The Role of Cdk5 and E2F1 in Cell Cycle Regulation of the Post-Mitotic

Home , CDK5R1, Cell cycle, Cyclin B1, Cyclin E2, G2 phase, MYT1

CELL CYCLE REGULATION IN THE POST-MITOTIC NEURONAL CELLS

LI WANG

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Dissertation Adviser: Dr. Karl Herrup

Department of Genetics

CASE WESTERN RESERVE UNIVERSITY

August, 2007

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the dissertation of

______

candidate for the Ph.D. degree *.

(signed)______(chair of the committee)

______

(date) ______

*We also certify that written approval has been obtained for any proprietary material contained therein.

Dedication

To my family, especially to my parents

Table of Contents

Dedication………………………………………………………………………………….i

Table of Contents………………………………………………………………………….ii

List of Figures and Tables………………………………………………………………..iv

Acknowledgements…………………………………………………………………….....vi

Abstract…………………………………………………………………………………..vii

Chapter1: introduction…………………………………………………………………….1

The Normal Cell Cycle Regulation…………………………………………………...2

The overview of E2F family………………………………………………………….7

Background of CDK5……………………………………………………………….27

Figures……………………………………………………………………………….40

Chapter2: E2F1 works as a cell cycle suppressor in mature neurons …………………...43

Abstract……………………………………………………………………………..44

Introduction…………………………………………………………………………45

Materials and Methods……………………………………………………………...47

Results………………………………………………………………………………53

Discussion…………………………………………………………………………..62

Figures………………………………………………………………………………68

Chapter3:CDK5 Nuclear export of CDK5 is required for cell cycle re-entry in neuronal cells stressed by β- amyloid……………………………………………………………...86

Abstract……………………………………………………………………………...87

Introduction………………………………………………………………………….88

Materials and Methods………………………………………………………………92

Results………………………………………………………………………………96

Discussion…………………………………………………………………………107

Figures……………………………………………………………………………..113

Chapter4: Discussion…………………………………………………………………...131

Chapter5: Literature cited………………………………………………………………148

iii

List of Figures and Tables

Chapter1: introduction

Figure 1…………………………………………...... 40

Figure 2…………………………………………...... 41

Table 1………………………………………………………………………..42

Chapter2: E2F1 works as a cell cycle suppressor in mature neurons

Figure 1……………………………………………………………………….69

Figure 2……………………………………………………………………….71

Figure 3……………………………………………………………………….73

Figure 4……………………………………………………………………….75

Figure 5……………………………………………………………………….77

Figure 6……………………………………………………………………….79

Figure 7……………………………………………………………………….81

Figure 8……………………………………………………………………….83

Figure 9……………………………………………………………………….85

Chapter 3: Nuclear export of CDK5 is required for cell cycle re-entry in neuronal cells stressed by β- amyloid

Figure 1……………………………………………………………………...114

Figure 2……………………………………………………………………...116

Figure 3……………………………………………………………………...118

Figure 4……………………………………………………………………...120

Figure 5……………………………………………………………………...122

Figure 6……………………………………………………………………...124

Figure 7……………………………………………………………………...126

Figure 8……………………………………………………………………...128

Figure 9……………………………………………………………………...130

Acknowledgements

I would like to thank my thesis advisor, Dr. Karl Herrup, for his guidance over the years.

As a successful, passionate scientist, he has given me unfaltering support and guidance throughout the years. Not only did I learn science from him, I have also learned from him which is important in life

I would also like to thank the members of my thesis committee, Drs. Gary Landreth,

Bruce Lamb, Robert Miller and Peter Harte. Their support and guidance over the years is greatly appreciated.

Thanks to each member of the Herrup lab and the Landreth lab. They have created a unique and wonderful place for my work. I really appreciate your kind help and have enjoyed the discussions with everyone. A lot of happy and precious moments will be in my memory forever.

Finally, I would like to thank my family and my friends, especially my parents, my sister, my two daughters and my husband. Without your love and support, there is no way I could have done this.

Cell Cycle Regulation in the Post-Mitotic

Neuronal Cells

Abstract

LI WANG

A normal cell cycle is essential for cell proliferation in embryonic development. Once the appropriate numbers of cells are generated, however, there are various populations, e.g. neurons, which will exit permanently from the cell cycle in order to undergo differentiation. It has been widely believed that mature CNS neurons are regarded as permanently post-mitotic and unable to reenter cell cycle. However, emerging data suggests that in different neurodegenerative disease and their mouse models there is a significant deregulation of cell cycle control in specific neuronal populations in precisely those regions where cell is lost.

It seems that the neurons must constantly hold the cell cycle in check after their differentiation. This raises the important but often ignored question: how do adult neurons regulate their cell cycle machinery? Our studies indicate the certain molecules might take on an apparently different role in neurons than they do in other types of cells.

In Chapter 2, I report on the role of E2F1 in cell cycle regulation of adult neurons. E2F1 has been described as an important cell cycle initiator because of its ability to positively

vii

regulate the transcription of genes that are important for normal cell cycle progression.

As part of my thesis work, I have found that, in adult neurons, E2F1 plays a role as a cell cycle suppressor. In Chapter 3, I report on my analysis of a unique cyclin dependent kinase, CDK5, in neuronal cell cycle regulation. Following up on previous work showing that CDK5 is important in suppressing cell cycle progression in post-mitotic neurons, we have further shown that it is the nuclear localization of CDK5 that is important for holding the cell cycle in check in post-mitotic neurons stressed by Aβ.

Overall, our findings indicate that it is crucial for post-mitotic neurons to hold their cell

cycle in check. Both E2F1 and CDK5 apparently play central roles in this process. The

investigation of those signaling pathways in the future has the potential not only to

improve our understanding of the basic biology of neuronal degenerative diseases, but

also offer new pathways to plan for therapeutic interventions.

viii Chapter 1: Introduction

Normal cell cycle regulation

To successfully replicate itself, a eukaryotic cell must complete a proliferative cell cycle.

This progression includes four sequential and tightly regulated phases: G1 phase (Gap1),

S phase (DNA synthesis), G2 (Gap2) phase, and M phase (Mitosis).

During G1 phase, cells receive signals from the extracellular environment and prepare for

proliferation or an alternate fate. Cyclin dependent kinases 4 and 6 (CDK4/CDK6) and

their regulatory partner – cyclin D are expressed in response to certain extracellular

stimuli. At the beginning of this phase, the two proteins, retinoblastoma (Rb) and the

DNA binding protein E2F are associated in a heterodimer. The Rb/E2F complex acts as a

transcription repressor for the genes that are crucial for completion of G1 and the S phase

entry. When CDK4/6 forms an active complex with cyclin D, It phosphorylates Rb. This

hyperphosporylated Rb will release E2F from the Rb/E2F repressor complex. The freed

E2F now can activate the expression of many genes that are crucial for the transition

from G1 to S phase such as cyclin E and cyclin A. The expression of cyclin E activates

CDK2 to effect further phosphorylation of pRb, thereby enabling the cells to cross the G1

restriction point. During S phase, cyclin E is replaced by cyclin A, which binds to and

activates CDK2. The CDK2:cyclin A complex controls the progression through S phase

by phosphorylation of a series of proteins that are necessary for DNA synthesis such as

DP-1 subunits (inhibitors of DNA binding) and CDC6 (an initiator of DNA replication).

After DNA replication is completed, cells enter the second Gap phase (G2) phase and prepare for the mitosis. CDK2:cyclin A is still important for the G2/M transition. When cells enter M phase, cyclin A is degraded and the level of cyclin B increases. Cyclin B

binds and activates CDK1. CDK1:cyclin B controls many genes that are required for M phase progression and exit. At the end of M phase, one cell separates into two daughter cells and replication is complete (Figure 1).

In addition to positive regulators of the cell cycle such as CDKs, there are also some negative regulators such as cyclin dependent kinase inhibitors (CKIs) and cell cycle checkpoint proteins. They control the cell cycle progression by specifically inhibiting the

CDKs at specific phase and allow cell cycle progress sequentially and rightly go through

G1 to M phase.

Initiation a cell cycle does not always mean a successful replication. The cells will stop at specific check points and repair DNA damage or other problems if they encounter them.

If the damage cannot be repaired, cells will die through activation of apoptosis.

The normal neuronal generation

Before the formation of nervous system, three main cell layers have been generated in embryogenesis – the endoderm (the innermost layer), the mesodum (the middle layer) and the ectoderm (the outermost layer). Different layer will give rise to different organs.

Along the development, one special sheet of ectodermal layer begins to acquire neuronal properties and this special sheet is called neural plate. Neuronal and glia cells are generated from precursor cells in the neural plate. Soon after neural plate has formed, it begins to fold into a tubular structure called the neural tube. The caudal region of the neural tube will give rise to spinal cord and the rostral region becomes the brain.

During the early stage of neural development, cells in neural plate expand very quickly

and predominantly in the ventricular zone, a dense layer of cell which lines the surface of

the ventricles. In early divisions the fate of all cells is same; they behave as stem cells for

formation the nervous system. As the development progress, a separate population of

progenitor cells accumulates above the ventricular surface and forms the subventriculare

zone. The progenitor cells in this layer will also generate committed cells.

When ventricular and subventricular cells divide, their replication can be either

symmetric or asymmetric. If symmetric replication, two daughter cells are produced. If

asymmetric division, then one daughter cell is committed to differentiation and it stops to

divide. Once they exit from the cell cycle, the neurons in their life never go back to the replication progress for the rest of the life of the organism. How do the progenitor cells

exit the cell cycle before they differentiate to neurons? How do the neurons hold the cell

cycle in check for the life long time? The answers to these questions are still a mystery.

After neurons are born, they leave the ventricular zone and migrate to their final position,

often along the processes of radial glia cells, and mature. In cerebral cortex, the neurons

migrate in an inside-out sequence to form the cortex: the later born neurons will migrate

through the existing cells of the cortical plate before they stop and form at the specific

layer of the cortex. In this way, multiple signals can influence what kind of neuron each

progenitor will become. After their fate is determined, neurons will grow axons to form

specific connections with their target - either related neurons or other effecter organs.

Ectopic cell cycle events and neuronal death

For neuronal cells, it was believed that cell cycle events only happen in the prenatal

development stage. At that time, the replication of stem cells and progenitor cells is

required for the nervous system formation. Recently, the discovery of the stem cell pool in the adult mammalian brain has provided evidence of small numbers of cell cycle events in the adult nervous system (Gage, 2002). However, the cell cycle events involve only previously undifferentiated stem cells, not mature neurons. For mature neurons, it has been thought that they are post mitotic and permanently exited from cell cycle after they differentiate. This would imply that, in adult brain, only non-neuronal cells

proliferate in response to CNS damage and the neuronal cells can not replenish lost cells.

If forcing a neuron into a cell cycle process, what will happen? In past years, more and

more evidence indicates that neurons can go back to the cell cycle. However, this event is

generally related with cell death, not cell proliferation. The over-expression of an

oncogene can force a quiescent cell into a cell cycle. SV40 T-antigen is such an oncogene.

It can induce an unscheduled cell cycle by blocking the function of Rb family members.

In SV40 T-antigen expressing postnatal Purkinje cells, Bromodeoxyuridine (BrdU) is

incorporated which indicates that those cells have undergone DNA replication and re-

entered an abnormal cell cycle. The BrdU-labeled Purkinje cells die instead of dividing

(Feddersen et al., 1992; Feddersen et al., 1995). The induction of DNA synthesis and the

death of neuronal cells are also found in the T-antigen expressing post-mitotic retinal

photoreceptor neurons (al-Ubaidi et al., 1992a; Al-Ubaidi et al., 1997). Consistent with

T-antigen transgenic mice, mice deficient in the Rb gene have ectopic cell cycle entry

and mitotic figures in several populations of early differentiated neurons, and the fate of

those neurons is death (Clarke et al., 1992; Jacks et al., 1992; Lee et al., 1992). Taken

together, these mouse models are the first indications that there might be correlation

between ectopic cell cycle events and cell death in multiple types of differentiated

neurons.

In addition to forced cell cycle re-entry, three examples of situations in which cerebellar

granule neurons (CGN) are lost to a natural process of target-related cell death reveal that

these more physiological deaths also proceed through a cell cycle related pathway during

development. In staggerer mice, the Purkinje cells never fully develop because of the

deletion of the orphan nuclear receptor (RORα) gene. This leads to the loss of 100% of the cerebellar granule cells because of a deficiency of trophic support. In those granule cells, there is DNA synthesis in reported and expression of cell cycle protein markers – cyclin D and PCNA – appear before they die. In another mutant mouse, Lurcher, there is

a similar phenotype. Purkinje cells die due to a point mutation in the gene encoding an

ionotropic glutamate receptor (GRID2) (Zuo et al., 1997). This results in the death of

most CGNs, which incorporate BrdU and express cell cycle proteins before they die

(Herrup and Busser, 1995). Recently, in mice with targeted deletion of cyclin-dependent

kinase inhibitor (CKI) Ink4d, hair cells are observed to aberrantly re-enter a cell cycle and

subsequently undergo apoptosis, resulting in progressive hearing loss (Chen et al., 2003).

Together, the studies strongly suggest that post-mitotic neurons are vulnerable to the cell cycle related death (CRND).

In vitro, a similar cell cycle re-entry has been shown in different type of neuronal cultures in response to different treatments. Cultured cerebellar granule neurons incorporate

BrdU and/or express cell cycle related proteins prior to death induced by different stimuli such as β-amyloid or K(+) deprivation (Giovanni et al., 1999; O'Hare et al., 2000).

Cultured cortical neurons re-enter a lethal cell cycle following DNA damage and Aβ- stimulated neuronal death (Copani et al., 1999; Kim et al., 2001; Kruman et al., 2004). In addition to these studies, several groups have shown that neuronal death are effectively blocked or delayed by blocking the ectopic cell cycle events through application of several different cell cycle blocking methods (Park et al., 1996; Park et al., 1997a;

Appert-Collin et al., 2006). These in vitro results provide direct evidence that ectopic cell cycle events are caused in all of these different examples of the neuronal death.

How and why the post-mitotic neurons re-enter a cell cycle and the mechanisms of this cell cycle related neuronal death are unclear. From the evidence cited above, it seems like that normal cell cycle machinery such as Rb/E2Fs and CDKs play some role in this pathway. To approach a molecular understanding of the ectopic cell cycle re-entry in post-mitotic neurons, we selected two proteins - E2F1 and CDK5 - to explore their role in this pathway.

Overview of the E2F Family

The members of E2F family

The E2F family is a group of transcription factors that is recognized for playing a pivotal role in cell proliferation. E2F was originally discovered as the cellular mediator of the effects of the adenovirus E1A oncogene (Arroyo et al., 1993). Since the first member of

E2F family, E2F1, has been identified, the mammalian E2F family has become larger by the addition new members identified from the last two decades. Now there are eight structurally related E2F members (E2F1-E2F8). Each member has distinct regulatory functions, transcriptional activities and biological activities. E2F1-E2F6 must heterodimerize with DP proteins (DP1 or DP2) to give rise to functional E2F activity

(Girling et al., 1993). Subsequent studies have shown that E2F controls the transcription of cellular genes that are essential for cell division. More recently, that clears that the spectrum of E2F-mediated transcriptional control may extend well-beyond the set of genes related to the cell cycle.

On the basis of their homology, the proteins of the E2F family can be divided into three distinct subgroups: group1 (E2F1-E2F3a); group 2 (E2F3b, E2F4, E2F5) and group 3

(E2F6, E2F7 and E2F8) (Fig 1).

Among the group 1 proteins, E2F1-3 are highly homologous to each other; they have homologous domains that are responsible for DNA binding, DP dimerization and Rb binding. E2Fs in this group are primarily activated in dividing cells. They are exclusively associated with and specifically regulated by Rb protein, but not regulated by other

pocket proteins (e.g.p107 and p103). In addition to structural similarity, they also show similar transactivation function. They can transcriptionally activate many of the genes that are essential for S phase entry. These include cyclin E, cyclin A and more. Triple knock-outs of E2F1, E2F2 and E2F3 can completely block cellular proliferation (Wu et al., 2001). By contrast single knock-outs have minor phenotypes. This indicates that these activators are likely to have overlapping roles in triggering cell cycle entry. In addition to a function in cell division, deregulation of proteins in this subgroup, especially E2F1, is found in the apoptosis pathway (DeGregori et al., 1995; Hsieh et al., 2002). While E2Fs in this subgroup are thought act as transcriptional activators, however, additional studies indicate that they may also act as transcriptional repressors in specific contexts (Field et al., 1996; Ishida et al., 2001; Muller et al., 2001; Vernell et al., 2003; Young et al., 2003).

The proteins in this subgroup have a nuclear localization signal (NLS). Despite the presence of a NLS in E2F1; other factors are still required to transport E2F1 into the nucleus.

The members of the E2F second subgroup (E2F3b, E2F4 and E2F5) are quite different from those in group 1: they lack most of the sequence that is amino-terminal to the DNA- binding domain. E2F3b is a new addition of this group. It is transcribed from a previously unrecognized promoter in the first intron of E2F3a (He et al., 2000; Leone et al., 2000).

Its function has been suggested to include transcriptional repression of the expression of p19Arf (Aslanian et al., 2004). E2F4 and E2F5 are older members of this subgroup. They are mostly expressed in quiescent cells. E2F4 can interact with all three pocket proteins

(Rb,p107 and p103) while E2F5 mainly interacts with p107 and p103 (Dyson, 1998;

Nevins, 1998). The best characterized function of this subgroup of E2Fs is to induce cell cycle exit and terminal differentiation.

The third subgroup of the E2F family is made up of E2F6, E2F7 and E2F8. E2Fs in this

group are different from the previously described E2Fs. They lack the transcriptional activation and pocket protein-binding domains. This structure abuse them to act as repressors of E2F responsive genes, but by a mechanism that does not involve interaction with pocket proteins (see reviews (Dimova and Dyson, 2005; Fang and Han, 2006).

The DNA-binding and dimerization domains of other E2Fs are conserved in E2F6. E2F6

has to dimerize with DP protein to act as a repressor. It is expressed in all stages of cell

cycle. It has been suggested that it plays a role in regulation of E2F target genes in G0

(Ogawa et al., 2002). However, the E2F6 null MEFs show no cell cycle defects (Storre et

al., 2002). This indicates that E2F6 might have overlapping roles with other repressive

E2Fs.

E2F7 and E2F8 are new members of this subgroup. They contain two DNA-binding

domains and lack the DP dimerization domain. So they bind to DNA in a DP-

independent manner, possibly as a homodimer. They are expressed in a cell cycle

dependent manner and peak level found during S phase. Both are expressed in the same

adult tissue of mice. Ectopic expression of them repressed the E2F target genes and block

the cell proliferation (de Bruin et al., 2003a; Di Stefano et al., 2003; Logan et al., 2004;

Christensen et al., 2005; Logan et al., 2005; Maiti et al., 2005). Those data suggest that

E2F7 and E2F8 may repress some specific E2F target genes during the cell cycle when other E2Fs de-repressed from their regulatory proteins. The regulation and target genes of those two E2F members remain to be determined.

The regulation of E2F activity

The regulation of E2F activity by Pocket family

Regulation of E2F activity is achieved through multiple mechanisms. The first is interaction with the pocket proteins --- pRb, p107 and p130. As mentioned above these proteins function as negative regulators of E2Fs. Different pocket proteins interact with different E2F members and regulate different cellular programs.

The first subgroup of E2Fs (E2F1-E2F3) is specifically regulated by pRb protein, but not by other pocket proteins (p107 and p103). E2Fs in group 1 are freed from hyperphosphorylated pRb and activate or derepress multiple E2F-responsive genes.

Because of this relationship, inactivaton of Rb has a phenotype similar to the overexpression of E2F --- Rb-null mice have excessive apoptosis of cells in the nervous systems, lens and skeletal muscles (Huang et al., 1992). The apoptosis phenotype of Rb null embryos is correlated with ectopic S-phase entry in Rb-null neurons and muscles. As would be predicted double mutants of both Rb and E2F1 or E2F3 have reduced ectopic

S-phase entry and apoptosis of the Rb null neurons in CNS and epithelial cells in the lens

(Tsai et al., 1998) (Wu et al., 2001).

E2F4 and E2F5 are mainly regulated by p107 and p130. The complexes containing E2F4

(or E2F5) together with p107 or p130 are found in all tested conventional E2F target

genes in quiescent cells. They function as a transcriptional repressor for those target genes. P107-/-; p130-/- MEFs show accelerated entry into S phase and deregulation of a

subset of E2F target genes (Dyson, 1998). Another level of p107 and p130 regulates the

transcriptional activity of E2F4/5 is by controlling their cellular localization. E2F4 and

E2F5 are translocated from the nucleus to the cytoplasm during G0 to S phase progress.

In contrast to the activating E2Fs, however, E2F4 and E2F5 do not have a nuclear

localization signal (NLS). So interaction of these E2Fs with p107 and p103 has been

proposed to be required of their nuclear localization (Lindeman et al., 1997; Verona et al.,

1997). Traslocation of p107/E2F repressor complexes to the nucleus results in

transcriptional repression of E2F responsive genes by recruitment of histone deacetylase

(HDAC) and changes the chromatin structure (Altiok et al., 1997; Slomiany et al., 2000;

Porse et al., 2001).

Studies show that there are several ways in which pocket proteins such as pRb regulate of

E2F activity. First, pRb can directly bind to the activation domain of the activator E2fs

(E2F1-E2F3). So it directly blocks the activity of this domain (Flemington et al., 1993;

Helin et al., 1993). Second, the recruitment of pRb to a promoter can block other

transcription activators to bind to the promoters and thus blocks the formation of

transaction complexes (Ross et al., 1999). Third, after it binds to E2Fs, pRb can recruit

additional proteins to form complexes that can modify chromatin structure. It can also

associate with chromatin modifying enzymes such as histone deacetylase (HDAC) to

change the structure of E2Fs target promoters (Brehm et al., 1998; Luo et al., 1998; Ross

et al., 2001). Similar regulation mechanisms are found for p107 and p130.

