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Exploring the role of cooperative and competitive interactions in oligodendrocyte differentiation and myelination

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Exploring the role of cooperative and competitive interactions in oligodendrocyte differentiation and myelination

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Content  
EXPLORING THE ROLE OF COOPERATIVE AND COMPETITIVE
INTERACTIONS IN OLIGODENDROCYTE DIFFERENTIATION AND
MYELINATION

by

Sheila Sara Rosenberg

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(NEUROSCIENCE)

May 2010

Copyright 2010 Sheila Sara Rosenberg
 
 
ii 
DEDICATION

The teacher who is indeed wise does not bid you to enter the house of his wisdom but
rather leads you to the threshold of your mind.
~Kahlil Gibran

To everyone who has acted as a true teacher in my life, thank you.

 
 
iii 
ACKNOWLEDGEMENTS
There are many experiences and interactions that greatly impact our lives and influence
who we become. I know that the words I put on this page will fall dreadfully short both
of expressing my true feelings about the profound effect that people have had on my life
and of acknowledging everyone who has contributed to who I am and where I am today. I
hope that my actions and my relationships now and in the future will be the best indicator
of the way I truly feel about the indelible mark that so many of you have made on my life
and who I have become.
First I want to acknowledge my family and the way you have shaped my life from
the very beginning. Thank you to my parents, Linda and George, for the countless
sacrifices you have made to give me so many wonderful opportunities to learn and to
grow and to follow my passions for the things that I love. Thank you for all the things
you have taught me, and the values you instilled. Thank you for raising me to believe in
my ability to pursue my dreams and for giving me the encouragement and the freedom to
discover what I wanted to do with my life. Thank you for your support and your
unending faith in me. And above all, thank you for your love and for all you have done
that has made me believe that your deepest desire is for me to be happy.
Thank you to my sister Melissa for your incredible loyalty, genuine support and
endless love and concern. Thank you for being there for me in so many different ways-
especially when I needed you the most. Thank you for making me think and most of all,
thank you for making me laugh like no one else can. I am so proud and so grateful to
have you as my sister and my friend.
 
 
iv 
Thank you to my godmother Laura, for your thoughtfulness and your insight and
for always making time. Thank you for your love, your encouragement and advice, and
thank you to the entire Kikawa family for truly providing me a home away from home.
Thank you to all of my friends for supporting me and showing me how much you
care. Thank you for listening to me and for believing in me and reminding me to believe
in myself.
Thank you to everyone at USC who has contributed positively to my graduate
experience and has challenged me to think in different ways. Thanks especially to my
committee members and to all of the professors who have taken the time to talk to me
about science and a career in academics and shared with me your own thoughts,
experiences and advice.
Thank you to all of the past and present members of the Chan lab for your help
along the way and for all that I have gained as a result of my opportunity to work with
you. Thank you also for the scientific contributions that each of you have made to the
work that I will discuss here. In particular, thank you to Ben Ng for your sincere
kindness and all of the time that you spent listening to me when I first joined the lab.
And thank you to Christin Chong for the things that you have shown me and for
everything you have done to make our project the best that it can be. I have learned so
much as a result of our close collaboration for the past year and a half.
Thank you to my advisor, Jonah Chan, for all of the opportunities that you have
given me during the past five years. I have learned a great deal about both myself and
other people as a result of the time spent in your lab. Thank you for your extreme
 
 
v 
generosity, especially for all of the conferences that you made it possible for me to
attend. These are experiences that I truly love and will always be grateful for. Thank you
for teaching me the importance of asking alternative conclusions and for constantly
stressing the value of asking good questions. And thank you for the advice you gave me
long ago to strive to be myself.
Lastly, I want to offer sincere thanks for the blessing of discovering a life that I
love and my deepest gratitude for all of the little things that gave these years added
meaning for me-like the sunsets, the mountains and the Magnolia trees.

 
 
vi 
TABLE OF CONTENTS

Dedication ii
Acknowledgements iii
List of Tables vii
List of Figures viii
Abstract ix
Chapter 1: In Search of Wisdom and Enlightenment 1
Chapter 2: The Geometric and Spatial Constraints of the Microenvironment 17
Induce Oligodendrocyte Differentiation

Introduction 18
Methods 20
Results 25
Discussion 43
Chapter 3: Nogo-A establishes the spatial segregation and extent of 47
myelination in the developing CNS

Introduction 48
Methods 53
Results 57
Discussion 73

Chapter 4: Looking Ahead 76
References 91

 
 
vii 
LIST OF TABLES

Table 1. Effects of membrane-coated and Fc-coated beads 69
on oligodendrocyte differentiation and myelination.

Table 2. Nogo deletion does not significantly affect OPC and 72
oligodendrocyte numbers.

 
 
viii 
LIST OF FIGURES

Figure 2.1: Temporal regulation of oligodendrocyte differentiation 26
Figure 2.2: The induction of differentiation at a critical density of 31
OPCs does not require dynamic axonal signaling

Figure 2.3: The inhibition of proliferation is not sufficient to 34
induce differentiation

Figure 2.4: An increase in available mitogens is not sufficient 35
to delay differentiation
Figure 2.5: Spatial constraints along an axon are sufficient to 37
induce differentiation

Figure 2.6: Secreted factors are not responsible for the 40
induction of oligodendrocyte differentiation by Schwann cells

Figure 2.7: Dynamic axonal signaling is not required for the induction 41
of differentiation by spatial and geometric constraints.
Figure 2.8: The induction of differentiation is dependent on the 42
geometry of the microenvironment
Figure 3.1: Oligodendrocytes exhibit striking diversity in the 59
number and length of myelin internodes in vivo
Figure 3.2: Variations in myelinogenic potential exist both 61
within and between brain regions

Figure 3.3: Membrane-bound inhibitory cues expressed by 63
oligodendroglial decrease the number of myelin
internodes formed per oligodendrocyte

Figure 3.4: Nogo-A is necessary and sufficient for the regulation 68
of myelinogenic potential
Figure 3.5: Increased myelination in Nogo knock-out mice 71

 
 
ix 
ABSTRACT

As humans, a swift and highly accurate means of neuronal communication is required for
both our basic survival, and our unique capacity for traits such as creativity and critical
thinking. Communication between the nervous system and all other systems, as well as
within the nervous system itself, depends on the transmission of neuronal action
potentials, electrical signals required for the generation of functional outputs. The
efficient transmission of the neuronal action potential is greatly enhanced by the
insulating properties of the myelin sheath. Destruction of the myelin membrane, as a
result of nerve injury or disease, significantly impairs the ability of the nervous system to
communicate and can lead to a host of debilitating symptoms, as well as an ultimate loss
of function. The development of novel strategies to promote remyelination is essential to
limit the extent of sensory, motor and cognitive deficits that occur following
demyelination. It is our belief that understanding how myelination occurs during
development could provide insight into efforts to promote remyelination.
In the central nervous system (CNS), myelin is formed by glial cells known as
oligodendrocytes. During development, oligodendrocyte precursor cell (OPCs)
proliferate and migrate throughout the CNS. Upon reaching a final destination, an OPC
will either differentiate into a myelinating oligodendrocyte or remain as a precursor cell
into adulthood. Oligodendrocytes that differentiate must then coordinate the appropriate
non-overlapping placement of myelin internodes along axonal tracts. Each differentiated
oligodendrocyte is capable of forming multiple myelin internodes. Akin to other aspects
 
 
x 
of nervous system development, proper myelination requires a precise match between the
number of myelinating cells and myelin internodes, and the number of axonal segments
requiring myelination. Understanding how the extent of myelination is precisely
coordinated between axons and oligodendrocytes requires investigation of the
mechanisms governing both oligodendrocyte differentiation and myelin internode
formation. We find that both cooperative and competitive interactions between
oligodendroglial cells play an instrumental role in regulating the fate and function of
these myelinating cells.

 
 
1 
CHAPTER ONE
In Search of Wisdom and Enlightenment

The philosopher Lao-Tzu wrote, “He who knows others is wise; He who knows himself
is enlightened”. One could argue that the acquisition of both wisdom and enlightenment
is intimately tied to a better understanding of nervous system function. The nervous
system is responsible for receiving and integrating a massive amount of sensory stimuli,
organizing and interpreting these external cues, and then consolidating information and
generating a response. All of these tasks are dependent on the formation of neural circuits
that facilitate electrochemical communication between neurons. While it is the neuronal
cell that is responsible for the actual conduction of signaling currents, the rate at which
the signal travels is greatly enhanced by the insulating properties of the glial-derived
myelin sheath. The functional role of myelin is the same in both the central nervous
system (CNS) and the peripheral nervous system (PNS), however, the composition of the
myelin membrane, the signals regulating myelination and the timing of myelination differ
somewhat between the two systems (Schwab & Schnell 1989). In the PNS, myelination
begins immediately after birth, while the onset of myelination in the CNS varies,
beginning anywhere from two days until approximately two weeks postnatal, or even
later depending on the specific brain region (Schwab & Schnell 1989). In the PNS,
myelin is formed by Schwann cells. Each Schwann cell ensheaths and myelinates a single
axon (Chan 2007). In the CNS, myelin is formed by terminally differentiated
oligodendrocytes. Each myelinating oligodendrocyte has the capacity to extend multiple
 
 
2 
processes that contact nearby axonal segments. Upon stabilization of axonal contact, each
process will then wrap a membranous sheet concentrically around the axon, culminating
in the formation of a myelin internode. The presence of myelin internodes facilitates a
rapid and efficient manner of signal propagation known as saltatory conduction
(Rosenberg et al 2006). When myelin internodes are lost due to disease or nerve damage,
the capacity for saltatory conduction is greatly inhibited or completely lost. This process
of demyelination results in severe deficits including sensory, motor and cognitive
impairments (Franklin 2002). It is therefore absolutely imperative to understand how to
promote remyelination in the damaged or diseased nervous system.
It is our belief that efforts to promote remyelination could be greatly enhanced by
establishing a better understanding of the manner in which myelin formation occurs
during development. We hypothesize that the cues responsible for regulating
developmental myelination could also play a role in the remyelination process. Indeed,
recent studies have shown that transcription factors involved in developmental
myelination are upregulated in demyelinated lesions in human MS patients (Fancy et al
2009; Rosenberg & Chan 2009). It is our hope that studying the developmental process of
myelination will help elucidate the factors that are necessary and sufficient to promote
remyelination and help alleviate the devastating effects of demyelinating injuries and
diseases.

 
 
3 
Obstacles Preventing Remyelination
The primary requirement for maintaining and regaining function after demyelination in
the CNS is the presence of new myelinating cells to replace lost oligodendrocytes.
Transplantation of exogenous cells has long been considered a possible approach to
enhance remyelination following CNS injury or disease (Franklin & Kotter 2008).
However, recent studies suggest that the use of transplanted cells serves primarily to
reduce inflammation, not to promote remyelination (Franklin & Kotter 2008). While
transplantation efforts may still prove invaluable in an immunomodulatory capacity,
alternative strategies will likely be required to promote remyelination and to restore
function. For this reason, mobilization of endogenous precursor cells represents an
attractive alternate approach for the treatment of demyelinating conditions.
During development, the majority of oligodendrocyte precursor cells (OPCs)
differentiate into myelinating oligodendrocytes. However, a small but significant number
of OPCs remain in an undifferentiated state into adulthood (Chong & Chan 2010). In
demyelinating conditions, these adult OPCs are capable of differentiating into
oligodendrocytes with a limited ability to remyelinate axons (Franklin & Ffrench-
Constant 2008). Upon the initial onset of demyelination, adult OPCs are induced to
differentiate and remyelinate, effectively replacing lost oligodendrocytes. Unfortunately,
the capacity for remyelination is limited, and ultimately fails in the presence of chronic
demyelination. Current studies suggest two possible explanations for the failure of the
nervous system to facilitate long-term remyelination. One possibility is that adult OPCs
fail to proliferate continuously to an extent sufficient to repopulate demyelinated lesions.
 
 
4 
Another explanation is that adult OPCs ultimately lose their ability to differentiate into
remyelinating oligodendrocytes (Franklin 2002). It is also possible that a combination of
these two deficits contributes to the ultimate loss of function seen in demyelinating
conditions.
What factors are responsible for inhibiting differentiation and what is required to
overcome this inhibition in order to allow remyelination to proceed? To properly address
these questions, two specific hypotheses need to be considered. One possible explanation
is that environmental cues are preventing the OPCs from successfully differentiating and
remyelinating. Both the presence of inhibitory cues and an absence of inductive cues
could result in limited differentiation. Equally important to consider is the possibility that
intrinsic changes to the OPCs themselves are responsible for the failure to successfully
transition into oligodendrocytes. Recent work suggests that the reduced capacity for
regeneration may be due in part to age-related epigenetic changes (Shen et al 2008). This
study suggests that adult OPCs exhibit a decreased efficiency for recruitment of histone

deacetylases (HDACs), which ultimately results in permanent inhibition of myelin gene
expression (He et al 2007). Most likely, it is a combination of both environmental and
intrinsic factors that is responsible for the arrested development of adult OPCs. It is
important to note that adult OPCs are capable of extensive myelination when transplanted
into congenitally dysmyelinated brains (Windrem et al 2004). These results suggest that
adult OPCs retain a limited intrinsic capacity for differentiation and remyelination that is
extrinsically inhibited by the presence of a demyelinated environment. Stimulating
endogenous OPCs to remyelinate will likely require the removal of inhibitory
 
 
5 
environmental cues and the restoration of environmental signals that promote
differentiation and myelination. It is possible that the components of a continuously
demyelinating environment, which can occur in diseases such as MS, will ultimately
prove too inhibitory to allow for the ongoing differentiation of adult OPCs. This
unfortunate scenario may help explain in part why the capacity for successful
remyelination decreases over time. It is therefore crucial to examine the environmental
impact on normal OPC development. It is our goal to characterize the components of the
developmental microenvironment that are required for effective myelination. Considering
the tremendous diversity of cells present in the CNS, it is likely that heterogeneous
microenvironments exist that could differentially regulate myelination during
development. Identifying the nature of these microenvironments may allow for the
possibility to target inhibitory cues and to recreate permissive and instructive
environmental components in the aftermath of nerve injury and disease.

Developmental Differentiation and Myelination
A great deal remains unknown about the precise mechanisms governing differentiation
and myelination during development in vivo. It is generally accepted that an intrinsic
program contributes to the differentiation of individual OPCs cultured at clonal density,
and that these OPCs will undergo a set period of proliferation before differentiating (Raff
2006). However, the relevance of these findings to an in vivo environment is complicated
by the fact that both mature oligodendrocytes and OPCs continue to exist even in the
adult CNS. Therefore, not all OPCs in vivo are subject to an intrinsic developmental
 
 
6 
program that ultimately results in differentiation. How then is it possible to reconcile the
idea of an intrinsic developmental program with the fact that OPCs have more than one
fate choice? A possible explanation is that OPCs possess the intrinsic capacity to make a
multitude of choices, but their surrounding environment heavily influences which choice
they actually make. Previous findings demonstrate that OPCs are capable of
differentiating not only into oligodendrocytes, but also into astrocytes and neurons,
depending on their local environment (Zhao et al 2006). Interestingly, the ability of an
OPC to adopt either an astrocytic or neuronal fate in vitro is demonstrated by studies in
which OPCs are cultured in the absence of axons. We therefore speculate that while
OPCs may have the intrinsic ability to adopt multiple cell fates, in the context of
development, the axon may act as a type of instructive niche, maintaining OPCs as
precursors until other environmental factors induce differentiation.
Understanding how differentiation is regulated during development could offer
insight into both the intrinsic and extrinsic requirements for differentiation following
nerve injury or disease. Oligodendrocytes represent a terminally differentiated, non-
proliferative population of cells. For this reason the number of differentiated
oligodendrocytes is highly dependent on the extent of proliferation that occurs at the level
of the oligodendrocyte precursor cell (OPC). During development, OPCs are exposed to
proliferative signals as they migrate along axons throughout the CNS. In vitro studies
demonstrate that one of the main promoters of OPC proliferation is platelet-derived
growth factor (PDGF), which is secreted by astrocytes and neurons (Levine 1989; Noble
et al 1988; Raff et al 1988; Richardson et al 1988; Yeh et al 1991). In addition,
 
 
7 
transgenic mice with neurons overexpressing PDGF exhibit a significant increase in the
number of OPCs (Calver et al 1998). OPCs cultured in the presence of PDGF continue to
divide indefinitely, unless factors regulating cell cycle arrest are also present (Barres et al
1994; Durand et al 1998). For instance, the addition of growth factors such as thyroid
hormone (TH) and retinoic acid (RA) limits the proliferative capacity of cultured OPCs
(Barres et al 1994; Gao et al 1998). During development, migratory and proliferative
OPCs will eventually reach a final destination and the vast majority will differentiate into
myelinating oligodendrocytes. The final number of myelinating cells is determined by the
extent of OPC proliferation, as well as the number of OPCs that ultimately differentiate.
If proliferation terminates prematurely, the number of OPCs available for differentiation
will not be sufficient to successfully populate the CNS and deficits may result. On the
other hand, delayed differentiation can also lead to hypomyelination and impaired
signaling. It is therefore essential that the transition from OPC to oligodendrocyte be
temporally coordinated. Identifying the factors that regulate this transition is important
not just for understanding development but also for therapeutic treatment of
demyelinating diseases, particularly in light of growing evidence suggesting that OPCs
are involved in remyelination (Dawson et al 2003; Gensert & Goldman 2001; Horner et
al 2000; Levine et al 2001; Levison et al 1999; Nishiyama et al 1999; Rivers et al 2008).

