THE ROLES OF ERK1 AND ERK2 MAP KINASE IN
NEURAL DEVELOPMENT AND DISEASE
by
IVY SAMUELS
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Thesis Advisor: Dr. Gary E. Landreth
Department of Neurosciences
CASE WESTERN RESERVE UNIVERSITY
August 2008
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
Ivy S. Samuels ______candidate for the Ph.D. degree.*
Stephen O’Gorman ______chair of the committee
Gary E. Landreth ______
Robert H. Miller ______
Ruth E. Siegel ______
06/19/2008
* We also certify that written approval has been obtained for any proprietary material contained therein.
For my teachers, past and present, who inspired me.
TABLE OF CONTENTS
Table of Contents………………………………………………………………….1
List of Tables………………………………………………...……………………4
List of Figures……………………………………………………………………..5
Acknowledgements……………………………………………………………..…7
Abstract…………………………………………………………………………....9
Chapter 1: Introduction..……………………………………………………….11
Mitogen-Activated Protein Kinase Signal Transduction……………………..12
Structural Deformities of the Cerebral Cortex and MR: A Historical
Perspective…………………………………………………………………....17
Controlling Cortical Development: The cell cycle…………………………...19
ERK1/2 and the Cell Cycle…………………………………………………...22
Cortical Development I: The Expansion of Progenitor Cells………………...26
Cortical Development II: Neurogenesis……………………………………...28
Modes of Neuronal Generation…………………………………………..29
Cortical Malformations and Disrupted Division…………………………30
Cortical Development III: Migration………………………………………....32
Interkinetic migration during Neurogenesis……………………………...32
Cortical Malformations and Disrupted Migration………………………..34
Cortical Development IV: Differentiation of Neurons and Synaptogenesis....35
Molecular Mechanisms of Cortical Development…………………………....36
MAPK and Cell Fate Determination…………………………………………39
Disruptions in upstream elements of the MAPK signaling cascade lead to
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anatomical and behavioral impairments in mice………………………....41
Aberrant ERK1/2 Signaling leads to Developmental Disorders…………45
ERK in Learning and Memory……………………………………………….47
Individual Roles for the ERK1 and ERK2 Isoforms………………………....49
Research Goals……………………………………………………………….54
Literature Cited……………………………………………………………….56
Chapter 2: Conditional Inactivation of ERK2 Identifies Its Key Roles in Cortical
Neurogenesis and Cognitive Function………………….……………………...68
Abstract………………………………………………………………………69
Introduction…………………………………………………………………..69
Materials and Methods.………………………………………………………72
Results………………………………………………………………………..81
ERK2 is required for normal development of the cerebral cortex……….81
ERK2 inactivation alters the cellular composition of the cortex ………..83
ERK2 inactivation alters the dynamics of neurogenesis………………...86
ERK2 CKO neural progenitors generate fewer neurons in vitro…...... 88
Mature ERK2 CKO mice display impaired associate learning………….89
MAPK1/ERK2 deficiency is associated with learning deficits………….90
Discussion…………………………………………………………………...92
Figures………………………………………………………………………98
Literature Cited……………………………………………………………..128
Chapter 3: ERK1 and ERK2 are required for cell cycle regulation of neural progenitor cells………………………………………………………………...132
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Abstract……………………………………………………………………..132
Introduction…………………………………………………………………133
Materials and Methods………...……………………………………………136
Results……..………………………………………………………………..140
ERK1/2 DKO mice die embryonically, but display normal brain
morphology……………………………………………………………..140
Loss of both ERK1 and ERK2 does not exacerbate the alterations in cellular
composition identified in the ERK2 CKO cortex……………………....141
ERK1/2 DKO NPCs display inhibited cell cycle progression attributable to
reduced in cyclin D2 and increased p27kip1 expression…...…………..142
Discussion…………………………………………………………………..144
Figures………………………………………………………………………150
Literature Cited……………………………………………………………...166
Chapter 4: Discussion…………………..……………………………………...169
Isoform specific expression regulates isoform specific function…………...169
ERK and Neurogenesis/the Cell Cycle……………………………………..174
ERK and Cognitive Function/Mental Retardation…………………………179
ERK and 22q11 Deletion Syndrome……………………………………….182
22q11 and psychopathology……………………………………………183
22q11 physical pathology………………………………………………185
ERK and Affective Function/Mood Disorders……………………………..187
Literature Cited……………………………………………………………..191
Chapter 5: Literature Cited…………………………………………………..198
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LIST OF TABLES
Table 3-1 Inactivation of ERK2 does not alter the morphology of Layer III cortical neurons………………………………………...………………………………..127
Table 3-2 Inactivation of ERK2 does not alter the morphology of Layer V cortical neurons………………………………………………………………………….127
4
LIST OF FIGURES
Figure 1-1 MAPK Signal Transduction…………………………………………13
Figure 1-2 General cycle cycle regulation……………………………………….20
Figure 1-3 Temporal development of the murine cerebral cortex...……………..21
Figure 1-4 ERK1/2 regulates cell cycle entry and progression………………….23
Figure 1-5 Modes of Cellular Generation in the Developing Cerebral Cortex….30
Figure 1-6 ERK1/2 Activity Regulates Cell Fate Determination………………..40
Figure 1-7 Mutations in elements of the MAPK Signaling Cascade lead to MR and CFC Syndromes………………………………..……………………………46
Figure 2-1 Generation of ERK2 Conditional Knockout mice…………………...99
Figure 2-2 ERK2 CKO mice display reduced cortical thickness………………101
Figure 2-3 Loss of ERK2 expression and activity in ERK2 CKO cortices…….103
Figure 2-4 Inactivation of ERK2 in neural progenitor cells results in generation of fewer cortical neurons…………………………………………………..……105
Figure 2-5 Inactivation of ERK2 in neural progenitor cells results in the presence of more astrocytes within the cerebral cortex…………………………107
Figure 2-6 ERK2 CKO mice display changes in the dynamics of NPC proliferation……………………………………………………………………..109
Figure 2-7 ERK2 CKO NPCs exhibit reduced neuronal generation…………....111
Figure 2-8 ERK2 CKO cortical progenitors generate more astrocytes in the presence of gliogenic stimuli…………………………………………………...113
Figure 2-9 Male ERK2 CKO mice have deficits in associative learning……....115
5
Figure 2-10 ERK2 expression is abolished in NPCs beginning at E13.5.………117
Figure 2-11 ERK2 expression is reduced in the hippocampus, but not the amygdala……………………………………………………………….………..119
Figure 2-12 The ERK2 CKO cortex does not exhibit apoptosis……….……….121
Figure 2-13 The number of Pax6 and Tbr2 immunoreactive cells at E14.5 is not altered by conditional inactivation of ERK2…………………………………....123
Figure 2-14 Patients with deletions of distal 22q11 have reduced ERK2 levels..125
Figure 3-1 Generation of the ERK1/2 Double Knockout Mouse………………151
Figure 3-2 ERK1/2 Double knockout mice die embryonically but display normal brain morphology...……………………………………………………………..153
Figure 3-3 Loss of ERK1/2 expression and activity in DKO cortices………….155
Figure 3-4 Inactivation of ERK1 and ERK2 in neural progenitor cells results in the generation of fewer cortical neurons…………………………………..……157
Figure 3-5 DKO NPCs exhibit reduced neuronal generation…………………..159
Figure 3-6 DKO neural progenitor cells exhibit reduced proliferation throughout neurogenesis………………………………………………………………….…161
Figure 3-7 DKO mice exhibit reductions in number of mitotic Tbr2+ intermediate progenitor cells………………………………………………………………….163
Figure 3-8 Decreased proliferation of DKO NPCs is mediated by reductions in cyclin D2 and increased p27 expression……………...………………………...165
Figure 4-1 Mutations in elements of the MAPK Signaling Cascade lead to MR
and CFC Syndromes…………...……………………………………………….179
6
ACKNOWLEDGEMENTS
To Dr. Gary Landreth. Without his unwavering belief in my ideas, his enthusiasm for science, and his ability to follow the biology, this dissertation would not have been possible. His patience, understanding and caring have nurtured my life and my career.
To my thesis committee, Dr. Robert Miller, Dr. Stephen O’Gorman, Dr. Ruth Siegel and
Dr. Karl Herrup, their ideas and advice were extremely helpful.
To the Landreth Lab. It has been said that this is a special place to work. It is so much more, because it is a home and a family. I can’t thank them enough for their advice, challenges and support in all endeavors. I would especially like to express my gratitude to Colleen Karlo for her initial and continuous contributions to this project, her endless support, and her contagious laughter.
To the Alzheimer Laboratory. I didn’t study AD, but they didn’t care. I would like to thank them for all their advice, help and the fun times.
To my parents, who recognized and nurtured my potential and encouraged me to become the first doctor in our family. And to Vicki and Mike. The love and support of my family inspires me everyday.
7
To Randall, who is not only an amazing scientist, but also my best friend and cheerleader. I am most appreciative of his support, advice, and love. And to Zander,
Malone and Callie, my munchkins.
To my immediate, extended, and “Cleveland” families and friends. They fill my life with happiness and were always interested, supportive and encouraging. I would specifically like to thank Jessica Lerch, Brandy Wilkinson, Barbara Schwartz, the Kossiver’s, and the
York’s for helping to make Cleveland a wonderful place to work and live.
8
The Roles of ERK1 and ERK2 MAP Kinase in Neural Development and Disease
Abstract
by
IVY SAMUELS
The Mitogen Activated Protein kinases, ERK1 and ERK2 are critical intracellular signaling intermediates in proliferation and differentiation. ERK1/2 are also essential for memory, learning, and synaptic plasticity in the brain. ERK1/2 activity in neural progenitor cells (NPCs) is required for neuronal cell fate determination; yet little is known about their isoform-specific functions during neural development. We have previously shown that inactivation of ERK1 does not initiate severe phenotypic or
behavioral deficits in mice, suggesting a redundant role for ERK2. Inactivation of ERK2 causes mice to die during embryogenesis. We have therefore examined the role of ERK2 and the combined actions of both isoforms in neural development by targeted and conditional inactivation of mapk3/ERK1 and mapk1/ERK2 in murine NPCs. Loss of
ERK2 alone resulted in a reduction in cortical thickness attributable to impaired proliferation of NPCs and the generation of fewer neurons. Mutant NPCs remained in an undifferentiated state until gliogenic stimuli induced their differentiation, resulting in the generation of more astrocytes. The ERK2 mutant mice displayed profound deficits in associative learning. Importantly, we identified patients with a 1 Mb microdeletion on chromosome 22q11.2 encompassing the MAPK1/ERK2 gene. These children have reduced ERK2 levels and exhibit microcephaly, impaired cognition, and developmental delay. These findings demonstrate an important role for ERK2 in cellular proliferation
9
and differentiation during neural development as well as in cognition and memory formation.
Simultaneous inactivation of both ERK1 and ERK2 (ERK1/2 DKO mice) lead to a
slightly more severe phenotype than loss of ERK2 alone; however, the loss of both isoforms did not significantly alter the composition of the cortex. ERK1/2 DKO mice
display similar changes in cell fate determination as ERK2 conditional knockouts where
fewer neurons are generated and a greater number of NPCs remain undifferentiated.
These changes are due to elevated levels of p27 and reductions in cyclin D2. Our
findings indicate that in vivo, ERK1 can compensate for ERK2 late in neurogenesis, but
that the normal function of ERK1 in neural development is subordinate to ERK2.
Moreover, ERK1/2 are indispensable for proper neural progenitor cell function as well as
learning and cognition.
10 INTRODUCTION
Congenital developmental abnormalities within the brain are the predominant cause of mental retardation and intellectual disabilities in humans. The incidence of these
disorders is between 1 and 3 percent of children and young adults as classified by the
Diagnostic and Statistical Manual of Mental Disorders- 4th Ed. (DSM-IV). The diagnosis
for mental retardation is defined by an overall intelligence quotient lower than 70, motor
dysfunction, and a lack of adaptive behavior. Investigation into the mechanisms leading
to cognitive impairment has yielded a collection of almost 300 causative genes (Inlow
and Restifo 2004). Importantly, nearly all of the genetic mutations giving rise to varying
degrees of mental retardation are from the inactivation or functional impairment of
proteins involved in cortical development and plasticity (Chelly et al., 2006).
Furthermore, recent evidence suggests that size and gyrification of the cerebral cortex is
associated with intellectual capacity (Shaw et al., 2006; Witelson et al., 2006). These
data suggest that a delineation of the causes of abnormal cortical development and
function is the key to understanding mental retardation.
The mammalian cerebral cortex is produced through strict temporal and spatial control of
cellular proliferation. Approximately 21.5 billion neurons (Williams and Herrup, 1988)
populate the mature human cortex, and about 35 million neurons in the mouse (Vaccarino
et al., 1999). The generation of these cells depends on an intricate choreography of the
proliferation of founder progenitor cells and committed neuronal progenitors within the
germinal centers of the brain. Therefore, control of the cell cycle in each cellular
11 population is extremely important in development of the cerebral cortex, and more broadly, to cognitive function. A mitogen is defined as a substance which induces a cell to commence cellular division, triggering mitosis. The canonical Mitogen-Activated
Protein Kinases were therefore the target of our investigation. This thesis demonstrates
that the prototypical mitogen activated protein kinases, ERK1 and ERK2, act to regulate
proper cortical development through maintenance of the cell cycle in neural progenitor
cells and that ERK2 is the predominant isoform responsible for this function. Moreover,
this work demonstrates that reduced expression of only the ERK2 isoform is sufficient to
induce physical and behavioral characteristics of mental retardation.
Mitogen-Activated Protein Kinase Signal Transduction
The Mitogen-Activated Protein Kinase (MAPK) superfamily is comprised of a conserved
group of enzymes which regulate a diverse range of cellular processes including
proliferation, migration, differentiation, stress, and survival. In mammals, the MAPKs
are divided into three major groups based on sequence similarity, activity, and substrate
specificity. They are the Extra-cellular signal Regulated protein Kinases (ERK 1 and 2;
also termed p44 and p42 MAPK, respectively), the Stress activated protein kinases, also
known as the Jun N terminal kinases (JNK1, 2 and 3), and the p38 MAPKs (α, β and γ).
Enzymatic activity of the MAPKs is tightly regulated by phosphorylation status of the proteins, which is mediated by kinases and phosphatases. Activation of the MAP Kinase superfamily occurs through a classical three-tier kinase cascade whereby extracellular cues stimulate cell surface receptors which then propagate intracellular signaling to the cytoplasm and nucleus (Figure 1-1).
12 In the developing organism, growth factors are a primary mitogenic stimulus and act largely through receptor tyrosine kinases which signal to elements that control the cell cycle. Receptor tyrosine kinases are located on the plasma membrane and
Figure 1-1: Signal transduction from the plasma membrane to the nuclueus is mediated by the canonical Mitogen Acticated Protein Kinases, ERK1/2.
become activated upon ligand binding, provoking them to undergo dimerization, autophosphorylation for full transactivation, and subsequent conformational changes.
These changes result in the recruitment of scaffolding proteins which bind the newly
phosphorylated tyrosine residues located on the intracellular domain of the receptor.
Activation of the ERK signaling pathway leads to recruitment of signaling molecules that
possess Src homology 2 (SH2), phosphotyrosine (PTB), and PDZ domains that recognize
13 and associate with phospho-tyrosine residues. These binding domains enable the formation of signaling complexes through the association of various adaptor proteins, namely Shc and Grb2, which bind additional scaffolding proteins (FRS2, Crk, CrkL) near the plasma membrane to ultimately activate guanine nucleotide exchange factors (SOS,
C3G). Subsequently, members of the Ras family of small G-proteins such as Ras and
Rap1 are activated to facilitate intracellular signaling. The stimulation of these small G- proteins initiates a downstream chain of phosphorylation events whereby MAP Kinase kinase kinases (MAP3Ks: c-Raf, B-Raf, A-Raf) phosphorylate and activate the dual specificity MAP Kinase Kinases (MAP2K: MEK1/2) which in turn finally activate the
MAP Kinases. The prototypical signaling elements of the mitogen-activated protein kinase superfamily are ERK1 and ERK2, proline-directed serine/threonine kinases which become activated by phosphorylation on specific threonine and tyrosine residues
(Threonine 183, Tyrosine 185) located in the activation loop of the enzymes (Pearson et al., 2001; Roux and Blenis, 2004; Rubinfeld and Seger, 2005). The amplification of the signaling cascade is demonstrated by the ability of activation of just 5% of cellular Ras molecules to induce full activation of ERK1/2 (Hallberg et al., 1994).
The three tier structure of the MAP Kinase cascade allows for both diversity in signaling and specificity. Depending on the stimulus, the engaged receptor tyrosine kinase (RTK) recruits an appropriate set of scaffolding proteins to initiate signaling through specific
MAPK3Ks and MAP2Ks, generating a ligand-specific downstream effect and biological response (Pouyssegur et al., 2002).
14 Due to the variety of stimuli which induce MAPK activity, cell type and signaling kinetics generate different cellular responses. For example, rat pheochromocytoma cells,
PC12 cells, have the capacity to differentiate into mature adrenal chromaffin cells by dexamethosone (a MAPK independent event) or depending on the level of MAPK stimulation, differentiate into sympathetic-like neurons or continue to proliferate. When stimulated with either EGF or insulin, PC12 cells rapidly proliferate due to a transient increase in ERK1/2 activity. Alternatively, when stimulated with NGF, FGF or IL-6, sustained ERK1/2 activation leads to differentiation of these cells into sympathetic-like neurons as evidenced by neurite extension and electrical excitability (Marshall 1995).
Additionally, a single stimulus can initiate different biological responses based on the required duration of MAPK activation. Again in PC12 cells, Nerve Growth Factor recruits either a Crk-C3G-Rap1 complex to RTKs which signal through B-Raf for prolonged MAPK activation and differentiation, or RTKs can assemble with a Shc-Grb2-
SOS-Ras complex to signal through c-Raf for transient MAPK activation and proliferation (Marshall, 1995; York et al., 1998; Kao et al., 2001). Other cell types such as T lymphocytes and megakaryocytes, are similarly receptive to transient or prolonged
MAPK activity for generation of distinct biological responses (Katz et al., 2007).
Importantly, the response is cell type specific as NGF-driven prolonged MAPK activity induces differentiation in PC12 cells, but leads to proliferation in olfactory sensory neuron precursors (Simpson et al., 2007). A similar proliferative response to sustained
ERK1/2 activity occurs in fibroblasts as well (Katz et al., 2007). These studies suggested that cell type, regulation of duration, and intensity of ERK1/2 activity is important in differentiation or proliferative responses. Moreover, the signaling complex which leads
15 to the activation of ERK1/2 often dictates biological outcomes and serves as a modulator
of MAPK activity in time and space (Pouyssegur et al., 2002; Sweatt et al., 2003).
Given the differential response that can occur based on MAPK activity it is not surprising
that ERK1/2 plays a role in cell fate decisions. The induction of ERK1/2 activity in embryonic stem (ES) cells dictates the alternative fates of self renewal or lineage commitment, during embryogenesis. Autocrine stimulation of ES cells by LIF leads to proliferation and ES cell renewal, whereas FGF4 stimulation results in cell commitment, prompting naïve ES cells to exit self-renewal and differentiate into neural and mesodermal cell fates (Kunath et al., 2007). Interestingly, the complete lack of ERK1/2
activity or removal of ERK2 expression does not affect ES cell proliferation (Meloche et al., 2004).
It is hypothesized that ERK1/2 similarly affects cellular function and cell fate decisions within the developing telencephalon. Rossant and colleagues (2003) developed two
specific antibodies which recognize the di-phosphorylated form of ERK1/2. These tools
were used to map the spatial and temporal domains of FGF activity in the developing
mouse embryo through visualization of phosphorylated ERK1/2 (Corson et al., 2003).
Importantly, they found that among the most reproducible and prominent domains of
sustained ERK1/2 activity from E8.5 to E10.5 were the frontonasal passages, the
forebrain, and the midbrain-hindbrain boundary. This places ERK1/2 at the correct time
and location to influence cortical development.
16 Structural Deformities of the Cerebral Cortex and MR:
A Historical Perspective
Mental retardation was first linked to changes in brain structures as early as the 1600s
when Thomas Willis initiated the field of clinical neurology by dissecting human brains
and mapping their cellular connections. He recognized that patients with congenital
mental retardation have abnormalities in their brain size and/or shape (Molnar, 2004).
However, it was not until the mid 1900s that clinically-defined changes in brain structure
became associated with mental retardation. Patients with mental retardation were initially classified on the basis of known causes, such as chromosomal abnormalities, infection, and identified syndromes (i.e. phenylketonuria, hydrocephalus or tuberous sclerosis), or undetermined causes. Between 15% and 50% of patients identified with mental disabilities fell into one of the classified causes. Remarkably, of the unclassified cases of MR, 90% were attributable to malformations of the brain or gliotic encephalopathies (Crome, 1972). The malformations documented to be associated with mental retardation included microcephaly which is literally “small brain.” It was thought that “the smaller the brain, the less intelligent the individual and the shorter expectation his life” (Crome, 1972). Another malformation associated with the microcephaly was unclassified hydrocephaly, whereby the lateral ventricles are enlarged and the overlying tissue is smaller, producing an overall larger head. Prosencephaly, or the union of the two telencephalic hemispheres due to the lack of formation of two separate ventricles, was also associated with mental retardation. It was further recognized at this time that neuronal migration defects give rise to mental retardation. The cortical deformities
17 specifically associated with migration were microgyria (smaller brain convolutions or
gyri), pachygyria (diminuation of gyri and sulci) and nodular ectopias (extra-cellular spots within the brain). This group of malformations is now commonly grouped as various forms of lissencephaly, or smooth brain disorders.
In the 1980s, Magnetic Resonance Imaging (MRI) became available and revolutionized diagnosis of MR. Scientists and physicians found that even subtle defects in cortical structures are associated with decreased mental capacity and developmental delays. The question then became one of mechanism and function. How do structural changes of the cerebral cortex arise? And, how do they translate into reduced mental capacity? Thus, the generation of the cortex became the focus of attention.
Cortical development is separated into four major stages: (1) expansion, the exponential
proliferation of founder progenitor cells by symmetric division necessary to generate
sufficient cells to populate the cortex; (2) neurogenesis, the generation of the correct
number of neurons and glia to produce and fill the cortex; (3) neuronal migration, the
traveling of each neuron to its proper location within the cortex; and (4) neuronal
differentiation, the maturation of neurons and establishment of connections to their
correct targets. It is now recognized that reductions in numbers of neurons populating the
cortex (from aberrant neurogenesis or neuronal death) gives rise to cortical hypoplasias
which include microcephaly, and cortical ectopias. Migration defects can also result in
cortical ectopias and lissencephaly. And finally, cortical dysplasias (abnormalities in
number or structure of dendrites, i.e. maturation) are due to abnormal neuronal migration
18 or neuronal maturation (Berger-Sweeney and Hohmann, 1997). The observations and
linkages made early in clinical neuroscience and later with more advanced imaging capabilities have established the connection between malformations of the cerebral cortex and congenital mental retardation. Proper cortical development is crucial for normal
cognitive function, and mutations in the genes fundamental for cortical development
result in mental retardation.
Controlling Cortical Development: The cell cycle
The process by which all eukaryotic cells divide is termed the cell cycle. It is comprised
of 4 major phases, namely mitosis (M phase), DNA synthesis (S phase), gap 1 (G1 phase)
and gap 2 (G2 phase). When cells are not in the process of division they are said to be in
the G0 phase. G0 is a quiescent stage whereby cells cease division or have terminally
differentiated. The G1 phase of the cell cycle is critical for a cell as it is the time in
which a cell surveys its environment and decides whether to commit to another round of
division or undergo differentiation, exiting into G0. As such, a restriction checkpoint
occurs at the end of G1. If a cell passes the checkpoint, it will proceed to the S phase and
prepare to complete another round of division. A second checkpoint is located at G2. If
a cell has undergone faulty replication of its DNA, the cell will not complete division and
will undergo apoptosis. Mitosis occurs through a precise and defined sequence of events,
in which structures within the nucleus of a cell separate both the nucleus and cytoplasm
into two compartments providing a means for division by cytokinesis.
19 Progression between the stages of the cell cycle is controlled by regulatory mechanisms which integrates extracellular signals that control proliferation with the intricate orchestration of events within the cell cycle. The induction of a cell cycle is tightly
regulated through the expression of cyclins which act to initiate the cell cycle and sustain
Figure 1-2: Progression through the cell cycle is controlled by inductive cyclin-dependent kinases and antiproliferative cyclin dependent kinase inhibitors.
its progression (Ohnuma and Harris, 2003). In order for a mammalian cell to transition from the cellular growth phase or first gap, G1, to DNA synthesis (S), cyclins D1-3, and cyclin E are required. The G2-M checkpoint is controlled by a separate group comprised of cyclin A, B1 and B2, in an intricate signaling cascade leading to mitosis and resulting
20 in proliferation (Ross, 1996). The function of the cyclins is regulated by the formation of a complex with specific cyclin-dependent kinase (CDK) molecules. This interaction controls the transition between the phases of the cell cycle. Cell cycle kinetics can further be modulated by tumor suppressors, p53 and pRb as well as the Cyclin-dependent
Kinase Inhibitors (CKIs), p15, p16, p21 and p27 (Dehay and Kennedy, 2007). These antiproliferative genes suppress progress through the cell cycle by sequestering Cyclin
D/E from their Cdk binding partners, and prevent phosphorylation of Rb, thus inhibiting
E2F-dependent transcription of additional cyclins required for G1 to S transition.
