Download mecp2 and the epigenetic regulation of excitatory synaptic

MECP2 AND THE EPIGENETIC REGULATION OF EXCITATORY SYNAPTIC
TRANSMISSION
APPROVED BY SUPERVISORY COMMITTEE
__________________________________
Ilya Bezprozvanny, Ph.D.
__________________________________
Lisa Monteggia, Ph.D.
__________________________________
Ege Kavalali, Ph.D.
__________________________________
Weichun Lin, Ph.D.
Dedicated to my parents,
Ann & Steve Cunningham and Jon Nelson,
and to the rest of my family and friends
for their continuing love and support.
ii
MECP2 AND THE EPIGENETIC REGULATION OF EXCITATORY SYNAPTIC
TRANSMISSION
By
ERIKA DAWN NELSON
DISSERTATION
Presented to the Faculty of the Graduate School of Biomedical Sciences
The University of Texas Southwestern Medical Center at Dallas
In Partial Fulfillment of the Requirements
For the Degree of
DOCTOR OF PHILOSOPHY
The University of Texas Southwestern Medical Center at Dallas
Dallas, Texas
June 2007
iii
Copyright
by
Erika Dawn Nelson, 2007
All Rights Reserved
iv
ACKNOWLEDGEMENTS
I would like to thank my mentor, Dr. Lisa Monteggia, for all of her support and
guidance over the last 4 years. I would also like to thank Dr. Ege Kavalali for teaching
me all about synaptic transmission and for his advice and expertise in the field. I would
like to acknowledge all past and present members of the Monteggia lab for all of their
help and encouragement. Especially Terry Gemelli and Sunbola Ashimi for their senses
of humor and Megha Upadhyaya for being an awesome friend and a great cheerleader. I
would also like to acknowledge all past and present members of the Kavalali lab, in
particular, Tuhin Virmani, Yildrim Sara and Mert Ertunc for their extraordinary
knowledge of electrophsiology and their patience in sharing some of that with me. Also, I
thank Catherine Wasser for being a fortunate person to work so closely with and for
being an inspirational friend.
I’d like to thank the other members of my thesis committee, Ilya Bezprozvanny
and Weichun Lin, for all of their helpful suggestions over the years. I must also thank our
collaborators Eric Olson and Rusty Montgomery for the use of their HDAC mice and to
Tina Han and Leeju Wu in the McKnight lab for their assistance in measuring DNA
methylation. I would also like to thank my fellow researchers Tom Green, Imran Alibhai,
Brian Potts, Victor Galanis and Herb Covington for being great people to work with and
to get to know. Finally, I would like to thank the University of Texas Southwestern
Medical Center for their monetary support through the University training grant.
As for my family and friends, I don’t have enough thanks for my mother who is
always there to calm me down after a bad day and who loves me unconditionally. To
Christi Edwards, thanks for continuing to be my best friend even when I found myself too
v
busy to spend time with you. Thank you to John Lacy, to whom I owe my new optimistic
outlook on life. Last but not least, I’d like to thank my pug, Brownie, for always making
me smile and helping me to get out of bed in the mornings.
You all mean so much to me… I am truly blessed to have you in my life!
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MECP2 AND THE EPIGENETIC REGULATION OF EXCITATORY SYNAPTIC
TRANSMISSION
Erika Dawn Nelson, Ph.D.
The University of Texas Southwestern Medical Center at Dallas, 2007
Supervising Professor: Lisa Monteggia, Ph.D.
Accurate regulation of gene expression is critical for normal brain function. Many
human neurodevelopmental and neurodegenerative disorders are associated with
mutations in genes important for controlling transcription. Mutations in one such gene,
the transcriptional repressor methyl-CpG-binding protein 2 (MeCP2), lead to a form of
mental retardation called Rett Syndrome (RTT). Though the MeCP2 protein is expressed
ubiquitously, symptoms of RTT patients are primarily neurological, which include
reduced mental capacity, autistic-like behavior and autonomic dysfunction. In addition, a
mouse model with reduced MeCP2 expression specifically in postnatal, forebrain neurons
recapitulates many of the phenotypes seen in human patients. These findings, among
others, lead to interest in MeCP2’s function in the brain. Our research has focused on the
transcriptional repression activity of MeCP2 and its role in the regulation of synapse
vii
function. Using mainly electrophysiological techniques, we found that the loss of MeCP2
in hippocampal neurons results in deficits in both spontaneous and evoked excitatory
synaptic transmission. Using pharmacological manipulations, we were able to attribute
these deficits to the loss of transcriptional repression by MeCP2. By utilizing a
conditional knockout approach, we found that these effects were not due to the loss of
MeCP2 during neurodevelopment and that they were primarily due to a deficiency in
presynaptic vesicle release. We further extended these findings by looking at two
mechanisms for controlling the repression of gene expression, DNA methylation and
histone deacetylation, both of which are important for MeCP2’s function as a
transcriptional repressor. Using inhibitors of DNA methyltransferases, we discovered that
synaptic activity-dependent decreases in DNA methylation occur in post-mitotic neurons,
and that these changes in DNA methylation can regulate spontaneous synaptic
transmission. We were also able to rescue the MeCP2-dependent decrease in spontaneous
activity by treating neurons with the methyl donor, S-adenosyl-L-methionine. Finally, we
addressed the role of histone deacetylation in synapse function by conditionally deleting
histone deacetylases (HDACs) 1 and 2 from mature hippocampal neurons. HDAC1 and 2
are present in the transcriptional repressor complex containing MeCP2. After acute
knockdown of HDAC1 or HDAC2, we found deficits in excitatory synaptic transmission
that mimicked the defects seen after the constitutive loss of MeCP2. In summary, we
have discovered a role for the transcriptional repressor, MeCP2, and two components of
its repressor complex, DNA methylation and HDACs, in the control of excitatory
synaptic transmission between hippocampal neurons.
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TABLE OF CONTENTS
Dedication ………………………………………………………………………………...ii
Acknowledgments…………………………………………………………………………v
Abstract…………………………………………………………………………………..vii
Table of Contents…………………………………………………………………………ix
Prior Publications………………………………………………………………………….x
List of Figures…………………………………………………………………………….xi
List of Abbreviations……………………………………………………………………xiv
Chapter 1: Introduction...………………………………………………………………...17
MeCP2…………………………………………………………………………...20
DNA Methylation………………………………………………………………..26
Histone Deacetylation…………………………………………………………....32
Chapter 2: MeCP2-dependent Transcriptional Repression Regulates Excitatory
Neurotransmission……………………………………………………….37
Introduction……………………………………………………………………....37
Results…………………………………………………………………………....40
Discussion………………………………………………………………………..46
Chapter 3: Activity-dependent Suppression of Excitatory Miniature
Neurotransmission Through the Regulation of DNA Methylation………......56
Introduction……………………………………………………………………....56
Results…………………………………………………………………………....58
Discussion………………………………………………………………………..68
Chapter 4: Loss of HDAC1 and HDAC2 in Hippocampal Neurons
Results in Specific Alterations in Excitatory Synaptic Transmission…………...79
Introduction……………………………………………………………………....79
Results…………………………………………………………………………....82
Discussion………………………………………………………………………..87
Chapter 5: Conclusions and Future Directions…………………………………………..96
Materials and Methods………………………………………………………………….104
References………………………………………………………………………………114
Vitae…………………………………………………………………………………….126
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PRIOR PUBLICATIONS
Nelson, E.D., Kavalali, E.T., Monteggia, L.M. (2006). MeCP2-dependent transcriptional
repression regulates excitatory neurotransmission. Current Biol. 16, 710-716.
Gemelli, T., Berton, O., Nelson, E.D., Perrotti, L.I., Jaenisch, R., and Monteggia, L.M.
(2005). Postnatal loss of MeCP2 in the forebrain is sufficient to mediate behavioral
aspects of Rett Syndrome in mice. Biol Psychiatry. 59, 468–476.
Nelson, E.D., and Monteggia, L.M. (2007). Activity-dependent regulation of gene
expression. Encyclopedia of Neuroscience. (In press). Elsevier.
Nelson, E.D., Kavalali, E.T., Monteggia, L.M. (in revision) Activity-dependent
suppression of excitatory miniature neurotransmission through the regulation of DNA
methylation.
Nelson, E.D., Montgomery, R., Olson, E.D., Kavalali, E.T., Monteggia, L.M. (in
preparation) Loss of HDAC1 and HDAC2 in hippocampal neurons results in specific
alterations in excitatory synaptic transmission.
x
LIST OF FIGURES
Figure 1-1. Transcriptional repression by MeCP2 is relieved following synaptic
activity…………………………………………………………..…………....20
Figure 1-2. Common MeCP2 mutations in Rett Syndrome……………………..……….22
Figure 1-3. DNA methylation and demethylation……………………………………….26
Figure 1-4. Histone acetylation and deacetylation……………………………………….33
Figure 2-1. Spontaneous miniature synaptic currents in cultured hippocampal neurons
from MeCP2 knockout and control mice…………………………………….49
Figure 2-2. Miniature EPSC frequency is reduced in older MeCP2 KO neurons……. ...50
Figure 2-3. Total pool and readily releasable pool of synaptic vesicles from MeCP2
knockout and control neurons………………………………………………..51
Figure 2-4. Evoked synaptic responses in MeCP2 knockout and control neurons
during 10 Hz field stimulation immediately followed by 1Hz recovery
stimulation……………………………………………………………………52
Figure 2-5. Spontaneous miniature synaptic responses from wild type C57BL/6 and
MeCP2 knockout hippocampal cultures after a 24-hour treatment with
inhibitors of transcriptional repression and activation……………………….53
Figure 2-6. High and low titer lentiviral infection of dissociated hippocampal
cultures………………………………………………………………………54
Figure 2-7. Spontaneous miniature excitatory synaptic currents in floxed MeCP2
neurons infected with a lentivirus expressing Cre recombinase……………..55
Figure 3-1. Inhibiting DNMT activity in neurons causes a deficit in excitatory
synaptic transmission………………………………………………………...71
xi
Figure 3-2. DNMT inhibition for 24 hours specifically affects spontaneous
presynaptic function………………………………………………………….72
Figure 3-3. Changes in spontaneous excitatory neurotransmission after DNMT
inhibition are mediated by the loss of function of the transcriptional
repressor, MeCP2…………………………………………………………….73
Figure 3-4. 24-hour treatment of hippocampal cultures with DNMT inhibitors
reveals demethylation of genomic DNA……………………………………..74
Figure 3-5. Schematic of the CpG island in the BDNF gene…………………………….75
Figure 3-6. Treatment of hippocampal cultures with DNMT inhibitors reveals
activity-dependent demethylation of DNA and concurrent alterations
in synaptic transmission……………………………………………………...76
Figure 3-7. BDNF mRNA expression after chronic manipulation of DNA
methylation in hippocampal cultures………………………………………...77
Figure 3-8. Model for activity-dependent demethylation of genomic DNA in
post-mitotic neurons………………………………………………………….78
Figure 4-1. Treatment with HDAC inhibitors results in increased acetylation of H4
and defects in both spontaneous and evoked synaptic transmission…………90
Figure 4-2. Quantification of HDAC1 and 2 mRNA and protein expression levels one
week after infection with Cre-recombinase lentivirus……………………….91
Figure 4-3. Loss of HDAC2, but not HDAC1, results in decreased frequency
of spontaneous mEPSCs……………………………………………………..92
Figure 4-4. Miniature inhibitory synaptic currents from conditional HDAC1 and 2
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KO neurons ………………………………………………………………….93
Figure 4-5. Increased evoked EPSC amplitudes and decreased paired pulse ratios
in HDAC1, HDAC2, and MeCP2 KO neurons……………………………...94
Figure 4-6. Evoked inhibitory postsynaptic currents from HDAC1, HDAC2, and
MeCP2 KO cultures………………………………………………………….95
Figure 5-1. List of possible presynaptic gene targets of MeCP2………………………...99
Figure 5-2. Quantitative Real-Time PCR results from MeCP2 KO cultures…………..100
xiii
LIST OF ABBREVIATIONS
5azaC – 5-azacytidine
Act D – Actinomycin D
AMP – Adenosine monophosphate
AP5 - 2-amino-5-phosphonovaleric acid
BDNF – Brain-derived neurotrophic factor
CaMK – Calcium/calmodulin-dependent protein kinase
CBP – CREB binding protein
CNQX - 6-cyano-7-nitroquinoxaline-2,3-dione
CNS – central nervous system
Cplx -Complexin
CREB - Cyclic AMP response element-binding protein
DG – dentate gyrus
DIV – days in vitro
DMSO – dimethyl sulfoxide
DNA – deoxyribonucleic acid
DNMT – DNA methyltransferase
EPSC – excitatory postsynaptic current
FMR – Fragile X mental retardation
Gadd45α – Growth arrest and DNA damage 45α
GFP – Green fluorescent protein
HAT – Histone acetyltransferase
HDAC – Histone deacetylase
xiv
ICF - Immunodeficiency, centromeric region instability, facial anomalies
IPSC – inhibitory postsynaptic current
KO - knockout
LTD – long-term depression
LTP – long-term potentiation
MAP2 – Microtubule associated protein 2
MBD – methyl binding domain
MeCP2 – Methyl-CpG binding protein 2
mEPSC – miniature EPSC
mIPSC – miniature IPSC
mRNA – messenger RNA
MS-AFLP – Methylation-sensitive amplified fragment length polymorphism
NBQX - 2,3-dihydroxy-6-nitro-7-sulfamoyl-benzo[f]quinoxaline-2,3-dione
Nlgn - Neuroligin
NMDA - N-methyl-D-aspartic acid
Nrxn - Neurexin
PCR – Polymerase chain reaction
PP1 – Protein phosphatase 1
PSD-95 - postsynaptic density protein of 95 kDa
PTX - Picrotoxin
RE1 - repressor element 1
REST – RE1 silencing transcription factor
RNA – ribonucleic acid
xv
RTS – Rubinstein-Taybi syndrome
RTT – Rett syndrome
SAM – S-adenosyl-L-methionine
Synpor -Synaptoporin
Syp1 – Synaptophysin 1
Syt7 – Synaptotagmin 7
TRD – transcriptional repression domain
TSA – Trichostatin A
TTX - Tetrodotoxin
VPA – Valproic acid
WT – wildtype
Zeb – Zebularine
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CHAPTER 1
INTRODUCTION
Epigenetics is the study of stable, and potentially heritable, changes in gene
expression without alteration of the actual DNA sequence. It is also believed to be a way
that environment might control the phenotype of a cell. One epigenetic example is the
ability of each cell in an organism, all genetically identical, to differentiate into a
particular cell type to serve a specific purpose. Our kidney, skin, and blood cells, and
even our neurons, all share the same genetic makeup, but all perform significantly
different functions within our bodies due to the differential expression of cell-type
specific genes. The expression of genes is partially controlled by how tightly DNA is
bound around a core group of proteins called histones, and epigenetic phenomena are part
of what regulates that association. The epigenetic control of gene expression is mediated
by two main mechanisms: DNA methylation and histone modifications. DNA
methylation is the covalent addition of methyl groups to the 5-position of cytosine
residues within CpG dinucleotides and an important molecular tag for the repression of
gene transcription. Certain proteins, such as the methyl-CpG-binding protein 2 (MeCP2),
contain methyl-binding domains that recognize methylated DNA and repress
transcription. There are a number of different histone modifications that are important for
either the activation or repression of gene expression. Histone acetylation occurs on the
amino side chain of specific lysine residues found on histone tails, which loosens the
association between DNA and histones and allows for the activation of transcription.
17
Conversely, histone deacetylation refers to the removal of these acetyl groups, which
causes the repression of gene expression by tightening the DNA-histone complex. My
work has concentrated on both DNA methylation and histone deacetylation as means of
repressing transcription due to their association with the repressor protein MeCP2.
The study of epigenetics has focused mainly on its roles in cancer. Promoter
hypermethylation of tumor suppressor genes and altered levels of individual histone
modifications are found in many types of tumor cells. Outside of cancer, epigenetics has
received less attention. Recent accumulating evidence is pointing towards a role for
epigenetics in neurons. Some believe that these stable changes in gene expression might
be the underlying basis for learning and memory and long lasting forms of synaptic
plasticity in the brain. For example, extensive research has established multiple
interdependent activities of the transcriptional activator cyclic-AMP-response element
binding protein (CREB) and synaptic transmission. Expression levels of CREB have been
shown to regulate certain types of late long-term synaptic potentiation (LTP) and longterm depression (LTD) (Ahn et al., 1999). Recent data suggests that mutations in its
coactivator CREB binding protein (CBP), a histone acetyltransferase (HAT) enzyme that
can influence CREB function, exert profound effects in human brain by causing a form of
mental retardation, Rubinstein-Taybi syndrome (Petrij et al., 1995).
While there is ample research connecting transcriptional activation and synaptic
transmission, less attention has been focused on the role of transcriptional repression. The
association of mutations in the methyl-binding protein MeCP2 gene and the human form
of mental retardation called Rett Syndrome strongly suggests that transcriptional
repression plays an important role in the CNS. My thesis work has been concerned with
18
studying the role of transcriptional repression in the control of basal synaptic function,
with my focus centering on the repressor protein MeCP2.
19
MeCP2
The methyl-CpG-binding protein 2 (MeCP2) gene encodes a DNA binding
protein that belongs to a family of proteins that bind methylated cytosines in the
mammalian genome (Nan et al., 1996). The other members include methyl-binding
domain (MBD) proteins 1-4. MeCP2 acts as a transcriptional repressor by binding to
target gene promoters via its MBD and silences transcription via its transcriptional
repression domain (TRD) by recruiting additional silencing proteins such as Sin3a and
histone deacetylases (HDACs) 1 and 2 (Nan et al., 1998). MeCP2 protein is expressed at
high levels in the brain, specifically in neurons, not in glia, and correlates with neuronal
maturation (Akbarian et al., 2001; Shahbazian et al., 2002b). Recent evidence suggests
that MeCP2 is phosphorylated in response to synaptic activity (Chen et al., 2003; Zhou et
al., 2006). Under unstimulated conditions, MeCP2 associates with methylated CpG
dinucleotides in the promoters of target genes and recruits Sin3a and HDAC1 or 2. In
response to neuronal activity, the kinase CaMKII phosphorylates MeCP2 at serine 421,
causing its release from methylated promoters and relieving its repression of target genes
(Figure 1-1).