The Relationship between Proliferation and Differentiation
In order to understand how OPCs transition to oligodendrocytes, it is necessary to
determine whether the factors regulating the cessation of proliferation are distinct from
 
 
8 
those that control the initiation of differentiation. Normal progression through the cell
cycle is modulated by interactions between cyclins and cyclin-dependent kinases (cdks).
The formation of cyclin-cdk complexes, such as the cyclinE-cdk2 complex, activates
signaling cascades that promote cell division. Formation of the cyclinE-cdk2 complex is
regulated by the cell cycle inhibitor protein, p27
Kip1
(Belachew et al 2002).
Overexpression of p27
Kip1
in OPCs in vitro arrests the cell cycle by inhibiting cdk2
activity (Tang et al 1999). The control of cell division by p27
Kip1
has been shown to play a
regulatory role in the proliferation of OPCs. Interestingly, in the presence of PDGF,
actively proliferating OPCs exhibit a gradual increase in p27
Kip1
expression (Durand et al
1997). In the absence of PDGF, when OPCs differentiate within 48 hours, the rate at
which p27
Kip1
expression increases is upregulated dramatically. This increase in p27
Kip1

expression correlates with withdrawal from the cell cycle (Casaccia-Bonnefil et al 1997;
Durand et al 1998; Tang et al 1998). However, experience a premature induction of cell
cycle arrest. However, instead of differentiating immediately, OPCs overexpressing
p27
Kip1
remain in an extended period of quiescence, and eventually differentiate at the
same time as control OPCs (Tang et al 1999). These findings suggest that the inhibition
of proliferation, as a result of cell cycle arrest, is not sufficient to induce differentiation of
OPCs. Corresponding in vivo experiments examine how the loss of p27
Kip1
affects
differentiation. In p27
Kip1-/-
mice, the number of proliferating OPCs increases (Casaccia-
Bonnefil et al 1999). However, the timing of differentiation is similar between knock-out
mice and wild-type mice, despite the increased proliferative capacity of p27
Kip1-/-
OPCs.
Together, these studies support the idea that proliferation and differentiation are
 
 
9 
regulated through distinct mechanisms. Similar results were obtained after transfection of
purified OPCs with a dominant-negative cdk2 (dn-cdk2), which inhibits the interaction
between endogenous cdk2 and cyclinE (Belachew et al 2002). Because an upregulation
of p27
Kip1
also inhibits the formation of the cyclinE-cdk2 complex, one would expect the
dn-cdk2 vector to inhibit proliferation. As expected, transfection with dn-cdk2
significantly decreased the proliferation of OPCs. However, the dn-cdk2 did not affect
OPC differentiation as compared to controls (Belachew et al 2002). These findings
indicate that proliferation and differentiation are regulated by distinct mechanisms.

Extrinsic Regulation of Oligodendrocyte Differentiation
Uncoupling proliferation and differentiation could potentially offer great insight into
general development and even into the treatment of demyelinating diseases. If the factors
that inhibit proliferation are not sufficient to induce differentiation, then it is essential to
identify the mechanisms responsible for initiating differentiation. Recent findings have
highlighted the specific effects of transcription factors such as Nkx2.2, Sox10 and Olig2
on the differentiation of OPCs (Qi et al 2001; Stolt et al 2002a; Zhou et al 2001). Equally
important is to understand the role of extrinsic factors in the regulation of differentiation
and myelination. Various studies have identified TH as an extrinsic regulator of
differentiation in vitro. TH may mediate the expression of the cell cycle inhibitor protein
p21
cip1
(Tokumoto et al 2001), which is required for the differentiation of OPCs (Zezula
et al 2001). The role of p21
cip1
is specific to the differentiation process, as cells isolated
from p21
cip1-/-
mice exit the cell cycle in a timely manner. This is in direct contrast to
 
 
10 
cells purified from p27
Kip1-/-
mice, which proliferate longer than wild-type and p21
cip1-/-

cells (Zezula et al 2001). This finding demonstrates that while p27
Kip1
is necessary for cell
cycle arrest, p21
cip1
is not. However, myelination in the cerebellum of p21
cip1-/-
mice is
decreased compared to wild-type mice (Zezula et al 2001). This result suggests that
p21
cip1
may be important for the proper differentiation of OPCs. These experiments
further suggest that proliferation and differentiation can be uncoupled by demonstrating
that the p27
Kip1
pathway is responsible for inhibiting proliferation, whereas a mechanism
mediated by p21
cip1
plays a role in initiating differentiation. How is the regulation of
differentiation by p21
cip1
affected by the presence of TH? Interestingly, treatment of
OPCs with TH was shown to promote an increase in the expression of p21
cip1
(Tokumoto
et al 2001). Additional experiments suggest that the effects of TH on p21
cip1
expression
are mediated through the tumor suppressor protein, p53, which has been shown to
activate transcription of p21
cip1
(Brugarolas et al 1995; Deng et al 1995). The infection of
purified OPCs with a dominant-negative form of p53 substantially inhibits TH-induced
differentiation (Tokumoto et al 2001). Based on this finding, the authors of this study
suggest that TH may regulate a p53 pathway that initiates differentiation by increasing
transcription of p21
cip1
. Together, these results provide a potential mechanism for the role
of TH in promoting differentiation.
TH represents one example of an extracellular signal that promotes OPC
differentiation under certain conditions. In contrast, Jagged1 and Delta1 represent
extracellular signals that can inhibit the differentiation of OPCs through activation of the
Notch signaling pathway (Wang et al 1998). OPCs from the optic nerve express the
 
 
11 
Notch1 receptor both in vivo and in vitro. In cultures of purified OPCs, addition of the
soluble Notch ligand Delta1 significantly inhibits the appearance of oligodendrocytes.
Purified OPCs cultured on top of cells expressing Jagged1, another Notch receptor
ligand, also fail to differentiate. Importantly, the expression of Jagged1 does not impair
the ability of OPCs to divide, thereby demonstrating that activation of the Notch pathway
specifically inhibits differentiation without affecting the proliferation process. These
findings correlate with in vivo studies examining oligodendrocyte development in Notch1
conditional knock-out mice. These mice exhibit premature oligodendrocyte
differentiation in multiple regions of the CNS (Genoud et al 2002). Additionally, the gene
inactivation of Notch1 results in the ectopic appearance of oligodendrocytes in the gray
matter of the spinal cord. Together, these findings suggest that activation of the Notch
pathway by extracellular signals can specifically inhibit OPC differentiation. In addition,
these studies suggest that oligodendrocyte differentiation may be subject to extrinsic
regulation from neurons. This is not surprising given the intimate nature of the
intercellular interaction between axons and myelinating oligodendrocytes.
Indeed, one landmark study suggests that axon diameter may represent a crucial
regulator of myelination (Voyvodic 1989). In these studies, an increase in the size of an
axonal target was shown to promote a corresponding increase in axon diameter. Results
from this work demonstrate that an increase in axon diameter can promote the
myelination of previously unmyelinated axons. Although these experiments were
performed on peripheral axons, changes in axon diameter were also regarded as a likely
regulator of CNS myelination. More recent studies suggest that an increase in axon
 
 
12 
diameter is not the only axonal factor responsible for myelination by oligodendrocytes.
This was demonstrated by experiments in which the addition of nerve growth factor
(NGF) to neuron-OPC cocultures inhibited myelination (Chan et al 2004). Importantly,
the effect of NGF was mediated through modulation of an axonal signal, and not through
a direct effect on oligodendrocytes. This was demonstrated by experiments examining the
effects of NGF on myelination of TrkA-expressing neurons as compared to TrkB-
expressing neurons. NGF failed to inhibit myelination of TrkB-expressing neurons,
suggesting that the effect of NGF was mediated through a specific interaction with
neuronal TrkA. These results therefore imply that activation of TrkA by NGF modulates
an axonal signal that controls oligodendrocyte myelination. It is possible that this effect is
mediated in part through LRR and Ig domain-containing, Nogo receptor-interacting
protein (LINGO-1). LINGO-1 was recently identified as an inhibitor of oligodendrocyte
myelination that is expressed on both oligodendrocytes and axons (Lee et al 2007; Mi et
al 2005). NGF signaling through TrkA promotes an increase in the axonal expression of
LINGO-1 (Lee et al 2007). Inhibition of LINGO-1 using either a DN-LINGO-1 lentivirus
or LINGO-1-Fc greatly enhances oligodendrocyte differentiation and myelination. These
studies suggest that LINGO-1 is capable of inhibiting both differentiation and
myelination. However, because the myelination studies were performed on OPC-DRG
cocultures, it is possible that the effects of LINGO-1 are specific to differentiation, and
that the observed reduction in myelination is simply a result of decreased numbers of
oligodendrocytes. If signals such as LINGO-1 are to be used as targets for treating
 
 
13 
demyelinating diseases, it will be important to determine the specific developmental
process that is impacted by manipulation of these factors.
Another possible candidate for an axonal signal that regulates myelination is the
cell adhesion molecule, polysialic acid-neural cell adhesion molecule (PSA-NCAM).
This is suggested by experiments in which dissociated cultures of cerebral hemispheres
were treated with an anti-PSA-NCAM antibody. Addition of the antibody resulted in an
increase in the number of myelinated axons (Charles et al 2000). This effect appears to be
specific to the process of myelination, as the addition of the anti-PSA-NCAM antibody
had no effect on the number of MBP-expressing oligodendrocytes. These results suggest
that PSA-NCAM represents an axonal inhibitor of oligodendrocyte myelination. While
the findings described here are intriguing, further studies are required to confirm the role
of axonal cues in regulating oligodendrocyte differentiation and myelination.

The Importance of Interactions between Oligodendroglial Cells
Clearly, the neuronal-glial interaction is an example of a particularly intimate
intercellular interaction that would be expected to play an important role in the regulation
of oligodendrocyte development and myelination. However, interactions between
oligodendroglial cells themselves represent an additional element that could also be
involved in the extrinsic regulation of the myelination process. Surprisingly, very little is
known about how interactions between oligodendroglial cells contribute to the regulation
of differentiation and myelination. It is possible that these types of intercellular
interactions could be important for both the spatial and temporal regulation of the
 
 
14 
myelination process. Interactions within a myelinating population of cells could play an
instrumental role in shaping processes such as the appropriate placement of non-
overlapping myelin internodes along axonal tracts. In the work presented here, we seek
to address a fundamental developmental question, “How does the developing CNS
generate the appropriate number of myelin internodes to perfectly myelinate all
necessary axonal segments? Akin to other aspects of nervous system development,
proper myelination requires a precise match between the number of myelinating cells and
myelin internodes, and the number of axonal segments requiring myelination. How does
this coordination occur? The precise and perfect myelination of the CNS will be a
function of both the number of oligodendrocytes and the number of myelin internodes
that each oligodendrocyte forms. The ability of oligodendrocytes to form multiple and
variable numbers of internodes represents an intriguing example of intrinsic plasticity
that has not yet been extensively explored. Here, we refer to this dynamic and inherent
plasticity as the myelinogenic potential of oligodendrocytes. In the work presented here,
we identify a role for intercellular interactions between oligodendroglial cells in
regulating both differentiation and myelinogenic potential. We propose a novel
mechanism by which the axons act indirectly to influence the onset of oligodendrocyte
differentiation through the regulation of oligodendrocyte precursor cell numbers. We then
extend this paradigm to suggest that the number of myelin internodes formed per
oligodendrocyte is dependent on the number of oligodendroglial cells.

 
 
15 
In the work presented here, we sought to address the potential importance of population-
based mechanisms in the following two questions:
1) How does the developing nervous system determine when a sufficient number of
OPCs has been generated and differentiation should begin?
2) What factors regulate the myelinogenic potential of individual oligodendrocytes?
In Chapter 2, we examine the factors that govern the coordinated differentiation of
oligodendroglia. Our studies identify a potential role for biomechanical interactions in the
temporal coordination of differentiation within a population of OPCs. In Chapter 3, we
investigate the role of population-based interactions in shaping the extent of myelination
by individual oligodendrocytes. We find that contact-mediated interactions between
oligodendroglial cells, mediated in part by Nogo-A, are involved in the regulation of
myelinogenic potential.
In addition to the relevance to developmental myelination and to remyelination,
we believe that the questions addressed here also have the potential to provide new
insight into general principles of nervous system development. We hope that the studies
in Chapter 2 may help explain how cells in a seemingly homogeneous population can
adopt two divergent fates. In addition, we believe that these studies demonstrate a novel
mechanism responsible for coordinating fate decisions within a given population of cells.
The studies outlined in Chapter 3 may also provide insight into population-based
interactions. We hypothesize that the mechanisms governing the myelinogenic potential
of oligodendrocytes may be reminiscent of those that regulate branching, arborization and
pruning patterns utilized in the refinement of developing neuronal projections (Luo &
 
 
16 
O'Leary 2005). We propose that the number of myelin segments formed per individual
cell may be determined through competition-based mechanisms similar to those involved
in repulsive guidance signaling and growth cone collapse. It is our hope that our studies
will provide insight into generalizable mechanisms that may govern arborization patterns
and process refinement in both glial and neuronal cells. Finally, we believe that
collectively, the studies described here can also provide insight into how ratios between
groups of cells are appropriately matched. While our studies are focused specifically on
matching the number of oligodendrocytes and myelin segments to the number of axons
requiring myelination, these findings could have implications for how numbers of
neurons, axons and dendrites are appropriately matched to their targets- be it other
populations of neurons or target populations such as muscle, retina, or skin cells. We
believe that understanding the environmental regulation of differentiation and
myelination has direct relevance to both the treatment of demyelinating conditions, and to
the principles that govern nervous system development.

 
 
17 
CHAPTER TWO
The Geometric and Spatial Constraints of the Microenvironment Induce
Oligodendrocyte Differentiation

The oligodendrocyte precursor cell arises from the subventricular zone during early
vertebrate development to migrate and proliferate along axon tracts before differentiating
into the myelin-forming oligodendrocyte. We demonstrate that the spatial and temporal
regulation of oligodendrocyte differentiation is intimately dependent on the axonal
microenvironment and the density of precursor cells along a specified axonal area.
Differentiation does not require dynamic axonal signaling, but instead is induced by
packing constraints resulting from intercellular interactions. Schwann cells and even
artificial beads bound to the axonal surface can mimic these constraints and promote
differentiation. Together these results describe the coordinately controlled biophysical
interaction of oligodendrocyte precursors within an axonal niche leading to self-renewal
and differentiation.

 
 
18 
Introduction
Damage to the myelin membrane, as a result of nerve injury or disease, significantly
impairs the ability of the nervous system to communicate and can lead to a host of
debilitating symptoms, as well as an ultimate loss of function. In the central nervous
system (CNS), demyelination is accompanied by the loss of oligodendrocytes, the
terminally differentiated cells responsible for the formation of the myelin sheath.
Following the initial onset of demyelination, oligodendrocyte precursor cells (OPCs) are
induced to differentiate and remyelinate, effectively replacing lost oligodendrocytes.
Unfortunately, the capacity for remyelination is limited, and ultimately fails in the
presence of chronic demyelination. It remains unclear why the CNS cannot sustain this
initial ability to repair the myelin sheath. One possible explanation is that adult OPCs
eventually lose their ability to differentiate into remyelinating oligodendrocytes (Franklin
2002). It is plausible that the continuous presence of a demyelinating environment is
responsible for inhibiting the differentiation process. If this is true, then it is imperative to
identify the environmental conditions conducive to the ongoing production of
oligodendrocytes.
Examining oligodendrocyte generation during development could prove useful for
determining the role of the environment in the induction of differentiation. Developing
OPCs are proliferative and self-renewing cells that originate in the subventricular zone
(SVZ) and migrate along axons throughout the CNS. During development, an OPC must
decide how many times it will divide, and where it will migrate. Additionally, an OPC
must choose whether to remain as a precursor cell into adulthood or to differentiate into
 
 
19 
a myelinating oligodendrocyte. Such complex decisions are likely to be heavily
influenced by the nature of the surrounding environment, and by the behavior of
neighboring cells. Based on these assumptions, it is our goal to identify the
environmental factors that influence the decision of an OPC to differentiate into an
oligodendrocyte. To accomplish this goal, we first looked at the developing rat spinal
cord to examine the temporal regulation of oligodendrocyte differentiation in vivo.

 
 
20 
Methods

Immunopanning Protocol
Oligodendrocyte precursor cells (OPCs) were purified from 6-7 day old (P6-P7) rat brain
cortices with a panning protocol adapted from one previously described (Chan et al
2004). Briefly, petri dishes containing a goat anti-mouse IgG +IgM secondary antibody
solution (Jackson Laboratories) were incubated overnight. Dishes were rinsed and
incubated with primary antibody solutions containing either GalC or A2B5 hybridoma
supernatants. Rat brain cerebral hemispheres were first diced and then dissociated with
papain at 37° C. Following trituration, cells were resuspended in a panning buffer and
then incubated at room temperature sequentially on three immunopanning dishes:
IgG+IgM, GalC and A2B5. A2B5
+
OPCs were released from the final panning dish using
trypsin (Sigma).