In the mouse, the neurogenic phase of cortical development includes the neural progenitor cells which have been shown to undergo 11 cell cycles spanning 6 embryonic days (E11-E17, Figure 1-3). Each cell cycle lengthens in duration through regulation of
Figure 1-3: Development of the Murine Cerebral Cortex occurs over a 6 day interval during which successively longer cell cycles generate the majority of cells.
21 G1 phase, so that the first cycle lasts 8.1 hours and the final cycle lasts 18.4 hours
(Caviness et al., 1995; Takahashi et al., 1995). The increase in cell cycle length is attributed to increases in the length of G1 as S phase is continuously about 4 hours long and G2-M is consistently 2.5hrs. Post-mitotic neurons exit the cell cycle and enter G0, where they become quiescent and unable to further divide. The exquisite regulation of both the progenitor cell expansion and neuronal proliferation to populate the cerebral cortex ensures its proper generation and function.
ERK1/2 and the Cell Cycle
As the original nomenclature implies, ERK1 and 2 were identified as mitogen-activated protein kinases which play a substantial role in mitotic regulation. Before the proteins were cloned and identified as protein kinases, it was recognized that they were phosphorylated and activated upon the presentation of mitogenic stimuli and their long- term phosphorylation through the G0 and G1 phases in the cell cycle were mandatory for cell cycle entry (Pages G 1993). The extent of ERK1/2 activity required for S phase entry has been debated. It is generally accepted that sustained activity promotes progression through G1, although ERK1/2 activity early in G1 is dispensable for transition in fibroblasts (Villanueva et al., 2007). Treatment of mouse embryonic fibroblasts with the MEK inhibitor U0126 at varying timpoints post serum-stimulation demonstrated that ERK1/2 activity is critical from mid-G1 to the end of G1 (as marked by cyclin A expression). Treatment of FGF-stimulated NIH3T3 cells with U0126 similarly showed that sustained ERK1/2 activity is not required until 2-3 hours prior to S phase entry (Yamamoto et al., 2006; Villanueva et al., 2007). These studies demonstrate
22 that ERK1/2 activity should be sustained through late G1 to ensure cell cycle progression into S phase. Further analysis of cell cycle progression by in vitro siRNA studies have demonstrated that ERK1 is necessary for completion of the G2 phase of the cell cycle, while ERK2 is reported to be required for G1 progression in HeLa cells (Liu et al., 2004), suggesting different roles for the ERK isoforms in cellular proliferation.
There are many known targets of ERK1/2 activity which are important for cell proliferation (Figure 1-2, 1-4). The primary mechanism by which ERK1/2 regulates cell cycle entry and progression is through induction of the critical G1 restriction checkpoint
Figure 1-4: ERK1/2 regulates cell cycle entry and progression by phosphorylation and transcription of numerous targets.
23 cyclin, Cyclin D1 (Roovers and Assoian, 2000; Wilkinson and Millar, 2000). In response
to serum or growth factor stimulation, ERK1/2 activation leads to phosphorylation of Ets
transcription factors such as Elk-1 and SRF which form the AP-1 transcriptional complex
with c-myc. This ternary complex binds the Serum Response Element on the cyclin D1
promoter to facilitate transcription of the cyclin. ERK1/2 can also regulate cyclin D1 by
phosphorylation of the transcription factor, c-myc. ERK-dependent expression of cyclin
D1 and its complex formation with cdk4/6 is therefore capable of stimulating cells to exit from G0 and promote entry into the cell cycle. Although cyclin D1 is an essential component of cell cycle progression and the primary target of ERK activity in response to mitogens, as cells prepare for replication and division, ERK activity also contributes to the production of multiple constituents of cellular growth. ERK1/2 facilitates the generation of new DNA and rRNA through the phosphorylation of carbamoyl phosphate synthetase, an enzyme which catalyzes the synthesis of new pyrimidine nucleotides
(Graves et al., 2000). ERK1/2 aids in transcription of ribosomal RNA genes through its phosphorylation of HMG box DNA binding domains on the transcription factor UBF
(Stefanovsky et al., 2006a; Stefanovsky et al., 2006b). ERK1/2 also directs protein translation by the phosphorylation of serine 209 on the translation initiation factor eIF4E through its immediate downstream target, the protein kinase, MNK1 (Chambard et al.,
2007).
ERK1/2 activity is critical for both cell cycle entry and progression through the cell cycle. Once in G1, cells will not enter the S phase if ERK activity is abrogated. This action is mediated by the suppression of antiproliferative genes (Yamamoto et al., 2006).
24 ERK1/2 can therefore regulate cell cycle entry and progression through phosphorylation
of both pRb and p27. ERK1/2 acts to regulate p27KIP1 levels at multiple levels. ERK1/2
directly phosphorylate p27KIP1, inhibiting its function, and stimulating its degradation
through induction of the ubiquitin E3 ligase, SKP-2 (Aktas et al., 1997; Kawada et al.,
1997; Takuwa and Takuwa, 1997; Villanueva et al., 2007). ERK1/2 have also been
reported to regulate p27KIP1 translation (Miskimins et al., 2001) or mRNA stability
(Sakakibara et al., 2005). ERK1/2 activation is correlated with reduced cellular p27KIP1
levels (Teixeira et al., 2000; Sakakibara et al., 2005) and enhanced cellular proliferation.
Conversely, inhibition of ERK activity results in elevation of p27KIP1 levels and cell cycle arrest (Gysin et al., 2005). Another potential level of ERK regulation on cell cycle entry and progression is through phosphorylation of the antiproliferative gene, tis21. It is of particular importance due to its expression in cells which are undergoing terminal mitosis
(Iacopetti et al., 1999) and downregulation in post-mitotic neurons. Overexpression of tis21 inhibits G1-S progression through transcriptional regulation of cyclin D1 in the presence of Rb, while in the absence of Rb, tis21 downregulates cyclinE-cdk4. Of particular relevance is the fact that ERK1/2 phosphorylates tis21 (Hong et al., 2005), providing a potential mechanism by which ERK affects terminal mitosis of neural progenitor cells.
The ERKs were first identified as microtubule associated protein (MAP), kinases owing to their phosphorylation of MAP itself. ERK1/2 has also been implicated in regulation of
the cell cycle through phosphorylation of and association with cytoskeletal proteins such
as neurofilament and paxillin (Chambard et al., 2007). ERK activity has been shown to
25 be important in regulating mitotic progression at the G1/S boundary (Guadagno and
Ferrell, 1998; Roberts et al., 2002; Horne and Guadagno, 2003), as well as in microtubule
reorganization during M phase (Gotoh et al., 1991). ERK has also been localized to spindle poles during pro-metaphase, the mid-body during cytokinesis and is generally
thought to contribute to normal mitotic progression in somatic cells (Willard and Crouch,
2001).
Cortical Development I: The Expansion of Progenitor
Cells
Cortical development begins upon closure of the neural tube. Initially, the single layer of
multipotent neural progenitor cells which define the proliferative neuroepithelium
(neuroepithelial cells) must undergo an exponential series of symmetrical divisions. This
expansion in the number of neuroepithelial cells in turn increases the number of
committed neural precursors that will comprise the telencephalic ventricular zone. The
neuroepithelial or founder cell expansion is a critical component of cortical development
as insufficient proliferation results inadequate numbers of committed neural progenitors
and the generation of a microcephalic brain (Rakic, 1988). Furthermore, the expansion of
neuroepithelial cells is responsible for the evolutionary enlargement of brains between
species (Caviness et al., 1995; Rakic, 1995).
Neuroepithelial cells are highly polarized in their apical-basal axis and are identifiable
due to their epithelial features including expression of nestin and the presence of tight
26 junctions. The symmetrical division of the neuroepithelial cells is ensured by the
orientation of the cleavage plane in cytokinesis (Figure 1-5, division A). A vertical cleavage plane allows for the equal distribution of apical and basal cellular matter and the production of two equal neuroepithelial daughter cells (Gotz and Huttner, 2005). After expansion of the progenitor pool is complete, the neuroepithelial cells transition into a committed neural progenitor cell, downregulating some of their epithelial features and adopting astroglial properties such as glycogen granules (Figure 1-5, division B). This cell has also been termed a radial glial cell. Committed neural progenitor cells are marked by the expression of numerous proteins and their maturation into restricted- precursors and can be identified by the inclusion or exclusion of a subset of these markers. Most of the markers for radial glial cells are intermediate filament proteins including nestin, (which is maintained from the neuroepithelial cell), vimentin, and the glial fibrillary acidic protein (GFAP). Additional markers of radial glia include expression of the transcription factor Pax6, the astrocyte-specific glutamate transporter,
GLAST, and the brain-lipid binding protein, BLBP (Gotz and Huttner, 2005). A population of secondary or subventricular progenitors is further derived from neuroepithelial cells and the radial glia (Fishell and Kriegstein, 2005). These cells are successively committed intermediate progenitor cells identifiable by downregulation of
Pax6, and expression of the transcription factor Tbr2 (Englund et al., 2005; Gal et al.,
2006); (Figure 1-5, division C). The expansion of the founder neuroepithelial cell population through self-renewal and generation of intermediate progenitor cells is essential for the generation of the millions to billions of neurons which populate the mature cortex (Rakic, 1995; Martinez-Cerdeno et al., 2006). It is of significance that
27 IPCs are the only symmetrically neurogenic progenitor cells. Furthermore, they
contribute neurons to all cortical lamina and are postulated to underlie the amplification
of the cortex throughout development and evolution (Pontious et al., 2008).
Cortical Development II: Neurogenesis
Upon completion of the expansion phase and concomitant with the appearance of radial
glia, successive groups of neural progenitors re-enter the cell cycle and subsequently
initiate neurogenesis. The first glimpses of the cellular dynamics and the level of
complexity involved in cortical neurogenesis came in the late 19th century when neuroanatomists such as Wilhelm His recognized that two populations of cells within the
embryonic cerebral cortex underwent mitosis. One was located at the ventricular surface
and the other away from the ventricular lumen. Each of these cell types divided,
producing the non-mitotic cells which resided in the cortical plate. Rudolf Albert von
Kolliker further identified the progenitor cells which lined the ventricles in the pseduostratified epithelium and ultimately changed their shape. These were the surface dividing cells that His described as germinal cells and are now known as the radial glia.
The non-surface dividing or abventricular mitotic cells, which His also documented, are now thought to be the intermediate progenitor cells that reside in the sub-ventricular zone. Notably, it has been recognized that the excitatory projection neurons which populate the cerebral cortex are derived from and can migrate along His’ germinal cells
(radial glia). Extensive lineage mapping has shown that nearly all divisions producing
the excitatory projection neurons are derived from a radial glia. Studies by Messing and
colleagues revealed the multipotential properties of these cells following generation and
28 analysis of transgenic mice expressing the cre-recombinase by the human GFAP promoter (hGFAP-cre) (Zhuo et al., 2001). Subsequently, the hGFAPcre/+ transgenic mice have been used for a detailed fate mapping of the developing telencephalon (Malatesta et al., 2003; Anthony et al., 2004). Malatesta et al. have shown that over 90% of neurons in the neocortex and hippocampus are derived from hGFAP-cre expressing progenitors, including all long range projection neurons, but few interneurons (Malatesta et al., 2003).
Modes of neuronal generation
After the initial expansion phase of cortical development, whereby radial glia are produced, these cells then begin to produce neurons. Multiple modes of division underlie the generation of cortical projection neurons (see Figure 1-5): (1) Radial glia which are located in the ventricular zone undergo asymmetric divisions producing one neuron and maintaining one progenitor cell (65.8%). (2) Radial glial cells divide to produce one self-renewed radial glia and a cell which becomes an intermediate progenitor cell and goes on to divide away from the ventricular zone (7.3%). (3) Radial glia symmetrically divide into two radial glia,
29 Figure 1-5: Modes of Cellular Generation in the Developing Cerebral Cortex.
one which divides again and one which will translocate to the cortical plate and differentiate (26.9%). (4) Radial glia symmetrically divide to produce two more radial
glia which will both undergo asymmetric divisions (9.6%) (5) Abventricular
intermediate progenitors symmetrically divide to produce two neurons (89.5% of
abventricular divisions). (6) Abventricular intermediate progenitors divide, producing
two cells which will each divide again (10.5% of abeventricular divisions) (Haydar et al.,
2003; Noctor et al., 2004; Gotz and Huttner, 2005).
Cortical malformations and disrupted division
Disruption in either phase of proliferation, expansion of the progenitor pool or
neurogenesis, leads to significant structural changes in the cerebral cortex and are
30 associated with cognitive deficits. For example, Autosomal Recessive Primary
Microcephaly, MCPH, is characterized by reduced growth of the cerebral cortex, albeit with normal lamination, a small head circumference, and mental retardation. Four genetic loci have been mapped and each gene is important in the centrosomal regulation of neural progenitor cellular mitosis (Bond and Woods, 2006). The genes,
Microcephalin, ASPM (the abnormal spindle-like microcephaly associated protein),
Cdk5Rap2, and CenpJ are highly expressed and localized to the neuroepithelium lining the lateral ventricles of the developing cortex. Mutations in each of these genes individually results in a reduction in numbers of neurons stemming from either a reduction in neural progenitor cells themselves or an inability of the progenitor to divide appropriately. Nde1 (a Lis1-interacting protein) mutants are also microcephalic due to a loss of microtubule organization and spindle pole orientation which results in aberrent neuroepithelial cell mitosis (Feng and Walsh, 2004). One of the most recognizable forms of mental retardation, Down’s syndrome, is also characterized by small brain size stemming from reduced numbers of neural progenitors due to an increase in cell cycle length (Haydar et al., 2000). The conditional knockout of the cell cycle regulator Rb in the developing telencephalon also causes a robust (7 fold) increase in progenitor proliferation resulting in a dramatic increase in telencephalon size (Ferguson et al., 2002).
This and previous studies therefore demonstrate that the Rb protein is critical for commited neural progenitors to undergo terminal neurogenic mitotsis and survive (Slack et al., 1998). The CKI, p27KIP1 is of particular interest and importance due to its role in cortical development (Delalle et al., 1999; Mitsuhashi et al., 2001; Goto et al., 2004;
Lukaszewicz et al., 2005; Tarui et al., 2005). Animals in which p27KIP1 has been
31 knocked out exhibit increased cortical thickness resulting from a 27% increase in neurons
in upper cortical lamina (Caviness et al., 2003; Goto et al., 2004). Conversely,
overexpression of p27KIP1 in the developing telencephalon slows progenitor proliferation with fewer neurons populating the upper cortical layers, resulting in reduced cortical thickness (Tarui et al., 2005).
Cortical Development III: Migration
Newly generated neurons and self-renewed progenitor cells each have a separate fate.
This is most obvious in the third stage of neurogenesis, neuronal migration.
Interkinetic migration during neurogenesis:
Within the ventricular zone, radial glial cells begin the cell cycle located immediately
adjacent to the ventricle. During S phase, the cells migrate away from the ventricle and subsequently return to their original position along the ventricle to undergo mitosis
(Takahashi et al., 1993; Noctor et al., 2004). After division, committed neural progenitor
cells or radial glia can maintain their position within the apical ventricular zone and
divide again, or they can migrate away from the ventricle where they can mature into an intermediate or basal progenitor cell. In contrast to radial glia, intermediate progenitor
cells do not undergo interkinetic nuclear movement. Upon their generation from a radial
glial division, they migrate away from the ventricle and undergo all phases of the cell
cycle in an abventricular position. Despite the progenitor from which a neuron is born,
both the ventricular neurons and abventricular-derived neurons migrate toward the pial
surface of the brain upon their birth. The laminar organization of the cerebral cortex is
32 derived from the migration of these newly born neurons in a stereotypical “inside-out”
pattern. The neurons utilize radial glial fibers or self-generated radial fibers as scaffolds
to migrate toward their final locations. The earliest generated neurons split the preplate and subplate and remain closest to the ventricle, forming layer VI. Later born cells migrate outward past the earlier migrants and are positioned at successively more superficial positions (demonstrated in Figure 1-3).
Within each discrete wave of division, newly born neurons carry intrinsic signals which dictate their location in the cortical plate. If neurons are harvested from a mouse embryo at E14 and transplanted into a distinct ventricular zone, they can migrate to their correct location. However, after gliogenesis has begun, the newly born neurons can maintain their transcription factor identity but they do not migrate to the proper position
(McConnell and Kaznowski, 1991; Desai and McConnell, 2000). Takahashi et al. found that a cell cycle counting mechanism is intrinsic to the laminar fate of the neuron
(Takahashi et al., 1999).
The generation of neurons and glia which fill the cortical plate therefore arises from a highly regulated process of symmetrical division, asymmetrical division and migration.
The radial glia which do not divide either differentiate into neurons early in neurogenesis or eventually translocate into the cortical plate where they encounter extrinsic cues to direct their differentiation into astrocytes or oligodendrocyte precursors (Qian et al.,
2000; Sauvageot and Stiles, 2002). GABAergic interneurons, which also reside within the cortex, are not descended from GFAP-expressing radial glia. They are derived
33 principally from the ganglionic eminence and migrate tangentially into the cortical plate
(Malatesta et al., 2003; Anthony et al., 2004).
Cortical malformations and disrupted migration
Aberrations in neuronal migration have been shown to be responsible for lissencephaly and developmental disorders which are characterized by cognitive impairment (Rakic,
2000). When neurons are not properly positioned after migration, heterotopias of neurons are formed and numerous clinical syndromes can result. In humans, these disorders are associated with the cortical malformations of microgyria, pachygria, and nodular ectopias, all of which are more generally classified as lissencephaly. Mutations in Reelin, Lis1 and DCX lead to lissencephaly and mental retardation by affecting the process of neuronal migration or the substrates on which neurons migrate (Vallee and
Tsai 2006). Lis1 and doublecortin encode proteins which associate with microtublules and cytoskeletal elements like the microtubule motor protein dynein. These proteins ensure that neuronal migration is possible and that neurons migrate properly to form the laminar organization of the cortex. Reelin is an extracellular matrix glycoprotein which is important in defining the pial surface and generating a stop signal for neurons migrating out of the germinal zones (Haydar, 2005). When neuronal migration is faulty, heterotopias often form, leading to improper synaptic connectivity. This often results in cortical malformations associated with epilepsy (Rakic, 2000; Guerrini and Marini,
2006).
34 Cortical Development IV: Differentiation of Neurons
and Synaptogenesis.
Upon reaching its final destination, a newly-born cortical neuron differentiates into a
specific cell-type by expressing neurotransmitters and corresponding receptors,
elaborating dendrites and dendritic spines, and sending axons to their synaptic targets.
These processes are important and ensure that communication between cells occurs properly and efficiently. Mutations in FMRP, the fragile X mental retardation protein, is responsible for abnormal dendritic spines found in patients with Fragile X MR due to disrupted translation of mRNA at synapses. Disruption of dendritic arborization and spine density is also associated with Rett syndrome and Down syndrome. The mechanisms underlying the malformations seen in these diseases is unknown, however, synapse formation and elaboration of dendrites is critically linked to neuronal activity and therefore both neuronal proliferation and migration (Haydar, 2005). This phase of development will not be covered in depth.
In summary, cortical development can be divided into four very broad stages, founder cell expansion, neurogenesis, neuronal migration and differentiation. When neurogenesis is disrupted, the result is cortical hypoplasia or microcephaly; when migration is disrupted, cortical ectopias or lissencephaly results, and when differentiation is disrupted, cortical dysplasia or a loss of dendritic arborization results. Abrogation of any stage of cortical development through cell cycle machinery or directive proteins leads to alterations in cortical structures associated with cognitive impairment, providing
35 convincing evidence that the two are coupled (Berger-Sweeney and Hohmann, 1997).
Accordingly, the genetic mutations linked to structural defects and MR are involved in
regulation of mitotic structural proteins or the intrinsic and extrinsic cues required for
proper progenitor expansion and division. It is clear that normal division of these cells
through the cell cycle is critical to cortical development.
The Molecular Mechanisms of Cortical Development
Following the neurogenic phase of cortical development, a gliogenic phase arises and peaks at birth to produce first astroglia, followed by oligodendroglia (Qian et al., 2000).
This transition from expansion of progenitor pools to neurogenesis and then gliogenesis
is regulated by extrinsic and instrinsic factors (Cameron et al., 1998; Ross et al., 2003).
Basic Fibroblast Growth Factor (bFGF or FGF2) and Epidermal Growth Factor (EGF)
are responsible for directing neural progenitor cell expansion in concert with the basic
Helix Loop Helix (bHLH) transcription factors (Vaccarino et al., 1999). The principal
bHLH factors that act to regulate early stages of cortical development are members of the
Hes (Nakamura et al., 2000; Ohtsuka et al., 2001) and Id (Lyden et al., 1999) families.
These transcription factors act by promoting the progenitor proliferation through
suppression of neuronal differentiation. They functionally antagonize the actions of the
proneural bHLH factors, Ngn1/2 and Mash1 (Nieto et al., 2001). Neurogenesis is
induced when the Hes/Id factors are dowregulated and the proneural factors dominate.
Inactivation of the proneural bHLH factors Ngn2 and Mash1 results in a smaller cortex
with fewer neurons and many more astrocytes (Nieto et al., 2001). Inactivation of either
the Hes1 or Hes5 gene leads to premature differentiation of neurons due to the absence of
36 this intrinsic brake on neural differentiation. This results in a two-fold increase in the
number of neurons in the brain during early development (Ishibashi et al., 1995;
Nakamura et al., 2000; Ishibashi, 2004). Double knockout mice in which both Hes1 and
Hes5 are inactivated results in an even more severe phenotype, in which radial glia
differentiate prematurely and lose their structural integrity to organize the neural tube.
This phenotype results from the loss in compensation by the other gene. To further
assess compensation by Hes3, triple knockout mice were generated and demonstrate that
these Hes genes are required for maintenance of radial glia as their ablation leads to
complete premature neuronal generation and loss of all progenitors (Hatakeyama et al.,
2004). Conversely, overexpression of Hes1 increases the number of mitotically active progenitors in the developing telencephalon (Ishibashi et al., 1994) stemming from the delayed arrest of proliferation, and causing hypercellularity of the cortex.
The Id family of bHLH factors consists of 4 members and is functionally similar to the
Hes family in their ability to promote progenitor proliferation and suppress the action of
proneural factors (Lyden et al., 1999). Moreover, Id sustains Hes1 expression to allow
for expansion of the progenitor pool and suppress precocious neuronal differentiation
(Bai et al., 2007). Specifically, Id2 overexpression in chick neural tube enhances Hes1
expression and inhibits expression of the proneural genes and therefore neuronal
differentiation. Moreover, Id1/3 double knockout mice or chicks electroporated for Id1-3
RNAi display precocious neurogenesis whereby Hes1 expression is suppressed and
proneural gene expression expands (Bai et al., 2007). In addition to regulation of bHLH
transcription factors, Id proteins stimulate cell cycle progression by inhibiting the action
37 of the retinoblastoma protein (Toma et al., 2000). Rb-deficient neural progenitor cells exhibit significant delays in cell cycle exit compared with wildtype cells, suggesting that
Rb is essential for cell cycle control of E2F activity required for cell cycle progression
(Callaghan et al., 1999). Mutation of the Rb protein leads to hypercellularity of the cortex due to increased proliferation of the neural progenitor cells (Ferguson et al., 2002).
The expression of Id1/4 is high in neural precursors and expression is extinguished upon exit from the cell cycle and differentiation (Iavarone and Lasorella, 2004). Interestingly,
Id1-4 are targets of MeCP2, the mutated gene in Rett Syndrome which is characterized by MR. In both a Mecp2 deficient mouse model, and human brain tissue from Rett
Syndrome patients, all four Id proteins were found to be elevated (Peddada et al., 2006).
The transition into neurogenesis is critically dependent upon expression of the proneural genes Mash1 and Ngn1/2. When the activity of these transcription factors is inhibited, a loss of neurons and neural progenitors results, accompanied by the precocious generation of glial-restricted progenitors and astrocytes (Guillemot, 2007). Interestingly, a novel role for p27 has recently identified its ability to promote neuronal differentiation through stabilization of Neurogenin 2 (Ngn2), and migration of post-mitotic projection neurons through inhibition of RhoA. Although both of these activities are independent of p27’s role in cell cycle regulation, they demonstrate a previously unappreciated role for p27 in neuronal differentiation at the level of bHLH transcription factors (Nguyen L 2006).
The transition from neurogenesis to gliogenesis is largely controlled by JAK-STAT signaling. Gliogenesis is suppressed during early neurogenic periods due to epigenetic factors such as DNA methylation, but more importantly, by proneural gene-mediated inhibition. Ngn1/2 inhibits STAT phosphorylation by sequestering CBP/p300 complexes
38 away from STAT activation sites. Once Hes/Id bHLH transcription factors dominate over the proneural transcription factors, STAT1/3 is phosphorylated and activated to promote astrogliogenesis through transcription of gfap (He et al., 2005).