Members of the third subgroup of E2Fs (E2F6-8) lack the pocket protein binding domain,

and as a consequence they are not regulated by Pocket proteins. It has been shown that

E2F6 represses its target genes through binding to Polycomb group (PcG) proteins or through the formation of a large transcriptional repression complex (Trimarchi et al.,

2001; Ogawa et al., 2002). The association of E2F6 with PcG proteins suggests the

possibility that E2F6 might regulate a different population of target genes. PcG proteins are important for the regulation of Hox gene expression (Gould, 1997; Bantignies and

Cavalli, 2006). In fact, the posterior homeotic transformation of the axial skeleton in

E2F6-/- mice is similar to the phenotype observed in mice deficient in several different

PcG proteins (Storre et al., 2002). The regulation and target genes of E2F6 are remain to

be determined.

Transcriptional regulation and post-transcriptiondal modifications of E2Fs

Transcriptional regulation

In addition to regulation of E2Fs by pocket proteins, the levels of E2Fs are also regulated

at the transcriptional and post-transciptional levels.

The levels of E2Fs are transcriptionally regulated by their respective promoters (Neuman et al., 1994). The mouse E2F1 promoter contains an E2F-reposive sit (Hsiao et al., 1994).

Thus the transcription of activating E2Fs is regulated by a E2F-dependent self-inhibitory

loop (Leone et al., 1997; Sears et al., 1997; Adams et al., 2000; Leone et al., 2001; Araki et al., 2003). In activator E2F promoter regions, there are Myc binding sites. These are believed to enhance the transcription of the adjacent genes at critical points in the cell cycle (Sear R, 1997; Leone G, 1997; Leone G, 2001).

Post-transcriptional regulation —phosphorylation and acetylation

Phosphorylation

E2Fs can be modified by both phosphorylation and acetylation, both of which will affect

E2F trancriptional activity. Among them, the post-transcriptional regulation of E2F1 has

been well studied.

E2F1 can be phosphorylated on serine-375 by cyclin A:CDK2. Phosphorylation of E2F1

on S375 greatly increases its binding affinity for pRb (Peeper et al., 1995), thus

potentially limiting the activity of E2F1 during the cell cycle progression and facilitating

a smooth transition to mitosis. Phosorylation at serine 403 and threonine 433 of the

transactivity domain of E2F1 triggers E2F1 degeradation during S phase (Vandel and

Kouzarides, 1999). On the other hand, phosphorylation of E2F1 by ATM/ATR increases

the levels of E2F1 by inhibiting its degradation after the DNA damage (Lin et al., 2001).

This stabilization of E2F1 might work through the phophoserine protein-14-3-3 tau

(Wang et al., 2004).

Acetylation

After they are freed from their pocket proteins, E2F1-6 will bind to DP protein to form

heterodimer. This association will help E2Fs bind to the promoters of their target genes.

Besides association with DP proteins, acetylation of E2F1-3 on three lysine residues

outside its DNA-binding domain by P/CAF and p300/CBP increases its DNA-binding

activity (Marzio et al., 2000). Mutation of the acetylation sites in E2F1 reduces its

transcriptional activity, and there is additional specificity implied because the binding

ability of the mutant protein is reduced for some promoters (Martinez-Balbas et al., 2000;

Pediconi et al., 2003). This raises the possibility that acetylation of E2F1 might be important for its recruitment to specific promoters. pRb-associated HDAC activity is sufficient to deacetylate E2F1 (Marzio et al., 2000). This is one mechanism by which pRb represses the transcriptional activity of E2F1. No acetylation of other E2Fs has been found. Thus acetylation might be a specific regulation method for “activator” E2Fs. In fact, E2Fs itself is not the only element that is regulated by acetylation. The target promoters of those E2Fs are also regulated by acetylation and deacetylation.

Degradation of E2F

Only free E2Fs are degraded by the ubiquitin system. The binding of E2Fs with pocket

proteins could stabilize them and increase their half-life while at the same time blocking their synthesis. This will form a negative feed-back loop. The best study of degradation pathway of E2F is also that of E2F1. It depends upon the ubiquitination by multi-part ubiquitin ligases. The F-box protein p46skp2, the E3 component of the ubiquitin conjugating complex , selectively binds to E2F1 (Campanero and Flemington, 1997).

E2F1 and E2F3 can be degraded in the nuclear proteosome via interaction with p19ARF

(Martelli et al., 2001). Recent studies has shown a possible role for ROC-culin ligases in the degradation of E2F1 process (Ohta and Xiong, 2001). The importance of this pathway, however, remains to be investigated.

The Function of E2Fs

The role of E2fs in S-phase entry

The role of E2F family in cell proliferation has been well documented. pRb/E2F proteins control the expression of many genes required for S-phase entry including genes controlling cell cycle regulation, DNA synthesis and DNA replication.

In G0 and early G1 phase of the cell cycle, the promoters of cell cycle genes are mainly regulated by repressor E2Fs: E2F4 and E2F5, which are bound preferentially to p130, exert an inhibitory effect on those E2F target genes. The repressive complexes formed by

E2F4 or E2F5 with p130 are disrupted in the late G1 phase. Consistent with this model,

E2F4 and E2F5 are important for cell cycle exit and differentiation, and cells lacking repressor E2Fs fail to respond to certain cell cycle arrest signals (Gaubatz et al., 2000).

For S-phase entry, the activator E2Fs (E2F1-E2F3a) are central player. In late G1, pRb and p130 are phosphorylated by cyclin D:CDK4, 6 and cyclin E:CDK2, thus, releasing activator E2Fs. At the same time, p130 is degraded through a ubiquitin-mediated pathway

(Tedesco et al., 2002; Bhattacharya et al., 2003). The repressor, E2F4 (and possibly E2F5) is removed from the nucleus (Allen et al., 1997; Apostolova et al., 2002). Activator E2Fs turn on the genes that are required for S phase entry and cell cycle initiation such as

cyclin E and cyclin A. Consistent with this, overexpression of activator E2Fs can drive

quiescent cells into S-Phase (Johnson et al., 1993; Lukas et al., 1996). Further studies

have found that CDC25A phosphatase and cyclin E are important for this deregulated S

phase re-entry (Vigo et al., 1999). Moreover, the over-expression of E2F1, E2F2, and

E2F3a can induce transformation of primary cells (Johnson et al., 1994; Singh et al.,

1994; Xu et al., 1995). In some situations, expression activating E2Fs can override

various growth-arrest signals , including the CDK inhibitors and TGF-b (DeGregori et al.,

1995; Schwarz et al., 1995; Mann and Jones, 1996). On the other hand, blocking

endogenous E2F3 either by antibody or by removal of the E2F3 gene, affects cell cycle

progression in primary cells (Leone et al., 1998; Humbert et al., 2000a). Finally, the

combined mutation of E2F1, E2F2 and E2F3 is sufficient to completely block cellular

proliferation (Wu et al., 2001). This also suggests that those activators might have

overlapping roles in the induction of cell cycle entry. The role of those activators in S- phase entry is conserved in Drosophila (Frolov et al., 2001; Dimova et al., 2003) and

Xenopus (Tanaka et al., 2003).

Clearly, the activator E2Fs has very important roles for S-phase entry of cells. Whether the cell will actually complete the cell cycle and divide or die depends on the cellular context, because E2Fs do not always promote cell division.

The Role of E2Fs in apoptosis

In addition to inducing proliferation, de-regulated E2Fs can induce apoptosis when over- expressed or when activated in response to DNA damage. Studies show that only

activator E2Fs have roles in inducing apoptosis. For example, ectopic expression of E2F1

leads to apoptosis in cultured cells (Qin et al., 1994; Shan and Lee, 1994; Wu and Levine,

1994; Kowalik et al., 1995; DeGregori et al., 1997) and transgenic mice (Guy et al., 1996;

Holmberg et al., 1998). In E2F1 deficient mice, there is an excess of mature T cells due to a defect in thymocyte apoptosis. Ectopic expression of E2F2 and E2F3 can induce

apoptosis in some experimental systems (Vigo et al., 1999) , but not in others

(DeGregori et al., 1997; Kowalik et al., 1998). This indicates that their action might be

context dependent for promoting apoptosis.

Both cell culture experiments and mouse models, however, indicate that E2Fs induced

apoptosis through several different pathways. First, E2F can induce apoptosis in a p53-

dependent manner. E2F can stabilize and activate p53 through direct transactivation of the p14ARF tumor suppressor gene (Bates et al., 1998; Dimri et al., 2000; Elliott et al.,

2001; Aslanian et al., 2004; Harris and Levine, 2005). However, E2F can also induce p53-dependent apoptosis in the absence of ARF, which indicates the existence of alternative pathways for E2F1-dependent p53 induction. E2F1 can trigger the phosphorylation of p53 on a site that is also phosphorylated in response to DNA damage

(Rogoff et al., 2002; Russell et al., 2002). After DNA damage , the level of E2F1 is increased by phosphorylation of ATM/ATR; blocking the activity of the ATM/ATR by caffeine blocks both the E2F1-dependent induction of apoptosis and p53 phosphorylation (Rogoff et al., 2002). This suggests that the modification of E2F1 by

ATM might alter the list of E2F target genes. It is also interesting to note that E2f1 can

physically interact with p53 through its cyclin A binding domain and enhance the

apoptotic activity of p53. This function of E2F1 is independent of its DNA binding or

transcriptional activity and it is shared by E2F2 and E2F3 (Hsieh et al., 2002). So E2Fs

can affect p53-dependent apoptosis through several different pathways.

The apoptotic activity of E2F1 can also proceed through p53-independent pathways. The

p53 family member, p73, has been shown to play a role in E2F-induced death (Irwin et

al., 2000; Lissy et al., 2000; Stiewe and Putzer, 2000). E2F1 also can directly activate the

expression of the Apaf-1 gene, which activates procaspase-9, leading to the activation of

downstream effector- caspases (Moroni et al., 2001; Furukawa et al., 2002). DNA

microarray studies show that ectopic expression of E2Fs up-regulates pro-apoptotic

members of Bcl-1 and caspase families (Muller et al., 2001; Polager et al., 2002; Stanelle

et al., 2002).

Deregulated E2Fs can also trigger apoptosis through the inhibition of anti-apoptotic

signals. This occurs through the disruption of NF-kB signaling by down-regulating

TRAFa protein levels and inhibiting NF-kB DNA binding ability (Phillips et al., 1999;

Tanaka et al., 2002).

Overall, the induction of apoptosis by deregulated E2Fs is complicated and context

dependent. Of the pathways that have been studied, some directly depend on E2F

transcriptional activation; some just depend on interaction with other proteins involved in

the apoptotic pathway. Whether other E2Fs involved in the apoptosis pathway remains to

be investigated. Importantly, those E2Fs are also involved in the regulation of cell

proliferation. This means it is important understand the mechanisms that will determine

the final outcome of E2F activity-cell survival or cell death.

E2Fs in development and differentiation

The role of pRbs/E2Fs in the development process has become known through the analysis the mutant animals and their specific developmental defects. Rb -/- mice die between embryonic days 14 and 15 with defects in differentiation of multiple tissue.

Some of the tissue specific defects observed in Rb -/- mice are due to the defects in extraembryonic development. E2F1 knockout mice have defects in T-cell apoptosis and testicular atrophy and, surprisingly, develop a broad spectrum of tumors (Field et al.,

1996; Yamasaki et al., 1996). E2f2-/- mice have increased proliferation of hematopoietic cells and also have problems of tumorigenesis (Lin et al., 2002; Martinez et al., 2002).

Some E2f3-/- mice die at embryonic state while other die in adulthood because of heart defects (Humbert et al., 2000a; Rempel et al., 2000). The disruption of E2F4 results in defects in hematopoietic, craniofacial and intestinal tissue (Humbert et al., 2000a;

Rempel et al., 2000). E2F5 mutant mice develop hydrocephalus and are perinatal lethal

(Lindeman et al., 1998). E2f6-/- mice display overt homeotic transformations of the axial

skeleton (Storre et al., 2002). In summary, different E2Fs play important role in different

aspects of tissue development and differentiation. The importance of E2Fs in

development is conserved in flies, worms and frogs.

The role of E2Fs in cell differentiation is also context dependent. In vitro studies show that inhibition of E2F1 is necessary for cell differentiation in several different cell types

(Paramio et al., 2000; Porse et al., 2001; Scheijen et al., 2003; Wong et al., 2003). Mice lacking functional E2F4, E2F5, p107, p130 and Rb all have defective differentiation of various cell lineages (Cobrinik et al., 1996; Lindeman et al., 1998; Humbert et al., 2000b;

Rempel et al., 2000). Overexpression of E2F4 is sufficient to trigger the differentiation of neuronal precursors (Persengiev et al., 1999), but loss of E2F4 increases adipogenesis

(Fajas et al., 2002). In conclusion it would appear that different cells need different E2Fs for their final differentiation, and aberrant expression of different E2Fs might have different effects on differentiation.

The mechanism by which pRb/E2F complexes affect differentiation is not clear. One proposal might be that those repressive complexes simply block the transcription of cell cycle genes and thus facilitate cell cycle exit. Logically, this would prepare the intracellular environment for cell differentiation. Second the pRb/E2Fcomplex might permanently arrest the cell cycle by recruitment of chromatin modification enzyme or by repressive E2F6 - E2F8 occupying cell cycle gene promoters. Third, microarray data suggest that the complex can directly induce the expression of genes which are necessary for differentiation (Muller et al., 2001).

In summary, E2Fs/Rbs complex play important roles in mammal development and differentiation. The detailed mechanism, by which this is accomplished, however, remains unclear. It is thus important to identify the role of individual E2F in specific tissue development.

E2F in DNA damage response and checkpoint control

A role for E2Fs in DNA damage response pathway is suggested by studies showing that the level of E2F1 is increased in cells treated with DNA-damaging agents (Blattner et al.,

1999; Hofferer et al., 1999; O'Connor and Lu, 2000). So far, this defect has only been shown for E2F1 not for other members of the E2F family until now. The signaling pathways that regulate E2F1 stability and connect E2F1 induction with apoptosis in response to DNA damage are becoming clear from recent studies. E2F1 is phophorylated and stabilized by ataxia-telangiectasia mutated (ATM) and ataxia-telangiectasia and

RAD3-related (ATR) checkpoint kinases. It is also phosphorylated by Checkpoint-2

(Chk2) protein kinase. The acetylation of E2F1 has also been connected with phosphorylation by ATM/ATR (Pediconi et al., 2003). So the increase in E2F1 level might be the result of a phosphorylation-acetylation cascade that prevents the normal turnover of E2F1. As the status of the acetylation of E2F1 also affects its transcriptional activity and specifity, the suggestion is that E2F1 might turn on different target genes after DNA damage than it does in a normal cell cycle.

After a “cycling” cell detects DNA damage, it will trigger a cell cycle checkpoint, stopping cell cycle and repairing this DNA damage. If the repair fails, it will undergo programmed cell death. What is the role of E2F1 in these pathways? First, E2F1 might play a role in the progress of DNA repair. Many checkpoint and DNA repair genes are regulated by E2F1, including ATM, BRCA-1, Msh2 andMsh6, DNA replication factor C and PCNA. E2F1 also regulates some genes that function in non-stressed cells to suppress genomic rearrangements during normal cell cycle. The increased level and

specific modifications of E2F1 during DNA damage probably help guide a specific

cellular response to DNA damage. E2F1 localizes to distinct nuclear bodies with intense staining after it is phophorylated by Chk2. Once located in these nuclear foci, it associates with the BRCA-1-repair complex and TobBP1 (the DNA topoisomeraseIIb binding protein1). E2F1 also associates with MRN (Mre11- Rad50- Nbs1) checkpoint protein complex through the N-terminal region of Nbs1. Induced Nbs1/E2F1 interaction occurs close to the site of DNA damage. This suggests that E2F1 is required to target the

MRN complex to origins of replication (Maser et al., 2001). Together, these findings suggest that E2F1 has a role in the detection of DNA damage and its subsequent repair.

DNA damage-induced E2F1 activity may also play a role in apoptosis. Thymocytes derived from E2F1 mutant mice, like those from ATM deficient animals, show defects in

DNA-damage induced apoptosis (Lin et al., 2001). E2F1 can specifically induce the expression of p73 - an apoptosis-inducing gene after DNA damage. The response is specifically related to an increased level of E2F1 acetylation and phosphorylation at S31 by ATM/ATR. It is possible that the high levels of E2F1 that are induced after DNA damage might turn on a different spectrum of genes that are needed for in DNA-damage induced apoptosis. Taken together, E2F1 is involved in multiple aspects in the DNA damage response.

The genes regulated by E2F family

The E2F family was initially recognized for its pivotal role in cell cycle progression.

More and more evidence suggests, however, that the E2Fs regulate genes involved in multiple biological processes. How many genes are regulated by E2Fs? What processes

are these genes involved in? Through the use of microarray technology or computer-

assisted identification of consensus E2F recognition promoter genes, researchers have

identified many additional transcriptional targets of E2F. These targets are involved in

mitosis, chromosome segregation, mitotic spindle checkpoints, DNA repair, chromatin

assembly /condensation, apoptosis, differentiation, and development, etc.(Muller et al.,

2001) (Table1). These studies confirm the role of E2F in the regulation of genes involved

in the G1/S transition, S phase and DNA replication. For example, cyclin E is involved in

the G1/S phase transition; MCMs and CDC6 are involved in the assembly of the pre-

replication complex at origins of replication; other enzymes are directly needed in DNA

systhesis such as the DNA polymerase complex. Beyond this, some genes involved in the

progression through M-phase are also the targets of E2F.These include CDC2 and cyclin

B1. These data suggest the important role of E2F family in cell cycle events. On the other

hand, several key regulators of apoptosis are also in the spectrum of E2F controlled genes.

These include Apaf-1, caspase-3 and caspase-7. E2Fs also down regulate the expression

of anti-apoptosis factors such as Bcl-3, a known co-activator of NFκB. Most of these

apoptosis genes are induced by E2F1. The discovery of these pro-apoptotic genes regulated by E2F1 strongly support the apoptotic effects of E2F activity.

The significance of E2F1 in the nervous system

In the nervous system, cell proliferation and apoptosis are a normal part of development.

These two opposing processes are also indispensable in most tissues for normal maturation and homeostasis. Once neurons are fully differentiated, they will never go back into a cell cycle. In many neurodegenerative diseases, neuron loss is a key feature.

Although the mechanisms of this death include an evolutionarily conserved core

apoptotic pathway, it appears that many instances of neuron death also need the

transcription-dependent induction of proapoptotic molecules. Curiously, in post-mitotic

neurons many of these molecules are involved in a typical cell cycle. For example, in

human AD brain tissue and mouse AD models, cyclins are over-expressed in the neurons

at risk for death (Nagy et al., 1997b; Busser et al., 1998; Yang et al., 2001; Yang and

Herrup, 2005; Yang et al., 2006). As discussed above, transcription factor E2Fs can

regulate expression of various genes that are involved in both the cell cycle and cell death pathways. This indicates that they may be involved in these two opposing processes in

one cell. The outcome of this specific function will determine the fate of the cell. Due to

the closely relationship between cell cycle re-entry and cell death in neurons, many

studies have been performed to examine the role of E2Fs in neuronal death.

As we know pocket proteins inhibit expression of E2F regulated genes by directly

binding to and blocking the E2F activation domains or by recruiting chromatin-

modifying proteins (such as HDAC and polycomb group proteins). The inactivation of

one major pocket protein, pRb, induces ectopic S-phase entry and apoptosis in both CNS

and PNS neurons (Huang et al., 1992). Deletion of E2F1 or E2F3 can partially rescue the

CNS phenotype of Rb null mice (Tsai et al., 1998; Ziebold et al., 2001). Although conditional knock out Rb does not promote death of postmitotic neurons in the CNS, it causes defects in cell cycle and enhanced neurogenesis (de Bruin et al., 2003b). P107 or p130 null mice show strain dependent neuronal death, and all neuron death induced by disruption of pocket proteins involves the release of E2Fs and the activation of E2F

responsive genes. The activation of these genes may result in ectopic S phase entry and

the initiation of the apoptosis pathway. This intersection of cell cycle and cell death

pathways is an attractive mechanistic explanation of the many descriptive studies linking

the two. It also is a major impector for examining.

Over-expression of E2F1 causes neuron death in several model systems. E2F1 is up-

regulated in neurons subjected to cell cycle related neuron death (CRND) stimuli in vitro

and in vivo, and is also elevated in degenerating neurons of patients with AD and Down’s

syndrome (Liu and Greene, 2001). The reverse is also true, cultured cortical neurons or

cerebellar granule neurons from E2F1 knockout mice are resistant to stimuli leading to

CRND (Giovanni et al., 1999; O'Hare et al., 2000). Green and colleagues, working

primarily with PNS neurons, have proposed a model that links E2F1 to the neuron

apoptotic pathway. In their proposal, E2F1 induces the transcription of CDK1, which

contributes to death by phosphorylating BAD and promotes its translocation from the

cytoplasm to mitochondria. Subsequently, b-myb and c-myb (transcription factors with

proapoptotic activity that are repressed by Rb/E2F complex) are up-regulated. It is known

that over-expression of Myb proteins can induce neuron death. The likely target of the

myb proteins is BIM - a proapoptotic member of the Bcl2 family(Estus et al., 1994; Liu

and Greene, 2001). Another attractive E2F1 targets is Apaf-1, which is regulated by E2F1

and p53. Rb-/- mice embryos exhibit increased levels of Apaf-1. Analysis of compound mutant embryos lacking both Rb and Apaf-1 revealed that Apaf-1 is absolutely required for apoptosis in the central nervous system and lens. This dependence on Apaf-1 should be considered in light of the requirement, documented previously, for E2F1 and p53

induced cell death in the respective tissues. Loss of Apaf-1 specifically suppresses

apoptosis, but not the proliferation and differentiation defects in Rb-mutant embryos. It

should be noted, however, that most of findings come from in vitro culture systems or from embryonic stage cells. The role of E2F1 in the cell cycle regulation of the adult neurons and the cell cycle related neuronal death (CRND) is not known. My thesis work provides some clues in this direction.

As we talked in the very beginning, there is another group of proteins which are very important for cell cycle progress – cyclin dependent kinases (CDKs). In post-mitotic neurons, they do not need the normal CDKs, however, they have their unique CDK –

CDK5, which plays very important role for the normal neuronal function.

Background of CDK5

CDK family and normal regulation

Cyclin dependent kinases (CDKs) are a family of serine/threonine kinases that are

important for the progression of normal cell cycle events. Eleven members of the CDK

family have been identified to date. They are well conserved at the level of primary

amino acid sequence (sharing 40-75% identity), and 6 of them, CDK1-4, 6 and 7 are

considered important for the cell cycle regulation (Morgan, 1997). Of the others, CDK5

is known to be important for nervous system development and function, while CDK8 and

CDK9 are appreciated as regulators of transcription (Akoulitchev et al., 2000; Sano and

Schneider, 2003). The function of CDK10 is not known, but it has been shown to interact

with the transcription factor Ets2 (Kasten and Giordano, 2001).