Purified OPC/DRG Cocultures
OPC-DRG cocultures were prepared as described previously (Chan et al 2004). Briefly,
DRG neurons from E13-15 Sprague-Dawley rats were dissociated, plated and purified on
collagen coated coverslips in the presence of NGF (100 ng/ml). Neurons were maintained
for 2-3 weeks prior to the addition of OPCs. Either the TrkA-Fc receptor chimera (1
ug/ml; Regeneron Pharmaceuticals) or the anti-TrkA antibody (RTA, 50 ug/ml) was
added to the DRG cultures one week prior to the addition of OPCs. OPCs were seeded at
either low (20,000 OPCs), medium (200,000 OPCs), or high (2 million OPCs) density
 
 
21 
onto coverslips containing purified DRG neurons. Coverslips were incubated in a small
volume of MEM medium overnight to facilitate OPC attachment. The following day,
coverslips were transferred into wells with MEM medium containing 10% FBS and either
anti-NGF or TrkA-Fc. All OPC/DRG coculture experiments were conducted using MEM
medium containing 10% FBS and either anti-NGF or TrkA-Fc, with the exception of the
experimental condition in Supplemental Fig 2 in which 50 ng/ml of PDGF-AA
(Peprotech) was added to the medium.

Western Blot Analysis
Samples from cocultures and rat spinal cords were prepared for Western blot as
previously described (Chan et al 2006). The proteins were transferred to pure
nitrocellulose membranes and probed with specific antibodies. Antibodies for Westerns:
rabbit polyclonal anti-PDGFRa antibody (Santa Cruz), rabbit polyclonal anti-NG2
antibody (Chemicon), rat monoclonal anti-MBP antibody (Chemicon), mouse
monoclonal anti-GFAP antibody (Chemicon), mouse anti-MAG antibody (Chemicon),
and mouse monoclonal anti-b-Actin (Sigma). The Alexa Fluor goat anti-rabbit, anti-
mouse, and anti-rat 680 IgG antibodies were used as secondary antibodies for near-
infrared fluorescent detection performed on the LI-COR Odyssey Infrared Imaging
System.

 
 
22 
Immunostaining
Immunostaining of rat spinal cord sections and cocultures was performed as previously
described (Lee et al 2007). Briefly, cocultures were fixed using 4% paraformaldehyde
and dehydrated and then permeabilized and blocked by incubation with 20% goat serum
and 0.2% Triton X-100 in PBS. Differentiated oligodendrocytes and myelin were
detected with a rat monoclonal anti-MBP antibody (Chemicon). OPCs were detected
using a rabbit polyclonal anti-PDGFRa antibody (Santa Cruz). Astroyctes were detected
using a mouse monoclonal anti-GFAP antibody (Chemicon). Axons were detected using
a mouse monoclonal antibody to neurofilament (American Type Culture Collection).
Schwann cells were detected using a rabbit polyclonal antibody to S100
(DakoCytomation). The Alexa Fluor anti-rat 594, anti-rabbit 488, anti-mouse 350, 488
and 594 IgG antibodies (Invitrogen) were used as secondary antibodies for fluorescence
detection. Cell nuclei were examined with DAPI.

Axonal Fixation
Prior to the seeding of OPCs, coverslips containing purified DRG neuronal cultures were
washed twice gently in PBS. Neurons were then fixed with 2 mls of a 4%
paraformaldehyde (PFA) solution for 10 minutes. Following the removal of PFA,
neurons were gently rinsed multiple times with PBS and then transferred to fresh wells
with MEM medium containing 10% FBS. All fixed axon coculture experiments were
conducted using MEM medium containing 10% FBS and either anti-NGF or TrkA-Fc.

 
 
23 

Electron Microscopy
High density OPCs co-cultured with live or fixed DRGs for 7 days were fixed in 2%
glutaraldehyde, stained with 1% osmium tetroxide and counterstained with 1% uranyl
acetate overnight. Co-cultures were subsequently rinsed with distilled water, dehydrated
in ethanol and embedded in resin (EMBed-812, Electron Microscopy Sciences).
Ultrathin sections (70nm) were obtained and visualized with a JEOL JEM 1400 Electron
Microscope.

Schwann Cell/OPC/DRG Cocultures
Schwann cells were collected from postnatal day 2 rats as described previously (Chan et
al 2006). Schwann cells were purified with cytosine arabinoside and then seeded at either
low (200,000 Schwann cells) or high (2 million Schwann cells) density onto live purified
DRG neurons. 200,000 OPCs were added to Schwann cell/DRG cocultures 2-3 days after
Schwann cells were seeded. All Schwann Cell/OPC/DRG coculture experiments were
conducted using MEM medium containing 10% FBS and either anti-NGF or TrkA-Fc.

Bead/OPC/DRG Cocultures
5, 20, and 100 mm Protein A beads were incubated overnight with an mouse monoclonal
antibody (hybridoma supernatant) to p75 neurotrophin receptor (p75
NTR
). The beads were
then washed multiple times with MEM medium. In initial experiments (Fig 3), 20 mm
beads were added at either low or high density to live purified DRG neurons. In
 
 
24 
subsequent experiments (Fig 4), 5, 20, and 100 mm beads were each added at high
density to live purified DRG neurons. The attachment of the beads to the axons was
facilitated by the antibody-mediated interaction with neuronal p75
NTR
. Beads were
incubated on the neurons for 1-3 hours before the addition of 200,000 OPCs. 5 and 20
mm polystyrene beads were obtained from G. Kisker-Products for Biotechnology. 100
mm sepharose beads were obtained from Zymed Laboratories, Inc. All Bead/OPC/DRG
coculture experiments were conducted using chemically defined medium (Chan et al
2004) to avoid the potential for IgGs in serum-containing medium to disrupt the
antibody-mediated binding of the beads to the axons.

 
 
25 
Results
Temporal Regulation of Oligodendrocyte Differentiation
OPCs appear in the rat spinal cord as early as embryonic day 15 (E15). These precursor
cells can be seen to increase in number as they migrate throughout the spinal cord (Fig
2.1A). OPCs continue to proliferate until shortly after birth, at which time the number of
OPCs becomes relatively constant (Fig 2.1 A and C). Beginning around postnatal day 8
(P8), we observe a gradual decline in the number of OPCs (Fig 2.1 A and C). This
decrease in OPC numbers is followed by the appearance of differentiated
oligodendrocytes, which are undetectable in the rat spinal cord until approximately one
week postnatal (Fig 2.1 B and C). These in vivo observations suggest that developing
OPCs make a temporally synchronized transition from proliferating progenitors to
differentiated oligodendrocytes. Using an in vitro coculture system involving sensory
dorsal root ganglion (DRG) neurons and purified OPCs (Chan et al 2004), we can
consistently replicate this temporally specific pattern of differentiation. In our system, we
find that after seeding a standard density of OPCs (200,000), the precursor cells undergo
a fixed period of proliferation prior to differentiation. Differentiation begins
approximately ten days after the OPCs are seeded onto the neurons (Fig 2.1 D and E).
This sequence of events closely approximates the pattern of OPC development that we
observe in the rat spinal cord. What mechanisms coordinate the initiation of
oligodendrocyte differentiation that we see both in our cocultures and in vivo? In other
words, how is the timing of oligodendrocyte differentiation synchronized between OPCs?
 
 
26 
Figure 2.1: Temporal regulation of oligodendrocyte differentiation
(A) Immunostaining in rat spinal cord sections from embryonic day 10 (E10) through postnatal
day 20 (P20). OPCs, identified by immunostaining for platelet-derived growth factor receptor α
(PDGFRα), appear initially at E15 and then proliferate as they migrate throughout the spinal
cord. (B) Immunostaining in rat spinal cord sections from P0 through P10. Astrocytes (blue) are
identified by immunostaining for glial fibrillary acidic protein (GFAP). Oligodendroyctes (red),
identified by immunostaining for myelin basic protein (MBP), do not appear in the spinal cord
until P5, almost two weeks after the initial appearance of OPCs. (C and D) Western blots of rat
spinal cords (C) and OPC-DRG cocultures (D) probed for proteins expressed by OPCs (NG2),
oligodendrocytes (MBP), and astrocytes (GFAP). Actin serves as a loading control. Cocultures
replicate the temporal expression pattern seen in vivo, in which the appearance of
oligodendrocytes is delayed in comparison to the appearance of OPCs. (E) In OPC-DRG
cocultures, immunostaining for MBP labels differentiated oligodendrocytes (red). Nuclei are
stained using DAPI (blue). Differentiation begins 10 days after a standard density of 200,000
OPCs is seeded onto neurons. After 15 days, a robust and reproducible amount of myelination is
observed. The onset of differentiation is preceded by a period of OPC proliferation, a pattern
which approximates the behavior of OPCs in the spinal cord. (F-H) Immunostaining and Western
blots of OPC-DRG cocultures in which OPCs are seeded at (F) low density (20,000 OPCs), (G)
standard density (200,000 OPCs), and (H) high density (2 million OPCs) onto DRG neurons.
Immunostaining for MBP labels differentiated oligodendrocytes (red). Nuclei are stained using
DAPI (blue). Western blots of cocultures are probed for proteins expressed by OPCs (PDGFRα),
oligodendrocytes (MBP), and astrocytes (GFAP). Actin serves as a loading control. These results
demonstrate that an increase in cell density accelerates the onset of oligodendrocyte
differentiation, but does not affect the differentiation of OPCs into astrocytes.
 
 
27 

Figure 2.1: Continued

 
 
28 
Induction of Differentiation Requires a Critical Density of OPCs
It is possible that the global synchronicity of this cell fate decision can be explained by
previous studies which suggest that OPCs differentiate based on the presence of an
intrinsic timer (Raff 2006). Essentially, OPCs are programmed to undergo a set period of
division and then differentiate. Related OPCs possess highly similar programs and are
coordinated in the timing of their differentiation, even when they are relocated to separate
environments. It is important to note that the intrinsic timer was identified in studies
performed at clonal density in the absence of other cell types (Gao et al 1997; Temple &
Raff 1986). In contrast, an OPC in our system is subject to the potential external
influences of axons and neighboring OPCs. Can an intrinsic program explain the timing
of oligodendrocyte differentiation in the presence of these extrinsic influences? To test
this possibility, we seeded neurons with three different densities of OPCs (Fig 2.1 F-H).
If the initiation of differentiation is regulated by an intrinsic timer, then OPCs in all three
experimental conditions should differentiate simultaneously. Instead, we find that the
time at which differentiation is initiated depends on the density at which OPCs are
seeded. OPCs seeded at high density (2 million OPCs) start to differentiate after only five
days in culture (Fig 2.1H), whereas OPCs seeded at our standard density (200,000 OPCs)
and low density (20,000 OPCs) must be cultured for approximately two (Fig 2.1G) and
three weeks (Fig 2.1F), respectively, before differentiation begins. These results suggest
that an intrinsic timer is not responsible for the initiation of differentiation in our
coculture system. Instead, it appears that a critical density of OPCs must be reached in
order for differentiation to begin. This conclusion is supported by Western blots (Fig 2.1
 
 
29 
F-H) showing the expression profiles of the OPC marker, platelet-derived growth factor
receptor α (PDGFRα), and the oligodendrocyte marker, myelin basic protein (MBP).
Regardless of the initial density of OPCs seeded, the expression of PDGFRα must reach
a threshold level before MBP expression is detected. In addition, these findings
demonstrate that in our cocultures, the extent of oligodendrocyte differentiation is not
proportional to the number of OPCs initially seeded. Together these studies support the
hypothesis that extrinsic factors must regulate the onset of oligodendrocyte
differentiation.

Differentiation and Myelination in the Absence of Dynamic Axonal Signaling
It is likely that neurons serve as extrinsic regulators of OPC development. What role does
the axon play in the process of oligodendrocyte differentiation? Similar to previous
studies (Zhang & Miller 1996), we find that oligodendrocyte differentiation is not density
dependent in the absence of axons (not shown). These results suggest that interactions
between axons and OPCs may be responsible for the density-dependent coordination of
oligodendrocyte differentiation. To test this possibility, we treated our cocultures with
either conditioned medium from axons or with purified axonal membranes (not shown).
Because these classic experiments failed to induce oligodendrocyte differentiation, we
took an alternative approach to clarify the role of the axon. To eliminate dynamic
changes in axonal signaling, we fixed axons with paraformaldehyde prior to seeding
OPCs. We find that high-density OPCs seeded onto fixed axons differentiate with the
same timing and robustness as OPCs on live axons (Fig 2.2 A and B). To our surprise,
 
 
30 
the OPCs seeded onto fixed axons could also form compact myelin (Fig 2.2 C-E). These
results clearly indicate that dynamic interactions between oligodendrocytes and axons are
not required for either differentiation or myelination.
 
 
31 
Figure 2.2: The induction of differentiation at a critical density of OPCs does not require
dynamic axonal signaling
(A and B) Immunostaining of cocultures 5 days after seeding a high density of OPCs. Axons
(green) are identified by immunostaining for neurofilament (NF). (A) Live axons. (B) Axons
fixed with 4% paraformaldehyde to eliminate dynamic axonal signaling. Immunostaining with
MBP (red) demonstrates that fixed axons support oligodendrocyte differentiation and myelination
in a manner comparable to live axons. Nuclei are stained using DAPI (blue). (C-E) Electron
micrographs of oligodendrocyte myelination. The compact, multi-layered myelin formed by
oligodendrocytes on live axons (C-D) is also seen in fixed axon cocultures (E). (F-I)
Quantification of the critical density of OPCs required to induce population-wide differentiation
after 5 days in culture. A density of approximately 500-600 PDGFRα
+
OPCs on axons per
millimeter squared is required for the induction of oligodendrocyte differentiation on both fixed
axons (F and G) and on live axons (H and I). Here we define population-wide differentiation as
approximately 60-100 MBP
+
oligodendrocytes per millimeter squared. Note that because OPCs
seeded onto fixed axons (F and G) fail to proliferate, population-wide differentiation is induced
only when eight million (8M) OPCs are initially plated. In contrast, OPCs seeded onto live axons
(H and I) at an initial density of 2M, 4M, or 8M cells will all reach the critical density required
for differentiation. Note that after 5 days, OPCs seeded onto live axons at an initial density of 2M
have just reached the critical density and are just beginning to differentiate. Error bars represent
standard deviation. MBP
+
oligodendrocytes were quantified by counting 20 fields/coverslip, 3
coverslips/density. *p<0.01 versus 1M density cultures (Student-Newman-Keuls post hoc
comparison after one-way ANOVA).
 
 
32 
Figure 2.2: Continued
 
 
33 
Inhibition of Proliferation is Not Sufficient to Induce Differentiation
Could differentiation result from the presence of a permissive environment rather than
from induction by an instructive cue? Our findings suggest that the onset of
differentiation is somehow inextricably tied to the process of OPC proliferation. Perhaps
differentiation is merely the result of inhibited proliferation that occurs once a critical
density of OPCs on axons is reached. After all, it is plausible that achieving a high
density of OPCs could inhibit proliferation by depleting available mitogens or through
contact-mediated interactions. If this is true, than an experimental inhibition of
proliferation should induce oligodendrocyte differentiation. To test this possibility, we
seeded our standard density of OPCs onto fixed axons (Fig 2.3). It is known that axons
secrete OPC mitogens such as platelet-derived growth factor, PDGF (Calver et al 1998).
In our cocultures, axonal fixation eliminates the secretion of these mitogenic factors, and
greatly reduces the extent of OPC proliferation (Fig 2.3). These findings are in line with
previous studies demonstrating that axonal fixation selectively halts proliferation, but
does not affect the viability of glial cells (Salzer et al 1980). Despite the inhibition of
proliferation in our fixed axon cocultures, the OPCs failed to differentiate in advance of
OPCs on live axons. In fact, even after 20 days, very few differentiated oligodendrocytes
are detectable in these fixed axon cocultures (Fig 2.3). These results suggest that the
absence of mitogenic factors prevents OPCs from reaching the critical density required
for differentiation.
 
 
34 
Figure 2.3: The inhibition of proliferation is not sufficient to
induce differentiation
(A and B) Immunostaining of cocultures 15 days after seeding a
standard density of 200,000 OPCs onto DRG neurons. Axons
(blue) are identified by immunostaining for neurofilament (NF).
(A) Live axons. (B) Axons fixed with paraformaldehyde to
eliminate dynamic axonal signaling. OPCs (green) are
identified by immunostaining for platelet-derived growth factor
receptor α (PDGFRα). The number of OPCs is reduced on
fixed axons as compared to live axons. Differentiated
oligodendrocytes (red), identified by immunostaining for
myelin basic protein (MBP), are largely absent from fixed axon
cocultures. (C) Quantification of changes over time in the
number of PDGFRα+ OPCs per millimeter squared on live and
fixed axons. Compared to live axon cocultures, the proliferation
of OPCs is greatly reduced on fixed axons. (D) Quantification
of changes over time in the number of MBP+ oligodendrocytes
per millimeter squared on live and fixed axons. Compared to
live axon cocultures, the extent of differentiation on fixed axons
is greatly reduced, suggesting that an inhibition of proliferation
is not sufficient to induce differentiation. Error bars represent
standard deviation.
 
 
 
35 
Additionally, we tested whether it was possible to delay differentiation by increasing the
amount of available mitogens. The addition of exogenous PDGF to live axon cultures
does not inhibit OPC differentiation and may actually advance the differentiation process
(Fig 2.4). Taken together, these results demonstrate that the induction of differentiation at
a critical density of OPCs is not the result of an inhibition of proliferation.