Figure 1-1. Transcriptional repression by MeCP2 is relieved following synaptic activity. In
the absence of activity, MeCP2 is bound to methylated CpG sequences in the promoters of
certain genes. The expression of these genes is repressed due the inability of transcriptional
activators to bind to promoter regions as well as to the deacetylation of histones by HDAC1 and
2 which are recruited to chromatin by MeCP2 and the corepressor Sin3a. Upon stimulation,
CaMKII phosphorylates MeCP2 causing release of the repressor complex from CpG islands,
allowing subsequent binding of transcriptional activators that can enhance gene expression.
20
A distinctive characteristic of MeCP2 is that, in humans, mutations in the MeCP2
gene result in a neurodevelopmental disorder called Rett Syndrome (RTT). RTT is a
childhood neurological disorder that accounts for one of the leading causes of mental
retardation and autistic behavior in females (Hagberg et al., 1983). In general,
individuals affected with RTT experience normal development up to the age of 6-18
months at which time they fail to acquire new skills and enter a period of motor skill
regression. With time, the RTT defects become more pronounced and include a wide
range of neurological defects (mental retardation, autism-like behavior, seizures,
disturbances of sleep, problems with gait, decelerated head growth, and stereotypical
hand movements). In addition, most children afflicted with RTT show a loss of social
and cognitive abilities. Recent work has demonstrated that RTT is an X-linked dominant
disorder that results from mutations in the MeCP2 gene found on the X chromosome
(Amir et al., 2000; Amir et al., 1999; Bienvenu et al., 2000; Huppke et al., 2000; Wan et
al., 1999). Classical RTT is found primarily in females because they are heterozygous for
MeCP2 mutations and the presence of a normal MeCP2 in approximately half of their
cells allows them to survive past birth. The amount of X chromosome inactivation in
these females accounts for the severity of an individual’s phenotypes (Shahbazian et al.,
2002c; Wan et al., 1999). There are a few cases of males that present classical symptoms
of RTT, but they are usually Klinefelter’s (XXY) or have greatly skewed somatic
mosaicism (Armstrong et al., 2001; Clayton-Smith et al., 2000; Leonard et al., 2001;
Schwartzman et al., 2001).
21
Figure 1-2. Common MeCP2 mutations in Rett Syndrome. Missense
(below) and nonsense (above) mutations are found both in and around the
functional domains of MeCP2.
In RTT patients, several mutations in the MeCP2 gene have been identified (Figure
1-2). These mutations can be missense mutations that result in single amino acid changes,
frameshift mutations that cause a shift in the reading frame of the gene, or nonsense
mutations that result in a truncated protein. Most of the mutations affect either the MBD
of MeCP2 that mediates its binding to DNA or the TRD that recruits co-repressor
complexes after DNA binding. The disease causing mutations found in RTT patients are
predicted to result in the loss of function of MeCP2 that then results in genes turned on in
an inappropriate manner. However, only the expression of a few genes have been shown
to be regulated by MeCP2, including brain-derived neurotrophic factor (BDNF) and the
imprinting genes Dlx5 and Dlx6, although the direct relationship between these changes
in gene expression and functional consequences in synaptic transmission have not been
shown (Chen et al., 2003; Horike et al., 2005). Microarray analysis of RTT patients and
mouse models of RTT have revealed little about possible genes whose expression levels
are controlled by MeCP2 (Colantuoni et al., 2001; Tudor et al., 2002). One study tried to
narrow down the possible list of MeCP2 target genes by suggesting that MeCP2
22
specifically binds to A/T rich sequences near methylated CpG dinucleotides, while
another methylated DNA binding protein, MBD2, does not (Klose et al., 2005). Using a
candidate gene approach, MeCP2 was found to control BDNF expression in mammalian
cultured cortical neurons, in an activity-dependent manner (Chen et al., 2003;
Martinowich et al., 2003). However, it is thought that there are a number of unidentified
gene targets of MeCP2 whose expression levels are subtly increased in the absence of
functional MeCP2 (Bienvenu et al., 2000). Therefore, how the loss of function of MeCP2
leads to the neurological phenotypes seen in RTT patients is of great interest.
Attempts to model the disease by generating constitutive MeCP2 knockout (KO)
mice results in the recapitulation of many of the neurological symptoms of RTT, although
these mice die early in postnatal development (Chen et al., 2001; Guy et al., 2001).
MeCP2-null mice develop normally until about 6 weeks of age and then begin to show a
variety of defective neuronal phenotypes, such as ataxic gait, tremors, and hindlimb
clasping (Guy et al., 2001). Studies examining the brains of RTT patients, as well as
mouse models of the disease, have not found any major neuropathological abnormalities
or neuronal loss but rather only subtle changes in neuronal morphology (Kaufmann and
Moser, 2000; Shahbazian et al., 2002a). To more fully characterize the role of MeCP2 in
the brain on behavioral phenotypes, our lab used conditional MeCP2 KO mice in which
MeCP2 was selectively deleted in broad forebrain regions of the brain during early
postnatal development (Gemelli et al., 2005). We found that these conditional MeCP2
KO mice have many of the behavioral abnormalities that are reminiscent of the
symptoms seen in RTT patients, including, impaired motor coordination, increased
anxiety, and abnormal social interaction with other mice. These data suggest that
23
expression of MeCP2 in postnatal neurons is crucial for normal development and that the
loss of MeCP2 in postnatal neurons of broad forebrain areas is sufficient to recapitulate
many features of RTT.
Based on the CNS-specific phenotypes of RTT, and the necessity of neuronal
expression of MeCP2, it is plausible that the mechanism underlying the disease
progression of RTT involves defects in neurotransmission in the CNS. Indeed, there is
evidence that individuals affected with RTT, as well as mouse models of the disorder,
have alterations in dendritic and synaptic spine structure indicative of a malfunction of
synaptic development and plasticity (Kaufmann and Moser, 2000). Recent work has
demonstrated that alterations in dendritic structure can result from changes in synaptic
activity (Zito and Svoboda, 2002). Therefore, these morphological changes in dendritic
structure of RTT patients, and of mouse models of the disease, may suggest an
underlying synaptic deficit.
Recent data suggests that MeCP2 may serve as a link between synaptic activity and
transcriptional regulation. Over the past couple of years, researchers have discovered a
number of defects in synaptic function in different mouse models of Rett Syndrome. It
appears that the loss of MeCP2 function can lead to changes in spontaneous synaptic
transmission as well as in short- and long-term synaptic plasticity. Brain slices from mice
overexpressing MeCP2 displayed an increase in paired pulse facilitation and long-term
potentiation (LTP) (Collins et al., 2004), while MeCP2 null mice exhibited the converse
(Asaka et al., 2006), suggesting that MeCP2 expression may contribute to functional
alterations in synaptic transmission resulting in the disease phenotype. Two studies, one
using an MeCP2-null mouse (Asaka et al., 2006) and another using a mouse expressing a
24
truncated form of MeCP2 (Moretti et al., 2006), found deficits in both LTP and long-term
depression (LTD) in hippocampal slices from these mice compared to littermate controls.
Interestingly, while the first study saw these changes only in older, symptomatic mice
(Asaka et al., 2006), the second found them also in younger, asymptomatic mice,
suggesting the possibility that these synaptic deficits may be occurring before the
manifestation of RTT-like behaviors (Moretti et al., 2006). Patients with RTT are normal
for the first 6-18 months of age, therefore much research is focusing on the role of
MeCP2 in neurodevelopment in hopes that an early intervention might lessen some of the
behavioral deficits.
Since microarray studies have been inconclusive in identifying large-scale gene
changes in RTT patients, and in mouse models of the disease, MeCP2’s role as a
transcriptional repressor has come into question. There is new evidence that MeCP2 may
also play a role in RNA splicing (Young et al., 2005). Another recent study found MeCP2
localized to the postsynaptic compartment, suggesting some additional role for MeCP2
outside of the nucleus (Aber et al., 2003). Despite this small amount of evidence to the
contrary, research continues to focus mainly on MeCP2’s role as a transcriptional
repressor, though no study has directly linked this function to the neurological deficits
seen in RTT patients and mouse models.
25
DNA Methylation
DNA methylation refers to the covalent addition of methyl groups to the C5position of cytosine residues within CpG dinucleotides. It is an important cellular
instrument by which cells control the repression of gene expression. Methylated DNA
can be used by the cell to exclude transcription factors important for activating gene
expression (Hark et al., 2000) or to recruit transcriptional repressor proteins like MeCP2
(Nan et al., 1996). In addition, methylation of DNA plays roles in processes such as X
chromosome inactivation, chromosome stability, and genomic imprinting.
Figure 1-3. DNA methylation and demethylation. DNMTs 3a and 3b add de novo methyl groups to
DNA. DNMT1 is responsible for copying these methyl groups onto newly synthesized DNA strands
after replication. Without DNMT1, DNA becomes passively demethylated during DNA replication.
Active demethylation is also believed to occur, though the enzyme responsible is still unknown.
26
DNA methyltransferases (DNMTs) are the enzymes responsible for adding
methyl groups to CpG islands found in mammalian genomic DNA. There are three main
DNMTs expressed in mammals: DNMT1, 3a and 3b. DNMT3a and DNMT3b are de
novo methyltransferases that establish methylation patterns at specific sites within the
genome (Okano et al., 1999). DNMT1 is responsible for the maintenance of these
methylation patterns during DNA replication (Hermann et al., 2004) (Figure 1-3). The
initiation signals for DNA methylation, and how DNMTs are targeted to specific gene
promoters remains unclear.
A few studies have found relationships between DNA methylation and histone
modifications. DNA methylation has been linked with histone H3 lysine 9 methylation in
silenced heterochromatin, though these modifications have been found to precede one
another in reverse order in different species (Johnson et al., 2002; Lehnertz et al., 2003).
Also, quite a few associations between DNMTs and HDACs have been discovered within
transcriptional corepressor complexes (Geiman et al., 2004; Rountree et al., 2000).
Nevertheless, there is still much to be discovered about what governs the control of DNA
methylation to specific CpG sites.
Much research has focused on DNA methylation changes during mammalian
development. Both male and female primordial germ cells undergo methylation
reprogramming via a putative active process of genome wide demethylation followed by
de novo remethylation carried out by DNMT3a (Hajkova et al., 2002; Kaneda et al.,
2004). In pre-implantation embryos, the paternal genome is reprogrammed by active
demethylation while the maternal is done by a passive demethylation process that is
dependent on DNA replication (Figure 1-3) (Haaf, 2006; Mayer et al., 2000). Once the
27
embryo reaches the blastocyst stage, genome-wide de novo methylation is performed by
both DNMT3a and 3b (Okano et al., 1999).
Following cellular differentiation, DNA methylation changes are less numerous
and thought to control the tissue-specific gene expression required to maintain the
identity of cells from one generation to the next. One example of this is seen with maspin,
a human cancer gene that is unmethylated in tumor cells that express it and methylated in
normal cells that don’t (Futscher et al., 2002). Evidence for DNA demethylation in
differentiated cells suggests only a passive mechanism by which methylation patterns are
lost during the DNA replication that occurs with each cellular division. The idea that an
active mechanism of demethylation can occur in a replication-independent manner, or in
non-dividing cells such as neurons, has garnered plenty of controversial attention.
However, evidence for this type of demethylation does exist. For example, demethylation
of transfected DNA into non-replicating cells has been shown to occur (Paroush et al.,
1990).
Many researchers have searched for a possible enzyme that can remove
established methylation patterns and thereby turn on specific gene expression. One group
discovered that the methyl-binding protein MBD2b can demethylate DNA by removing
the methyl moiety on cytosine residues and replacing it with a hydrogen atom
(Bhattacharya et al., 1999; Ramchandani et al., 1999), though this finding has been
strongly contested by others (Ng et al., 1999; Wolffe et al., 1999). Instead, most
demethylase supporters have focused on DNA damage and repair related-mechanisms. 5methylcytosine-DNA glycosylase can remove 5-methylcytosines in chick embryos,
allowing them to be replaced by unmethylated cytosines via DNA repair enzymes (Jost et
28
al., 1995), though there are many caveats to this reaction. The same group also
discovered that the protein MBD4 possesses this same demethylase activity (Zhu et al.,
2000), while another group has suggested that the function of MBD4 is to remove G-T
mismatched bases following 5-methylcytosine deamination (Hendrich et al., 1999). Most
recently, the growth arrest and DNA damage protein Gadd45α was discovered to promote
DNA repair and thereby erase methylation marks in non-dividing cells (Barreto et al.,
2007).
Interest in DNA methylation alterations in post-mitotic neurons has risen due its
association with a number of neurodevelopmental disorders. As previously mentioned,
mutations in the DNA methyl-binding protein MeCP2 result in Rett Syndrome, a form of
mental retardation in humans. Two other neurodevelopmental disorders that manifest
symptoms of mental retardation, Fragile X and ICF (Immunodeficiency, Centromeric
region instability, Facial anomalies) syndromes, arise from malfunctions in the
establishment of normal DNA methylation patterns. Fragile X mental retardation (FMR)
is brought about by the expansion of CGG or CCG triplet repeats in the promoters of the
FMR1 or FMR2 genes causing an increase in DNA methylation and therefore a decrease
in the expression of these genes (Turner et al., 1996). ICF syndrome is caused by
mutations in the DNMT3b gene resulting in decreased DNA methylation throughout the
genome (Hansen et al., 1999). Defects in DNA methylation have also been suggested to
play a role in schizophrenia, a serious cognitive disorder in humans. Evidence indicates
that deficiencies in the protein reelin, an extracellular matrix protein shown to play a role
in synaptic plasticity, may be responsible for the etiology of schizophrenia (Costa et al.,
29
2002). These deficiencies in reelin expression are believed to result from DNA
hypermethylation of the gene’s promoter region.
DNMTs are known to be highly expressed in adult brain neurons along with high
patterns of DNA methylation. DNMT mRNA expression and enzymatic activity levels
are both high in mature neurons (Brooks et al., 1996; Goto et al., 1994), although specific
DNMT isoforms are expressed at different levels in distinct areas of the brain. Both
DNMT1 and DNMT3a are expressed in post-mitotic neurons found in the olfactory bulb
(MacDonald et al., 2005). DNMT3a expression was also found in neurons of the cortex,
hippocampus, striatum, and cerebellum (Feng et al., 2005). And although one study
found DNMT1 expression in the adult cerebellum to be localized to the cytoplasmic
compartment (Inano et al., 2000), research continues to focus on the functions DNMTs
might have for controlling gene expression in the nuclei of mature, post-mitotic neurons.
Currently, there is little understood about the possible functions of DNMTs and
DNA methylation in mature, post-mitotic neurons. Some evidence points to them playing
a role in the control of long-term synaptic plasticity and memory formation. Treatment of
hippocampal slices with inhibitors of DNMT activity blocks both long-term potentiation
and memory formation following contextual fear conditioning, a commonly used test for
hippocampal-dependent associative learning and memory (Levenson et al., 2006; Miller
and Sweatt, 2007). Interestingly, DNMT expression and promoter methylation levels of a
specific learning and memory-suppressing gene, protein phosphatase 1 (PP1), are
upregulated following fear conditioning, while methylation of the reelin promoter is
decreased, suggesting that mechanisms controlling both DNA methylation and DNA
demethylation are altered in correlation with hippocampal learning and synaptic plasticity
30
(Miller and Sweatt, 2007). These studies suggest that DNA methylation plays an
important role in the control of synaptic transmission and indicate that changes can occur
in the DNA methylation patterns found in post-mitotic neurons in response to synaptic
activity. However, the exact mechanisms leading to these changes in DNA methylation
and their effects on synaptic function remain unclear, and whether or not they are actually
occurring in non-dividing neurons and not in the dividing glial cells surrounding the
neurons remains unresolved.
31
Histone Deacetylation
In the nucleus, DNA is wrapped around a core group of proteins called histones.
These proteins contain histone tails whose amino-termini of specific residues can receive
the covalent addition of a number of different post-translational modifications, such as
phosphorylation, methylation, ubiquitination, SUMOylation, and acetylation. The
addition of specific groups controls how tightly DNA is associated with histones, which
in turn defines how readily accessible gene sequences are to the transcriptional machinery
and other DNA-binding proteins. For example, acetylation of multiple lysine residues on
the tail of histone H4, acetylation of two lysine residues (9 and 14) on the tail of histone
H3, and dimethylation of lysine4 on H3 are all modifications commonly associated with
active gene transcription, while dimethylation of lysine 9 on H3 is associated with
transcriptionally silent chromatin (Turner, 2002). The combination of specific
modifications and their resulting effects on gene expression are collectively referred to as
the “histone code”. Many of the functions of these modifications are not completely
understood, however the detailed role of histone acetylation and deacetylation is more
apparent.
Histone acetylation occurs on the amino side chain of specific lysine residues
found within histone tails, thereby neutralizing their positive charge and interrupting their
association with negatively-charged DNA, allowing for the activation of transcription
(Figure 1-4). Histone acetyltransferases (HATs) are the enzymes responsible for utilizing
the cofactor acetyl-CoA in this acetylation reaction (Varga-Weisz and Becker, 1998).
Histone deacetylases (HDACs) repress transcription through removal of the acetyl group,
32
which then strengthens the histone-DNA interaction and blocks access of the
transcriptional machinery to the DNA template (Fischle et al., 2003).
Figure 1-4. Histone acetylation and deacetylation. Gene transcription is activated by
histone acetylation via histone acetyltransferases (HATs). Removal of the acetyl groups
by histone deacetylases (HDACs) results in repressed transcription.
There are 3 distinct classes of HDACs in mammalian cells, referred to as Class I,
II and III. Class I HDACs, which consist of HDACs 1, 2, and 3, are widely expressed.