The foregoing description makes it plain that a well regulated cell cycle requires the

activity of the mitotic CDKs to be strictly regulated. The regulation occurs through both

activation and inhibition. Monomeric, unmodified CDKs have no kinase activity. As

their name implies, they must bind to their regulatory cyclins as an initial step in their

activation. The cyclins are themselves a diverse group of proteins. There are 8 different type cyclins, distinguished by the letters A to H. They bind and activate distinct CDKs at distinct cell cycle stages (Malumbres and Barbacid, 2005; Bloom and Cross, 2007). Even after cyclin binding, however, CDKs are only partially activated. To be fully activated, they must be phosphorylated at a conserved threonine residue located in a region known as the catalytic T loop. Phosphorylation is accomplished by the CDK activating kinase

(CAK) - CDK7:cyclin H – a distantly related CDK:cyclin pair (Shuttleworth, 1995).

This modification flattens the T-loop resulting in critical changes in the substrate binding

site. Phosphorylation acts primarily to enhance the binding of protein substrates (Russo

et al., 1996), but it also enhances the affinity of the CDK:cyclin interactions (Desai et al.,

1995).

CDKs are also regulated by inhibition. The activity of CDK:cyclin complex can be

reduced by the phosphorylation of the CDK subunit on inhibitory sites by dual specificity

kinases (such as WEE1 and MYT1). This inhibition is relieved when the CDC25

phosphatases dephosphorylate these residues. Second, the binding of CDK inhibitors

known as CKIs to the CDK:cyclin complex also can inhibit the kinase activity. There are

two types of CKIs –the INK family and the WAF/KIP family. There are four members in

INK family (p16INK4a, p15INK4b, p18INK4c and p19INK4d). They serve as competitive

inhibitors of the the CDK4/6:cyclin D complex, from which CDK4 dissociates when they

bind. They can also bind CDK4 directly and prevent its binding to cyclin D (Lees, 1995).

There are three members of the WAF/ KIP family--- p21WAF1, p27 KIP1 and p57 KIP2.

They share a conserved region at their N- terminus, which is important for binding and

inhibition of CDK:cyclin complexes. Unlike the INK4 family, WAF/KIP family

members bind to the entire CDK:cyclin complex, inhibiting its activity. They are able to inhibit all of the G1 CDK:cyclin complexes as well as the CDK1:cyclin B complex

(Harper et al., 1993; Xiong et al., 1993; Harper et al., 1995; Matsuoka et al., 1995).

Each cyclin has a unique expression pattern during the cell cycle. This determines the activity of the CDKs at the different stage of the cell cycle. Cyclin D bind and activate

CDK4/6 in the early G1 phase (Meyerson and Harlow, 1994; Jinno et al., 1999). The D type cyclins are important integrators of mitogenic signaling, as they are the main product of RAS/RAF/ MAPK pathway. They are unstable and transported from the nucleus to the cytoplasm where it is targeted for proteolysis. They are essential for the cells through G1 to S phase. CDK2 is sequentially activated by the E-type cyclins (cylinE1 and cyclin E2) during G1/S phase transition and the A-type cyclins (cyclin A1, A2) during S phase.

CDK2 is required for the completion G1 and initiation S phase. The role of CDK3 is not clear until now, mainly due to its low expression levels. It binds with cyclin C and might promote the S phase transition like CDK2 (Hofmann and Livingston, 1996). In several strains of mice, however, it is truncated and presumably inactive, indicating that it might

be dispensable for the cell cycle (Ye et al., 2001). CDK1 pairs with cyclin A2 and cyclin

Bs. It is required to complete the G2/M phase transition.

CDK5 is unique in the CDK family

As their name suggests, most CDKs are involved in the cell cycle progression. From this

viewpoint, CDK5 is an atypical member of the CDK family. It was originally identified

by its close sequence homology (60%) to CDC2 (Lew et al., 1992; Meyerson et al., 1992).

Yet ectopic expression of CDK5 in mammalian cells or yeast does not promote cell cycle

progression (Meyerson et al., 1992; van den Heuvel and Harlow, 1993). Nonetheless,

CDK5 phosphorylates many of the same substrates as CDK2 or CDK1, making the

absence of a role in cell cycle control a curious observation.

Regulation of CDK5

Like the traditional CDKs, the activation of CDK5 requires its association with a

regulatory partner. It can bind several of the normal cyclins, but it is not activated by

them (Xiong et al., 1992; Zhang et al., 1993; Miyajima et al., 1995). It is activated instead

by its specific partners – p35 and p39. These two proteins are structurally similar to

cyclins yet they share no homology to cyclins at the amino acid level. And since the

phenotype of p35-/- ;p39-/- double knockout mice is virtually identical to the CDK5 knockout itself (Ko et al., 2001), the suggestion is that p35 and p39 are the only two relevant activators of CDK5 during development.

A high level of CDK5 kinase activity is primarily detected in the nervous system (Lew et al., 1994; Tsai et al., 1994; Tang et al., 1995) as this where the levels of p35 and p39 are highest. Nonetheless, the CDK5 protein itself is found in all tissues. Besides the brain, low levels of CDK5 kinase activity are also present in adult mouse prostate and embryonic limb buds (Zhang et al., 1997) The expression of p35 and p39 in brain is found in overlapping but distinct sub-cellular compartments including growth cones and synapses (Humbert et al., 2000d; Humbert et al., 2000c; Niethammer et al., 2000; Fu et al., 2001). Evidence points to the fact that p39 can compensate for most of the functions of p35 since in p35-/- mice CDK5 kinase activity is present, albeit reduced (Hallows et al.,

2003). The reverse is true as well; there are no obvious abnormalities in p39-deficient mice (Ko et al., 2001).

Phosphorylation of CDK5

As discussed above, full activation of mitotic CDKs depends on the phosphorylation of a conserved residue in its activating T loop. Phosphorylation of the homologous loop of

CDK5, however, is not required for its maximal activation (Poon et al., 1997). One study shows that the structure of the CDK5:p25 complex (p25 is a caRboxy-terminal proteolytic product of p35) is in a fully active conformation which is indistinguishable from phosphorylated CDK2 bound to cyclin A. This offers a hint as to why CDK5 does not need to be phosphorylated to achieve full activity (Tarricone et al., 2001). As mentioned above, the mitotic CDKs are also regulated by the phosphorylation of Thr14 and Thr15 by kinases Wee1 and Myt1 (which serves to inhibit their activity). In CDK5, these two sites are conserved, but are not phosphorylated by Wee1 in vitro (Poon et al.,

1997). Phosphorylation of Thr14 on CDK5 by an inhibitory protein kinase purified from

bovine thymus cytosol inactivates CDK5 (Matsushita et al., 1996; Matsuura and Wang,

1996). Thr15 on CDK5 is phosphorylated by c-Abl, but rather than blocking CDK5 activity, this phosphorylation increases CDK5 kinase activity (Zukerberg et al., 2000).

Inhibitors

As discussed above, the activities of traditional mitotic CDKs are inhibited by several different CKIs. For example, p21 and p27 can effectively inhibit CDK2, the homologue of CDK5. CDK5 activity, however, escapes inhibition when bound to p35 (Lee et al.,

1996). Whether there are other inhibitory subunits to regulate CDK5:p35 complex is unknown at this time.

Subcellular localization

Another mechanism of regulation of traditional CDKs is the subcellular localization change of their regulatory partner cyclins. P35 and p39 have an animo-terminal myristoylation signal motif that targets them to cell membranes. The sub-celluar distribution of CDK5 is therefore controlled by p35 and p39. In p35-/- mice brain, the subcellular distribution of CDK5 is altered and phosphorylation status of its substrates is also altered (Hallows et al., 2003).

Function of CDK5

The roles of CDK5 in development

Migration

CDK5-deficient mice reveal an important role for CDK5 in CNS development. Cdk5-/- homozygotes die around birth with severe disruptions in the neuronal layering of cerebral cortex, hippocampus, cerebellum and olfactory bulb (Ohshima et al., 1996; Gilmore et al.,

1998; Ohshima et al., 1999). In the cerebral cortex of Cdk5-/- mice, after the early

arriving neurons of layer VI split the preplate, the later born neurons cannot migrate past

their predecessors to their appropriate superficial positions. Instead they stall underneath

the subplate.

Neuronal migration depends on the regulation of actin, microtubule and intermediate-

filament cytoskeletal components, and modulation of the cell adhesion. The kinase

activity of CDK5 is involved in all these processes. CDK5 can phosphorylate and

facilitate the degradation of nestin, an intermediate filament protein found in progenitor

cells and down-regulated after neuronal differentiation begins (Sahlgren et al., 2003).

Nestin continues expressing in Cdk5-/- deficient cortex. Consistent with the delayed nestin

degradation, there is also reduced expression of neuron-specific class III β-tubulin protein

(an early neuronal marker) and a very low level of expression of Map2 (a mature

neuronal marker). These data suggest that CDK5 is involved in cytoskeletal maturation

and neuronal differentiation (Cicero and Herrup, 2005). CDK5 is also involved in the

post translational modification of microtubule-associated proteins such as tau (Patrick et

al., 1999). In CDK5-deficient neurons, the basal level of tau phosphorylation is reduced

compared with the wild type neurons (Cicero, Wang et al, and unpublished observations).

Recent studies have identified several novel CDK5 substrates that are involved in

neuronal migration: focal adhesion kinase (FAK) and doublecortin (DCX).

Phosphorylation of FAK by CDK5 at Ser732 is involved in the regulation of a

centrosome-associated microtubule structure to promote nuclear translocation, which is

important for neuronal positioning (Xie et al., 2003). Phosphorylation of DCX by CDK5

at Ser297 decreases its ability to bind and stabilize microtubules and this phosphorylation is necessary for DCX induced migration (Tanaka et al., 2004). These studies indicate that

CDK5 governs neuronal migration in the developing neocortex by phosphorylation of various proteins involved in these processes.

Survival

Several lines of evidence indicate that CDK5 activity is also needed for neuronal survival.

In Cdk5-/- mice, swelling of cell soma and nuclear marginalization is observed in

brainstem and spinal cord neurons (Ohshima et al., 1996). In CDK5 chimeric mice, the

percentage of Cdk5-/- neurons present in the cerebral cortex is much lower than would be

predicted if CDK5 deficiency had no effect on cell survival (Gilmore and Herrup, 2001).

Further evidence comes from large number of cells that are stained with either TUNEL or

activated caspase-3 in the cortical plate of Cdk5-/- brains. This indicates substantial neuronal loss during the final maturation of even those neurons that migrate as far as cortical plate (Cicero and Herrup, 2005).

The roles of CDK5 in the adult

The role of CDK5 in the synaptic transmission

CDK5 and its activators p35 and p39 are present in sub-cellular fractions that are enriched for synaptic membranes, where they localize to both pre-and postsynaptic

compartments (Humbert et al., 2000c; Niethammer et al., 2000). CDK5 has the ability to

phosphorylate pre- and post synaptic proteins such as snapsin1,MUNC18, amphiphysin,

synaptojanin1, PSD-95 as well as the NMDA receptor (Matsubara et al., 1996; Shuang et

al., 1998; Fletcher et al., 1999; Floyd et al., 2001; Li et al., 2001; Morabito et al., 2004).

This activity has functional consequences as neurotransmitter release through P/Q type

voltage-dependent calcium channels and NMDA receptors is down-regulated by

phosphorylation (Tomizawa et al., 2002; Wang et al., 2003).

CDK5 also modulates CNS dopamine signaling through phosphorylation of DARPP-32.

Phosphorylation of DARPP-32 by Protein kinase A (PKA) on Thr34 promotes dopamine

signaling by inhibition of protein phosphatase-1(PP-1). By contrast, when CDK5

phosphorylates DARPP-32 on Thr75, it becomes an inhibitor of PKA, thus reducing the

efficiency of dopamine signaling (Bibb et al., 1999). Moreover, CDK5 is also involved

in the processes of the dopamine synthesis and releasing (Chergui et al., 2004; Moy and

Tsai, 2004) . Those studies revealed the important role of CDK5 in the dopamine

signaling. CDK5 is also required for the normal development of the neuromuscular junction (Fu et al., 2001).

CDK5 also functions in the synapse to regulate endocytosis, an important step in synaptic

transmission. CDK5 can phosphorylate dynamin and other dephosphins, which mediate

synaptic vesicle endocytosis (Tomizawa et al., 2002; Tan et al., 2003). Further, CDK5 is

involved in the modulation of phosphoinositide (PI) signaling. The regulation of level

and localization of PI(4,5)P2 is important for endocytosis, and CDK5 phosphorylates

both PiPKIr – the kinase involved in the generation of PI(4,5)P2 – and synaptojanin1 – a

PI(4,5)P2 phosphotase (Lee et al., 2004; Lee et al., 2005).

Neuronal survival

In human Alzheimer’s disease, it has been reported that there is an unusual accumulation

of p25 due to the calpain-mediated cleavage of p35. This accumulated p25 results in

elevated CDK5 activity (Grynspan et al., 1997; Lee et al., 2000). Since CDK5 can

function as a tau kinase and since hyper-phosphorylated tau is the major constituent of

the neurofibrillary tangle, a pathological hallmark of AD, it has been proposed that

elevated CDK5 activity contributes to AD pathology, including neuronal degeneration.

CDK5 interacts closely with early neurofibrillary tangles in AD patients, further

supporting this idea. Further, in two different lines transgenic mice that over-express p25

or hyper-phosphorylated tau, neuorofibrillary tangles and neurodegeneration are found

(Ahlijanian et al., 2000; Van den Haute et al., 2001; Bian et al., 2002; Cruz et al., 2003;

Noble et al., 2003). It should be noted, however, that the activity of other kinases such as

Gsk3β is also increased (Noble et al., 2003), leaving a level of uncertainty as to which

kinase plays the main role in Aβ-related tau hyper-phosphorylation and neuronal death

(Bian et al., 2002; Hallows et al., 2003; Tandon et al., 2003; Giese et al., 2005; Hallows et al., 2006; Plattner et al., 2006).

In addition to AD, elevated levels of CDK5 and p35 are observed in the cell body of

apoptotic cells following ischemia (Hayashi et al., 1999) and in models of Parkinson’s

disease (Neystat et al., 2001). Elevated p25 and CDK5 kinase activity are also found in

the mouse model of ALS as well as in human ALS patients (Nguyen et al., 2001; Nguyen and Julien, 2003). Together, these observations suggest an important and continuous role for CDK5 in the physiology of the adult neurons.

Deregulation of CDK5 (p25/p35, levels)

As mentioned above, CDK5 deregulation has been implicated in several neurodegenerative diseases, including Alzheimer disease, amyotrophic lateral sclerosis,

Parkinson’s Disease and progressive supranuclear palsy (Borghi et al., 2002; Tseng et al.,

2002; Nguyen and Julien, 2003; Smith et al., 2003a).

Now it is thought that the deregulation of CDK5 activity is caused by the p35 or p39 proteolytically cleaved into p25/p29 by calpain(Lee et al., 2000; Patzke and Tsai, 2002) .

P25 has all the elements necessary for CDK5 binding and activation (Qi et al., 1995;

Poon et al., 1997). It is different with p35. It has a longer half-life than p35. Second, it localizes in the cell body and nucleurs because it lacks the amino-terminal myristoylation site which directs p35 to associate with memebrane. So the generation of p25 is likely to disrupt the normal regulation of CDK5.

The phenotype of p25 over-expressing mice suggests that too much CDK5 activity may be as bad as too little. As discussed, CDK5/p25 has increased kinase activity compared to CDK5/p35, and the generation of the p25 fragment of p35 often accompanies apoptosis upon challenge by a variety of apoptotic stimuli, like neurotoxicity, ischemia and oxidative stress (Lee et al., 2000; Nath et al., 2000; Fu et al., 2002; Zhu et al., 2002).

Further, the mis-localization of CDK5, as well as its hyperactivation in neurons, may

alter substrate specificity and trigger various events that induce neurodegeneration.

Indeed, introduction of p25 into cultured primary neurons induces neurite retraction,

microtubule collapse and apoptosis (Patrick et al., 1999). In part, the consequences of

p25 over-expression might depend on the specific neuronal subtype and the nature of the

death inducing stimulus. For example, p25, p35 and CDK5 are all down-regulated in

hippocampal neurons after treatment with various apoptotic stimuli (Kerokoski et al.,

2001).

CDK5 and the cell cycle regulation in neurons

The lack of a role of CDK5 in normal cell cycle regulation is still a mystery given its high

level of structural homology with traditional CDKs. It was reported that CDK5 could

bind with normal cyclins such as cyclin D, cyclin E and PCNA (Xiong et al., 1992;

Zhang et al., 1993; Miyajima et al., 1995), but no CDK5 kinase activity is detected in

those complexes. The mystery may be beginning to be solved, however, and CDK5 may

yet take its place in the list of CDKs that help to regulate the cell cycle. Cicero et al,

2005 report a novel and unexpected role for CDK5 in neuronal cell cycle control.

Analysis of Cdk5-/- mice found that loss of CDK5 leads to loss of cell cycle control in

cortical plate cells of E16.5 mouse neocortex. This includes the abnormal expression of

cell cycle proteins such as cyclin D, cyclin A, and PCNA as well as BrdU incorporation.

These unexpected cell cycle events were found in multiple layers of Cdk5-/- neocortex.

Further examination revealed that the “cycling” neurons were dying, as indicated by their staining positive for DNA breakdown (TUNEL) and activated caspase-3. In vitro cultures of primary Cdk5-/- neurons continue to express cell cycle related protein such as

PCNA and continue to incorporate BrdU, consistent with the in vivo data. Importantly,

introduction of expression plasmids encoding wild-type CDK5 into Cdk5-/- neurons stops cell cycle re-entry. Further, both in vitro and in vivo, the abnormal cell cycle events are

coupled with a failure of neuronal differentiation in CDK5-deficient neurons. In the

aggregate, therefore, these data strongly suggest that CDK5 plays a dual role as a cell cycle suppressor and as a facilitator of cell differentiation during neuronal development.

The important thing is to illustrate that how CDK5 control the neuronal cell cycle. Our studies in Chapter 3 suggest that localization of CDK5 might play important role in this cell cycle deregulation.

Figures

G0 G1

Trimarchi and Lees, 2002 M

Cyclin B G2 Cdk1

Cyclin A Cdk1

www.fhcrc.org/science/ labs/fero/CellCycle.html

Figure 1: Normal cell cycle regulation.

Figure 2: The structure of E2Fs. (From DeGregrie and Johnson et al, Current

Molecular Medicine, 2006, 6, 739-748)

Genes involved in cell cycle regulation and P107, Rb, cyclinE1,cyclinD3 , cyclinA,

DNA replication CDC2,CDC6,CDC24A,MCMs,ORC1L,

POLA2,Top2A,PCNA, TK1,MAD2,c-

Myb,b-Myb, E2F1-3,E2F7,Rb1,CDKN2C

and CDKN2D,.

Genes involved in apoptosis Apaf-1, caspase3, caspase7, caspase8, Bcl-

1, BAD, BAK1, TP73, Ask1.

Genes involved in DNA repair P53, BRCA1,chk1, MSH2, MShH6,MLH1,

UNG, RPA3, PMS2,RAD51 and RAD54L,

Genes involved in differentiation and Homeobox, PcG genes and TGF-beta development

Table 1: Selected genes regulated by E2F

Chapter 2: E2F1 works as a cell cycle suppressor in mature neurons

Abstract

Neurons are highly differentiated cells that normally never enter a cell cycle; if they do

the result is usually death, not division. For example, cerebellar granule neurons in staggerer and lurcher mutant mice initiate a cell cycle-like process just before they die.

E2F1 is a transcription factor that promotes cell cycle progression. As E2F1 is also involved in apoptosis we bred double mutants (E2f1-/-; staggerer and E2f1-/-; lurcher) to assess its role in the cell cycle-related death of cerebellar granule cells in vivo. We found

neither granule cell cycle initiation nor cell death was significantly altered in either

double mutant. However, after postnatal day 10, neurons throughout the CNS of

E2f1-/- and E2f1+/- animals were found to express cell cycle proteins and replicate their

DNA. The appearance of Purkinje cell dendrites at one year of age suggests that the mutant cells also undergo a slow atrophy. These events are cell autonomous, as cultured

E2f1-/- cortical neurons "cycle" in vitro while wild-type neurons do not. Our results suggest that in mature CNS neurons E2F1 functions as a cell cycle suppressor.

Introduction

In the developing CNS, once young neurons leave the neurogenic regions of the ventricular and subventricular zone, they will never again complete a full cell cycle. Yet evidence is mounting that adult nerve cells are not permanently postmitotic (reviewed in

Herrup and Yang, 2007). Target related cell death, for example, is preceded by the dying neurons entering a cell cycle-like process (Herrup and Busser, 1995). Similarly in adult onset neurodegenerative diseases neuronal death is often preceded by the neurons re- engaging a cell cycle-like process. In human Alzheimer disease (AD) and its mouse models, many cell cycle proteins are over-expressed in the neurons at risk for death

(Nagy et al., 1997a; Busser et al., 1998; Herrup and Yang, 2001; Yang and Herrup, 2006).

The E2F family of transcription factors plays a pivotal role in cell proliferation,

differentiation and apoptosis through transcription regulation (Ishida et al., 2001; Muller

et al., 2001; Weimann et al., 2001; Ren et al., 2002; Stanelle et al., 2002; Weimann et al.,

2002; Yang et al., 2003). Among the eight structurally related E2F members, E2F1,

E2F2 and E2F3a are transcriptional activators, an activity that is blocked by their

association with the retinoblastoma (RB) tumor suppressor. This association is lost after

RB is phosphorylated by cyclin dependent kinases. Cell cycle promotion is the most

frequently cited function of E2F, but deregulation of E2F1 can also induce apoptosis in

different cellular contexts (Wu and Levine, 1994; Johnson, 2000; Heldin, 2001; Kaelin,

2003; Mundle and Saberwal, 2003; Vermeulen et al., 2003). For example, E2F1 is

elevated in the brains of AD and Down’s syndrome patients (Liu and Greene, 2001), and

E2f1-deficient neurons in culture are resistant to stimuli leading to cell cycle related

neuronal death (CRND) (Giovanni et al., 2000; O'Hare et al., 2000). Similarly, in RB-

deficient mice there is profound neuronal loss in the CNS, and E2f1-/-; Rb1-/- double

mutants show that this loss is E2F-dependent (Tsai et al., 1998; Ziebold et al., 2001).