Figure 2.4: An increase in available mitogens is not sufficient to delay
differentiation
(A) Immunostaining of live axon cocultures 15 days after seeding a standard density
of 200,000 OPCs. Oligodendrocytes (red) are identified by immunostaining for
myelin basic protein (MBP). OPCs are identified by immunostaining for platelet-
derived growth factor receptor α (PDGFRα). Nuclei are stained using DAPI (blue).
The addition of 50 ng/ml of platelet-derived growth factor AA (PDGF-AA) does not
delay the onset of differentiation as compared to control cultures. (B) Western blots
of live axon cocultures seeded with 200,000 OPCs. Cocultures with (+) and without
(-) the addition of 50 ng/ml of exogenous PDGF-AA, as well as sensory DRG
neurons cultured alone (DRG), are probed for proteins expressed by OPCs [NG2
and PDGFRα], oligodendrocytes [myelin-associated glycoprotein (MAG) and
myelin basic protein (MBP)], and astrocytes [glial fibrillary acidic protein (GFAP)].
Actin serves as a loading control. These results suggest that the addition of excess
mitogen does not delay the onset of differentiation and may actually accelerate the
initiation of this process.
 
 
 
36 
Spatial and Geometric Constraints Induce Differentiation
Why do OPCs need to reach a critical density in order to differentiate? Quantification of
the critical density suggests that differentiation is induced once OPCs reach a density of
approximately 500-600 platelet-derived growth factor receptor α-positive (PDGFRα
+
)

cells per millimeter squared. This density is not dependent on the number of OPCs
initially seeded and is highly conserved between live and fixed axon cultures (Fig 2.2 F-
I). Additionally, quantification of MBP
+
cells suggests that the number of differentiated
oligodendrocytes is not proportional to the number of OPCs initially seeded. These
results confirm that a significant amount of differentiation is induced only after a critical
density of OPCs is reached. Is it possible that the induction of differentiation at a critical
density requires density-dependent interactions between OPCs? To test this possibility,
we attempted to induce differentiation in cocultures containing both OPCs and Schwann
cells seeded onto live axons. When our standard density of OPCs is seeded onto neurons
either alone (Fig 2.1 E and G) or in the presence of a low density of Schwann cells (Fig
2.5A), we fail to see any differentiated oligodendrocytes after 5 days in culture. However,
when seeded with a high density of Schwann cells, we see a significant amount of
differentiation after 5 days (Fig 2.5B). These findings suggest that differentiation is not
controlled by the expression of a signal exclusive to OPCs. Instead, the addition of a high
density of Schwann cells is sufficient to mimic the critical density required for the
initiation of differentiation.
 
 
37 
Figure 2.5: Spatial constraints along an axon are sufficient to induce differentiation
(A and B) Immunostaining of OPC-DRG live axon cocultures 5 days after seeding a standard
density of 200,000 OPCs. As shown by staining for S100 (green), cocultures are also seeded with
either a low (A) or high (B) density of Schwann cells. Oligodendrocytes (red) are identified by
immunostaining for MBP. Nuclei are stained using DAPI (blue). OPCs are induced to
differentiate only in the presence of a high density of Schwann cells (B). (C) Phase contrast
microscopic images of polystyrene beads. Beads are 20 µm in diameter, comparable in size to the
cell body of an OPC (left panel) and Schwann cell (not shown). Beads were coated with an
antibody to an axonal protein (p75 neurotrophin receptor) and then conjugated at either low
density (middle panel) or high density (right panel) along the length of axons. (D and E)
Immunostaining of OPC-DRG cocultures 5 days after seeding 200,000 OPCs and either a low (D)
or high (E) density of polystyrene beads. OPCs are induced to differentiate only in the presence
of a high density of beads (E). Oligodendrocytes are identified by immunostaining for MBP (red).
Axons are identified by immunostaining for NF (green). Nuclei are stained using DAPI (blue).
Beads are visualized using DIC microscopy.

 
 
38 
Figure 2.5: Continued

 
 
39 
Because Schwann cells and OPCs originate from distinct cell lineages and do not
interact in vivo, it is not clear why Schwann cells would express a signal that induces
OPCs to differentiate. We therefore propose an alternative explanation for the ability of
Schwann cells to induce oligodendroycte differentiation. We hypothesize that an OPC
differentiates not because of instructive intercellular signaling, but instead because of a
restriction in axonal space proximal to the OPC. To test this hypothesis, Schwann cells
were fixed and seeded onto live axons along with a standard density of OPCs (Fig 2.6).
When seeded at a high density, fixed Schwann cells were sufficient to induce
oligodendrocyte differentiation after only 5 days. These results suggest that secreted
factors do not control the ability of Schwann cells to induce differentiation, adding
support to the hypothesis that differentiation is induced through a restriction of available
axonal space.

 
 
40 

To rule out the possibility that a membrane-bound factor is required to induce
oligodendrocyte differentiation, we conjugated axons with polystyrene beads measuring
20 µm in diameter, which is comparable in size to both OPCs and Schwann cells (Fig
2.5C). Coating the beads with an antibody against an axonal protein allowed us to bind
the beads at low and high density along the surface of live axons (Fig 2.5C). When a
standard density of OPCs was seeded onto neurons with a low density of beads, no
oligodendrocytes were observed after 5 days in culture (Fig 2.5D). However, we find that
an increase in the density of beads along axons is sufficient to induce differentiation on
either live (Fig 2.5E or fixed axons (Fig 2.7).
Figure 2.6: Secreted factors are not responsible for the induction of
oligodendrocyte differentiation by Schwann cells.
High density Schwann cells (2 million) were fixed with 4%
paraformaldehyde and seeded onto live axons. Cocultures were then seeded
with a standard density of OPCs (200,000). (A and B) Phase contrast
microscopic images of cocultures 5 days after seeding OPCs. (B and C)
Oligodendrocytes (red) are identified by immunostaining for myelin basic
protein (MBP). (C) Nuclei are stained using DAPI (blue). These results
suggest that the induction of differentiation is dependent on a contact-
mediated interaction between neighboring cells.
 
 
 
41 

These findings indicate that the induction of differentiation at a critical density of OPCs
is dependent on a restriction of available axonal space. However, in the absence of
axons, the polystyrene beads do not influence differentiation (not shown). Additionally,
we find that the induction of differentiation is dependent not only on the density of beads
along an axon, but also on their dimensions. Either an increase or decrease in the
diameter of the beads eliminates the potential to induce differentiation (Fig 2.8). These
results suggest that the induction of oligodendrocyte differentiation depends on both the
spatial and geometric parameters of the surrounding microenvironment.
Figure 2.7: Dynamic axonal signaling is not required for the induction of
differentiation by spatial and geometric constraints.
(A and B) Axons were fixed with 4% paraformaldehyde. 20 mm polystyrene
beads were then coated with an antibody to the neurotrophin receptor p75 and
conjugated at high density to the fixed axons. A standard density of OPCs were
seeded onto the axons and immunostained after 5 days. Polystyrene beads were
visualized using DIC microscopy. (A) Axons are identified by immunostaining
for neurofilament (green). (B) Oligodendrocytes (red) are identified by
immunostaining for myelin basic protein (MBP). Nuclei are stained using DAPI
(blue).
 
 
 
42 

Figure 2.8: The induction of differentiation is dependent on
the geometry of the microenvironment
(A) Phase contrast microscopic images of beads measuring
either 5 (left), 20 (center), or 100 (right) µm in diameter. Beads
were coated with an antibody to an axonal protein (p75
neurotrophin receptor) and then conjugated at high density
along the length of axons. (B-D) Immunostaining of OPC-
DRG cocultures 5 days after seeding a standard density of
200,000 OPCs and a high density of beads measuring either 5
(B), 20 (C), or 100 (D) µm in diameter. OPCs are induced to
differentiate only in the presence of the 20 µm beads (C).
Oligodendrocytes are identified by immunostaining for MBP
(red). Axons are identified by immunostaining for NF (green).
Nuclei are stained using DAPI (blue). Beads are visualized
using DIC microscopy.
 
 
 
43 
Discussion
Our results suggest that physical characteristics of the microenvironment can influence
cell fate decisions. We hypothesize that spatial and geometric constraints along an axon
may induce differentiation through lateral compression of OPCs. How does a mechanical
stimulus promote a change in cell fate? One possibility is that contractile forces can
physically alter the size or shape of an OPC. Changes in cell shape have previously been
shown to regulate processes such as cell survival (Chen et al 1997) and cell fate decisions
(McBeath et al 2004). It has been suggested that changes in cell shape may facilitate
structural rearrangement within the cell (Ingber 1997). This intracellular reorganization
could initiate interactions between upstream mediators and downstream effectors that are
responsible for the induction of differentiation (Boudreau & Jones 1999). Additionally, it
has been demonstrated that changes in cell shape can be directly linked to structural
changes in the nucleus (Maniotis et al 1997). In our system, we observe that an increase
in cell density correlates with a decrease in nuclear size, as shown by staining with DAPI.
It is possible that changes to the size and structure of the nucleus induce transcriptional
activity necessary for oligodendrocyte differentiation. Perhaps an increase in cell density
affects transcription factors known to regulate differentiation, such as Nkx2.2, Sox10,
Olig2 and the recently identified Yin Yang 1 (He et al 2007; Qi et al 2001; Stolt et al
2002b; Zhou et al 2001). Understanding the relationship between intrinsic and extrinsic
factors could help identify environmental conditions that will promote oligodendrocyte
differentiation in the presence of chronic demyelination.
 
 
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Treatment of demyelinating conditions may also require elucidation of the
relationship between the various environmental stimuli that have been shown to regulate
differentiation. In our studies we find that the presence of the axon is required for the
induction of differentiation through geometric and spatial constraints. Our results are in
line with previous studies, which suggest that the nature of the cellular substrate can
influence the effect of a mechanical stimulus on cell fate decisions (Ingber 1997). For
instance, the fate of mesenchymal stem cells can be modulated by the flexibility of the
substrate on which the cells are cultured (Engler et al 2006). It is possible that biophysical
characteristics of the axon, such as its size, shape, and tensile strength could be
instrumental in the density-dependent induction of differentiation. After all, the caliber of
the axon has previously been shown to influence multiple aspects of the myelination
process. Studies suggest that axon diameter can regulate whether axons are myelinated
(Voyvodic 1989), the thickness of the myelin membrane, and the distribution of
internodes along an axon (Trapp & Kidd 2000). However, recent studies suggest that in
the peripheral nervous system the regulation of myelin sheath thickness may be due
solely to the fact that an increase in axon diameter correlates with an increase in the
expression of the membrane-bound axonal factor neuregulin 1 (NRG1) type III
(Michailov et al 2004; Taveggia et al 2005). As illustrated by these findings, it will be
important to determine whether it is biophysical or biochemical attributes (or a
combination of both) that make the axon a necessary component of density-dependent
differentiation.
 
 
45 
It is possible that the induction of differentiation through environmental
constraints is dependent not on the structural dimensions of the axon, but instead on the
expression of membrane-bound axonal signals. Perhaps these signals facilitate the ability
of an OPC to transduce mechanical forces into the biochemical initiation of
oligodendrocyte differentiation. This type of context-dependent regulation could
represent an alternative explanation for the role of an axonal substrate in the density-
dependent induction of oligodendrocyte differentiation. It is possible that the mechanical
induction of differentiation depends on the formation of an adherens junction between the
axon and the OPC. This theory is supported by studies that implicate extracellular matrix
(ECM) receptors and cell-cell adhesion molecules as key mediators of
mechanotransduction (Alenghat & Ingber 2002). Interestingly, adhesion molecules have
previously been shown to play a role in oligodendrocyte differentation and myelination
(Buttery & ffrench-Constant 1999; Charles et al 2000; Fewou et al 2007; Schnadelbach et
al 2001; Tait et al 2000). Additionally, ECM receptors such as integrins are also known
to modulate the effects of extrinsic growth factors on various aspects of oligodendrocyte
development (Baron et al 2005; ffrench-Constant & Colognato 2004). It is therefore
plausible that the neuronal-glial interaction facilitates the formation of an adhesion
complex that is responsible for the integration of various chemical and mechanical factors
regulating differentiation (Schwartz & Ginsberg 2002). This hypothesis could help
explain how the axon can regulate differentiation through the expression of putative
factors such as Jagged-1 (Genoud et al 2002; Wang et al 1998) and Lingo-1 (Lee et al
2007; Mi et al 2005), as well as through its role in mechanotransduction. As novel
 
 
46 
differentiation factors continue to be identified, it is essential to understand how a
multitude of diverse extrinsic factors can be properly synthesized in the coordination of a
single cell fate decision. Identifying the mechanisms responsible for the integration of
extrinsic signals may be crucial for the establishment of an environment promoting
remyelination.

 
 
 
 
 
 
 
 
 
 
 
 
47 
CHAPTER THREE
Nogo-A establishes spatial segregation and extent of myelination
in the developing CNS

A requisite component of nervous system development is the achievement of cellular
recognition and spatial segregation through competition-based refinement mechanisms.
Competition for available axon space by myelinating oligodendrocytes ensures that all
relevant CNS axons are myelinated properly. To ascertain the nature of this competition,
we generated a transgenic mouse with sparsely labeled oligodendrocytes and establish
that individual oligodendrocytes occupying similar axon tracts can greatly vary the
number and lengths of their myelin internodes. Here we show that intercellular
interactions between competing oligodendroglia influence the number and length of
myelin internodes and identify the amino-terminal region of Nogo-A, expressed by
oligodendroglia, as necessary and sufficient to inhibit this myelinogenic potential.
Together, these findings suggest that myelination is a graded process, subject to
competition within the microenvironment and identify a potential novel physiological
role for Nogo-A in the precise myelination of the developing CNS. Maximizing the
myelinogenic potential of oligodendrocytes may offer an effective strategy for repair in
future therapies for demyelination.

 
 
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Introduction
A need for precise coordination underlies the formation of a functional nervous system.
This can be seen in many developmental processes, ranging from the perfect match
between the number of innervating axons and the size of a given target, to the match
between neurotransmitters and their appropriate postsynaptic receptors. Here we focus on
the need for precise coordination in the developmental process of myelination. What
mechanisms coordinate the precise myelination of axons in the nervous system? Or in
other words, what mechanisms help ensure the perfect match between axons and the cells
that myelinate them? In the central nervous system (CNS), myelin is formed by glial cells
known as oligodendrocytes. These terminally differentiated cells arise from
oligodendrocyte precursor cells (OPCs). Myelination of all necessary axonal segments
will be a function of the number of differentiated oligodendrocytes and the number of
myelin internodes that each oligodendrocyte forms. During development, each
differentiated oligodendrocyte is capable of myelinating multiple axonal segments,
ranging from 10 to 50 myelin segments per cell (Baumann & Pham-Dinh 2001). The
periodic placement of these internodes along axons facilitates the process of saltatory
conduction, which allows for the rapid and efficient propagation of action potentials
throughout the nervous system. Amazingly, developing oligodendrocytes somehow
manage to myelinate all necessary axonal segments in a non-overlapping fashion. This
type of accuracy necessitates the presence of extremely precise spatial and temporal cues
that allow for coordination between myelinating oligodendrocytes. These cues are
important to coordinate both the placement of myelin internodes and the generation of
 
 
49 
the appropriate number of myelinating cells and their associated internodes. In order to
achieve the proper spatial alignment, it is essential that no extraneous overlapping
internodes are formed. How is this extremely precise process orchestrated within a
population of oligodendroglial cells?
We predict that this precise spatial alignment is achieved in part through a direct
competition between oligodendrocytes for available axonal space. Our findings suggest
that the competition-based mechanisms that dictate neuronal connectivity paradigms may
be conserved in the establishment of proper CNS myelination. We propose a direct
relationship between oligodendroglial cell numbers and the number of myelin segments
formed per individual oligodendrocyte. This relationship suggests the possibility of a
repulsion-mediated mechanism by which individual oligodendrocytes compete with one
another for available axonal space. The ultimate spatial segregation of myelin internodes
is reminiscent of the phenomenon of dendritic tiling (Gao 2007) and may require a
similar strategy of homotypic repulsion mechanisms to achieve this non-overlapping
placement of myelin internodes along axons. We sought to identify molecular cues that
could influence both the number and length of myelin internodes. Precise regulation of
these features is an essential component of proper spatial segregation. If competitive
interactions between oligodendroglial cells regulate internode formation, it would suggest
that the ultimate capacity to form myelin, referred to here as the myelinogenic potential
of individual cells, is shaped in large part by the nature of the extrinsic environment.
This possibility holds great promise for efforts directed at promoting remyelination,
following the loss of myelin internodes due to nerve injury or disease. The ability of
 
 
50 
oligodendrocytes to form multiple and variable numbers of myelin internodes represents
an intriguing example of intrinsic plasticity that has not yet been extensively explored.
Maximizing the myelinating capacity of individual cells could be important for
therapeutic intervention. Here we will refer to this dynamic and inherent plasticity as the
myelinogenic potential of oligodendrocytes.
Currently, little is known about the nature of the cues responsible for regulating
internode formation and spatial segregation along axons. How might this precisely
coordinated task be achieved? Here we investigate the possibility that the proper
generation and alignment of internodes results from competitive interactions between
oligodendroglial cells. We hypothesize that coordination between oligodendroglia may
actually begin at the level of the precursor cell. Examination of cultured OPCs suggests
that the development of a multipolar morphology may be heightened when cells are
cocultured in the presence of neurons. While somewhat cursory in nature, these
observations suggest the potential presence of axonal cues that promote the outgrowth of
oligodendroglial processes. Static images of oligodendroglial/neuronal cocultures
frequently reveal a network of interdigitated OPC processes with significant overlap
between neighboring cells. In addition, live-imaging studies in developing zebrafish
suggest that OPCs are extremely dynamic cells (Kirby et al 2006). These studies
demonstrate that OPCs continuously extend and retract their processes. In addition, these
live-imaging studies provide evidence that the retraction or realignment of processes can
result from interactions with neighboring cells. Arguably, this type of dynamic
intercellular interaction makes the ultimate alignment and spatial segregation of myelin
 
 
51 
internodes even more impressive. This developmental progression can be seen as
somewhat reminiscent of the eye-specific segregation of initially overlapping retinal
ganglion cell projections seen in visual processing pathways. In these pathways, these
initially overlapping projections are eventually resolved into beautifully patterned ocular
dominance columns (Crowley & Katz 2002; Feller & Scanziani 2005; Katz & Crowley
2002). In a similar fashion, this initially overlapping organization of OPC processes is
eventually resolved into a nicely aligned and spatially segregated pattern of myelin
internodes along axons.
Based on these studies and preliminary observations, it is plausible that
myelinogenic potential will be dependent on three factors, all of which remain relatively
unexplored. Currently, we know very little about the upper limitations to the intrinsic
capacity of oligodendrocytes to form myelin. This includes both the length, the number,
and the thickness of myelin internodes formed. Intriguing studies of sensory neurons in
developing zebrafish suggest that these cells can compensate for the loss of neighboring
cells in vivo by extensive arborization of axonal projections-suggesting an extremely
high intrinsic capacity to form extended projections (Sagasti et al 2005). The
implications of these findings lie in the possibility that the ultimate patterning of these
projections results from extrinsic regulation of this intrinsic capacity. More specifically,
arborization patterns appear to be influenced by competitive interactions between
neighboring cells. Do similar principles apply to the establishment of the morphological
phenotype of myelinating oligodendrocytes? It seems possible that an inherent capacity
to form multiple internodes is likely to be modified by the surrounding environment.
 
