DH domain was necessary for Rac activation (Kanekura et al., 2005), which confirms the findings reported here that the DH domain mediates ALS2 Rac GEF activity, as discussed below.
In addition, studies using in vitro GDP-dissociation assays have shown that ALS2 is a
GEF for Rab5 (Otomo et al., 2003; Topp et al., 2004). The stimulatory effect of ALS2 on Rab5 required its VPS9 domain (Otomo et al., 2003). The authors of this paper also
146 studied the effect of ALS2 on Rac activity but were unable to detect any stimulation of
Rac. This discrepancy is probably due to the different assays used. Topp et al. (2004),
Kanekura et al. (2005) and the studies reported in this thesis all utilised cellular Rac activation assays whereas Otomo et al. (2003) used in vitro exchange assays. Several
GEFs require specific components for their regulation that are only present in a cellular environment. For example the Rac GEF activity of SWAP-70 is minimal in in vitro
GDP dissociation assays but is greatly enhanced upon addition of phosphatidylinositol
3,4,5-triphosphate (PtdIns(3,4,5)P3) (Shinohara et al., 2002), and the activity of several other GEFs, including Vav, Sos and Tiam1, is enhanced by binding to phospholipids such as PtdIns(3,4,5)P3 (Das et al., 2000; Fleming et al., 2000; Han et al., 1998).
Therefore, it is possible that ALS2 may only act as a Rac GEF in vivo.
ALS2-dependent activation of Rac was found to require the DH domain, which has previously been identified as the region responsible for catalytic GEF activity in other
Rac GEFs such as Dbl (Hart et al., 1994). However, overexpression of the isolated
DH/PH region of ALS2 did not result in stimulation of Rac activity. This may be due to autoinhibition of the catalytic DH domain by residues of the PH domain. Indeed, this is known to occur for a number of GEFs including Sos1 and P-Rex1 (Nimnual et al.,
1998; Welch et al., 2002) although some isolated DH/PH constructs can act in a constitutively active manner; for example Fgd1(DH/PH) is a constitutively active Cdc42
GEF (Olson et al., 1996). Alternatively, regions of the full-length ALS2 protein other than the DH/PH domain may be necessary for regulation of the Rac GEF activity. For example, MORN motifs are implicated in association with the plasma membrane
(Takeshima et al., 2000) and this may be necessary for the Rac GEF activity of ALS2, as the activity of several Rac GEFs is dependent upon membrane-targeting (Whitehead et al., 1995; Whitehead et al., 1999). Although the regulatory regions of ALS2 are as yet
147 unidentified, this study shows that the full-length ALS2 protein is necessary for activation of Rac.
Finally, overexpression of ALS2 was found to activate PAK1, a major downstream effector of Rac1. The level of activation of PAK1 observed, as measured by phosphorylation of MBP, was approximately 1.5-fold. This is similar to the level of
PAK1 activation resulting from overexpression of the Rac1/Cdc42 GEF α-PIX in 293T human embryonic kidney cells (Yoshii et al., 2001). Thus, ALS2 functions as a Rac
GEF to activate PAK1. This activation requires a functional DH domain, as was demonstrated by overexpression of the non-functional ALS2ΔDH mutant.
Rac1 is involved in a variety of cellular activities including organisation of the actin cytoskeleton, regulation of cell proliferation, motility and survival, generation of ROS, and induction of both JNK and p38 signalling pathways (for reviews see Nobes and
Hall, 1995a; Van Aelst and D'Souza-Schorey, 1997). Of particular relevance to motor neuron disease is the role Rac plays in cell survival and death. Rac activates both apoptotic (through the activation of JNK) and anti-apoptotic (through direct interaction with PI(3)K and subsequent activation of the serine/threonine kinase Akt/PKB) pathways (Lee et al., 2004; Murga et al., 2002). Indeed, it has recently been reported that ALS2 mediates neuroprotection by sequential activation of Rac1, PI(3)K and Akt3 and this protection is specific against mutant SOD1 toxicity (Kanekura et al., 2005).
Six members of the PAK family (PAK1-6) have been identified; PAK1-3 are major downstream effectors of Rac and Cdc42, possessing a CRIB domain/p21 binding domain (PBD) which binds specifically to active Rac and Cdc42, whereas PAK4-6 differ significantly in their structure and regulation. For a review see (Bokoch, 2003).
148 Of the three serine/threonine kinases PAK1, PAK2 and PAK3, PAK1 has so far been the most widely studied, and is an important regulator of many cellular events including cytoskeletal dynamics and cell motility, transcription through SAPK/JNK and p38 kinase cascades, cell death and survival signalling and cell-cycle progression (Daniels et al., 1998; Frost et al., 1998; Frost et al., 2000; Frost et al., 1996; Robinson and Cobb,
1997; Schurmann et al., 2000; Sells et al., 1999; Sells et al., 1997; Zhang et al., 1995).
Several downstream effectors of PAK1 have been identified including Filamin,
Paxillin/PIX/PKL complex, the adaptor protein Nck, Merlin (a member of the
Ezrin/Radixin/Moesin family, which is thought to link actin filaments to the plasma membrane), Stathmin (a protein that regulates microtubule stability), and LIM kinase 1
(LIMK1) and myosin light chain kinase (MLCK), which control actin dynamics via their substrates Cofilin and myosin light chain, respectively (Chew et al., 1998; Manser et al., 1997). More recently, PAK1 has also been implicated in the regulation of microtubule dynamics by phosphorylating tubulin cofactor B (Vadlamudi et al., 2005).
Thus the combined studies presented here and in other reports show that ALS2 is a GEF for Rab5 and Rac and that ALS2-mediated Rac activity leads to activation of one of its downstream effectors PAK1 (Otomo et al., 2003; Topp et al., 2004; Kanekura et al.,
2005). Since ALS2 disease-causing mutations are all predicted to result in a loss of its function, it seems likely that perturbation of Rab5 and/or Rac/PAK signalling induces disease in the ALS2 kindreds.
149 Figure 4.1 ALS2 stimulates Rac activity GTPase activation assays were conducted in transfected CHO cells. (A, B, C) show representative assays for the three GTPases RhoA, Rac1 and Cdc42 respectively. Active RhoA was pulled-down from the lysates using GST-RBD and active Rac1 and Cdc42 were pulled-down with GST-PBD. The bound Rho, Rac and Cdc42 were then detected by immunoblotting (Active). The amounts of Rho, Rac and Cdc42 transfected into the cells were also detected by immunoblotting so as to demonstrate equal transfection efficiencies (Total). GST alone did not bind any GTPase in the transfected cells (No RBD/ No PBD). CHO cell samples were incubated with GDP as a negative control and with GTPγS as a positive control. CHO cells were transfected with the constitutively- active GEF mutants Net1ΔN, VavΔN and Fgd1DH/PH as positive controls for the Rho, Rac and Cdc42 assays respectively. (D) shows immunoblots from a Rac1 activation assay demonstrating that, unlike full-length ALS2, the isolated DH/PH domain of ALS2 does not activate Rac1. (E) shows histogram of fold-increases in GTPase activity, obtained from the average of 4 separate experiments for each GTPase. Error bars are SEM. One-Way ANOVA tests showed that ALS2 significantly increased Rac activity by 3.3 fold (p=<0.001) but not Rho (p=0.113) or Cdc42 (p=0.858) activities.