Their localization within cells is strictly nuclear, their simple structures contain only a
catalytic domain, and they are frequently found to be associated with large corepressor
complexes. Class II HDACs (4, 5, 7, and 9) are a bit more sophisticated in that, in
addition to their catalytic domain, they contain an N-terminal extension that allows
interaction with coactivator and corepressor proteins and also includes serine/threonine
residues that can be phosphorylated by kinases in response to calcium. Most of the Class
II HDACs can be shuttled into and out of the nucleus in response to phosphorylation or
dephosphorylation (Fischle et al., 2001). The third class of HDACs is believed to be
primarily responsible for the deacetylation of tubulin in microtubules, which gives them a
very specific role in the modulation of cytoskeletal dynamics (Kovacs et al., 2004).
33
The association of Class I HDACs with corepressor protein complexes is widely
corroborated. Both HDACs 1 and 2 are associated with the corepressor, Sin3a, a protein
included in complexes with a number of different transcriptional repressors, such as Mad,
REST, and MeCP2. HDACs 1 and 2’s involvement with Mad is important for the control
of cellular proliferation by blocking cell cycle progression (Laherty et al., 1997). The
REST, Sin3a, and HDAC1/2 complex is important in nonneuronal cells for repressing
genes responsible for inducing a neuronal phenotype (Huang et al., 1999). The
association of HDACs 1 and 2 with MeCP2 is clearly defined (Nan et al., 1998), though
the functional consequences of transcriptional repression by these proteins are not
entirely clear. Evidence is pointing to this complex playing a role in everything from
cancer to neurodevelopment (Harikrishnan et al., 2005) (Amir et al., 1999). Therefore, it
is becoming increasingly apparent that HDACs can serve as transcriptional repressors
that modulate a variety of cellular functions.
While most HDACs are expressed ubiquitously, interest in the role of histone
acetylation and deacetylation in the brain has arisen due to a number of serendipitous
findings. First, it was discovered that a widely-used drug to treat epilepsy and bipolar
disorder, valproic acid (VPA), is also a potent inhibitor of HDACs (Phiel et al., 2001),
suggesting that inhibitors of these proteins, and likely the proteins themselves, play an
important role in the function of the CNS, perhaps in cellular excitability. Another link
between histone acetylation and the brain came from the discovery that Rubinstein-Taybi
syndrome, a form of mental retardation, is caused by mutations in the CREB-binding
protein (CBP), a protein known to have intrinsic HAT activity (Kalkhoven et al., 2003).
Finally, as was previously mentioned, mutations in the MeCP2 gene cause another form
34
of mental retardation called Rett syndrome, and it has been shown that part of MeCP2’s
function as a transcriptional repressor is dependent on the recruitment of HDACs 1 and 2
(Nan et al., 1998).
In neurons, histone acetylation has been shown to play roles in both neural
development and synaptic plasticity. The association of HDACs with the transcriptional
repressor REST has obvious implications towards having an effect on neuronal
development. Overexpression of REST in neuronal precursor cells represses the
expression of a voltage-gated Na channel, Nav1.2, and reduces neurite outgrowth,
demonstrating its important function in repressing the induction of a neuronal phenotype
in nonneuronal cells during development (Ballas et al., 2001). It has also been shown that
HDAC inhibitors can induce neuronal differentiation, suppress glial differentiation and
decrease proliferation in neural progenitor cells (Hsieh et al., 2004).
A role for HATs and HDACs in learning and memory and long-term synaptic
plasticity is quickly being established. To begin with, CBP heterozygous mice, models of
the human mental retardation disorder Rubinstein-Taybi syndrome, display deficits in
long-term memory after fear conditioning as well as in maintenance of the late phase of
LTP, known to be dependent on gene transcription (Alarcon et al., 2004). These defects
correlated with decreased acetylation of histone H2B, and the LTP deficit could be
rescued by treatment with HDAC inhibitors. Treatment of Aplysia sensorimotor neurons
with the HDAC inhibitor Trichostatin A (TSA) resulted in increased sensitivity for the
induction of long-term facilitation (Guan et al., 2002), while treatment of rodent
hippocampal slices caused an enhancement of LTP (Levenson et al., 2004). In addition,
HDAC inhibitors have been shown to augment the locomotor and reward responses to
35
cocaine while overexpression of HDAC4 in the striatum had the opposite effect (Kumar
et al., 2005), and HDAC5 overexpression in the hippocampus is able to block the
antidepressant effects of imipramine following defeat stress (Tsankova et al., 2006).
Though all of these studies point to a definite role for histone deacetylation in neuronal
function, there is currently no information about the roles of specific HDACs in synaptic
transmission. Therefore, there is still much to be discovered about the functional
consequences of epigenetic alterations, in particular histone acetylation and DNA
methylation, in neurons.
36
CHAPTER 2
MECP2-DEPENDENT TRANSCRIPTIONAL REPRESSION REGULATES
EXCITATORY NEUROTRANSMISSION
Introduction
Mutations in the transcriptional repressor, methyl-CpG-binding protein 2 (MeCP2), result
in a neurodevelopmental disorder called Rett Syndrome (RTT) (Amir et al., 2000; Amir
et al., 1999; Bienvenu et al., 2000; Huppke et al., 2000; Wan et al., 1999). RTT is an Xlinked dominant disorder in which defects are predominantly expressed in the central
nervous system, including stereotypical hand movements, mental retardation, autismrelated behavior, seizures, disturbances of sleep, and problems with gait (Hagberg et al.,
1983). The MeCP2 gene encodes a DNA binding protein that binds to methylated
cytosines in the mammalian genome. Normally, MeCP2 acts as a transcriptional
repressor by binding to target gene promoters and silencing their transcription through the
recruitment of additional silencing proteins such as Sin3a and histone deacetylases
(HDACs) 1 and 2. The disease causing mutations found in RTT patients are predicted to
result in the loss of function of MeCP2 that then results in genes turned on in an
inappropriate manner (Ballestar et al., 2000; Yusufzai and Wolffe, 2000). It is intriguing
how mutations in MeCP2, a gene present in many tissues, leads to the wide array of
neurological phenotypes observed in RTT patients. Studies examining the brains of RTT
patients, as well as recent mouse models of the disease, have not found any major
neuropathological abnormalities or neuronal loss but rather only subtle changes in
neuronal morphology (Kaufmann and Moser, 2000; Shahbazian and Zoghbi, 2002).
37
Attempts to model the disease by generating MeCP2 knockout (KO) mice results in the
recapitulation of many of the neurological symptoms of RTT (Chen et al., 2001; Guy et
al., 2001).
There is evidence that individuals affected with RTT have alterations in dendritic
and synaptic spine structure indicative of a malfunction of synaptic development and
plasticity (Kaufmann and Moser, 2000). Recent work has demonstrated that alterations
in dendritic structure can result from changes in synaptic activity (Zito and Svoboda,
2002). Therefore, it is possible that the MeCP2 mutations contribute to functional
alterations in synaptic transmission resulting in the disease phenotype. Based on the
neurological phenotypes observed in Rett patients, we examined the potential role of
MeCP2 in synaptic function. We compared elementary properties of synaptic
transmission between cultured hippocampal neurons from MeCP2 knockout and wild
type littermate control mice and found a decrease in the frequency of spontaneous
excitatory synaptic transmission (mEPSCs) in neurons lacking MeCP2. We also detected
a significant increase in the rate of short-term synaptic depression. To explore whether
these functional effects can be attributed to MeCP2’s role as a transcriptional silencer, we
treated cultures with a drug that impairs histone deacetylation and examined spontaneous
synaptic transmission. Treatment with this compound induced a similar decrease in
mEPSC frequency in wild type control cultures but this decrease was occluded in
MeCP2-deficient neurons. Interestingly, neither the loss of MeCP2, nor the drug
treatment resulted in changes in mIPSC properties. Finally, using a lentivirus expressing
Cre recombinase, we show that loss of MeCP2 function after neurodevelopment and
synaptogenesis was sufficient to mimic the decrease in mEPSC frequency seen in
38
constitutive MeCP2 KO neurons. Taken together, these results suggest a role for MeCP2
in control of excitatory presynaptic function through regulation of gene expression.
39
Results
To elucidate the role of MeCP2 in the regulation of synaptic transmission, we studied
functional alterations of synapses in 11-14 days in vitro hippocampal cultures made from
newly born MeCP2 KO mice. Recent electrophysiological measurements of synaptic
plasticity in MeCP2-deficient mice were performed on both hippocampal and cortical
slices (Asaka et al., 2006; Dani et al., 2005). Dissociated primary cultures allow
examination of synaptic function independent of potential general alterations in brain
homeostasis, thus enabling a distinction between cell autonomous defects and global
systemic dysfunction. We quantified the frequency and amplitude of spontaneous
miniature excitatory postsynaptic currents (mEPSCs) in MeCP2 KO and wild type (WT)
littermate control cultures using whole-cell recordings performed in the presence of
tetrodotoxin (TTX) to block action potential firing and picrotoxin to block inhibitory
activity. We found a significant decrease in the frequency of mEPSCs in the KO neurons
compared to WT controls (Figure 2-1A-C). The decrease was also observed in older
cultures (>20 DIV) (Figure 2-2), suggesting that the loss of MeCP2 produces long-term
alterations in excitatory synaptic transmission. This alteration in mEPSC frequency may
implicate a presynaptic deficit in the MeCP2 KO neurons. In contrast, the frequency of
spontaneous miniature inhibitory postsynaptic currents (mIPSCs) was unaffected,
suggesting a specificity of MeCP2 function in excitatory neurotransmission (Figure 2-1EG). The amplitudes of individual synaptic events in both mEPSCs and mIPSCs were also
unaffected by the loss of MeCP2, indicating no potential change in the number of
postsynaptic receptors at either excitatory or inhibitory synapses (Figure 2-1D, H).
40
To better understand how the loss of MeCP2 may contribute to the alteration in
mEPSC frequency, we examined the number of presynaptic terminals formed on
pyramidal neuron dendrites in culture. Neurons were immunostained for microtubuleassociated protein (MAP2) and Synapsin, a synaptic vesicle protein, to identify
presynaptic terminals on the dendrites and soma. This analysis revealed that the number
of presynaptic terminals were unchanged in MeCP2 KO neurons compared with WT
neurons, suggesting that the decrease in spontaneous synaptic events is not the result of a
decreased number of presynaptic terminals (Figure 2-1I, J).
We next examined whether there was a decrease in the size of the total recycling
pool and the number of readily releasable vesicles. To probe the total pool size, we
stimulated cultures with 47 mM K+ solution for 90 seconds in the presence of the styryl
dye FM1-43 (Betz et al., 1996). This strong stimulation normally labels all recycling
vesicles within a presynaptic terminal (Harata et al., 2001). After dye wash out (~10
minutes), we stimulated the cultures with 90 mM K+ solution applied four times, the first
for 90s followed by three applications of 60s each (each separated by 60s intervals), to
release all the dye trapped within presynaptic terminals. The kinetics of dye loss from
synaptic terminals (Figure 2-3A), and the total amount of dye trapped in individual
synapses was indistinguishable between KO and control neurons (Figure 2-3A insert),
indicating that the sizes of the total vesicle pools were unchanged. To quantify
differences in the number of readily releasable vesicles, we stimulated cultures with a
brief hypertonic sucrose application, which selectively releases vesicles from the readily
releasable pool (Rosenmund and Stevens, 1996). We found no difference between the
MeCP2 KO and WT neurons, suggesting that the changes in spontaneous release
41
frequency in the KO cultures cannot be attributed to a reduction in the number of readily
releasable vesicles (Figure 2-3B and C). However, we cannot fully exclude the possibility
that the numbers of spontaneously recycling vesicles are reduced, but we consider this
unlikely given that sizes of distinct vesicle pools are usually highly correlated in central
synapses (Sara et al., 2005).
We next examined the properties of evoked neurotransmission in response to
action potential stimulation. Trains of action potentials applied at 10 Hz typically depress
neurotransmission. This depression is, at least in part, elicited by a rapid decrease in the
number of vesicles available for release within a synaptic terminal. When the kinetics of
synaptic depression and recovery in the MeCP2 KO cultures were examined, we
observed a more rapid depression during 10 Hz stimulation and a slower response
recovery when the stimulation frequency was switched to 1 Hz at the end of the 10 Hz
train, compared to WT cultures (Figure 2-4A, C, and D). The KO cultures also showed a
slight increase in first response amplitudes and significantly smaller paired pulse ratios
during high stimulation frequencies compared to controls (Figure 2-4B, C insert). These
findings suggest that the loss of MeCP2 may contribute to an increase in neurotransmitter
release probability leading to a faster depletion of releasable vesicles, as well as a delay
in synaptic vesicle recycling after stimulation retarding the recovery of responses. Taken
together with the decrease in spontaneous event frequency, these findings suggest a role
for MeCP2 in presynaptic control of neurotransmitter release and vesicle recycling
specifically in excitatory synapses.
To examine whether the alterations in excitatory neurotransmission of neurons
lacking MeCP2 are the result of impairments in gene silencing, we treated wild type
42
C57BL/6 hippocampal cultures chronically with drugs that inhibit either transcriptional
activation or repression and then measured synaptic transmission. To suppress
transcription, we treated cultures with the RNA polymerase inhibitor Actinomycin D (Act
D). To suppress transcriptional repression, we used Trichostatin A (TSA), a histone
deacetylase inhibitor. Cultures were treated with these individual drugs (and DMSO as a
control) for 24 hours and then synaptic activity was measured. We preferred chronic
rather than acute treatments because earlier studies did not reveal a significant change in
baseline synaptic transmission after acute TSA application, although they reported a
strong augmentation of long term synaptic plasticity (Levenson et al., 2004). We initially
examined cell viability in the presence of these chronic treatments and found that it was
not compromised (see Experimental Procedures). In these experiments, we focused on
potential alterations in the frequency and amplitude of spontaneous miniature events,
which is a more direct measurement of presynaptic machinery, since evoked transmission
may be vulnerable to alterations in membrane excitability and Ca2+ signaling, two aspects
of neuronal function that may also be targeted by transcriptional regulation. We found a
significant decrease in mEPSC frequency in TSA treated cultures compared to DMSO
treated cultures, while Actinomycin D did not have an effect (Figure 2-5A and B). The
average amplitudes of individual events were not significantly affected by these
treatments (data not shown). We also determined the number of presynaptic terminals in
these cultures and found no changes between the control and TSA treatments (Figure 25C and D). Similarly to what was seen with the MeCP2 KO cultures, both the frequency
and amplitudes of mIPSCs were unchanged between the DMSO and TSA treated neurons
(Figure 2-5E, F and data not shown). To investigate whether newly transcribed genes are
43
involved in the suppression of synaptic function, we treated cultures with both
Actinomycin D and TSA and found a reversal of the mEPSC deficits seen in the TSA
treated neurons (Figure 2-5B). These findings indicate a selective impairment of
excitatory presynaptic function after suppression of transcriptional repression but not
transcriptional activation.
To examine whether the TSA-mediated decrease in mEPSCs was related to
MeCP2 function, we chronically treated MeCP2 KO cultures with TSA and did not detect
a reduction in the frequency of spontaneous mEPSCs (Figure 2-5G and H). This result
strongly suggests that the decrease in spontaneous miniature frequency we observed in
the presence of this transcriptional activator was in significant part due to inhibition of
MeCP2 function and thus was occluded in the absence of MeCP2. Interestingly,
Actinomycin D did not rescue the reduction in mEPSC frequency seen in MeCP2 KOs
back to control levels (Figure 2-5G and H). This may not be surprising since degradation
of abnormally expressed presynaptic proteins may take longer than 24 hours.
We also investigated whether an acute loss of MeCP2 function after
neurodevelopment and synapse maturation would result in similar phenotypes found in
the constitutive MeCP2 KO neurons. We made primary dissociated hippocampal cultures
from newborn floxed MeCP2 mice and allowed them to age 7 days in vitro before
infecting them with high or low titer lentivirus expressing the gene Cre recombinase, or
GFP as a control (Figure 2-6). One week later, we found a significant decrease in the
frequency of mEPSCs in high titer Cre-infected floxed MeCP2 neurons compared to
GFP-infected neurons (Figure 2-7A and B). This significant frequency decrease was also
seen when mEPSCs from Cre-infected floxed MeCP2 neurons were compared with Cre-
44
infected wild type littermate neurons, ruling out a nonspecific effect of Cre expression on
mEPSCs (Figure 2-7A and B). These data suggest that MeCP2 acts as a regulator of
synaptic transmission even in mature neurons and the loss of MeCP2 may have profound
effects on synaptic function after neurodevelopment. We also recorded mEPSCs from
uninfected floxed MeCP2 neurons receiving the majority of their inputs from Creinfected neurons and found the same decrease in event frequency, supporting the
presynaptic origin of this observation (Figure 2-7A and B). In accordance with this
premise, mEPSC frequency was not altered in floxed MeCP2 neurons infected with a low
titer Cre-expressing lentivirus compared to GFP controls (Figure 2-7D and E). This data,
as well as the fact that we saw no significant changes in mEPSC amplitudes among any
of the lentiviral-infected cultures (Figure 2-7C and F), strongly suggests that MeCP2
plays a specific role in presynaptic function.
45
Discussion
Taken together, these findings suggest that synaptic transmission, in particular
presynaptic function, is under transcriptional control and that MeCP2-dependent
transcriptional repression is a critical component of this regulation. In our experiments,
we detected MeCP2 dependent alterations in spontaneous neurotransmission as well as in
short-term synaptic depression. Spontaneous neurotransmission is important for a number
of neuronal processes, which include maturation and stability of synaptic networks
(McKinney et al., 1999; Zucker, 2005) and inhibition of local dendritic protein synthesis
(Sutton et al., 2004). Therefore, the decrease in mEPSCs we describe here may well
underlie some of the neuromorphological abnormalities seen in RTT patients as well as
RTT mouse models (Chen et al., 2001; Guy et al., 2001; Kaufmann and Moser, 2000;
Shahbazian and Zoghbi, 2002). However, our data also suggest genes involved in evoked
transmission and short-term plasticity as potential transcriptional targets under MeCP2
control. Short-term synaptic depression is a fundamental synaptic mechanism, which
underlies key brain functions such as sound localization and sensory adaptation (Abbott
and Regehr, 2004; Chung et al., 2002; Cook et al., 2003). Therefore, changes in synaptic
depression may have important implications for the synaptic basis of RTT, as well as
other neurodevelopmental disorders.