MAPK and Cell Fate Determination
ERK1/2 is critical for development of the neocortex and specifically in cell fate
determination of neural progenitor cells. A series of elegant experiments by Miller and
colleagues have delineated the pathway by which endogenous growth factors such as
bFGF, NT-3 and BDNF activate a MEK-ERK-RSK-C/EBP signal transduction cascade
within neural progenitor cells to induce neuronal differentiation. Specifically, MEK
activation leads to activation of both ERK1/2 and RSK, which ultimately phosphorylates
the C/AAT Enhancer Binding Protein (C/EBP) to induce transcription of the neuron-
specific Tα1 tubulin isoform. The expression of Tα1 tubulin represents the initial phase
of cellular differentiation and specifies neuronal fate. MEK regulation of C/EBP activity through ERK1/2 and RSK phsophorylation promotes neurogenesis and inihbits
gliogenesis without affecting progenitor proliferation or survival (Menard et al., 2002;
Barnabe-Heider and Miller, 2003; Paquin et al., 2005; Bartkowska et al., 2007; Calella et
al., 2007). In neural progenitor cells, ablation or mutation of a single member of the
scaffolding complex which links receptor tyrosine kinases to the activation of the MAP
Kinase cascade, such as SHP2 (Gauthier et al., 2007; Ke et al., 2007), FRS2 (Yamamoto
et al., 2005) or ShcA (McFarland et al., 2006) leads to reduced ERK1/2 phosphorylation and reduced neuronal differentiation. These data will be discussed in detail below.
39 Greenberg and colleagues (Bonni et al., 1997) have demonstrated that CNTF promotes
differentiation of cortical precursors into astrocytes through JAK-STAT and transient
ERK1/2 signaling; ERK1/2 inhibition was shown to augment CNTF-mediated GFAP
expression. C/EBP inhibition also leads to enhanced CNTF-mediated GFAP
Figure 1-6: Neurogenic versus gliogenic cell fate determination is regulated by ERK1/2 Activity.
transcription. These data demonstrate that ERK1/2 serves to mediate neurogenic cell fate determination and actively suppresses gliogenesis until the end of neurogenesis when proliferative and neurotrophic growth factors are downregulated. The neurogenic/
gliogenic switch identified by ERK1/2 activity is illustrated above in Figure 1-6.
40 ERK1/2 has also been shown to regulate bHLH transcription factors. Inhibition of
Ras/MAPK activity with MEK inhibitors reduces TGFβ-stimulated Hes1 transcription
(Stockhausen et al., 2005). Of particular importance is that Id 1/3 expression is positively regulated by phosphorylation (Deed et al., 1997; Hara et al., 1997) and specifically by
ERK activity (Bain et al., 2001; Ruzinova and Benezra, 2003). As Hes1 and the Id transcription factors suppress neurogenic bHLH transcription factors during the proliferative phases, these data support the idea that ERK1/2 may also play a role in ensuring sufficient proliferation of founder progenitor cells to enable the generation of a fully populated cortex.
Together, these data define a role for ERK1/2 in cell fate determination within the developing cerebral cortex whereby ERK1/2 activation promotes neurogenesis and suppresses gliogenesis to ensure the proper timing and generation of cells in the neocortex.
Disruptions in upstream elements of the MAPK signaling cascade lead to anatomical and behavioral impairments in mice
The effect of targeted mutations or deletions in the components of the MAPK cascade on cell fate determination leads to significant repercussions in formation and function of the cerebral cortex. Disruption or mutation of elements at each level of the MAPK cascade, from receptors at the plasma membrane to guanine nucleotide exchange factors, results in cortical malformations. Mice lacking the extrinsic ligand, FGF2, display reductions in neuronal density and cortical thickness due to a severe reduction in the number of neural
41 progenitor cell divisions. The loss of progenitors limits the numbers of neurons that are
able to be generated (50% fewer in FGF2 nulls as compared to wildtypes) (Dono et al.,
1998; Ortega et al., 1998; Vaccarino et al., 1999; Qian et al., 2000; Raballo et al., 2000;
Korada et al., 2002). Likewise, mice overexpressing a mutated FGF2 receptor, kinase
domain deficient FGFR1 mice, exhibit similar phenotypes with drastically reduced numbers of neural progenitors (Shin et al., 2004). Interestingly, the converse experiment
whereby a constitutively activated FGFR3 mutation was expressed in cortical progenitor
cells revealed that the activated receptor induced the generation of enlarged brains due to increased progenitor cell proliferation (Inglis-Broadgate et al., 2005; Thomson et al.,
2007). This effect was due to ERK1/2 activity, as treatment with MEK inhibitiors reversed the proliferative effects. These data reveal that ERK1/2 are critical components of FGF2 signaling necessary for the expansion of the neural progenitor cell pool and subsequent neurogenesis.
Recently, Miller and colleagues have demonstrated that inhibition or downregulation of the TrkB/C receptors also abrogates the proliferation and neuronal differentiation of neural progenitor cells (Bartkowska et al., 2007). These data elucidated a role for the neurotrophins BDNF and NT-3 in neurogenesis through their activation of TrkB and
TrkC. Trk activation leads to signaling through the MEK-ERK-RSK-C/EBP cascade.
Downstream of the RTKs, mutation and inactivation of the MAP3Ks, C-Raf/B-Raf/A-
Raf results in neurological perturbations. Activation of each isoform has been implicated in the neuron-like differentiation of PC12 cells, and is coupled to signaling from RTKs through to MEK1/2 and ERK1/2 (Wixler et al., 1996). Therefore, it is of importance to note the effects of inactivation of each. B-Raf knockouts die during embryogenesis and
42 display specific loss in both sensory and motoneurons (Wiese et al., 2001). A-Raf
knockout mice survive birth; however, they display distinct motor deficits and die in
early perinatal life (Pritchard et al., 1996). C-Raf knockouts also die embyronically,
displaying gross growth retardation (Wojnowski et al., 1998; Mikula et al., 2001). B-Raf
and C-Raf can compensate for each other, however. When A-Raf is expressed from the
B-Raf locus (in a B-Raf null mouse) although lethality is rescued, neural defects remain.
These mice continued to display deficits in neural progenitor cell proliferation, neuronal migration and dendritic maturation, all of which contribute to reductions in cortical thickness and disorganization of cortical lamination. These data demonstrate that B-Raf is specifically responsible for regulation of neural progenitor cell differentiation associated with cortical layering (Camarero et al., 2006). In a separate attempt to circumvent the lethality of B-Raf knockout mice, behavioral assessment of a spatially restricted B-Raf mutant, where inactivation occurs in the forebrain (predominantly hippocampus), spatial learning and hippocampal-dependent memory deficits were identified by the Morris Water Maze and context discrimination tasks, but not the formation or extinction of cued or contextual fear conditioning (Chen et al., 2006). These data further show that B-Raf has specific roles in cortical function. The generation of additional B-raf and Raf-1 (C-Raf) conditonal knockouts, where these genes are lost after recomibination driven by a nestin-cre transgene, lead to thinner cortices because of hypothalamic dysfunction and embryonic death (Zhong et al., 2007).
Further insight into the mechanism behind the cortical phenotypes associated with reductions in Raf function come from the inactivation of the MAP2Ks, MEK1/2. The
43 expression of dominant negative MEK constructs in cortical progenitor cells either
through in utero electroporation or by transfection/infection techniques demonstrates that
MEK-ERK activity is required for neuronal differentiation but not neural progenitor cell
survival or proliferation. Progenitors remain in an undifferentiated state (as defined by
nestin immunoreactivity) until gliogenic stimuli induce their differentiation into
astrocytes (Menard et al., 2002; Paquin et al., 2005). These data demonstrate a direct role
for MEK-ERK activity in promoting neuronal fate. In behavioral assays, mice which
express dominant negative MEK1 under the Tα1 tubulin promoter (expression occurs in
post-mitotic neurons) have poor performance on contextual fear conditioning paradigms,
despite displaying normal performance in motor coordination and learning, and cued fear conditioning (Shalin et al., 2004). These data demonstrate that MEK1 activity is required for hippocampal-dependent learning and memory. Ablation of various components of the signaling cascade which couples extracellular stimuli to MAPK activity supports the importance of MAPK signal transduction in cortical development and function.
Inactivation of the adaptor protein ShcA in the developing forebrain by Nestin-cre
recombination leads to a normally organized cortex of significantly reduced size. This
microcephalic brain arises from apoptosis of neural progenitor cells during early
neurogenesis (E10.5-12.5) and the specific reduction in neurons of the superficial layers
of the cortex. Interestingly, the neural progenitor cells do not display proliferation
defects in the ventricular or subventricular zones, nor do they undergo premature
differentiation (McFarland et al., 2006). SHP2 is the Src-homology 2 domain containing
tyrosine phosphatase which also acts as an adaptor protein for ERK activation. Two
independent reports demonstrate that Shp2 ablation or loss of function elicits impaired
44 corticogenesis, whereby neural progenitor cells lose their capacity to proliferate and
generate neurons, therefore giving rise to an abnormally large number of astrocytes
(Gauthier et al., 2007; Ke et al., 2007). Both studies identify a loss of ERK1/2 activation
as a result of Shp2 inactivation. Mice which have mutation in the Shp2 binding sites on
the FRS2α adaptor protein (FRS2α2F/2F mice) have normal gross morphology, but exhibit
a 10% reduction in brain size compared to wildtype mice, or those with mutations in
Grb2 binding sites on FRS2. These mice also have specific deficits in cortical thickness and lamination of the cortical plate at E18.5, which was attributed to the presence of many fewer cells. The number of intermediate progenitor cells and mitotic cells found in the subventricular zone were both found to be significantly reduced in FRS2α2F/2F mice.
Importantly, the level of ERK1/2 activity was dramatically lower throughout neurogenesis in FRS2α2F/2F mice (Hadari et al., 2001; Yamamoto et al., 2005).
Mice hypomorphic for the guanine nucleotide exchange factor, C3G, die at E14.5 due to
defects in blood vessel maturation; however, even at this age they exhibit increased neural progenitor proliferation and decreased neuronal differentiation (Voss et al., 2003;
Voss et al., 2006). These results were due to a delay or failure of the progenitors to exit the cell cycle.
Aberrant ERK1/2 Signaling leads to Developmental Disorders
45 Significantly, many components of the MAPK superfamily are responsible for genetic
Figure1-7: Mutation of nearly every component of the MAPK signaling pathway leads to a congenital disorder characterized by mental retardation and/or cardio-facio-cutaneous abnormalities.
developmental disorders, each of which is characterized by a form of mental retardation
(See Figure 7). As reviewed by Weeber and Sweatt (2002), Rsk2 mutations leads to
Coffin-Lowry Syndrome (Zeniou 2002, Dufresne 2000) and Rsk4 mutations are associated with X-linked MR. Mutations in the Ras-GAP, NF1Gap, leads to
Neurofibromatisis1 MR, while CREB Binding Protein (CBP) mutations cause
Rubenstein-Taybi Syndrome, and MSK2 has been mapped within the Bardet Biedl locus.
46 Mutations in MNK also contribute to Fragile X Syndrome. Furthermore, it has recently been appreciated that mutations in genes at each level of the MAPK signaling cascade are associated with Neuro-Cardio-Facio-Cutaneous (NCFC) Syndromes (Duesbery N and
Woude 2006, Bentires-Alj 2006). These include neurofibromatosis 1, NF1
(neurofibromin), Noonan Syndrome (SHP2), LEOPARD Syndrome (SHP2 and K-Ras),
Costello Syndrome (H-Ras), and CFC Syndrome (B-Raf, MEK1/2, and K-Ras).
Importantly, each of these syndromes presents with varying degrees of mental retardation.
ERK in Learning and Memory
There is now a significant body of work demonstrating the critical role the ERKs play in synaptic plasticity, learning and memory (reviewed in Adams and Sweatt, 2002). Sweatt and colleagues performed a series of seminal experiments demonstrating the requirement for ERK activation in synaptic plasticity by showing that electrical stimulation of the hippocampus resulted in ERK activation, the development of hippocampal LTP and expression of long term memory (English and Sweatt, 1996, 1997; Atkins et al., 1998).
Subsequent studies have demonstrated that ERKs are required for other forms of plasticity including the development of LTP in the dentate gyrus (Coogan et al., 1999), amygdala (Huang et al., 2000; Schafe et al., 2001), insular cortex (Berman et al., 1998) and visual cortex (Di Cristo et al., 2001). ERK signaling has been shown to be essential for spatial learning (Blum et al., 1999; Hebert and Dash, 2002), fear conditioning, (Atkins et al., 1998; Schafe et al., 2001) and conditioned taste aversion (Berman et al., 1998;
47 Schaeffer et al., 1998; Sharma et al., 2004; Vomastek et al., 2004). The molecular basis
of these effects is the subject of intense interest and a number of mechanisms have been
investigated, including ERK-dependent regulation of AMPA receptor insertion into the postsynaptic membrane (Lu et al., 2001; Shi et al., 2001), physical remodeling (Wu et al.,
2001; Goldin and Segal, 2003; Arendt et al., 2004) and generation (Futter et al., 2005) of dendritic spines, Kv4.2 potassium channel function (Yuan et al., 2002; Morozov et al.,
2003) and the local regulation of protein synthesis (Kelleher et al., 2004). The ERKs also serve to regulate the expression and function of transcription factors such as CREB, c-fos,
Zif268, Arc and Homer that regulate synaptic plasticity (Rosenblum et al., 2002;
Vazdarjanova et al., 2002; Bozon et al., 2003; Igaz et al., 2004). Of particular relevance is the report that mice expressing a dominant negative MEK1, the immediate upstream regulator of the ERKs, in cortical neurons display deficits in contextual memory formation, spatial reference memory, contextual fear conditioning (Shalin et al., 2004), as well as the late, translation-dependent, phase of LTP induction (Kelleher et al., 2004). It is unclear whether structural abnormalities within the cortex, or the molecular mechanism by which ERK regulates synaptic plasticity in mature neurons underlies these conditions.
As it has been recognized that mutation in both upstream and downstream components of
ERK1/2 are involved in cognitive ability, and ERK1/2 is a critical component of the mechanism underlying memory formation and retrieval, it seems likely that ERK1/2 are also involved in higher order cognition. Intriguingly, the ERK2 locus is at chromosome
22q11.2 in the human. This locus is responsible for 22q11Deletion Syndrome which is a
1.5 or 3 Mb microdeletion resulting in a congenital disorder that has been variously
48 termed DiGeorge Syndrome, Velocardiofacial Syndromes, Conotruncal Anomaly Face
Syndrome (and others) all manifesting as a Cardio-Facio-Cutaneous syndrome. In addition to craniofacial patterning and cardiac defects, these patients present with varying levels of cognitive deficits ranging from mild to severe mental retardation, a dramatically increased risk of schizophrenia or major depressive disorder and they exhibit small but significant reduced cortical volumes (Swillen et al., 2000; Maynard et al., 2002;
Campbell et al., 2006). ERK2 maps to a locus just distal to the typical 3Mb deletion breakpoint critical to DiGeorge Syndrome (Saitta et al., 1999) but is similarly flanked by low copy repeats which facilitate the larger microdeletion (Edelmann et al., 1999). A small cohort of individuals with a 1 Mb micodeletion encompassing ERK2 and who have a classic DiGeorge Syndrome phenotype has been identified by 2 independent studies
(Saitta et al., 1999; Shaikh et al., 2007; Ben-Shachar et al., 2008). The identification of these individuals is of relevance to the idea that ERK1/2 signaling is critical for cortical development and the subsequent learning deficits and cognitive disabilities associated with these syndromes.
Individual roles for the ERK1 and ERK2 isoforms?
The MAPKs, ERK1 and ERK2, share 84% sequence identity and are conserved from yeast through man (Boulton et al., 1990). Both isoforms are ubiquitously expressed, have identical substrate specificity and kinetics of activation. The ERKs have been implicated in an extraordinary range of biological phenomena and are among the best studied protein kinases (Pearson et al., 2001; Roux and Blenis, 2004). Our lab (Selcher et al., 2001;
Selcher et al., 2003; Nekrasova et al., 2005) and others (Pages et al., 1999) have
49 generated and characterized ERK1 knockout mice. These animals exhibit no overt
phenotype, are viable, fertile and have normal life spans. Pages et al. reported a modest
reduction in T-cell differentiation (Pages et al., 1999), which we have been unable to confirm in our ERK1 null animals (Nekrasova et al., 2005). We have found that ERK1 null animals exhibit a greater sensitivity to experimental allergic encephalitis (Nekrasova et al., 2005) and defects in innate immune system responses (Dillon et al., 2004). ERK1 is specifically required for adipogenesis (Bost et al., 2005). The ERK1 null animals are behaviorally normal with respect to acquisition and retention of contextual or cued fear conditioning and passive avoidance (Selcher et al., 2001) and did not have any deficits in their ability to induce hippocampal LTP (Selcher et al., 2001; Mazzucchelli et al., 2002).
These animals are hyperactive. A detailed examination of the ERK1 null animals by
Brambilla and colleagues revealed that the mutant animals showed a paradoxical improvement in a striatal-based long term memory task and facilitation of LTP in the nucleus accumbens (Mazzucchelli et al., 2002). ERK2 activation in the brains of the mutant animals was only modestly greater, but was sustained for longer periods. This was interpreted as a compensatory action of ERK2 within specific brain regions and that
ERK1 deletion enhances the action of ERK2 in the brain. ERK2 activity in the absence of ERK1 is clearly illustrated in ERK1 knockout mouse models which display enhanced behavioral plasticity and gene expression in response to cocaine (Ferguson et al., 2006).
The ERK pathway had been previously implicated in the behavioral and molecular responses of cocaine as the MEK selective inhibitor SL327 abolished cocaine-induced hyper-locomotion, reward-seeking behaviors, and the expression of c-fos after acute administration of cocaine to mice (Valjent et al., 2000); however, data from the ERK1
50 knockout mouse line suggests that this response can be facilitated solely by ERK2.
ERK1 null mice were also noted to exhibit behavioral excitement profiles similar to that evoked by psychostimulants. When these mice were treated with lithium, olanzapine, or valproate, drugs commonly used to treat manic phases of ADHD or bipolar disorder, this behavior was reversed (Engel et al., 2008). These data suggest that ERK1 and ERK2 may have specific roles in behavior, or that a threshold of ERK activity may be required for proper signaling which is not fulfilled in the absence of ERK1.
Several groups have generated animals in which the ERK2 gene has been knocked out.
Inactivation of the ERK2 gene results in embryonic lethality as early as E6.5 (Hatano et al., 2003; Saba-El-Leil et al., 2003; Yao et al., 2003); our unpublished observations. Two of these recent reports identify the cause of lethality as a defect in trophoblast development, resulting in failure of placenta formation. In contrast, Yao et al. found that in their ERK2 null line, embryonic lethality occurred by E11.5 resulting from the failure to induce mesoderm formation. Examination of ERK2 null embryonic stem cells and chimeric embryos revealed that ERK2 is not required for stem cell proliferation but has been argued to act principally to promote cellular differentiation (Kuida and Boucher,
2004; Meloche et al., 2004). A recent report has suggested that ERK1 and ERK2 specific actions may not exist and the localization and expression level of each isoform has made it appear that each isoform has specific functions (Lefloch et al., 2008). This hypothesis stems from the identification of skewed expression patterns of ERK1 and ERK2 in specific tissues and cell types.
51 Localization of the ERK isoforms is conferred by scaffolding proteins and specific
docking sites which hold the kinases at particular locations. For example, KSR1 binds
both ERK isoforms to ensure their localization at the plasma membrane where they can
be phosphorylated by Raf isoforms. Furthermore, translocation of ERK to the nucleus
generally leads to gene transcription and differentiation. In this regard, PEA-15 is an adaptor protein which binds to ERK1/2 exclusively in the cytoplasm and prevents
ERK1/2 nuclear translocation. This is of biological relevance due to the ability of PEA15
to prevent proliferation which requires the nuclear translocation of ERK1/2. β-arrestin is
another scaffolding protein which binds ERK1/2 to facilitate signaling by the kinases
through the generation of signaling complexes at the endosome. The localization of
ERK1/2 at yet a different cellular compartment, the Golgi apparatus, is mediated by its
interation with Sef, which captures activated MEK-ERK1/2 complexes at the Golgi and
restricts signaling to cytosolic targets. The MEK partner-1, MP-1, is a small scaffolding
protein which also localizes ERK1/2 to the endosome, and facilitates nuclear signaling.
It is of interest that MP-1 is selective for association with ERK1 thereby restricting
signaling by isoform specificity (Kolch, 2005). These data demonstrate that both tissue
expression of each isoform as well as the compartmentalization of ERK1/2 helps to
regulate the intensity and duration of ERK activity as well as the target specificity.
It has been of substantial interest to determine the specific targets and actions of the
ERKs. ERK1/2 activation in neurons is elicited during the normal processes of synaptic
transmission by membrane depolarization following neurotransmitter receptor activation
(Adams and Sweatt, 2002; Thomas and Huganir, 2004). Importantly, although both
52 ERK1 and ERK2 are highly expressed in hippocampus and cortex, ERK2 is the
predominant isoform that is expressed (in a 13:1 ratio in nude mice (Lefloch et al., 2008)) and activated by these stimuli, with very little activation of ERK1 observed (English and
Sweatt, 1996, 1997). In contrast, both ERK isoforms are activated by neurotrophin stimulation. The recent generation of transgenic mice which are hypomorphic for only
ERK2 has implicated this isoform in a dominant role for memory and behavior. This mouse loses 20% of its ERK2 expression within the hippocampus and displays significant deficits in cued and contextual fear conditioning. These data suggest that a minor loss in ERK2 levels is sufficient to induce measurable behavioral changes.
53 Research Goals:
The evidence reviewed above demonstrates a gap in the understanding of how
ERK2 regulates cortical development. Three facts about ERK1 and ERK2
underscore the importance of this research. First, the ERKs regulate both signaling
components and machinery important to the cell cycle. Moreover, ERK activity is a
critical determinant in neural progenitor cell fate determination. And, mutations in
both upstream and downstream components of the MAPK cascade are causative for
MR. Because ERK1 knockout animals are largely normal and ERK2 activity is
prevalent throughout the developing telencephalon, it is likely that ERK2 plays a significant role in contributing to normal cognitive function through development of the cerebral cortex and postnatally, through downstream signaling. The goal of this dissertation is to discern if loss of only the ERK2 MAP Kinase isoform affects cortical development and cognitive function.
We have investigated the specific actions of ERK1 and ERK2 in newly developed animal
models. Initially, we investigated cortical development of a mouse in which ERK2 is
deleted in neural progenitor cells. We have found that the deletion of ERK2 results in
profound perturbations of cortical development and learning/memory tasks.
Furthermore, we have identified individuals with chromosomal microdeletions including
the MAPK1 gene that exhibit neurocognitive deficits. These observations provide the
first definitive evidence for ERK2-dependent regulation of neural progenitor cell
proliferation and differentiation, and emphasize its critical role in these processes.
Second, we assessed the outcome of loss of ERK2 in neural progenitor cells in an animal
54 which has a complete loss of ERK1. Our findings suggest that ERK2 plays a dominant
role over ERK1 in processes critical to cortical development as these brains are largely similar to those missing only ERK2.
55
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67 DELETION OF ERK2 MAP KINASE IDENTIFIES ITS KEY ROLES IN
CORTICAL NEUROGENESIS AND COGNITIVE FUNCTION
Ivy S. Samuels1, J. Colleen Karlo1, Alicia N. Faruzzi2, Kathryn Pickering3, Karl Herrup4,
J. David Sweatt2 , Sulagna C. Saitta3, and Gary E. Landreth1
1Department of Neurosciences
Case Western Reserve University
Cleveland, OH 44106
2Department of Neurobiology and McKnight Brain Institute
University of Alabama, Birmingham
Birmingham, AL 35294
3Division of Human Genetics
The Children's Hospital of Philadelphia
University of Pennsylvania School of Medicine
Philadelphia, PA 19104
4Department of Cell Biology & Neuroscience
Rutgers University
Piscataway, NJ 08855-8082
68 Abstract
The MAP kinases, ERK1 and ERK2 are critical intracellular signaling intermediates;
however, little is known about their isoform-specific functions in vivo. We have examined the role of ERK2 in neural development by conditional inactivation of the murine mapk1/ERK2 gene in neural progenitor cells of the developing cortex. ERK
MAPK activity in neural progenitor cells is required for neuronal cell fate determination.
Loss of ERK2 resulted in a reduction in cortical thickness attributable to impaired proliferation of neural progenitors during the neurogenic period and the generation of fewer neurons. Mutant neural progenitor cells remained in an undifferentiated state until gliogenic stimuli induced their differentiation, resulting in the generation of more
astrocytes. The mutant mice displayed profound deficits in associative learning.
Importantly, we have identified patients with a 1 Mb microdeletion on chromosome
22q11.2 encompassing the MAPK1/ERK2 gene. These children, who have reduced
ERK2 levels, exhibit microcephaly, impaired cognition, and developmental delay. These
findings demonstrate an important role for ERK2 in cellular proliferation and differentiation during neural development as well as in cognition and memory formation.