 
52 
These extrinsic influences could include both inductive cues, likely expressed by axons,
and inhibitory cues presumably expressed by neighboring oligodendroglial cells. Based
on this hypothetical model, we predict that oligodendrocytes in vivo may exhibit regional
differences in myelinogenic potential. This potential heterogeneity could result both from
differences in axonal cues and from the relative density of oligodendroglial cells in a
given region. Efforts to promote remyelination are hindered in part by the presence of
inhibitory environmental factors (Franklin 2002). Previous work suggests that the
diseased or post-injury milieu is prohibitive to both axon regeneration and the
differentiation of novel oligodendrocytes (Franklin 2002; Yiu & He 2006). It is possible
that demyelination also involves an upregulation of factors that inhibit myelinogenic
potential. Identifying extrinsic regulators of myelinogenic potential could prove
extremely useful for promoting enhanced remyelination following nerve injury or
disease.

 
 
53 
Methods
Immunopanning Protocol. OPCs were purified from P6-7 rat brain cortices with a
panning protocol adapted from one previously described. Petri dishes containing a goat
anti-mouse IgG/IgM secondary antibody solution (Jackson Laboratories) were incubated
overnight. Dishes were rinsed and incubated with primary antibody solutions containing
Ran-2, GalC or A2B5 hybridoma supernatants. Rat brain cerebral hemispheres were first
diced and then dissociated with papain at 37°C. After trituration, cells were resuspended
in a panning buffer and then incubated at room temperature sequentially on three
immunopanning dishes: Ran-2, GalC, and A2B5. A2B5+ OPCs were released from the
final panning dish by using 0.25% trypsin-EDTA (Invitrogen).

Purified OPC/DRG Co-cultures. OPC-DRG co-cultures were prepared as described
previously. DRG neurons from E15 Sprague–Dawley rats were dissociated, plated, and
purified on collagen-coated coverslips in the presence of NGF (AbD Serotec, 100 ng/ml)
in MEM with 10% FBS (Hyclone). Neurons were maintained for 3 weeks before the
addition of OPCs. Coverslips were incubated in a small volume of chemically defined
medium overnight to facilitate OPC attachment. The day after, coverslips were
transferred into wells with chemically defined medium.

Western Blot Analysis. OPCs were grown on poly-L-lysine-coated wells in chemically
defined medium with PDGF (Peprotech, 25 ng/mL) or T3 (Sigma, 30 ng/mL) and CNTF
(Peprotech, 10 ng/mL) for inducing differentiation into oligodendrocytes. Samples were
prepared for Western blot analysis as previously described. The proteins were transferred
to pure nitrocellulose membranes and probed with specific antibodies. Antibodies for
Western blot analysis: rabbit polyclonal anti-PDGFR antibody (Santa Cruz), rat
 
 
54 
monoclonal anti-MBP antibody (Millipore), rabbit polyclonal anti-p75NTR (Promega),
mouse monoclonal anti-MAG antibody (Millipore), goat polyclonal anti-OMgp (R&D
Systems), goat polyclonal anti-Ephrin-B3 (R&D Systems), goat polyclonal anti-Nogo-A
(R&D Systems) and mouse monoclonal anti-Actin (Sigma). The Alexa Fluor donkey
anti-goat, goat anti-rabbit, anti- mouse, and anti-rat 680 IgG antibodies were used as
secondary antibodies for near-infrared fluorescent detection performed on the Odyssey
Infrared Imaging System (LI-COR).

Immunostaining. Immunostaining of tissue sections and co-cultures was performed as
previously described. Co-cultures were fixed by using 4% paraformaldehyde and
dehydrated and then permeabilized and blocked by incubation with 20% goat serum and
0.2% Triton X-100 in PBS. Differentiated oligodendrocytes and myelin were detected
with a rat monoclonal anti-MBP antibody (Chemicon). OPCs were detected by using a
rabbit polyclonal anti-PDGFR antibody (Santa Cruz). Axons were detected by using a
mouse mAb to neurofilament (NF) (American Type Culture Collection). In vivo staining
of oligodendrocytes cell bodies through antibody to APC (Calbiochem) and OPCs was
conducted on 16 μm thick tissue sections. The Alexa Fluor anti-rat, -rabbit, and -mouse
488, 594 and 647 IgG antibodies (Invitrogen) were used as secondary antibodies for
fluorescence detection. Cell nuclei were examined with DAPI.

Bead/OPC/DRG Co-cultures. 20 μm protein A coated beads (G. Kisker) were incubated
overnight with 20 μg of the appropriate Fc (R&D Systems with the exception of
p75NTR-Fc, a generous gift from Regeneron) and washed once in DMEM before seeding
onto the neurons for 1–3 h before the addition of 200,000 OPCs. All Bead/OPC/DRG co-
culture experiments were conducted by using chemically defined medium to avoid the
 
 
55 
potential for IgGs in serum-containing medium to disrupt the binding of Fcs to protein A
on the beads.

Membrane purification. Cells were grown in culture and collected through mechanical
extraction in DPBS. A predetermined portion of the dish was collected separately and
resuspended in RIPA buffer for protein quantification using a BCA assay. Membrane
extract was subjected to 3 freeze-thaw cycles before ultracentrifugation at 540,000 g for 1
h at 4 °C. The pellet was resuspended in chemically defined medium and sonicated in
brief pulses until no longer visible. A normalized portion of this membrane preparation as
standardized through protein quantification is added to NH
2
-group coated 20 μm
polystyrene beads (G. Kisker) and incubated overnight before seeding onto neurons for 1-
3 h before the addition of 200,000 OPCs.

Histology. Animals were anesthetized and perfused transcardially with PBS followed by
4% paraformaldehyde (Electron Microscopy Sciences) in sodium phosphate buffer, pH
7.4 and post-fixed for 2 h before cryoprotection with 30% sucrose in PBS overnight. The
brains and spinal cords were embedded in 1 part O.C.T. compound (Tissue-Tek) with 2
parts 30% sucrose and snap-frozen before sectioning.

In vivo myelin internode quantification. 100 μm coronal and sagittal sections of the
brain and spinal cord were collected and coverslipped with Vectorshield containing DAPI
(Vector Labs) for anatomical identification. 40X z-stack images (z distance = 1 μm) were
acquired using the Zeiss Axio Imager Z1 ApoTome. Myelin internode numbers were
quantified manually, using the cell body as a guide for determining individual
oligodendrocytes.
 
 
56 
In vitro myelin internode quantification. At least 10 20X z-stack images were acquired
using the Zeiss Axio Imager Z1 ApoTome for each condition. Myelin internode numbers
were quantified manually, using the cell body as a guide for determining individual
oligodendrocytes. The percentages of myelinating, non-myelinating oligodendrocytes and
total number of differentiated cells per field were also noted.

RNAi-mediated Nogo-A knockdown. After immunopanning as described, OPCs were
resuspended in rat oligodendrocyte nucleofector solution (Lonza) and transfected with the
appropriate RNAi (Invitrogen, 4 μM) using Amaxa program O-17. They are then seeded
onto poly-L-lysine coated dishes for 3 days before extraction.

 
 
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Results
Assessing Myelinogenic Potential In Vivo
Investigating the myelinogenic potential of oligodendrocytes requires the ability to
examine individual oligodendrocytes in vivo. Effective characterization of this potential
is hampered by the high density of myelinating cells in the CNS. Existing methods only
approximate the number of myelin internodes produced by individual oligodendrocytes
and are limited by sample size and brain region. To investigate the myelinogenic
potential of individual oligodendrocytes, we generated a transgenic mouse with sparsely
labeled oligodendrocytes (0.1-0.5%, in collaboration with Dr. Richard Lu, UT
Southwestern). Efforts to generate transgenic mouse reporter lines have revealed the vast
variability in expression patterns that can result from the use of a single reporter
construct. This effect, known as position effect variegation, has provided an exciting and
unexpected benefit for the generation of sparsely-labeled transgenic mouse reporter lines
(Feng et al 2000). On the basis of this phenomenon, transgenic reporter mice were
generated using a construct expressing GFP under the control of the myelin basic protein
(MBP) enhancer. Use of this construct ensures the restriction of GFP expression to
myelinating cells. Using this strategy, a mouse line was successfully generated with
approximately 0.1% of oligodendrocytes expressing GFP (Fig 3.1B). In contrast to
transgenic lines in which all oligodendrocytes are labeled, (Fig 3.1A) this sparsely
labeled mouse line allows for the successful investigation and characterization of
individual oligodendrocytes and their associated myelin internodes (Fig 3.1C-G). Using
this mouse line, we have systematically reconstructed single oligodendrocytes from
 
 
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multiple regions within the CNS, including the cerebral cortex, corpus callosum and
cerebellum. To our knowledge, this approach is one of the first examples of quantitative
analysis of individual oligodendrocytes in vivo.
 
 
59 

Figure 3.1: Oligodendrocytes exhibit striking diversity in the
number and length of myelin internodes in vivo. (A) Representative
section from the brain of a CNP-GFP transgenic mouse. GFP is
expressed only in mature oligodendrocytes and their associated myelin
internodes. (B) Representative section of a brain from a transgenic
mouse expressing GFP in 0.1-0.5% of oligodendrocytes. GFP
expression is driven by the MBP enhancer. (C) Individual
oligodendrocytes from different brain regions of the transgenic mouse
with sparsely labelled oligodendrocytes. Arrows point to cell bodies.
Scalebar = 20 μm.
 
 
 
60 
The Heterogeneity of Myelinogenic Potential In Vivo
Using our newly developed transgenic mouse line, we have systematically reconstructed
single oligodendrocytes from multiple regions within the CNS, including the cerebral
cortex, corpus callosum and cerebellum (Fig 3.1C). Quantification of myelin internodes
suggests that oligodendrocytes can form between 15 and 60 myelin segments per
individual oligodendrocyte (Fig 3.1C and Fig 3.2). We also find remarkable variation in
the length of myelin segments, with internodes ranging from 100-400 um in length (not
shown). Importantly, this marked variability does not appear to be region-specific.
Instead, we find that variations in both length and internode number occur not only
between brain regions, but also within the same local brain region and along the same
axonal tracts (Fig 3.1C-G). This intriguing finding suggests that the myelinogenic
potential of individual oligodendrocytes is not determined by their gross spatial
localization within the CNS. Indeed, the variation found within brain regions suggests
that localized microenvironmental interactions may regulate myelinogenic potential.
Variation within a given axonal tract further suggests that neither axonal density nor
molecular axonal determinants are the sole factors influencing the number of myelin
internodes formed. We propose that such localized variation results in part from contact-
mediated interactions between neighboring oligodendrocytes. We hypothesize that local
repulsive interactions between oligodendroglial cells play a pivotal role in shaping the
number of myelin internodes that each oligodendrocyte will ultimately form. We next
turned to an in vitro coculture system to test the validity of this hypothesis.
 
 
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Dynamic Variations in Myelinogenic Potential Exist In Vitro
Our coculture system is a reduced system that represents an ideal strategy to examine the
role of neuronal-glial interactions in the regulation of myelinogenic potential. The
coculture system consists of purified embryonic sensory dorsal root ganglion neurons
(DRGs) and purified OPCs. Initially, DRGs are purified and maintained in culture for
approximately three weeks, at which point axon density is highly uniform throughout the
coverslip. Our cocultures are therefore characterized by minimal variability in both
Figure 3.2: Variations in myelinogenic
potential exist both within and between
brain regions. Quantification of the
number of myelin internodes formed per
oligodendrocyte for each region. Error bars
represent range.
 
 
62 
axonal density and the molecular identity of cultured neurons. Despite this highly
homogeneous environment, our coculture system exhibits dynamic variability in the
myelinogenic potential of individual oligodendrocytes (Fig 3.3B). This variability is
comparable to the variation we see in vivo (Fig 3.3A). Together, these observations
underscore the likelihood that interactions between oligodendrocytes are responsible for
helping to shape the number of myelin segments formed by individual cells.
 
 
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Figure 3.3: Membrane-bound inhibitory cues expressed by oligodendroglial decrease the
number of myelin internodes formed per oligodendrocyte. (A) 2D compilations of Z-stacks of
individual oligodendrocytes from different brain regions of the sparsely labelled transgenic
mouse exhibit marked variability in both the number and length of myelin internodes. (B)
Oligodendrocytes from OPC-DRG co-cultures exhibit a broad range of internode dimensions that
are comparable to cells found in vivo. Oligodendrocytes are visualized by immunostaining with
an antibody against MBP. Arrows point to cell bodies. Scale bar = 20 μm. (C) In OPC-DRG co-
cultures seeded with a high density of OPCs (left panel), oligodendrocytes form fewer myelin
internodes per cell (right panel) as compared to (D) co-cultures seeded with a low density of
OPCs with polystyrene beads (left panel) in which the number of myelin internodes formed per
oligodendrocyte is increased (right panel). OPCs are visualized by immunostaining for PDGFRα.
Axons are labelled with an antibody against neurofilament (NF). Scale bar = 50 μm. (E-G) High
magnification images of individual oligodendrocytes from (E) high density OPC co-cultures, (F)
low density OPC co-cultures + beads, and (G) low density OPC co-cultures with OPC-membrane
coated beads. Arrows point to cell bodies. Scale bar = 20 μm. (H) To test for the presence of
inhibitory cues expressed on oligodendroglial membranes, OPC membranes were subsequently
extracted and coated onto NH
2
-coated positively charged polystyrene beads prior to being seeded
onto co-cultures. (I) Quantification of the effect of various types of cell membranes on myelin
internode numbers. Only oligodendroglial membranes are sufficient to decrease the number of
internodes in a manner comparable to high density OPC co-cultures. Error bars represent standard
deviation. (p<0.01, Tukey post hoc comparison after one-way ANOVA).
 
 
64 

Figure 3.3: Continued

 
 
65 
Density of Oligodendroglial Modulates Myelinogenic Potential
If interactions between oligodendroglial cells are involved in the regulation of
myelinogenic potential, then the density at which OPCs are seeded onto neurons in vitro
could have a direct impact on the number of myelin segments formed by individual
oligodendrocytes. We sought to test the hypothesis that OPCs and/or oligodendrocytes
express a repulsive cue mediating the collapse of nearby processes and ultimately
dictating the myelinogenic potential of neighboring oligodendrocytes. If this hypothesis
is correct, then an increase in the density of either OPCs or oligodendrocytes should
provide a corresponding increase in the expression of a repulsive cue(s). Therefore, by
increasing the density of OPCs or oligodendrocytes, we would expect to see an overall
decrease in the average number of myelin internodes formed per oligodendrocyte. Our
identification of the density-dependent nature of differentiation in vitro (Rosenberg et al
2008) provides us with a unique opportunity to test this possibility. The use of
polystyrene beads conjugated with antibodies against axonal proteins allows us to create
spatial constraints along axons that are sufficient to induce differentiation in the absence
of a critical density of OPCs. This phenomenon allows us to compare the average
myelinogenic potential of cells in cultures that have reached a critical density versus
cultures with a low density of OPCs in the presence of polystyrene beads. We find, as
hypothesized, that an increase in OPC density leads to a corresponding decrease in the
average number of myelin segments formed per cell (Fig 3.3C-F, and I). This finding is
consistent with the possibility that repulsive interactions between OPCs or
oligodendrocytes mediate myelinogenic potential.
 