A CAT ALS2 Net1ΔN GDP GTPγS No RBD Active RhoA Total
B CAT ALS2 GDP GTPγS No PBD VavΔN Active Rac1 Total
C CAT ALS2 GTPγS GDP Fgd1DH/PH No PBD Active Cdc42 Total
D E CAT DH/PH ALS2 Active Rac1 Total
150 Figure 4.2 ALS2ΔDH does not activate Rac1 A mutant ALS2 (ALS2ΔDH), in which the central region of the Dbl-homology (DH) domain (residues 747-826) was deleted, was created by PCR-based site-directed mutagenesis. PCR products were analysed by agarose gel electrophoresis before (A; lane 1) and after (A; lane 2) digestion with Dpn1 to remove template DNA. The PCR products migrate at the predicted size of 10,185 bp. (A; lane 3) is linearised (by digestion with Xho1) 10,426 bp template DNA (pCIneo-ALS2). L is DNA ladder λHindIII. Following ligation, clones were isolated and subjected to restriction enzyme digest with Sal1 (B). Lane 1 is template (pCIneo-ALS2). Fragment sizes are 7,697 kb and 2,729 kb. Lane 2 is an incorrect clone, with fragments migrating at the same sizes as the template DNA. Lane 3 is a successfully-mutagenised clone (fragment sizes are 7,697 kb and 2,488 kb). L is DNA ladder λHindIII. The successfully-mutagenised clone (ALS2ΔDH) was transfected into CHO cells and the CHO cell lysate was subjected to SDS-PAGE and immunoblotting with anti-myc (9B11) antibody. (C) shows an immunoblot of lysates of CHO cells transfected with either ALS2 or ALS2ΔDH. ALS2ΔDH migrates slightly faster than ALS2. Rac activation assays were carried out in CHO cells co-transfected with Rac1 and either pCIneo-CAT (CAT) as a negative control, ALS2 or ALS2ΔDH. (D) shows an immunoblot of amounts of Rac1 bound to GST-PBD (Active). The amounts of Rac1 transfected into the cells were also detected by immunoblotting so as to demonstrate equal transfection efficiencies (Total). In contrast to ALS2, ALS2ΔDH did not activate Rac1. A B L 1 2 3 L 1 2 3 bp bp
23,130 23,130 9416 9416 6557 6557 4361 4361 2322 2027 2322 2027
C D kDa ALS2 ALS2ΔDH
CAT ALS2ΔDH ALS2 Active 181.8 Rac1 Total
115.5
151 Figure 4.3 ALS2 activates PAK1 PAK1 in vitro kinase assays were performed from CHO cells co-transfected with PAK1, Rac1 and either ALS2, ALS2ΔDH or vector pCIneo-CAT (CAT) as a negative control. For a positive control, cells were co-transfected with constitutively active Rac1 (L61Rac1). (-) and (+) refer to absence or presence of PAK1 immunoprecipitating antibody in the reactions; reaction mix (RM) contains no immunoprecipitation sample. (A) shows the Coomassie Blue-stained gel and (B) the corresponding autoradiograph of the samples. (C) is an immunoblot detecting the levels of PAK1 in the immunoprecipitation samples. The assays were repeated a further three times and one- way ANOVA tests revealed that the stimulatory effect of ALS2 and L61Rac1 on PAK1 activity was significant (p<0.01). (D) is a histogram showing fold increase in MBP phosphorylation with ALS2 and ALS2ΔDH compared with control (CAT). Data is pooled from five independent experiments. Error bars are SEM.
A RM CAT ALS2 ALS2ΔDH L61Rac kDa 115 - + - + - + - + - + 82 64 48.8 IgG 37.1 25.9 IgG 19.4 14.8 MBP
B Phospho-MBP
C PAK1
D
152
CHAPTER 5: ALS2 IS PRESENT IN NEURONAL
GROWTH CONES AND PROMOTES NEURITE
OUTGROWTH
153 5.1 Introduction
Rho family GTPases play a major role in organising the cytoskeleton and in particular, the actin cytoskeleton. In developing neurons, Rho, Rac and Cdc42 are present in growth cones of axons and dendrites where they regulate actin-myosin contractility to control neurite outgrowth and growth cone guidance (Li et al., 2000b; Threadgill et al.,
1997; Dickson, 2001; Nikolic, 2002). In some paradigms, Rac and Cdc42 promote whereas Rho inhibits neurite outgrowth. For example, in cell lines such as N1E-115 neuroblastoma cells and PC12 pheochromocytoma cells, and in primary cell systems such as chick retinal neurons and rat hippocampal neurons, Rac1 and Cdc42 promote neurite outgrowth whereas RhoA causes growth cone collapse and neurite retraction
(Albertinazzi et al., 1998; Da Silva et al., 2003; Jalink et al., 1994; Kita et al., 1998;
Schwamborn and Puschel, 2004; Sebok et al., 1999). However, a few studies using constitutively active (CA) and dominant negative (DN) mutants of Rac1 have revealed effects that contrast with those described above. For example, CA Rac1 has been found to decrease the length of the longest neurite in cultured rat cortical neurons (Kubo et al.,
2002) and DN Rac1 promotes neurite outgrowth in chick dorsal root ganglion (DRG) neurons (Fournier et al., 2003). These differential effects of GTPases on neurite development may be due to the type or age of the cells used in these studies and/or culture conditions. Additionally, it has been suggested that the varying results observed with the use of CA or DN GTPase mutants may reflect the importance of GTPase cycling between active and inactive states in the regulation of neuritogenesis. This idea is supported by studies in which both CA and DN mutants result in the same effect. For example, both CA and DN Rac1 mutants inhibit growth cone advance and neurite outgrowth in primary chick embryo motor neurons (Kuhn et al., 1998). Therefore, it is evident that tight regulation of GTPase activation (by GEFs and GAPs) is required for normal neuronal outgrowth.
154 A number of GEFs for Rho family GTPases have now been shown to be involved in the development of axons and dendrites including GEFT, Kalirin, Trio, Tiam1, Vav2, Vav3 and STEF (Aoki et al., 2005; Bryan et al., 2004; Chakrabarti et al., 2005; Estrach et al.,
2002; Kunda et al., 2001; Leeuwen et al., 1997; Matsuo et al., 2002; May et al., 2002;
Penzes et al., 2001). Therefore, since ALS2 functions as a Rac GEF (Chapter 4), its role in neurite initiation and outgrowth was investigated.
5.2 Materials and Methods
5.2.1 Antibodies and immunofluorescence microscopy
Affinity-purified polyclonal ALS2 antibody (see Chapter 3) was used at 1:100 dilution in 5% FBS/PBS for immunofluorescence. See Table 2.3 for details and working concentrations of Rac and α-tubulin antibodies. For staining of endogenous F-actin,
Alexa Fluor568-phalloidin was added to the coverslip (5 U/ml working concentration in
5%FBS/PBS) 30 minutes after incubation with ALS2 primary antibody and Alexa Fluor
488 secondary antibody. The coverslips were then washed in PBS for 3x 10 minutes and mounted in Vectashield. Conventional images were captured using a Zeiss Axioscop microscope and CCD camera (Princeton Instruments), and confocal images captured using a Zeiss LSM 510 META confocal microscope.
5.2.2 Preparation of mouse brain homogenates
Brain tissue samples from E15, E18, P1, P7, P11, P21 and 1yr old mice were homogenised in 1 ml of ice-cold tissue homogenisation buffer (see section 2.1.5.1) per
0.1 g of tissue using a dounce homogeniser, passed through a 28-gauge needle three times and centrifuged at 15,000xg for 30 minutes. Samples were taken for protein concentration determination following which a quarter volume of 5x SDS protein sample buffer was added to the supernatant and the samples were boiled for 10 minutes,
155 aliquoted and stored at -70°C until required. Protein samples were analysed as described in section 2.2.4.
5.2.3 Neurite outgrowth measurements
Cortical neurons grown on PDL-coated coverslips were co-transfected (using 1 µl
Lipofectamine 2000™ reagent per coverslip) at 2 DIV, with 1 µg of plasmid pEGFPC.1
(Clontech) expressing GFP plus 3µg of experimental or control plasmids (see section
2.2.6.3). Experimental and control plasmids included ALS2, ALS2ΔDH, dominant negative Rac1 (N17Rac), dominant negative Rab5 (N39Rab5), and pCIneo-CAT which was used to balance transfections so that all cells received the same numbers and amounts of plasmid. 24 hours after transfection, cells were fixed in 4% (w/v) paraformaldehyde (PFA) in TBS for 20 minutes, and processed for immunofluorescence
(to ensure all plasmids were expressing in the cells examined) as described in section
2.2.11. Images were analysed by counting the number of neurites, and measuring the length of the longest neurite per cell, using Metamorph® image analysis software.