Introduction
The Extracellular signal-Regulated Kinases, ERK1/2, are amongst the most prominent
signal transduction molecules through which extracellular stimuli are propagated from
the cell surface to cytoplasmic and nuclear effectors. ERK1 (mapk3) and ERK2 (mapk1)
exhibit 84% sequence identity and are uniquely and coordinately activated through the
sequential phosphorylation of the classical three-tier mitogen-activated protein kinase
69 (MAPK) cascade. Significantly, although both ERK1 and ERK2 are expressed
throughout the brain (Selcher et al., 2001; Mazzucchelli et al., 2002), genetic inactivation
of ERK1 has only subtle phenotypic effects while ERK2 inactivation results in early
embryonic lethality (Aouadi et al., 2006).
The orchestration of cortical development occurs primarily through the actions of growth
factors which signal through receptor tyrosine kinases (RTKs) to control cell cycle
initiation, progression, and cell fate decisions in neural progenitor cells (NPCs). The
mammalian neocortex arises from self-renewing multipotent NPCs that initially generate
neurons and subsequently produce glia. During neurogenesis, NPCs within the
ventricular zone (VZ) divide symmetrically to produce two neurons, or asymmetrically to
generate a self-renewing progenitor and a daughter cell, which becomes either a neuron
or an intermediate progenitor cell (IPC). The nascent IPC continues to undergo
asymmetric renewal or, more frequently, divides symmetrically yielding two neurons
(Noctor et al., 2004; Pontious et al., 2008). Gliogenesis commences late in
embryogenesis, with undifferentiated NPCs responding to stimuli secreted by newly born
neurons (Miller and Gauthier, 2007). Disruption of the dynamics of NPC or IPC
proliferation and differentiation result in alterations in cell number, cellular identity and
organization of the cortex (Dehay and Kennedy, 2007).
Signal transduction through the sequential activation of elements of the ERK MAPK cascade directs neurogenesis while concurrently suppressing gliogenesis (Miller and
Gauthier, 2007). It has been demonstrated that alteration of MAPK signaling abrogates the generation of a fully populated, normal size cortex (Vaccarino et al., 1999; Menard et
70 al., 2002; Barnabe-Heider and Miller, 2003; Dono, 2003; Ohkubo et al., 2004; Zheng et
al., 2004; Paquin et al., 2005; Thomson et al., 2007). Inhibition of the upstream activator
of the MAPKs, MEK1, causes NPCs to remain in the SVZ/VZ in an undifferentiated
state, blocking neurogenesis (Menard et al., 2002; Barnabe-Heider and Miller, 2003;
Paquin et al., 2005). Similarly, mutation or inactivation of scaffolding proteins such as
FRS2 and SHP-2 which link RTKs to MAPK activation have analogous effects on
cortical progenitors (Hadari et al., 1998; Yamamoto et al., 2005; Gauthier et al., 2007; Ke
et al., 2007).
It has recently been appreciated that a group of genetic disorders termed neuro-cardio-
facial-cutaneous syndromes (including Cardio-Facio-Cutaneous, Costello, LEOPARD
and Noonan Syndromes) are caused by mutations in upstream elements of the
ERK/MAPK signaling cascade (Bentires-Alj et al., 2006; Roberts et al., 2006) and are
collectively characterized by distinctive cardiac and craniofacial defects, developmental
delay, and mental retardation. The mutation of downstream elements in the ERK cascade
has similarly been associated with mental retardation syndromes (Weeber and Sweatt,
2002). These observations suggest that perturbations in ERK MAPK signaling underlie a
diverse range of neurodevelopmental syndromes.
In order to determine the role of ERK2 in cortical development and function, we generated a murine model in which mapk1/ERK2 undergoes conditional inactivation at the peak of neurogenesis. The loss of ERK2 resulted in the generation of fewer neurons and many more astrocytes in the cortex. This change in cell fate is due to alterations in
71 the dynamics of NPC proliferation and specifically with a reduction in IPC division.
Behaviorally, the mutant mice display profound impairments in cued and contextual fear
conditioning. The identification of individuals with haploinsufficiency for the
MAPK1/ERK2 gene as a result of distal microdeletions at 22q11.2 (Shaikh et al, 2007),
enabled the first analysis of humans with reduced expression levels of ERK2. These
patients exhibit microcephaly and neurodevelopmental deficits, consistent with the
phenotype observed in the murine models presented here. Taken together, these studies
provide direct evidence for the unique actions of the ERK2 isoform in neural
development and suggest its involvement in cognitive function.
Materials and Methods
Generation of a conditional ERK2 allele. A targeting construct was designed using the
loxP system to excise exon 2 in the mapk1 gene, resulting in a null allele. A neo cassette
flanked by loxP sites was inserted into an ApaI site upstream of exon 2, and a loxP site
was cloned into an Xba I site between exons 2 and 3. The βADT diphtheria toxin
cassette was used for negative selection. The construct was linearized and electroporated
into 129 ES cells, and clones were selected following treatment with G418. Clones
positive for homologous recombination were further transfected with a Cre-expressing
vector, and resultant clones were analyzed for deletion of the Neo cassette. One clone
had deletion of only the Neo cassette and was expanded and injected in C57BL/6
blastocysts. Resultant chimeras were mated to establish germline transmission, and
heterozygote (Mapk1flox/+) animals were intercrossed to establish a Mapk1flox/flox line of mice. hGFAP-cre mice on an FVB/N background were obtained from Dr. Albee
72 Messing (University of Wisconsin-Madison) and interbred with the ERK2 floxed line to
generate Mapk1flox/flox;hGFAPcre/+ animals in which the ERK2 gene has been
conditionally knocked out (ERK2 CKO). All experiments were performed in accordance
with the Case Western Reserve University Institutional Animal Care and Use Committee.
Southern blotting. Genomic DNA derived from cortex was digested with Xba, resolved
on a 0.8% agarose gel and transferred to a nylon membrane. The DNA was crosslinked
and hybridized to a 32P radiolabelled, 500 bp 3’ probe (located distal to the targeted region of the mapk1 locus, between exons 3 and 4) using ExpressHyb (Clonetech,
Mountain View, CA).
Cell Culture.
Cortical Progenitor Cell Culture. Cortical progenitor cells were cultured from embryonic day 14.5 (E14.5) mouse embryos using protocols previously described by
Miller and colleagues (Menard et al., 2002; Barnabe-Heider and Miller, 2003). Briefly,
cortices were dissected from E14.5 mouse embryos in ice-cold HBSS and transferred to
Neurobasal medium containing 500 µM L-glutamine, 2% B27 supplement, 1% penicillin-
streptomycin (Invitrogen, Gaithersburg, MD) and 40 ng/ml FGF2 (BD, Bedford, MA).
The tissue was mechanically triturated with a plastic pipette and plated into eight-well
chamber slides (Nunc, Naperville, IL) precoated with 2% laminin (BD) and 1% poly-D-
lysine (Sigma, St Louis, MO) at a cell density of 50,000 cells per well. Cells were fed
every 2 days. For experiments with CNTF, cells were plated in FGF2 for 12 hours and
73 then CNTF (3 ng/ml, Peprotech, Rocky Hill, NY) was added to the media for an additional 2 or 4 days.
Astrocyte Culture. Astrocytes were derived from the brains of neonatal mice as previously described but with some modifications (Giulian and Baker, 1986). Cerebral cortices were isolated from postnatal day 1-3 mice. Cortices were minced and cells were dissociated in PBS containing 0.25% trypsin and 1 mM EDTA. Digestion was terminated by adding an equal volume of DMEM/F12 medium (Life Technologies, Gaithersburg,
MD) containing 20% FCS, and cells were triturated to obtain a single-cell suspension.
Cells were plated and media was replaced the next day with DMEM/F12 containing 20%
FCS.
Cell Counts. Quantification of immunocytochemical staining was performed on dissociated cells or cortical progenitor cell cultures. Counts were made from at least 3 embryos of each genotype obtained from a minimum of 3 litters. For each condition,
>300 cells from 9 fields of a randomly selected reference space per well were counted and analyzed from 2 wells per embryo. Digital image acquisition was performed on a
Leitz DM R B microscope with Leica DC 500 camera and manufacturer’s software
(Cambridge, UK).
Viability Assay. Cellular viability was assessed by 3-[4,5-dimethylthiazol-2-yl]-2,5- diphenyltetrazolium bromide (MTT; Sigma) assay. In brief, primary CKO and WT astrocytes (5 x 103/well) were seeded in a 96-well plate. At 2 and 4 days, 10 µl of MTT
74 (5 mg/ml PBS) was added to each well, and plates were incubated at 37°C for 2 h.
Medium was then removed and cells were resuspended in 100 µl of DMSO. Cell
viability was assessed by colorimetric change using the SpectraMax 340 PC plate reader
(Molecular Devices, Sunnyvale, CA) at = 550 nm.
Immunocytochemistry. Immunocytochemistry of cultured cells was carried out by
washing the cultures with HEPES-buffered saline (HBS), followed by fixation with 4%
paraformaldehyde for 15 min. The cells were permeabilized with 0.2% NP-40 in HBS,
followed by incubation in blocking with buffer containing 6% normal goat serum (NGS) and 0.5% bovine serum albumin (BSA) for 1-2 h at room temperature. Cells were then
incubated with primary antibodies in HBS containing 3% NGS and 0.25% BSA at 4°C
overnight. After washing with HBS, cells were incubated with secondary antibodies
prepared in HBS containing 3% NGS and 0.25% BSA at room temperature for 1 h. Cells
were then washed with HBS, and counterstained with 1 µg/ml 4',6'-diamidino-2-
phenylindole (DAPI) for 2 min. Primary antibodies used were anti-ERK2 (BD, 1:200),
anti-Nestin (Chemicon, Temecula CA 1:200), and anti-βIII tubulin (1:4000), anti-MAP2
(1:200), anti-S100β (1:200), anti-Vimentin IgM (1:200, each from Sigma).
Immunohistochemistry. For immunohistochemistry, 10 µm cryosections were rinsed in
PBS, blocked, and permeabilized with 0.1% Triton X-100, 10% goat serum in PBS.
Sections were incubated with primary antibodies at 4°C overnight, washed with PBS, and incubated with secondary antibodies at room temperature for 1 h. The sections were then
counterstained with DAPI for 2 min. The primary antibodies used were anti-Tbr1 (a gift
of Dr. Robert Hevner, 1:2000), anti-Otx1 (Developmental Hybridoma Bank, 1:10), anti-
75 Brn1 (Santa Cruz,1:50), anti-GFAP (DAKO, 1:500), anti-NeuN (Chemicon, 1:300), anti-
phosphoHistone H3 (Millipore, Billerica, MA,1:200), anti-Zebrin (1:50), anti-Reelin
(Santa Cruz, 1:50), anti-GABA (Sigma, 1:500), anti-Caspase 3 (Cell Signaling 1:100).
The secondary antibodies used were Alexa fluorophore-conjugated IgG or IgM. TUNEL staining was performed as described by the manufacturer’s protocol (DeadEnd
Fluorometric TUNEL staining kit, Promega). Golgi staining was carried out using the
Rapid GolgiStain kit from FD Neurotechnologies (Ellicot City, MD).
For confocal analysis of sections, images were obtained using a Zeiss LSM 510 confocal laser microscope (Zeiss, Gottingen, Germany) using Argon and HeNe lasers
(excitation lines, 488 and 594 respectively) and Plan-Neofluar 10x, N.A. 0.3, Plan-
Neofluar 20x, N.A. 0.5, and C-Apochromat 40x, N.A. 1.2W corr objectives.
Photomicrographs were processed using Adobe Photoshop and cell number quantified
with Image ProPlus software.
Cell Counts. Analysis was performed on at least 8 tissue sections obtained from
four mice in each group. Counts were made within the frontal approximately 2.0mm
proximal to Bregma) and parietal lobes (approximately 1.25mm distal to Bregma), owing to the robust recombination in these areas. The primary motor cortex (“medial”) and primary somatosensory cortex (“lateral”) were identified in the same section of each lobe
using standard stereotaxic coordinates (Paxinos and Franklin, 2001). The reference space
was fixed by the reticule of the microscope objective, measuring 150 µm in width. Cell
layers were identified based on morphological distinctions and distance from the pial
76 surface. Quantification of cell numbers was performed using the Image ProPlus software
by blinded observer.
Western blot Analysis. Cells were washed with ice-cold HBSS and lysed by sonication in
lysis buffer (20mM Tris pH 7.5, 150mM NaCl, 1% NP-40, 10% glycerol, 1mM EDTA,
1.5mM MgCl2, 20mM NaF, 20mM β-glycerophosphate) supplemented with protease
inhibitors (1µg/ml leupeptin, 1µg/ml aprotinin, 1mM PMSF, 1mM Na3VO4). Lysates
were cleared by centrifugation and protein concentration was determined using the
bicinchoninic acid assay (Pierce, Rockford, IL) using BSA as a standard. Equal amounts
of protein were boiled in sample buffer, separated by SDS-PAGE gels, and transferred to
Immobilon-P PVDF membranes (Millipore). Membranes were blocked in 3% BSA (or
5% skim milk powder for anti-G3PDH) in TBS and 0.1% Tween 20 (TBS-T) for 2 h at
room temperature and incubated overnight at 4°C with primary antibodies. The primary
antibodies used were rabbit anti-pERK (Cell Signaling, Danvers, MA 1:1000), mouse
anti-ERK2 (BD, 1:3000) directed to the C-terminus of ERK2 or mouse anti-ERK2
antibody that detects the N-terminus (Acris Antibodies, Herford Germany, 0.5 µg/ml),
mouse anti-ERK1 (Zymed 1:1000), anti-CrkL (Santa Cruz, 1:500), anti-G3PDH
(Trevigen, Gaithersburg, MD, 1:5000), anti-β-tubulin (Santa Cruz, 1:5000) and anti-c-src
(Santa Cruz, 1:500). After washing with TBS-T, membranes were incubated with HRP-
conjugated goat anti-mouse or anti-rabbit (1:5,000, GE Healthcare) secondary antibodies
in TBS-T with 5%milk for 2 h at room temperature. Blots were washed with TBS-T and
detection was performed using either Millipore or Pierce chemiluminescence reagents
using BioMax MR X-ray film (Eastman Kodak, Rochester, NY). Densitometric analysis
77 was performed using Adobe Photoshop histogram function, and statistical analysis was
done with GraphPad Prism software. Patient samples and clinical data were collected
after informed consent under an IRB approved protocol at the Children’s Hospital of
Philadelphia.
BrdU Labeling and Analysis. Pregnant mice were injected intraperitoneally with BrdU at
100 mg/kg of body weight. The animals were sacrificed 30 min after the injection.
BrdU-positive cells were detected by immunostaining as described by the manufacturers’
instructions with heat-mediated antigen retrieval in citrate buffer followed by 2N HCl-
treatment. Sections were incubated with a rat anti-BrdU antibody (Abcam, 1:1000)
overnight at 4oC. Quantification of BrdU+ cells during embryogenesis was performed by
dissociation of individual cortices and fixation in 4% paraformaldehyde immediately after sacrifice. Cells were plated at 50,000 cells per well and immunostained and counted as described above. For BrdU labeling of astrocytes, cells were plated at 50,000 cells per well in 8 well chamber slides overnight. Astrocytes were then treated with 1µg/ml BrdU for 24 hours and processed as described above.
Statistical Analysis. Statistical analysis was performed using either two-way ANOVA
with the Bonferroni post hoc test or the Student's t test, as indicated in the figures. In all graphs, the error bars indicate SD.
Behavioral Analysis. Animals used for behavioral analysis were bred at the University of
Alabama at Birmingham. They were housed with same sex siblings and with ad libitum
78 access to food and water, and they were maintained on a 12 hr light:dark cycle. Male
wild-type and ERK2 CKO littermates were compared in all behavioral experiments. All experiments were performed in accordance with the University of Alabama at
Birmingham Institutional Animal Care and Use Committee. All experiments were
performed with the experimenter blind to genotype.
The following assessments were performed similarly to those previously described in
(Selcher et al., 2001). Adult mice (14-40 weeks of age were tested for baseline
locomotor behavior and anxiety-like behavior using an open field test (Med-Associates,
Georgia, VT) which consisted of a 17 x 17 inch square polycarbonate box with three 16-
beam I/R arrays (measuring X, Y, and Z axis movement). Mice were placed into the
center of the open field to start the test, and activity was recorded for 15 min under
fluorescent lighting. Total distance moved was used as a measure of locomotor activity
and thigmotaxis (quantified by calculating the percent of total ambulatory distance that
was traveled in the center zone only of the open field) was used as a measure of baseline
anxiety-like behavior. Nociception and visual ability were performed as previously described (Crawley, 1999). Nociception was measured with a shock threshold test. One- second duration shocks were administered in increasing increments (0.08 mA, 0.10 mA,
0.15 mA, 0.25 mA, 0.35 mA, 0.45 mA, and 0.50 mA), each separated by 15-sec, and mice were observed for flinching, jumping, running, and vocalization at each increment.
The test was terminated when mice performed all four behaviors or when the highest shock increment was reached, whichever came first. Visual ability was measured using a
79 visual cliff test. All mice were observed during fear conditioning experiments for their
startle response to the onset of the tone used as a conditioned stimulus.
Fear Conditioning. Mice were tested for cued and contextual fear conditioning as previously described (Selcher et al., 2001) with some modifications. For training, animals were placed in a conditioning chamber with a metal grid floor (Video Fear
Conditioning, Med-Associates) for eight minutes. Training consisted of a 3-min exploration period followed by 3 tone-shock pairings consisting of a 30 sec-long 90dB tone with a 1-sec 0.5 mA shock occurring during the last second of the tone. Each tone- shock pairing was followed by a 60-sec rest except after the third pairing, which was followed by a 90-sec rest period leading up to the conclusion of the training session.
Twenty-four hours following training, each mouse was placed back into the training chamber for a 5-min contextual learning test, during which time freezing behavior was assessed in the absence of a tone or foot shock. One hour following the contextual test, cued learning was assessed by placing the mouse back into the chamber after the chamber’s context had been masked by altering its size, shape, color, and odor. After a
3-min exploration period of the novel context, the 90 dB conditioned stimulus tone was presented for 3 mins, during which time freezing behavior was assessed. For the repeated-training paradigm, animals were treated as noted, except that the training procedure was performed three times in the same day. Contextual and cued learning was then assessed 24 hours after the second training trial, using the procedure described above.
80 Results
ERK2 is required for normal development of the murine cerebral cortex
To directly assess the role of ERK2 in neural development, we generated a conditional mapk1/ERK2 null mutation. The mapk1flox allele was produced by introducing loxP sites into positions flanking exon 2 of the mapk1/ERK2 gene (Fig. 2-1A). Exon 2 is within the
ERK2 open reading frame and contains the highly conserved subdomain II, which is essential for catalytic activity of the kinase (Robinson et al., 1996). Transgenic mice in which cre recombinase expression is driven by the human glial fibrillary acidic protein
(hGFAP) promoter (Zhuo et al., 2001) were interbred with the mapk1flox/flox mice, resulting in deletion of ERK2 within cortical NPCs. Upon activation of the hGFAP promoter in these animals, cre expression led to the deletion of a 2kb genomic region that includes exon 2 of the mapk1/ERK2 gene. Cre-mediated recombination resulted in production of a 9.5kb fragment on the Southern blots (Fig. 2-1B). Western blot analysis of adult cortical brain homogenates confirmed that ERK2 protein levels in the cortex were reduced. No truncated fragments of ERK2 were detected by immunoblot analysis using an antibody directed to the N-terminus of the protein, confirming that the targeting strategy resulted in a null allele (Figure 2-1D). Expression of ERK1 was analyzed as a control and was not altered (Fig. 2-1C).
Mapk1flox/flox;hGFAPcre/+ mice are viable and fertile mice with normal life spans.
Macroscopic examination of these ERK2 Conditional Knockout (CKO) brains at P2 revealed no gross abnormalities (Fig. 2-2A) or differences in brain mass (CKO= 0.157g, n=6; WT= 0.167g, n=7; unpaired t-test, p=0.1466). Despite the normal overall
81 morphology and size of the ERK2 CKO cortex, we observed a significant reduction in cortical thickness throughout the dorsal telencephalon of CKO mice compared to wild- type littermates. A detailed regional analysis revealed a reduction in thickness of primary motor (0.8968), somatosensory (0.8673), auditory (0.8872) and visual (0.9042) cortices
(Fig. 2-2B). No deficits in the organization of specific cortical lamina were recognized; however, cellular density appeared greater throughout the ERK2 CKO cortex (Fig 2-2C).
ERK1 null mice were analyzed and were not different from wild-type controls (Selcher et al., 2001), establishing that specific inactivation of ERK2 causes the cortical thinning
(Fig. 2-2D).
The hGFAPcre/+ transgene is expressed in NPCs beginning at embryonic day 13.5 (Zhuo et al., 2001; Malatesta et al., 2003). This population of NPCs gives rise to >90% of
projection neurons and astrocytes populating the neocortex and hippocampus (Malatesta
et al., 2003), which we verified by interbreeding of Rosa26 reporter (R26R) mice
(Soriano, 1999) with mice carrying the hGFAPcre/+ transgene and subsequent LacZ
staining in the adult brain of cre+ progeny (Fig. 2-10). ERK2 is deleted throughout the dorsal telencephalon with only a minority of cells retaining expression. The ERK2 expressing cells include Tbr1+ cells populating layer VIb, and Reelin+ Cajal Retzius cells, both of which are generated prior to induction of cre expression. The majority of
ERK2-expressing cells in the mature cortex of the CKO mice are GABAergic interneurons, which arise and migrate from the ganglionic eminence that does not express the hGFAPcre/+ transgene (Supplemental Fig. 2-10). ERK2 expression was absent in the
hippocampus of the mutant mice; however, expression in the amygdala was similar to
82 that of wild-type animals, as the hGFAPcre/+ transgene is not expressed in this structure
(Supplemental Fig. 2-11).
We quantified cortical ERK2 protein levels at E14.5 and found a 50% reduction in ERK2
levels and a corresponding decrease in the levels of activated, phosphoERK2 in the
mutant mice. We did not detect any change in ERK1 expression (Fig. 2-3A,B) or
activation status at this time (Fig. 2-3C,D). Evaluation of these animals at E16.5 revealed
that the expression levels of ERK2 were reduced to 25% of wild-type levels, reflecting
the ongoing recombination of the mapk1/ERK2 gene and the half-life of the protein. The
level of active ERK2 was found to be 75% lower in the ERK2 CKO cells than those from
the wild-type animals (Fig. 2-3A,C) at E16.5. Although no compensatory increase in
ERK1 expression was found, phosphorylated ERK1 levels were dramatically (4-fold)
higher in the ERK2 CKO as compared to the wild-type cortex at this time (Fig. 2-3B,D).
These data provide evidence that while ERK1 expression was not elevated to compensate
for the loss of ERK2, there was a significant increase in ERK1 activity at E16.5.
ERK2 inactivation alters the cellular composition of the cortex
To determine the basis for the cortical thinning and assess the cellular density within the cortex (Fig. 2-2B,C), we quantified both total number of DAPI+ cells per cortical region
as well as the number of neurons (NeuN+ cells) in ERK2 CKO and wild-type littermates at postnatal day 10. The area from the VZ/SVZ boundary to the pial surface of the cortex
was analyzed in 150 µm sectors from sections of the frontal and parietal lobes. We found
no difference in DAPI+ cells per cortical region between genotypes, a finding consistent
83 with the observation of increased cellular density and reduced cortical thickness.
Strikingly, we found 10% fewer NeuN+ neurons per region in the ERK2 CKO as compared to wild-type littermates (Fig. 2-4A). However, there was a 40% increase in non-neuronal (DAPI; NeuN-), presumptive glial cells. These data indicate that ERK2 inactivation resulted in the production of fewer neurons and more non-neuronal cells, which was associated with an overall reduction in cortical thickness.
To determine if neuronal identity was affected by the loss of ERK2, a detailed examination of the number of neurons within each cortical lamina was performed.
Cortices from P2 wild-type and ERK2 CKO littermates were stained with antibodies to transcription factors whose expression is limited to specific cortical layers. We found a
35% reduction in number of Tbr1+ neurons in layer VI and the cortical preplate (Fig. 2-
4B) of ERK2 CKO animals. Similarly, we detected approximately half as many Otx1+ neurons populating layer V in the ERK2 CKO as compared to wild-type animals (Fig. 2-
4C). Evaluation of layers II-IV using Brn1 revealed an analogous, genotype-dependent
30% decrease in neurons populating this layer (Fig. 2-4D). No significant differences were found between medial and lateral regions of the cortex. Thus, inactivation of ERK2 was associated with an overall reduction in the numbers of neurons populating the cortex.