Recent evidence suggests that synaptic vesicles giving rise to evoked and
spontaneous neurotransmission originate from different pools (Sara et al., 2005).
Therefore it is possible to envision that simultaneous changes in expression levels of
multiple synaptic proteins, which are under MeCP2 regulation, may have opposing
effects (or a complex additive effect) on the two forms of neurotransmission.
46
Understanding the mechanisms underlying these synaptic alterations will require
identification of synaptic molecules that are MeCP2-dependent in their transcription.
Could the alterations in synaptic transmission produced by mutations in the
MeCP2 gene underlie the behavioral phenotypes observed in RTT patients? Recent
studies have shown that the loss of MeCP2 selectively in the brain is sufficient to
recapitulate many features of RTT including impaired motor coordination, increased
anxiety-related behavior and social deficits (Gemelli et al., 2005). In related studies,
brain slices from mice overexpressing MeCP2 displayed an increase in paired pulse
facilitation and long-term potentiation (LTP) (Collins et al., 2004), while MeCP2 null
mice exhibited the converse (Asaka et al., 2006), suggesting MeCP2 expression may
exert a profound role on synaptic plasticity. Our data indicates a specific deficit in
excitatory synaptic transmission upon the loss of MeCP2 function. A recent study of
cortical slices from 4-5 week old MeCP2 knockout mice also found changes in
excitatory, as well as inhibitory, transmission (Dani et al., 2005). These additional
inhibitory changes may be specific for cortical neurons, or they may just occur later in
development. Nevertheless, it appears that an imbalance in excitatory and inhibitory
input, as we see in hippocampal cultures, may underlie some of the neurological deficits
in RTT, and possibly other related neurodevelopmental disorders. Indeed, other recent
studies have shown that the neuroligin genes, a family of proteins implicated in autism
spectrum disorders, are important for maintaining a balance of excitatory and inhibitory
neurotransmission (Chih et al., 2004; Chih et al., 2005).
It is worthwhile noting that we did not observe any morphological changes in
neurons lacking MeCP2. This may well be due to the fact that our studies were done in
47
young cultures and may reflect a more immature neuronal population. This suggests that
the functional deficits we observed in MeCP2 KO neurons may preclude the neuronal
structural abnormalities observed in the disease state.
These data suggest that transcriptional repression is important in regulating
presynaptic function of hippocampal neurons, even after neurodevelopment. This
provides insight into how the loss of function of the transcriptional repressor, MeCP2,
may contribute to the disease state and may have profound implications, especially if
additional neuronal populations are under MeCP2 control. This information is important
in delineating the cellular and functional abnormalities that lead to the wide array of
neurological deficits observed in RTT patients.
48
Figure 2-1. Spontaneous miniature
synaptic currents in cultured
hippocampal neurons from MeCP2
knockout and control mice
(A-D) Spontaneous excitatory synaptic
currents. Representative recordings of
miniature excitatory events in control (A)
and MeCP2 knockout (B) neurons
recorded in 1 μM TTX and 50 μM
picrotoxin. (C) Bar graph showing a
decrease in the frequency of spontaneous
excitatory events in knockout neurons
compared to controls (*; p<0.05). (D)
Cumulative histogram of mEPSC
amplitudes. (E-H) Spontaneous
inhibitory synaptic currents.
Representative recordings of miniature
inhibitory events in control (E) and
MeCP2 knockout (F) neurons recorded in
1 μM TTX and 10 μM NBQX. (G) Bar
graph showing similar frequencies of
spontaneous inhibitory events in control
and knockout neurons. (H) Cumulative
histogram of mIPSC amplitudes. (I and
J) Immunostaining of cultured neurons.
(I) Dissociated neurons from control and
knockout mice were labeled with primary
antibodies to MAP2 (green) and
Synapsin (red). (J) Bar graph depicts the
number of presynaptic terminals found in
control and mutant neurons. There was
no significant difference in the number of
presynaptic terminals between control
and MeCP2 knockout neurons (p>0.2).
49
Figure 2-2. Miniature EPSC frequency is reduced in older MeCP2 KO neurons
(A) Representative recordings of miniature excitatory events in control and MeCP2
knockout neurons >20 DIV. (B) Bar graph showing a decrease in the frequency of
spontaneous excitatory events in older knockout neurons compared to controls (*;
p<0.05).
50
Figure 2-3. Total pool and readily releasable pool of synaptic vesicles from MeCP2
knockout and control neurons
(A) Fluorescence destaining of MeCP2 knockout and control synapses in response to
high K+ stimulation. Synapses were loaded with FM1-43 by 47 mM K+ induced
depolarization and destained using 90 mM K+. Insert, bar graph depicts the total vesicle
pool sizes in control and mutant synapses measured by the total change in fluorescence
during 90 mM K+ destaining. (B and C) Synaptic responses to hypertonic sucrose
stimulation. (B) Representative traces from control and knockout neurons showing
synaptic responses to 500 mM sucrose. (C) Bar graph depicting the average amplitudes of
the peak currents during hypertonic sucrose stimulation.
51
Figure 2-4. Evoked synaptic responses in MeCP2 knockout and control neurons
during 10 Hz field stimulation immediately followed by 1Hz recovery stimulation
(A) Representative whole-cell recordings of the first 20 responses. (B) Paired pulse
ratios of the first two responses during various stimulation frequencies were measured
from control and knockout neurons. The paired pulse ratios recorded from MeCP2
knockout neurons were significantly less than control at both 30 Hz and 10 Hz
frequencies (*, p<0.05, **, p<0.01). (C) Average response amplitudes of control and
mutant neurons measured during the first two seconds of 10 Hz stimulation. Recorded
responses from field stimulated knockout neurons depressed significantly faster than
control responses (*, p<0.05, **, p<0.01). Insert is a bar graph depicting similar average
first peak amplitudes of these responses. (D) Average response amplitudes measured
during 1 Hz stimulation following the 10 Hz depression. Knockout response amplitudes
did not recover as quickly as those recorded from control neurons (*, p<0.05).
52
Figure 2-5. Spontaneous miniature
synaptic responses from wild type
C57BL/6 and MeCP2 knockout
hippocampal cultures after a 24-hour
treatment with inhibitors of
transcriptional repression and
activation
(A and B) Spontaneous excitatory
synaptic currents from C57BL/6 neurons.
(A) Representative miniature excitatory
synaptic currents from drug treated
neurons recorded in 1 μM TTX and 50
μM picrotoxin. (B) Bar graph of the
significant decrease in mEPSC
frequencies in neurons treated for 24
hours with the inhibitor of transcriptional
repression, TSA (*; p<0.05). This
decrease was reversed when treatment
included both TSA and the inhibitor of
transcriptional activation, Act D. (C and
D) Immunostaining of C57BL/6 cultured
neurons. (C) Control and TSA treated
dissociated neurons were labeled with
primary antibodies to MAP2 (green) and
Synapsin (red). (D) Bar graph depicting a
similar number of presynaptic terminals
found in control and TSA treated
neurons. (E and F) Spontaneous
inhibitory synaptic currents from
C57BL/6 neurons. (E) Representative
recordings of miniature inhibitory events
from control and TSA treated neurons
recorded in 1 μM TTX and 10 μM
NBQX. (F) Bar graph showing the
frequencies of spontaneous miniature
inhibitory events. (G and H) Spontaneous
excitatory synaptic currents from MeCP2
knockout neurons. (G) Representative
traces from drug treated knockout
neurons recorded in the presence of 1 μM
TTX and 50 μM picrotoxin. (H) Bar
graph showing that mEPSC frequencies
in knockout neurons were not
significantly affected by 24-hour
treatment with either Act D or TSA.
53
Figure 2-6. High and low titer lentiviral infection of dissociated hippocampal
cultures
(Top) Light and fluorescent microscopic images and an overlay of the two
demonstrate the low titer (<20% of neurons infected) Cre lentivirus used to
knockdown the expression of MeCP2 in floxed MeCP2 neurons. (Bottom) Light,
fluorescent and overlaid images of neurons infected with the high titer (>80%
infected) Cre lentivirus.
54
Figure 2-7. Spontaneous miniature excitatory synaptic currents in floxed MeCP2 neurons
infected with a lentivirus expressing Cre recombinase
(A-C) Miniature excitatory events from floxed neurons infected with high titer lentivirus. (A)
Representative traces from infected neurons recorded in the presence of 1 μM TTX and 50 μM
picrotoxin. (B) Bar graph revealing a significant decrease in mEPSC frequencies from floxed
MeCP2 neurons infected after synapse formation with high titer lentiviral Cre recombinase (*;
p<0.05). This decrease was seen in recordings from both Cre infected and uninfected postsynaptic
floxed neurons and is compared to both floxed neurons infected with GFP and wildtype neurons
infected with Cre. (C) Cumulative histogram showing no significant changes in mEPSC
amplitudes. (D-F) Miniature excitatory currents from floxed neurons infected with low titer
lentivirus. (D) Representative traces of mEPSCs recorded from infected neurons. (E) Bar graph
revealing similar mEPSC frequencies from floxed MeCP2 neurons infected after synapse
formation with low titer lentiviral Cre recombinase (*; p<0.05). (F) Cumulative histogram showing
no significant changes in mEPSC amplitudes.
55
CHAPTER 3
ACTIVITY-DEPENDENT SUPRESSION OF EXCITATORY MINIATURE
NEUROTRANSMISSION THROUGH THE REGULATION OF DNA
METHYLATION
Introduction
DNA methylation is a key cellular mechanism used to repress gene expression and
promote genome stability in various species. In mammals, methylation of DNA plays
roles in many processes, such as X chromosome inactivation, genomic imprinting and the
control of tissue-specific gene expression. DNA methyltransferases (DNMTs) are the
enzymes responsible for adding methyl groups at the 5-position of cytosine residues
within CpG dinucleotides. During development, widespread methylation changes occur
in primordial germ cells and pre-implantation embryos (Jaenisch and Bird, 2003; Reik et
al., 2001). After differentiation, alterations in DNA methylation are less abundant and
primarily control local gene expression to preserve cellular identity. These methylation
changes are commonly believed to occur only in actively dividing cells, where specific
methylation sites on genomic DNA are established and maintained by DNMTs during
replication. More recently however, the Gadd45a protein was discovered to play a role in
active demethylation of DNA in proliferating as well as non-dividing cells (Barreto et al.,
2007).
The role of DNA methylation in neurons has recently become of interest due to
alterations in the methylation status of genes associated with a number of mental
retardation syndromes including, Rett, ICF and Fragile X (Robertson and Wolffe, 2000).
56
DNMTs are highly expressed in neurons in the adult brain suggesting they may have a
functional role in postmitotic neurons (Brooks et al., 1996; Feng et al., 2005; Goto et al.,
1994; Inano et al., 2000). Methylcytosine analogs, including 5-azacytidine, can inhibit
DNA methylation in many cell types (Robertson and Jones, 2000). Recent studies suggest
that inhibiting DNA methylation in hippocampal slices blocks long-term synaptic
plasticity (Levenson et al., 2006). In other studies, prolonged depolarization of cultured
cortical neurons has been reported to result in a decrease in methylation in the promoter
region of BDNF, a neurotrophin important for synaptic plasticity (Martinowich et al.,
2003). These studies suggest that there may be a relationship between synaptic activity
and DNA methylation in mature neurons. However, the mechanisms behind these
changes in DNA methylation and synaptic activity are unknown. We are interested in
looking more closely at the role of DNA methylation in synaptic function, therefore we
used an array of pharmacological approaches to impair DNMT activity in hippocampal
neurons and tested basic properties of neurotransmission using electrophysiological and
imaging techniques.
57
Results
To examine whether alterations in DNA methylation influence synaptic transmission,
hippocampal neurons were cultured from newborn C57BL/6 mice, matured (13-21 DIV)
(Mozhayeva et al., 2002), and then treated for 24 hours with two different DNMT
inhibitors, the methylcytosine analogs 5-azacytidine (5azaC, 2.5 μM) and Zebularine
(Zeb, 50 μM) (Tawa et al., 1990). Whole-cell voltage clamp recordings were performed
on cultured hippocampal neurons immediately following 24-hour chronic treatments.
Spontaneous miniature excitatory postsynaptic currents (mEPSCs) were measured in
neurons treated with 5azaC, Zeb or vehicle (DMSO). These recordings were done in the
presence of TTX to block action potential firing and picrotoxin to block inhibitory
activity. We found a significant decrease in the frequency of mEPSCs in both the 5azaC
and Zeb treated neurons compared to controls, but no change in the amplitudes of these
events (Fig. 3-1A-C). The changes in mEPSC frequency, but not amplitudes, point to a
possible presynaptic deficit following DNMT inhibition. To test whether this effect on
spontaneous activity is specific for excitatory synapses, miniature inhibitory postsynaptic
currents (mIPSCs) were recorded in neurons in the presence of TTX and the AMPA-type
glutamate receptor blocker, NBQX. No changes were observed in the frequency or
amplitudes of mIPSCs in neurons with either 5azaC or Zeb treatments compared to
controls, suggesting that DNMT inhibition is affecting only excitatory synaptic activity
(Fig. 3-1D-F). The fact that both DNMT inhibitors caused similar deficits in excitatory
synaptic transmission suggests that inhibiting DNA methylation produces specific
alterations in synapse function and not generalized, global nonspecific effects.
58
The deficits in mEPSC frequency produced by DNMT inhibition could be caused
by decreases in excitatory synapse number and/or release probability from excitatory
presynaptic terminals. Therefore, the number of functional excitatory synapses formed
onto cultured hippocampal neurons was assessed following 24 hours of DNMT
inhibition. Neurons were immunostained for Synapsin, a presynaptic vesicle protein, and
PSD-95, a postsynaptic scaffolding protein found specifically at excitatory synapses. The
analysis of colocalized Synapsin and PSD-95 revealed that the number of excitatory
synapses was unaffected by treatment with either 5azaC or Zeb, indicating that the
decreases in spontaneous mEPSCs are not the result of a decreased number of excitatory
inputs (Fig. 3-1G, H).
We also examined the properties of evoked excitatory neurotransmission in
response to action potential stimulation to determine if there were deficits in release
probability after inhibition of DNMT activity. After treatment with 5azaC or Zeb for 24
hours, there were no changes in excitatory postsynaptic response depression during a 10
Hz field stimulation (Fig. 3-2A, B) or in the paired pulse ratios of these responses
recorded at various stimulation frequencies (Fig. 3-2C). These observations are consistent
with a previous experiment which did not find effects of these drugs on basal synaptic
transmission or short term synaptic plasticity in hippocampal slices (Levenson et al.,
2006). These findings suggest that inhibition of DNA methyltransferases by
methylcytosine analogs specifically affects the release probability of spontaneous
miniature excitatory events.
To further explore the presynaptic properties of these neurons, we used FM dye
imaging to measure aspects of both evoked and spontaneous synaptic vesicle recycling.
59
To label all recycling vesicles within presynaptic terminals, cultures were stimulated with
47 mM K+ solution for 90 seconds in the presence of the styryl dye FM1-43 (Betz et al.,
1996; Harata et al., 2001). After a brief wash, the kinetics of release of these vesicles was
measured by destaining synapses with 90 mM K+ solution for 90 seconds. The rates of
dye loss from total vesicle pools in control and DNMT inhibitor treated synapses was
similar (Fig. 3-2D), as were the total amounts of dye trapped within individual synapses
(Fig. 3-2E), indicating that the sizes of the total vesicle pools were unchanged. We then
assessed the properties of spontaneously recycling vesicles from neurons treated with
inhibitors of DNMT activity. Vesicles were loaded for 15 minutes in the presence of TTX
to block activity, washed, and then the destaining of individual synapses was measured
for 20 minutes, again in the presence of TTX to block synaptic activity. Treatment with
5azaC resulted in slower destaining kinetics compared to controls (Fig. 3-2F), suggesting
a specific deficit in spontaneous vesicle fusion following DNMT inhibition. The total
amount of dye released during destaining was similar between control and 5azaC treated
synapses (Fig. 3-2G), implying that the number of vesicles that recycle spontaneously
was unaffected. Together, these results support a specific role for DNA methylation in
the control of spontaneous vesicle release from presynaptic terminals.
The molecular substrate for the DNMT methylation reaction is S-adenosyl-Lmethionine (SAM). The interaction of methylcytosine analogs with DNA
methyltransferases interferes with the enzymes’ abilities to transfer methyl groups from
this endogenous substrate to cytosines in CpG dinucleotides. However, in the presence of
excess SAM, DNMTs can actually methylate 5azaC-containing DNA (Gabbara and
Bhagwat, 1995). Therefore, we surmised that the addition of excess SAM may reverse
60
the DNMT inhibitor-induced deficits in spontaneous synaptic transmission. The
previously seen decrease in mEPSC frequency after 24-hour 5azaC and Zeb treatments
was no longer seen after the addition of SAM (Fig. 3-3A). The proposed functions of
5azaC and Zeb are to inhibit DNMT-dependent methylation of appropriate CpG
dinucleotides thus resulting in the improper transcription of specific genes. Therefore, the
effects of these two drugs should be dependent on active gene expression. To investigate
whether the deficit in mEPSC frequency caused by inhibition of DNMT activity is
dependent on transcription, we treated cultures with the RNA polymerase inhibitor,
Actinomycin D (Act D), in combination with either 5azaC or Zeb. Act D treatment was
able to block the decrease in mEPSC frequency seen in neurons treated with DNMT
inhibitors (Fig. 3-3B). These results demonstrate that the defects in miniature excitatory
event frequency seen after blocking the ability of DNMTs to add methyl groups to DNA
is, in fact, a result of increased gene transcription and not some nonspecific effect of the
drugs.