 
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Membrane-bound Oligodendroglial Factors Regulate Myelinogenic Potential
We next sought to identify cues that could mediate repulsive interactions between
oligodendroglial cells. The expression of a membrane-bound cue responsible for either
inhibitory or repulsive signaling would require contact-mediated intercellular
interactions, allowing for highly localized regulation of myelin internode numbers. To
confirm that a membrane-bound factor is sufficient to modulate myelinogenic potential,
we coated custom designed polystyrene beads with OPC and/or oligodendrocyte
membranes (Rosenberg et al 2008). Using this approach, we can essentially create
artificial cells and examine whether membrane-tethered molecules are responsible for the
decrease in the number of myelin segments that occurs in the presence of a high density
of oligodendroglial cells. We find that robust inhibitory signals on the oligodendroglial
membranes diminish the number and length of myelin segments (Fig 3.3G-I), whereas
control membranes from other cell types, such as astrocytes, do not alter myelinogenic
potential. These findings suggest that a membrane-bound cue(s) expressed by
oligodendroglial is sufficient to regulate myelinogenic potential. It is plausible that
remodeling of oligodendroglial processes occurs continuously at both the OPC and the
oligodendrocyte stages of development, prior to the determination of the final number of
myelin internodes that each cell will form.

 
 
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Regulation of Myelinogenic Potential by Nogo-A
To identify specific membrane-bound oligodendroglial factors capable of regulating
myelinogenic potential, we performed a candidate screen to test factors previously
identified as repulsive guidance cues or regulators of axon outgrowth. We reasoned that
when expressed on developing oligodendroglial cells, these cues may act to regulate the
number of myelin internodes through an inhibitory mechanism reminiscent of either
repulsive guidance or growth cone collapse. We propose that a correlation exists between
the number of stabilized oligodendroglial processes and the ultimate number of myelin
segments formed. To determine the validity of this hypothesis, it is necessary to identify
the expression of repulsive inhibitory cues that could mediate the collapse of
oligodendroglial processes. Our candidate screen included molecules previously
identified as myelin-based inhibitors of axon outgrowth such as MAG, Nogo-A, Nogo-
66, p75NTR, Ephrin-B3 and OMgp (Yiu & He 2006). Initially we verified the expression
of these inhibitory cues on cultured OPCs and oligodendroyctes (Fig 3.4A). To test the
effect of these candidates on myelinogenic potential, we conjugated soluble Fc-linked
candidate proteins to Protein A-coated polystyrene beads. With this strategy it is possible
both to test for sufficiency, and to mimic the physiological effect of varying cell density
by presenting candidate molecules locally to myelinating cells. The majority of factors
screened individually were not sufficient to modulate myelinogenic potential (Fig 3.4B-
C). However, the amino-terminal of Nogo-A is sufficient to significantly inhibit both the
number and length of myelin segments formed by individual oligodendrocytes (Fig 3.3B-
C).
 
 
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The results of the candidate screen suggest that the exogenous expression of Nogo-A is
sufficient to inhibit the number of myelin internodes formed by individual
oligodendrocytes. We next sought to examine whether the expression of Nogo-A in
oligodendroglial cells is necessary to inhibit myelin internode formation. To test this
hypothesis, we used siRNA to knock-down Nogo-A expression in cultured OPCs and
Figure 3.4 Nogo-A is necessary and sufficient for the regulation of myelinogenic
potential. (A) Western blot probing for expression of candidate molecules on OPCs,
oligodendrocytes and control cells. (B) Candidate molecule Fc- fusion proteins were
conjugated to Protein A coated polystyrene beads. Beads were seeded onto DRGs and
co-cultures were maintained for 10 DIV. Arrows indicate cell bodies. Scale bar = 50
μm. (C) Quantification of the effect of candidate molecules on myelinogenic
potential. Only Nogo-A/Fc significantly decreases the number of myelin internodes
formed per oligodendrocyte across all conditions (p<0.01, Tukey post hoc
comparison after one-way ANOVA). Error bars represent standard deviation. (D-E)
Knockdown of Nogo-A in OPC membranes prior to adsorption onto polystyrene
beads leads to an increase the number of myelin internodes formed per cell. OPCs
were transfected with either a control siRNA (D, left panel) or siRNA targeting the
expression of Nogo-A (D, right panel). (E) Quantification of internode formation per
oligodendrocyte following treatment of OPC membranes with siRNA.
 
 
 
69 
oligodendrocytes prior to membrane extraction (Grimes et al 1996; Yamauchi et al 2004).
Membranes were then coated around beads and presented to myelinating cells as
described previously. We find that a reduction in Nogo-A expression leads to an increase
in the myelinogenic potential of individual oligodendrocytes. While these results are
preliminary and experiments are currently ongoing, these findings suggest that Nogo-A is
necessary for the effect of oligodendroglial membranes on myelinogenic potential (Fig
3.4D-E).
Together our findings suggest that Nogo-A is both necessary and sufficient to
regulate myelinogenic potential in vitro. However, Nogo-A alone does not affect
internode formation with the same potency as the oligodendroglial membranes,
suggesting that other membrane-bound cues are also likely to be involved in the
regulation of myelinogenic potential (Table 1).
Table 1. Effects of membrane-coated and Fc-coated beads on oligodendrocyte
differentiation and myelination.
Membrane expt
Low
density
High
density
OPC
mem
Oligo
mem
COS
mem
3T3
mem
Astro
mem
% Myelinating 67.6 ± 0.5 90.9 ± 1.5 54.9 ± 0.3 50.0 ± 2.2 87.7 ± 1.9 84.8 ± 1.5 75.7 ± 0.9
% Non-
myelinating
32.4 ± 0.5 9.1 ± 0.3 45.1 ± 0.5 50.0 ± 4.6 12.3 ± 0.4 15.2 ± 0.5 24.3 ± 0.9
# Oligos/field 12 ± 5 10 ± 3 11 ± 5 12 ± 6 10 ± 4 9 ± 2 9 ± 2
Fc expt
p75NTR
Fc
MAG
Fc
OMgp Ephrin-B3
Fc
Nogo-A
Fc
Nogo-66
Fc
% Myelinating 77.8 ± 0.3 78.6 ± 0.7 76.7 ± 1.3 87.5 ± 4.6 77.1 ± 0.6 80.1 ± 0.6
% Non-
myelinating
22.2 ± 0.2 21.4 ± 0.4 23.3 ± 0.7 12.5 ± 2.0 22.9 ± 0.3 19.9 ± 0.4
# Oligos/field 10 ± 4 11 ± 6 9 ± 4 17 ± 7 12 ± 5 11 ± 3

OPC and oligodendrocyte membrane coated beads significantly increase the percentage of non-
myelinating oligodendrocytes present within the OPC-DRG-bead co-culture across all conditions.
Abbreviations: mem = membrane, oligo = oligodendrocyte, astro = astrocyte.
 
 
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Regulation of Myelinogenic Potential In Vivo
Based on our in vitro experiments, we are eager to examine the role of Nogo-A in
mediating myelinogenic potential in vivo. Currently we are breeding our mouse line
expressing membrane GFP in 0.1-0.5% of oligodendrocytes with existing Nogo knockout
mice (Lee et al 2009). By reconstructing and quantifying the number of myelin segments
in single oligodendrocytes, this strategy will allow us to initially examine the role of
Nogo-A in the formation of myelin segments in vivo. If Nogo-A is involved in regulating
competitive interactions between oligodendrocytes and inducing collapse of growth cone-
like processes, then in mice that globally lack these molecules we should see an overall
increase in the average number of myelin internodes formed per individual
oligodendrocyte. Intriguing preliminary data suggests that in regions of the CNS such as
the corpus callosum and layer 1 of the somatomotor cortex, the extent of myelination is
increased in Nogo knock-out mice as compared to littermate controls (Fig 3.5).
Preliminary findings suggest this increase in myelination is not due to an increase in
oligodendrocyte numbers (Fig 3.5C-F and Table 2). It will be exciting to confirm whether
this increase in myelination results instead from an increase in the myelinogenic potential
of individual oligodendrocytes. Ultimately, the in vivo approach described here could be
applied to test any additional membrane-bound factors that are shown to have an effect
on myelinogenic potential in vitro. We believe that this strategy will help us to elucidate
the mechanisms that govern intercellular interactions and the generation of myelin
segments by oligodendrocytes.
 
 
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Figure 3.5 Increased myelination in Nogo knock-out mice
(A and B) The extent of myelination in layer 1 of the somatomotor cortex is increased in Nogo
knock-out mice (B) as compared to wild-type littermate controls (A). Lower panel is a high
magnification image of the boxed area in the top panel. Myelin is visualized by immunostaining
for MBP (green). (C-D) While the extent of myelination is increased in the somatomotor cortex
(D left and right panels) and the corpus callosum (F left and right panels) of Nogo knock-out
mice as compared to wild-type littermate controls (C and E left and right panel), the number of
OPCs and oligodendrocytes is not significantly different (C-F middle and right panels) between
Nogo knock-out mice and wild-type littermate controls. Myelin is visualized by immunostaining
for MBP (green). Immunostaining for PDGFRα labels OPCs (red) and immunostaining for CC1
labels oligodendrocytes (blue).
 
 
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Figure 3.5: Continued

Cortex Corpus callosum
Cells/mm
2
WT Nogo KO WT Nogo KO
OPCs/PDGFRα+ 326 ± 22 303 ± 28 673 ± 71 731 ± 132
Oligos/CC1+ 544 ± 77 510 ± 53 1592 ± 366 1736 ± 425
Table 2. Nogo deletion does not significantly affect OPC and
oligodendrocyte numbers.
 
The number of PDGFRα+ OPCs and CC1+ oligodendrocytes is not
significantly different in the somatomotor cortex or the corpus callosum
of Nogo knock-out mice as compared to wild-type littermate controls.
 
 
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Discussion
In the studies described here, we present one of the first demonstrations of the remarkable
heterogeneity that exists in both the length and number of myelin internodes associated
with individual myelinating oligodendrocytes in vivo. The development of a novel
transgenic mouse with GFP expressed in a sparse population of myelinating cell provides
a unique opportunity to begin to examine the factors that regulate myelinogenic potential.
Here we find that factors expressed on oligodendroglial membranes are sufficient to
regulate internode formation in vitro. Specifically we find that Nogo-A appears to be
both necessary and sufficient to regulate the number of internodes formed per individual
cell. These findings are the first to suggest a physiological role for Nogo in regulating
myelination during development. The role of Nogo in regulating axon regeneration and
outgrowth has been well-described (He & Koprivica 2004). Additional studies suggest
that Nogo may be important for the regulating closure of the critical period, particularly
in the visual cortex (McGee et al 2005). However, our findings are the first to suggest
that Nogo may function during development to regulate internode formation through its
role in mediating intercellular interactions between neighboring oligodendrocytes. These
results support the hypothesis that competition for axonal space between neighboring
oligodendrocytes contributes to the precise myelination of the CNS.
Efforts to understand developmental myelination may provide new insight and
ideas for novel strategies to promote remyelination. Current research efforts have focused
primarily on addressing the initiation, rather than the modulation, of myelination. Our
application aims to shift the focus solely from those factors that initiate myelination, to
 
 
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also include those factors that can enhance the number of myelin internodes formed-
specifically in the presence of a demyelinating environment.
Regulation of myelination at the level of internode numbers and lengths
represents a novel conceptual approach to examining myelination. Currently, it remains
unclear what factors are responsible for regulating the number of myelin segments
formed by individual oligodendrocytes. Here we sought to use novel strategies to
investigate and identify factors regulating myelinogenic potential. We believe this
objective could contribute to remyelination strategies in two ways. First, we propose that
enhancing the myelinogenic potential of existing oligodendrocytes could represent a
novel approach for remyelination and repair. In conditions such as MS, demyelinating
lesions are localized and do not necessarily affect all proximal oligodendrocytes
(Franklin 2002; Franklin & Ffrench-Constant 2008). Is it possible that surviving
oligodendrocytes could be induced to form additional myelin internodes along
demyelinated axonal segments? While a demyelinated environment is not conducive to
ongoing differentiation, this environment may be less inhibitory to the actual formation
of myelin internodes. It is possible that promoting differentiated oligodendrocytes to form
additional myelin segments could represent a novel and attractive alternative strategy for
promoting remyelination. By blocking the expression or activity of factors that inhibit
internode formation, it may be possible to induce healthy neighboring oligodendrocytes
to extend additional myelin segments and effectively replace lost oligodendrocytes. Is
this strategy feasible or realistic? Currently, it remains unclear whether myelination by a
single oligodendrocyte happens all at once, or is instead a continuous process that can
 
 
75 
occur over time. While some studies suggest that myelination represents a single
temporally coordinated event, this finding requires further investigation and relevance to
an in vivo setting remains unaddressed (Watkins et al 2008). The advent of advanced
live-imaging studies will no doubt prove essential to further examination of this
fundamental characteristic of myelination.
It is exciting to speculate that differentiated oligodendrocytes might be induced to
form additional myelin internodes to compensate for oligodendrocytes lost to nerve
injury or disease. However, our investigations could be relevant to remyelination, even if
differentiated oligodendrocytes do not retain the capacity to generate additional myelin
segments. Harnessing the myelinogenic potential of newly differentiatied adult OPCs
could also prove effective in promoting remyelination. In other words, if adult OPCs can
be induced to differentiate and form higher numbers of myelin segments, then a smaller
total number of newly differentiated cells would be required to facilitate recovery and
repair. A reduced need for newly differentiated cells would be extremely beneficial in
light of the apparent decrease in differentiation capacity that accompanies ongoing
demyelination (Franklin 2002; Franklin & Ffrench-Constant 2008). For all of these
reasons, we believe that examining the mechanisms regulating myelinogenic potential of
oligodendrocytes is extremely pertinent to efforts aimed at promoting remyelination.

 
 
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CHAPTER 4
Looking Ahead

A journey of a thousand miles begins with a single step. ~Lao-Tzu

When I look back on my graduate career, I will always remember this as a time when I
first truly fell in love with being in science. Witnessing the beauty of a myelinating cell
is a profound and unique experience that I will always treasure. Jonah was the first one
to teach me that serendipity is a word that belongs in research, and I consider myself
extremely fortunate to have experienced firsthand the truth of this lesson. I have truly
enjoyed the projects I have worked on and the serendipitous discoveries in which I have
had the chance to participate. Some of the greatest joy associated with these projects
comes from the hope and belief that the work we have done has potential implications for
exciting future directions.
Our initial studies on oligodendrocyte differentiation took an unexpected and
exciting turn when they lead to the discovery that spatial constraints along axons are
sufficient to induce differentiation, even in the absence of a critical density of cells.
These findings suggest that biomechanical interactions could play a role in regulating the
differentiation process. It is extremely intriguing to consider the potential downstream
signaling events responsible for this effect, as well as the potential relevance of this
finding to differentiation in vivo. One potential explanation for our results is that
 
 
77 
biomechanical intercellular interactions initiate differentiation by inducing changes in the
shape of OPCs.

Density Dictates Shape
A significant factor influencing cell shape is the interaction of a cell with its
environment- both the substrate to which it adhers and the cells that surround it on all
sides. It is possible that the density of cells can indirectly regulate cell shape by
influencing the extent of interaction between a precursor cell and its substrate. To address
this possibility, it is first important to understand the manner by which a substrate can
influence cell shape and cell fate. Some insight into this relationship can be gained from
an intriguing recent study by Engler et al. This group examined the effect of matrix
stiffness on stem cell shape and fate (Engler et al 2006). The stiffness of a matrix can be
measured by the elastic modulus, E, which indicates the elasticity- or potential of the
substrate to be deformed. This elegant study used varying concentrations of bis-
acrylamide to alter the elasticity of collagen-coated gel culture substrates. The authors
show that changing the elasticity of the substrate correlates with a change in cell shape.
Cells cultured on softer gels exhibit a more branched neuron-like morphology whereas
cells cultured on stiffer substrates adopt a spindle-shaped morphology with minimal
branching- reminiscent of muscle cells. Finally, mesenchymal stem cells (MSCs) cultured
on extremely stiff substrates adopt a polygonal shape that approximates that of
osteoblasts (Engler et al 2006). These studies suggest that the shape of a cell can be
modulated by the stiffness and deformation capacity of the substrate. Interestingly, these
 
 
78 
findings also suggest that the stiffness of the culture matrix can act as a critical
determinant in the ultimate fate adopted by MSCs (Engler et al 2006). How do changes in
substrate stiffness affect the shape of the cell? An increase in matrix stiffness is sufficient
to increase the formation of focal adhesions (Engler et al 2006), which facilitate the
attachment of cells to their substrate (Bershadsky et al 2006). Multiple studies have
shown a correlation between focal adhesion formation and cell shape (Boudreau & Jones
1999; Chen et al 2003). For example, cells lacking focal adhesion proteins exhibit
abnormal cell spreading (Chen et al 2003). Additionally, focal adhesions mediated by
interactions between integrins and ECM substrate proteins can promote cytoskeletal
reorganization, thereby facilitating alterations in cell shape (Boudreau & Jones 1999). It
is important to note that integrin-mediated signaling has also been implicated in cell fate
decisions. As evidenced by the findings listed above, the shape of a cell can be influenced
by both the nature of the substrate, and by the level of interaction between the substrate
and the cell. Intuitively, I would expect that an increase in OPC cell density within a
given area ultimately requires compression of the cells as a result of spatial constraints.
As a result, individual cells must compensate for a decrease in available area by
becoming increasingly constrained and minimizing the extent of their interactions with
the substrate.
Additionally, an increase in cell density can modulate cell shape through more
direct means as well. Physical interactions between cells can directly modulate cell shape
to optimize cellular packing. A study performed using retinal epithelial cells as a model
suggests that interacting cells tend to cluster together in formations reminiscent of soap
 
 
79 
bubble associations (Hayashi & Carthew 2004). The results of this study suggest that
cells tend to interact in a manner that reduces the overall surface area of the group of
cells. Compression from all sides, resulting from a high density of neighboring cells,
could effectively inhibit cell spreading and adhesion, as well as compacting the cells into
polygonal or hexagonal shapes closing approximating a spherical appearance. Consistent
with this theory are studies that induce a similar manner of constraint by culturing cells
on various sizes of micropatterned islands coated with ECM proteins. These studies
demonstrate that cells cultured on smaller islands adopt a round, spherical cell shape,
whereas those cultured on larger islands exhibit a more flattened and elongated
phenotype (Chen et al 2003; McBeath et al 2004). I would hypothesize that the
constriction experienced by cells on small islands is comparable to the constriction that
cells would undergo at a high density. Together, these studies support the idea that cell
density can influence cell shape, which may ultimately result in changes in cell fate.