To confirm the identity of the non-neuronal (DAPI+; NeuN-) cells, we performed immunohistochemistry with astrocyte-specific markers. We found a dramatic increase in
Zebrin II+ (Fig. 2-5A) and GFAP+ astrocytes (Fig. 2-5B) throughout the cortex and subventricular zone. In order to assess the proliferative capacity of ERK2 CKO
84 astrocytes as compared to wild-type astrocytes, we cultured these cells from each genotype and evaluated their proliferation by measuring BrdU incorporation, Ki67 immunoreactivity and change in cell number (Fig. 2-5C,D,E respectively). We also assessed astrocyte metabolism and viability using an MTT assay (Fig. 2-5F). No significant differences were found in any of these parameters. We interpret these data to suggest that loss of ERK2 does not significantly affect astrocyte viability or their intrinsic ability to proliferate. We did not observe increasing numbers of astrocytes in older animals.
Thus, these data demonstrate that the loss of ERK2 from NPCs results in the reduction of neurons populating those cortical layers that are generated subsequent to its inactivation.
We further document that the increase in cellular density is caused by reductions in cortical thickness as the total number of DAPI+ cells is equivalent between genotypes.
We also demonstrate that many more astrocytes are present in the ERK2 CKO cortex.
We next investigated whether the reduction in neuronal cell number resulted from enhanced apoptosis. Analysis of E17 embryos by immunostaining for cleaved caspase 3 and by TUNEL assay revealed no differences between genotypes (Fig. 2-12). These data suggest that loss of ERK2 does not provoke neuronal death. Additionally, we assessed whether incomplete neuronal differentiation might contribute to the reduction in cortical thickness. Brains from four month-old CKO and wild-type littermates were Golgi- stained, and individual projection neurons within layers 3 and 5 of the primary motor cortex were traced and reconstructed with Neurolucida. We did not observe any
85 differences in dendritic length, branching, volume, or surface area (Supplemental Table 1
and 2). We also did not identify any structural changes in axonal projections.
ERK2 inactivation alters the dynamics of neurogenesis
The ERKs play critical roles in regulating cellular proliferation (Chambard et al., 2006)
and neuronal differentiation of cortical NPCs (Menard et al., 2002; Barnabe-Heider and
Miller, 2003; Paquin et al., 2005). To determine when the loss of ERK2 affects cell fate
determination, we investigated the proliferation of NPCs by evaluation of BrdU
incorporation. Pregnant dams were injected with BrdU, embryos were harvested 0.5 hr
post injection, and coronal sections of embryonic brains were immunostained with anti-
BrdU and anti-phosphohistone H3 antibodies. During S phase, BrdU is incorporated into
NPCs that are actively replicating DNA. Phosphorylated-Histone H3 (pH3) marks
mitotic cells during G2 and M phase of the cell cycle. At E12.5 (Fig. 2-6A), prior to the
induction of the hGFAPcre/+ transgene, the incorporation of BrdU and the number of ventricular cells undergoing mitosis was indistinguishable between genotypes. However,
when pregnant dams were injected on E14.5, the peak of the neurogenic period and one
day following initiation of hGFAP-cre expression, we observed significantly less BrdU
incorporation and pH3 immunoreactivity in the ERK2 CKO cortex (Fig. 2-6B). To
quantify the difference, individual embryonic cortices were dissected 30 minutes post-
injection, dissociated, plated, and immediately processed for immunostaining. We found
a 44% reduction in the number of BrdU+ cells in the E14.5 ERK2 CKO cortex
(CKO=6.7%; WT=11.87%, n=3 for each; unpaired t-test, p<0.0001). Importantly, we
observed a dramatic reduction in the number of abventricular pH3+ cells following ERK2
86 inactivation. These cells were identified as intermediate progenitor cells by co-labeling with Tbr2, a marker of this cellular population (Hevner et al., 2006) (Fig. 2-6D). There was no significant genotype-related difference in the total number of mitotic pH3+ cells
(CKO=101; WT=104.3, for each n=3, unpaired t-test, p=0.6005) as counted in tissue sections. Similarly, we did not observe any significant difference in the number of Tbr2+
IPCs or in Pax6+ progenitors (Fig. 2-13). However, the number of dividing IPCs (pH3+;
Tbr2+) was significantly reduced (by 37%) in the ERK2 CKO cortex (CKO=13%; WT=
20.67% per section, n=3 animals per genotype, unpaired t-test, p=0.0192).
In contrast to the results at E14.5, analysis of NPC proliferation at E16.5 revealed
dramatically more BrdU incorporation (Fig. 2-6C). Quantification of dissociated cortices
revealed 2.5 fold more BrdU+ cells in the CKO at E16.5 when compared to wild-type
littermates (CKO=9.41%; WT= 4.18%, n=4 animals per genotype, unpaired t-test,
p=0.005). A moderate increase in the numbers of pH3+, mitotic cells lining the ventricle
(Fig. 2-6C, E) was also observed in the ERK2 CKO; however, no increase in
abventricular pH3+ IPCs was identified (Fig 2-6E). Wild-type animals have passed
through the neurogenic period and exhibit few proliferating or dividing cells at this time.
Taken together, these findings suggest that ERK2 inactivation delays NPC proliferation
until E16.5, and suppresses the division of IPCs at the peak of the neurogenic period,
resulting in fewer neurons populating the ERK2 CKO cortex. The data also suggest a
reinitiation of progenitor proliferation during the period of normal gliogenesis in the
mutant cortex, resulting in more astrocytes.
87 ERK2 CKO neural progenitors generate fewer neurons in vitro
In order to evaluate whether ERK2 inactivation affected intrinsic cell fate determination
by NPCs, we performed in vitro analysis of cortical progenitor cell cultures (Menard et al., 2002; Paquin et al., 2005). Cortical progenitor cells from ERK2 CKO and wild-type
littermate embryos were cultured at E14.5 and grown under proliferating conditions in
Neurobasal media supplemented with FGF2 for 2 days. Importantly, we observed a
significant increase (1.3 fold) in the number of uncommitted, Nestin+ NPCs (Fig. 2-7A1-
2, E) in the ERK2 CKO cultures compared to those of wild-type littermates.
Furthermore, there were fewer immature βIII-tubulin+ (0.8 fold) and mature MAP2+ (0.7
fold) neurons in the ERK2 CKO cultures compared to those from wild-type animals (Fig.
2-7B, C, E). To assess if ERK2 CKO progenitors became committed to glial fates in
vitro, we counted S100β+ cells. ERK2 CKO cultures have 20% fewer S100β+ cells than
wild-type cultures (Fig. 2-7D-E), indicating that NPCs lacking ERK2 persist in an
undifferentiated state. Our finding is consistent with a previous report demonstrating that
NPCs transfected with a dominant negative MEK1-construct remain in an
undifferentiated state until gliogenic signals induce astrocyte differentiation (Paquin et al., 2005).
It has been suggested that ERK1/2 activation actively suppresses gliogenesis elicited by
CNTF/LIF (Bonni et al., 1997; Paquin et al., 2005). To assess if the loss of ERK2 affects cellular fate in a gliogenic environment, we added CNTF to cultures 12 hrs after plating.
A significant (1.4 fold) increase in vimentin+ astrocytes in the ERK2 CKO cultures was observed after treatment for either 2 or 4 days with CNTF (Fig. 2-8A1-4, D). Similar
88 results were found at both 2 and 4 days in vitro when cultures were assessed with the
mature astrocyte marker, GFAP (1.7 fold at 2DIV, 1.6 fold at 4DIV) and the immature
astrocyte marker, S100β (1.6 fold at 2DIV, 1.3 fold at 4DIV), (Fig 2-8B-F). These data
indicate that ERK2 normally acts to suppress gliogenesis and that this isoform alone may
be sufficient for directing cell fate decisions. In summary, our in vitro analysis confirms
and validates the conclusion that ERK2 participates in neurogenic cell divisions.
Furthermore, loss of ERK2 does not change the differentiative potential of the NPCs, as
they remain in an undifferentiated nestin+ state until gliogenic stimuli induce astrocytic
differentiation.
Mature ERK2 CKO mice display impaired associative learning
Due to the changes in cellular composition observed in the ERK2 CKO cortex, and the
critical roles the ERKs play in learning and memory (Selcher et al., 2001; Mazzucchelli
et al., 2002), we tested whether ERK2 CKO mice showed impairments in long-term
memory using two associative learning tests. Cued and contextual fear conditioning are
sensitive to disruption of the MAPK signaling pathway as MEK or MAPK inhibitors
have been shown to attenuate both forms of memory (Walz et al., 1999; Walz et al.,
2000; Sweatt, 2001; Shalin et al., 2004). ERK2 CKO mice exhibited significant deficits
in long term memory for both contextual and cued conditioning compared to littermate
wild-types (Fig. 2-9A,B). Further testing was conducted to determine whether
overtraining could rescue the deficits in contextual and cued conditioning; however, despite repeated training, ERK2 CKO mice continued to display significantly less freezing behavior during testing as compared to littermate wild-types (t15 = 3.65 and 3.35,
89 respectively, p<0.05; Fig. 2-9B). These data indicate a striking deficit in associative fear
memory in ERK2-deficient animals. Differences in freezing behavior could not be
explained by deficits in baseline sensory responses and baseline activities, as wild-type
and CKO mice showed no differences in locomotor behavior (CKO = 3300 cm, WT =
2450 cm, t15 = -1.536) or general anxiety-like behavior measured by thigmotaxis
(CKO=73.386%, WT=77.116%, t15 = 0.027) in the open field, in threshold to respond to
a footshock (flinch: CKO= 0.083 mA, WT= 0.087 mA, t15 = 0.98; jump: CKO= 0.369 mA, WT= 0.389 mA, t15 = 0.42; run: CKO= 0.141 mA, WT= 0.183 mA, t15 = 1.27;
vocalize: CKO= 0.275 mA, WT= 0.2560 mA, t15 = -0.45), or in visual ability (CKO=
71.429%, WT= 66.6670%, t15 = -0.19), which was measured with a visual cliff test; n=8
for CKO, n=10 for WT and p>0.05 for all baseline experiments. We conclude that the
selective loss of the ERK2 gene results in a pronounced associative memory deficit, in
the absence of generalized alterations in sensory responses.
MAPK1/ERK2 deficiency in humans is associated with learning deficits
The human MAPK1 gene encoding ERK2 is positioned on the long (q) arm of
chromosome 22. Chromosomal microdeletions frequently occur at 22q11.2 due to the
presence of highly homologous intrachromosomal low copy repeats (LCRs) that mediate
aberrant recombination events (Emanuel et al., 2001). The most common recurrent
deletion syndrome in humans involves the proximal LCRs (A-D) and leads to 22q11
Deletion/DiGeorge Syndrome (Fig. 2-14). Recently, a number of unrelated patients have
been identified with a 1 Mb deletion mediated by the distal LCRs, D-E (Rauch et al.,
1999; Saitta et al., 1999; Shaikh et al., 2007; Ben-Shachar et al., 2008). These deletions
90 include MAPK1/ERK2, and while they do not overlap the proximal 3Mb deletion causing
DiGeorge Syndrome, they independently exhibit a similar spectrum of craniofacial abnormalities, cardiac defects and neurodevelopmental defects. Several other genes including HIC2, YPEL1, PPIL2, UBE2L3, SDF2L1, and PPM1F occupy the D-E interval. A cluster of genes encoding the immunoglobulin lambda light chain are also located in the distal part of the deletion as well; however, none of these genes have been associated with human disorders (SCS, unpublished data). Thus far, the characteristics of
8 individuals that have DiGeorge-like symptoms but which carry atypical distal deletions have been identified (Rauch et al., 1999; Saitta et al., 1999; Shaikh et al., 2007; Ben-
Shachar et al., 2008). We have analyzed two patients with identical distal 1 Mb deletions. Patient 1 is a male with microcephaly, (head circumference consistently <3% for age and sex-matched norms measured over 4 years) and neurocognitive deficits
(Saitta et al., 1999) that include delays in fine motor function, expressive speech development, and in complex language comprehension. Patient 2 is a recently diagnosed female with microcephaly (head circumference <3% for age and sex-matched norms), a history of delays in expressive speech development and decreased academic performance overall. Analysis of cellular levels of ERK2 in lymphoblasts obtained from these patients with the distal deletion exhibit decreased levels of ERK2 expression relative to either normal individuals (n=4), or those with a typical 3 Mb DiGeorge deletion (n=1).
Densitometric analysis of western blots revealed that ERK2 expression in the distal microdeletion patients was 0.60876 of normal levels (p=0.0026). In contrast, levels of
CRKL, encoded by a gene located within the 3Mb classical deletion interval of DiGeorge syndrome were normal in patients with the distal 22q11 deletions. These data
91 demonstrate that patients with chromosomal microdeletions of distal chromosome 22q have a phenotype that includes microcephaly, neurodevelopmental and neurocognitive deficits, but is not due to genes in the DiGeorge interval. These children instead, have a small microdeletion that includes the MAPK1/ERK2 gene, show reduced ERK2 levels and display cognitive deficits. These findings are in line with our data from the murine model generated in this study where cortical hypoplasia and cognitive deficits in associative learning are found with genetic deletion of mapk1/ERK2.
Discussion
We have identified a specific role for ERK2 in the development of the cerebral cortex and shown that its deletion in the developing telencephalon is associated with the loss of higher order functionality in the mature brain. These findings provide concrete evidence for isoform-specific actions of the ERKs and follow from our previous studies demonstrating that mice in which the mapk3/ERK1 gene was knocked out did not have an overt phenotype and exhibited only modest behavioral changes (Pages et al., 1999;
Selcher et al., 2001; Mazzucchelli et al., 2002).
The developing cortex is exquisitely sensitive to perturbations of the tightly regulated patterns of cellular proliferation and differentiation. The conditional inactivation of
ERK2 within neural progenitors at the beginning of the neurogenic period resulted in fewer neurons populating the cortex. This effect resulted from the dramatic reduction in both the number of progenitors proliferating within the ventricular zone and the number of mitotic abventricular intermediate progenitor cells at E14.5. Importantly, the loss of
92 ERK2 appears only to delay initiation or stall the progression of neural progenitors through the cell cycle during the neurogenic period, as these cells neither die, nor migrate prematurely. It is of importance in this regard that isolated NPCs maintain their ability to undergo properly timed mitotic divisions, generating successive and appropriately specified lineages of neurons (Shen et al., 2006). Our findings demonstrate that loss of
ERK2 does not disrupt the lineage specification of neurons populating any of the cortical lamina. These findings are consistent with in vitro studies that have shown that ERK2 regulates initiation and progression through the G1 phase of the cell cycle (Liu et al.,
2004; Chambard et al., 2006; Meloche and Pouyssegur, 2007). The suppression of NPC proliferation as a consequence of ERK2 inactivation is transient, as we observed a robust increase in S phase-cells two days later (E16.5). The effects observed in the ERK2 CKO may reflect specific actions of this isoform or alternatively might result from a net reduction in overall ERK activity. Of particular relevance, Brambilla and colleagues have argued that ERK1 acts to functionally antagonize the actions of ERK2 in stimulating cellular proliferation, thus slowing progression of the cell cycle (Vantaggiato et al., 2006). However, Lenormand and colleagues have shown that each isoform is indiscriminately activated, and contend that the overall abundance of ERK1 and ERK2 dictates biological outcomes including cellular proliferation. Interestingly, the same report demonstrates that ERK2 is 13 times more abundant than ERK1 in the superficial cortex of nude mice (Lefloch et al., 2008). Our findings are consistent with the hypothesis that net ERK1/2 activity governs cellular behaviors. The striking increase in the levels of activated forms of ERK1 in the cortex of our ERK2 CKO mice at E16.5, following the near complete loss of ERK2 protein, suggests that the increase in ERK1
93 activity might reflect a compensatory response to the loss of ERK2, and this is correlated
with a resumption of NPC proliferation. An analogous compensatory effect on ERK2
activation was reported in a number of tissues in the ERK1 knockout mice, including the
brain (Pages et al., 1999; Selcher et al., 2001; Mazzucchelli et al., 2002). These findings
support the possibility that the biological actions of the ERKs are dictated by their spatial
and temporal expression, rather than the specific activity of one isoform or the other.
However, there is compelling evidence that isoform-specific actions of the ERKs may
arise from their selective association with molecular scaffolds that direct their interaction
with protein substrates and with phosphatases (Pouyssegur et al., 2002). These issues are
under active investigation.
The change in cellular generation by NPCs upon mapk1/ERK2 inactivation confirms and expands the findings from experimental manipulation of other components of the MAPK signaling cascade. Inhibition of the upstream activator of the ERKs, MEK1, causes
NPCs to remain in the SVZ/VZ in an undifferentiated state, blocking neurogenesis
(Menard et al., 2002; Barnabe-Heider and Miller, 2003; Paquin et al., 2005). Similarly, mutation or inactivation of scaffolding proteins which link receptor tyrosine kinases to
ERK activation have analogous effects on cortical progenitors. Mutations in the ERK cascade scaffolding protein FRS2 result in animals that exhibit nearly 30% reductions in cortical thickness and fewer cortical neurons (Hadari et al., 1998; Yamamoto et al.,
2005). This study confirmed that loss of cortical neurons was due to reduced numbers of proliferating IPCs and was a consequence of dramatically reduced levels of ERK activation. Two independent studies have recently found that deletion or mutation of
94 SHP-2, an upstream element of the ERK cascade, in cortical progenitors leads to the
inhibition of neuronal differentiation through ERK-dependent mechanisms (Gauthier et al., 2007; Ke et al., 2007). It has recently been proposed that IPCs are the only progenitors to undergo symmetric neurogenic divisions, contributing neurons to all cortical lamina throughout neurogenesis (Pontious et al., 2008). Our findings which document a decrease in neurons occupying each cortical lamina and a specific reduction in mitotic IPCs indicate that ERK2 may play a critical role in regulation of IPC division.
These data warrant further investigation into the precise roles that ERK2 plays in IPC cell cycle progression and terminal differentiation.
The conditional knockout of ERK2 also resulted in more astrocytes populating the neonatal and mature cortex. Bonni and colleagues originally demonstrated that ERK activation blocked the differentiation of NPCs into astrocytes (Bonni et al., 1997) and this conclusion has been validated both in vitro (Menard et al., 2002) and in animal models
(Paquin et al., 2005). Our findings are therefore consistent with the postulated action of the ERK cascade in suppressing glial fate and promoting neurogenesis in the developing cortex (Menard et al., 2002; Barnabe-Heider and Miller, 2003; Paquin et al., 2005;
Gauthier et al., 2007).
The ERKs are known to play critical roles in learning and memory and the absence of a robust behavioral phenotype in ERK1 null mice has suggested that ERK2 is the principal isoform responsible for learning behaviors which are dependent on the neocortex and associated structures (Selcher et al., 2001; Mazzucchelli et al., 2002). Fear conditioning
95 requires ERK-dependent interactions within and between the hippocampus, cortex and
amygdala. We demonstrate that the loss of ERK2 within the telencephalon resulted in
significant impairment on cued and contextual fear conditioning tests. Our findings are
similar to effects of pharmacological inhibition of the ERKs (Sweatt, 2004) or expression
of dominant negative MEK on fear conditioning (Shalin et al., 2004). Thus, the present
findings provide direct evidence that ERK2 is the principal isoform required for learning
in this behavioral task; however, it is unclear if the deficit is due to developmentally-
related structural changes of the cortex or the loss of ERK2 in mature neurons. It is
important to note that glutamatergic or electrical stimulation of neurons selectively
stimulates ERK2 activation (English and Sweatt, 1996; Mazzucchelli et al., 2002). Our
results compliment a recent report by Satoh and colleagues describing the behavior of an
ERK2 hypomorphic mouse that exhibits deficits in long-term memory in contextual and
cued fear conditioning paradigms (Satoh et al., 2007).
We have identified individuals with deletion of a single MAPK1/ERK2 allele and exhibit
decreased ERK2 protein levels. These patients’ clinical presentation include congenital
anomalies and neurocognitive deficits reflected by developmental delays, microcephaly,
and learning disabilities (Shaikh et al., 2007). Our findings in mutant mice suggest that
these abnormalities may arise from perturbation of neural developmental as well as
impaired ERK2 signaling required for memory and cognition. We postulate that the
cognitive deficits observed in the patients and in the mice result from a perturbation in
signal transduction through the ERK signaling cascade. We further suggest that these findings may have broader significance given the recent recognition that genetic
96 mutations within either upstream elements (Bentires-Alj et al., 2006) or downstream
targets (Weeber and Sweatt, 2002) of the ERK cascade are associated with developmental delay or mental retardation.
97 Figure 2-1. Generation of ERK2 Conditional Knockout mice. A, Schematic representation of gene targeting strategy. The shaded boxes on the mapk1/ERK2 locus and the targeting construct designate homologous regions identified for recombination.
Numbered boxes represent mapk1 Exons 2 through 4. The targeting construct contains a floxed Neo cassette upstream of Exon 2 and a loxP site between Exon 2 and 3. A βADT cassette was inserted downstream of Exon 3 for negative selection. Relevant restriction sites and the 3’ probe used for Southern blot analysis are indicated. Recombinant ES cells were transfected with a Cre plasmid to remove the Neo cassette, giving rise to the floxed mapk1 allele. Cre recombinase expression from the hGFAP-cre transgene produces a null allele. B, Southern blot of genomic brain DNA. Following Xba digestion of DNA, the 3’ probe identifies the wild-type allele as a 4.7kb fragment and the mutant allele as 11.5kb before and 9.5kb after recombination. C, Western blot analysis of adult cortical brain homogenates. Tissue was homogenized, separated by SDS-PAGE and immunoblotted with anti-ERK1, anti-ERK2 and β-tubulin antibodies. D, Western blot analysis of embryonic cortical homogenates. Tissue was lysed, separated by SDS-
PAGE and immunoblotted with an antibody directed to an epitope at the N-terminus of
ERK2. A lack of bands below 42 kD in both flox/flox and flox/flox cre+ samples demonstrates that no truncated protein fragments were produced by recombination. An antibody for c-src was utilized as a loading control.
98 Figure 2-1
99 Figure 2-2. ERK2 CKO mice display reduced cortical thickness. A, Dorsal view of P2
CKO (bottom) and wild-type (top) littermates. B, Quantitative analysis of cortical thickness. Corresponding coronal sections from CKO and wild-type brains were used to measure thickness of the frontal, parietal, temporal and occipital lobes corresponding to the motor, somatosensory, auditory and visual cortices. Measurements were taken from the ventricular zone to the pial surface (n=5), Two-way ANOVA p<0.0001 with
Bonferroni post-tests,***p<0.001, **p<0.01. C, Representative coronal sections of CKO and wild-type brains from P2, stained with DAPI. Note the difference in cellular density apparent between genotypes. D, Coronal sections of adult ERK1 -/- and wild type brains stained with the fluorescent Nissl stain, Neurotrace.
100
Figure 2-2
101 Figure 2-3. Loss of ERK2 expression and activity in ERK2 CKO cortices. A-D, E14.5
and E16.5 cortices were microdissected and prepared for Western blot analysis immediately upon dissection. The levels of ERK2, ERK1 and phospho-ERK1/2 were detected by immunoblotting and representative blots are shown. Densitometric analysis of the bands, normalized to β-tubulin, was performed for quantification and expressed as fold difference relative to wild-type. For A, quantification of ERK2 levels at E14.5
(CKO n=15, WT n=21, p<0.0001) and E16.5, (CKO n=7, WT n=8, p<0.0001). B,
Quantification of ERK1 expression levels at E14.5, (CKO n=13, WT n=18, p=0.4019) and E16.5, (CKO n=7, WT n=8, p=0.7772). C, Relative phospho-ERK2 activation at
E14.5, (CKO n=12, WT n=18, p=0.0093) and E16.5, (CKO n=7, WT n=8, p<0.0001), and D, relative phospho-ERK1 activation at E14.5, (CKO n=11, WT n=17, p=0.3594) and E16.5, (CKO n=4, WT n=4, p<0.0001).
102 Figure 2-3
103 Figure 2-4. Inactivation of ERK2 in neural progenitor cells results in the generation of
fewer cortical neurons. A, Analysis of cell density and cell fate in CKO and wild-type
littermates at P10. Corresponding coronal sections of P10 cortices were immunostained
with NeuN (green) and counterstained with DAPI (blue). Total cell number per cortical
region from the VZ to the pial surface was calculated by counting DAPI+ cells in both
medial (primary motor cortex, CKO=1386.167, WT=1398.167 avg cells per region, for
each n=3) and lateral (primary somatosensory cortex, CKO=1534.833, WT= 1571.167 avg cells per region, for each n=3) sections. Two-way ANOVA p=0.6179 and Bonferroni post-test, p>0.05. The fold difference in NeuN+ cells was calculated by Two-way
ANOVA p<0.0001, Bonferroni post-test p<0.0001. Fold difference in DAPI;NeuN- cells was calculated, Two-way ANOVA p=.0004, Bonferroni post-test p<0.01. B-D,
Corresponding coronal sections of P2 CKO (left) and WT (right) littermates were stained with anti-Tbr1 (B, green) anti-Otx1 (C, red) and anti-Brn1 (D, red) antibodies and counterstained with DAPI. Cortical lamina were identified based on morphology and distance from the pial surface. Immunoreactive cells were counted from 2 independent reference spaces of the cortex from at least 8 sections per genotype. For each, Two-way
ANOVA p<0.0001 and Bonferroni post-tests p<0.0001 were performed (n=4).