In mammalian cells, one primary role of DNA methylation is to promote the
binding of transcriptional repressor proteins to the promoters of specific genes. One such
DNA methyl-binding protein is methyl-CpG binding protein 2 (MeCP2). Mutations in the
MeCP2 gene are associated with the neurological disorder, Rett syndrome. We
previously reported a defect in excitatory neurotransmission in cultured hippocampal
neurons from MeCP2-deficient mice, of which included a specific decrease in the
frequency of mEPSCs (Nelson et al., 2006). We were intrigued by the similar decrease in
mEPSC frequency seen after DNMT inhibition, suggesting a putative shared mechanism
for the control of excitatory spontaneous transmission by DNA methylation and MeCP2.
61
To investigate this possibility, primary hippocampal cultures were made from newborn
MeCP2 knockout mice and mature cultures were treated with DNMT inhibitors. No
further reduction in the frequency of mEPSCs from these neurons was detected after the
addition of either 5azaC or Zeb (Fig. 3-3C, D), effectively denoting that the loss of
MeCP2 function, either in the knockout or by eliminating DNA methylation, is the
primary cause of this deficit. These results also suggest that the effects of DNMT
inhibition occur in post-mitotic neurons since MeCP2 is not expressed in glial cells
(Shahbazian et al., 2002b).
Next, we examined the possibility that the methyl donor, SAM, may also alleviate
the synaptic defects seen in MeCP2-deficient neurons. A decrease in mEPSC frequency
was still seen after a 24-hour treatment with the DNMT substrate, however, after 48
hours, SAM was able to rescue the defect in spontaneous event frequency seen in MeCP2
knockout neurons (Fig. 3-3C, D). The need for a longer exposure to S-adenosyl-Lmethionine is not surprising since previous work has shown that an increase in DNA
methylation does not occur until after >36 hours treatment with SAM (Noh et al., 2005).
The transcriptional repressor, MeCP2, is the only methyl-binding protein that can bind to
singly methylated CpG dinucleotides (Lewis et al., 1992), while other proteins containing
a methyl-binding domain (MBDs) require additional sites (Hendrich and Bird, 1998). It is
possible that the addition of excess SAM leads to an increase in the number of
methylated CpGs on specific promoters, allowing for other MBDs to bind in the absence
of MeCP2. We saw no effect of SAM treatment, for either 24 or 48 hours, on wildtype
neurons (Fig. 3-3A, C, D), suggesting that MeCP2’s binding to sufficiently methylated
promoter regions has already repressed the expression of genes responsible for
62
suppressing synaptic function. The ability of SAM to rescue the synaptic deficits in the
MeCP2 knockout is intriguing and future studies will be important to determine whether
SAM can rescue the behavioral deficits in mouse models of Rett Syndrome.
Our data, thus far, support a role for DNA methylation in synaptic function, which
overlaps with MeCP2’s role as a transcriptional repressor. To ensure that the DNMT
inhibitors were indeed effective at blocking methylation, we employed the use of
methylation-sensitive restriction endonucleases on genomic DNA extracted from 24-hour
treated hippocampal cultures. Using the technique, NotI-MseI methylation-sensitive
amplified fragment length polymorphism (MS-AFLP) we looked for decreases in the
methylation patterns of total genomic DNA compared to controls (Yamamoto et al.,
2001). NotI is a restriction enzyme that cleaves unmethylated DNA in CpG islands, or
areas rich in G+C content; NotI is unable to cleave these sites when methylated. With
NotI-MseI MS-AFLP, hypomethylation of DNA is depicted by an increase in band
intensity from a fluorescent DNA fingerprint covering >95% of the entire genome. A
comparison of the methylation patterns from 5azaC and Zeb treated cells revealed that
~1% of the increased band intensities overlapped (Fig. 3-4A, B and data not shown).
Though these changes are small, they demonstrate the successful demethylation of
genomic DNA by DNMT inhibitors. The modest size of this decrease in methylation is
consistent with the extensive stability of overall DNA methylation patterns in postmitotic
neurons in the adult brain.
Recently, MeCP2’s function was shown to be susceptible to activity-dependent
regulation via phosphorylation in response to Ca2+ (Chen et al., 2003; Zhou et al., 2006).
There is recent evidence that DNA methylation can also be regulated by neuronal
63
activity. A commonly used learning and memory paradigm, contextual fear conditioning,
increases the levels of DNMT enzymes in the brain and alters promoter methylation
patterns in two genes associated with synaptic plasticity (Miller and Sweatt, 2007),
however the molecular signaling mechanisms that drive these changes in DNA
methylation are still not understood. To examine whether DNA methylation indeed forms
an additional target for activity-dependent regulation of gene expression, we focused on
the BDNF gene, due to its earlier association of high potassium depolarization mediated
demethylation in its promoter region (Martinowich et al., 2003). After exposing
dissociated hippocampal cultures to a chronic, 24-hour treatment with the DNMT
inhibitor, 5azaC, we isolated genomic DNA and processed it for bisulfite modification.
Using quantitative Real-Time PCR with primers specific for a CpG island located in the
promoter region for exon 1 of the BDNF gene (Fig. 3-5), we found a significant increase
in unmethylated DNA after 5azaC treatment compared to vehicle controls (Fig. 3-6A, B),
suggesting that demethylation of this BDNF promoter occurs in mature, hippocampal
cultures. We hypothesized that the spontaneous network activity present in these mature
cultures (Virmani et al., 2006) may exert control over BDNF expression by regulating
methylation at the BDNF promoter I. Therefore, we measured the methylation status of
the CpG island after a 24-hour co-treatment with 5azaC and either tetrodotoxin (TTX) or
2-amino-5-phosphonopentanoic acid (AP5). The increase in demethylation after 5azaC
treatment was prevented when either evoked synaptic activity was blocked with TTX or
when NMDA receptor-activity was blocked using AP5 (Fig. 3-6A, B). We also measured
the amount of unmethylated BDNF in hippocampal cultures treated with SAM in
combination with 5azaC. The addition of SAM was able to reverse the effects of 5azaC
64
on DNA methylation (Fig. 3-6A, B), indicating a rescuing effect of the substrate in the
presence of DNMT inhibition. These results demonstrate that spontaneous network
activity present in these cultures is sufficient to render the BDNF promoter susceptible to
demethylation and alter BDNF expression. The mechanism by which neuronal activity
regulates the methylation status of genes, such as BDNF, may be that activity makes
DNA susceptible to demethylation by causing damage/repair or perhaps facilitates
putative demethylation activity.
To determine whether the deficit in synaptic transmission caused by DNMT
inhibitors is dependent on activity, mEPSCs were recorded from cultures silenced with
TTX during the entire 24 hours they were exposed to either 5azaC or Zeb. In the absence
of action potential firing, treatment with these DNMT inhibitors no longer resulted in
decreased frequency of spontaneous miniature excitatory events (Fig. 3-6C). Thus, the
ability of DNA methylation to influence synaptic transmission requires neuronal activity,
suggesting that the methylation status of genes that play a role in controlling synapse
function may be regulated by synaptic activity.
A well-known mechanism by which synaptic activity can drive changes in gene
expression is by calcium influx through postsynaptic N-methyl-D-aspartate-type
(NMDA) glutamate receptors (Bito et al., 1997). Since we observed a requirement of
synaptic activity for DNA methylation-induced alterations in synaptic function, we tested
the importance of NMDA receptor-dependent calcium signaling in this regulation. We
exposed our cultures to the NMDA receptor antagonist, AP5, and measured the effects of
DNMT inhibition on mEPSCs. The previously seen decrease in mEPSC frequency after
treatments with either 5azaC or Zeb was blocked in the presence of AP5 (Fig. 3-6D),
65
suggesting that calcium influx through NMDA receptors is, at least in part, responsible
for the effects DNMT inhibition has on excitatory synaptic transmission. Taken together,
these data indicate that the regulation of DNA methylation in mature neurons is
dependent on neuronal activity and NMDA receptor-dependent calcium signaling, and
that these activity-dependent changes in DNA methylation play a fundamental role in
controlling basal synaptic function.
Finally, we assayed whether the changes in methylation of the BDNF promoter I
resulted in changes in BDNF mRNA expression. Using quantitative Real-Time PCR with
primers specific to BDNF exon 5, the mRNA coding exon of the gene, we determined the
levels of BDNF expression in our cultures following treatment with the DNMT inhibitor,
5azaC. Interestingly, there was no change in the level of BDNF expression following 24
hour treatment with 5azaC (Figure 3-7), as would not have been expected given the
increase in BDNF promoter demethylation seen previously. However, 5azaC treatment
for 48 hours did lead to a significant increase in BDNF mRNA levels compared to
controls. This treatment also resulted in demethylation of BDNF promoter I (data not
shown). One caveat of this experiment is that BDNF mRNA expression is controlled by
four different promoters, I-IV, in mice. We may not have been able to detect changes in
expression due to differential regulation of these three promoters, at least not until longer
exposure to 5azaC possibly leads to additional decreases in promoter methylation. Future
analyses of methylated cytosines within the different BDNF promoters following DNMT
inhibition should help clarify these discrepancies. Next, we further tested the role of
synaptic activity in controlling changes in BDNF expression in response to alterations in
DNA methylation by combining the 5azaC treatments with either TTX or AP5. Both the
66
blocker of action potentials and the antagonist of NMDA receptors significantly
decreased the expression of BDNF (Figure 3-7). These results were expected given that
there is ample evidence indicating an activity-dependent regulation of BDNF gene
expression through the regulation of promoters III or IV (Tao et al., 1998). Again, our
assay cannot differentiate between specific promoters driving changes in BDNF
expression. In conclusion, our studies suggest a novel mechanism by which synaptic
activity can drive an increase in BDNF transcription through the demethylation of BDNF
promoters.
67
Discussion
Collectively, our findings suggest that spontaneous synaptic transmission between
postmitotic neurons is regulated by alterations in DNA methylation that occur in response
to synaptic activity. Spontaneous synaptic currents are known to be important for
controlling synapse maturation and the stability of neuronal networks (McKinney et al.,
1999; Zucker, 2005). In addition, spontaneous neurotransmission can be an important
read-out for homeostatic mechanisms such as synaptic scaling (Kilman et al., 2002;
Turrigiano et al., 1998). In response to 24-hour DNMT inhibition, we found a significant
decrease in the frequency of mEPSCs, which correlates with a decrease in BDNF
promoter I methylation. Interestingly, an increase in BDNF mRNA expression was seen
only after longer exposure to 5azaC treatment. Previous work has established a role for
BDNF in the induction of homeostatic plasticity. A chronic decrease in endogenous
BDNF signaling in cortical cultures increases the intrinsic excitability of pyramidal
neurons, a similar effect to what occurs following chronic activity blockade (Desai et al.,
1999). In our hands, the deficit in neurotransmission in response to DNMT inhibition is
seen prior to the increase in BDNF expression. This suggests that the regulation of
homeostatic plasticity can act in two ways: neurons can respond to decreases in the
expression of normal activators of synaptic plasticity, like BDNF, and consequently
increase excitatory synaptic function, or they can respond to decreases in
neurotransmission by increasing the expression of genes that will enhance neuronal
excitability.
We also demonstrate that both effects on synaptic transmission and BDNF
promoter methylation are dependent on the spontaneous neuronal activity occurring in
68
our cultures. This activity-dependent regulation is likely via increased postsynaptic
calcium influx through NMDA receptors and presumably leads to the demethylation of
specific gene promoters. Alas, the presence of an active demethylating enzyme in
neurons has yet to be discovered which leads to the possible conclusion that activity is
driving demethylation through a process utilizing DNA damage and repair machinery
(Brooks et al., 1996). Nevertheless, there is additional support for activity-driven changes
in DNA methylation. Both increases and decreases in DNA methylation patterns of two
genes involved in synaptic plasticity were found in the hippocampus following fear
conditioning (Miller and Sweatt, 2007). While this study does not explore a molecular
mechanism controlling activity-dependent alterations in DNA methylation, the
association of learning and memory behaviors with postsynaptic signaling cascades
activated by synaptic inputs implicates potential pathways that may be able to drive these
changes.
Furthermore, our studies demonstrate an intimate relationship between DNA
methylation in neurons and the transcriptional repressor MeCP2. The deficit seen in
spontaneous synaptic transmission following 5azaC and Zeb treatments was occluded in
the absence of MeCP2, the loss of which causes a similar defect in synapse function
(Nelson et al., 2006). In addition, treatment of MeCP2 KO neurons with the methyl donor
SAM was able to reverse this decrease in miniature neurotransmission. An interesting
future avenue to pursue is the possible affects of SAM treatment on many of the
behavioral deficits seen in MeCP2-deficient mice. These findings suggest that MeCP2 is
the methyl-binding protein responsible for mediating the effects of DNA methylation on
69
neuronal function. Our results also have implications towards treatment of RTT patients
with SAM in an attempt to ameliorate some of their neurological symptoms.
In conclusion, our data indicate a role for DNA methylation in the control of
synaptic function, which shares a common pathway with the methyl-binding protein,
MeCP2. Furthermore, our data suggest that neuronal activity can drive the transcription
of genes important for controlling spontaneous neurotransmitter release by regulating the
methylation status of these genes (Fig. 3-8). Previous work has demonstrated that activity
can also induce the phosphorylation of MeCP2, causing its dissociation with target genes
and relieving its repression of transcription (Chen et al., 2003). Activity-dependent loss
of DNA methylation may be an additional mechanism by which MeCP2 is released from
the promoters of target genes. Our results concerning DNA methylation and its impact on
neurotransmission may suggest a homeostatic mechanism by which neuronal nuclei can
monitor alterations in activity levels and adjust neurotransmitter output via altering gene
expression and thus impact network excitability.
70
Figure 3-1. Inhibiting DNMT activity in neurons causes a deficit in excitatory
synaptic transmission.
(A) Sample traces of miniature EPSCs recorded from mature hippocampal cultures
treated 24 hours with DNMT inhibitors, 5azaC and Zeb. (B) Bar graph showing a
decrease in the frequencies of mEPSCs recorded from 5azaC and Zeb treated neurons
compared to controls (**p<0.01; *p<0.05). (C) Histograms of mEPSC amplitudes show
no differences among control, 5azaC, or Zeb treatments. (D) Sample recordings of
miniature IPSCs from cultures treated with inhibitors of DNMT activity. (E) Bar graph
depicts no change in mIPSC frequencies from 5azaC or Zeb treated neurons compared
with controls. (F) Cumulative histograms of mIPSC amplitudes show no differences
between control and DNMT inhibitor treated neurons. (G) Dissociated hippocampal
cultures were immunostained with antibodies to Synapsin (red) and PSD-95 (green) to
determine the number of excitatory synapses. (H) Bar graph shows no alterations in
excitatory synapse number among control, 5azaC, or Zeb treated cultures.
71
Figure 3-2. DNMT inhibition for 24 hours specifically affects spontaneous
presynaptic function.
(A) Normalized sample traces of the first 20 evoked EPSCs in response to 10 Hz field
stimulation from neurons treated with inhibitors of DNMTs. (B) Average normalized
EPSC amplitudes from treated neurons measured during 20 sec of 10 Hz stimulation
show no alterations in response depression after treatment with DNMT inhibitors
compared to controls (DMSO n=15, 5azaC n=14, Zeb n=12). (C) Paired pulse ratios of
the first two evoked EPSCs in response to various stimulation frequencies were not
significantly different in DNMT inhibitor treated neurons compared with controls. (D)
Total synaptic vesicle pools were loaded with FM1-43 by 47 mM K+ induced
depolarization and destained using 90 mM K+. The kinetics of destaining were not
different between control and 5azaC treated neurons. (E) Bar graph depicts no change in
total recycling synaptic vesicles from control and 5azaC treated synapses measured by
the total change in fluorescence during 90 mM K+ destaining (DMSO n=8 coverslips,
5azaC n=8). (F) Spontaneously recycling synaptic vesicles were loaded for fifteen
minutes in the presence of TTX and destained for twenty minutes in the same manner.
Destaining of spontaneous vesicles was slower in 5azaC treated synapses compared to
controls (**p<0.01). (G) Bar graph indicates no difference in the numbers of
spontaneously recycling synaptic vesicles between control and DNMT inhibitor treated
neurons (DMSO n=6, 5azaC n=5).
72
Figure 3-3. Changes in spontaneous excitatory neurotransmission after DNMT
inhibition are mediated by the loss of function of the transcriptional repressor,
MeCP2.
(A) Bar graph demonstrating that cotreatments with DNMT inhibitors and the methyl
donor, SAM, or treatment with SAM alone, results in no alterations in mEPSC frequency
compared with controls. (B) Bar graph revealing no changes in mEPSC frequencies when
neurons were treated with DNMT inhibitors in combination with the inhibitor of
transcriptional activation, Act D, suggesting that gene transcription is required for the
deficit seen with DNMT inhibitor treatment alone (black bar +/- gray S.E.M.). (C)
Representative traces of mEPSCs recorded from MeCP2 deficient neurons. (D) Bar graph
showing a decrease in mEPSC frequency in MeCP2 knockout neurons that is not
significantly affected by 24-hour treatments with 5azaC, Zeb or SAM. However, a 48hour application of SAM onto MeCP2 deficient neurons was able to reverse this
frequency deficit (**p<0.01; *p<0.05).
73
Figure 3-4. 24-hour treatment of hippocampal cultures with DNMT inhibitors
reveals demethylation of genomic DNA.
(A) Example NotI-MseI methylation-sensitive amplified fragment length polymorphism
(MS-AFLP) fingerprints of genomic DNA samples from control and DNMT inhibitor
treated neurons. (B) Plot profiles of average band intensities from MS-AFLP example in
(A) (n=2 for each treatment). Blue arrows indicate increases in band intensities with both
5azaC and Zeb treatments compared with controls. Red arrowheads indicate examples
where band intensities increased with 5azaC treatment, but not Zeb.
74
Figure 3-5. Schematic of the CpG island in the BDNF gene.
After bisulfite modification of genomic DNA, primers specific for a CpG island located
in the promoter region for exon I were used with quantitative Real-time PCR to measure
the amount of unmethylated DNA.