Neurite lengths were determined as the distance from the edge of the cell body to the growth cone tip, as visualised by GFP fluorescence. Measurement of the longest neurite of each cell has been used in numerous studies on the effects of signalling cascades on neurite outgrowth (e.g. (Nikolic et al., 1996; Penzes et al., 2001)). Cells were analysed without knowledge of the transfected plasmids, and only healthy cells as judged by morphology (including nuclear staining with Hoechst 33258 (Sigma) to confirm that nuclei had a non-apoptotic appearance) were analysed.
156 5.2.4 Statistics
Statistical significance was determined using one-way ANOVA tests followed by Tukey post-hoc tests for pair-wise comparison. Differences were considered significant at p<0.05.
5.3 Results
5.3.1 Developmental expression of ALS2
The developmental expression of ALS2 was studied in mouse brain homogenates and in cultured cortical neurons using immunoblotting. Immunoblots of mouse brains aged from embryonic day 15 (E15) to 1 year revealed the presence of similar levels of ALS2
(Figure 5.1 A). Total protein levels in the samples were indicated by re-probing the immunoblots with α-tubulin antibody (DM1A). Others have also reported that ALS2 expression does not change markedly in the rodent brain post E10 (Devon et al., 2005).
However, in immunoblots of rat cortical neuron lysates there was a small but consistent decrease in ALS2 levels in >7 DIV cultures, even though levels of tubulin remained equal (Figure 5.1 B). This suggests that although levels of ALS2 do not vary in whole mouse brain samples, ALS2 may be developmentally regulated in specific neuronal populations such as cortical neurons.
5.3.2 ALS2 is present in growth cones of hippocampal neurons, where it is co- localised with Rac1 and F-actin
Application of the ALS2 antibody to 2 DIV rat hippocampal and cortical neurons produced prominent staining within cell bodies; labelling of punctate structures within neurites was also detected (Figure 5.2). These results are similar to those described by others in 7 DIV rat hippocampal neurons (Topp et al., 2004). However, in the 2 DIV
157 neurons, labelling within growth cones of both axons and dendrites was also detected
(Figure 5.2 and Figure 5.3).
Growth cones are highly specialised structures at the leading edge of developing neurites, which control the rate and direction of neurite outgrowth in response to environmental cues. Growth cones comprise two domains: the central (C) domain and the peripheral (P) domain. The C domain contains microtubules whereas the P domain is actin-rich containing the most motile structures, the lamellipodia and filopodia
(Letourneau, 1983; Tosney and Wessells, 1983). Co-staining for ALS2 and tubulin or actin revealed that ALS2 was present throughout the whole growth cone (Figure 5.3 B,
C). Double labelling for ALS2 and Rac1 was also performed, and this revealed a close overlap in the distributions of the two proteins in growth cones (Figure 5.3 A). The findings reported here complement the studies in Chapter 4 which demonstrate that
ALS2 is a Rac GEF and together they suggest that ALS2 may function in growth cone dynamics and neurite outgrowth.
5.3.3 Overexpression of ALS2 (but not ALS2ΔDH) promotes neurite outgrowth in rat embryonic cortical neurons via a Rac-dependent mechanism
A number of Rac GEFs are involved in various aspects of neuronal development, including neurite outgrowth and branching (Bryan et al., 2004; Leeuwen et al., 1997;
Lundquist et al., 2001; Penzes et al., 2001; Shin et al., 2002). As ALS2 has been shown to activate Rac (see Chapter 4), overexpression of myc-tagged ALS2 and ALS2ΔDH in
2-3 DIV cultured embryonic rat cortical neurons was utilised in order to investigate the involvement of ALS2 in neurite outgrowth and branching.
158 Prior to these experiments, the subcellular localisation of transfected ALS2 and
ALS2ΔDH was investigated in 3 DIV neurons (Figure 5.4). Both proteins showed a similar subcellular distribution to that of endogenous ALS2 in 2 DIV hippocampal neurons (Figure 5.2) where they localised to cell bodies, axons, dendrites and growth cones. Transfected ALS2 therefore localised to the appropriate cellular compartments where endogenous ALS2 resides.
In order to investigate the role of ALS2 in neurite outgrowth, ALS2 was overexpressed in 2 DIV embryonic rat cortical neurons together with GFP as a marker for neuronal cell shape. Neurite outgrowth was analysed 24 hours later, by comparison of the length of the longest neurite per cell with controls (cells that had been co-transfected with pCIneo-CAT and GFP). Comparison of the length of the longest neurite per cell has been used in several similar studies, for example (Nikolic et al., 1996) and (Penzes et al., 2001).
Overexpression of ALS2 in these neurons resulted in a significant (1.5-fold) increase in length of the longest neurite compared with control cells (Figure 5.5). This level of stimulation is similar to that observed by other Rac GEFs, for example GEFT (Bryan et al., 2004). As this stimulation of neurite outgrowth was hypothesised to be due to
ALS2’s Rac GEF activity, the mutant ALS2ΔDH, which is not functional as a Rac
GEF, was overexpressed in place of ALS2. In contrast to the effect seen with ALS2,
ALS2ΔDH did not stimulate neurite outgrowth (Figure 5.5). Therefore, ALS2 is thought to promote neurite outgrowth via activation of Rac. To pursue this hypothesis further, the experiments were repeated with co-expression of a dominant negative Rac1
(N17Rac1). This GTPase contains a substitution of Asn for Ser at position 17, and acts as a dominant negative protein by competing with endogenous Rac for binding to
159 upstream GEFs, but not interacting with downstream targets, thereby blocking Rac- dependent signalling pathways (Ridley et al., 1992). Such dominant negative GTPases have been used in numerous studies to dissect out pathways by which other GEFs stimulate neurite outgrowth (Bryan et al., 2004; Penzes et al., 2001). Co-expression of
N17Rac with ALS2 effectively blocked the ALS2-induced stimulation of neurite outgrowth (Figure 5.5), which supports the hypothesis that promotion of neurite outgrowth by ALS2 is via a Rac-dependent pathway.
ALS2 has also been shown to activate Rab5 via its VPS9 domain (Otomo et al., 2003).
To investigate whether stimulation of neurite outgrowth ALS2 could be linked to its ability to activate Rab5, a dominant negative Rab5 (N39Rab5) protein was co-expressed with ALS2 in these cells. Co-expression of N39Rab5 did not alter the stimulatory effect of ALS2 on neurite outgrowth (Figure 5.5). Thus, ALS2 stimulates neurite outgrowth via activation of Rac and independently of its function as a Rab5 GEF.
Expression of N17Rac alone had no effect on neurite outgrowth which is consistent with a number of other reports (Arakawa et al., 2003; Bryan et al., 2004). However, Rac has been shown to both promote and inhibit neurite outgrowth, and it is likely that such conflicting results are due to the different types of neurons, ages and culture conditions used for experimentation (e.g. (Jin and Strittmatter, 1997; Kita et al., 1998; Kuhn et al.,
1998; Lamoureux et al., 1997; Luo et al., 1994; Ruchhoeft et al., 1999)). Similarly, expression of N39Rab5 alone had no effect on neurite outgrowth.
160 5.3.4 Overexpression of ALS2 does not affect the number of neurites per cell, or the extent of neurite branching, in cultured cortical neurons
In some paradigms, Rac and PAK signalling has been shown to affect the initiation of neurite growth and branching (Allen et al., 2000; Hayashi et al., 2002; Leeuwen et al.,
1997; Threadgill et al., 1997). Indeed, several Rac GEFs have been found to affect the number of neurites per cell and/or the extent of neurite branching, including Tiam1
(Kunda et al., 2001) and GEFT (Bryan et al., 2004). Furthermore, expression of dominant negative Rac1 leads to a significant reduction in the number of primary dendrites in rat cortical neurons (Threadgill et al., 1997) and the extent of branching in
Xenopus retinal ganglion cells (Ruchhoeft et al., 1999), whereas Rac1 induces neuritogenesis and neurite branching in chick neural retinal cells (Albertinazzi et al.,
1998). Therefore, the effect of ALS2 on the number of neurites per cell and the extent of neuronal branching was investigated. This was measured by counting the total number of neurites emanating from the cell body, and by counting the number of neurite tips per cell, respectively. Overexpression of ALS2 did not affect the number of neurites, or the extent of branching (Figure 5.6). However, the results were only obtained from one timepoint during development (at 3 DIV) and therefore it remains possible that ALS2 may affect neurite promotion and/or branching at other stages in development.