Given the key role played by these proteins in cell division and apoptosis, it is surprising

in my many ways that E2F1, E2F2 or E2F3 knockout mice are born as viable if abnormal

animals. In older E2f1-/- mice, paradoxically, cellular dysplasia and tumor formation has

been described in several tissues.

In both staggerer (Rorasg) and lurcher (Grid2Lc) mice, there are well-defined CRNDs in

post-mitotic cerebellar granule cells (Herrup and Busser, 1995) because of the failure of

Purkinje cell trophic support. Because of the results from the RB-deficient mice (Tsai et

al., 1998; Ziebold et al., 2001) we predicted that E2F1 might be involved in this neuronal

death, so we created both E2f1-/-; staggerer and E2f1-/-; lurcher double mutant mice. We

observed the rescue of neither cell cycle activity nor cell death in these animals. Instead,

we found ectopic cell cycle events in many other neurons throughout E2f1-/- mutant brain.

These events include constitutive expression of proteins normally only found in cycling cells as well as DNA replication. We suggest that E2F1 works as cell cycle suppressor in differentiated neurons and other E2F members cannot compensate this role.

Experimental Procedures

Animals

A breeding colony of mice with a targeted disruption of the E2f1 gene (Field et al., 1996) was established from founders obtained from The Jackson Laboratory. Generation of mutants was achieved through the mating of heterozygous E2f1 males and females. The mice were maintained on a mixed (C57BL6/Jx129/S4) background. Genotyping was performed on DNA extracted from tail biopsies using PCR techniques as described previously (Field et al., 1996).

For E2f1-/-; Rorasg/sg double mutant mice, we first crossed E2f1-/- mice with Rorasg/+ mice to get E2f1+/-; Rorasg/+ mice, then intercrossed the double heterozygotes to get double

mutants. We used a similar method to create E2f1-/-; Grid2+/Lc mice. We first crossed

E2f1-/- mice with Grid2+/Lc mice to get E2f1+/-; Grid2+/Lc mice. Then we crossed E2f1+/-;

Grid2+/Lc with E2f1-/-; Grid2+/+ mice to get E2f1-/-; Grid2+/Lc mice. Genotyping of the staggerer locus was performed on DNA extracted from tail biopsies using PCR techniques described previously. Grid2+/Lc mice were identified based on their ataxia.

All animals were housed in the Case Western Reserve University Medical School Animal

Resource Center, a facility fully accredited by the Association for Assessment and

Accreditation of Laboratory Animal Care. All procedures for animals were approved by the Institutional Animal Care and Use Committee of the Case Western Reserve

University.

Histology

After animals were deeply anesthetized with Avertin (0.02 cc/mg bodyweight), they were transcardially perfused with PBS, followed by 4% paraformaldehyde in 0.1 M sodium phosphate buffer (PB). The brain was dissected and transferred to fresh 4% paraformaldehyde at 4°C overnight. The brains were then cryoprotected by sinking in

30% sucrose/PB at 4°C overnight. After bisecting along the midline, the brains were embedded in OCT, and 10 µm cryostat sections were cut and allowed to air dry on

SuperPlus glass slides overnight. All slides were stored at -80°C.

Single and double immunocytochemistry

Antibodies The proliferating cell nuclear antigen (PCNA) mouse monoclonal antibody and rabbit polyclonal antibody (Dako, High Wycombe,UK) were diluted 1:300 in 10% goat serum/PBS blocking buffer before use. Rat anti-BrdU (1:100; Abcam, Cambridge,

UK) was used to dectect DNA replication. A rabbit polyclonal cyclin A antibody raised against the C-terminal domain of cyclin A2 was used at a dilution of 1:500 (ab 7956;

Abcam, Cambridge, UK). The mouse anti-Tuj1 (1:1000; Covance, Princeton, NJ) was used as neuronal marker. The mouse monoclonal calbindin antibody at a dilution of 1:500

( Sigma, St.Louis, MO) was used as Purkinje cell marker.

Double fluorescence immunocytochemistry was used on the mouse cryosections.

Sections were first rinsed in PBS, followed by pretreatment in 0.1M citrate buffer for 6–8

min at 95°C. After the slides had cooled in buffer for 45 min at room temperature, slides

were rinsed in PBS. Sections were incubated in 10% goat serum in PBS to block

nonspecific binding for 1 h at room temperature. All primary antibodies were diluted in

PBS containing 0.5% Triton X-100 and 10% goat serum and then were applied to sections and incubated overnight at 4°C. After rinsing in PBS, they were incubated for 2 h with secondary antibody, which was conjugated with various fluorescent Alexa dyes

(dilution, 1:1000; Molecular Probes, Eugene, OR). The sections were then rinsed in PBS and reincubated in 10% goat serum blocking solution for 1 h, followed by addition of the second primary antibody (raised in a different species from the first primary antibody) for a second overnight incubation at 4°C. Sections were then rinsed in PBS, and both secondary antibodies conjugated with different fluorescence dyes were applied to the sections for 2 h at room temperature. After rinsing, all sections were mounted in

PBS/glycerol under a glass coverslip.

Fluorescent in situ hybridization - FISH

Three mouse-specific DNA probes were generated from bacterial artificial chromosomes

(BACs), which carried unique mouse genomic DNA sequences. One of the DNA probes

(480C6) was made from the region that encodes the beta-amyloid precursor protein located on mouse chromosome 16. The other two probes were generated from overlapping BACs (170L21 and 566) containing the structural gene for mouse aldolase C on mouse chromosome 11. Each of the three probes covers 100–300 kb. They were labeled by standard nick translation protocols using digoxygenin-labeled dUTP.

After labeling, probes were concentrated with mouse Cot-1 DNA (Invitrogen, San Diego,

CA) to block hybridization to repetitive sequences. Before hybridization, all tissue sections were rinsed in PBS and then pretreated with 30% pretreatment powder (Oncor,

Gaithersburg, MD) for 15 min at 45°C followed by treatment with protease (0.25 mg/ml;

Oncor) for 25 min at 45°C. After rinsing in 2x SSC, the slides were dehydrated through graded alcohols and allowed to air dry. Labeled probe was applied to the individual

sections, which were then covered with a glass coverslip and sealed with rubber cement.

To denature DNA, slides were heated at 90–92°C on a heated block for 12 min and

hybridized with probe overnight at 37°C. After rinsing in 50% formamide/2x SSC at

37°C for 15 min, the slides were transferred to 0.1x SSC buffer for 30 min at 37°C and

then rinsed in 0.5 M phosphate buffer with 0.5% NP-40 (PN buffer) at room temperature.

To block nonspecific antibody binding, 10% goat serum in PBS was applied to the

sections, followed by incubation in mouse anti-digoxygenin primary antibody (dilution,

1:200; Boehringer Mannheim, Indianapolis, IN) for 30 min at 37°C. After rinsing,

secondary anti-mouse antibody conjugated with Alexa 488 (dilution, 1:250) was applied

to the sections for 30 min at 37°C. Slides were rinsed three times in PN buffer, and the

sections were counterstained with either 4’, 6’-diamidino-2-phenylindole (DAPI) or

propidium iodide and covered with a coverslip.

The number of spots of hybridization in each nucleus was determined at 1000x

magnification under fluorescent illumination. Images were captured on a Leitz (Wetzlar,

Germany) research microscope equipped with a digital camera (Prog C14). Purkinje

cells were identified on the basis of the size of their nucleus and their position in the

Purkinje cell layer. Cortical neurons were identified on the basis of the size and the

position of their nucleus. For each of genotypes (P30 E2F1+/+, and E2f1-/- ), 60 cortical

neurons from layer II and layer III were randomly picked and scored from six different

areas for each section. A total of 12 sections was scored for each genotype.

Cell counts

For each of the three genotypes (P30 E2f1+/+, E2f1+/-, and E2f1-/-), we began at one end of the Purkinje cell layer within a given section and continued counting until the other end of layer was reached. All calbindin positive cells were scored for the presence or absence of the cell cycle marker. Only cells with a discernable portion of their nucleus in the

section were scored. Two animals were scored for each genotype.

Primary Neuronal Cultures

Pregnant dams were sacrificed by cervical dislocation and the uteri dissected and placed

on ice during the embryo harvest procedure. For E2F1-deficient cultures, all embryos

from an E2f1-/- x E2f1-/- mating were treated separately. Embryos were collected in ice cold PBS-glucose, and the cortical lobes were dissected out. Meninges were removed and the cortices were placed in 1X trypsin-EDTA for 15 minutes at 37ºC. The tissue was removed from the trypsin solution and placed in DMEM with 10% FBS to inactivate the trypsin, followed by transfer to Neurobasal media supplemented with B-27, penicillin/streptomycin (1X), and L-glutamine (2mM). Tissue was triturated 10X through a 5mL pipette and allowed to settle to the bottom of a 15mL conical tube for approximately one minute. Cells in solution above the tissue pellet were removed and used in all subsequent procedures. Aliquots of the suspension were stained with Trypan

Blue to allow counts of live and dead cells. The culture substrate used was poly-L-lysine

(0.05mg/mL) coated on plates or glass coverslips. Cells were grown in 24 well plates at a

density of 50,000 cells/well. All cultures were grown for a minimum of 5 days in vitro

(DIV) before any treatment. To assess cell cycle activity, BrdU was added to the culture

to obtain a final concentration of 10 µM. After 5 days, cultures were fixed with 4%

paraformaldehyde in phosphate buffer at room temperature for 30 minutes, then washed

and stored in PBS. All experiments were performed on a minimum of three litters; each

condition was examined in triplicate.

Immunoblotting

E2f1-/- littermates were killed by cervical dislocation. Brains were rapidly dissected and

placed in lysis buffer (20 mM Tris, 1% Triton X-100, 100 mM NaCl, 40 mM NaF, 1 mM

EDTA, 1 mM EGTA, 1 mM Na3VO4, aprotinin, leupeptin, and PMSF) on ice and homogenized. Lysates were then sonicated for 15 s and centrifuged at 13,000 rpm for 15 min at 4°C. Supernatant was collected and protein was measured by the Bradford method.

Equal amounts of protein (10 or 15 µg) in Laemmli buffer were boiled for 7 min. The samples were run on SDS-polyacrylamide gels and transferred onto polyvinylidene

difluoride membranes as described previously (Combs et al., 2001). Antibodies used for immunoblotting were as follows: mouse anti-E2F2 (1:200; Santa Cruz), mouse anti-E2F3

(1:200; upstate), mouse anti-actin (1:5,000; sigma), donkey anti-mouse HRP (1:20,000;

Jackson Immuno-Research, West Grove, PA), and donkey anti-rabbit HRP (1:1500;

Amersham Biosciences). Blots were developed with ECL (Amersham Biosciences)

according to the instructions of the manufacture

Results

Granule cell death-by-cycle in staggerer and lurcher mice is

E2F1-independent

Staggerer and lurcher are two mutations of mice that suffer postnatal granule cell loss in

the cerebellum (Sidman et al., 1962; Caddy and Biscoe, 1979). The recessive mutation

‘staggerer’ is the result of the inactivation of the Rora gene encoding an orphan retinoid

receptor (Hamilton et al., 1996). The dominant mouse mutation lurcher is caused by a

point mutation in the orphan glutamate receptor gene Grid2 (Zuo et al., 1997). Both

mutants have severe ataxia due to the cell-autonomous loss and/or deformation of

Purkinje cells. In both mutants, there is a well-defined cell cycle re-entry process that

occurs before the granule cells die (Herrup and Busser, 1995). As the E2F1 protein is

involved in both cell cycle and cell death we sought to learn whether it played a role in

staggerer or lurcher granule cell death. We produced double homozygous animals

(E2f1-/-; Rorasg/sg) by the crosses described in Methods. We used a similar mating

strategy to produce E2f1-/-; Grid2+/Lc mice. The cerebella of these animals and their littermate controls were examined at P30. At this age, the internal granule cell layer (IGL) of single mutant mice (staggerer or Lurcher) is thin and sparsely populated because of

the death of most of the granule cells (Figure 1A, B). Significantly, the size of

cerebellum of the E2f1 double mutant mice was similar to that seen in staggerer or

Lurcher alone (Figure 1C, D). Thus, the deletion of the E2f1 gene provides no protection

against granule cell loss in either mutation.

Although the absence of E2F1 does not rescue the granule cells in either staggerer or

lurcher, we wanted to know whether the cell cycle events (CCEs) preceding the cell

death were affected. We immunostained sections from double mutant animals with

antibodies against a Purkinje cell marker, calbindin, and the proliferating cell nuclear

antigen (PCNA) – a commonly used cell cycle marker and a component of the DNA

polymerase complex that is elevated in S-phase and remains high through G2 (Bravo and

Macdonald-Bravo, 1987). As reported (Herrup and Busser, 1995), in Rorasg/sg or

Grid2+/Lc mouse cerebella there are PCNA positive cells in IGL (Figure 2A, G). We also saw PCNA staining in some Purkinje cells, but this was quite weak (Figure 2C, I). In the cerebellum of E2f1 double mutant mice; the pattern of PCNA staining in the IGL was unchanged, suggesting that E2F1 deficiency does not alter CCEs in staggerer or lurcher mice. We noticed, however, that PCNA staining of Purkinje cells were much brighter in double mutant mice compared with that in the single mutant mice (Figure 2F, L).

Cell cycle related protein expression in E2f1-/- mouse brain

The enhanced cell cycle protein expression in Purkinje cells prompted us to investigate

whether loss of E2F1 alone has any effect on cell cycle control in the cerebellar neurons.

Mice of three genotypes – E2f1+/+, E2f1+/-, and E2f1-/- – were examined by

immunohistochemistry. As expected, we saw no PCNA staining in any wild-type

Purkinje cell (Figure 3A). To our surprise, however, we found PCNA expression in the

nuclei of substantial numbers of Purkinje cells in the E2f1-/- mouse brain (Figure 3C).

The same unexpected expression of PCNA was also found in heterozygous, E2f1+/-, animals (Figure 3E). This indicates that the loss of even one allele of the E2f1 gene can

induce ectopic CCEs in mouse Purkinje cells. To determine the extent of this

phenomenon, we counted the number of calbindin/PCNA double-positive cells. We

found that nearly half of the E2f1-/- Purkinje cells had high levels of PCNA protein in

their nuclei, and further that there was no significant difference between E2f1 heterozygous and null mice (Figure 3G). To expand this finding, we used a second cell cycle maker, cyclin A (a partner of the Cdk2 and Cdk1 kinases and an S-G2 phase marker). As with PCNA, cyclin A appeared in both E2f1 heterozygous and null Purkinje cells. Unlike PCNA, however, the sub-cellular localization of cyclin A was not nuclear, as expected, but rather cytoplasmic (Figure 3D, F). This can be seen as cell body staining

(asterisks) as well as in a punctate configuration suggestive of a process such as an axon.

No cyclin A or PCNA-labeled Purkinje cells were found in control animals (Figure 3B).

The unusual pattern of cell cycle protein expression led us to examine other regions of

the E2f1-/- mouse brain. We found PCNA and cyclin A expressed in the nerve cells of

several neuronal populations in E2f1-/- animals and in heterozygotes. Perhaps the most

prominent of these sites was the neocortex (Figure 4D-I). The subcellular localization of

the cell cycle proteins was similar to that seen in Purkinje cells: PCNA in the nucleus as

expected and cyclin A ectopically expressed in the perikaryon and cell process. We also

found variation among the six layers of neocortex. There were more cell cycle positive

neurons in Layers II and III than in Layers V and VI. We found no cell cycle re-

expression in the neurons of hippocampus (data not shown). No cyclin A and PCNA

positive cells were found in the wild type cortex (Figure 4A-C). All of the cells in

Figures 3 and 4 are from brains of one-month old animals. The unusual protein

expression patterns do not appear to be transient, however, as we saw similar quantitative and qualitative PCNA and cyclin A staining patterns in adult mice (see below).

Neuronal DNA replication in the E2f1-/- brain

Cell cycle related protein re-expression alone is not sufficient to rigorously distinguish

DNA replication from DNA repair (Herrup and Yang, 2007; (Kuan et al., 2004).

Therefore, we used fluorescence in situ hybridization (FISH) to probe the ploidy of the

neuronal nuclei. We used 3 different mouse BAC probes to detect unique genomic

sequences on two different mouse chromosomes, 16 and 11. The results with probe

480C6, which recognizes sequences on chromosome 16, are shown in Figure 5. We

found many Purkinje cells and neocortical neurons with three or four hybridization

signals (three to four spots) in both mutant and heterozygous mice (Figure 5B, C, E, F).

In no case did we find an example of a neuron in which there were more than four spots

of hybridization. In wild type animals, all neurons in comparable regions had two or

fewer hybridization signals (Figure 5A, D). This indicates that there has been true DNA

replication in E2f1+/- and E2f1-/- neurons. The distribution of the ‘hyperploid’ neurons

matched our immunohistochemistry findings well. For example, there were substantially

more neurons with three or four hybridization signals in Layers II and III than in Layers

V and VI of neocortex.

Cell counts (details in Methods) further validate these impressions (Figure 4H). In wild-

type animals the number of cells with three or four spots in Layers II and III was about

5% of the total nuclei. In the same location in the E2f1-/- brains, over 20% of the nuclei had three or four spots of hybridization. We were unable to achieve reliable

immunocytochemistry in conjunction with our FISH protocol, precluding the positive identification of the cell type in which the hyperploid signals were found. Our qualitative impression is that many, if not most; of the three- and four-spot cells in the wild-type animals were smaller and likely to be non-neuronal. Some of the E2f1-/- hyperploid cells

were also of this class, but many more were found in large nuclei that are characteristic of

neurons. In the cerebellum, the position and large size of the Purkinje cell nuclei made

positive identification of the cell type more reliable than in neocortex. On occasion, in

both wild type and E2F1-deficient mice, we found cells with more than two hybridization

signals. The nuclei of these cells tended to be quite small, however, and their location

was outside the Purkinje cell layer. We are confident that these are not Purkinje cell

nuclei. FISH data obtained with another pair of BAC probes (170L21 and 566)

recognizing sequences on mouse chromosome 11 demonstrated similar results (data not

shown). In the aggregate, our data demonstrate that nerve cells in the E2f1-/- brain have undergone DNA replication.

Compensation by E2F3 in the E2f1-/- brain

The observation of cell cycle events in E2f1-/- neurons in the absence of any obvious behavioral or anatomical phenotype prompted us to ask whether one of the other E2F family members might be compensating for the role of E2F1 after its germline deletion.

Previous work has shown that the levels of E2F3 often increase in E2F1 depletion (Kong et al., 2007). We therefore assayed, by Western blot, the levels of other E2F family members in homogenates of P30 E2f1-/- brain. The levels of E2F2 do not change in the

E2f1-/- brain (data not shown). As shown in Figure 4 J and K, however, the levels of

E2F3 are increased. It is unlikely that this increase in E2F3 is responsible on it own for

the neuronal cell cycle activity we see. We base this on our observation that the levels of

E2F3 are not altered in the E2f1+/- heterozygote despite the fact that the neurons in the

brains of these animals are as susceptible as those in the homozygote to initiation of cell cycle activity.

Age of onset of the cell cycle events in the E2f1-/- brain

The high prevalence of the CCEs in the one-month E2f1-/- brain raises the possibility that

these nerve cells had never left the cell cycle and thus represent a developmental problem

rather than a problem of cell cycle regulation in the postnatal CNS neuron. To address

this question, we examined E2f1+/+; E2f1+/-; E2f1-/- mice at different ages from birth to adulthood. We used double-labeled immunohistochemistry to detect PCNA and cyclin A

in those mice brains. As late as P10 we found no evidence of ectopic neuronal CCE in

either cerebellum or neocortex. This was the case for all three genotypes (Figure 6). In

these younger animals, the robust mitotic activity in the cells of the external granule cell

layer (EGL) of the cerebellum serves as a positive control. In this mitotically active layer

we found strong PCNA and cyclin A staining. There were also some positive cells in the

internal granule cell layer at P10 (Figure 6A-C), but these likely reflect the normal CCEs

of granule cells during target related cell death (Herrup and Busser, 1995). By P20,

however, there was a dramatic increase of PCNA and cyclin A positive Purkinje cells in

both the E2f1-/- mutant and the heterozygote (Figure 6D-F). The number of positive cells was comparable to that found in P30 brain (Figure 3D, F). The subcellular localization of

PCNA and cyclin A staining was also same as that found in the P30 mice – PCNA in the nucleus and cyclin A in the cytoplasm.

In neocortex, the picture was similar to that found in the cerebellum. At P10, we could

see some PCNA and/or cyclin A positive cells in all three genotype, but these did not

appear to be neuronal cells (as assessed by their nuclear morphology). Furthermore, in

these cells, cyclin A was nuclear in subcellular localization as it should be in normal

cycling cells (Figure 7A-C). By P20, both mutant and heterozygous mice had abundant

numbers of PCNA and cyclin A double positive neuronal cells. In these cells, the

localization of the protein was the familiar pattern of PCNA in the nucleus and cyclin A

in the cytoplasm including some process (Figure 7E, F). We found no PCNA or cyclin A

immunostaining in wild type neuronal cells (Figure 7D).

As discussed above, the appearance of these cell cycle proteins is not a time-limited developmental event. PCNA and cyclin A are found in neurons of both the cerebellum and cortex of E2f1-/- animals as old as one year (Figures 8D, E). As at one month, no cell cycle positive cells were detected in the control brain. The strong suggestion of these images is that the ‘cycling’ cells of the E2f1-/- brain maintain abnormally high levels cell cycle proteins throughout their life. The normal size of the adult brain and the apparently

normal neuronal density suggests that these cells neither progress to finish division, thus

increasing neuronal cell number, nor die. To determine whether the ‘cycling’ cells

suffered consequences from the ectopic protein expression, we examined the dendrites of

the E2f1-/- Purkinje cells closely using calbindin. At low magnification, the calbindin

staining of the E2f1-/- Purkinje cells (Figure 8F) looks less intense than that in wild type

(Figure 8E). At high magnification in the confocal microscope, there is a clear loss of

dendritic mass that appears to be accompanied by a decrease in the density of the

dendritic spines found on the tertiary branches (Figure 8G-J).