75
Figure 3-6. Treatment of hippocampal cultures with DNMT inhibitors reveals
activity-dependent demethylation of DNA and concurrent alterations in synaptic
transmission.
(A) Bisulfite modification of genomic DNA followed by Quantitative PCR to measure
levels of unmethylated BNDF promoter I. Representative gel electrophoresis of PCR
products following control and 5azaC treatments ± TTX, AP5, or SAM. (B) Bar graph
shows a 4-fold increase in the level of unmethylated BDNF promoter I following 5azaC
treatment compared to controls (black bar ± gray S.E.M.), but no changes in the level of
demethylation when 5azaC treatments were in combination with TTX, AP5, or SAM
(*p<0.05). (C) Bar graph shows no changes in mEPSC frequency when cultures were
treated with DNMT inhibitors in the presence of TTX, suggesting that neuronal activity is
required for the decrease in frequency seen with DNMT inhibitor treatment alone (black
bar +/- gray S.E.M.). (D) Bar graph indicates no difference in the frequencies of mEPSCs
when 5azaC and Zeb treatments included the NDMA antagonist, AP5.
76
Figure 3-7. BDNF mRNA expression after chronic manipulation of DNA
methylation in hippocampal cultures.
Expression of BDNF was unchanged in cultures treated 24 hours with the DNMT
inhibitor, 5azaC, compared to controls. However, after 48 hr exposure to 5azaC, BDNF
mRNA levels were significantly increased. Blocking activity with TTX, or calcium influx
through NDMA receptors using AP5, caused a significant decrease in the amount of
BDNF mRNA compared to controls (*, p<0.05; **, p<0.01; ***, p<0.001).
77
Figure 3-8. Model for activitydependent demethylation of
genomic DNA in post-mitotic
neurons.
(A) In the absence of activity,
MeCP2 is able to bind to
methylated CpG dinucleotides
and repress the transcription of
target genes. (B) In the presence
of activity, demethylation occurs
(possibly through DNA
damage/repair) and MeCP2 can
no longer bind to the promoter
unless remethylation occurs via
DNMT activity. (C) In the
presence of activity and 5azaC,
DNMTs become covalently
bound to the methylcytosine
analog and can no longer
methylate the promoter. (D) In the
presence of excess SAM, 5azaC
can be methylated by the
covalently bound DNMT enzyme,
thereby allowing MeCP2 to bind
and repress transcription. (E) In
MeCP2 deficient neurons, the
addition of excess SAM results in
multiple methylated sites at the
promoter that allows the binding
of other MBD transcriptional
repressors.
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CHAPTER 4
LOSS OF HDAC1 AND HDAC2 IN HIPPOCAMPAL NEURONS RESULTS IN
SPECIFIC ALTERATIONS IN EXCITATORY SYNAPTIC TRANSMISSION
Introduction
Post-translational modifications of chromatin, the complex of DNA wound around a core
group of proteins called histones, are fundamental in controlling gene expression in all
eukaryotes. These modifications can occur on cytosine nucleotides within the DNA itself
or on numerous amino acid residues found in histone tails. The addition of different
combinations of these modifications leads to either the activation or repression of gene
transcription, a system that is commonly referred to as the “histone code” (Turner, 2002).
Acetylation of histones, via histone acetyltransferase (HAT) activity, allows for the
activation of gene expression by interfering with the electrophilic interaction between
histones and DNA that then permits binding of the transcriptional machinery to specific
gene promoters (Varga-Weisz and Becker, 1998). Conversely, the removal of acetyl
groups by histone deacetylases (HDACs) results in the tighter compaction of DNA within
chromatin, which therefore leads to the repression of gene transcription.
There are 3 distinct classes of HDACs in mammals. Class I HDACs, consisting of
HDACs 1, 2, and 3, are known to interact with multiple corepressor complexes, like those
containing Mad, REST, and MeCP2, via their direct binding to Sin3a (Huang et al., 1999;
Laherty et al., 1997; Nan et al., 1998). The presence of particular corepressor proteins
within these complexes dictates the repression of specific target genes. Class II HDACs
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(4, 5, 7, and 9) are unique in that they are shuttled back and forth from the nucleus to the
cytoplasm in response to activity-driven phosphorylation or dephosphorylation (Fischle
et al., 2001). Finally, the Class III HDACs are responsible for the deacetylation of
tubulin, among other proteins, and are thereby believed to regulate the cytoskeletal
dynamics of cells (Kovacs et al., 2004).
Most HDACs are expressed ubiquitously within an organism, and histone
acetylation and deacetylation have been shown to play important roles in a number of
biological disorders, from multiple forms of cancer to neurodegenerative diseases such as
Huntington’s. It is increasingly becoming apparent that HAT and HDAC activities are
fundamental for normal brain function. In humans, mutations in the histone
acetyltransferase CREB-binding protein (CBP) cause Rubinstein-Taybi syndrome (RTS),
a disorder associated with mental retardation (Kalkhoven et al., 2003). Mouse models of
RTS, in particular mice heterozygous for CBP, show increased acetylation of histone
H2B that correlates with decreases in both long-term memory and synaptic plasticity
(Alarcon et al., 2004). Interactions of HDACs1 and 2 with the corepressor complex
containing MeCP2 (Nan et al., 1998), a gene in which mutations lead to the
neurodevelopmental disorder Rett syndrome, suggest an important role for histone
deacetylases in the brain. MeCP2 KO mice have defects in excitatory spontaneous
neurotransmission and short and long-term synaptic plasticity (Dani et al., 2005; Moretti
et al., 2006; Nelson et al., 2006). They also display deficits in behavior tests of learning
and memory, as well as anxiety and social interaction (Gemelli et al., 2005; Moretti et al.,
2006).
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Recent studies suggest that small molecule inhibitors of HDAC activity can have
profound effects on neuronal function. In humans, valproic acid (VPA) is commonly
utilized for patients in order to treat both epilepsy and bipolar disorder (Phiel et al.,
2001). Treatment of rodent hippocampal slices with Trichostatin A (TSA) results in
enhanced long-term potentiation (LTP) (Levenson et al., 2004), while TSA treatment of
dissociated hippocampal cultures causes a decrease in excitatory spontaneous synaptic
currents, an effect that was occluded in MeCP2 KO neurons (Nelson et al., 2006). Since
most research regarding HDAC activity in the brain has been done using these broadscale inhibitors, evidence for the functions of specific HDAC proteins in neurons is
considerably lacking. Using mice, two studies were able to demonstrate that
overexpressing HDAC4 in the striatum can have effects on cocaine reward (Kumar et al.,
2005), while HDAC5 overexpression in the hippocampus can alter the effects of
antidepressants on stress (Tsankova et al., 2006). Unfortunately, many questions are left
unanswered about the exact functions these HDACs are carrying out in those particular
areas of the brain. Are they somehow controlling cellular excitability via the regulation of
genes involved in synaptic transmission or membrane polarity? If so, how is their
deacetylase activity being regulated? In an attempt to delve further into the functional
roles for histone deacetylation in neurons, we have measured various aspects of synaptic
function in hippocampal neurons lacking specific HDAC proteins. Given our previous
interest in MeCP2 and the association of HDACs 1 and 2 with the MeCP2 transcriptional
repressor complex, we have focused on these two HDACs and their potential roles in the
regulation of synapse function.
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Results
We previously discovered a role for HDAC function in the control of spontaneous
excitatory synaptic transmission using the HDAC inhibitor TSA (Nelson et al., 2006). To
further support this finding, we treated dissociated hippocampal cultures for 24 hours
with an additional inhibitor of HDAC activity, Valproic acid (VPA), and then used
whole-cell voltage clamp electrophysiology to measure miniature excitatory postsynaptic
currents (mEPSCs). Both TSA and VPA treatments resulted in a specific decrease in the
frequency of mEPSCs, but had no effects on event amplitudes (Figure 4-1A, B). To
determine if HDAC inhibition had any additional effects on synapse function, we
measured the short-term synaptic plasticity of TSA treated neurons in response to 10 Hz
field stimulation. Chronic treatment with TSA caused a significantly faster depression of
evoked EPSCs, indicating an increase in the release probability of evoked synaptic
vesicles following HDAC inhibition (Figure 4-1C). To be certain that TSA and VPA
were indeed inhibiting HDAC activity, we measured the amount of histone acetylation
following 24-hour treatment with these drugs. Both HDAC inhibitors caused a significant
increase in the amount of acetylated histone H4 (AcH4) immunoreactivity (Figure 4-1D,
E), indicating that the effects of these drugs on synaptic transmission are being mediated
by increases in histone acetylation.
Following this initial investigation of synaptic properties in response to treatment
with broad scale HDAC inhibitors, we went on to explore the roles of specific HDACs in
synapse function. Due to our previous discovery of a relationship between HDAC
inhibition and the loss of MeCP2’s effects on synaptic transmission, we decided to focus
our study on HDACs 1 and 2, the two HDACs known to interact with MeCP2 in a
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transcriptional repressor complex (Nan et al., 1998). Since constitutive HDAC1 and
HDAC2 knockout mice die during embryonic development, we decided to use a viralmediated approach to knockout these individual HDACs, which would also allow us to
study the importance of these proteins after neurodevelopment. To investigate whether an
acute loss of HDAC function would result in alterations in synaptic transmission, we
made primary dissociated hippocampal cultures from newborn floxed HDAC1 or
HDAC2 mice and allowed them to age 7 days in vitro before infecting them with a
lentivirus expressing the gene Cre recombinase, or GFP as a control. One week later, we
assayed the amount of HDAC1 and 2 mRNA levels in both knockout cultures. In Creinfected floxed HDAC1 (KO) neurons, we found a significant decrease in HDAC1
mRNA levels, indicating successful knockout of the gene (Figure 4-2A). Furthermore,
the expression level of HDAC2 mRNA was unchanged, arguing against any
compensation for the loss of HDAC1 (Figure 4-2B). In floxed HDAC2 (KO) neurons
infected with Cre-lentivirus, we successfully induced a significant decrease in HDAC2
mRNA and saw no difference in the level of HDAC1 mRNA compared to controls
(Figure 4-2A, B). We also looked at protein levels in our cultures using western blotting
techniques. We found a significant decrease of HDAC1 in HDAC1 KO cultures and
HDAC2 protein was significantly decreased in the HDAC2 KO cultures (Figure 4-2C,
D).
After establishing that we had induced substantial knockdowns of HDAC1 and 2,
we went on to measure various aspects of synaptic transmission. We found a significant
decrease in the frequency of spontaneous mEPSCs in HDAC2 KO neurons compared to
GFP-infected neurons, but not in HDAC1 KO neurons. (Figure 4-3A, B). The amplitudes
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of these events were unchanged in both HDAC1 and 2 KO cultures (Figure 4-3C). The
alteration in mEPSC frequency may implicate a presynaptic deficit in the HDAC2 KO
neurons and suggests that HDAC2 may play more of a role than HDAC1 in the control of
excitatory spontaneous transmission in hippocampal neurons. A recent report has
demonstrated higher HDAC2 expression, compared to HDAC1 levels, in the
hippocampus (DG), suggesting that HDAC2 may be the predominant histone deacetylase
in hippocampal neurons (Cassel et al., 2006). In addition, both control and HDAC1 and 2
KO cultures were treated for 24 hours with TSA to determine the effect of this broadscale HDAC inhibitor on mEPSC properties within each genetic background. Similarly to
the previous TSA results, treatment with this HDAC inhibitor caused a significant
decrease in mEPSC frequency in all three cultures compared to untreated controls
(Figure 4-3A, B). The fact that we saw a small, but more severe, decrease in the HDAC2
KO after TSA treatment suggests that there may be additional HDACs causing this
deficit, or perhaps that the knockdown of HDAC2 was not complete enough to cause the
same level of decrease in event frequency.
To better understand how the loss of HDAC2 may contribute to the alteration in
mEPSC frequency, we examined the number of functional excitatory synapses formed
onto cultured hippocampal neurons after acute knockdown of HDAC2. Neurons were
immunostained for Synapsin, a presynaptic vesicle protein, and PSD-95, a postsynaptic
scaffolding protein found specifically at excitatory synapses. The analysis of colocalized
Synapsin and PSD-95 revealed that the number of excitatory synapses was unaffected by
the loss of HDAC2, indicating that the decreases in spontaneous mEPSCs are not the
result of a decreased number of excitatory inputs (Fig. 4-3D, E).
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To determine whether the loss of either HDAC1 or HDAC2 resulted in deficits in
inhibitory synaptic transmission, we measured miniature inhibitory postsynaptic currents
(mIPSCs) from these neurons. The frequencies of mIPSCs were unaffected by the loss of
either HDAC1 or HDAC2, suggesting a specificity for HDAC2’s function in excitatory
spontaneous neurotransmission (Figure 4-4A, B). The amplitudes of individual synaptic
events were also unaffected by the loss of HDAC1 and HDAC2, indicating no potential
change in the number of postsynaptic receptors at inhibitory synapses (Figure 4-4C).
We next examined the properties of evoked neurotransmission in response to
action potential stimulation. We analyzed the amplitudes and paired pulse ratios of
EPSCs evoked with a number of different stimulation frequencies from the conditional
HDAC1 and HDAC2 KO cultures, as well as from constitutive MeCP2 KO neurons for
comparison. Both HDAC1 and 2 KO cultures showed a significant increase in first
response amplitudes and significantly smaller paired pulse ratios at the 20Hz stimulation
frequency compared to controls (Figure 4-5A-C). Interestingly, these results are very
similar to those seen in the constitutive MeCP2 KO neurons. MeCP2 deficient neurons
showed a significantly larger first EPSC amplitude and significantly smaller paired pulse
ratios at 20, 10 and 5 Hz stimulation frequencies (Figure 4-5A-C) It is not surprising that
the viral-mediated knockdowns of HDAC1 and HDAC2 did not cause the same severity
of deficits as the constitutive loss of MeCP2 since they were knocked down after
neurodevelopment. Taken together with the decrease in spontaneous event frequency,
these findings suggest a role for both HDAC1 and 2 in the presynaptic control of
neurotransmitter release at excitatory synapses.
85
To be certain of this specific effect on excitatory neurotransmission, we analyzed
the amplitudes and paired pulse ratios of evoked IPSCs from the HDAC1, HDAC2, and
MeCP2 KO cultures. No differences in IPSC amplitudes or paired pulse ratios were seen
as a result of the loss of any of these components of the transcriptional repressor complex
(Figure 4-6A-C). Therefore, it appears that the loss of either HDAC1 or HDAC2 results
in specific defects in excitatory synaptic function, very similar to those seen after the
constitutive KO of MeCP2. These data suggest that the transcriptional repression
complex containing MeCP2 and HDACs 1 or 2 acts to regulate excitatory synaptic
transmission in mature neurons and that the loss of these proteins may have profound
effects on synaptic function after neurodevelopment.
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Discussion
New studies are beginning to reveal a role for histone deacetylation in the central nervous
system. The availability of HDAC inhibitors has allowed for simple but important
discoveries concerning histone deacetylation and its implications in brain function.
Treatment with these compounds can enhance long-term memory and LTP (Fischer et al.,
2007; Levenson et al., 2004) and decrease spontaneous excitatory synaptic transmission
in mice (Nelson et al., 2006). We have further extended these findings with the discovery
that TSA can induce a faster synaptic depression of EPSCs in response to train
stimulation, indicating that decreases in HDAC activity, and therefore increases in
histone acetylation, can also effect basal synaptic transmission. Major concerns of these
studies arise from the fact that these are nonspecific HDAC inhibitors, and they give no
clear indication as to the molecular mechanisms behind the effects on synaptic function.
While there has been a small amount of work done looking at the roles of Class II
HDACs in animal behavior (Kumar et al., 2005; Tsankova et al., 2006), no studies have
yet to explore the regulation of synaptic transmission by specific HDACs. We decided to
focus our research on the Class I HDACs 1 and 2 due to their associations with the
corepressor protein MeCP2 (Nan et al., 1998). MeCP2’s function in the brain is clearly
significant given that mutations in the gene lead to a form of mental retardation, Rett
syndrome, and that MeCP2 KO mice show deficits in spontaneous neurotransmission and
short- and long-term synaptic plasticity (Dani et al., 2005; Moretti et al., 2006; Nelson et
al., 2006). Interestingly, the loss of MeCP2 causes a decrease in LTP while HDAC
inhibition results in enhanced LTP (Levenson et al., 2004; Moretti et al., 2006), but both
are believed to result in the increased expression of certain genes important for
87
controlling synapse function. Again, these differences could be due to the nonspecificity
of the HDAC inhibitors used that would result in increased transcription of a number of
genes, many of which may not be regulated by MeCP2 function.
To reconcile these differences, we measured spontaneous and evoked synaptic
transmission in neurons lacking either HDAC1 or 2, the two HDACs known to bind
MeCP2. We used a viral-mediated KO approach for these studies, which allowed us to
determine the roles of these HDACs in mature synapse function, separate from any
neurodevelopmental defects that might be caused by the loss of these proteins at an
earlier time point. With respect to spontaneous synaptic activity, the loss of HDAC2, but
not HDAC1, resulted in a decrease in the frequency of mEPSCs. This specific alteration
in mEPSC frequency in the HDAC2 KO neurons suggests that HDAC2 may play more of
a role than HDAC1 in the control of excitatory spontaneous transmission in hippocampal
neurons. In fact, HDAC2 mRNA expression is more prevalent in the hippocampus than
HDAC1 (Cassel et al., 2006). However, evoked synaptic transmission was effected by the
loss of both HDACs 1 and 2, resulting in increased EPSC amplitudes and decreased
paired pulse ratios at high stimulation frequencies. All of these defects were specific for
excitatory synaptic activity as both miniature and evoked IPSCs were unaltered in
HDAC1 or 2 KO neurons compared to controls. These results are very similar to those
seen in constitutive MeCP2 KO neurons which show defects in mEPSC frequency
(Nelson et al., 2006) and evoked excitatory synaptic transmission but no alterations in
inhibitory synapse function.