5.4 Discussion
The studies presented here demonstrate that endogenous ALS2 is present in neuronal growth cones where it is co-localised with F-actin. This suggests that ALS2 may be involved in actin dynamics during neurite formation, outgrowth and/or pathfinding.
Furthermore, ALS2 co-localised with Rac1 which supports the hypothesis that ALS2 exerts its Rac GEF activity within the growth cone. Similarly, others have shown that
161 overexpressed ALS2 co-localises with Rac1 and with actin at leading membrane edges and membrane ruffles of NIH3T3 cells (Topp et al., 2004).
In the studies presented here, overexpression of ALS2 resulted in stimulation of neurite outgrowth (as measured by the length of the longest neurite per cell, compared with controls) and this effect was not seen with the non-functional mutant ALS2ΔDH.
Similarly, the promotion of outgrowth by ALS2 was not observed upon co-expression of dominant negative Rac1 (N17Rac1). Therefore, it can be concluded that ALS2 stimulates neurite outgrowth via its Rac GEF activity. In contrast, the effect of ALS2 on neurite length was not altered by co-expression of dominant negative Rab5 (N39Rab5), which suggests that the Rab5 GEF activity of ALS2 is not involved in this signalling pathway. Although Rac signalling has been shown to affect the initiation of neurite growth and branching, overexpression of ALS2 did not affect the number of neurites or extent of branching in these neurons. However, it is possible that ALS2 may be involved in such cellular events at a different stage in development.
In addition to Rac1, its downstream effector PAK1 is also present within the growth cone, where it too can function in neurite outgrowth, although (as in the case of Rac1) this may depend upon the stage of development and type of neuron (Daniels et al.,
1998). Rac1-PAK1 signalling has additionally been implicated in dendrite initiation in cultured embryonic mouse cortical neurons (Hayashi et al., 2002). ALS2 activates both
Rac1 and PAK1, and therefore the ALS2-mediated outgrowth observed in these experiments may involve PAK1 activation. This could be confirmed by use of dominant negative PAK1 proteins in the neurite outgrowth experiments to determine whether
PAK1 activation is essential for ALS2-mediated outgrowth to occur. Although overexpression of PAK1 has been reported to induce neurite outgrowth, it has been
162 suggested that this may be independent of its kinase activity (Daniels et al., 1998) and that PAK may be involved upstream of Rac1 (Obermeier et al., 1998). Similarly another
PAK subtype, PAK5, is highly expressed in brain, acts downstream of active Rac and
Cdc42, and leads to neurite outgrowth in a kinase activity-dependent manner (Dan et al.,
2002).
Molecular mechanisms of neurite outgrowth
Neurite (axon and dendrite) outgrowth defines neuronal shape, mediates neuronal pathfinding, and is essential for the establishment of synaptic connections during development, as well as being an important factor in neuronal regeneration following injury or neuropathological conditions. This complex process requires both co-ordinated cytoskeleton (actin and tubulin) remodelling and membrane expansion (mediated by vesicle targeting to the plasma membrane) in the growth cone.
Assembly and disassembly of polymeric actin filaments and polymerisation of tubulin into microtubules in the growth cone are essential for neurite extension. Several studies have demonstrated that polymerisation of actin occurs at the periphery of the growth cone, and depolymerisation occurs in the central domain, resulting in a retrograde flow of actin. Neurite elongation is driven by co-ordination of these actin forces with protrusion of microtubules into the central domain (Baas and Buster, 2004; Bradke and
Dotti, 1999; Li et al., 1994; Mallavarapu and Mitchison, 1999; Mitchison and
Kirschner, 1988; Suter and Forscher, 2000). Microtubules are transported into neurites and growth cones as short polymers, and live-cell imaging studies have shown that this transport is intermittent and infrequent, occurring at rates consistent with known molecular motor proteins and in both anterograde and retrograde directions (Wang and
Brown, 2002). Several studies have recently highlighted the importance of molecular
163 motors, in particular cytoplasmic dynein and myosins, in neurite outgrowth. It has been found that dynein transports microtubules from the centrosome into developing neurites
(Ahmad et al., 1998) as well as playing an important role in the retrograde transport of membranous vesicles, many of which contain growth factors (Karki and Holzbaur,
1999). Kinesins have also been implicated in neurite outgrowth (Sharp et al., 1997).
Myosins have been found to fuel retrograde actin flow within growth cones and cause microtubules to move retrogradely down the neurite, leading to neurite retraction
(Ahmad et al., 2000; Schaefer et al., 2002), as well as transporting vesicular elements to the growing neurite tip (Evans and Bridgman, 1995). Therefore, it is believed that a balance between the forces mediated by dynein and kinesins on microtubules and actin- myosin contractility controls neurite outgrowth.
Rho family GTPases are key regulators of actin-myosin contractility and are thought to exert their role in neurite outgrowth by controlling the retrograde flow of actin within the growth cone. Downstream targets of Rho family GTPases have been identified which are involved in actin polymerisation (N-WASP, SCAR/WAVE and VASP/Ena), actin depolymerisation (PAK, ROCK and LIMK) and myosin activity (PAK, ROCK and MLCK). For a review see (Huber et al., 2003). Additionally, it has been suggested that Rac-PAK signalling may affect microtubule dynamics, as PAK phosphorylates and inhibits the microtubule stabilising protein Stathmin in response to Rac signalling (Daub et al., 2001), although the role of this signalling pathway in neurite outgrowth has not yet been investigated. The involvement of Rac in signalling to both actin and microtubules therefore raises the possibility that Rac is involved in co-ordination between the actin and microtubule arrays within the growth cone.
164 Evidence is emerging that neurite outgrowth is also dependent on exocytic addition of plasma membrane at the leading edge of the growth cone (Dai and Sheetz, 1995).
Inhibitors of vesicle fusion proteins such as tetanus neurotoxin-insensitive vesicle- associated membrane protein (TI-VAMP), SNAP-25 and Syntaxin inhibit neurite outgrowth, whereas overexpression of TI-VAMP or Syntaxin promotes neurite outgrowth (Grosse et al., 1999; Hirling et al., 2000; Igarashi et al., 1996; Martinez-Arca et al., 2001; Osen-Sand et al., 1996), which suggests that exocytosis is a rate-limiting step. The Rab family of GTPases are important regulators at various stages of membrane traffic, and antisense oligonucleotides of the Rab guanine nucleotide dissociation inhibitor (GDI) inhibit axonal outgrowth of hippocampal neurons
(D'Adamo et al., 1998), although so far the only Rab GTPase that has been implicated in neurite outgrowth is Rab8 (Huber et al., 1995).
Neurite outgrowth and MND
Several models of MND show impaired neurite outgrowth. For example, embryonic motor neurons isolated from pmn mutant mice show severely impaired axonal growth
(axonal length is reduced by more than 50% compared to controls) (Bommel et al.,
2002). As described in Section 1.1.3.1, these mice have a homozygous mutation in the
Tbce gene involved in tubulin assembly and develop a MND resembling spinal muscular atrophy (SMA) (Matsuo et al., 2002). Interestingly, the morphology of the growth cones of pmn mouse motor neurons appears normal, whilst axonal swellings are present with an irregular pattern of immunoreactivity for tubulin and tau (Jablonka et al., 2004). Additionally, knockdown of survival motor neuron protein (Smn), which is mutated in SMA, causes defects in motor axon outgrowth and pathfinding in zebrafish
(McWhorter et al., 2003).