Cell cycle events in cultured E2f1-/- primary neurons

Studies of the ectopic appearance of neuronal cell cycles have shown that they can be

triggered by extrinsic events such as the failure of trophic support (Herrup and Busser,

1995; Park et al., 1997a) or by intrinsic problems such as a compromised cell cycle

checkpoint system (Yang and Herrup, 2005) or both (Lipinski et al., 2001; MacPherson et

al., 2003). Since the activation of cell cycle events in an E2F1-deficient brain is

unexpected, we sought to determine whether this was an intrinsic, cell autonomous response of the mutant neurons or instead a response to an extrinsic failure elsewhere in the animal. Cultures of dissociated E16.5 neurons were plated in minimal medium

(Neurobasal) at relatively low density on poly-L-lysine coated coverslips. Identical cultures were prepared from wild-type and homozygous E2f1-/- embryos. Replicate coverslips were analyzed every 5 days. For each time-point, BrdU (10µM) was added to the medium 5 days before fixation in order to assess DNA synthesis. Cultures were stained by immunocytochemistry for either PCNA expression or BrdU incorporation as well as either TuJ1 or Map2 as neuronal cell type markers.

In vivo (Figure 7) cell cycle events appear in cortical neurons between P10 and P20. For neurons harvested at E16.5, these dates are the equivalent of 13 and 23 days in vitro

(DIV). In cultured neurons, at 5 DIV, there is no neuronal PCNA expression apparent in any culture and only a small amount of BrdU incorporation, which we believe is a reflection of residual mitotic activity from the newly harvested ventricular zone cells in

wild-type cycle (Figure 9A). The neurons in the wild-type cultures remain mitotically

inactive for the entire culture period (Figure 9 C, E). In the E2f1-/- cultures, however, significant amounts of PCNA synthesis and BrdU incorporation are detected at 5, 10 and

15 DIV (Figure 9B, D, F, G). These are the temporal equivalents of days P2, P7 and P12 in vivo. Thus in dissociated cultures the appearance of cell cycle events is triggered by a factor intrinsic to the mutant neurons themselves, but the timing is set by an extrinsic cue, which appears on an accelerated time scale in vitro. Cell counts, performed on cultures harvested at several different times in vitro, revealed no significant loss of total TuJ1- positive cells.

Discussion

Mature neurons represent a population of differentiated cells that are believed to have

exited from the cell cycle forever once they have left the ventricular or subventricular

zone and begun differentiation. Yet ectopic cell cycle events in mature neurons do occur

and are related to neuronal degeneration in human neurodegeneration diseases such as

Alzheimer’s (Nevins et al., 1997; Busser et al., 1998; Vincent et al., 1998; Yang et al.,

2001), amyotrophic lateral sclerosis (Ranganathan et al., 2001) and others. Abortive

neuronal cell cycle events (CCEs) have also been found in mouse neurodegenerative

models. These include the retinoblastoma knockout mouse (Jacks et al., 1992; Lee et al.,

1992), T-antigen transgenic mice (al-Ubaidi et al., 1992b; al-Ubaidi et al., 1992a;

Feddersen et al., 1992), as well as in granule cells in staggerer and Lurcher mice (Herrup and Busser, 1995). In other cases the neurons in question engage in cell cycle re-entry but no neuronal cell loss is reported (Yang and Herrup, 2005; Yang et al., 2006). In the aggregate, the data indicate that neurons need to constantly hold their cell cycle in check in order to sustain their normal function and possibly their viability.

In a typical cycling cell, E2F1 helps to control the G1/S transition. When it is

deregulated, by over-expression or unscheduled release from its binding to RB, it can

initiate cell cycle re-entry both in vivo and in vitro. Mutation of E2f1 can reduce ectopic

S-phase entry and apoptosis in Rb-null neurons in the embryonic CNS (Tsai et al., 1998).

Our results suggest that E2F1 plays a role in the regulation of the cell cycle in CNS neurons as well, but the direction of its influence – cell cycle suppression – appears reversed from that found in other systems.

Cultured E2f1-/- granule cells have been reported to be slower to die when exposed to low

K+ (O’Hare et al, 2000), toxic concentrations of the Aβ peptide (Giovanni et al., 2000) or kainite (Smith et al., 2003b). These findings of a protective influence of the E2F1- deficient state are in contrast to our findings in staggerer and Lurcher mice. Here, target- related granule cell death is preceded by a well characterized ectopic S phase entry, yet deletion of E2F1 in either mutant had no effect on either granule cell cycle re-entry or cell death. The different conclusions arrived at in these reports merit consideration. One obvious difference is the nature of the assays. All other tests examined granule cells in vitro while we analyzed the same cells in vivo. Also, the triggers for granule cell death in the various reports are different. In staggerer and lurcher mice, granule cells die because of a loss of trophic support from Purkinje cells. The other studies used specific toxins

(often at high concentrations) and this might well engage a different cell death pathway.

As it has been shown that the function of E2F1 is context dependent, this difference could be crucial. In vitro, for example, E2F1 has been shown to either increase or decrease oncogenic transformation, depending on the conditions of the assay (Johnson et al., 1994;

Melillo et al., 1994; Lee and Farnham, 2000).

Our finding of differentiated neurons with cell cycle related protein expression and

significant DNA replication in vivo is quite consistent with the original reports on the

E2f1-/- knockout mice (Field et al., 1996; Yamasaki et al., 1996). Both groups highlight the different roles played by E2F1 in different tissues. In exocrine tissues such as pancreas and salivary gland it appears to act as a tumor suppressor, while in other tissues

it functions more as an cell cycle promoter (Field et al., 1996; Yamasaki et al., 1996).

And the mice die with significant numbers of tumors in their bodies

In the A-T mouse model, only about 12% of the ATM-deficient Purkinje cells and a few

striatal neurons engage in cell cycle activity (Yang and Herrup, 2005), while in the

E2f1-/- mouse, nearly half of the Purkinje cells show cell cycle protein re-expression.

Despite these quantitative differences, there are similarities in the cell cycle phenotypes of the E2f1 and atm mutants that deserve special notice. There is a striking similarity in the time window during which cell cycle re-entry occurs in both the atm-/- and E2f1-/-

mouse. Both begin between P10 and P20 and quickly rise to a percentage involvement

that remains relatively constant throughout the life of the animal. An additional similarity is that with both the atm and E2f1 mutations ectopic cell cycle events are found in heterozygous and homozygous neurons. This implies that the tumor suppressor role of both genes functions as a recessive trait, possibly indicating a threshold of activity that is needed to fully suppress the neuronal cell cycle. The similarities between the neuronal phenotypes of the two mutations may have even deeper roots as studies have shown that

E2F1, like ATM is involved in DNA repair and cell cycle checkpoint functions. The level of E2F1 is increased when cells are treated with DNA damaging agents, and recent studies have shown that the stability of E2F1 is highly sensitive to phosphorylation by either ATM or checkpoint kinase 2 (Chk2) (Lin et al., 2001; Stevens and La Thangue,

2003). This suggests a model where ATM acts in part to regulate E2F1 stability in normal neurons. When the atm gene is deleted, the levels of E2F1 are reduced and in cells where the effective levels drop sufficiently (by about half, as suggested by the

heterozygote data) the nerve cells re-enter the cell cycle. Deletion of E2f1 itself simply

expands the extent of the ectopic cell cycle events. It would be predicted the levels of

E2F1 would be reduced in cell cycle positive atm-/- neurons.

FISH staining in cerebral cortex of the E2f1-/-mice showed that about 20% of the cells were hyperploid. This confirms a true cell cycle was at least begun in those postmitotic neurons. We note, however, that about 5% of cells in the wild type Layers II and III had three or four hybridization signals. This is not consistent with previous work on AD mouse models (Yang et al., 2006) or the results from human pathological material. In these earlier studies, wild type neurons displayed no multiple hybridization signals.

There are several differences between our systems with AD mouse model. First, the genetic background is different. Our E2f1 mice are maintained on a mixed (C57BL/6J x

129/S4) background. The earlier study used C57BL/6J as control. The age of the animals investigated was different as well – prepuberal (P30) in the current study, adult or aged mice (6-12 months) in Yang et al. The findings are in agreement, however, with those of Rehens et al. who found a comparable level of hyperploidy in their analysis

(Rehen et al., 2001)

Among the various E2F family members, E2F1, E2F2 and E2F3a are grouped because of their structural and functional similarities. While there are no changes in the levels of

E2F2 protein, the level of E2F3 was increased. This suggests that E2F3 may compensate for E2F1 in the mutant neurons, allowing them to function normally even though they are

involved in the early stages of a cell cycle process. This pattern of E2F3 filling in for

E2F1 is consistent with the findings of Kong et al (Kong et al., 2007). But elevated E2F3

levels are clearly not sufficient to restore the cell cycle suppression function of E2F1 in

postmitotic neurons. While it is possible that the ectopic cell cycle events we saw in

E2F1 deficiency mice might be directly caused by the increase of the level of E2F3, this

hypothesis is incompatible with our observation that the levels of E2F3 appear unchanged

in the heterozygous E2f1+/- brain even though the CCEs are fully expressed.

Cell cycle re-entry in cultured E2f1-/- primary neurons indicates that the ectopic cell cycles are a cell autonomous response to the deficiency of E2f1. However, the earlier initiation of the CCEs in vitro argues that E2F1 collaborates with some extrinsic factor(s) that normally holds the cell cycle in check for almost a week longer in vivo than in vitro.

We conclude from this that an ectopic cell cycle event is regulated by both intrinsic and extrinsic mechanisms, but the extrinsic factors can not compensate intrinsic failure in the end.

In summary, our findings indicate that E2F1 functions as a cell cycle suppressor in CNS neurons, and other E2F family members cannot compensate for this non-canonical role.

These findings help explain why, despite the predictions of the earlier tissue culture experiments, the absence of E2F1 does not protect granule cell neurons from target- related cell death in staggerer and lurcher mice. Tissue culture results suggest that while the timing of the cell cycle initiation might be advanced slightly, the failure to suppress the cycle is intrinsic to the mutant neurons. Cell cycle re-entry in E2f1-/- neurons is begun in the second and third postnatal weeks in vivo but then stalls and several proteins,

typically found only in actively cycling cells, continue to be expressed for many months with few harmful affects apparent in neuronal structure or function.

Figures

Figure 1: Deletion of the E2f1 gene does not protect granule cells from target-related cell death in either staggerer or lurcher cerebellum.

Sagittal sections of P30 cerebellum were stained by cresyl violet. Note the thinning of the granule cell layer of staggerer (A) and lurcher cerebellum (B) due to the death of the most of the granule cells. Double mutants with E2f1, either E2f1-/-; Rora sg/sg (C) or

E2f1-/-; Grid2+/Lc (D) leads to no change in the overall morphology of the cerebellum.

Scale bar: 200 µm

Figure 1: Deletion of the E2f1 gene does not protect granule cells from target-related cell death in either staggerer or lurcher cerebellum.

Figure 2: Deletion of the E2f1 gene does not suppress ectopic cell cycle events in either staggerer or lurcher cerebellum.

Sagittal sections of P30 cerebellum from different genotypes were double labeled with

the antibodies for the cell cycle marker PCNA (green) and the Purkinje cell marker

calbindin (red). A-C: E2f1+/+; Rorasg/sg. D-F: E2f1-/-; Rorasg/sg. G-I: E2f1+/+; Grid2+/Lc. J-

L: E2f1-/-; Grid2+/Lc. The pattern of PCNA staining in the granule cell layer of double

mutant mice is similar with that seen in single mutant animals. In the Purkinje cells,

however, the PCNA staining is much brighter in double mutant mice (D and J) than in

single mutant animals (A and G). Scale bars: 25 µm

Figure 2: Deletion of the E2f1 gene does not suppress ectopic cell cycle events in either staggerer or lurcher cerebellum.

Figure 3: Ectopic cell cycle protein expression in E2f1-deficient Purkinje cells.

Mice of three E2f1 genotypes were examined by immunostaining. A, B: E2f1+/+; C, D:

E2f1+/-; E, F: E2f1-/-. A, C, E: P30 sagittal cerebellar sections were labeled with the

Purkinje cell marker, calbindin (red), and the cell cycle marker, PCNA (green). No

PCNA staining is found in wild type animals (A). However, PCNA is expressed in the

nuclei of E2f1+/- and E2f1-/- Purkinje cells (C, E). Counts of the percentages of calbindin-

positive cells that were also PCNA-positive in three E2f1 genotypes are represented in

the histogram showed in Panel G. Panels B, D, and F are double immunostained with

PCNA (green) and cyclin A (red). Both PCNA and cyclin A staining were found in the

Purkinje of E2f1 heterozygotes (D) and mutant (F) mouse brains. No cyclin A or PCNA

was expressed in wild type animal (B). Scale bar = 25µm

Figure 3: Ectopic cell cycle protein expression in E2f1-deficient Purkinje cells.

Figure 4: Reappearance of cell cycle proteins in E2F1-deficient neocortex.

Sagittal sections of P30 mouse brain from three different E2f1 genotypes (E2f1+/+; E2f1+/-;

E2f1-/-) were double labeled with PCNA (green) and cyclin A (red) antibodies. As in cerebellum, PCNA and cyclin A were re-expressed in both the E2f1+/- and E2f1-/- brain

(D-F E2f1+/- and G-I E2f1-/-). There was no cell cycle protein detected in wild type animal (A-C). The subcellular localization of these two cell cycle proteins was the same as seen in cerebellum – PCNA in the nucleus and cyclin A in cytoplasm. Scale bar =25

µm.

Figure 4: Reappearance of cell cycle proteins in E2F1-deficient neocortex.

Figure 5: DNA replication occurs in neurons of the E2f1+/- and E2f1-/- brain, as revealed by fluorescent in situ hybridization (FISH).

A, D: E2f1+/+; B, E: E2f1+/-; C, F E2f1-/-. Evidence for DNA replication was readily observed in both Purkinje cells (A-C) and cells of neocortex (D-F). In wild type animals

(A, D) there were two or fewer hybridization signals while in either heterozygous (B, E) or homozygous (C, F) mutant mice, there were many cells with three or four hybridization signals (insets in B, C, E, and F: high magnification of the nuclei indicated with the arrows). Confocal microscopy reveals that the spots of hybridization are clearly within the nuclei. Wild type cells are illustrated in the left two figures while E2f1 mutant cells are shown in the right two figures. Counts of nuclei (performed as described in

Methods) show that there are many more cells with three or four spots in the E2f1-/- brain

(~20%) than that in wild type (~5%). ML: molecular layer. PC: Purkinje cell layer. IGL: internal granule cell layer. Scale bar = 10µm

Figure 5: DNA replication occurs in neurons of the E2f1+/- and E2f1-/- brain, as revealed by fluorescent in situ hybridization (FISH).

Figure 6: Ectopic cell cycle events in the developing cerebellum of E2F1-deficient mice.

Brains of P10 (A-C) and P20 (D-F) animals of three different genotypes were stained

with PCNA (green) and cyclin A (red) antibodies. A, D: wild type brain; B, E: E2f1+/-; C,

F: E2f1-/-. No cell cycle events are apparent in the young animals (A-C) even though the mitotic cells in the external granule cell layer serve as a positive control. By P20, PCNA and cyclin A stained many Purkinje cells in both E2f1+/- and E2f1-/- mice (but not in wild type controls). Scale bar = 25 µm.

Figure 6: Ectopic cell cycle events in the developing cerebellum of E2F1-deficient mice

Figure 7: Ectopic cell cycle events in the developing neocortex of E2F1-deficient mice.

P10 (A-C) and P20 (D-F) neocortical cells were stained with antibodies to cyclin A (red)

and PCNA (green). A, D: wild type brain; B, E: E2f1+/-; C, F: E2f1-/-. PCNA and cyclin

A were not expressed in any neuronal cells at P10 while by P20 PCNA and cyclin A were positive in both E2f1+/- and E2f1-/- brain. The subcellular localization of the cell cycle proteins was same as P30 – PCNA was nuclear and cyclin A was cytoplasmic. Scale bar

= 25 µm.

Figure 7: Ectopic cell cycle events in the developing neocortex of E2F1-deficient mice.

Figure 8: Cell cycle protein re-expression in one year old E2f1-/- brain.

Wild type (A, C, E, G, I) and E2f1-/- mutant (B, D, F, H, J) brains were immunostained to examine cell cycle activity and Purkinje cell dendritic structure. PCNA (green) and cyclin A (red) antibodies revealed that the mutant Purkinje cells remained robustly stained for both cell cycle proteins (B, D) while both were absent in comparable regions of wild type controls. A, B: cortex; C, D: cerebellum. Calbindin staining (E-J) reveals the fine details of the Purkinje cells. Even at one year of age, wild type Purkinje cells (E,

G) retain richly branched and densely spined dendritic arbors while the arbors of the

E2f1-/- dendrites (F, H, J) were clearly less well developed. G-J: confocal micrographs.

Scale bar = 25 µm in A-F; 10 µm in G-J.

Figure 8: Cell cycle protein re-expression in one year old E2f1-/- brain.

Figure 9: Ectopic cell cycle events in primary neurons from E16.5 mouse neocortex.

Neurons were cultured for 5 (A, B), 10 (C, D) and 15 (E, F) days in vitro (DIV) and

fixed. BrdU was added to the culture medium for 5 days prior to fixation. Fixed cells

were double-labeled with BrdU (green) and TuJ1 (red) antibodies. A, C, E: wild type; B,

D, F: E2f1-/-. In wild type cultures, few neuronal (TuJ1+) cells were positive for BrdU staining at 5 DIV while none were present at 10 or 15 DIV. In the E2f1-/- cultures, by contrast, there were substantial numbers of such cells at all three time points, with an apparent peak in DNA synthesis between 5 and 10 DIV. shown in panel G is the percentage of BrdU positive neurons (Tuj1positive cells) at 5, 10, 15 days of primary

neuron culture from E2F1 wildtype and E2F1 knockout mice. Scale bar = 100µm.

Figure 9: Ectopic cell cycle events in primary neurons from E16.5 mouse neocortex.

Chapter 3: Nuclear export of CDK5 is required for cell cycle re-entry in neuronal cells stressed by β- amyloid

Abstract

CDK5 is a non-traditional CDK that is primarily active in post-mitotic neurons. Its best

known substrates are cytoskeletal proteins. Less appreciated is its role in the maintenance of the neuronal post-mitotic state. We report here that Aβ treatment of neurons leads to loss of nuclear CDK5. Concomitant with nuclear export is an increase in cell cycle re- entry, which subsequently leads to death. Blocking CDK5 nuclear export abolishes cell cycle re-entry. Data from human and mouse tissues recapitulate these results. To explore the role of hyperphosphorylated tau in Aβ-induced neuronal death we used pharmacological and genetic manipulations. The data reveal that CDK5 is sufficient for tau phosphorylation but not necessary. Further, we provide evidence that cells with hyperphosphorylated tau are not irretrievably fated to die. The data suggest that loss of

nuclear CDK5 leads to a cell cycle related neuronal death that seems unrelated to tau

hyperphosphorylation.

Introduction

Cyclin dependent kinase 5 (CDK5) is a proline directed serine/threonine kinase.

Structurally, it is similar to cdc2 (CDK1) (Hellmich et al., 1994), but functions differently

from traditional cyclin dependent kinases (for review see (Dhavan and Tsai, 2001)). For

example, CDK5 does not have a cyclin as its activating partner. Instead, one of a unique pair of proteins, p35 or p39, binds to CDK5 and activates it (Lew et al., 1994; Lew et al.,

1995; Lee et al., 1996). Another example of the atypical nature of CDK5 is that, despite its membership the CDK family, it has no established role in a normal cell cycle. Indeed, its most prominent roles have been linked to developmental processes of cell differentiation and migration (Ohshima et al., 1996; Gilmore et al., 1998), as well as to specialized functions in neuronal cell synapses and analogous roles in other differentiated cells (Angelo et al., 2006). The paradoxical characteristics of CDK5 were emphasized yet again by recent observations from our lab suggesting that, unlike the cell cycle-promoting functions of its CDK relatives, CDK5 functions in normal neocortical neurons to hold the cell cycle in check (Cicero and Herrup, 2005). This is particularly intriguing because we

and others have recently shown a close association between neuronal cell death and

unscheduled cell cycle events, including DNA replication in several types of

neurodegenerative diseases, including Alzheimer’s disease (Vincent et al., 1996; Jordan-

Sciutto et al., 2003; Ranganathan and Bowser, 2003; Yang and Herrup, 2005).

Alzheimer Disease (AD) has a late age of onset and is characterized pathologically by

high densities of senile plaques and neurofibrillary tangles, mainly in hippocampus and

frontal cortex. In addition to the plaques and tangles, there is also significant neuronal

cell loss in different brain regions, including hippocampus, the dorsal raphe and locus

coeruleus (D'Amato et al., 1987; Whitehouse, 1988; Zweig et al., 1988, 1989). The senile

plaques are formed by high molecular weight aggregates of the β-amyloid peptide (Aβ).

The basic enzymology underlying the generation of Aβ, including the proteases involved, has been well worked out (Selkoe, 2000, 2001). The neurofibrillary tangles are formed by the aggregation of paired helical filaments of hyperphosphorylated tau (Grundke-Iqbal et al., 2006; Iqbal and Grundke-Iqbal, 2006). Tangle densities in limbic and frontal cortex correlate with the observed cognitive impairment better than the deposition of β -amyloid

(e.g., (Arriagada et al., 1992). With over 70 potential phosphorylation sites on the tau protein, it is small wonder that the specific pathways from tau to tangles to neuronal death are not known in any detail. It seems likely, however, that CDK5 has a significant role to play in the process (Monaco, 2004).