104 Figure 2-4
105 Figure 2-5. Inactivation of ERK2 in neural progenitor cells results in the presence of
more astrocytes within the cerebral cortex. A-B, Representative images of corresponding
coronal sections from neonatal (A) and mature (B) ERK2 CKO and wild-type littermates
immunostained with anti-Zebrin II (red) anti-GFAP (green) antibodies and counterstained
with DAPI. A higher magnification image is shown on the right. C, Primary astrocyte cultures from CKO and wild-type P2 pups were prepared and maintained for 5 DIV.
Cultures were incubated with BrdU overnight and fixed. Quantification of percent BrdU immunoreactivity (CKO n=4, WT n=3, p=0.9084). D, Quantification of Ki67 immunoreactive cells (CKO n=3, WT n=3, p=0.4166). E, Characterization of astrocyte proliferative index (CKO n=2, WT n=2, p=0.9827). F, MTT viability assay (CKO n=4,
WT n=5, p=0.4556).
106 Figure 2-5
107 Figure 2-6. ERK2 CKO mice display changes in the dynamics of NPC proliferation. A-
C, Pregnant mice were given intraperitoneal injections of BrdU at E12.5, E14.5 and
E16.5, respectively. Embryos were fixed 30 minutes after BrdU injection and
corresponding coronal sections of ERK2 CKO and wild-type littermates were double
labeled with anti-phospho-histone H3 (pH3+, red) and anti-BrdU (green) antibodies
followed by DAPI counterstaining (blue). Confocal images were taken at 10x and 20x
(A-B, scale bar=50 µm) and 10x and 40x (C, scale bar= 20 µm). D-E, Corresponding coronal sections of E14.5 (D) and E16.5 (E) embryos were immunostained with Tbr2 and phosphorylated Histone H3 followed by counterstaining with DAPI. Scale bar= 20 µm.
108 Figure 2-6
109 Figure 2-7. ERK2 CKO cortical progenitor cells exhibit reduced neuronal generation.
A, Cortical progenitor cell cultures from ERK2 CKO and wild-type E14.5 embryos were grown in Neurobasal with FGF2 for 2 days in vitro. Cells were then fixed and immunostained. Progenitors were identified by Nestin immunoreactivity (A, n=3).
Cultures were assayed for immature neurons with βIII Tubulin (B, n=3), mature neurons with MAP2 (C, n=4), and astrocyte precursors with S-100β (D, n=4). E, Analysis of cellular identity in culture with FGF2. Fold difference is the comparison of average of positive cells/total cell number relative to wild-type. Student’s t-test, p<0.001 for Nestin, p=0.014 for βIII-tubulin, p=0.0003 for MAP2 and p=0.0018 for S100β.
110 Figure 2-7
111 Figure 2-8. ERK2 CKO cortical progenitors generate more astrocytes in the presence of gliogenic stimuli. Cortical progenitor cell cultures from ERK2 CKO and wild-type E14.5 embryos were grown in Neurobasal media. Twelve hours after plating, cells were treated with CNTF to induce astrocyte differentiation. Cells were fixed and immunostained with anti-vimentin (A) anti-GFAP (B) and anti-S100β (C) antibodies after 2 days in vitro (1-2) and 4 days in vitro (3-4). D-F, Analysis of cellular identity with CNTF treatment. Fold difference in astrocyte generation was measured by the average of marker immunopositive cells/total cell number relative to wild-type At 2DIV and 4 DIV, (n=4 for each), Two-way ANOVA, p<0.0001 with Bonferroni post-tests, p<0.0001 for each.
112 Figure 2-8
113 Figure 2-9. Male ERK2 CKO mice have deficits in associative learning. Male ERK2
CKO mice displayed deficits in cued and contextual fear conditioning. A, 24-hr post- training, CKOs showed significantly less freezing behavior when tested for contextual and cued recall, t16 = 2.27 and 2.95, respectively, p<0.05. B, Repeated training, or
“overtraining,” did not overcome the deficit seen in cued and contextual fear conditioning. CKOs showed significantly less freezing behavior in both recall tests when tested the next day, t16 = 3.65 and 3.35, respectively, p<0.05.
114 Figure 2-9
115 Figure 2-10. ERK2 expression is abolished in neural progenitor cells beginning at
E13.5. A-B, Expression of hGFAP-cre activity. Sagittal sections of adult hGFAP- cre;R26R mice were stained for X-gal and counterstained with neutral red. High magnification inset in B demonstrates the extent of recombination throughout the cortex.
C, ERK2 expression in ERK2 CKO and wild-type mice at P2. Coronal sections of ERK2
CKO (left) and wild type littermates (right) were immunostained with an ERK2 specific antibody and visualized using DAB. D-I, Coronal sections of P2 ERK2 CKO and wild- type mice immunostained with an ERK2 specific antibody (red) and colabeled with antibodies directed to (D-E) Tbr1, the preplate/layer VIb marker (green), (F-G)
GABAergic interneuron marker, GABA (green) and (H-I) Reelin (green) which labels
Cajal-Retzius cells. The merged images and inset enlargements are shown in the far left panels of each.
116 Figure 2-10
117 Figure 2-11. ERK2 expression is reduced in the hippocampus, but not the amygdala in
ERK2 CKO mice. A-B, ERK2 expression was assessed by immunostaining with an anti-
ERK2 specific antibody. Corresponding coronal sections of adult ERK2 CKO (A) and
littermate wild-types (B) were evaluated for ERK2 expression in the hippocampus and
amygdala. Note that ERK2 is most highly expressed in the dentate gyrus (DG) and CA3
regions of the hippocampus and was lost in these areas in the mutant mice. No change in
ERK2 expression was identified within the basolateral amygdala (B/LA).
118 Figure 2-11
119 Figure 2-12. The ERK2 CKO cortex does not exhibit apoptosis. Analysis of apoptosis
was performed by immunostaining with anti-Caspase 3 antibody (A, DAB) and TUNEL
staining (B, green). Corresponding coronal sections from E17 ERK2 CKO (left) and wildtype (right) brains were stained with anti-Caspase 3 antibodies (note black arrow) and by TUNEL staining (white arrow).
120 Figure 2-12
121 Figure 2-13. The number of Pax6 and Tbr2 immunoreactive cells at E14.5 is not altered by conditional inactivation of ERK2. Cellular analysis was performed by immunohistochemistry of E14.5 cryosections with anti-Pax6 (red) and anti-Tbr2 (green) antibodies and counting of immunoreactive cells for each marker and for those cells expressing both markers. Two-way ANOVA, p>0.05 for each.
122 Figure 2-13
123 Figure 2-14. Patients with deletions of distal 22q11 have reduced ERK2 levels. A,
Schematic representation of Chromosome 22q11 with the centromere (CEN) to the left and telomeric (TEL) end to the right. Low copy repeats, LCRs are denoted by black boxes. The gene encoding CRKL is located between LCR-C and LCR-D and is included in the common 3Mb DiGeorge Syndrome deletion spanning from LCR-A to LCR-D.
The MAPK1 gene is located between LCR-D and LCR-E and is included in a 1 Mb distal
22q11 deletion occurring between these two LCRs. The D-E deletion interval is expanded to illustrate the other genes located within this 1Mb region. B, Lymphoblasts from normal subjects, distal 22q11 deletion patients, and DiGeorge Syndrome patients were lysed and prepared for Western blot analysis. The levels of ERK1, ERK2, and
CRKL were detected by immunoblotting and representative blots are shown. G3PDH levels were assayed for loading controls. C, Quantification of ERK2 levels from normal,
D-E deletion, and DiGeorge Syndrome patient lymphoblasts (normal, n=4, D-E, n=2,
DiGeorge, n=1, p=0.0026).
124 Figure 2-14
125 Table 2-1 and 2-2. Inactivation of ERK2 does not alter the morphology of cortical neurons.
126
Table 1- Parameters of basal dendrites of layer III pyramidal neurons
Order Quantity Length (µm) Surface area (µm2) Volume (µm3) ______CKO WT CKO WT CKO WT CKO WT ______1 47.67+ 1.76 60.67+ 5.69 122.72+ 9.80 156.52+ 2.62 910.44+ 101.61 1141.69+ 25.23 568.08+ 92.88 762.00+ 29.85 2 47.33+ 4.67 59.67+ 3.48 148.30+ 6.51 153.82+ 4.44 1085.68+ 75.67 901.99+ 48.78 660.29+ 71.69 515.32+ 32.39 3 38.00+ 1.15 25.67+ 0.33 * 158.15+ 3.65 135.16+ 6.58 1021.18+ 42.84 781.59+ 30.18 * 528.76+ 18.40 402.61+ 22.79 * ______
Mean values + SEM. * P< 0.05.
Table 2- Parameters of basal dendrites of Layer V pyramidal neurons
Order Quantity Length (µm) Surface area (µm2) Volume (µm3) ______CKO WT CKO WT CKO WT CKO WT ______1 42.67+ 5.90 61.67+ 1.76 132.53+ 24.96 156.30+ 26.47 1238.69+ 272.60 1151.01+ 279.62 994.20+ 228.48 728.64+ 199.65 2 41.67+ 5.49 42.33+ 3.93 142.68+ 32.83 169.34+ 31.41 1246.00+ 339.24 1323.47+ 310.27 922.62+ 260.06 882.67+ 219.33 3 14.00+ 1.15 18.33+ 1.20 149.35+ 22.53 181.39+ 37.52 1516.03+ 96.15 1409.52+ 328.76 978.03+ 161.41 905.58+ 219.87 ______
Mean values + SEM. * P< 0.05.
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131 ERK1 AND ERK2 ARE REQUIRED FOR CELL CYCLE REGULATION OF
NEURAL PROGENITOR CELLS
Ivy S. Samuels1, Joanna Pucilowska1, J. Colleen Karlo1, Natalie A. Cherosky1, Michiko
Watanabe2 and Gary E. Landreth1
1Department of Neurosciences
Case Western Reserve University
Cleveland, OH 44106
2Department of Pediatrics
Case Western Reserve University
Cleveland, OH 44106
Abstract
The canonical MAP Kinases, ERK1 and ERK2, are essential for fundamental cellular
processes such as proliferation and differentiation, as well as actions specific to the
nervous system such as long term potentiation and synaptic plasticity. We have
previously shown that the inactivation of ERK1 does not result in severe phenotypic or
behavioral deficits in mice, suggesting a redundant role for ERK2. Conditional
inactivation of the ERK2 isoform in murine neural progenitor cells causes significant
alterations in cell fate determination within the telencephalon and profound deficits in
associative learning tasks. The latter finding predicts that ERK2 plays a pivotal role in
cortical development and cognition which is not likely to be fully compensated for by
132 ERK1. To investigate the combined contribution of ERK1 and ERK2 to nervous system
development and function, we generated mice in which the genes encoding both ERK1
and ERK2 were inactivated in the telencephalon. ERK1/2 double knockout (DKO) mice
display a more severe phenotype than single knockout of either isoform and die during embryogenesis. ERK1/2 DKO mice exhibit changes in telencephalic cell fate determination similar to those of the ERK2 conditional knockout. We find that fewer
MAP2 neurons are generated and a greater number of neural progenitor cells remain undifferentiated during cortical development. Furthermore, we find a delay in S-phase entry as measured by acute BrdU incorporation during proliferative phases of
neurogenesis. These changes are associated with elevated levels of p27 and reductions in
cyclin D2 levels. Our results demonstrate that despite their role in cell cycle control, the
ERKs are not required for generation of the murine cerebral cortex.
Introduction
The Extracellular signal Regulated protein Kinases, ERK1 (mapk3) and ERK2 (mapk1)
are highly homologous protein kinases that play important roles in cellular proliferation
and differentiation. As the classical Mitogen Activated Protein Kinases (MAPKs), ERK1
and ERK2 are the central components of a canonical three-tier protein kinase cascade in
which extracellular signals emanating from a variety of cell surface receptors at the
plasma membrane stimulate cellular proliferation, differentiation, and survival
(Chambard et al., 2006).
133 The murine cerebral cortex is generated over a 6 day interval in which neural progenitor
cells undergo 11 cell cycles to produce 95% of neurons populating the adult neocortex.
Therefore, intricate regulation of cell cycle progression and termination are essential for
proper cortical development (Nowakowski et al., 2002). ERK1/2 activity has been
identified in the forebrain as early as E8.5 (Corson et al., 2003) and their actions within
neural progenitor cells have been shown to be essential for the proper generation of
pyramidal projection neurons as well as astrocytes that populate the cerebral cortex
(Menard et al., 2002; Paquin et al., 2005; Gauthier et al., 2007). Miller and colleagues have further demonstrated that ERK1/2 is not required for survival of NPCs, and proliferation of cultured NPCs in vitro is not strictly dependent on these kinases either
(Barnabe-Heider and Miller, 2003). Instead, neurotrophin signaling through MEK-
ERK1/2 is required for neuronal differentiation.
A differential role for ERK1 and ERK2 is implied from the individual knockout of each isoform. ERK2 activity in the extra-embryonic tissues at E6.5 is critical for survival as
ERK2 null mice die at this time due to the requirement for ERK2 in mesoderm and trophoblast formation as well as placental development (Hatano et al., 2003; Saba-El-Leil
et al., 2003; Yao et al., 2003). ERK1 knockouts do not exhibit phenotypic defects (Pages
et al., 1999; Selcher et al., 2001; Mazzucchelli et al., 2002). Conditional inactivation of
the ERK2 isoform in the mouse forebrain led to reductions in cortical thickness
associated with the generation of fewer neurons and more astrocytes. The dynamics of
cell cycle progression and exit were altered such that fewer proliferating neural
progenitor cells were identified at the peak of murine neurogenesis, E14.5. These results
134 indicated that ERK2 plays important roles in the control of neurogenic progenitor cell
divisions. However, we observed a dramatic increase in activated, phosphorylated ERK1
at E16.5 in the ERK2 mutant, suggesting that this isoform may have compensated for
ERK2 inactivation and alleviated a more severe developmental defect (Chapter 2).
Insight into the individual roles of each isoform in cell cycle regulation comes primarily
from in vitro studies. Within synchronized HeLa cells, downregulation of ERK1 induced
cell cycle arrest at G2, while loss of ERK2 led to cell cycle arrest and apoptosis in G1
(Liu et al., 2004). Furthermore, in NIH3T3 cells, ERK1 was postulated to suppress
ERK2 activity as the loss of ERK1 induced faster cell proliferation and facilitated cell
growth while down-regulation of ERK2 severely reduced cell proliferation (Vantaggiato et al., 2006). ERK1 null hepatocytes act essentially as wildtype cells; however, in double
null hepatocytes, cell proliferation is inhibited (Fremin et al., 2007).
These findings suggest that ERK2 plays key roles in the early stages of cell proliferation
which are not carried out by ERK1; however, Lenormand and colleagues have argued
that the expression of ERK2 and ERK1 dictates the specificity of isoform activation, as
opposed to individual activation and function of each (Lefloch et al., 2008). Their
analysis of isoform expression suggests that functions previously thought to be ERK isoform-specific may result from the lack of expression of one isoform or the other, not from differential activation.
135 In order to determine if combined inactivation of both ERK isoforms more severely
disrupts development of the cortex than loss of ERK2 alone, we generated a murine
model in which mapk1/ERK2 undergoes conditional inactivation at the peak of neurogenesis in a mapk3/ERK1 null background. We then analyzed cellular composition of the developing cortex of these animals in vivo and cell fate determination by neural progenitors in vitro. We further assessed neural progenitor cell proliferation throughout
neurogenesis and ERK1/2 dependent regulators of the cell cycle.
Materials and Methods
Generation of Mapk3-/-;Mapk1flox/flox;hGFAPcre/+ Mice. Mapk1flox/flox mice (Chapter 2)
were maintained on a mixed 129/B6 background by interbreeding. Congenic B6 mapk3-
/- mice (Nekrasova et al., 2005) were mated with mapk1flox/flox mice to generate doubly
hetereozgous mapk3+/-; mapk1flox/+ mice. These animals were intercrossed to generate
mapk3-/-;mapk1flox/flox mice. This line was maintained and DKO mice were generated by
first mating FVB/N hGFAPcre/+ mice with the mapk3-/-;mapk1flox/flox line. The progeny of
these matings (mapk3+/-; mapk1flox/+ ;hGFAPcre/+ ) were backcrossed to the mapk3-/-;
mapk1flox/flox line to garner litters carrying wildtype (mapk3+/-; mapk1flox/flox or flox/+),
ERK1 null (mapk3-/-;mapk1flox/fox or flox/+) and DKO (mapk3-/-; mapk1flox/flox; hGFAPcre/+) embryos. Primers for mapk3/ERK1 (5’-3’)
ExVI: CCAGGAGGACCTTAATTGCATCATT
3’ERK1: TTAGGGGCCCTCTGGCGCGGGTGGCTG.
Primers for mapk1/ERK2 (5’-3’) E2F-U: AGCCAACAATCCCAAACCTG, E2KOL:
CAACACCACCCCACTTACTA
136
Cortical Progenitor Cell Culture. Cortical progenitor cells were cultured from embryonic day 14.5 (E14.5) mouse embryos using protocols previously described by
Miller and colleagues (Menard et al., 2002; Barnabe-Heider and Miller, 2003). Briefly,
cortices were dissected from E14.5 mouse embryos in ice-cold HBSS and transferred to
Neurobasal medium containing 500 µM L-glutamine, 2% B27 supplement, 1% penicillin-
streptomycin (Invitrogen, Gaithersburg, MD) and 40 ng/ml FGF2 (BD, Bedford, MA).
The tissue was mechanically triturated with a plastic pipette and plated into eight-well
chamber slides (Nunc, Naperville, IL) precoated with 2% laminin (BD) and 1% poly-D-
lysine (Sigma, St Louis, MO) at a cell density of 50,000 cells per well. Cells were fed
every 2 days. For experiments with CNTF, cells were plated in FGF2 for 12 hours and
then CNTF (3 ng/ml, Peprotech, Rocky Hill, NY) was added to the media for an
additional 2 or 4 days.
Quantification of immunocytochemical staining was performed on dissociated cells or
cortical progenitor cell cultures. For each condition, >300 cells per well of at least two
wells were counted and analyzed. Digital image acquisition was performed on a Leitz
DM R B microscope with Leica DC 500 camera and manufacturer’s software
(Cambridge, UK). Photomicrographs were processed using Adobe Photoshop and cell
number quantified with Image ProPlus software.
Immunocytochemistry. Immunocytochemistry of cultured cells was carried out by
washing the cultures with HEPES-buffered saline (HBS), followed by fixation with 4%
137 paraformaldehyde for 15 min. The cells were permeabilized with 0.2% NP-40 in HBS,
followed by incubation in blocking with buffer containing 6% normal goat serum (NGS) and 0.5% bovine serum albumin (BSA) for 1-2 h at room temperature. Cells were then
incubated with primary antibodies in HBS containing 3% NGS and 0.25% BSA at 4°C
overnight. After washing with HBS, cells were incubated with secondary antibodies
prepared in HBS containing 3% NGS and 0.25% BSA at room temperature for 1 h.
Samples were then washed with HBS, and counterstained with 1 µg/ml 4',6'-diamidino-2- phenylindole (DAPI) for 2 min. Primary antibodies used were anti-Nestin (Chemicon,
Temecula CA 1:200), and anti-βIII tubulin (1:4000), anti-MAP2 (1:200), anti-S100β
(1:200). For immunohistochemistry, sections were rinsed in PBS, blocked, and
permeabilized with 0.1% Triton X-100, 10% goat serum in PBS. Sections were
incubated with primary antibodies at 4°C overnight, washed with PBS, and incubated with secondary antibodies at room temperature for 1 h. They were then counterstained
with DAPI for 2 min. The primary antibodies used were anti-Nestin, anti-MAP2, anti-
bIII-tubulin (all as described above), anti-Tbr1 (a gift of Dr. Robert Hevner, 1:2000),
anti-Tbr2 (Chemicon, 1:1000), anti-phosphoHistone H3 (Millipore, Billerica, MA,1:200)
and anti-p27 (BD Transduction Labs, 1:250). The secondary antibodies used were Alexa
fluorophore-conjugated IgG or IgM.
Western blot Analysis. Cells were washed with ice-cold HBSS and lysed by sonication in
lysis buffer (20mM Tris pH 7.5, 150mM NaCl, 1% NP-40, 10% glycerol, 1mM EDTA,
1.5mM MgCl2, 20mM NaF, 20mM β-glycerophosphate) supplemented with protease
inhibitors (1µg/ml leupeptin, 1µg/ml aprotinin, 1mM PMSF, 1mM Na3VO4). Lysates
138 were cleared by centrifugation and protein concentration was determined using the
bicinchoninic acid assay (Pierce, Rockford, IL) using BSA as a standard. Equal amounts
of protein were boiled in sample buffer, separated by SDS-PAGE gels, and transferred to
Immobilon-P PVDF membranes (Millipore). Membranes were blocked in 3% BSA (or
5% skim milk powder for anti-G3PDH) in TBS and 0.1% Tween 20 (TBS-T) for 2 h at
room temperature and incubated overnight at 4°C with primary antibodies. The primary
antibodies used were rabbit anti- pERK (Cell Signaling, Danvers, MA 1:1000), mouse
anti-ERK2 (BD, 1:3000), anti-β-tubulin (Santa Cruz, 1:5000), rabbit anti-G3PDH
(Trevigen, 1:5000), mouse anti-p27 (BD Transduction Labs, 1: 2500) and anti-Cyclin D2
(Thermo Scientific, 1: 1000). After washing with TBS-T, membranes were incubated
with HRP-conjugated goat anti-mouse or anti-rabbit (1:5,000; GE Healthcare) secondary
antibodies in TBS-T with 5%milk for 2 h at room temperature. Blots were washed with
TBS-T and detection was performed using either Millipore or Pierce chemiluminescence
reagents using BioMax MR X-ray film (Eastman Kodak, Rochester, NY). Densitometric
analysis was performed using Adobe Photoshop histogram function, and statistical
analysis was done with GraphPad Prism software.
BrdU Labeling and Analysis. Pregnant mice were injected intraperitoneally with BrdU at
100 mg/kg of body weight. The animals were sacrificed 30 min after the injection.
BrdU-positive cells were detected by immunostaining as described by the manufacturers’ instructions with heat-mediated antigen retrieval in citrate buffer followed by 2N HCl- treatment. Sections were incubated with a rat anti-BrdU antibody (Abcam 1:1000) overnight at 4oC.
139
Results
ERK1/2 DKO mice die embryonically, but display normal brain morphology.
To directly assess the combined roles of mapk3/ERK1 and mapk1/ERK2 in neural
development, we generated a mouse experiencing a conditional mapk1 null mutation with
a mapk3 null background. The mapk1flox/flox mouse line (previously described in Chapter
2) was interbred with a mapk3 null mouse line (Nekrasova et al., 2005) resulting in a
mouse with loxP sites surrounding exon 2 of mapk1 and a neo cassette replacing the span
between exons 1 and 7 of mapk3 (Fig. 3-1A). Transgenic mice in which cre recombinase expression is driven by the human glial fibrillary acidic protein (hGFAPcre/+) promoter
(Zhuo et al., 2001) were interbred with the mapk3-/-;mapk1flox/flox mice, resulting in the deletion of ERK2 within cortical NPCs in an ERK1 null environment at E13.5. PCR analysis confirms the recombination of exon 2 in mapk1/ERK2 represented by the presence of an amplified product, and the loss of mapk3/ERK1 in appropriate genotypes
(Fig 3-1B).
The mapk3-/-;mapk1flox/flox; hGFAPcre/+ (ERK1/2 Double Knockout, DKO) mice die
embryonically between E16.5 and birth. However, the DKO embryos display normal
morphology and size with no obvious phenotypic defects as compared to ERK1 knockout embryos or wildtype embryos at the same age (Fig 3-2A-C). It has been shown the the hGFAPcre/+ transgene is expressed in cardiac neural crest cells (Guris et al., 2001) and we have found that the hearts of the DKO mice have thin ventricular walls, and semi- penetrant ventricular septal defects which may cause the lethality (Fig. 3-2D-F). The
140 overall brain morphology and cortical structure at E14.5 is similar in the DKO, ERK1
knockout and wildtype (Fig. 3-2G-I).
Immunostaining of cortical cryosections from E16.5 DKO and ERK1 null littermates
demonstrated the loss of ERK2 in the developing neocortex (Fig. 3-3A). Western blot analysis of E14.5 cortical brain homogenates confirmed the activity of the hGFAPcre/+
transgene and revealed that ERK2 protein levels in the cortex were reduced to 25% of
mapk3/ERK1 -/- levels (Fig. 3-3B). Activity of ERK2 as measured by its
phosphorylation status was correspondingly reduced by 40% in the DKO at E14.5 (Fig.
3-3C). Cortical homogenates from ERK2 CKO mice revealed an increased level of
active, phosphorylated ERK1 at E16.5, suggesting a compensatory role for ERK1 with
the loss ERK2 (Chapter 2). As ERK1 is never expressed in the ERK1/2 DKO mice, this
phenomenon does not occur (Fig. 3-3C).
Loss of both ERK1 and ERK2 does not exacerbate the alterations in cellular
composition identified in the ERK2 CKO cortex.