165 Mice heterozygous for either of two ENU-generated mutations, Loa and Cra1, demonstrate a motor neuron degenerative phenotype, and positional cloning has revealed that both disease-causing mutations are in the cytoplasmic dynein heavy chain
1 gene (Dnchc1). Furthermore, mutations in dynactin (p150 subunit) have been found to cause an autosomal dominant form of lower motor neuron disease in humans (Puls et al., 2003). Aside from its role in retrograde transport of vesicles, dynein is also required for proper transport of microtubules into developing neurites (Ahmad et al., 1998).
Mutations in dynein may therefore cause motor neuron disease by perturbation of such functions. Studies of cultured spinal motor neurons from SOD1G93A mutant mice have also revealed neurite outgrowth defects in these cells (Azzouz et al., 2000). Similarly, motor neuron-neuroblastoma cells expressing SOD1G93A showed neurite outgrowth defects (Lee et al., 2002). Such defects were rescued by treatment with copper chelators
(Azzouz et al., 2000) and by expression of heat shock proteins HSP70 and HSP40 leading to reduced protein aggregation (Takeuchi et al., 2002b). Furthermore, the transport of tubulin is impaired and axonal levels of the motor protein kinesin are reduced in SOD1G85R mice months before disease onset (Warita et al., 1999; Zhang et al., 1997) which raises the possibility that microtubules may not be transported to the axonal tip during neuronal development in these mice.
Finally, mutations in L1CAM have been found to cause hereditary spastic paraplegia
(SPG1) (Jouet et al., 1994). L1CAM is an axonal glycoprotein involved in neuronal growth and guidance (Bixby et al., 1988; Castellani et al., 2000). Thus, aside from the findings described in this Chapter, a number of other studies have implicated defective neurite outgrowth in motor neuron disease.
166 Figure 5.1 Developmental expression of ALS2 in mouse brain and cultured rat embryonic cortical neurons (A) is an immunoblot to demonstrate developmental expression of ALS2 in mouse brain from embryonic day (E) 15 to post-natal (P) 1 year. (B) is a similar immunoblot of rat cortical neurons cultured for 2-21 DIV as indicated. The samples were also probed for tubulin to demonstrate equal protein loadings.
A E15 E18 P1 P7 P11 P21 1Yr
ALS2
tubulin
B 2 4 7 11 14 21 ALS2
tubulin
167 Figure 5.2 Subcellular localisation of ALS2 in cultured embryonic neurons Embryonic rat hippocampal neurons were cultured and at 2 DIV were subjected to immunofluorescence with ALS2 antibody (to detect endogenous ALS2) and Alexa Fluor 568-phalloidin (to detect filamentous (F)-actin). Pictures shown are confocal z- projections. ALS2 colocalises with F-actin (Overlay), and is present in both axonal and dendritic growth cones (arrows). Punctate ALS2 staining can be seen in the cell body and neurites. Scale bar is 10 µm.
168 Figure 5.3 Subcellular localisation of ALS2 in neuronal growth cones Endogenous ALS2 is co-localised with Rac (A) and F-actin (B) in the growth cone and with tubulin in the axon shaft (C) of 2 DIV rat embryonic hippocampal neurons. Overlays of co-staining are shown. A is a confocal slice; B and C are confocal z- projections. Scale bars are 5 µm.
A
B
C
169 Figure 5.4 Transfected ALS2 and ALS2ΔDH display identical subcellular localisation to endogenous ALS2 Embryonic rat cortical neurons were transfected with either myc-tagged ALS2 or ALS2ΔDH at 2 DIV and immunostained with the anti-myc antibody 9B11 24 hours later. Both of the transfected constructs localised to the cell body, processes and growth cones (arrows), which is an identical localisation to that of endogenous ALS2 (as seen in Figure 5.4). Scale bar is 10 µm.
170 Figure 5.5 ALS2 promotes neurite outgrowth in cultured rat cortical neurons 2 DIV rat cortical neurons were transfected with GFP+control or experimental plasmids and the length of the longest neurite measured using GFP as a marker. Transfections were balanced with pCIneo-CAT (CAT) so that all cells received the same numbers and amounts of plasmid. (A) shows histogram of mean neurite length for each transfection condition as indicated. Data was obtained from 40-50 cells per transfection and the experiments were repeated at least 3 times. ALS2 but not ALS2ΔDH stimulates neurite outgrowth by 1.5-fold (p=<0.001) and this effect is lost upon co-transfection with N17Rac but not N39Rab5. (B) shows representative images of cells in the different transfections: (a), GFP+CAT; (b), GFP+ALS2; (c), GFP+ALS2ΔDH; (d) GFP+ALS2+N17Rac; (e), GFP+ALS2+N39Rab5; (f), GFP+N17Rac; (g), GFP+N39Rab5. Scale bar is 10 µm.
A
B
171 Figure 5.6 Overexpression of ALS2 does not affect the number of neurites or the extent of neurite branching in cortical neurons (A) shows histogram of the mean number of primary neurites per cell for rat cortical neurons co-transfected with either GFP+pCIneo-CAT (CAT), GFP+ALS2 or GFP+ALS2ΔDH at 2 DIV. (B) shows histogram of the mean neurite tip number per cell, which is a measure of total neurite number and an indication of the extent of neurite branching, for the same transfection conditions as above. Data was obtained from 40-50 cells per transfection and the experiments were repeated three times to obtain mean values.
A
B
172
CHAPTER 6: ALS2 IS A PHOSPHOPROTEIN
173 6.1 Introduction
DH/PH (Dbl-family) GEFs activate Rho family GTPases (for example Rho, Rac and
Cdc42) by de-stabilising GDP interactions and stabilising the nucleotide-depleted
GTPase resulting in GTP binding (driven by GTP concentration being approximately
10-fold higher than GDP in the cell). GEFs are extremely important in the regulation and integration of Rho family GTPase signalling pathways and it is therefore not surprising that GEFs themselves are highly regulated. Several ways in which GEFs are positively regulated have been identified including activation by direct interaction with
G protein subunits. This activation has been found to occur in DH/PH GEFs that contain a regulator of G-protein signalling (RGS) domain, such as p115RhoGEF (Hart et al.,
1998). A second means by which GEFs are activated is by binding to phospholipids; for example the activities of Vav, Sos and Tiam1 are all enhanced by binding to
PtdIns(3,4,5)P3, which is thought to occur via the PH domain (Das et al., 2000; Fleming et al., 2000; Han et al., 1998). An additional and common form of positive regulation of the DH/PH GEFs is phosphorylation by protein kinases.
Many GEFs are phosphorylated and activated in vivo by tyrosine and/or serine- threonine protein kinases, for example Vav, p85 β-PIX and GEF-H1 (Das et al., 2000;
Shin et al., 2002; Zenke et al., 2004). Phosphorylation by protein kinases may elicit a variety of effects on the target protein and therefore could alter GEF activity, specificity, stability, association with other proteins or localisation within the cell. To begin to understand the upstream mechanisms that regulate ALS2 activity, studies were carried out to investigate whether ALS2 is phosphorylated in vivo.
174 6.2 Materials and Methods
6.2.1 Cell culture and transfection
CHO cells were cultured on 10 cm dishes and transfected using Lipofectamine™, as described in Section 2.2.6.3. For ALS2 immunoprecipitation, cells were transfected with 8 μg myc-tagged ALS2 cDNA. For Rac activation assays, cells were co- transfected with 2 µg of myc-tagged Rac1 and 8 µg of either ALS2, ALS2ala or pCIneo-CAT cDNA (see Table 2.2 for information about expression vectors used).
ALS2ala is a mutant ALS2 in which 5 identified serine and threonine phosphorylated residues were mutated to alanine to preclude phosphorylation. For PAK1 kinase assays, cells were co-transfected with 2 µg myc-tagged PAK1, 2 µg Rac1 and 6 µg of either pCIneo-CAT as a negative control, ALS2, ALS2ala or constitutively active Rac1
(L61Rac1) as a positive control.