In spite of its positive role in promoting differentiation and migration in developing brain, there is evidence that CDK5 might have a negative influence in the adult, aiding tangle formation in AD and other diseases (Patrick et al., 1999; Augustinack et al., 2002; Cruz et al., 2003; Hamdane et al., 2003; Noble et al., 2003; Hung et al., 2005; Zheng et al.,

2005). The CDK5 activators, p35 and p39, can be cleaved to the more active p25 and p29 respectively (Lee et al., 2000). The smaller peptides appear to increase CDK5 activity because the half-life of CDK5/p25 (or CDK5/p29) is greater than that of the unmodified

CDK5/p35 complex (Lee et al., 2000). It has been proposed that, during the course of AD, the calcium dependent protease, calpain, promotes p35 cleavage. This then leads to

increased CDK5 activity and hyperphosphorylation of tau protein ultimately leading to

the neurofibrillary tangles. Consistent with this idea, the p25 peptide has been reported to

be elevated in AD brain (Patrick et al., 1999). Further p25 over-expressing transgenic

mouse lines have significant neuronal tau hyper-phosphorylation (Ahlijanian et al., 2000;

Van den Haute et al., 2001; Bian et al., 2002; Cruz et al., 2003; Noble et al., 2003).

However, the activity of other kinases such as Gsk3β is also increased (Noble et al.,

2003). Thus there remains a certain level of uncertainty as to which kinase plays the main role in Aβ-related tau hyper-phosphorylation (Bian et al., 2002; Hallows et al., 2003;

Tandon et al., 2003; Giese et al., 2005; Hallows et al., 2006; Plattner et al., 2006) and neuronal death.

In the current report we have used 3 experimental paradigms. The first is cultured primary neurons from mouse embryonic neocortex. The second is human pathological material from individuals who have died with AD. The third is the AD mouse model known as R1.40. This line of mice carries the entire human APP gene including tens of kilobases of flanking 3' and 5' sequences on a yeast artificial chromosome. The transgene has been engineered to contain the dominant early-onset Swedish mutation –

K670M/N671L (Lehman et al., 2003). R1.40 mice develop classic plaques in cortical regions beginning at 13 postnatal months of age (Kulnane and Lamb, 2001). The study of

Yang, Varvel et al. Yang, 2006 revealed that cell cycle events also develop in this mouse model, and importantly, these events occur 7 months prior to the first Aβ deposit, in an anatomical pattern that mimics that found in the human condition. Against these various backdrops we have examined the behavior of CDK5 protein in Alzheimer disease like

situations. Our results indicate that CDK5 is sufficient but not necessary for Aβ induced tau hyper-phosphorylation, while Gsk-3β appears to play a major role as a tau kinase. We also report that CDK5 exits the nucleus of cultured neocortical neurons after Aβ treatment and that this occurs prior to, or concomitant with, the initiation of unscheduled cell cycle events. A similar nuclear exit is observed in vivo in both human and mouse AD.

Finally, in vitro, cell cycle re-entry is blocked by inhibition of CDK5 export from nucleus.

In the aggregate, the results suggest that CDK5 has an unexpected role as a negative cell cycle regulator in the post-mitotic cortical neuron.

Experimental Procedures

Animals. A colony of Cdk5+/-mice was maintained on a mixed (C57BL6/Jx129/S1) background (Gilmore et al., 1998). Homozygous Cdk5-/- mice are not viable, so mutant

embryos were produced by intercrossing Cdk5+/- mice. Timed pregnancies were

established from these mating; the date of appearance of a vaginal plug was considered embryonic day 0.5. Embryos were taken at embryonic day 16.5 (E16.5) for either cortical cultures or histology. Genotyping of embryos was performed by PCR of embryo tail

DNA as previously described (Cicero and Herrup, 2005). The primers used were

CDK5F1 (ATTGTGGCTCTGAAGCGTGTC), CDK5R1

(CTTGTCACTATGCAGGACATC) and PGK1 (TCCATCTGCACGAGACTAGT). All

animal procedures were carried out in accordance with and Case Western Reserve

University IACUC standards. The animal facilities at Case Western Reserve University

Medical School is fully AAALAC accredited.

Primary Neuronal Cultures

Embryonic cortical neurons were isolated by standard procedures. Pregnant dams were

sacrificed by cervical dislocation and the uteri dissected and placed on ice during the

embryo harvest procedure. For CDK5-deficient cultures, all embryos from a Cdk5+/- x

Cdk5+/- mating were treated separately; DNA was isolated from each embryo's tail for

genotyping. Embryos were collected in ice cold PBS-glucose, and the cortical lobes were

dissected out. Meninges were removed and the cortices were placed in 1X trypsin-EDTA

for 15 minutes at 37ºC. The tissue was removed from the trypsin solution and placed in

DMEM with 10% FBS to inactivate the trypsin, followed by transfer to Neurobasal

media supplemented with B-27, penicillinstreptomycin (1X), and L-glutamine (2mM).

Tissue was triturated 10X through a 5mL pipet and allowed to settle to the bottom of a

15mL conical tube for approximately one minute. Cells in solution above the tissue pellet

were removed and used in all subsequent procedures. Surviving cells stained with Trypan

Blue; live and dead cells were counted separately. Cells were plated on a substrate of poly-L-Lysine (0.05mg/mL) and laminin (5ug/mL) coated on plates or glass coverslips.

Cells were grown in 24 well plates at a density of 50,000 cells/well. All cultures were grown for a minimum of 5 days in vitro (DIV) before any treatment. To assess cell cycle activity, medium was changed to new medium containing 10uM BrdU. After 24 or 48 hours, cultures were fixed with 4% paraformaldehyde in phosphate buffer at room temperature for 30 minutes, then washed and stored in PBS. All experiments were performed on a minimum of three litters; each condition was examined in triplicate.

Pharmacological inihibition of nuclear export was done with 10uM leptomycin B added to cultures with Aβ. Inihibition of CDK5 by the selective CDK5 inhibitor roscovitine was done at a 10uM concentration added at time of Aβ treatment. 15 mM lithium was used to inhibit GSK-3β at time of Aβ treatment.

Preparation of b-amyloid: The 42 amino acid form of the beta-amyloid peptide

(Aβ1-42) was synthesized by American Peptide and re-constituted in sterile double distilled H2O (ddH2O). After re-constitution, Aβ was allowed to fibrillarize at 37º C for a period of 5 days. Before use, the solution containing the Aβ was vortexed and diluted to the equivalent of 10uM into culture media with additional vortexing.

Immunocytochemistry. Cells were rinsed once with PBS and then exposed to

4% paraformaldehyde in 0.1M phosphate buffer for 30 minutes at room temperature,

followed by three rinses with PBS. Immunohistochemistry of cell cultures was done

without antigen retrieval. For BrdU labeling, DNA was hydrolyzed by exposing the cells

to 2N HCl for 10 minutes. Specimens were then neutralizing in 0.1M sodium borate (pH

8.6) for 10 minutes and then rinsed extensively in PBS (3X) for 45 minutes before

treatment with blocking reagent. Non-specific antibody binding was blocked by exposing

the fixed cells to 5% normal goat serum in 0.1% Triton-X for 1 hour before application of

the primary antibody.

Antibody concentrations used for cell culture were: mouse anti-PCNA (Santa Cruz)

1:1000, rat anti-BrdU (Abcam) 1:100, rat anti-BrdU (Accurate) 1:6, mouse anti-Map-2a/c

(Sigma) 1:500, mouse anti-Tuj1 (Covance) 1:1000, rabbit anti-caspase-3 (Cell Signaling

Technologies) 1:500, rabbit anti-CDK5 (Santa Cruz, C-18) 1:500, rabbit anti-p35 (Santa

Cruz, C-19) 1:500, mouse anti-PHF1 (PHF1, Endogen Bioclear) 1:8000, mouse antiphosphotau (AT8, Endogen Bioclear) 1:1000. Secondary antibodies used were: Goat anti-rat Rhodamine (Jackson Labs) 1:300, and Goat anti-mouse Alexa 488 (Molecular

Probes) 1:1000, Goat anti-rat LRSC (Jackson Labs) 1:500, Goat anti-rabbit FITC

(Jackson Labs) 1:500. Cells were counterstained with 1ug/mL DAPI.

Human tissue: All human AD and age-matched control autopsy brain tissues were

obtained from the Neuropathology Core of the Alzheimer's Disease Research Center of

Cleveland (P50 AG08012). Brain tissue from 8 pathologically confirmed cases of AD was obtained. The specimens had been from whole brains routinely fixed in 10% formalin. The average of post-mortem interval time was 12.5 hours. Six age-matched controls were pathologically verified as free of central nervous system disease. The average of the postmortem interval time for the control tissue was 11 hours. All of the specimens used in this study were embedded in paraffin, sectioned at 10 μm.

Sections were deparaffinized with xylene and re-hydrated through graded ethanol to water. The sections were soaked in 0.3% hydrogen peroxide in methanol for 20 min to remove endogenous peroxides activity, rinsed in Tris-buffered saline (TBS), and pretreated in a solution of 0.1M citrate buffer (PH7.5) heated to 90-960C for 10 min.

Sections were cooled and rinsed in TBS. Slides were incubated in 10% goat serum in

PBS at room temperature for 1 hours to block non-specific binding. After overnight 40oC incubation with primary antibodies (mouse PCNA 1:100; rabbit CDK5 1:100; rabbit p25/35 1:100), the sections were rinsed 3 times in TBS before applying the secondary antibody (diluted in blocking solution at 1:300) for 1 hour. Afterwards, rinsed sections were incubated in ABC complex (VectaStain ABC Elite Kit) for 1 hour at room temperature, followed by 3 rinses in TBS. DAB was used as a substrate for visualization of the HRP-tagged ABC reagent, according to the manufacturer’s specifications (Vecta peroxidase substrate DAB Kit, Vector Laboratory). Some sections were counterstained with hematoxylin and all sections were covered with Permount. Immunohistochemical negative control sections are put through the identical staining procedure except for the omission of the primary antibody.

Results

Loss of nuclear CDK5 is concomitant with cell cycle re-entry

To explore the role CDK5 in blocking cell cycle re-entry, we investigated changes in

CDK5 expression after Ab stimulation of primary cortical neuron cultures. We cultured

E16.5 primary cortical neurons for 5 days in vitro (DIV). After 5 DIV, over 95% of the neurons (TuJ1-positive cells) in the untreated cultures were post-mitotic (BrdU-negative)

(Wu et al., 2000; Cicero and Herrup, 2005). In these untreated cells, strong CDK5 expression is visible in the nucleus and peri-nuclear region of the cell (Figure 1A-C). We then exposed cells to fresh medium with or without 10μM Aβ for 24-48 hours. After Aβ treatment, there were many cells in which the localization of CDK5 had changed dramatically. In these neurons, CDK5 immunostaining was reduced or totally lost in the nucleus (Figure 1D-F, arrows) and the distribution of the antigen in the peri-nuclear region appeared clumped. Western blot analysis of whole cultures (not shown) revealed unaltered levels of total CDK5 after treatment suggesting that redistribution rather than degradation was the major change.

The translocation of CDK5 protein from nucleus to cytoplasm led us to ask whether or

not this shuttled protein retained activity. Unfortunately, no in situ procedure to

specifically determine CDK5 activity has yet been described. To answer the question

indirectly, we immunostained our cultures for the CDK5 activator protein(s), p25/35,

using a polyclonal antibody that detected both forms. In untreated cells, the staining of

p25/35 was mainly located in the nucleus and peri-nuclear region of TuJ1 positive

neurons (Figure 1G-I), roughly similar to the distribution of CDK5. Unlike CDK5,

however, there was no change in the localization or distribution of p25/35 after Aβ

treatment (Figure 1J-L). This indicates that CDK5 and its activator proteins are separated after Aβ exposure and that each responds differently to challenge with the peptide. The results further suggest that CDK5 may lose its normal kinase activity after Aβ treatment.

To determine whether the localization change of CDK5 was related to cell cycle re-entry, we cultured E16.5 cortical neurons for five days. We then added BrdU to the culture media along with the Aβ treatment. After 24 hours, the cells were fixed and double immunolabeled for the presence of BrdU and CDK5. In untreated cells, there were very

few cells double-labeled by anti-BrdU and CDK5 antibodies (Figure 2A-C). After

Aβ treatment, however, the numbers of BrdU/CDK5 positive cells were significantly increased (Figure 3D-F). In addition, there were always a few non-neuronal cells in the culture incorporated BrdU during the course of the experiment and these served as an internal control. In both untreated and treated cells, CDK5 shifted from nucleus to cytoplasm in the BrdU-positive cells. In Aβ treated wells, BrdU/cytoplasmic CDK5 double-positive cells increased dramatically among the pycnotic (dying) cells (Figure 2E,

F). To confirm the BrdU findings, we used antibody to the proliferating cell nuclear antigen (PCNA) as an independent measure of cell cycle activity. PCNA, a component of the DNA polymerase complex that travels with the replication fork, is widely used as a marker of cells in S-phase. We saw the same pattern as with BrdU when we immunostained cells with mouse anti-PCNA antibody (Figure 2G-L). Together, these data indicate that loss of CDK5 from the nucleus occurs just as cell cycle control is lost in neurons.

Cell cycle re-entry is reduced after inhibiting CDK5 translocation

The correlation of CDK5 translocation with cell cycle activity implies that the two events

are causally linked. To validate this impression, we used the nuclear export inhibitor

leptomycin B (LMB) to block CDK5 export from the nucleus. Leptomycin B (LMB) is a

potent, specific inhibitor of the CRM1/exportin1-dependent nuclear export pathway. We

cultured E16.5 primary cortical neurons 5 DIV, with or without LMB. In all LMB treated

cells CDK5 remained in the nucleus (Figure 3B; compare with untreated cells, Figure

3A). This indicates that CDK5 relies either directly or indirectly on an active CRM1

nuclear export machinery to leave the nucleus. Cultures were then subjected to one of

four different treatments: medium change only, Aβ-only, LMB-only or Aβ plus LMB. All treatments were done in the presence of BrdU to detect cell cycle activity. We then fixed the cultures and stained them with BrdU and CDK5 antibodies.  After 24 hours 25% of

Aβ treated cells were lost (Figure 3C). Aβ only treatment led to a 3-fold increase in the percentage of BrdU-positive cells (Figure 3D). Although there was substantial cell death in LMB-only treated wells (Figures 3C) the number of BrdU positive cells was very low

(Figure 3D). Further, addition of Aβ (Aβ plus LMB treated wells) did not relieve this cell cycle blockage. Virtually all pycnotic cells in these cultures were BrdU-negative (Figure

3E) indicating that those cells did not die through cell cycle related pathway. When we quantified the number of BrdU positive cells that had CDK5 only in the cytoplasm (cyto-

CDK5), we found that more pycnotic cells were BrdU+/cyto-CDK5+ double positive than live cells in Aβ treated cultures (Figure 3E). In the Aβ treated wells, 13% of the pycnotic cells (visualized by DAPI staining), were BrdU+/cyto-CDK5+ (Figure 3E).

Counts of pycnotic BrdU+ cells showed that cyto-CDK5 cells are more abundant than

cells with CDK5 in both the nucleus and cytoplasm (nuc-CDK5), indicating that CDK5’s

localization is key to the cell loss (Figure 3E-F, compare Aβ treated groups). In live cells the opposite is the case: nuc-CDK5 cells are more prevalent among the BrdU+ cells than are cyto-CDK5 cells. However, as might be expected, global blockade of protein export from the nucleus with LMB is eventually toxic to nerve cells. Thus, the pharmacological approach alone is not sufficiently rigorous prove that Aβ induced nerve cell death is permanently blocked by retaining CDK5 in the nucleus. LMB does block CDK5 translocation from the nucleus and, as BrdU was present throughout the treatment period, the data indicate that preventing CDK5 nuclear export blocks the induction of cell cycle events in cultured neurons.

CDK5 localization changes in an AD mouse model

The results presented have implications for neurodegenerative conditions such as

Alzheimer's disease and suggest that the role of CDK5 in disease might be complex with location as well as activity playing a part in the final disease process. This prompted us to examine the relationship between CDK5 localization and cell cycle re-entry in vivo.

We first examined the location of CDK5 in the R1.40 mouse model. As reported, CDK5 is detected throughout the neuropil of the normal adult mouse brain (Gilmore et al., 1998).

Closer inspection reveals that the protein is distributed nearly uniformly between nucleus and cytoplasm (Figure 4 A-H). In R1.40 animals this pattern was largely maintained.

However, in brain regions where cell cycle events had begun, such as hippocampus and frontal neocortex, we found that the neurons that were 'cycling' (PCNA-positive) were

largely devoid of CDK5 protein in their nuclei (Figure 4M-P). As an internal control, we note that the nuclear/cytoplasmic translocation of CDK5 was rarely observed in R1.40 cerebellar Purkinje cells, which are unaffected in human AD and do not develop cell cycle events in the R1.40 mouse (data not shown). We saw little change in neuronal

CDK5 localization (and no cell cycle events) in age matched control mouse brains. This suggests that Aβ-induced CDK5 translocation is related to the cell cycle re-entry.

Analyses of Western blots of brain lysates (not shown) indicate that the level of total

CDK5 protein was largely unaffected in the R1.40.

CDK5 localization changes in human AD

In all stages of Alzheimer's disease, neurons at risk for death show cell cycle related protein expression (Yang et al., 2003) and DNA replication (Yang et al., 2001). We therefore sought to determine the localization of CDK5 in the brain cells of patients who had died with AD and to determine its relationship to the previously reported cell cycle events. As in the wild-type mouse, neuronal CDK5 expression is found ubiquitously in the brain regions we examined. Within most cells from control specimens, the protein is distributed nearly uniformly, although in mainly nuclear and peri-nuclear (cytoplasmic) locations (Figure 5A, C). This is true in both hippocampus and locus coeruleus. In the

AD brain, however, the level of CDK5 was markedly increased with a large part of that increase found in the nucleus (Figure 5B, D). This is different from our findings in treated cell cultures and in the Western analysis of the R1.40 mouse model. By

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immunocytochemical criteria, the majority of cells, which are unaffected by cell cycle

induced loss, continue to express high levels of CDK5. However, we consistently

observed that CDK5 was lost from the nucleus of neurons expressing PCNA (Figure 5F).

Further, as we had observed in cell culture, the staining levels of p25/35 were increased

but their localization remained mainly nuclear (Figure 5G, H). Again, this suggests that

CDK5 and its normal activating binding partners respond differently to the disease

process in the AD brain. Importantly, in all three systems that we examined – primary

neuronal cell culture, the R1.40 APP transgenic mouse model and the human AD brain

neurons that lose their nuclear CDK5 also lose cell cycle control.

CDK5 is not necessary for tau phosphorylation

The data reported here lend themselves to the conclusion that CDK5 functions as a

prosurvival factor in post-mitotic nerve cells. The suggestion would be that, by inhibiting neuronal cell cycle events, CDK5 is protective force acting against the progress of AD.

This perspective is in contrast to a growing interest in CDK5 as a potential pathogenic

force in AD, based on its known role as a tau kinase. Hyperphosphorylated forms of tau are the building blocks of the neurofibrillary tangles, and these are a classic pathological feature of the AD brain. To determine whether CDK5 is serving as an active tau kinase in

our model system, we established dissociated primary cortical neuron cultures from

Cdk5+/+, Cdk5+/-, and Cdk5-/- E16.5 embryos. After five days, cultures were treated with

Aβ for 24h then fixed and immunostained with mouse PHF-1 antibody. This antibody

detects tau phosphorylation at a pair of serine residues: S396 and S404. Both are known

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CDK5 phosphorylation targets (Sakaue et al., 2005) and PHF-1 epitopes are easily found

in neurons of AD patients.

We observed PHF-1 staining in cultured neurons from all 3 genotypes both before and

after Aβ treatment, but the pattern of staining was different for each genotype. The PHF-

1 staining of neurons in the Cdk5-/- cultures was weak and significantly less than that

found in the wild type cultures (Figure 6A). The levels of PHF-1 staining in the

heterozygous neurons was punctuated and appeared intermediate between the Cdk5-/- and

Cdk5+/+ cultures (Figure 6A, C, E). After Aβ treatment, the intensity of PHF-1 staining became stronger in cultures of all three genotypes (Figure 6B, D, and F). In Cdk5-/-

cultures there were indeed fewer PHF-1 positive neurons than in wild type cultures, but

the levels of PHF-1 staining was not visibly different between the genotypes after Aβ treatment.

These data suggest that other kinases can phosphorylate tau in the absence of CDK5 after

a neurotoxic insult. To ensure that our findings were not unique to the S396/S404

residues, we used a second phospho-tau antibody, AT8. AT8 recognizes tau

phosphorylated at sites S199/202, which is also a common tau modification in the AD

brain. In our cultures, the pattern of AT8 immunostaining was same as that for PHF-1.

Before Aβ treatment, AT8 immunostaining of Cdk5-/- neurons was significantly less

intense than that of Cdk5+/+ neurons. After Ab treatment, the level of AT8 was increased

in all three genotypes of neurons to almost equal levels (Figure 10).

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It is clear that Aβ reatment leads to neuronal cell death, but the PHF-1 and AT8 results raise the question of the relationship between phosphorylated tau residues and cell death.

If hyperphosphorylated tau were indeed an important pathway to death then one would expect to find that dying cells were well labeled with phospho-tau antibodies. Yet, after double labeling of our Aβ treated cultures, we found no neurons that were immunopositive for both PHF-1 and activated caspase-3 – a widely used cell death marker (see below). The reduced levels of PHF-1 and AT8 staining in the untreated

Cdk5-/- cultures strongly implicate CDK5 as a natural tau kinase in basal conditions.

However, the responses of both epitopes to Aβ exposure suggest that the enhanced phosphorylation of tau under the stress of toxic concentrations of Aβ1-42 neither requires the presence of CDK5 and nor leads directly to neuronal cell death.

Loss of CDK5 does not protect against neurodegeneration

CDK5 has been proposed to contribute to the death of neuronal cells in AD, but the nuanced role that it plays in tau phosphorylation led us to ask whether similar complexities could be uncovered in its pro-death functions as well. While pharmacological manipulations of wild type cultures tend to support the pro-death role of

CDK5 we reasoned that if this were the entire story then primary cultures of CDK5- deficient neurons should also be resistant to Aβ induced cell loss.