Histological analysis of DKO cortices by cresyl violet staining showed no obvious
defects in cortical lamination or morphology (Fig 3-2G-I). At E16.5, it is difficult to
assess the 10-15% reduction in cortical thickness identified within neonatal ERK2 CKO
mice (Chapter 2). However, analysis of the cellular composition of the DKO cortex
revealed that fewer neurons are produced compared to littermate mapk3/ERK1 nulls by
this time point, as displayed by reductions in MAP2 and Tbr1 immunostaining (Fig. 3-4A
and Fig. 3-4B, respectively). This finding is comparable to the 10% reduction in NeuN+
141 neurons identified in ERK2 CKO mice at postnatal day 10. Furthermore, DKO embryos
maintain significantly more Nestin progenitors as compared to mapk3/ERK1 null littermates (Fig. 3-4C), a finding first recognized in the ERK2 CKO. No significant difference in the number of immature neurons immunostained with βIII-tubulin was
revealed between DKO and mapk3/ERK1 null embryos at E16.5 however (Fig. 3-4D).
Cortical progenitor cells cultured at E14.5 and maintained in vitro for 2 days in the
presence of FGF2 revealed analagous differences to the CKO in cellular generation. The
1.418 fold increase in nestin+ progenitors from mapk3/ERK1 -/- cultures as compared to
DKO cultures (p=0.0022, Fig. 3-5A) was correlated with fewer MAP2+ neurons (0.6935
fold, p=0.0001, Fig. 3-5C) and fewer S100β+ glial progenitor cells (0.7904 fold,
p=0.0006, Fig 3-5D). No difference in the percentage of βIII-tubulin+ neurons was
identified between genotypes (0.9501 fold different, p=0.0548, Fig 3-5B). Together, our
in vivo and in vitro findings suggest that ERK1 does not compensate for ERK2 during
cortical development, as the DKO mice do not display a more severe cortical phenotype
than ERK2 CKO mice.
DKO neural progenitor cells display inhibited cell cycle progression attributable to
reductions in cyclin D2 and increased p27Kip1 levels.
Reductions in intermediate progenitor cell divisions and an inhibition of neural
progenitor cells to proceed through G1 to S phase within the ERK2 CKO mice resulted in
the production of fewer neurons and more astrocytes as compared to wildtype littermates
(Chapter 2). To investigate if these processes were similarly affected in ERK1/2 DKO
142 embryos, pregnant dams were injected with BrdU at E14.5 and E16.5 and harvested 0.5
hr post injection. Embryos were either immediately fixed by immersion in 4%
paraformaldehyde or individual embryonic cortices were dissected, dissociated, plated
and immediately processed for quantitative immunostaining. We found that at E14.5,
BrdU incorporation was reduced by 0.6815 fold in ERK1/2 DKO embryos as compared
to mapk3/ERK1 -/- embryos (p= 0.0008, Fig. 3-6C). Coronal sections from embryos
reflect this difference (Fig. 3-6A,B). Interestingly, we found that at E16.5, BrdU
incorporation was also reduced in ERK1/2 DKO embryos as compared to mapk3/ERK1 null embryos, although to a lesser extent, 0.8032 fold (Fig. 3-6D-F). This finding contrasts the observed increase in BrdU incorporation in the ERK2 CKO at E16.5. Our current data confirms the hypothesis that the increase in BrdU incorporation observed in the CKO was due to elevated phospho-ERK1 levels at this time. Our data further suggests that ERK1 and ERK2 play similar roles in proliferation of neural progenitor cells as targeted deletion of both isoforms exacerbates the phenotype found in the ERK2
CKO.
We next investigated the effect of the deletion of both isoforms on progenitor cell division by analyzing the number of mitotic neural progenitor cells and intermediate progenitor cells in ERK1/2 DKO embryos at E14.5. As seen in the ERK2 CKO, fewer
Tbr2+ intermediate progenitor cells were positive for phosphorylated Histone H3 (pH3+) in the DKO as compared to both ERK1 null and wildtype embryos (Fig. 3-7). Our data therefore confirms that the reduction in neuronal production identified in our ERK1/2
DKO embryos is due to slowed cell cycle dynamics and fewer mitotic divisions.
143
ERK1/2 directly promotes cell cycle progression from G1 to S phase by inducing
transcription of the D-type cyclins through the phosphorylation of AP1 and Ets
transcription factors, Elk-1 and c-myc. Cyclin D1 forms a complex with cyclin dependent kinase 4/6 (cdk4/6) to propel the cell past the G1 restriction checkpoint.
Concurrently, ERK1/2 promotes the degradation of the cyclin dependent kinase inhibitor, p27Kip1, through phosphorylation of the ubiquition ligase, SKP2. p27 degradation directly promotes progression to S phase late in G1 as it initially inhibits the rate-limiting enzymatic complex for progression, Cdk2-Cyclin E. To investigate the effect of ERK1/2 loss on transition between G1 and S phase, we measured both CyclinD1/2 and p27 levels at E14.5. We found that ERK1/2 DKO cortices had higher expression levels of p27 than mapk3/ERK1 -/- littermates (Fig. 3-8A). Immunostaining of coronal sections reflects this difference (Fig. 3-8B). We also found that Cyclin D2 levels were lower in ERK1/2
DKOs as compared to mapk3/ERK1 null animals (Fig. 3-8C). Expression of Cyclin D1 was not consistently different between DKO and ERK1 null embryos (Fig. 3-8D).
Discussion
We have investigated the individual roles of ERK1 and ERK2 during neural development through the conditional inactivation of mapk1/ERK2 in a mapk3/ERK1 null mouse model. We find that until embryonic lethality, the cortices of these mice display the same phenotype as ERK2 CKO mice, whereby fewer neurons and more progenitors populate the cortical plate. We further demonstrate similar changes in the dynamics of cell cycle progression, as less BrdU incorporation occurs at E14.5. However, we now establish that
144 the increase in BrdU incorporation identified at E16.5 in the ERK2 CKO is associated with compensatory activation and signaling by ERK1, as the DKO mouse continues to display less BrdU incorporation compared to ERK1 null littermate controls at this stage of neurogenesis.
We have found that the expression levels of two downstream targets of ERK1/2, the cell cycle progressive cyclin, D2 is reduced, and the cyclin dependent kinase inhibitor, p27Kip1 is elevated in the DKO, at E14.5. The disruption of these cell cycle regulators was correlated with a reduction in intermediate progenitor cell division. These findings provide direct evidence for the unique actions of the ERK2 isoform in neural development and demonstrate that the ERK1 isoform does not play a significant role in neurogenesis. Because the DKO mice die before gliogenesis, we were unable to assess the capacity for astrocyte generation and proliferation in vivo. However, we speculate that gliogenesis would occur undisrupted in the ERK1/2 DKO mouse as in the CKO and hypothesize that more astrocytes would populate the cortex.
It is widely accepted that sustained ERK1/2 activity is required for cell cycle progression through G1 phase and for passage past the G1 restriction checkpoint into S phase. The primary mechanism by which ERK1/2 exerts these effects is through production of the
D-type cyclins, a rate-determining molecule in G1 progression (Villanueva et al., 2007).
Cyclin D complexes with cdk4 and leads to both the sequestration and subsequent degradation of the cell cycle inhibitor p27Kip1 as well as the catalytic actions of Cyclin
D-Cdk4. This enzyme complex furthers G1 progression through phosphorylation of the
145 pocket protein pRb, allowing for the release of E2Fs and initiation of E2F dependent gene transcription. The E2F dependent gene Cyclin A then complexes with cdk2 to propel the cell into S phase. Inactivation of the D-type cyclins in mouse models does not cause significant phenotypic changes in the brain due to compensation by Cyclin E (Chen et al., 2005); however, these mice do exhibit neurological abnormalities, overall small size, hypoplastic retinas and pregnancy-insensitive mammary glands which are due to lack of proliferation (Fantl et al., 1995; Sicinski et al., 1995). Our findings demonstrate reductions in Cyclin D2 levels. It has recently been shown that Cyclin D1 and Cyclin D2 display differential expression profiles in forebrain progenitor cell subsets. Specifically,
Cyclin D1 is highly expressed throughout the ventricular zone in early to mid- neurogenesis and decreases as development progresses, while Cyclin D2 expression rises from mid-neurogenesis to late neurogenesis and is more highly expressed in intermediate progenitor cells (Glickstein et al., 2007). It is noteworthy that we find both reductions in
Cyclin D2 and the number of mitotic intermediate progenitor cells. These data suggest that the ERKs may play specific roles in proliferation of intermediate progenitor cells.
Macrophage-colony stimulating Factor-1 (CSF-1) induces cyclin D1 and D2 through combinatorial activation of src, myc, and ERK1/2. Complete inhibition of MEK/ERK
downstream of CSF-1 stimulation causes a 50% decrease in D2 expression, similar to the
level of reduction identified in our mouse model. Cyclin D2 expression is regulated by
the Myc/Max/Mad network (Bouchard et al., 2001) and it has been shown that activation
of myc’s transactivating activity selectively induces D2 but not D1 (Dey et al., 2000).
These findings provide a mechanism by which ERK1/2 inactivation could lead to the
146 selective reduction in Cyclin D2 but not D1. It is of significance that mice harboring a
null mutation in CSF-1 have alterations the cortical circuitry that balances excitation and
inhibition. (Michaelson et al., 1996)
Our data revealed elevations in p27kip1 in the DKO cortex. Animals in which p27Kip1
has been knocked out exhibit increased cortical thickness resulting from a 27% increase
in neurons in upper cortical lamina (Goto et al., 2004). Conversely, overexpression of
p27Kip1 in the developing telencephalon slows progenitor proliferation with fewer
neurons populating the upper cortical layers resulting in reduced cortical thickness (Tarui
et al., 2005). The conditional knockout of the cell cycle regulator pRb in the developing
telencephalon also causes a robust (7 fold) increase in progenitor proliferation resulting in
a dramatic increase in telencephalon size (Ferguson et al., 2002). Intriguingly, removal
of p27 in a cyclin D1 null animal restored cell cycle control and abolished the phenotypic
defects (Geng et al., 2001). Expression levels of p27 are also regulated by c-myc. Myc
stimulates the expression of Cyclin D2 and Cdk4 which in turn sequester p27 in a
p27/CyclinD2/Cdk4 complex, targeting it for degradation (Bouchard et al., 1999; Coller et al., 2000; Dey et al., 2000; Bouchard et al., 2001). These data support our identification of reduced Cyclin D2 levels and increased p27 levels in the DKO cortex.
Furthermore, the alteration in expression of these proteins is sufficient to induce the
change in cellular generation identified in the ERK1/2 DKO. It is of interest to determine
if a reduction in p27 levels would reverse these deficits in neuronal generation.
147 Our findings also reveal that the canonical ERK1/2 signaling pathway is not solely
responsible for generation and differentiation of cortical neurons. In the absence of
ERK1 and near complete loss of ERK2 at the peak of neurogenesis, neurons are still
made. It appears that the ERKs regulate cell fate through the critical maintenance of cell
cycle control. It has been shown that the non-classical ERK5 isoform is important for
neuronal generation and survival but does not regulate cortical progenitor cell
proliferation apoptosis (Liu et al., 2006; Nishimoto and Nishida, 2006). Furthermore,
ERK5 is able to regulate Cyclin D1 expression (Mulloy et al., 2003), providing an
additional potential explanation for the lack of reductions in this cell cycle regulator when ERK1/2 are inactivated. Additionally, Cyclin E can compensate for any reductions in Cyclin D activity, allowing for the generation of a generally normal cerebral cortex.
The ERK1/2 DKO mice are recoverable to E16.5 as ERK2 inactivation does not occur until E13.5 and the ventricular septal defects are not lethal until later in development. A lack of information regarding expression of the GFAP-cre transgene in the heart along with ambiguous expression of GFAP itself in this organ makes it difficult to ascertain exactly where recombination occurs to lead to the VSDs observed. Furthermore, the incomplete penetrance of the phenotype does not clarify the deficit. Importantly, mice in
which the ERKs have been inactivated in the developing cardiac neural crest develop
aortic arch defects and associating ventral septal defects (Newbern et al, submitted).
These mice have been generated as models of 22q11 Deletion Syndrome as they
phenocopy the disease phenotypes. We propose that the ERK1/2 DKOs generated here
can be useful in identifying and understanding the cortical phenotypes associated with
148 this disease. Because over 25% of 22q11 Deletion syndrome patients develop schizophrenia and affective mood disorders, this mouse model is also important for delineating the underlying mechanisms of these disorders.
149 Figure 3-1. Generation of ERK1/2 Double Knockout mice. A, Schematic representation of gene targeting strategy. The numbered boxes represent mapk3 exons 1 through 9 and mapk1 exons 2 through 6. A gene targeting strategy was designed to create a null allele for mapk3. Following homologous recombination, the mapk3 allele contains a floxed
Neo cassette replacing genomic sequence spanning from exon 1 to exon 7. For mapk1, we created a floxed allele with loxP sites flanking exon 2. Following hGFAP cre- mediated recombination, loss of exon 2 results in a null allele. B, PCR of genomic DNA isolated from embryonic cerebral cortices. An ERK2 KO reaction was designed to generate a product in the absence of Exon 2. For ERK1, the upper band represents the endogenous ERK1 allele. The lower band is a non-specific PCR product.
150 Figure 3-1
151 Figure 3-2. ERK1/2 DKO embryos die embryonically but display normal brain morphology. Lateral view of a wildtype (A), ERK1 null (B) and ERK1/2 DKO (C) littermate embryos at E14.5. Horizontal sections of a wildtype B6 mouse heart (D),
ERK1 null heart (E) and the heart from an ERK1/2 DKO (F) embryo at E14.5 stained with hematoxylin and eosin. Coronal sections of cortices from E14.5 wildtype (G),
ERK1 null (H) and ERK1/2 DKO (I) littermates stained with cresyl violet.
152 Figure 3-2
153 Figure 3-3. Loss of ERK2 expression and activity in ERK1/2 DKO cortices. A,
Immunostaining of coronal cortical sections from E16.5 ERK1/2 DKO and ERK1 null littermates using an anti-ERK2 antibody demonstrated the loss of ERK2 in the developing neocortex. B, E14.5 cortices were microdissected and prepared for Western blot analysis immediately upon dissection. The levels of ERK2 and phospho-ERK1/2 were detected by immunoblotting and representative blots are shown. Densitometric analysis of the bands, normalized to β-tubulin, was performed for quantification and expressed as fold difference relative to wild-type. For B, Quantification of ERK2 levels at E14.5 (DKO n=9, ERK1 null n=6, p<0.0001). For C, Relative phospho-ERK1/2 activation at E14.5, (DKO n=5, ERK1 null n=3, p=0.0444).
154 Figure 3-3
155 Figure 3-4. Inactivation of ERK1 and ERK2 in neural progenitor cells results in the generation of fewer cortical neurons. A-D, Analysis of cellular composition at E16.5.
Corresponding coronal sections of ERK1/2 DKO (left) and ERK1 null (right) littermates were immunostained with anti-MAP2 (A, green) anti-Tbr1 (B, red) anti-Nestin (C, green)
and anti-βIII tubulin (D, red) antibodies.
156 Figure 3-4
157 Figure 3-5. ERK1/2 DKO cortical progenitor cells exhibit reduced neuronal generation.
A, Cortical progenitor cell cultures from ERK1/2 DKO and ERK1 null E14.5 embryos were grown in Neurobasal with FGF2 for 2 days in vitro. Analysis of cellular identity in culture was performed by immunostaining with various cellular markers. Fold difference is the comparison of the average number of positive cells/total cell number relative to
ERK1 null. Progenitors were identified by Nestin immunoreactivity (A, n=5 for each, p=0.0022). Cultures were assayed for immature neurons with βIII Tubulin (B, n=5 for each, p=0.0548), mature neurons with MAP2 (C, for each n=4, p=0.0001), and astrocyte precursors with S-100β (D, n=5 for each, p=0.0006).
158 Figure 3-5
159 Figure 3-6. ERK1/2 DKO mice display reductions in proliferation. Pregnant mice were given intraperitoneal injections of BrdU at E14.5 (A-C) and E16.5 (D-F), respectively.
Embryos were fixed 30 minutes after BrdU injection and corresponding coronal sections of ERK1/2 DKO and ERK1 null littermates were double labeled with anti-phospho- histone H3 (pH3+, red) and anti-BrdU (green) antibodies. Cortices were similarly dissected, dissociated, plated and immediately fixed for immunostaining and quantification of BrdU immunoreactivity. For E14.5, DKO n= 5, ERK1 null n=5, p=0.0008 (C), For E16.5, DKO n=3, ERK1 null n=3, p= 0.0028 (F).
160 Figure 3-6
161 Figure 3-7. Fewer abventricular intermediate progenitor cells from ERK1/2 DKO embryos are mitotic at the peak of neurogenesis. Corresponding coronal sections of
E14.5 ERK1/2 DKO (A,D), ERK1 null (B,E) and wildtype (C,F) embryos were immunostained with Tbr2 (green) and phosphorylated Histone H3 (red).
162 Figure 3-7
163 Figure 3-8. DKO neural progenitor cells exhibit reductions in cyclin D2 and increased p27Kip1 levels. A-D, Analysis of cell cycle components. E14.5 cortices were microdissected and prepared for Western blot analysis immediately upon dissection or corresponding coronal sections from littermates were cryosectioned and immunostained.
The levels of p27Kip1 (A, B), Cyclin D2 (C) and Cyclin D1 (D) were detected by immunoblotting and representative blots are shown. Densitometric analysis of the bands, normalized to β-tubulin or G3PDH, was performed for quantification and expressed as fold difference relative to wild-type. For A, Quantification of p27 levels at E14.5 (DKO n=9, ERK1 null n=6, p=0.0829). B, Corresponding coronal sections were immunostained using an anti-p27 antibody (green). For C, Relative Cyclin D2 at E14.5, (DKO n=3,
ERK1 null n=2, p=0.0690) and D, representative western blot analysis of Cyclin D1 expression.
164 Figure 3-8
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168 DISCUSSION
ERK1 and ERK2 are ubiquitously expressed kinases which are activated by a diverse
variety of stimuli (Cobb et al., 1991). The importance of their expression and activation
is demonstrated by the existence of two isoforms which are 90% homologous when the
N-terminus is excluded from comparison (Boulton and Cobb, 1991). Both isoforms are
expressed in all species from yeast through human, with the exception of only the frog
and chick (which express only ERK2; (Lefloch et al., 2008). These facts suggest that either isoform specific functions exist, or that the two isoforms provide functional redundancy which ensures signaling fidelity.
Both ERK1 and ERK2 are expressed at high levels in the mature rodent cortex (Boulton et al., 1990; Boulton and Cobb, 1991; Ortiz et al., 1995). Furthermore, ERK signaling is required for both cortical development (Miller and Gauthier, 2007) and for long term potentiation and learning and memory in adult animals (Sweatt, 2001). Therefore, we utilized the murine cerebral cortex as a system in which to investigate the potential of isoform-specific actions of ERK1 and ERK2.
Isoform specific expression regulates isoform specific
function
We, and others, have previously demonstrated that ERK1 knockout mice do not display
significant phenotypic changes in the cortex (Pages et al., 1999; Selcher et al., 2001;
169 Mazzucchelli et al., 2002). However, it was established that ERK2 activity is elevated in
the absence of ERK1, demonstrating a potential compensatory role for ERK2. In ERK2 conditional knockout mice we found a reduction in cortical thickness which is associated with changes neural progenitor cell proliferation and differentiation such that fewer neurons and more astrocytes populate the cortex (Chapter 2). While no difference in
ERK1 activity at E14.5 was observed, we found a significant increase in ERK1 activity at
E16.5, three days after recombination was initiated. This elevated level of activity suggests that ERK1 may compensate for the absence of ERK2. However, the identification of specific perturbations in cortical cell fate which cause a defect in cortical thickness demonstrates the possibility of an ERK2-specific role in neural development.
To address this possibility, we generated double knockout mice, where both isoforms are inactivated in the developing cortex. The phenotype of the cortex in this mouse is slightly more perturbed than the single ERK2 conditional knockout at the time of embryonic death occurring in utero at about E17 (Chapter 3). The lethality of this mouse can be attributed to heart defects and is unlikely to be related to ERK1/2 inactivation within the nervous system. Because the loss of both ERK isoforms did not significantly exacerbate the cortical defects identified in the single ERK2 conditional knockout, it appears that ERK1 may not compensate for ERK2. This suggests that either ERK1 does not play a substantial role in cortical development, or that ERK1 is not expressed at high levels in the cells contributing to elaboration of the cortical mantle. We have visualized expression of ERK1 from dissociated cortices at E14.5 and E16.5 by western blotting techniques, but have been unable to observe ERK1 expression by immunohistochemistry on cryosections from animals of the same age. Therefore, it is difficult to assess the
170 location of ERK1 action at this time. The lack of a more severe phenotype in the DKO as
compared to the ERK2 CKO can be explained if ERK1 is expressed in post-mitotic
neurons but not neural progenitor cells. Lennormand et al., have recently assessed expression of ERK1 and ERK2 in adult nude mice. They demonstrate that ERK2 is 13 times more abundant than ERK1 in the prefrontal cortex (Lefloch et al., 2008).
Furthermore, studies of ERK1/2 protein expression in rats demonstrates that the
ERK1:ERK2 ratio is 0.16 and 0.28 in the frontal and parietal cortices, respectively (Ortiz et al., 1995). Analysis of ERK1 and ERK2 mRNA in the CNS by in situ hybridization revealed that there were no regions where ERK1 mRNA was present which did not also exhibit ERK2 mRNA localization. Furthermore, ERK2 mRNA was more widely distributed and displayed a higher level of expression than ERK1 throughout all cortical layers (Thomas and Hunt, 1993). These data argue that the modestly exacerbated phenotype that we documented in the double knockout as compared to the ERK2 CKO
(Chapter 3) may result from the low level of expression and activity of ERK1 at this time and place, not from an ERK2 specific function. However, within these experiments,
ERK1 and ERK2 mRNA was not measured at the critical developmental periods or locations.
A few examples of putative ERK1 and ERK2 specific functions have been described.
The most compelling evidence is the potent activation of ERK2 but not ERK1 in response to high frequency tetanic stimulation, which induces LTP in the hippocampus
(English and Sweatt, 1996, 1997). ERK1 deficient mice display deficits in thymocyte maturation (Pages et al., 1999), adipocyte differentiation (Bost et al., 2005a; Bost et al.,
171 2005b), and striatum-dependent synaptic plasticity (Mazzucchelli et al., 2002).
Additionally, ERK1 and ERK2 have been shown to play differential roles in certain phases of the cell cycle, depending on the cell line. In hepatocytes, ERK2 ablation inhibits proliferation while ERK1 silencing has no effect (Fremin et al., 2007). In HeLa cells, ERK1 inactivation arrests the cell cycle at the G2/M transition, while ERK2 inactivation results in cell arrest within G1 phase (Liu et al., 2004). Contradictory results arise from experiments performed with NIH 3T3 fibroblasts: silencing of ERK1 initially demonstrated that this isoform was involved in suppression of cell proliferation and
ERK2 was involved in the promotion of cell division; however, later studies demonstrated that differential expression of each isoform contributed to these results and that both ERK1 and ERK2 positively control cell proliferation (Vantaggiato et al., 2006;
Lefloch et al., 2008). In embryonic stem cells, ERK1/2 activity is required for self- renewal but only loss of ERK2 affects trophoblast stem cells (Meloche et al., 2004).
These data correspond with a specific role for ERK2 in trophoblast and mesoderm development of the embryo, as ERK2 knockout mice die during development and ERK1 cannot compensate (Hatano et al., 2003; Saba-El-Leil et al., 2003; Yao et al., 2003).
Both isoforms play a role in the positive selection for T-cell differentiation (Nekrasova et al., 2005; Agrawal et al., 2006), whereby inactivation of ERK1 and ERK2 results in more severe defects than loss of ERK1 alone. These results support the hypothesis that at least a significant proportion of presumed ERK1 and ERK2 specific functions are related to expression levels in a given environment (Shaul and Seger, 2007). It is important to note that the relative expression of each isoform has not been assessed in detail in many of the
172 tissues and cell types noted above, mainly due to the high sequence homology between the isoforms and antibody cross-reactivity.
It is of interest however, that the immediate activators of ERK1/2, MEK1 and MEK2 have both redundant and distinct functions. Like the ERKs, knockout of each isoform leads to vastly different results. MEK1 knockout mice die due to placental defects while
MEK2 knockouts are overtly normal in growth and development (Giroux et al., 1999;
Belanger et al., 2003). Furthermore, knockdown of each isoform leads to different outcomes in cell cycle control. MEK1 seems to promote proliferation while MEK2 inhibits this process through upregulation of the CKI, p21 (Ussar and Voss, 2004). A recent report has further demonstrated that MEK1 shuttles ERK2 to the nucleus and
MEK2 differentially sorts ERK2 to the cytoplasm (Skarpen et al., 2008). Although these data point out the differential roles for each isoform, it is important to note that MEK1 and MEK2 do not share the same extent of homology as ERK1 and ERK2, especially in their regulatory domains. These data also support the idea of differential activation of each ERK isoform in dictating specific functions.