Primary embryonic (E18) rat cortical cultures were prepared as described in Section
2.2.6.2 and cultured on PDL-coated 10 cm dishes. Cells were harvested for immunoprecipitation at 7 DIV in 1 ml of cell lysis buffer (see Section 2.1.5.1 and
2.2.8.1) and treated with λ protein phosphatase as described (Section 2.2.4.4). For neurite outgrowth assays, cells were transfected at 2 DIV with either ALS2, ALS2ala or pCIneo-CAT using Lipofectamine 2000™ and harvested at 3 DIV, essentially as described in Section 5.2.3.
6.2.2 Mass spectrometric sequencing of ALS2
ALS2 was immunoprecipitated from transfected CHO cells, as described in Section
2.2.8.1. Samples were subjected to SDS-PAGE and gels were stained with Colloidal
Brilliant Blue G (Sigma). The appropriate bands were excised from the gel and sequenced by mass spectrometry. Briefly, bands were reduced, alkylated and digested
175 with either trypsin, chymotrypsin or Asp-N (Roche Molecular Biochemical) and peptides extracted with two wash cycles of 50 mM NH4HCO3 and acetonitrile, and then lyophilised and resuspended in 20 μl of 50mM NH4HCO3. Peptide digests were analysed by on-line liquid chromatography tandem mass spectrometry (LC/MS/MS).
Peptides were ionised by electrospray ionisation using a Z-spray source fitted to a
QTof-micro (Micromass, UK). The instrument was set to run in automated switching mode, selecting precursor ions based on their intensity and charge state, for sequencing by collision-induced fragmentation. The MS/MS analyses were conducted using collision energy profiles that were chosen based on the m/z and the charge state of the peptide and optimized for phosphorylated peptides. The mass spectral data was processed into peak lists containing the mass/charge (m/z) value of each precursor ion and the corresponding fragment ion m/z values and intensities. Data was searched against a custom-built database containing the full-length sequence of ALS2 using the
Mascot searching algorithm (Matrix Science, UK). Peptides and phospho-peptides of
ALS2 were identified as described (Perkinton et al., 2004; Standen et al., 2003). Mass spectrometry studies were performed “in-house” by Helen Byers (see
Acknowledgements).
6.2.3 Rac activation assay
CHO cells were harvested in 1 ml MLB buffer 24 hours after transfection, following 16 hours of serum-starvation. Rac activation assays were carried out using GST-PBD, as described in Section 4.2.3. Immunoblotting revealed similar levels of ALS2ala and wild-type ALS2 in the cell lysates.
176 6.2.4 PAK1 kinase assay
CHO cells were harvested 24 hours after transfection, following 16 hours of serum- starvation and PAK1 in vitro kinase assays were carried out as described in Section
4.2.4.
6.2.5 Neurite outgrowth measurements
Neurite outgrowth measurements were carried out as described in Section 5.2.3.
Immunofluorescence revealed that the localisation of ALS2ala in cortical neurons was identical to that of wild-type ALS2.
6.3 Results
6.3.1 ALS2 is a phosphoprotein
To begin to understand the upstream mechanisms that regulate ALS2 activity, studies were carried out to investigate whether ALS2 is phosphorylated in vivo. Endogenous
ALS2 was immunoprecipitated from cultured embryonic rat cortical neurons at 7 DIV and samples were then incubated in buffer either with or without λ protein phosphatase
(Section 2.2.4.4). The samples were subjected to SDS-PAGE and immunoblotting with antibodies to ALS2 or to phospho-serine/threonine-proline (MPM-2). The ALS2 protein immunoprecipitated was found to be phosphorylated at serine/threonine-proline residues (Figure 6.1).
The ALS2 protein was then sequenced to identify phosphorylation sites. ALS2 is a particularly low abundance protein (Yamanaka et al., 2003) and so to obtain sufficient protein for sequencing, it was isolated from transfected CHO cells. Using a combination of trypsin, chymotrypsin and Asp-N protease digestion, 81 % sequence coverage was obtained. Serines 277, 492, 1335, 1464 and threonine 510 were all unambiguously
177 identified as phosphorylation sites but a number of other phosphopeptides were also detected although the responsible residues could not be identified (Figure 6.2). The identified residues all precede a proline making them candidates for phosphorylation by proline-directed kinases such as those of the MAP kinase superfamily, the cyclin- dependent kinases (Cdks) and glycogen synthase kinase-3α/β (GSK3α/β).
6.3.2 Mutation of serine/threonine-proline sites to alanine does not affect the activity of ALS2
To enquire whether phosphorylation of these sites influenced ALS2 regulation of Rac activity, mutants were constructed from myc-tagged ALS2 by Michael Perkinton
(Institute of Psychiatry) in which the identified serine/threonine residues were altered to alanine to preclude phosphorylation (ALS2ala). However, this mutant had no discernible effect on ALS2 Rac activity, PAK1 activity or neurite outgrowth (Figure
6.3).
6.4 Discussion
Several GEFs are known to be activated by phosphorylation. One way in which phosphorylation activates GEF activity is by inducing a change in conformation. For example, Vav is thought to be autoinhibited by binding of its N-terminal to the catalytic
DH domain and this intramolecular inhibition is relieved upon phosphorylation by Src family tyrosine kinases (Aghazadeh et al., 2000). Phosphorylation has also been shown to promote binding of proteins that recognise the phosphorylated epitope, for example
GEF-H1 is phosphorylated by the serine/threonine kinase PAK1, and this promotes binding to 14-3-3 adaptor protein (Zenke et al., 2004). Furthermore in some cases, for example Vav, phosphorylation is important in regulating GEF binding to its upstream regulators and/or adaptor proteins, thereby allowing further phosphorylation and
178 activation to occur (for a review see (Bustelo, 2000)). Therefore identification of ALS2 in vivo phosphorylation sites may provide important information on the upstream components of ALS2 signalling pathways.
Here, I have demonstrated that ALS2 is a phosphoprotein and report the identification of five cellular phosphorylation sites. All of these sites are ser/thr-pro motifs, which makes them targets for proline-directed kinases. Interestingly, aberrant activation of two proline directed kinases, p38 (a stress-activated kinase) and Cdk5/p35 (a neuronal cyclin-dependent kinase) have been described in human ALS cases and mutant SOD1 transgenic models of ALS (Ackerley et al., 2004; Hu et al., 2003a; Hu et al., 2003b;
Nguyen et al., 2001; Raoul et al., 2002; Tortarolo et al., 2003; Zhu et al., 2002).
However, mutation of the 5 ALS2 phosphorylation sites identified in the studies reported here did not noticeably alter the effect of ALS2 on Rac1 activity. Whether phosphorylation regulates the Rab5 GEF activity of ALS2 may be addressed in future studies and when the full complement of ALS2 phosphorylation sites have been identified.
179 Figure 6.1 ALS2 is phosphorylated on serine/threonine-proline residues in embryonic rat cortical neurons ALS2 was immunoprecipitated from rat cortical neurons (7 DIV) with affinity-purified ALS2 antibody and treated with λ protein phosphatase (λppase). Immunoprecipitations were carried out with (+) or without (-) the immunoprecipitating (IP) antibody and subjected to SDS-PAGE and immunoblotting with the phospho-serine/threonine-proline antibody MPM2 (Phospho SP/TP; upper panel). Blots were then re-probed for ALS2 to show the relative amounts of ALS2 in the immunoprecipitates (ALS2; lower panel).
IP antibody - + + λppase - - +
Phospho SP/TP
ALS2
180 Figure 6.2 Phosphorylation of ALS2 The amino acid sequence of ALS2 is shown; residues covered by mass spectrometry are shown in bold and identified phosphorylation sites are boxed. The Rac GEF domain including the pleckstrin homology region is underlined. Ser277 was covered by peptide DSHCCPLGVTLTEGQAENHASTALpSPSTETL; precursor ion [M+3H]3+, m/z=1131.82. Ser492 was covered by peptide RLSLPGLLSQVpSPR; precursor ion [M+2H]2+, m/z=841.89. Thr510 was covered by peptide TVVLpTPTYSGEADALLPSIR; precursor ion [M+2H]2+, m/z=1092.5. Ser1335 was covered by peptide QHRDpSPEILSR; precursor ion [M+3H]3+, m/z=472.2. Ser1464 was covered by peptide TGKSDSRSEpSPEPGYVVTSSGL; precursor ion [M+2H]2+, m/z=1160.61.