Cortical neurons from E16.5 embryos of all three genotypes were treated with Aβ for

24h, fixed and double-labeled labeled with antibodies against PHF-1 and activated caspase-3. DAPI staining was used to determine total cell number. Aβ treatment resulted

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in more activated caspase-3 positive neurons in the Cdk5-/- cultures than in comparably treated wild type ones (Figure 7A-F). These findings are difficult to interpret directly as

Cicero and Herrup (2005) have shown that the absence of CDK5 is itself a cell lethal

situation for a neuron. Nonetheless, when the number of surviving cells in the Aβ treated cultures was calculated as percentage of cells in the untreated culture, significant cell loss was found in all three genotypes. In wild type cultures, 19% of the cells were lost after 24 hours. By contrast, in heterozygous and homozygous CDK5 mutant cultures,

35% and 50% of the nerve cells were lost during the same treatment. Thus, if anything,

cell loss was greater in knockout cultures than in wild type (Figure 7G, H) and these

results extend even to a situation where the levels of CDK5 are only reduced, not

eliminated (the heterozygotes). The loss of only one allele leads to fewer viable cells after

5DIV (counted by DAPI labeling) even in the absence of Aβ treatment. These results

extend our previous observations that loss of one CDK5 allele leads to a subtle but dose

dependent phenotype both in vitro and in vivo (Cicero and Herrup, 2005). The results

suggest that, just as with tau phosphorylation, CDK5 may be necessary for basal neuronal

survival (fewer of the originally plated neurons survive in the untreated mutant cultures)

but it is not required for Aβ induced cell loss. Dissociated neuronal cultures respond to

the presence of Aβ1-42 and die, even in the complete genetic absence of CDK5.

Aβ-stimulated tau phosphorylation may be GSK-3β dependent.

Aβ exposure, even of Cdk5-/- neurons, leads to tau hyper-phosphorylation on several

different AD-relevant epitopes. This raises the question of the identity of the other

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kinase(s) that perform this function in Cdk5-/- neuronal cultures. GSK-3β is a likely candidate as an alternate tau kinase. We treated E16.5 primary cortical neurons with

Aβ in the presence and absence of lithium – a broad spectrum GSK-3β inhibitor. We also exposed neurons to Aβ in the presence and absence of roscovitine, a CDK5 inhibitor.

After 24h, we fixed cells and immunolabeled cells with the PHF-1 phosphotau antibody to determine tau phosphorylation. As expected, in untreated wells, there was an easily detectable basal level of PHF-1 immunostaining (Figure 8A, E). This basal level was inhibited by lithium (Figure 8C, G). The CDK5 inhibitor, roscovitine, also reduced the basal level of tau phosphorylation (Figure 8B, F), but less so than lithium. This suggests that both GSK-3β and CDK5 help to maintain normal tau phosphorylation. This result is consistent with our findings in Cdk5-/- cultures (Figure 6, 7).

After Aβ treatment, the intensity of PHF-1 positive immunostaining is enhanced (Figure

8I, M) and most of this tau phosphorylation can be blocked by lithium (Figure 8K, O), but not by roscovitine (Figure 8J, N). This indicates that GSK-3β induces the Aβ enhanced tau phosphorylation while CDK5 maintains the basal levels. These data are consistent with the data from cultures of Cdk5-/- neurons, where in the absence of CDK5 untreated neurons express few or no PHF-1 or AT8 epitopes. With the addition of Aβ,

however, Cdk5-/- neurons show an increased phosphorylation of tau at the PHF-1 and

AT8 phospho-epitopes. There were a few neurons that were still PHF-1 positive after

both lithium and roscovitine treatments (Figure 8K, O, L, P). This suggests that there are

still other kinases contributing to the Aβ response of tau (or that we did not achieve a full

block of these kinases by pharmacological means). In addition to lithium, we also used

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two other, more specific GSK3 b inhibitors – SB216763 and SB415286 – to verify our results. Both compounds gave results similar to those obtained with lithium (data not shown).

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Discussion

We have explored the roles of CDK5 as a cell cycle regulator in post-mitotic neurons.

Our findings suggest that CDK5 acts as a cell cycle suppressor under most circumstances

and that this function requires its presence in the nucleus. CDK5 also acts either directly

or indirectly as a tau kinase in basal situations, but under conditions meant to mimic those

of Alzheimer's disease our data suggest that other kinases, notably GSK-3β, are more active in producing hyperphosphorylated tau. Recent work, has indicated that CDK5 can phosphorylate tau at S235 and that this event primes T231 for phosphorylation by

GSK-3β (Sengupta et al., 2006). The relationship of tau phosphorylation events to the cell death process in our model systems remains unclear, but the evidence suggests that it is indirect at best. This is indicated by the fact that most of the caspase-3 positive neurons we examined after Aβ treatment were negative for phospho-tau and vice versa. In vitro, our model system consisted of dissociated E16.5 neocortical neurons grown in defined

Neurobasal™ medium. To mimic the pathogenic conditions of the AD brain, we exposed

5 day cultures to fibrillarized Aβ1-42. This exposure induced both neuronal cell cycle re-

entry and cell death. In the 'cycling' neurons, we found that the localization of CDK5

shifts – very little protein remains in the nucleus while the cytoplasmic protein tends to

be found in large aggregates. This result is consistent with our findings that cultured

Cdk5-/- neurons continue to incorporate BrdU (Cicero and Herrup, 2005), and it suggests

that the key to cell cycle arrest is not the total amount of CDK5, but rather its

subcellular location. To emphasize this point we used the nuclear export inhibitor,

leptomycin B, to block CDK5 translocation. BrdU incorporation was blocked in the

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presence of leptomycin B, consistent with our hypothesis that nuclear CDK5 holds the

cell cycle in check in a mature neuron.

It is of interest that the CDK5 activator protein(s), p25/35, did not exit the nucleus after

Aβ treatment. This suggests either that the anti-mitotic function of CDK5 in mature neurons does not rely on its kinase activity or that the process of cell cycle initiation begins with the dissociation of the initial complex. The former conclusion is similar to that put forward in our previous study (Cicero and Herrup, 2005). The protein complex by which the CDK5 activator protein is imported in the nucleus has been recently defined.

mportin-β -5, and -7 all function to import p35 into the nucleus and CDK5 is not required for this import mechanism (Fu et al., 2007). The corollary to this process may well be that export from the nucleus does not require interaction between CDK5 and its activator protein(s). It remains to be determined whether the nuclear pool of CDK5 includes bound p25/35 in the basal state in differentiated neurons, and whether other proteins are included in a larger macromolecular structure. It should be noted that bovine

CDK5 has been isolated in a macromolecular structure of approximately 670kD from whole brain lysates (Lee et al., 1996). These data, taken together, point to a more complex role for CDK5 in both normal differentiated neurons as well as neurons under toxic stress such as Aβ exposure. While the regulation of CDK5 activity is thought to occur by its interaction with its activator protein (for review see (Hisanaga and Saito,

2003), very little work has been done on the degradation of the CDK5 protein as an additional route of kinase activity control. Preliminary data (not shown) suggests that an increase in CDK5 ubiquitination occurs after toxic insult. It may be that the shift of

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CDK5 to the cytoplasm allows it to act as a tau kinase under stress conditions, and this

activity may be further regulated by ubiquitination. This shift of CDK5 from the nucleus potentially frees other unknown partners from their cell cycle suppressor function.

The exact mechanism by which CDK5 exerts its cell cycle suppression effect is currently

unknown. Possibilities include alterations in cytoskeletal functions as well as direct

action on proteins with cell cycle-specific functions. An example of the latter possibility

is the cell cycle inhibitor, p27. CDK5 can stabilize p27 (Kawauchi et al., 2006), and the

protein has been documented to affect multiple processes in neurons, including cell

migration (Nguyen et al., 2006b), cell cycle suppression and differentiation. In p27-/- mice there is an increase in numbers of later born cortical neurons which ultimately leads to a larger cortex, emphasizing its role in cell cycle inhibition(Goto et al., 2004).

However, because p27 elicits multiple effects on the cytoskeleton and on cell cycle dynamics

(Besson et al., 2004; Nguyen et al., 2006b), as does CDK5, more work is necessary to define the mechanism.

AD mouse models and post-mortem human AD brain sections add an in vivo dimension

to our studies. We found the expression of CDK5 is reduced or lost in the nucleus of

‘cycling’ neurons, such as neuronal nuclei of the frontal cortex and hippocampus, in both

mouse and human AD. No such change in localization is seen in controls or in neurons of

unaffected brain regions. In “non-cycling” neurons in vivo, the expression of CDK5 is as

same as in the neurons in our in vitro cell cultures – most of it is nuclear and perinuclear.

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Although CDK5 localization changes, p35/25 expression remains predominantly nuclear

and does not change significantly either in vivo or in vitro. This is consistent with the

findings of others who have shown that CDK5 and p35/25 can move in and out of the

nucleus independently (Fu et al., 2006).

The other pathogenic feature found AD is the presence of neurofibrillary tangles, which

are formed by higher order aggregates of hyper-phosphorylated tau. The work of Patrick

and coworkers first suggested that in AD it is the presence of the CDK5 activator p25 that

ultimately leads to tau hyper-phosphorylation (Patrick et al., 1999). Many transgenic

mice have been made to determine the role of p25/CDK5 in tau hyper-phosphorylation

(Ahlijanian et al., 2000; Van den Haute et al., 2001; Bian et al., 2002; Patzke et al., 2003;

Sakaue et al., 2005; Hallows et al., 2006; Plattner et al., 2006). The neuronal degeneration seen in some of these models implicates elevated CDK5 activity in AD nerve cell loss. The role of phosphotau itself can only be inferred from the results, however. The findings we report here suggest caution in the final interpretation of the

findings. First, the fact that activated capase-3 and PHF-1 tau are not found in the same

neurons suggests that the presence of hyper-phosphorylated tau is not enough to cause cell loss. Further, the results of Aβ treatment of Cdk5-/- primary cultures suggest that loss

of CDK5 is not protective for Aβ treatment (as predicted by the p25 transgenic

phenotype). While the interpretation of the Aβ induced loss of homozygous Cdk5-/- cells is confounded somewhat by the requirement of CDK5 for normal neuronal differentiation

(Gilmore et al., 1998; Gilmore and Herrup, 2001; Cicero and Herrup, 2005), the

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heterozygote data demonstrate that even with a normal differentiation program (Cicero

and Herrup, 2005), reducing the levels of CDK5 by half provides no protection against

the neuronal cell loss induced by Aβ.

In Cdk5-/- cultures tau phosphorylation at basal levels is significantly reduced from the levels seen in wild type neurons. This suggests that CDK5 is necessary to maintain a basal level of tau phosphorylation. After Aβ treatment of all cultures, tau phosphorylation was enhanced in neurons of all genotypes. Validation of this result comes from our finding that enhanced levels of phospho-tau in wild type cultures could not be blocked by roscovitine. These data suggest that other kinases are responsible for Aβ -induced tau hyper-phosphorylation. Thus, CDK5 may be sufficient for tau phosphorylation, but it is not necessary. Instead, GSK-3β may be the main tau kinase following Aβ

treatment.Three different GSK-3β inhibitors could block almost 90% of tau phosphorylation in our in vitro assay. This is consistent with other work that has shown that CDK5 can act as a priming kinase for GSK-3β on tau (Hallows et al., 2003; Hallows et al., 2006; Liu et al., 2006; Sengupta et al., 2006).

Previous work by our lab and others has shown that neurons at risk for death in AD exhibit cell cycle protein re-expression and DNA replication. Those phenomena are found in both late and early stages of human AD. Further, in AD mouse models, neurons begin cell cycle events in the areas that are known to be the most severely affected in the human disease. And the cell cycle events appear much earlier than the formation of the

Aβ deposits(Yang et al., 2006). This indicates that the neurons detect the stress of

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impending AD much earlier than the beginning of plaque deposition. There is little

known as to why or how neurons are stimulated to re-enter an unscheduled cell cycle

(Wu et al., 2000). It may be that the presence of soluble or oligomeric Aβ leads initially to synaptic loss. Loss of synapses in turn may lead to as stress signal in the neuron, leading to upregulation of CDK5 and an overall change in its homeostasis. In either of these cases, or in others, when the stress overwhelms the protective effect of nuclear

CDK5, it results in its transport to the cytoplasm where its ectopic activity might lead to further neuronal stress. With the loss of nuclear CDK5, the cell cycle is no longer effectively blocked and an unscheduled and ultimately lethal cell cycle process begins.

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Figures

Figure 1: CDK5 localization changes after Aβ1-42 treatment of primary neuronal cultures.

A-C: in untreated cultures of E16.5 wild type cortical neurons, CDK5 (red) is largely but not exclusively co-localized with the DAPI nuclear counterstain (blue). D-F: 48 hours after 10μM Aβ1-42 treatment, CDK5 moves to the cytoplasm of some cells where is

frequently appears clumped (white arrows). G-L: the response to Aβ1-42 treatment of the

CDK5 activator, p35/25 (red), is distinctly different. G-I: in untreated neurons (identified with TuJ1 immunostaining [G]), p35 (H) is largely nuclear. After Aβ1-42 treatment (J),

TuJ1 staining reveals the effect of the degeneration as the neuritic processes begin to fragment. The localization of p25/35, however, remains unchanged (K).

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Figure 1: CDK5 localization changes after Aβ1-42 treatment of primary neuronal cultures

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Figure 2: Cell cycle re-entry is marked by loss of CDK5 from the cell nucleus.

After 5 DIV, E16.5 cultured cortical neurons were exposed to 10μM BrdU for 48h in

either control conditions (A-C, G-I, M) or during treatment with 10μM Aβ1-42 (D-F, J-L,

N). Cells were stained for CDK5 (red) and a cell cycle marker (green) -- either BrdU (DF)

or PCNA (J-L). In untreated cells, we rarely saw BrdU or PCNA positive cells. After

Ab treatment, cells in which CDK5 staining had moved to the cytoplasm were BrdU or

PCNA positive. (D- F, J-L Arrows). This shift is made more apparent in confocal images of the same cultures (M, control; N, Ab treated).

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Figure 2: Cell cycle re-entry is marked by loss of CDK5 from the cell nucleus.

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Figure 3: Cell cycle re-entry is reduced after inhibiting CDK5 nuclear export.

E16.5 primary cortical neurons grown for 5 DIV before treatment with Leptomycin B

(LMB). In untreated cells (A), CDK5 immunoreactivity is both nuclear and cytoplasmic.

LMB treatment (B) eliminated virtually all of the cytoplasmic staining. C: After 48h,

cells counts (DAPI) were performed in untreated cultures (NTX) or cultures treated with

Aβ1-42 alone (Aβ), LMB alone (LMB) or both (Aβ + LMB). Counts are expressed as the percent survival of the cells originally plated. D: Counts of BrdU-positive cells are expressed as a percentage of the total DAPI cell count. Note that this percentage increases substantially after Ab1-42 treatment, but decreases in the presence of LMB – an effect that cannot be over-ridden by co-treatment with Aβ1-42 Before their complete degeneration, pycnotic cells are easily identified in the cultures by their condensed and often fragmented nuclei. Non-pycnotic cells were counted as “live”, a term that should be considered a working definition. E: This panel shows the percentage of the total cells in each category that were both BrdU-positive and in which CDK5 had shifted to a predominately cytoplasmic location. F: This panel shows the percentage of total cells that were CDK5/BrdU double-labeled and in which CDK5 expression was found in both cytoplasm and nucleus. Note that the number of BrdU+ live cells with CDK5 in the nucleus is higher than the number of BrdU+ live cells with CDK5 found only in the cytoplasm.

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Figure 3: Cell cycle re-entry is reduced after inhibiting CDK5 nuclear export.

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Figure 4: CDK5 absence in the nucleus is associated with cycle protein re-expression in R1.40 mouse brain.

Various cortical regions of the R1.40 APP transgenic mouse are shown stained for CDK5 location (red) and cell cycle activity (PCNA, green). DAPI (blue) was used as a nuclear counterstain. Non-transgenic animals show no PCNA staining either frontal cortex (A-D) or hippocampus (E-H). (Omission of the primary antibody eliminated the observed staining – not shown). In the 12 month old R1.40 animal, however, both frontal cortex

(I-L) and hippocampus (M-P) have numerous PCNA-positive neurons (arrows). None of these has CDK5 expression in its nucleus. Panels Q-T are confocal images of the same material. Panels Q and R are from the frontal cortex of non-transgenic (Q) and R1.40

(R) mouse brain while S and T are from the hippocampus of control (S) or R1.40 (T) animals.

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Figure 4: CDK5 absence in the nucleus is associated with cycle protein re-expression in R1.40 mouse brain.

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Figure 5: The expression of p35, CDK5 and PCNA in Alzheimer’s disease brain.

Paraffin embedded 10μm sections were immunostained for cell cycle activity and the location of CDK5 and its activator, p35. Although there was substantial case-to-case variability, immnostaining of CDK5 was substantially darker in neurons of hippocampus

(B) and locus coeruleus (D) in material from patients who died with Alzheimer’s disease

than in comparable material from age-matched non-demented individuals (A,

hippocampus: C, locus coeruleus). Double staining with CDK5 (brown) and PCNA (red)

revealed that in neurons that were PCNA positive CDK5 was almost exclusively

cytoplasmic. The level of p35 was increased in AD brain (H) compared with control

brain (G) but remains exclusively nuclear in location.

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Figure 5: The expression of p35, CDK5 and PCNA in Alzheimer’s disease brain.

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Figure 6: CDK5 is not necessary for tau hyperphosphorylation after Aβ treatment.

Neurons of three different genotypes (Cdk5+/+, Cdk5+/-, Cdk5-/-) were grown for 5 DIV and treated with 10μM Aβ1-42 for 24 hours after which PHF-1 antibody was used to detect

the levels of phosphorylated tau (at residues S396 and S404). Before treatment,

PHF-1 staining was present but not abundant in all genotypes (A, wild-type; C,

heterozygote; E, knockout). After treatment, staining intensity increased in all genotypes,

including the mutant Cdk5-/- (B, wild-type; D, heterozygote; F, knockout).

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Figure 6: CDK5 is not necessary for tau hyperphosphorylation after Aβ treatment.

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Figure 7: Loss of CDK5 does not protect against Aβ–induced neurodegeneration.

Wild type (A, D), heterozygote (B, E) and knockout (C, F) cortical neurons were grown

for 5 DIV then either treated with 10μM Aβ 1-42 (D, E, F) or left untreated (A, B, C).

After 48h, cells were labeled with PHF-1 (green), anti-activated caspase-3 antibody (red) and DAPI (blue). Aβ -treated cells, regardless of genotype, were positive for either phosphorylated tau or caspase-3, but not for both. Counts of all cells in culture revealed substantial cell loss after Ab treatment (G, H). This is apparent viewed as either the absolute number of cells remaining in the dish (G) or when the number of cells in the treated culture is expressed the percent of untreated (H).

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Figure 7: Loss of CDK5 does not protect against Aβ–induced neurodegeneration.

Figure 8: GSK-3β is crucial for Aβ -induced tau phosphorylation.

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E16.5 primary cortical neurons grown for 5 DIV were left untreated (A, E, I, M) or

treated with the CDK5 inhibitor, roscovitine (B, F, J, N), the GSK-3β inhibitor, lithium

(C, G, K, O), or both (D,H, L, P). There is a basal level of tau phosphorylation in the

cultured neurons that is detected with the PHF-1 antibody (A, E). Roscovitine (F) or lithium (G) treatment each results in only a partial reduction in this basal level.

Combined use of both drugs (D, H) is more effective, but still incomplete, suggesting that

a third kinase is available for tau phosphorylation. Addition of Aβ1-42 to untreated (I, M), roscovitine treated (J, N), lithium treated (K, O), or lithium plus roscovitine treated (L,P)

cultures for 24h was followed by double labeling with PHF-1 (green) and anti-TuJ1 (red)

with a DAPI counterstain (blue). In these cultures the level of PHF-1 staining was

increased in untreated cultures (I, M). This increase was substantially blocked by lithium

(K, O), not by roscovitine (J, N). The use of both inhibitors (L, P) did not significantly

differ from the results with lithium alone.

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Figure 8: GSK-3β is crucial for Aβ -induced tau phosphorylation

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Figure9:CDK5 is not necessary for tau hyperphosphorylation at the AT8 sites after Aβ

treatment.

Neurons of all three genotypes (Cdk5+/+, Cdk5+/-, Cdk5-/-) were grown for 5 DIV and treated with 10μM Aβ1-42 for 24 hours. The AT8 antibody was used to detect tau

phosphorylated at serine residues 199 and 202. The results were nearly identical to those

found with PHF-1. Before treatment, phosphorylated tau was not abundant in the cells of

any genotype (A, C, E). In knockout cultures, AT8 staining (E) is less than either

wildtype (A) or heterozygote (C) cultures. After treatment with Aβ, AT8 staining was increased in all 3 genotypes.

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Figure9:CDK5 is not necessary for tau hyperphosphorylation at the AT8 sites after Aβ treatment.

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Chapter 4: Discussion

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Research Conclusion

In most text books, neurons are described as permanently post-mitotic after they leave their birth place – the ventricular zone (VZ) or subventricular zone (SVZ). However, in the past decades, evidence has accumulated suggesting that this might not completely true. As already described, aberrant cell cycle events are found in many differentiated neuronal cells, such as Rb deficient neurons. These cell cycle events, however, tend to be related to neurodegeneration instead of division. My thesis work has provided further evidence for this unexpected linkage of cell cycle and cell death. I have found that there are multiple neuronal cell types re-enter a cell cycle like process in the E2f1-/- mice brain.

These processes include cell cycle related protein re-expression and DNA replication.

The later suggests that the neurons have already passed into S phase. The number of neurons in E2f1-/- mice does not obviously increase. Instead, there was neuronal atrophy

in older E2F1 knock out mice. This strongly suggests that the cycling neurons are

involved in a process of atrophy and degeneration instead of division. My study also suggests that conventional cell cycle proteins might play an unconventional role in neuronal cell cycle control. For example, E2F1 is a very important player in normal cell cycle progress. However, my results suggest that it plays the role of cell cycle suppressor

in neurons. My studies of the CDK5 protein also point to an abnormal role for this

protein in neuronal cell cycle control. The traditional CDKs are important drivers for

normal cell cycle progression. In contrast, the most important role of CDK5 in post-

mitotic neurons appears to be in processes such as neuronal migration, survival and

synaptic functions. Because it is not detected in most cells during a normal cell cycle, it

was thought that CDK5 was not involved in the cell cycle events. Our lab have shown

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that quite to the contrary, CDK5 is important for neuronal cell cycle regulation, further, it

is important for normal neuronal differentiation (Cicero and Herrup, 2005). My work has

significantly extended these observations by showing that the nuclear localization of

CDK5 is important for holding the cell cycle in check. This cell cycle suppression might

not require binding with its traditional activator p35 because the localization of p35 does

not change with CDK5 in “cycling” neurons. This finding has clinical relevance, because

ectopic cell cycle events are found in several different neurodegenerative diseases and

their mouse models.