In conclusion, our findings support a role for both HDACs 1 and 2 in the control
of excitatory synaptic transmission. It appears that HDAC2 may play a part in controlling
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both spontaneous and evoked synaptic activity, while HDAC1 is more specific in only
controlling evoked neurotransmission. The fact that these results occur in response to
acute knockdown of HDAC expression after neurodevelopment highlights the function of
these proteins in mature neurons. In addition, the loss of both HDACs closely mimicked
the defects seen in MeCP2 KO neurons indicating a shared mechanism for controlling
excitatory synapse function. More work needs to be done to explore this relationship and
to determine if the loss of these proteins results in the increased expression of similar
genes important for regulating synaptic transmission.
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Figure 4-1. Treatment with HDAC inhibitors results in increased acetylation of H4
and defects in both spontaneous and evoked synaptic transmission.
(A and B) Spontaneous miniature EPSCs recorded from neurons treated 24 hours with
inhibitors of HDAC activity. (A) Both TSA and VPA treatments caused a significant
decrease in the frequency of mEPSCs compared to controls (*, p<0.05). (B) No
alterations in mEPSC amplitudes were seen with these drugs. (C) TSA treatment caused a
significantly faster depression of evoked postsynaptic currents in response to 10 Hz field
stimulation (**, p<0.01). (D and E) Immunostaining for histone H4 acetylation. (D)
Images depicting an increase in fluorescence intensity after TSA and VPA treatments
compared to controls. (E) HDAC inhibitor treatments resulted in significant increases of
acetylated H4.
90
Figure 4-2. Quantification of HDAC1 and 2 mRNA and protein expression levels
one week after infection with Cre-recombinase lentivirus.
(A and B) HDAC1 and 2 mRNA measured by quantitative Real-Time PCR. (A) HDAC1
mRNA was significantly knocked down after lentiviral Cre infection in the floxed
HDAC1 neurons, while the levels of HDAC1 were unchanged in HDAC2 KO cultures
(***, p<0.001). (B) HDAC2 mRNA levels were significantly decreased in floxed
HDAC2, but not floxed HDAC1, neurons after lentiviral infection (**, p<0.01). (C and
D) Western blot analysis of HDAC1 and 2 protein expression. (C) Infection with Crerecombinase resulted in a significant decrease in HDAC1 protein in the floxed HDAC1
neurons (*, p<0.05). (D) Floxed HDAC2 neurons showed a significant knock down of
HDAC2 protein levels after infection (**, p<0.01).
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Figure 4-3. Loss of HDAC2, but
not HDAC1, results in decreased
frequency of spontaneous
mEPSCs.
(A-C) Miniature excitatory events
from floxed HDAC1 or 2 neurons
infected with lentivirus expressing
Cre-recombinase. (A)
Representative traces from infected
neurons recorded in the presence of
1 μM TTX and 50 μM picrotoxin.
(B) Bar graph revealing a
significant decrease in mEPSC
frequency from floxed HDAC2
neurons, but not floxed HDAC1
neurons, after infection with
lentiviral Cre recombinase. The
HDAC inhibitor, TSA, reduced
mEPSC frequencies in all neurons
compared to untreated controls (*,
p<0.05; **, p<0.01). (C)
Cumulative histogram showing no
significant changes in mEPSC
amplitudes. (D and E)
Immunostaining of cultured
neurons. (I) Dissociated neurons
from control and HDAC2 knockout
mice were labeled with primary
antibodies to PSD-95 (blue) and
Synapsin (red). (J) Bar graph
depicts the number of excitatory
synapses found in control and
HDAC2 neurons.
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Figure 4-4. Miniature inhibitory synaptic currents from conditional
HDAC1 and 2 KO neurons.
(A) Representative recordings of miniature inhibitory events from control,
HDAC1, and HDAC2 knockout neurons recorded in 1 μM TTX and 10 μM
NBQX. (G) Bar graph showing similar frequencies of spontaneous
inhibitory events in control and floxed HDAC1 and 2 neurons after Cre
infection. (H) Cumulative histogram of mIPSC amplitudes reveals no
differences between either HDAC1 or 2 neurons and controls.
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Figure 4-5. Increased
evoked EPSC amplitudes
and decreased paired pulse
ratios in HDAC1, HDAC2,
and MeCP2 KO neurons.
(A) Representative traces of
evoked excitatory currents
recorded in 50 μM PTX at 1
Hz and 20 Hz stimulation
frequencies. (B) First peak
amplitudes were significantly
increased in all three KO
neurons compared to controls
(*, p<0.05). (C) MeCP2 KO
neurons showed decreased
paired pulse ratios at 5, 10 and
20 Hz stimulation frequencies
compared to controls, while
both HDAC1 and HDAC2
neurons had reduced paired
pulse ratios at 20 Hz (*,
p<0.05; **, p<0.01).
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Figure 4-6. Evoked inhibitory postsynaptic currents from HDAC1, HDAC2, and
MeCP2 KO cultures.
(A) Representative traces of IPSCs recorded in 10 μM NBQX. (B) First peak
amplitudes of inhibitory currents were the same in all KO neurons compared with
controls. (C) Paired pulse ratios recorded at numerous stimulation frequencies were
unaffected by the loss of HDAC1, HDAC2, or MeCP2.
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CHAPTER 5
CONCLUSIONS AND FUTURE DIRECTIONS
An obvious role for transcriptional repression in the control of neuronal function
is quickly emerging. Upon the discovery of associations between a number of
neurological disorders and repressor proteins, like MeCP2 and DNMT3b, much research
has begun to focus on these different components of the transcriptional repression
machinery and their functions in the brain. One particular transcriptional repressor of
great recent interest is MeCP2. Mutations in this gene lead to a neurodevelopmental
disorder in humans called Rett syndrome (RTT), therefore much research has begun to
focus on MeCP2’s role in the brain. My thesis work was concerned with delineating
MeCP2’s function in the regulation of synaptic transmission. In addition, we looked at
two mechanisms important for directing and mediating the transcriptional repression
activity of MeCP2, DNA methylation and histone deacetylation.
In order to study the effects of these repressor proteins on synapse function, we
made dissociated hippocampal cultures from a number of different mouse strains and
recorded postsynaptic currents using whole-cell voltage clamp electrophysiology.
Overall, our findings agree significantly with what is known about the molecular
interaction between MeCP2, DNA methylation, and HDACs 1 and 2. MeCP2 knockout
neurons displayed deficits specifically in excitatory synaptic transmission, while showing
no alterations in inhibitory activity. Interestingly, there are a number of studies
suggesting an abnormal ratio of excitation/inhibition in the brains of autistic patients
(Purcell et al., 2001; Rubenstein and Merzenich, 2003; Serajee et al., 2003), and since
96
RTT is considered an autism-spectrum disorder it is not unreasonable to hypothesize that
something similar may be occurring in the brains of RTT patients. This specificity for
excitatory synaptic transmission was also seen after manipulation of DNA methylation as
well as after knockdown of HDAC1 and 2 in post-mitotic neurons. Alterations in
excitatory synapse function included both defects in spontaneous neurotransmission and
deficits in short-term synaptic plasticity. Interestingly, the loss of MeCP2, DNA
methylation, or HDAC2 caused defects in spontaneous synaptic currents, while the loss
of MeCP2 and both HDACs resulted in alterations in evoked neurotransmission. This
suggests that genes important for controlling spontaneous versus evoked synaptic activity
are differentially expressed depending on the components of the transcriptional repressor
complex. Evidence suggests a separation of presynaptic vesicles that is dependent on
whether they are released spontaneously or in response to activity (Sara et al., 2005).
Perhaps understanding the molecular mechanisms behind this distinction would help
determine possible genes regulated by the MeCP2 complex.
The identification of MeCP2 target genes has proved to be a challenging task.
Microarray analysis of RTT brains and MeCP2 KO mice revealed essentially nothing
about genes whose expression levels were increased after the loss of MeCP2-dependent
transcriptional repression (Colantuoni et al., 2001; Tudor et al., 2002). This has lead some
to believe that MeCP2 plays alternate functional roles in neurons, such as in the
regulation of RNA splicing (Young et al., 2005). The work presented in this thesis
demonstrates that MeCP2’s transcriptional repression activity is responsible for the
defects seen in synapse function. Future work will be directed towards identifying the
MeCP2 target genes responsible for these deficits.
97
For wide-scale analysis of MeCP2 target genes, a chromatin immunoprecipitation
(CHIP) assay will be utilized, followed by the identification of gene promoters using
microarray or sequence specific primers. A relatively new method by which to pull down
targets of MeCP2 is the use of the methyl-binding domain of the protein in CHIP
experiments (Ballestar et al., 2003). To expand the list of genes to include those also
affected by DNMT or HDAC inhibition, we can pull down using antibodies to
methylated cytosines or particular acetylated histones. Our hope is that by making use of
the CHIP technique, we can filter out many of the genes not regulated by MeCP2 and
hopefully resolve the question of MeCP2’s function as a transcriptional repressor.
We have also undertaken a more specified approach to identifying MeCP2 target
genes. Using viral-mediated knockdown of MeCP2, we were able to attribute some of the
synaptic deficits to the loss of MeCP2’s regulation of presynaptic function. Therefore, we
will attempt to identify presynaptic target genes using quantitative Real-Time PCR with
mRNA isolated from MeCP2-deficient neurons. We have comprised a list of possible
presynaptic genes based on their known localization and function at the synaptic terminal
(Figure 5-1). Real-Time PCR has been carried out on a number of these genes. In order to
narrow down the list for preliminary studies, we searched for DNA sequences thought to
be important for the specificity of MeCP2 to certain target genes. Previous work has
implicated an association of MeCP2 with the corepressor coREST, a protein that
recognizes RE1 sequences in the promoters of its target genes (Lunyak et al., 2002). We
narrowed down the list by looking for RE1 sequences in the promoters of these genes.
We then looked for A/T rich sequences located near CpG islands, since one study
98
suggests a need for A/T rich DNA in order to recruit MeCP2 to methylated promoters
(Klose et al., 2005).
Figure 5-1. List of possible
presynaptic gene targets of
MeCP2.
Genes with RE1 sequences
are highlighted as well as
those containing A/T rich
sequences near CpG islands.
The purple * indicates genes
whose expression levels
were previously reported to
be unaffected by the loss of
MeCP2 (Asaka et al., 2006).
Our Real-Time PCR results showed the significant upregulation of a couple of
possible MeCP2 targets, Complexin 2 (Cplx2) and Synaptoporin (Synpor) (Figure 5-2).
Not surprisingly, there were no dramatic alterations in the expression levels of any of the
genes, which may help explain the inability to use microarray analysis to identify these
targets. Cplx2 is an interesting candidate since Complexins have been shown to inhibit
fast neurotransmitter release in response to synaptic activity and have no effect on
spontaneous synaptic transmission (Reim et al., 2001; Tang et al., 2006). The fact that
Cplx2 differentially regulates evoked versus spontaneous activity makes it an intriguing
candidate. In addition, Cplx2 is specific for excitatory synapses, while Cplx1 is found at
inhibitory synapses, and the fact that we did not see changes in Cplx1 expression further
supports the specificity of MeCP2 for excitatory synapse function. Synpor, also called
Synaptophysin 2, is a little more difficult to understand because its function in the brain is
unknown, but we will continue to pursue it as a possible MeCP2 target gene.
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Figure 5-2. Quantitative Real-Time PCR results from MeCP2 KO cultures.
Only the mRNA expression levels of Cplx2 and Synpor were significantly increased
after the loss of MeCP2. Nrxns 1 and 3 appear to decrease in the MeCP2 KO.
We also looked at the expression of Neuroligins (Nlgn) and Neurexins (Nrxn) in
our MeCP2 KO cultures. Although Nlgns are postsynaptic scaffolding proteins, Nlgn3
mutations have been found in some cases of autism (Jamain et al., 2003). Nlgn1 is the
isoform specific for excitatory synapses and has been shown to modulate presynaptic
neurotransmitter release in a retrograde manner (Futai et al., 2007). While both of these
genes are promising candidates with respect to their functions and effects on the brain,
we found no changes in Nlgn1 or 3 mRNA expression in MeCP2 KO neurons (Figure 52). Nrxns are the presynaptic binding partners of Nlgns. Any regulation of synaptic
transmission by Nrxns would presumably be similar to that for Nlgns (Craig and Kang,
2007). Interestingly, we saw a decreasing trend in Nrxn1 and Nrxn3 expression (Figure
5-2), a change that may become significant with repeated experiments. Though this
argues against Nrxns being direct targets of MeCP2 repression, they may be indirectly
regulated in response to the loss of MeCP2 and may also play a role in the
neurophysiological defects seen in the KO.
100
Another exciting discovery from this work is the activity-dependent
demethylation of genomic DNA in mature, post-mitotic neurons. The practice of active
DNA demethylation in nondividing cells is quite controversial, mainly due to the lack of
convincing identification of a demethylating enzyme in mammals. Our data demonstrate
that DNA demethylation indeed occurs in post-mitotic neurons and that this process can
be used to regulate spontaneous synaptic transmission. Furthermore, our findings
demonstrate that these DNA methylation changes are regulated by synaptic activity. This
finding is intriguing because it adds to the many ways in which gene transcription can be
altered in response to synaptic input. Activity-dependent gene regulation is critical for the
brain to convert synaptic inputs into long-term alterations in synapse structure and
function. A recent study has suggested that learning can actually drive both DNA
demethylation and methylation, presumably through some activity-dependent
postsynaptic signaling cascade (Miller and Sweatt, 2007). Our work directly implicates
synaptic activity-driven changes in DNA methylation, possibly as a result of calcium
influx through postsynaptic NMDA receptors.
Finally, two of our most significant findings relate back to MeCP2 and RTT. Our
viral-mediated deletions of MeCP2, as well as HDAC1 and 2, after early maturation
demonstrate the importance of these proteins in the regulation of synapse function in
mature neurons. These proteins were present throughout neurodevelopment but the loss
of them after synapse formation resulted in synaptic deficits similar to those seen in the
constitutive MeCP2 KO. These data imply that MeCP2-, along with HDAC1- and 2-,
dependent transcriptional repression regulates excitatory synaptic transmission after
neurodevelopment. The data also suggest that RTT is not strictly a neurodevelopmental
101
disorder and indicate the possibility that late therapeutic intervention might reverse the
neurological symptoms seen in RTT patients. Along this idea, a recent study garnered a
great deal of attention by demonstrating a significant reversal of the behavioral
phenotypes and synaptic plasticity deficits in MeCP2 KO mice by returning MeCP2
expression to control levels (Guy et al., 2007). The authors generated a conditional
knockin mouse that did not express MeCP2 until the introduction of Cre-recombinase
caused the excision of a stop codon within the gene that then allowed for MeCP2
expression. Unfortunately, this study was problematical due to the method by which they
manipulated MeCP2 expression, but it supports the idea that RTT is not strictly a
neurodevelopmental disorder. In humans, using gene therapy to introduce MeCP2 to RTT
patients, who are mosaics of normal and mutant MeCP2-containing cells, would most
likely result in many of their cells expressing too much MeCP2. Both humans and mice
with MeCP2 overexpression display many of the same neurological deficits as RTT
patients (Collins et al., 2004; Luikenhuis et al., 2004; Van Esch et al., 2005), signifying
the importance for proper control of MeCP2 expression levels.
The discovery that RTT symptoms are not necessarily a result of
neurodevelopmental defects puts forward the idea that treatments intended to reverse
these phenotypic deficits may be successful later in life. While gene therapy to increase
MeCP2 expression levels may not be the answer, it may be possible to modulate the
expression levels of putative target genes of MeCP2 and ameliorate some of the
symptoms. One study attempted to do this by crossing mice overexpressing BDNF with
MeCP2 KO mice. The increased expression in BDNF was able to rescue some of the
deficits in the KO, however they provide no data on the effects of BDNF overexpression
102
in normal wildtype mice (Chang et al., 2006). Our results showing a reversal of the
mEPSC deficit seen in MeCP2 null mice using treatment with SAM could be another
therapeutic avenue. The next step would be to treat MeCP2 KO mice with SAM in vivo
and see if there is a rescue of any of their behavioral deficits. Interestingly, there has
already been a study in human RTT patients who received folinic acid treatments. Folinic
acid is the molecular precursor to methionine, among other reaction products. Folinic
acid is used to convert homocysteine to methionine, which is then activated by
methionine adenosyltransferase to make S-adenosylmethionine (SAM), the major methyl
donor for cellular methyltransferase reactions, including those involving DNMTs. RTT
patients treated with folinic acid showed improvements in social behavior and presented
less stereotypical “hand-washing” movements and fewer seizures (Ormazabal et al.,
2005), signifying folinic acid as a viable treatment for patients with Rett syndrome.
In conclusion, these studies implicate a role for the MeCP2 transcriptional
repressor complex in controlling excitatory synaptic transmission. Future work will be
directed towards the identification of genes whose expression is regulated by any or all of
the components of this complex. In addition, there will be continued investigation into
the association of MeCP2, DNA methylation, and HDACs 1 and 2 and their combined
effects on synapse function. Could the deficits caused by the loss of one protein, perhaps
MeCP2, be rescued by manipulating the amounts of another component like DNA
methylation or HDAC activity? Our findings support this idea and we look forward to
future work aimed at answering some of these questions.
103
MATERIALS AND METHODS
Cell Culture
Dissociated hippocamal cultures were prepared from the brains of MeCP2 null knockout
mice, (Jackson Laboratories) floxed MeCP2 mice, floxed HDAC1 mice, floxed HDAC2
mice, or C57BL/6 mice, according to previously published protocols (Kavalali et al.,
1999). Briefly, whole hippocampi were dissected from the floxed mice or C57BL/6 mice
on postnatal days 0-3, or MeCP2 knockout mice on postnatal day 0. Tissue was
trypsinized for 10 min at 37ºC, mechanically dissociated using siliconized glass pipettes,
and then plated onto matrigel-coated coverslips. For DNA methylation, RNA and protein
level measurements, neurons were plated directly onto matrigel-coated 6-well plates. All
experiments were done on cultures 11-22 days in vitro (DIV).