MDSKKRSSTE AEGSKERGLV HIWQAGSFPI TPERLPGWGG KTVLQAALGV KHGVLLTEDG EVYSFGTLLW RSGPVEICPS SPILENALVG QYVITVATGS FHSGAVTDNG VAYMWGENSA GQCAVANQQY VPEPNPVSIA DSEASPLLAV RILQLACGEE HTLALSISRE IWAWGTGCQL GLITTAFPVT KPQKVEHLAG RVVLQVACGA FHSLALVQCL PSQDLKPVPE RCNQCSQLLI TMTDKEDHVI ISDSHCCPLG VTLTESQAEN HASTALSPST ETLDRQEEVF ENTLVANDQS VATELNAVSA QITSSDAMSS QQNVMGTTEI SSARNIPSYP DTQAVNEYLR KLSDHSVRED SEHGEKPVPS QPLLEEAIPN LHSPPTTSTS ALNSLVVSCA SAVGVRVAAT YEAGALSLKK VMNFYSTTPC ETGAQAGSSA IGPEGLKDSR EEQVKQESMQ GKKSSSLVDI REEETEGGSR RLSLPGLLSQ VSPRLLRKAA RVKTRTVVLT PTYSGEADAL LPSLRTEVWT WGKGKEGQLG HGDVLPRLQP LCVKCLDGKE VIHLEAGGYH SL ALTAKSQV YSWGSNTFGQ LGHSDFPTTV PRLAKISSEN GVWSIAAGRD YSLFLVDTED FQPGLYYSGR QDPTEGDNLP ENHSGSKTPV LLSCSKLGYI SRVTAGKDSY LALVDKNIMG YIASLHELAT TERRFYSKLS DIKSQILRPL LSLENLGTTT TVQLLQEVAS RFSKLCYLIG QHGASLSSFL HGVKEARSLV ILKHSSLFLD SYTEYCTSIT NFLVMGGFQL LAKPAIDFLN KNQELLQDLS EVNDENTQLM EILNTLFFLP IRRLHNYAKV LLKLATCFEV ASPEYQKLQD SSSCYECLAL HLGRKRKEAE YTLGFWKTFP GKMTDSLRKP ERRLLCESSN RALSLQHAGR FSVNWFILFN DALVHAQFST HHVFPLATLW AEPLSEEAGG VNGLKITTPE EQFTLISSTP QEKTKWLRAI SQAVDQALRG MSDLPPYGSG SSVQRQEPPI SRSAKYTFYK DPRLKDATYD GRWLSGKPHG RGVLKWPDGK MYSGMFRNGL EDGYGEYRIP NKAMNKEDHY VGHWKEGKMC GQGVYSYASG EVFEGCFQDN MRHGHGLLRS GKLTSSSPSM FIGQWVMDKK AGYGVFDDIT RGEKYMGMWQ DDVCQGNGVV VTQFGLYYEG NFHLNKMMGN GVLLSEDDTI YEGEFSDDWT LSGKGTLTMP NGDYIEGYFS GEWGSGIKIT GTYFKPSLYE SDKDRPKVFR KLGNLAVPAD EKWKAVFDEC WRQLGCEGPG QGEVWKAWDN IAVALTTSRR QHRDSPEILS RSQTQTLESL EFIPQHVGAF SVEKYDDIRK YLIKACDTPL HPLGRLVETL VAVYRMTYVG VGANRRLLQE AVKEIKSYLK RIFQLVRFLF PELPEEGSTI PLSAPLPTER KSFCTGKSDS RSESPEPGYV VTSSGLLLPV LLPRLYPPLF MLYALDNDRE EDIYWECVLR LNKQPDIALL GFLGVQRKFW PATLSILGES KKVLPTTKDA CFASAVECLQ QISTTFTPSD KLKVIQQTFE EISQSVLASL HEDFLWSMDD LFPVFLYVVL RARIRNLGSE VHLIEDLMDP YLQHGEQGIM FTTLKACYYQ IQREKLN
181 Figure 6.3 Mutation of ALS2 phosphorylation sites does not affect ALS2 Rac GEF activity, PAK1 activation or neurite outgrowth Ser277, Ser492, Thr510, Ser1335 and Ser1464 in ALS2 were all mutated to alanine to preclude phosphorylation, generating the mutant “ALS2ala”. ALS2ala displayed the same functional properties as wild-type ALS2 in these studies: (A) shows Rac activation assays in CHO cells co-transfected with Rac1 and either pCIneo-CAT (CAT) as a negative control, ALS2 or ALS2ala. Active Rac was pulled down from lysates using GST-PBD and detected by immunoblotting (Active). The amounts of total Rac transfected into the cells were also detected by immunoblotting so as to demonstrate equal transfection efficiencies (Total). (B) shows PAK1 in vitro kinase assays that were performed from CHO cells co-transfected with PAK1, Rac1 and either CAT (as a negative control), ALS2 or ALS2ala. (-) and (+) refer to absence or presence of PAK1 immunoprecipitating antibody in the reactions; reaction mix (RM) contains no immunoprecipitation sample. “Total PAK1” is the relative amounts of PAK1 immunoprecipitated in the (+) samples. 2 DIV rat cortical neurons were transfected with either GFP+CAT (control condition), GFP+ALS2 or GFP+ALS2ala, and the length of the longest neurite was measured using GFP as a marker. (C) shows histogram of mean neurite length for each transfection condition as indicated. Data was obtained from 40- 50 cells per transfection and the experiments were repeated three times.
A B
C
182
CHAPTER 7: DISCUSSION & FUTURE
DIRECTIONS
183 7.1 Summary of findings
In the studies presented here, a polyclonal antibody to ALS2 was generated and used to investigate ALS2 localisation in the mammalian central nervous system. ALS2 was found to be present in a number of neuronal populations in the brain including neurons of the cortex, hippocampus and cerebellum, and in motor neurons of the spinal cord.
Since ALS2 contains a DH/PH domain, which is a hallmark of Rho family GEFs, assays were performed to investigate whether ALS2 activates the three most widely- studied Rho-family GTPases RhoA, Rac1 and Cdc42 in vivo. ALS2 was shown to activate Rac1 but not RhoA or Cdc42. Rac1 activation required the DH domain, as shown by use of an ALS2 mutant (ALS2ΔDH). Next, the activation of a major downstream effector of Rac1 (PAK1) by ALS2 was investigated. ALS2 was found to activate PAK1 in in vitro kinase assays. Rac1 is known to regulate neurite outgrowth during development. Experiments were therefore performed to investigate the role of
ALS2 in neurite outgrowth, and overexpression of ALS2 in rat embryonic cortical neurons was found to stimulate neurite outgrowth. Immunofluorescence showed that endogenous ALS2 is present in the cell body and processes, and is co-localised with F- actin and Rac in the growth cones, of rat embryonic cortical and hippocampal neurons, which further implicates a role for the protein in neurite outgrowth. Finally, the phosphorylation state of ALS2 was investigated, as the activity of GEFs is often regulated by means of phosphorylation. ALS2 was found to be a phosphoprotein in vivo and five serine/threonine phosphorylation sites were identified. Mutation of these sites to alanine did not affect the function of the ALS2 protein in Rac1 activation assays,
PAK1 kinase assays, or neurite outgrowth experiments. The findings reported in this thesis provide an insight into ALS2 function, which may contribute to our understanding of the molecular mechanisms involved in motor neuron disease.