While a definitive answer about isoform-specific functions cannot be gleaned from previous studies or the work in this dissertation, we propose that the regulation of isoform-specific expression and possibly activation is responsible for the apparent isoform specific functions for ERK1 and ERK2 identified to date. A detailed analysis of
ERK1 and ERK2 mRNA and protein expression throughout development and on a tissue and cell type basis is required for a complete understanding of ERK1/2 activity. An
173 additional line of experiments which would shed some light on the redundancy of the
isoforms would be the generation of knockin mouse lines in which each isoform was
expressed under the other’s promoter and regulatory elements. The phenotype of these
mice would reveal isoform-specific actions and redundant functions.
ERK and Neurogenesis/the cell cycle
Surprisingly, neither the loss of ERK1 or ERK2 alone, nor both ERK1 and ERK2 in the
developing cortex, lead to gross structural defects in the brain (Chapter 2 and 3
respectively). Furthermore, while we could not assess behavior in the double knockout
due to embryonic death, the ERK2 CKO displays behavioral impairments in fear
conditioning but not sensory tests, motor function, reproduction, or longevity. Due to the
relatively minor defects we did identify, two hypotheses can be entertained: (1) ERK1
and ERK2 do not play a significant role in neurogenesis or (2) Other signaling molecules
compensate for the loss of these isoforms.
To address the first possibility, the phenotype of the single and double knockouts must be
considered. The phenotype of the ERK2 CKO and ERK1/2 DKO cortex is similar to that
of other mouse models in which cell cycle components have been mutated or lost. The
knockout of the proto-oncogene n-myc in neural progenitor cells leads to microcephaly
due to severely disrupted proliferation. Like ERK2, n-myc is highly expressed in the
developing CNS. When inactivated in response to recombination with a Nestin-cre
trangene, reductions in neuroepithelial cell proliferation result, but not apoptosis
174 (Knoepfler et al., 2002). This mouse also exhibited high levels of the CKI, p27. A mutant mouse model in which p27 is over-expressed in neural progenitor cells is also phenotypically similar to the ERK2 CKO and ERK1/2 DKO mice. Specifically, significantly thinner cortices were identified which corresponded to fewer neurons populating the cortical lamina (Tarui et al., 2005). ERK2 CKO and ERK1/2 DKO (even more dramatically) mice display proliferation deficits, without apoptosis, and increased levels of p27. These results describe a mitogenic role for the ERKs in neurogenesis and predict that other cell cycle mutants will display similar findings.
Studies of mice in which all 3 D cyclins, or both cyclins E, or cdk4/6 and cdk2 have been knocked out demonstrate that these seemingly important proteins are dispensable for the majority of fetal development (Sherr and Roberts, 2004; Kozar and Sicinski, 2005).
Cyclin D1-/-D2-/-D3-/- mice are phenotypically similar to our ERK1/2 DKO mice as they die around embryonic day 16, but are of normal size and exhibit cardiac malformations characterized by ventricular septal defects and reductions in wall thickness (Kozar et al.,
2004). Furthermore, cdk4/6 double knockout mice display similar defects, owing to the ability of cyclin E-cdk2 and cyclin A-cdk2 to compensate for the loss of cyclin D or cdk4/6 (Malumbres et al., 2004). The role of cyclinD1-cdk4 activity is to phosphorylate and inactivate the retinoblastoma tumor suppressor gene, Rb. This subsequently allows
E2F transcription factors to induce S phase entry. It has been demonstrated in fibroblasts that loss of Rb allows for cell cycle progression in the absence of ERK1/2 activity
(D'Abaco et al., 2002). Individual silencing of each cyclin D isoform leads to specific phenotypic perturbations, where only cyclin D1 knockouts display neurological defects.
175 This finding was characterized as a cyclin D1-specific effect because mice expressing only cyclin D2, displayed the same phenotype (Ciemerych et al., 2002). Importantly,
deletion of p27 restores normal development (Geng et al., 2001) in cyclin D1 knockouts.
Furthermore, knockin of cyclin D2 into the cyclin D1 locus rescues most of the
abnormalities of the cyclin D1 knockout mouse, demonstrating that the tissue-specific
expression of each isoform mainly dictates their functions (Carthon et al., 2005).
Differential expression of the G1 phase cyclins, D1 and D2, in specific neural progenitor
subsets may contribute to the phenotype observed in both cyclin D1 knockouts and in our
mice. Different patterns of cyclin D1 and D2 expression in neurogenic proliferative zones was first identified in 1996 (Ross et al., 1996). A more recent detailed analysis in
the developing mouse demonstrated that cyclin D2 has a much more restricted pattern of
expression than cyclin D1 throughout neural development (Glickstein et al., 2007).
During early cortical neurogenesis, D2 is mainly expressed in cells of the subventricular zone. Subsequently, at E17.5, its expression becomes more widespread. These cells which make up the subventricular zone at E14.5 are mostly intermediate progenitor cells.
Importantly, we found a severe reduction in mitotic intermediate progenitor cells in the
ERK2 CKO mouse. We hypothesize that loss of ERK2 selectively perturbs cyclin D2 function and/or expression affecting mitotic progression.
The phenotype of the ERK2 CKO and ERK1/2 DKO is similar to other cell cycle component mutants. These data support a role for ERK1/2 in cell cycle control of neural progenitor cells and possibly more importantly, intermediate progenitor cells. To assess a
176 role for ERK1 and ERK2 in this latter cell type, ERK2 can be inactivated through lox/cre
mediated recombination under control of the transcription factor, Tbr2. Additionally,
Tbr2 expressing cells can be isolated and ERK1/2 expression and activity can be individually assessed. Selective inactivation of each isoform in the isolated cells can be achieved through transduction of shRNA or RNAi for each isoform and subsequent
analysis of cell cycle parameters and components.
Because fidelity in proliferation is critical for maintenance of cellular populations and
tissue development, fail-safe mechanisms ensure proper control of mitosis. Hence, the
compensation observed at multiple levels in cdk and cyclin knockouts. Therefore, it is likely that ERK1 and ERK2 are redundant isoforms who are expressed to ensure proper
proliferation of neural progenitor cells. This predicts that ERK1/2 are important for
neurogenesis and that additional levels of regulation may exist to ensure proper
proliferation in their combined absence or mutation.
In consideration of the compensation by other signaling molecules, several alternatively
spliced isoforms of ERK1 are generated in development. In rodents, ERK1b is a 46 kD
ERK isoform which is generated by a 26 amino acid insertion in ERK1. ERK1b has been shown to be activated and behave very similarly to ERK1 and ERK2 (Yung et al., 2000) and therefore may be able to compensate for the loss of each. ERK1c is another alternatively spliced variant which is found in primates but has been demonstrated to have unique functions such as golgi fragmentation during mitosis (Aebersold et al., 2004;
Shaul and Seger, 2006). Despite this latter finding, it remains possible that in the absence
177 of the prototypical ERK isoforms, the alternatively spliced forms may compensate in the normal functioning of ERK1 and ERK2.
Another ERK isoform which may be more relevant to issues of redundancy and
compensation is ERK5. Although ERK5 is similar in function to ERK1/2, it is not classified as a classic ERK MAP kinase due to differences in activation, structure and
identity as the Big MAP kinase (Bmk1). While ERK5 is activated by WNK instead of
Raf, its activity is similarly regulated by growth factor signaling and is important in cell
proliferation and differentiation (Nishimoto and Nishida, 2006). ERK5 shares many
downstream targets with ERK1/2, including phosphorylation of RSK, c-myc and
transcription of fos. Therefore, in the absence of ERK2 and/or ERK1 and 2, signaling
crosstalk to ERK5 can add a level of fidelity to cell cycle signaling. It has also been
recognized that ERK5 is also able to drive Cyclin D1 activity (Mulloy et al., 2003). In
this regard, it is important to note that studies have shown that ERK5 is both necessary
and sufficient for neuronal generation in the cortex (Liu et al., 2006). Knockout studies
have shown that ERK5 directs the survival of neural progenitor cells. Therefore it is
possible that ERK5 compensates for the loss of ERK2 and/or ERK1 in our knockout
models. Further analysis of neural progenitor cell survival and proliferation in a mouse
in which ERK5 is conditionally inactivated along with ERK1 and ERK2 would confirm
this proposal.
178 ERK and Cognitive Function/Mental Retardation
The molecular mechanisms underlying normal cognitive function and their dysfunction in mental retardation are unclear; however, it is well established that structure and plasticity of the cerebral cortex are associated with mental retardation. The ERKs have a long standing role in learning and memory. A substantial body of evidence in organisms ranging from Aplysia to humans implies that the Ras-MAPK pathway plays a role in cognitive function (Mazzucchelli and Brambilla, 2000). Furthermore, disruption of numerous elements both upstream and downstream in MAPK signal transduction leads to decreased intellectual ability and mental retardation in individuals (Thomas and Huganir,
2004).
Figure 4-1: Mutations in upstream and downstream members of the MAP Kinase signaling pathway lead to mental retardation and NCFC Syndromes.
179 Mutation of RSK2 leads to Coffin Lowry Syndrome, and is associated with reductions in total brain volume, with hippocampal and cerebellar volumes most
drastically affected, and significant mental retardation (Kesler et al., 2007; Poirier et al.,
2007). Other forms of X-linked mental retardation are also caused by downstream targets of the ERK signaling cascade as mutation of RSK4 causes X-linked Mental Retardation
(Yntema et al., 1999) and MNK1/2 mutations and aberrant ERK signaling has been associated with Fragile X Syndrome (Kim et al., 2008). Mutations in CREB Binding
Protein (CBP) are causative for Rubenstein Taybi Syndrome. Individuals with this disorder have severe mental retardation and characteristic dysmorphology including craniofacial abnormalities (Hallam and Bourtchouladze, 2006; Gervasini et al., 2007;
Roelfsema and Peters, 2007).
Our data extends these findings by demonstrating that the loss of ERK2 alone produces significant deficits in mouse models of learning. It has also recently been demonstrated that ERK2 hypomorphs display deficits in long term memory (Satoh et al., 2007).
These data provide a clear link between perturbations in ERK signaling and cognitive function. However, due to the limitations of our analysis, mechanistic questions remain.
Are the learning and memory deficits seen in ERK2 CKO and hypomorphic animals (as well as the other models of MAPK-related MR such as Coffin Lowry Syndrome and
Rubenstein Taybi Syndrome) developmental in nature (Do they result from improper connectivity or the reduction in neuron number identified in the CKO?), or do they result from impaired synaptic transmission and plasticity in post-mitotic neurons? Cognitive
180 function is thought to be based on both structural connectivity (cortico-thalamic cortico-
cortico, cortico-amygdalar) as well as neuronal activity via synaptic transmission.
Because ERK1 and 2 are involved in both, it is unclear to what the cognitive deficits are attributed.
Evidence from Down’s Syndrome (DS) patients suggests that this form of mental retardation is due to abnormalities in both structural connectivity and neuronal activity.
In DS patients, cortical size is reduced, and neuronal density is increased. Furthermore, small cerebellum, loss of cholinergic neurons of the basal forebrain, and alterations in dendritic spines have been reported (Branchi et al., 2003). These alterations are similarly found in the numerous trisomy 16 mouse models of DS in which reductions in brain weight, small cerebellum and reductions in specific neuronal populations are found
(Branchi et al., 2003). A subset of the models also revealed functional abnormalities in noradrenergic projections to the cortex and hippocampus. Ts65Dn mice, which have a large triplication of chromosome 16, exhibit learning and behavioral defects but normal morphology of the brain. These mice also have abnormal long-term potentiation (LTP) and depression (LTD) in the dentate gyrus and CA1 regions of the hippocampus. Mice with smaller triplications encompassing the Down’s Syndrome critical region, Ts1Cje mice, also exhibit behavioral and cognitive impairment as well as LTP and LTD. These reductions are generally attributed to decreases in dendritic spine density and spine neck length with an accompanying increase in spine head size. Connectivity was likewise abnormal with decreased inhibitory inputs to the dendritic shafts and spines with more inputs to the necks (Belichenko et al., 2007). Within the cortex, it was documented that
181 subplate neurons and neurons populating the cortical plate were generated simultaneously. This disruption in the temporal dynamics of cortical development was correlated with a significant decrease in thalamo-cortical innervation of the subplate in trisomy 16 mice (Cheng et al., 2004).
In our mouse model, we hypothesize that changes in both structural connectivity and synaptic plasticity play a role here as well. Intra-cortical and thalamocortical connectivity has not been assessed, nor has the excitability of ERK2 null neurons been measured. It is of great interest to assess the extent of thalamo-cortical and cortico- cortico connectivity through lipophilic dye labeling. To circumvent the possibility of abnormal connectivity as a cause for cognitive defects, analysis of mice in which ERK1/2 expression and activity is reduced shortly after development (by use of a CamKII-cre mouse line, currently being generated in collaboration with J. David Sweatt) will be illuminating. Additionally, analysis and comparison of LTP and LTD in CKO and DKOs should be performed. The activation of transcription factors (such as CREB, Elk-1 and c- myc) by luciferase reporter assay and the expression of immediate early genes (c-fos, zif268, BDNF), after glutamatergic stimulation can also be assessed to determine the contribution of the ERKs to these downstream targets.
ERK and 22q11 Deletion Syndrome
We have proposed for the first time a role specifically for ERK1/2 in the etiology of
22q11 Deletion Syndrome (DS). To date, numerous elements of the MAPK signaling
182 cascade have been implicated in the etiology of other Neuro-Cardio-Facial-Cutaneous syndromes. Noonan Syndrome occurs from mutations in K-Ras, SHP-2, C-Raf and
SOS1, while LEOPARD Syndrome results from SHP-2, K-Ras and C-Raf mutations.
Costello Syndrome results from H-Ras mutations, and MEK1/2, B-Raf and H-Ras mutations give rise to Cardio-facio-cutaneous Syndrome. This conglomerate of genotypically-related (MAPK) disorders shares phenotypic pathology which is also displayed by 22q11DS patients. These defects include conotruncal, cardiac and craniofacial anomalies, hypo- or aplastic thymus and thyroid, short stature, microcephaly and developmental delay.
22q11 and psychopathology
One of the most consistent features of 22q11 DS is impaired cognitive function. Easily recognizable in children, the cognitive defects displayed include developmental delays and learning disabilities (verbal – language, speech, reading; and nonverbal - motor skills, math, visuo-spatial organization), mild to moderate mental retardation, behavioral disorders (emotional instability, social withdrawal, anxiety disorder and depression) and psychiatric disorders (schizophrenia and bipolar disorder, schizoaffective disorder, schizotypal personality, Attention Deficit and Hyperactivity Disorder, and Obsessive
Compulsive Disorder). 22q11 DS patients usually have an IQ in the range of 70-85, which is one full standard deviation below the general population.
Children and adolescent 22q11 DS patients display social withdrawal, poor social skills, emotional instability, anxiety, ADHD, and autistic behaviors. As mature adults, their
183 behaviors develop into schizophrenia and bipolar spectrum disorders. The incidence of
22q11 DS patients developing one of these schizoaffective disorders as an adult is a staggering 25%-35%. One in 4000 live births results in a child with 22q11 DS. If 25% of these children develop a schizoaffective disorder, then 1 in every 16,000 people will have a 22q11-related psychiatric disorder. Therefore it is important to understand the basis for these disorders in the context of 22q11 Deletion Syndrome.
Intruigingly, the most effective treatment of the bipolar spectrum disorders in 22q11 DS patients is with valproic acid (valproate) (Vogels et al., 2002). This drug has been shown to activate ERK1/2 and induce neurogenesis in mice (Hao et al., 2004).
Psychiatric core traits of 22q11 DS can be assessed in mice. Paired pulse inhibition (PPI)
is a measure of sensorimotor gating, an established schizophrenia endophenotype. 22q11
DS patients exhibit reduced PPI (Sobin et al., 2005). Mouse models of 22q11 DS have also been tested for PPI and exhibit reductions which have been associated with the haploinsufficiency in Tbx1 (Paylor et al., 2006; Paylor and Lindsay, 2006). Analysis of psychiatric behavior profiles by paired pulse inhibition, contextual fear conditioning and working memory testing in ERK1/2 knockouts as well as the B-Raf/C-Raf and MEK1/2 knockouts would point toward a role for these proteins in cognitive function and disease states.
As numerous mouse models of 22q11 DS have already been generated, it is interesting to
assess ERK activity in both Df1/+ mice (a 21 gene hemizygous deletion of the DiGeorge
184 Critical region, including Tbx1 but not CrkL) and in a mouse model which possesses a deletion encompassing both CrkL and Tbx1. A rather intriguing set of experiments would be to feed pregnant females of the Df1/+ or CrkL/Tbx1 deletion mouse models clinically relevant doses of valproate and subsequently assess the pups for physical defects and psychological deficits.
22q11 physical pathology
One of the major physical impairments associated with 22q11 DS are structural defects of
the heart. They manifest as outflow tract defects such as overriding aortic arch, double
outlet right ventricle, Tetralogy of Fallot among others. These have been found to arise
from the deletion of Tbx1, the major causative gene for the disease pathology.
Importantly, Tbx1 is critical for Fgf8/10 expression. These growth factors are required
for the normal development of the pharyngeal arches and cardiac neural crest which give
rise to craniofacial structures, thymus, thyroid and parathyroid, in addition to the heart.
Recently, we (Newbern et al., submitted) have described the phenotype of B-Raf/C-Raf,
MEK1/2 and ERK1/2 single and double conditional knockout mice in which these genes
were targeted for deletion within the neural crest. They each present with a similar
spectrum of heart, craniofacial, and thymus/parathyroid defects. This represents a body
of work which confirms that the ERK signaling pathway is critical for proper neural crest
development, and its disruption recapitulates the physical characteristics of 22q11
Deletion Syndrome.
185 Due to the cognitive, psychiatric and behavioral impairments associated with 22q11 DS patients, structural abnormalities of the CNS have also been investigated. Within the brain, 22q11 DS is associated with overall reductions in brain volume (Deboer et al.,
2007). Neuroanatomical studies have reported multiple perturbations including changes in both gray and white matter in the temporal lobe (including the amygdala and hippocampus) lissencephaly, cerebral atrophy, cerebellar hypoplasia, ventricular enlargement, partial olfactory tract absence, and “simple gyral pattern” (Bird and
Scambler, 2000). Reductions in region specific cortical volumes, reductions in hippocampal volumes, and polymicrogyria are the most common abnormalities identified
(Bird and Scambler, 2000; Swillen et al., 2000; Campbell et al., 2006; Debbane et al.,
2006; Robin et al., 2006; Schaer et al., 2006; Deboer et al., 2007). Specific reductions in gyrification indices identified by MRI were found to be localized to the frontal and parietal lobes. While the prevalence and penetrance of each malformation is variable, it is generally accepted that brain structure and function is perturbed within these patients.
The mouse models described within this dissertation display reductions in thickness of the frontal and parietal lobes. Furthermore, the mouse models described in Newbern et al. demonstrate small or absent cerebella and olfactory bulbs. These data are the most compelling link to the cortical dysgensis observed in patients as cortical and hippocampal volume was not assessed and loss of gyrification or small gyri cannot be assessed in the mouse.
It has been proposed that reductions in cortical volume and polymicrogyria result from a lack of thalamic innervation. As noted above, Fgf8 expression is induced by Tbx1. It is
186 important to note that Fgf8 signaling controls both intracortical and cortico-thalamic
connectivity in the murine brain (Krubitzer and Kahn, 2003). It has generally been accepted that the cognitive disabilities displayed by afflicted individuals are associated with deficits in visuo-spatial processing and attention, both of which are mediated by cortical networks in the parietal and temporal lobes. It is of significant interest therefore to determine if these connections are intact in the ERK1/2 knockouts (as previously noted) as well as the B-Raf/C-Raf and MEK1/2 knockouts. Measurements of cortical and hippocampal volume would also be illuminating.
ERK and Affective Function/ Mood Disorders
Multiple findings lead one to consider ERK signaling as a component of affective
disorders. First, as mentioned above, the incidence of individuals with 22q11 DS and
schizoaffective disorder is extremely high. Second, there is a significant incidence of reduced ERK1/2 activity and expression (protein and mRNA) in the prefrontal cortex
(Brodmans area 8-10) and hippocampus of depressed suicide subjects. An ERK phosphatase, MKP2, was also elevated in these areas of suicide subjects (Dwivedi et al.,
2001; Dwivedi et al., 2006). And third, neurotrophic factor signaling through ERK1/2
has been shown to regulate the pathophysiology of mood disorders.
Given these facts, it is important to consider the role of ERK1/2 in both the development
and treatment of these illnesses. Chen and colleagues have demonstrated that ERK1 null
mice exhibit signs of mania (behaviors associated with antidepressant and
187 psychostimulant induced action) as they exhibit reduced immobility time in tail
suspension tests and increased swim time in forced swim tests (Engel et al., 2008). These
mice also demonstrate enhanced exploratory behavior in an open field as well as
increased home cage activity on a running wheel. When treated with amphetamines,
these actions were elevated even higher. Chronic treatment with clinically relevant does
of lithium or valproate as well as acute treatment with olanzapine attenuated the
amphetamine-induced hyperactivity, suggesting that the ERK1 null mice exhibit
behaviors that are characteristic of manic phases of bipolar disorder (Einat et al., 2003a;
Einat et al., 2003b; Engel et al., 2008). Independent studies have demonstrated that valproate treatment induces ERK activity and neurite outgrowth in vitro and in vivo
(Yuan et al., 2001; Hao et al., 2004; Rosenberg, 2007).
Others have demonstrated that inhibition of MEK with SL327 increases locomotion in a novel environment and reduces immobility in forced swim tests, further demonstrating that alteration in ERK signaling induces these phenotypes. Additionally, these data demonstrate that developmental perturbations may not be involved in these behaviors
(Einat et al., 2003a). Peripheral injection of MEK inhibitors produce similar helplessness phenotypes and eliminate the responsiveness to antidepressant treatments (Duman et al.,
2007).
Importantly, dysregulation of the FGF signaling cascade has been shown to be involved in Major Depressive Disorder. Specifically, a cohort of human subjects with major depressive disorder had significantly reduced expression levels of FGF1, FGF2, FGFR2
188 and FGFR3 in the dorsolateral prefrontal cortex and/or anterior cingulate cortex as
compared to control subjects (Evans et al., 2004). As the FGFs signal through their
cognate receptors to ERK1/2 these data support the notion that a disruption in ERK signaling underlies mood disorders. Depressive disorders can also be modeled in rats to further study mechanisms and drug therapies. In rats, chronic forced swim tests are used to induce depressive behaviors (less time in open arms, more time in closed arms; decreased locomotor activity, increased grooming, reduced weight). In animals which peformed the chronic forced swim task, phosphorylated ERK2 levels were reduced in both hippocampus and prefrontal cortex (Qi et al., 2006). Amitriptyline (antidepressant
treatment) restores ERK2 phosphorylation in the amygdala and hippocampal regions,
ERK1 phosphorylation in the striatum, and importantly, rescues depressive behaviors
after corticosterone-induced depression (Gourley et al., 2008).
Recently, mice which exhibit a polymorphism in DISC1, disrupted in schizophrenia,
demonstrate reduced ERK phosphorylation and reductions in gray matter volume
associated with Major Depressive Disorder (Hashimoto et al., 2006). These findings
suggest that disruptions in ERK1/2 signaling or improper a loss of ERK1/2 activity could
underlie depressive disorders as well as be causative in the manic and depressive
episodes in bipolar disorder. Taken together, these findings demonstrate that not only do
we need to consider the developmental effects of improper ERK activity on mental
retardation, but we should consider ERK2 and ERK1 hypomorphic effects on other
mental illnesses, especially bipolar disorder and major depressive disorder.
189 An interesting and unexplored line of investigation is the role of the ERK MAP kinases
in autism. Interestingly, two independent autism consortiums have found that
microdeletions and duplications on chromosome 16p11.2 are linked to hereditary forms
of autism (Lucarelli et al., 2003; Kumar et al., 2008; Weiss et al., 2008). Of significance
is the fact that mapk3/ERK1 is located within these chromosomal abnormalities.
Autistism spectrum disorders are generally characterized by language delay, repetitive
actions and deficits in socio-communicative behavior, psychological manifestations
identified in 22q11 DS children as well. These children also have phenotypic
craniofacial defects common to neuro-cardio-facio-cutaneous syndrome patients (Ballif et
al., 2007; Ghebranious et al., 2007; Weiss et al., 2008). Moreover, recent reports suggest
that improper and under-connectivity in thalamocortical and intracortical circuits may be
associated with the disorder (Mizuno et al., 2006; Hughes, 2007; Just et al., 2007). It
may be noteworthy that thinning of the corpus collosum is present in both autism and
22q11DS. Given the possibility of under or improper connectivity due to the loss of
ERK2, it is also worth investigating connectivity and synaptic plasticity during development of ERK1 null mice. This is an intriguing line of investigation as autism and
schizophrenia share common social-behavioral characteristics, and they are thought to
potentially share common etiology (Pinkham et al., 2008). The underlying signaling
pathway leading to both disorders may be the same, namely ERK MAP kinase signal
transduction.
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