Drug treatments and cell viability
24 or 48-hour treatments of hippocampal cultures were done with the following drugs:
dimethyl sulfoxide (DMSO) (1:1000), Trichostatin A (TSA) (1 μM), 5-azacytidine
(5azaC) (2.5 μM), Zebularine (Zeb) (50 μM), S-Adenosyl-L-methionine (SAM) (100
ug/ml), Actinomycin D (Act D) (2.5 μM), Tetrodotoxin (TTX) (1 μM), Picrotoxin (PTX)
(50 μM), Vaproic Acid (VPA) (0.8mM), or 2-amino-5-phosphonopentanoic acid (AP5)
(50 μM). SAM (50 ug/ml) treatment was also done for 48 hours where indicated. After
drug treatments, cell viability was checked using Trypan Blue exclusion (Sigma). For the
first study, the percentages of dead cells were not significantly different between control
cultures (6.8±1.8% S.E.M., DMSO treated) and cultures treated with drugs (6.8±1.1%
TSA and 3.5±1.8% Act-D). For the second study, the percentages of dead cells were not
104
significantly different between control cultures (7.4±1.5% S.E.M., DMSO treated) and
cultures treated with DNMT inhibitors (8.1±2.4% 5azaC and 7.4±2.1% Zeb).
Immunocytochemistry
For the first study, dissociated hippocampal neurons were fixed for 30 min with 4%
paraformaldehyde, rinsed twice with 1X PBS/Glycine, then blocked in 2% goat serum for
1 hour. The cells were then incubated with primary antibodies, anti-MAP2 monoclonal
(1:200, Chemicon), and anti-synapsin polyclonal (1:1000, Synaptic Systems) overnight at
4oC. The next day the cells were washed then incubated with fluorescent secondary
antibodies, goat-anti-rabbit (1:200, Molecular Probes), and goat-anti-mouse (1:200,
Molecular Probes). For the second and third studies, dissociated hippocampal neurons
were fixed for 2 min at room temperature in PBS with 2% formaldehyde and 2% sucrose
followed by treatment with cold methanol for 10 min at -20 oC. Then, the neurons were
blocked in 2% goat serum for 1 hr at room temperature. The cells were then incubated
with primary antibodies, anti-PSD-95 monoclonal (1:200, Affinity Bioreagents), antiAcH4 polyclonal (1:500, Upstate), or anti-Synapsin polyclonal (1:1000, Synaptic
Systems) added to 0.02% gelatin and 0.5% Triton X-100 in PBS overnight at 4oC. The
next day, neurons were washed with PBS and then incubated with fluorescent secondary
antibodies, goat-anti-rabbit (1:200, Molecular Probes), and goat-anti-mouse (1:200,
Molecular Probes). Coverslips were mounted with Vectashield (Vector Laboratories)
and neurons were visualized on a Zeiss Confocal microscope.
105
Electrophysiology
Synaptic activity was recorded from hippocampal pyramidal cells using a whole-cell
voltage clamp technique. Data were acquired using an Axopatch 200B amplifier and
Clampex 9.0 software (Axon Instruments). Recordings were filtered at 2 kHz and
sampled at 200 μsec. A modified Tyrode solution containing (in mM): 150 NaCl, 4 KCl,
2 MgCl2, 2 CaCl2, 10 Glucose, 10 HEPES, pH 7.4) was used as external bath solution for
all experiments unless otherwise noted. A hypertonic Tyrode solution was made by
adding 500 mM sucrose. The pipette internal solution contained (in mM): 115 CsMeSO3, 10 CsCl, 5 NaCl, 10 HEPES, 0.6 EGTA, 20 TEA-Cl, 4 Mg-ATP, 0.3 Na3GTP,
pH 7.35 (300 mOsm). The pipette solution for field stimulation also contained 10 mM
QX-314. Field stimulation was applied through parallel platinum electrodes immersed in
the perfusion chamber delivering 20 mA pulses.
Fluorescence Imaging
For high potassium stimulation, synaptic boutons were loaded with FM1-43 during a 90 s
incubation in Tyrode solution containing 47 mM K+. After washing with a dye-free
Tyrode solution for 10 min, synaptic terminals were destained using a 90 mM K+ Tyrode
solution for 90 s followed by 3 applications of 60 s (each separated by 60 s intervals). For
spontaneous recycling pool experiments, boutons were loaded with FM2-10 during a 15
min incubation in Tyrode solution containing tetrodotoxin (TTX), washed for 10 min in
dye-free Tyrode, then destained with Tyrode solution containing TTX for 20 min. This
was followed by 3 applications of 90 mM K+ Tyrode solution for 60 s (each separated by
60 s) to release all dye from synapses. All staining and washing solutions contained 10
106
μM CNQX and 50 μM AP-5 to prevent recurrent activity. Isolated boutons were selected
during the wash and fluorescence changes were measured during destaining. Images
were obtained by a cooled, intensified digital CCD camera (Roper Scientific) during
illumination (1 Hz and 40 ms) at 480 nm via an optical switch (Sutter Instruments).
Images were acquired and analyzed using Axon Imaging software (Axon Instruments).
Lentivirus Production
HEK 293 cells were transfected using the Fugene 6 transfection system (Roche
Molecular Biochemicals) with the expression plasmid, pFUGW or pFUGW-Cre and two
helper plasmids, delta 8.9 and vesicular stomatitis virus G protein, at 3 µg of each DNA
per 75 cm2 flask (Dittgen et al., 2004). After 48 hours, lentivirus containing culture
medium was harvested, filtered at a 0.45-µm pore size, and immediately used for
infection. Hippocampal cultures were infected at 7 DIV by adding 300 μl of viral
suspension to each well and recordings were done 13-17 DIV. Titer was determined by
counting the number of infected neurons per coverslip (high: >80%; low: <20%).
Methylation-Sensitive Amplified Fragment Length Polymorphism
Genomic DNA was extracted from hippocampal cultures (DNeasy tissue kit; Qiagen,
Valencia, CA) and then 1 μg DNA was processed for fluorescent NotI-MseI MS-AFLP
according to previous protocols (Yamamoto et al., 2001). DNA was digested for 2 hr at
37ºC with 10 units each of NotI and MseI. NotI adaptor sequences (5’CTCGTAGACTGCGTACC-3’ and 5’-GGCCGGTACGCAGTCTAC-3’) and MseI
adaptor sequences (5’-GACGATGAGTCCTGAG-3’ and 5’-TACTCAGGACTCAT-3’)
107
were duplexed together by adding RNase-Free Duplex Buffer (IDT) and incubating at 94º
C for 2 min. Then, 1.5 μL each of 50 μM NotI and 50 μM MseI adaptors were ligated to
the digest product by adding 2 μL T4 DNA ligase, 4 μL 10x buffer, 11 μL H2O and
incubating overnight at 16ºC. Two PCR reactions were then run. For PCR1, 5 μL 1:2.5
dilution of ligation product was added to 0.5 μL Taq Polymerase (Invitrogen), 5 μL 10x
PCR buffer, 3 μL MgCl2 (25 mM), 4 μL dNTPs (2.5 mM), 1 μL formamide, 30 μL H2O
and 0.75 μL of both NotI and MseI PCR1 primers (20 μM). PCR1 primer sequences:
Not1 5’-GACTGCGTACCGGCCGC-3’ and Mse1 5’-GATGAGTCCTGAGTAA-3’.
PCR1 program: 72ºC for 1 min, 94ºC for 2 min, 35 cycles of 94ºC for 30s, 52ºC for 1
min, and 72ºC for 2 min, then 72ºC for 7 min and 10ºC to end. For PCR2, 5 μL 1:15
dilution of PCR1 product was added to 0.25 μL Taq Polymerase (Invitrogen), 2.5 μL 10x
PCR buffer, 1.5 μL MgCl2 (25 mM), 3 μL dNTPs (2.5 mM), 0.5 μL formamide, 11 μL
H2O, 0.25 μL fluorescently labeled NotI PCR2 primer (20 μM), and 1 μL MseI PCR2
primer (20 μM). PCR2 primer sequences were the same for PCR1+N (A/C/T/G) added to
the 3’ ends of both Not1 and MseI primers and 56-FAM custom synthesized (IDT) to the
5’ ends of each Not1 primer, giving a total of 4 Mse1+N primers and 4 labeled Not1+N
primers. A total of 16 PCR2 reactions were run with different combinations of Mse1+N
and Not1+N primers. PCR2 program: 72ºC for 1 min, 94ºC for 2 min, 12 cycles of 94ºC
for 30s, touchdown 64ºC to 52ºC for 1 min, and 72ºC for 2 min, the 23 cycles of 94ºC for
30s, 52ºC for 1 min, and 72ºC for 2 min followed by 72ºC for 7 min and 10ºC to end.
PCR products were purified by centrifugation through Sephadex G-75 beads (Sigma)
108
then 10 μL were loaded onto an ABI 377 DNA sequencer and electrophoresed according
to manufacturer’s protocols.
Measurement of Unmethylated DNA
Genomic DNA was extracted from hippocampal cultures (DNeasy tissue kit; Qiagen,
Valencia, CA) and then bisulfite modification of 0.4 μg DNA was performed (CpGenome
DNA modification kit; Chemicon, Temecula, CA). Quantitative Real-Time PCR was
used to determine the amount of unmethylated CpG island present in the BDNF promoter
I according to previously published work (Levenson et al., 2006). Briefly, 2 μL DNA was
added to 10 μL iQ SYBR Green Supermix (Bio-rad, Hercules, CA), 7 μL DEPC H2O and
1 μL of each primer. The following primers were used at 18 uM concentration: forward
(5’-GGGTAGTGATTTTGGGGAGGAAGTAT-3’) and reverse (5’CAACCTCTATACACAACTAAATCCACC-3’). GAPDH primers were used as
controls: forward (5’-AGGTCGGTGTGAACGGATTTG-3’) and reverse (5’TGTAGACCATGTAGTTGAGGTCA-3’). Each sample was run in triplicate. Reactions
were run on an Mx3000P real-time PCR machine (Stratagene, La Jolla, CA) with the
following cycling program: 95°C for 3 min, 40 cycles of 95°C for 15 s, 60°C for 1 min,
and 74°C for 15 s. Detection of fluorescent products was at the end of the last step. For
each sample, a ΔCt value was determined (Ct BDNF – Ct GAPDH) followed by a ΔΔCt
value relative to DMSO controls (ΔCt Experimental treatment - ΔCt Control treatment).
Fold changes were determined by taking 2 to the power of ΔΔCt values. PCR products
were run out on an agarose gel and visualized using ethidium bromide.
109
RNA Isolation and Reverse Transcription
Hippocampal cultures were washed once with PBS. Neurons were scraped from 6-well
plate in 500 μL RNA STAT-60 reagent (Tel-Test, Inc., Friendswood, TX) and transferred
to 1.5 mL eppendorf tubes. Tubes were incubated on ice for 5 min, then 100 μL
chloroform was added, mixed thoroughly and incubated on ice for 2 min. Tubes were
spun for 15 min at 12,000xg at 4°C. Upper, aqueous layer was transferred to new tubes,
then 250 μL isopropanol and 4 μL linear acrylamide (Ambion, Austin, TX) was added,
mixed and incubated at -80°C for 1 hour. Tubes were spun for 15 min at 12,000xg at 4°C.
Pellets were washed with 1 mL 70% ethanol and then resuspended in 20 μL DEPCtreated H20. 0.8 μg RNA was brought up to a total volume of 17 μL with DEPC H20,
treated with 1 μL TURBO DNase (Ambion, Austin, TX) in 2 μL DNase buffer, mixed
and then incubated at 37°C for 25 min. 5 μL DNase Inactivation Reagent was mixed in
for 2 min at 23°C, then pelleted by spinning at 10,000xg for 1 min and supernatant was
transferred to new tubes. 2 μL each of Random Hexamers (50 ng/uL), dNTPs (10 μM),
and DEPC H20 was mixed in with the samples and then incubated at 65°C for 5 min.
Tubes were put on ice for 1 min, then 8 μL 5x 1st Strand Buffer (Invitrogen) and 2 μL
each of DTT (0.1M) (Invitrogen), RNase OUT (Invitrogen), and Superscript III Reverse
Transcriptase (Invitrogen) was mixed in and incubated for 5 min at 25°C, 60 min at 50°C,
then 15 min at 70°C.
Quantitative RT-PCR
1 μL cDNA was added to 10.5 μL iQ SYBR Green Supermix (Bio-rad, Hercules, CA),
7.5 μL H2O and 1 μL of each primer. The following primers were used at 10 μM
110
concentration: HDAC1 forward (5’-TCTACCGCCCTCACAAAGC-3’) and reverse (5’ACAGAACTCAAACAAGCCATCA-3’), HDAC2 forward (5’-GCGTACAGTCAAGGAGGCGG-3’) and reverse (5’-GCTTCATGGGATGACCCTGGC-3’), Syt7
forward (5’-ACGGCCACTACCCTTGAGT-3’) and reverse (5’-AAGGATTTCATGTCCAAGCCTC-3’), Cplx1 forward (5’-AGTTCGTGATGAAACAAGCCC-3’) and reverse
(5’-TCTTCCTCCTTCTTAGCAGCA-3’), Cplx2 forward (5’-AAGAGCGCAAGGCGAAAC-3’) and reverse (5’-TGGCAGATATTTGAGCACTGT-3’), Syp1 forward (5’AGTGCCCTCAACATCGAAGTC-3’) and reverse (5’-CGAGGAGGAGTAGTCACCAAC-3’), Synpor forward (5’-GGCACCTTTCGGGCATTGA-3’) and reverse (5’CCTCCTGAATAGCCACCACA-3’), Rab3a forward (5’-ACCACAGAATATTACCGAGG-3’) and reverse (5’-GCATTGTCCCACGAGTAAGTTTT-3’), Nlgn1 forward (5’CTTGGGGTACTTGAGAAAGAGAC-3’) and reverse (5’-CTTGTTTGGGTATAAAGCCTCCA-3’), Nlgn3 forward (5’-TCGCCACTTATATCCAGGAGC-3’) and reverse (5’ATCCCCGTCATTATCCGCTAA-3’), Nrxn1 forward (5’-AATCTGCGTCAGGTGACAATAC-3’) and reverse (5’-GCCACCACACCGTGAATCTT-3’), Nrxn3 forward (5’AGACCCCAGAGGCTTACATCA-3’) and reverse (5’-CGTGAGTGAAGAGAATCAGGC-3’), and BDNF forward (5'-CCTGCATCTGTTGGGGAGAC-3') and reverse
(5'-GCCTTGTCCGTGGACGTTTA-3'). GAPDH primers were used as controls: forward
(5’-AGGTCGGTGTGAACGGATTTG-3’) and reverse (5’- TGTAGACCATGTAGTTGAGGTCA-3’). Each sample was run in triplicate. Reactions were run on an Mx3000P
real-time PCR machine (Stratagene, La Jolla, CA) with the following cycling program:
95°C for 10 min, 40 cycles of 95°C for 20 s, 59°C for 30 s, and 72°C for 20 s. Detection
of fluorescent products was at the end of the second step.
111
Protein Extraction and Immunoblotting
Hippocampal cultures were washed once with PBS. Neurons were scraped from 6-well
plate in 500 μL PBS and transferred to 1.5 mL eppendorf tubes. Tubes were spun for 5
min at 1,000xg and pellets were washed in cold PBS. Pellets were resuspended in 30 μL
Lysis buffer (25 mM HEPES pH 7.9, 150 mM NaCl, 2 mM EDTA, 1mM DTT, 0.1%
NP-40, 1 mM PMSF, 1 μM aprotinin, 1 μM leupeptin, 20 mM NaF, 1X protease
inhibitor) and tubes were put on ice for 15 min. Incubated tubes for 1 hour at -20°C.
Tissue was homogenized using a 25 gauge needle and syringe. Tubes were spun for 5
min at 15,000xg at 4°C, then supernatant was kept for protein. 10 μg protein was mixed
with 5x loading dye (0.4 M Tris pH 6.8, 10% SDS, 60% glycerol, 2g bromophenol blue)
and loaded onto 10% 19:1 acrylamide stacking gels (1.5 M Tris pH 8.8, 10% SDS, 10%
ammonium persulfate, 0.04% TEMED). Gels were run at 100V for 1.5 hours, then
protein was transferred to nitrocellulose membranes at 4°C at 100V for 1.5 hours.
Membranes were blocked in 3% milk for 1 hour, then incubated in primary antibodies
made in 3% milk overnight at 4°C: rabbit anti-HDAC1 (1:2000, Sigma) and rabbit antiHDAC2 (1:2000, Sigma). Membranes were washed in TBS-10% Tween for 1 hour, then
incubated in secondary antibody, anti-rabbit (1:2000) in 3% milk, for 1.5 hours.
Membranes were washed in TBS-10% Tween for 1 hour, then incubated in ECL-Plus
reagent (Amersham Biosciences) for 5 min and exposed to film.
112
Statistical Analysis
All error bars represent the standard error of the mean (S.E.M) and all data was tested for
statistical significance by means of a two-tailed Student’s t test.
113
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VITAE
Erika Dawn Nelson was born in Kansas City, MO on July 28, 1980 to the parents of Ann
Cunningham and Jon Nelson. In 1998, she graduated in the top ten percent of her class at
Olathe South High School in Olathe, KS. Following graduation, she attended Southern
Methodist University in Dallas, TX for two years and then transferred to the University
of Kansas in Lawrence, KS in the fall of 2000. In May of 2002, she graduated from the
University of Kansas with honors and distinction and was awarded a Bachelor of Science
in Biology with an emphasis in Genetics. In the fall of 2002, she came back to Dallas, TX
where she entered graduate school at the University of Texas Southwestern Medical
Center in Dallas, TX. She did her graduate research in the lab of Dr. Lisa Monteggia,
funded by the University Cell and Molecular Biology training grant. She was awarded
her Doctorate of Philosophy in Neuroscience in June, 2007.
Permanent address: 2502 Live Oak St. #308
Dallas, TX 75204