What pathways are downstream of cell shape in regulating cell fate?
Once a role for cell shape in regulating cell fate has been established, it will be
important to determine precisely how the shape of a cell can influence cell fate decisions.
What are some of the signaling pathways downstream of cell shape that could influence
cell fate? One mechanism that has been shown to link cell shape and cell fate is the
signaling pathway mediated by the Rho GTPase, RhoA (McBeath et al 2004). As
described earlier, elegant studies employing micropatterned islands of ECM substrate
were used to demonstrate that manipulating cell shape is sufficient to regulate cell fate.
 
 
80 
This same technique- in which individual cells were plated on islands of various sizes,
was used to examine the role of RhoA signaling in cell fate determination. These authors
found that the activity of Rho Kinase (ROCK), a downstream effector of RhoA, is greater
in spread cells as compared to round cells. This is consistent with additional data from
this group demonstrating that RhoA activity is high in low density cultures, which are
comprised of spread cells, and low in high density cultures, which contain predominantly
round cells. Together these results suggest that RhoA activity can be correlated with cell
shape, and that spread cells exhibit increased RhoA signaling as compared to round cells.
While these findings do not show that RhoA signaling is directly regulated by cell shape,
they do demonstrate a correlation between changes in cell shape and changes in Rho
signaling. To test whether RhoA signaling is downstream of the effect of cell shape, the
authors expressed a dominant-negative (DN) RhoA construct in spread cells and an
constitutively active (CA) RhoA construct in round cells. In these studies, RhoA
signaling was not sufficient to override the effects of cell shape. However, expression of
a CA-ROCK construct was sufficient to induce a similar fate in both round and spread
cells. These findings suggest that ROCK-mediated Rho signaling has the potential to
influence cell fate and may be downstream of cell shape. This example raises two
intriguing questions about cell shape signaling. First, how could changing cell shape alter
Rho-ROCK signaling? And two, how might Rho-ROCK-mediated signaling regulate
changes in gene transcription?
A recent study examining cell shape changes provides a fascinating and beautiful
explanation for the question of how cell shape can elicit general changes in signaling
 
 
81 
pathways. The study by Meyers et al. provides a model for the activation of signaling
cascades based on the physical distance between positive and negative regulators of the
pathway (Meyers et al 2006). For instance, in the case of Rho signaling, the activity of
Rho could be dependent on the relative locations of factors activating Rho signaling, such
as guanine nucleotide exchange factors (GEFs) and factors inhibiting Rho activity such as
GTPase-activating proteins (GAPs) (Jaffe & Hall 2005). If we make the assumption that
activating factors are primarily localized to the membrane and inhibitory factors are
found in the cytoplasm, then we can postulate that the greater the cytoplasmic distance
between an active signaling molecule and its downstream effector, the greater the
likelihood that the molecule will be deactivated before reaching its downstream target
(Meyers et al 2006). This theory is quite intriguing in its own right, but how does this
relate to the effect of cell shape on RhoA signaling? First of all, the above mentioned
assumption is consistent with research demonstrating that post-translational modifications
target Rho GTPases to the plasma membrane, facilitating the interaction between Rho
GTPases and activating GEFs (Schmidt & Hall 2002). Next, the model generated by this
group suggests that cell shape-dependent variability could affect RhoA activity and
subsequent interactions with downstream targets. This variability could occur as a result
of shape-dependent changes in the distance between the plasma membrane and
downstream cytoplasmic targets. Based on this model, one would predict that flat cells
are more likely to have increased activation of signaling proteins because in a flat spread
cell, the distance between the plasma membrane and the majority of the cytoplasm is
greatly reduced as compared to round cells where a greater proportion of the cytoplasm
 
 
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is located more distal to the plasma membrane. This prediction is consistent with the
results of McBeath et al., which demonstrate increased Rho-ROCK activity in flat cells as
compared to round cells. (McBeath et al 2004). This exciting correlation presents the
intriguing possibility that cell shape may regulate cell fate by influencing the relative
physical proximity of intracellular factors. In fact, it has previously been suggested that
changes in cell shape may facilitate structural rearrangements within the cell (Ingber
1997). This intracellular reorganization could initiate interactions between upstream
mediators and downstream effectors that are responsible for the induction of
differentiation (Boudreau & Jones 1999). Additionally, it has been demonstrated that
changes in cell shape can be directly linked to structural changes in the nucleus (Maniotis
et al 1997). This possibility is intriguing because it provides an additional explanation for
the effect of cell shape on transcriptional programs affecting cell fate. For example, it has
been postulated that integrins may mediate changes in nuclear structure that facilitate
chromatin remodeling and the access of transcription factors to their respective targets
(Boudreau & Jones 1999). In fact, it has been shown that the mechanical manipulation of
integrins is sufficient to induce changes in nuclear structure. In this intriguing study,
micropipettes were coated with ECM-ligands such as fibronectin, allowing for
interactions with integrin-receptors on the surface of endothelial cells (Maniotis et al
1997). The micropipettes were then pulled in different directions to induce changes in the
shape of the cell. Manipulation of the cell through integrin-mediated interactions was
sufficient to induce changes not only in the structure of the cell, but also in nuclear
structure, as demonstrated by phase imaging. Importantly, in cells treated with
 
 
83 
cytochalasin D, micropipette manipulation was sufficient to deform the cell surface but
failed to induce the corresponding changes in nuclear structure. These results suggest that
actin-mediated interactions are necessary to mediate the effects of integrin-surface
receptor orientation on nuclear structure. These studies support the hypothesis that the
nucleus is physically coupled to the cell surface through integrin-actin-based interactions.
These findings lend support to the hypothesis that changes in cell shape can promote
physical changes in the nucleus that influence cell fate.
As mentioned earlier, Rho-ROCK signaling has also been implicated as a putative
mediator of cell shape-dependent cell fate. RhoA signaling has the potential to affect cell
fate decisions by modulating the activity of the transcription factor, Serum Response
Factor (SRF). SRF-dependent transcription requires the activity of a cofactor known as
MAL, which is translocated to the nucleus in response to Rho activation (Miralles et al
2003). Interestingly, this shift in MAL localization is dependent on the release of MAL
from G-actin, which occurs following RhoA-dependent actin assembly (Vartiainen et al
2007). Together, these findings suggest that ROCK could mediate cell fate decisions and
changes in transcription indirectly by mediating changes in actin assembly that allow for
the nuclear transport of MAL. In addition, Rho signaling can also regulate gene
transcription through signaling pathways that are not dependent on cytoskeletal
reorganization. Rho signaling is capable of regulating JNK and p38 MAP kinase
pathways, both of which regulate the activity of multiple transcription factors (Mackay &
Hall 1998). In addition, Rho has also been shown to activate the transcription factor NF-
kappa B (Perona et al 1997). Together these studies suggest multiple mechanisms by
 
 
84 
which a single signaling pathway could mediate the effects of cell shape on cell fate. It is
likely that there are many other mechanisms besides RhoA that could also provide this
link between morphological rearrangements and changes in transcriptional programs
leading to differentiation. Studies demonstrating the effect of cell shape on differentiation
highlight the fact that in addition to biochemical signaling pathways, biophysical
mechanisms can also play an important role in regulating cell fate. Indeed, considering
the morphological contortions that occur in a developing organism, it is not surprisingly
that physically-mediated interactions could influence cell development and fate (Montell
2008). It will be exciting to see if future studies can elucidate a mechanistic role for
biomechanical interactions and cell shape changes in the regulation of oligodendrocyte
differentiation.

Implications of Competitive Signaling and Myelinogenic Potential
Our studies of oligodendrocyte differentiation were exciting in their own right, but they
also played a pivotal and serendipitous role in the evolution of our second project
involving the investigation of myelinogenic potential. Both the observation of variations
in myelinogenic potential in vitro and the techniques used to investigate and manipulate
internode formation would not have been possible without the initial differentiation
studies and the unexpected results.
The studies presented here will hopefully provide significant insight into the
regulation of the myelinogenic potential of individual oligodendrocytes. Currently, this
question remains largely unaddressed, due in part to experimental limitations. We
 
 
85 
believe that our in vivo and in vitro systems provide us with a unique opportunity to begin
to characterize and manipulate the myelinogenic potential of individual cells. Our
preliminary studies suggest that variations in the number of myelin segments formed per
cell may be due largely to localized interactions between neighboring oligodendrocytes.
Defining the mechanistic basis for these interactions could help explain how the
developing nervous system perfectly myelinates all necessary axonal segments. Shedding
light on these developmental mechanisms could prove extremely valuable for harnessing
the myelinogenic potential of oligodendrocytes in promoting optimal recovery in a
remyelination setting. Additionally, the studies proposed here may offer new insight into
general principles that govern arborization and pruning mechanisms in the developing
nervous system. The refinement of axonal projections is characterized by a selective
elimination of individual branches, without activation of apoptotic mechanisms that
would result in the programmed death of the entire cell (Luo & O'Leary 2005). In a
similar fashion, variations in myelinogenic potential suggest that oligodendrocytes can
also undergo a selective loss of individual branches without affecting the survival of the
cell. In light of this similarity, it is possible that common and conserved mechanisms may
regulate competition for targets and subsequent pruning of extraneous branches in both
neuronal and glial cells. It is our hope that these studies will shed light into these
potentially redundant mechanisms.
It is also exciting to hope that future studies will delineate the mechanistic basis
for these competitive intracellular interactions. For example, if repulsive or inhibitory
factors such as Nogo-A are expressed on all oligodendroglial processes, then what
 
 
86 
mechanisms ensure that one process will be eliminated while another process is
maintained? In other words, what mechanisms are in place to give certain processes a
selective advantage? These questions are essential to address in order to really understand
how competitive interactions between oligodendroglial cells could ultimately shape the
myelinogenic potential of individual cells. In the case of Nogo-A, a mechanistic
understanding will no doubt require the identification of a receptor capable of mediating
downstream effects, which presumably lead to cytoskeletal rearrangement and either
collapse or turning of the extended process. Intriguingly, the receptor responsible for
mediating the effect of the amino-terminal of Nogo-A on axonal outgrowth remains
uncharacterized. This is in contrast to Nogo-66, a 66 amino acid portion of the Nogo
protein which has been shown to signal through the receptor complex comprised of
NogoR, Taj/Troy, Lingo-1, and p75 (McGee & Strittmatter 2003). Identification of an
oligodendroglial-expressed Nogo-A receptor will be informative for determined the
downstream signaling events initiated by Nogo-A. Repulsive signaling mediated through
ligand-receptor complexes such as Ephrin-EphRs exhibit the potential for bidirectional
signaling (Kullander & Klein 2002). This represents an attractive strategy by which a
single ligand-receptor pairing could simultaneously mediate the stabilization of one
process and the retraction or realignment of an extended oligodendroglial process from a
neighboring cell. In addition, it will be interesting to examine the possible existence of
feedback loops that provide one cell with a competitive edge over a neighboring cell.
Evidence of this type of competition-based feedback signaling can be seen in the case of
competition between sensory neurons for target-derived survival factors (Deppmann et
 
 
87 
al 2008). Alternatively, it is possible, but perhaps less likely, that factors such as Nogo-A
are differentially expressed within the oligodendroglial population, a phenomenon that
could also provide for a competitive edge between neighboring cells. For this reason, it
will be important to examine the spatial expression pattern of Nogo-A and other
candidate factors in vivo.
A parallel future direction will be the identification of factors that allow not only
for interactions between neighboring oligodendroglia, but also for spatial segregation of
processes from a single cell. This is again reminiscent of dendritic tiling, in which
signaling pathways exist to establish both self-identification and avoidance (Parrish et al
2007). The non-overlapping alignment of myelin internodes along axons suggests that
similar types of mechanisms are likely to exist in oligodendroglial cells.
Additional follow-up studies will hopefully identify additional candidate
molecules responsible for the strong effects of oligodendroglial membranes on
myelinogenic potential. Ultimately, it will be important to test for cell-specific effects of
Nogo-A and additional candidates in vivo. This could be accomplished using a cre-lox
strategy to selectively target the expression of candidate molecules specifically in
oligodendroglial cells. We are currently breeding our mouse line expressing GFP in 0.1%
of oligodendrocytes with mice expressing Cre recombinase under the control of the CNP
promoter. The CNP-Cre mouse line expresses Cre recombinase selectively in
myelinating cells. After successfully breeding these two lines, it will be possible to cross
their progeny with existing transgenic mouse lines expressing floxed versions of
candidate molecules. This strategy will generate mice in which the expression of Cre
 
 
88 
recombinase excises the desired candidate molecule selectively in oligodendrocytes.
Using the sparse expression of GFP in oligodendrocytes, it will then be possible to
evaluate the relative importance of cell-autonomous candidate molecules in regulating the
myelinogenic potential of individual oligodendrocytes. This strategy is particularly
important to examine candidate molecules where expression is not restricted exclusively
to oligodendroglial cells. This approach should provide additional insight into the role of
inhibitory cues in the regulation of myelinogenic potential. In addition, it will be
interesting to investigate potential extrinsic inductive cues, most likely expressed by
axons. The identification of these factors could also play a crucial role in the
development of therapeutic strategies. Finally, it is intriguing to speculate about the
intrinsic capacity of myelinating cells and their ability to modulate the length, number
and thickness of myelin internodes in the absence of any extrinsic cues. Strategies to test
this potential could ultimately depend on the ability to induce oligodendroglial to form
myelin around articifical nanotubes or fibers-allowing for the examination of myelination
in the absence of extrinsic factors. Previous efforts in this direction have been limited
primarily to oligodendroglial ensheathment, rather than myelination (Howe 2006).
However, preliminary experiments suggest that this exciting possibility may soon
become a reality.
An intriguing aspect of the studies presented here has been the relatively novel
focus on interactions between oligodendroglial cells. I have enjoyed these studies
immensely, and my experience has certainly helped to cultivate a deep interest in, and
appreciation for understanding how cells work together as a population to achieve a
 
 
89 
given task and functional outcome. This growing fascination has contributed to my
interest in systems neuroscience and a desire to understand the formation, function and
modification of neural circuits. As described previously, the maintenance of functional
circuits is intimately tied to the presence of myelin internodes and myelinating cells.
Examining myelinogenic potential and the appropriate spatial alignment of myelin
internodes naturally brings up many questions about the intimate relationship between
axons and myelinating glia. It is interesting to speculate that oligodendroglial may be
involved not only in the maintenance of neural circuits, but also in regulating both the
formation and the function of these circuits. A remaining question is how
oligodendroglial choose which axons to myelinate out of the many available axons to
choose from? An interesting corollary to this question is whether any coordination exists
between the axons myelinated by a single cell. Naturally, one could speculate that axons
myelinated at a similar time would be likely to signal in parallel, increased the likelihood
that these neurons will wire together and potentially leading to stabilization of synaptic
connections as a result of correlated inputs. In addition, it is intriguing to wonder
whether the length of myelin internodes could also contribute to temporal coordination
between neighboring axons, perhaps also influencing where and when synaptic
connections are formed. It is possible that myelin could also play a role in spike-timing
dependent plasticity (Fields 2005). Recent findings suggest that myelin, and specifically
Nogo-66 receptor (NgR) signaling plays an important role in the regulation of critical
period closure in the visual cortex (McGee et al 2005). It will be extremely exciting to
follow what appears to be a burgeoning interest in the relationship between myelination
 
 
90 
and novel influences on circuit formation, function and plasticity. While it is my desire to
investigate circuitry from a different perspective, I will also retain a keen interest in the
role myelination plays in this process. Ultimately, my experience in graduate school and
my opportunity to examine oligodendroglial differentiation and myelination has greatly
influenced both my development as a scientist and as a person. I will always be grateful
for the lessons I have learned through this experience. I feel lucky to have the opportunity
to endeavor to contribute to our understanding of the fundamental mechanisms that
govern both the development and the function of the nervous system. I hope that
ultimately these efforts will have some sort of translational relevance and will also help
contribute to our individual capacity to truly understand both others and ourselves.