184 7.2 The role of alsin/ALS2 in motor neuron disease
Nine disease-causing mutations in ALS2 have been described in nine different autosomal recessive kindreds. All of the affected individuals are homozygous for the mutation and develop a slowly progressive ascending upper motor neuron disorder which presents with a lower limb spasticity and can have onset in infancy, childhood or adolescence. All mutations result in premature translational termination and truncation of the full-length native protein (Devon et al., 2003; Eymard-Pierre et al., 2002; Gros-
Louis et al., 2003b; Hadano et al., 2001; Yang et al., 2001). The neuropathology has not been described but in a clinically similar genetic disorder, hereditary spastic paraplegia due to mutations in SPG4, upper motor neurons projecting in the corticospinal tract develop a dying-back axonopathy (Wharton et al., 2003). The recessive nature of ALS2 and truncation mutations suggest that the disorder is caused by a loss of normal ALS2 function, but the function of ALS2 is unclear and the precise mechanisms by which this leads to clinically selective motor neuron degeneration are unknown.
One possibility is that loss of ALS2 Rab5 GEF function perturbs membrane trafficking so as to induce disease. Indeed, disruptions to membrane trafficking and the Golgi apparatus are seen in mutant SOD1 transgenic mouse models of ALS and recently mutations in the vesicle-trafficking protein VAPB have been shown to cause late onset spinal muscular atrophy and ALS (Mourelatos et al., 1996; Nishimura et al., 2004).
There may even be mechanisms linking mutant SOD1 and ALS2 forms of ALS
(Kanekura et al., 2005; Kanekura et al., 2004).
Another possibility is that loss of ALS2 Rac1 GEF function compromises proper development of motor neurons making them more susceptible to later toxic insults.
185 Indeed, upper motor neurons are the largest in the central nervous system with the longest axons, and so any defect in axonal growth induced by loss of ALS2 function is likely to be most severe in these cells. Interestingly, Rho family GTPase and PAK family kinase signalling pathways have both been implicated in another nervous system disorder that is thought to result from disorganisation of neuronal network formation.
Mutations in Oligophrenin1 (which encodes p190 RhoGAP) and the Rac GEF α-PIX are both involved in X-linked mental retardation (MRX), which is classed as a developmental disorder, and is associated with an immature morphology of synaptic spines (Billuart et al., 1998; Xiao et al., 2003). MRX is also caused by point mutations in the brain-specific PAK isoform PAK3. One such mutation, R67C, lies in the conserved region reported to be critical for GTPase binding and PAK activation, which suggests that GTPase-dependent PAK activation is affected in the disease (Allen et al.,
1998; Bienvenu et al., 2000; Knaus et al., 1998). Furthermore, William's syndrome is a neurological condition characterized by mild mental retardation and defects in visuo- spatial cognition, and this syndrome results from deletions of the gene for LIM kinase.
LIM kinase acts downstream of both Rac-PAK and Rho-Rho kinase (ROCK/ROK) to phosphorylate Cofilin, regulating reorganization of the actin cytoskeleton within the growth cone (Frangiskakis et al., 1996; Kuhn et al., 2000; Maekawa et al., 1999).
Additionally, it is now known that endocytic trafficking and cytoskeletal dynamics are intimately linked and therefore it is possible that ALS2 may co-ordinate both Rac and
Rab5 activities to regulate actin remodelling in endocytic events in motor neurons
(Lanzetti et al., 2004; McPherson, 2002; Schafer, 2002). Such events may include endocytosis of AMPA receptors as this process is thought to involve actin reorganisation as well as endocytic trafficking (Zhou et al., 2001) and recently research was presented at the Society for Neuroscience 2004 meeting (Washington DC, USA)
186 stating that ALS2 interacts with GRIP1, an adaptor protein that specifically binds to the
C-terminals of AMPA receptor subtypes GluR2 and GluR3, via its RCC1 domain (Cai et al., 2004).
However, none of the above hypotheses are mutually exclusive. Whatever the mechanisms by which mutations in ALS2 induce disease, a proper understanding of
ALS2 function is likely to assist in unravelling the aberrant molecular processes by which motor neurons die in ALS, and therefore the work presented in this thesis provides a valuable contribution to this area of research.
7.3 Future directions
The results presented in this thesis provide novel information on the function of ALS2 and provide a starting point for several lines of study. Some suggestions for future research are given below.
1. All of the ALS2 disease-causing mutations described so far result in a predicted loss of function of the protein, as indicated by the recessive nature of inheritance and the finding that the mutant proteins are rapidly degraded when overexpressed in human cells (Yamanaka et al., 2003; Yang et al., 2001). Therefore, RNAi or knockout rodent models will be particularly useful tools in the analysis of the role of ALS2 in motor neuron disease. Although it has recently been reported that motor neurons of 20 month- old ALS2 knockout mice do not display any obvious morphological defects (Cai et al.,
2005), it would be interesting to investigate various aspects of neurogenesis in these cells, including growth cone morphology, neurite outgrowth and branching, and synapse formation.
187 2. The results in Chapter 5 show that overexpression of ALS2 enhances neurite outgrowth in a Rac1-dependent manner. It is possible that PAK1 is involved in this signalling pathway, especially as ALS2 activates PAK1 (as shown in Chapter 4). Co- expression of a dominant negative PAK1 construct with ALS2 in the neurite outgrowth experiments could be used to investigate this possibility further.
3. In Chapters 4 and 5 it was shown that ALS2 is a GEF for Rac1 and is present at all developmental stages in cultured cortical neurons, where it exhibits a punctate staining in the cell body, axon and dendrites. Furthermore, it is co-localised with Rac1 and F- actin in the neuronal growth cone, which raises the possibility that it is involved in regulation of actin assembly via activation of Rac1. Indeed as shown in Chapter 5, overexpression of ALS2 affects neurite outgrowth, and this is likely to be mediated by actin reorganisation. Dendritic spine formation and stability are also largely determined by the actin cytoskeleton, and numerous studies have implicated Rac1 in promotion of these events (Luo et al., 1996; Nakayama et al., 2000). Several Rac GEFs are enriched in spines, and their overexpression often results in an increase in the number and size of spines, for example Kalirin-7 (Penzes et al., 2001), GEFT (Bryan et al., 2004) and α-
PIX (Zhang et al., 2005). In contrast, the Rac GEF still life (SIF) localizes in the presynaptic periactive zone of the Drosophila neuromuscular junction; this suggests that the SIF-Rac pathway is involved in synaptic development (Sone et al., 2000).
Differential fractionation studies performed on extracts from rat cerebellum have revealed that a proportion of endogenous ALS2 protein is localised to the fraction containing synaptosomal membranes (Topp et al., 2004) and therefore it would be interesting to investigate whether ALS2 is present in dendritic spines or in the presynaptic terminal, and whether it functions to regulate actin-based events such as receptor endocytosis or dendritic spine morphogenesis.
188 4. The results in Chapter 6 show that ALS2 is a phosphoprotein in vivo, and five serine/threonine-proline phosphorylation sites were identified by mass spectrometry.
Although it was shown that mutation of these sites to alanine (to preclude phosphorylation) did not affect the ability of ALS2 to activate Rac1 or PAK1, or to enhance neurite outgrowth, there is still potential for these sites to be important in the regulation of ALS2 function. For example, phosphorylation of these sites may affect
ALS2 Rab5 GEF function or other cellular events mediated by its Rac GEF activity such as its recently discovered involvement in an anti-apoptotic pathway (Kanekura et al., 2005). These sites are phosphorylated in ALS2 expressed in CHO cells, however it would be interesting to identify further sites (or to confirm phosphorylation of these sites) in ALS2 obtained from either rodent brain or cultured neuronal cells.
Furthermore, the activity of many GEFs is regulated by tyrosine phosphorylation.
Interestingly, tyrosine phosphorylation activates several GEFs that are involved in growth cone motility and neurite outgrowth, for example Trio, Ephexin1 and p85 β-PIX
(Lanier and Gertler, 2000; Sahin et al., 2005; Shin et al., 2004). Although no phosphorylated tyrosines have as yet been identified in ALS2, the full-length protein contains 57 tyrosine residues. Therefore, it would be interesting to discover whether
ALS2 is also activated by tyrosine phosphorylation in vivo. Initial studies to investigate this possibility could include probing of immunoprecipitated ALS2 on immunoblots with antibodies that detect phosphotyrosine residues.
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