Aberrant cell cycle events in neurodegenerative diseases and their

mouse model

The best studied neurodegenerative disease which is associated with the aberrant cell

cycle events in their disease related brain area is Alzheimer’s disease (AD). Several groups have shown that there is increased expression of multiple cell cycle related protein

such as cyclin E, PCNA, Ki67, cyclin B1, cyclin D and CDK4 in AD related brain

region – including hippocampus, locus coeruleus and dorsal raphe (Nagy et al., 1997a;

Vincent et al., 1997; Busser et al., 1998). The neuronal loss is severe in these regions in

AD patients. However, in unaffected regions such as the cerebellar contex, the expression

of cell cycle related proteins is not found. This suggests that the ectopic cell cycle events are associated with neuronal death. This conccept is consistent with the cell cycle related neuronal death in T antigen transgenic mice and granule cell death in staggerer and

Lurcher mice as discussed in Chapter 1. In addition to the presence of these proteins,

There is also increased activity of CDC2:cyclin B complex and its activating phosphatase,

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CDC25, in the AD brain (Vincent et al., 1997; Ding et al., 2000; Vincent et al., 2001).

Further, Yang et al offer strong support for the presumption that this ectopic cell cycle

activity is productive through the use of fluorescent in situ hybridization (FISH) to show

that actual DNA replication has taken place in select regions of the AD brain. Even

though both DNA replication and G2/M phase marker expression can be documented, no

cell division is found. Instead, this ectopic cell cycle progression accompanies neuronal

cell death at all stages of AD disease (Yang et al., 2001; Yang et al., 2003). Recent

studies show that forcing a neuron back to cell cycle could induce a neurodegenerative

changes which is similar to those seen in AD both in vivo and in vitro. These data provide

support for the ectopic cell cycle events are involved in the generation of the

characteristic pathological hallmarks of AD (McShea et al., 2007; Park et al., 2007).

The ectopic cell cycle protein expression is not only found in AD. It has been reported in

the brains of patients with Parkinson’s disease (PD), Ataxia-telangiectasis (AT),

amyotrophic lateral sclerosis (ALS), Down’s syndrome , Pick’s Disease, Niemann-Pick

syndrome type C and brain injury induced by focal ischemia (Nagy et al., 1997b;

Husseman et al., 2000; Love, 2003; Nguyen et al., 2003; Ranganathan and Bowser, 2003;

Wen et al., 2005; Yang and Herrup, 2005). In no case is this ectopic cell cycle found to progress to M-phase in these diseases. It would appear that in the affected brain area, death rather than a full cell cycle is the result of the events. Recent results have shown that blocking CDK activity blocks focal ischemia induced neuronal death. This suggests that even though we refer to the proteins as cell cycle proteins they must also play important roles in neuronal cell death (Rashidian et al., 2005; Wen et al., 2005).

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Cell cycle events in mouse models

Consistent with the results for these human diseases, expression of cell cycle proteins is

found in the mouse models of AD and AT disease. Yang et al reported that abnormal cell

cycles occur in 3 different transgenic model of AD—(R1/40, Tg2576 and App23) (Yang

et al., 2006) as well as in one model of AT (Yang and Herrup, 2005). Ectopic cell cycle

protein expression and DNA replication has been documented in these transgenic lines.

In the AD mouse, ectopic cell cycle events first appear at 6 months of age, which is much

earlier than the earliest Aβ deposits. These are not found in these animals until the mouse is one year old. Despite the accurate mimicking of several phenotypes of the human disease, however, there is no neuronal cell loss in the relevant region of these mouse models. The appearance of cell cycle events in these neurons does not lead immediately to neuronal death. In this behavior, the disease models are similar to the situation in E2f1-

/- mice we report in Chapter2. This suggestion is that the cell cycle events indicate that

these neurons are under some stress and may be unhealthy. This implies that loss of cell

cycle control in neurons might be a sensitive detector of problem in the neuronal

environment.

Thus ectopic cell cycle events open a new view on neurodegenerative disease. Its early

appearance in the human neurodegenerative diseases and their mouse models might be of

considerable value in finding the early factors that induces these diseases. It urges us to

ask several important questions---what are the factors which induce this aberrant cell

cycle process? What is the difference between this abnormal cell cycle event and normal

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cell cycle process? What is the possible signaling pathway for this process and what is

the result of this process?

The factors that might induce the cell cycle re-entry

The factors that might induce the cell cycle re-entry are undoubtedly as numerous as the

reasons for the various neurodegenerative diseases. From the in vitro and in vivo data,

however, we have begun to identify some factors that might be important triggers for the

cell cycle re-entry in neurons.

Inflammation

Many neurodegenerative diseases such as AD, stroke and PD involve chronic

inflammation as part of their etiology. In these diseases, microglial induced inflammation is implicated in CRND related disorders (Bamberger and Landreth, 2002; McGeer and

McGeer, 2002a). In AD disease, Aβ can activate microglia and lead to secrete a diverse array of bioactive molecules including proinflammatory cytokines and mitogens. These secreted factors will further activate microglia as well as neighboring astrocytes. In the end, this local inflammatory environment will be part of processes that triggers induce the loss of neurons. In vitro, cultured primary neurons express several cell cycle related proteins such as PCNA, cyclin D and engaged in BrdU incorporation when treated with the conditioned media from Aβ -activated microglia (Wu et al., 2000). This suggests that activated microglia may release some factors that serve as the source of mitogens that induce cell cycle re-entry in post-mitotic neurons. The identity of those factors and the mechanism by which they induce cell cycle re-entry in a neuron are unknown. It is likely that multiple pathways involved in the balance between maintaining quiescence and cell

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cycle re-entry. It has been shown that beside the activated microglia, AD and other neurodegenerative disease brains also express other markers of inflammation including elevated levels of immune receptors, cell surface proteins and cytokines (Bamberger and

Landreth, 2002; McGeer and McGeer, 2002a, b, c, 2004). Some of these, such as nuclear factor kB (NF-kB), reactive oxygen and nitrogen species (RONS) and growth factors are involved in both cellular proliferation and carcinogenesis (Hofseth et al., 2005; Ying et al., 2005; Karin, 2006a, b). Among them, as discussed in the following section, reactive oxygen species (ROS) are neurotoxic. Whether those inflammation factors are related to cell cycle re-entry is unknown now. Their presence and association with neuronal disease should motivate researchers to explore their role in the cell cycle re-entry.

Oxidative stress and DNA damage

Oxygen is necessary for cells. However, its metabolites – reactive oxygen species (ROS)

- are toxic to cells. ROS can cause extensive cellular damage involving lipids, proteins and DNA. Neurons are thought to be particularly vulnerable to ROS because of their high metabolic rate. In several neurodegenerative diseases, markers of oxidative stress are found in patients as well as their animal models. Because both cell cycle deregulation and oxidative stress are associated human neurodegenerative disease and their mouse model, the relationship between them is interesting to investigate. Recently, the harlequin (Hq) mouse mutant has offered new insight the role of oxidative stress in cell cycle related neuronal death. Hq mice develop progress ataxia associated with the death of both granule and Purkinje cells in cerebellum. The Hq mutation is caused by a pro-viral insertion on the first intron of the apoptosis-inducing factor (Aif) gene. This results in an

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80%-90% reduction of Aif protein. Down regulation of Aif in Hq mutant mice is

associated with oxidative damage followed by the onset of neurodegeneration several

months later. This indicates a central role for oxidative stress in the progression of

neurodegeneration. In addition to the changes in oxidative damage, neurons in Hq mice

show the expression of cell cycle protein including PCNA and CDK4. These neurons also

incorporate BrdU. Unlike the neurons in E2f1-/- or in the mouse models of AD or AT we discussed above, this cell cycle re-entry is tightly associated with cell death. Most of these “cycling” cells are positive for apoptotic markers such as activated caspase-3 and

display pycnotic nuclei. All neurons which have mitotic figures had oxidized DNA;

however, not all neurons with oxidized DNA were cell cycle positive. This suggests that

the oxidative damage precedes cell cycle reentry.

The mechanism by which oxidative stress induced deregulation of the cell cycle in

postmitotic neurons is not clear. Ros has been implicated to change the mitogenic

signaling of several pathways by abnormally activating growth factor receptors or other

signaling molecules (Klein and Ackerman, 2003). Another possibility is that oxidative stress-induced DNA damage might contribute directly to the cell cycle re-entry in mature neurons. Recent data suggest that DNA damage itself may be the trigger for cell cycle activation. DNA damage agents are known induce cell cycle re-entry and subsequent apoptosis in cultured primary neurons (Kruman et al., 2004). ATM may be involved in this DNA damage induced neuronal death following S-phase re-entry because atm-/-

neurons are resistant to DNA-damage induced death. This model links with the E2F1

results reported in Chapter 2. The level of E2F1 is increased after it is stabilized by DNA

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damage induced ATM phosphorylation (Lin et al., 2001). As E2F1 plays important roles

in the regulation of both cell cycle events and apoptosis, elevated level of E2F1 might

well contribute to the DNA damage induced cell cycle re-entry and cell death.

Unconventional expression of cell cycle proteins in

degenerative neurons

In a normal cell cycle, the sub cellular localization of cell cycle regulators such as CDK-

cyclins is essential for proper cell cycle coordination. Distinct CDK-cyclin complex appear at the specific stages of the cell cycle to insure the normal cell cycle progress properly. For example, cyclin A and cyclin E are predominantly nuclear from the time of their initial appearance in G1 through M phase, when it is degraded (Pines and Hunter,

1991). The constitutive nuclear localization of these two cyclins and their associated

CDKs are important for DNA replication and prevention of re-replication. CDK-cyclin D only remains in the nuclear only during G1. As S phase begins, the complex is exported to the cytoplasm for targeted proteolysis (Baldin et al., 1993; Diehl et al., 1998). Cyclin

B1 and cyclin B2 are synthesized in the cytoplasm and they enter the nucleus only as cells enter prophase. Taken together these distinct localization patterns suggest that the different mitotic CDK-cyclin complexes have distinct substrates that promote mitotic changes in specific subcellular localizations.

In degenerating neurons of human, mouse models and in vitro culture, the expression of

some cell cycle proteins appears to be “mislocalization” (Vincent et al., 1997; Busser et

al., 1998; Yang et al., 2003; Yang et al., 2006). For example, nuclear accumulation of

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cyclin D1 may regulate neuronal death and survival. As discussed above, that cyclin D1

only appears in the nucleus during G1 phase in normal proliferating cells and cyclin D-

associated kinase activators are down regulated in differentiated neurons (Kranenburg et al., 1995). In degenerating neurons, however, the level of cyclin D1 and the activity of its associated kinase are increased upon CRND stimuli. In untreated cultures, cyclin D1 mainly localizes in the cytoplasm of differentiated neurons. After several CRND- inducing stimuli, cyclin D1 accumulates in the nuclear. This is accompanied by phophorylation of pRb and subsequent neuronal death. This cell death is blocked by blocking the kinase activity (Park et al., 1997a; Park et al., 1997b; Park et al., 1998a; Park et al., 1998b; Giovanni et al., 1999; Ino and Chiba, 2001; Zhang et al., 2006; Alvira et al.,

2007; Rashidian et al., 2007). Recently Park and colleagues have reported that neurons in dominant negative CDK4 transgenic mice were resistant to hypoxia induced cell death.

Also cultured cerebellar granule neurons (CGNs) from cyclin D1-deficient mice were much more resistant to hypoxia than control (Rashidian et al., 2005). These in vivo and in vitro data strongly suggest a role for CDK4:cyclin D in neuronal death, and that this role is affected by their cellular localization. The implication is that neurons normally prevent nuclear accumulation of cyclin D. Under some stimuli, however, neurons lose this ability and cyclin D changes its localization from cytoplasm to nucleus. This might trigger cell cycle activation and neuronal death probably through the pRb/E2F pathway. Interestingly, in some neurodegenerative diseases and their mouse models, the localization of cyclin D1 is cytoplasmic. This might explain why stressed neurons donot die even after they initiate a cell cycle re-entry process, but DNA synthesis still happen. Another example is cyclin

A. Generally, cyclin A is located in the nucleus of cycling cells in S phase. In stressed

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neurons of human neurodegenerative diseases and their mouse models, it has been consistently reported that cyclin A is expressed in the cytoplasm. Our preliminary data indicate that cyclin A moves into the nuclei when those “stressed” neurons get second hit and undergo apoptosis (unpublished data). It is not clear whether the presence of those proteins in abnormal site is toxic or protective for neurons. Given that “cycling” neurons does not die immediately, it seems like that they are not toxic for neurons when they are in the cytoplasm. It will be important to investigate the function of those proteins in each location. It will be interesting to know, for example, whether they can activate their kinase partners in the abnormal sites and what would happen if they were forced into nuclei.

Mechanism for the ectopic cell cycle re-entry

It has been more than a decade since the first report of ectopic cell cycle events in adult neurons and its association with neuronal degeneration. While much has been learned about the process, it is still unclear what signals drive a post mitotic neuron to re-enter a cell cycle. Less clear still is the nature of the signaling pathway leading from this ectopic cell cycle event to neuronal death. It seems likely that there are multiple mechanisms involved, and the complexity is probably magnified because different neurons may well use different cell cycle control machineries. In atm-/- mice, only Purkinje cells (and an occasional striatal neuron) show cellcycle deregulation. Other neurons apparently tolerate the deletion of the atm gene yet control their cycle properly. In AD mice, only neurons in specific AD related brain regions show cell cycle deregulation. In E2f1-/- mice, however, there are more neurons that are involved in the cell cycle de-regulation phenomenon. This

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implies that the normal cell cycle regulation machinery, important for holding the

neuronal cell cycle in check, is different in different neurons and is compromised by

different insults. In fact, interruption of pRb/E2F pathway, either by oncogene expression or targeted deletion of the genes involved in this pathway such as pRb, induces ectopic S

phase entry in both developing and mature neurons. Our studies show that both the

nuclear localization of CDK5 and the presence of E2F1 in mature neurons are important

for holding the cell cycle of mature neurons in check. This could imply either that there

are multiple pathways controlling the post mitotic neuronal cell cycle, or CDK5 and

E2F1 participate in a single master pathway. To investigate whether CDK5 and E2F1 participate in same pathway or parallel pathway, we first should know whether the

“cycling” neurons in E2f1-/- or neurons with CDK5 translocation are the same cell type or

not. Next we will exam the localization of CDK5 in E2f1-/- neurons and the level of E2F1

in neurons with CDK5 translocation. By using E2F1 and CDK5 double mutant neurons,

the “cycling” status of those double mutant neurons might give us some interesting

information about the interaction of these two proteins in the controlling of neuronal cell

cycle.

For pRb/E2F1 pathway, now we know deletion of either pRb or E2F1 can induce

neuronal cell cycle re-entry (MacPherson et al., 2003). In both cases there is cell cycle

protein expression and DNA replication. However, we know little about whether this is

accompanied by a change in all normal G1 and S phase proteins such as the CDKs.

Normally, the pRb/E2F1 complex represses gene transcription in differentiated neurons.

When either pRb or E2F1 is deleted, those genes might be de-repressed and affect the

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cycle status of neurons. Another question that needs to be resolved is how pRb/E2F1 complex represses cell cycle genes in normal neuronal cells and how pRb or E2F1 behave when separated from the complex or in situations where its normal partner is absent. Neuronal stem cell system might be a good way to study of the changing role of

E2F1 in proliferating cells and differentiated cells. By using protein-bond assays, we could detect normal cycle proteins as they move into and out of interaction with E2F1.

When the stem cells are induced to differentiation, the key protein interaction should shift and presumably the repression complex will form. If we used RNAi to knock down E2F1 or pRb, the inhibition complex might not form or form incompletely. In this way, we wish we could make progression in unraveling the regulation of cell cycle events in mature neurons.

For CDK5, although it is not involved in normal cell cycle progress, it is expressed immediately after neurons exit the cell cycle. Several studies show CDK5 is important for neuronal cell cycle exit as well as for differentiation. Our recent studies, discussed in

Chapter 3, show that the nuclear localization of CDK5 is an important part of the execution of preventing a cell cycle in post-mitotic neurons. The next step in this project might be to resolve the mechanisms by which CDK5 holds the cell cycle in check in differentiated neurons. First, we should determine whether this function of CDK5 is related in its kinase activity. Second we should block p35 and p39 interaction to find out whether these normal CDK5 activators are involved in this process. Third would be to explore the signaling pathways used by CDK5 in repressing the cell cycle.

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There are several possible models of CDK5 action that would explain the data accumulated so far. First, CDK5 might directly act through proteins which are involved in the cell cycle control. For example, CDK5 can stabilize p27 (Kawauchi et al., 2006), a

CDK suppressor. In p27-/- mice, there is an increase in the number of late born cortical neurons, which ultimately leads to a larger cortex. This emphasizes the role of p27 in normal neuronal cell cycle inhibition (Goto et al., 2004). It would be interesting to determine the p27 expression levels in Cdk5-/- neurons. P27 like CDK5 is involved in multiple neuronal processes such as cell migration and differentiation (Besson et al., 2004;

Nguyen et al., 2006b; Nguyen et al., 2006a). Clearly more work is needed to define the interaction of these two systems. A different model could be built around the idea that

CDK5 can bind to normal cyclins. This raises the possibility that CDK5 might bind them buffering their ability to activate cell cycle kinases. It is also reported that CDK5 could phosphorylate pRb and tighten its binding of E2F family members and /or promotes its differentiation functions (Lee et al., 1997; Kastner et al., 1998; Hamdane et al., 2005).

The relationship between CDK5 and other pocket protein members is not clear. It has been found that both CDK5 and p130 are increased when neuronal cells stop cell cycle and begin to differentiation in the development of the quail neuroretina(Kastner et al.,

1998). So it is deserved to explore the relationship between CDK5 and other pocket proteins. Another pathway is that CDK5 might work through extracelluar mitogen- activated protein kinase (MAPK) signaling pathway. This signaling pathway affects cell proliferation and differentiation (Strok, 2002). CDK5 could down-regulated this pathway through phosphorylating several proteins on this pathway such as MEK1, Ras guanine

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nucleotide exchange factor (RasGEF) (Sharma et al., 2002; Alexander et al., 2004;

Kesavapany et al., 2004; Hung et al., 2005; Kesavapany et al., 2006).

The fate of “cycling” neurons

A final important but unresolved question is what happens to that the neurons after they make the decision to enter an ectopic cell cycle. It is clear that “cycling” neurons can have at least two different fates: one in development and one in adulthood. In Cdk5-/- and

Rb-/- and T-antigen transgenic mice, forced cell cycle re-entry of the differentiating neurons clearly leads to neuronal death. This is not the whole story, however, becasue in

Rb conditional knock out mice, where Rb deleted from the nervous system at later developmental stages, the neurons undergone ectopic S phase entry, but they do so in the absence of cell death (MacPherson et al., 2003). Because this conditional mice lack the erythropoiesis defects that lead to CNS hypoxia in Rb-/- mice, the authors speculate that

hypoxia is a necessary cofactor to neuronal death. The Leone lab leads support to this

idea by identifying a placental defect in the Rb null mice that they can be rescued by

either tetraploid aggregation or conditional knockout strategies. In both cases, restoration

of normal erythropoiesis suppressed most of the defects seen in Rb-null animals. These

animals still have ectopic cell cycle re-entry in CNS, but they lack neuronal death. The

observations are consistent with our own findings that adult neurons can re-enter a cell

cycle without dying. This is the state of the neurons in both the AD and AT mouse

models as well as in the E2F1 deficient mice. The exact linkage between these two data

sets is not clear but the analogy might shed some light on the relationship between the

ectopic cell cycle events and neuronal cell death in immature versus adult neurons. Fully

differentiated neurons appear to have much more ability to control the initiation of

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ectopic cell cycle events than do neurons that are still in development. Perhaps adult

neurons detect stress and responded by re-entering some cell cycle like process, but then

are able to hold the cell cycle in check. They can do so until forced to initiate the cell

death program, perhaps in response to strong second “hits”.

The unconventional expression of cell cycle proteins in stressed neurons gives us some

clues as to the reason for the deregulated cell cycle events in neurons such as the E2f1-/- ,

AT and AD mouse brain. These neurons are in a highly unusual state. They express cell cycle proteins, all or part of their DNA is replicated but they do not die, and they do not divide. When other stimuli hits those stressed neurons, those cycling proteins might move to the nucleus and activate their associated CDKs.

Even these CDKs are in the right place and have normal function as they in the normal proliferating cells, it has less chance for a fully differentiated neuron to finish a normal cell cycle and divide again. There are several reasons about this issue. First, as discussed in Chapter 1, a cell cycle is a very complicated process that is tightly regulated at each step by many factors. Cells will be well prepared before entering a proliferating cell cycle.

For a post-mitotic neuron, however, its cycle machinery is probably shut down to allow

for full differentiation. Thus it is not ready for division and cannot complete each of steps

required to successfully finish a cell cycle. Further as part of the normal mechanism that

is in place to insure that differentiated neurons do not go back to the cell cycle, cell cycle

machinery itself might change its role in a stressed neuron. For example, E2F1 is a

necessary regulator for proper cell cycle progression in normal cycling cells. However, it

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also controls the transcription of other genes which are not involved in the cell cycle.

Studies in a variety of different cell types have shown that which signaling pathway will

be induced by E2F is very context dependent. Base on my work we now know that in

neurons E2F1 functions as a cell cycle suppressor and promotes differentiation. Thus

while deregulated E2F1 in mature neurons might be able to activate some cell cycle genes, it may also signal to the apoptosis pathway. Overall, it is impossible for an adult

neuron in a context to finish a cell cycle. When under stress, its cell cycle machinery is

activated; the fate of this neuron might be death. In developing neurons that death is

immediately, for example, the neurons in Rb-/- mice or Cdk5-/- mice or the cerebellar granule cells depleted of target. In adult neurons, the ability to control the cell cycle machinery and cell death progress is more complicated. These more established neurons can survive for months, but we believe they are very vulnerable to environment stimuli.

When one of other hits comes, they might initiate a death progress.

For the post-mitotic neurons, once they exit from the last cell cycle events, they will need to suppress their cell cycle machinery for the rest of whole life. If some factors re-activate their cell cycle machinery, they can go back to the cell cycle in some degree such as they can replicate their DNA, but it is hard for them to finish this kind of aberrant cell cycle.

The final fate of those “cycling” neurons will end at death.

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