 
 
91 
REFERENCES
Alenghat FJ, Ingber DE. 2002. Mechanotransduction: all signals point to cytoskeleton,
matrix, and integrins. Sci STKE 2002:PE6
Baron W, Colognato H, ffrench-Constant C. 2005. Integrin-growth factor interactions as
regulators of oligodendroglial development and function. Glia 49:467-79
Barres BA, Lazar MA, Raff MC. 1994. A novel role for thyroid hormone, glucocorticoids
and retinoic acid in timing oligodendrocyte development. Development 120:1097-
108
Baumann N, Pham-Dinh D. 2001. Biology of oligodendrocyte and myelin in the
mammalian central nervous system. Physiol Rev 81:871-927
Belachew S, Aguirre AA, Wang H, Vautier F, Yuan X, et al. 2002. Cyclin-dependent
kinase-2 controls oligodendrocyte progenitor cell cycle progression and is
downregulated in adult oligodendrocyte progenitors. J Neurosci 22:8553-62
Bershadsky A, Kozlov M, Geiger B. 2006. Adhesion-mediated mechanosensitivity: a
time to experiment, and a time to theorize. Curr Opin Cell Biol 18:472-81
Boudreau NJ, Jones PL. 1999. Extracellular matrix and integrin signalling: the shape of
things to come. Biochem J 339 ( Pt 3):481-8
Brugarolas J, Chandrasekaran C, Gordon JI, Beach D, Jacks T, Hannon GJ. 1995.
Radiation-induced cell cycle arrest compromised by p21 deficiency. Nature
377:552-7
Buttery PC, ffrench-Constant C. 1999. Laminin-2/integrin interactions enhance myelin
membrane formation by oligodendrocytes. Mol Cell Neurosci 14:199-212
Calver AR, Hall AC, Yu WP, Walsh FS, Heath JK, et al. 1998. Oligodendrocyte
population dynamics and the role of PDGF in vivo. Neuron 20:869-82
 
 
92 

Casaccia-Bonnefil P, Hardy RJ, Teng KK, Levine JM, Koff A, Chao MV. 1999. Loss of
p27Kip1 function results in increased proliferative capacity of oligodendrocyte
progenitors but unaltered timing of differentiation. Development 126:4027-37
Casaccia-Bonnefil P, Tikoo R, Kiyokawa H, Friedrich V, Jr., Chao MV, Koff A. 1997.
Oligodendrocyte precursor differentiation is perturbed in the absence of the
cyclin-dependent kinase inhibitor p27Kip1. Genes Dev 11:2335-46
Chan JR. 2007. Myelination: all about Rac 'n' roll. J Cell Biol 177:953-5
Chan JR, Jolicoeur C, Yamauchi J, Elliott J, Fawcett JP, et al. 2006. The polarity protein
Par-3 directly interacts with p75NTR to regulate myelination. Science 314:832-6
Chan JR, Watkins TA, Cosgaya JM, Zhang C, Chen L, et al. 2004. NGF controls axonal
receptivity to myelination by Schwann cells or oligodendrocytes. Neuron 43:183-
91
Charles P, Hernandez MP, Stankoff B, Aigrot MS, Colin C, et al. 2000. Negative
regulation of central nervous system myelination by polysialylated-neural cell
adhesion molecule. Proc Natl Acad Sci U S A 97:7585-90
Chen CS, Alonso JL, Ostuni E, Whitesides GM, Ingber DE. 2003. Cell shape provides
global control of focal adhesion assembly. Biochem Biophys Res Commun
307:355-61
Chen CS, Mrksich M, Huang S, Whitesides GM, Ingber DE. 1997. Geometric control of
cell life and death. Science 276:1425-8
Chong SY, Chan JR. 2010. Tapping into the glial reservoir: cells committed to remaining
uncommitted. J Cell Biol 188:305-12
Crowley JC, Katz LC. 2002. Ocular dominance development revisited. Curr Opin
Neurobiol 12:104-9
 
 
93 
Dawson MRL, Polito A, Levine JM, Reynolds R. 2003. NG2-expressing glial progenitor
cells: an abundant and widespread population of cycling cells in the adult rat
CNS. Mol Cell Neurosci 24:476-88
Deng C, Zhang P, Harper JW, Elledge SJ, Leder P. 1995. Mice lacking p21CIP1/WAF1
undergo normal development, but are defective in G1 checkpoint control. Cell
82:675-84
Deppmann CD, Mihalas S, Sharma N, Lonze BE, Niebur E, Ginty DD. 2008. A model
for neuronal competition during development. Science 320:369-73
Durand B, Fero ML, Roberts JM, Raff MC. 1998. p27Kip1 alters the response of cells to
mitogen and is part of a cell-intrinsic timer that arrests the cell cycle and initiates
differentiation. Curr Biol 8:431-40
Durand B, Gao FB, Raff M. 1997. Accumulation of the cyclin-dependent kinase inhibitor
p27/Kip1 and the timing of oligodendrocyte differentiation. Embo J 16:306-17
Engler AJ, Sen S, Sweeney HL, Discher DE. 2006. Matrix elasticity directs stem cell
lineage specification. Cell 126:677-89
Fancy SP, Baranzini SE, Zhao C, Yuk DI, Irvine KA, et al. 2009. Dysregulation of the
Wnt pathway inhibits timely myelination and remyelination in the mammalian
CNS. Genes Dev 23:1571-85
Feller MB, Scanziani M. 2005. A precritical period for plasticity in visual cortex. Curr
Opin Neurobiol 15:94-100
Feng G, Mellor RH, Bernstein M, Keller-Peck C, Nguyen QT, et al. 2000. Imaging
neuronal subsets in transgenic mice expressing multiple spectral variants of GFP.
Neuron 28:41-51
Fewou SN, Ramakrishnan H, Bussow H, Gieselmann V, Eckhardt M. 2007. Down-
regulation of polysialic acid is required for efficient myelin formation. J Biol
Chem 282:16700-11
 
 
94 
ffrench-Constant C, Colognato H. 2004. Integrins: versatile integrators of extracellular
signals. Trends Cell Biol 14:678-86
Fields RD. 2005. Myelination: an overlooked mechanism of synaptic plasticity?
Neuroscientist 11:528-31
Franklin RJ. 2002. Why does remyelination fail in multiple sclerosis? Nat Rev Neurosci
3:705-14
Franklin RJ, Kotter MR. 2008. The biology of CNS remyelination: the key to therapeutic
advances. J Neurol 255 Suppl 1:19-25
Franklin RJM, Ffrench-Constant C. 2008. Remyelination in the CNS: from biology to
therapy. Nat Rev Neurosci 9:839-55
Gao FB. 2007. Molecular and cellular mechanisms of dendritic morphogenesis. Curr
Opin Neurobiol 17:525-32
Gao FB, Apperly J, Raff M. 1998. Cell-intrinsic timers and thyroid hormone regulate the
probability of cell-cycle withdrawal and differentiation of oligodendrocyte
precursor cells. Dev Biol 197:54-66
Gao FB, Durand B, Raff M. 1997. Oligodendrocyte precursor cells count time but not
cell divisions before differentiation. Curr Biol 7:152-5
Genoud S, Lappe-Siefke C, Goebbels S, Radtke F, Aguet M, et al. 2002. Notch1 control
of oligodendrocyte differentiation in the spinal cord. J Cell Biol 158:709-18
Gensert JM, Goldman JE. 2001. Heterogeneity of cycling glial progenitors in the adult
mammalian cortex and white matter. J Neurobiol 48:75-86
Grimes ML, Zhou J, Beattie EC, Yuen EC, Hall DE, et al. 1996. Endocytosis of activated
TrkA: evidence that nerve growth factor induces formation of signaling
endosomes. J Neurosci 16:7950-64
 
 
95 
Hayashi T, Carthew RW. 2004. Surface mechanics mediate pattern formation in the
developing retina. Nature 431:647-52
He Y, Dupree J, Wang J, Sandoval J, Li J, et al. 2007. The transcription factor Yin Yang
1 is essential for oligodendrocyte progenitor differentiation. Neuron 55:217-30
He Z, Koprivica V. 2004. The Nogo signaling pathway for regeneration block. Annu Rev
Neurosci 27:341-68
Horner PJ, Power AE, Kempermann G, Kuhn HG, Palmer TD, et al. 2000. Proliferation
and differentiation of progenitor cells throughout the intact adult rat spinal cord. J
Neurosci 20:2218-28
Howe CL. 2006. Coated glass and vicryl microfibers as artificial axons. Cells Tissues
Organs 183:180-94
Ingber DE. 1997. Tensegrity: the architectural basis of cellular mechanotransduction.
Annu Rev Physiol 59:575-99
Jaffe AB, Hall A. 2005. Rho GTPases: biochemistry and biology. Annu Rev Cell Dev
Biol 21:247-69
Katz LC, Crowley JC. 2002. Development of cortical circuits: lessons from ocular
dominance columns. Nat Rev Neurosci 3:34-42
Kirby BB, Takada N, Latimer AJ, Shin J, Carney TJ, et al. 2006. In vivo time-lapse
imaging shows dynamic oligodendrocyte progenitor behavior during zebrafish
development. Nat Neurosci 9:1506-11
Kullander K, Klein R. 2002. Mechanisms and functions of Eph and ephrin signalling. Nat
Rev Mol Cell Biol 3:475-86
Lee JK, Chan AF, Luu SM, Zhu Y, Ho C, et al. 2009. Reassessment of corticospinal tract
regeneration in Nogo-deficient mice. J Neurosci 29:8649-54
 
 
96 
Lee X, Yang Z, Shao Z, Rosenberg SS, Levesque M, et al. 2007. NGF regulates the
expression of axonal LINGO-1 to inhibit oligodendrocyte differentiation and
myelination. J Neurosci 27:220-5
Levine JM. 1989. Neuronal influences on glial progenitor cell development. Neuron
3:103-13
Levine JM, Reynolds R, Fawcett JW. 2001. The oligodendrocyte precursor cell in health
and disease. Trends Neurosci 24:39-47
Levison SW, Young GM, Goldman JE. 1999. Cycling cells in the adult rat neocortex
preferentially generate oligodendroglia. J Neurosci Res 57:435-46
Luo L, O'Leary DD. 2005. Axon retraction and degeneration in development and disease.
Annu Rev Neurosci 28:127-56
Mackay DJ, Hall A. 1998. Rho GTPases. J Biol Chem 273:20685-8
Maniotis AJ, Chen CS, Ingber DE. 1997. Demonstration of mechanical connections
between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear
structure. Proc Natl Acad Sci U S A 94:849-54
McBeath R, Pirone DM, Nelson CM, Bhadriraju K, Chen CS. 2004. Cell shape,
cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev Cell
6:483-95
McGee AW, Strittmatter SM. 2003. The Nogo-66 receptor: focusing myelin inhibition of
axon regeneration. Trends Neurosci 26:193-8
McGee AW, Yang Y, Fischer QS, Daw NW, Strittmatter SM. 2005. Experience-driven
plasticity of visual cortex limited by myelin and Nogo receptor. Science
309:2222-6
Meyers J, Craig J, Odde DJ. 2006. Potential for control of signaling pathways via cell
size and shape. Curr Biol 16:1685-93
 
 
97 
Mi S, Miller RH, Lee X, Scott ML, Shulag-Morskaya S, et al. 2005. LINGO-1 negatively
regulates myelination by oligodendrocytes. Nat Neurosci 8:745-51
Michailov GV, Sereda MW, Brinkmann BG, Fischer TM, Haug B, et al. 2004. Axonal
neuregulin-1 regulates myelin sheath thickness. Science 304:700-3
Miralles F, Posern G, Zaromytidou AI, Treisman R. 2003. Actin dynamics control SRF
activity by regulation of its coactivator MAL. Cell 113:329-42
Montell DJ. 2008. Morphogenetic cell movements: diversity from modular mechanical
properties. Science 322:1502-5
Nishiyama A, Chang A, Trapp BD. 1999. NG2+ glial cells: a novel glial cell population
in the adult brain. J Neuropathol Exp Neurol 58:1113-24
Noble M, Murray K, Stroobant P, Waterfield MD, Riddle P. 1988. Platelet-derived
growth factor promotes division and motility and inhibits premature
differentiation of the oligodendrocyte/type-2 astrocyte progenitor cell. Nature
333:560-2
Parrish JZ, Emoto K, Kim MD, Jan YN. 2007. Mechanisms that regulate establishment,
maintenance, and remodeling of dendritic fields. Annu Rev Neurosci 30:399-423
Perona R, Montaner S, Saniger L, Sanchez-Perez I, Bravo R, Lacal JC. 1997. Activation
of the nuclear factor-kappaB by Rho, CDC42, and Rac-1 proteins. Genes Dev
11:463-75
Qi Y, Cai J, Wu Y, Wu R, Lee J, et al. 2001. Control of oligodendrocyte differentiation
by the Nkx2.2 homeodomain transcription factor. Development 128:2723-33
Raff M. 2006. The mystery of intracellular developmental programmes and timers.
Biochem Soc Trans 34:663-70

 
 
98 
Raff MC, Lillien LE, Richardson WD, Burne JF, Noble MD. 1988. Platelet-derived
growth factor from astrocytes drives the clock that times oligodendrocyte
development in culture. Nature 333:562-5
Richardson WD, Pringle N, Mosley MJ, Westermark B, Dubois-Dalcq M. 1988. A role
for platelet-derived growth factor in normal gliogenesis in the central nervous
system. Cell 53:309-19
Rivers L, Young K, Rizzi M, Jamen F, Psachoulia K, et al. 2008. PDGFRA/NG2 glia
generate myelinating oligodendrocytes and piriform projection neurons in adult
mice. Nat Neurosci
Rosenberg SS, Chan JR. 2009. Modulating myelination: knowing when to say Wnt.
Genes Dev 23:1487-93
Rosenberg SS, Kelland EE, Tokar E, De la Torre AR, Chan JR. 2008. The geometric and
spatial constraints of the microenvironment induce oligodendrocyte
differentiation. Proc Natl Acad Sci U S A 105:14662-7
Rosenberg SS, Ng BK, Chan JR. 2006. The quest for remyelination: a new role for
neurotrophins and their receptors. Brain Pathol 16:288-94
Sagasti A, Guido MR, Raible DW, Schier AF. 2005. Repulsive interactions shape the
morphologies and functional arrangement of zebrafish peripheral sensory arbors.
Curr Biol 15:804-14
Salzer JL, Bunge RP, Glaser L. 1980. Studies of Schwann cell proliferation. III. Evidence
for the surface localization of the neurite mitogen. J Cell Biol 84:767-78
Schmidt A, Hall A. 2002. Guanine nucleotide exchange factors for Rho GTPases: turning
on the switch. Genes Dev 16:1587-609
Schnadelbach O, Ozen I, Blaschuk OW, Meyer RL, Fawcett JW. 2001. N-cadherin is
involved in axon-oligodendrocyte contact and myelination. Mol Cell Neurosci
17:1084-93
 
 
99 
Schwab ME, Schnell L. 1989. Region-specific appearance of myelin constituents in the
developing rat spinal cord. J Neurocytol 18:161-9
Schwartz MA, Ginsberg MH. 2002. Networks and crosstalk: integrin signalling spreads.
Nat Cell Biol 4:E65-8
Shen S, Sandoval J, Swiss VA, Li J, Dupree J, et al. 2008. Age-dependent epigenetic
control of differentiation inhibitors is critical for remyelination efficiency. Nat
Neurosci 11:1024-34
Stolt CC, Rehberg S, Ader M, Lommes P, Riethmacher D, et al. 2002a. Terminal
differentiation of myelin-forming oligodendrocytes depends on the transcription
factor Sox10. Genes & Development 16:165-70
Stolt CC, Rehberg S, Ader M, Lommes P, Riethmacher D, et al. 2002b. Terminal
differentiation of myelin-forming oligodendrocytes depends on the transcription
factor Sox10. Genes Dev 16:165-70
Tait S, Gunn-Moore F, Collinson JM, Huang J, Lubetzki C, et al. 2000. An
oligodendrocyte cell adhesion molecule at the site of assembly of the paranodal
axo-glial junction. J Cell Biol 150:657-66
Tang XM, Beesley JS, Grinspan JB, Seth P, Kamholz J, Cambi F. 1999. Cell cycle arrest
induced by ectopic expression of p27 is not sufficient to promote oligodendrocyte
differentiation. J Cell Biochem 76:270-9
Tang XM, Strocchi P, Cambi F. 1998. Changes in the activity of cdk2 and cdk5
accompany differentiation of rat primary oligodendrocytes. J Cell Biochem
68:128-37
Taveggia C, Zanazzi G, Petrylak A, Yano H, Rosenbluth J, et al. 2005. Neuregulin-1 type
III determines the ensheathment fate of axons. Neuron 47:681-94
Temple S, Raff MC. 1986. Clonal analysis of oligodendrocyte development in culture:
evidence for a developmental clock that counts cell divisions. Cell 44:773-9
 
 
100 
Tokumoto YM, Tang DG, Raff MC. 2001. Two molecularly distinct intracellular
pathways to oligodendrocyte differentiation: role of a p53 family protein. Embo J
20:5261-8
Trapp BD, Kidd GJ. 2000. Axo-glial septate junctions. The maestro of nodal formation
and myelination? J Cell Biol 150:F97-F100
Vartiainen MK, Guettler S, Larijani B, Treisman R. 2007. Nuclear actin regulates
dynamic subcellular localization and activity of the SRF cofactor MAL. Science
316:1749-52
Voyvodic JT. 1989. Target size regulates calibre and myelination of sympathetic axons.
Nature 342:430-3
Wang S, Sdrulla AD, diSibio G, Bush G, Nofziger D, et al. 1998. Notch receptor
activation inhibits oligodendrocyte differentiation. Neuron 21:63-75
Watkins TA, Emery B, Mulinyawe S, Barres BA. 2008. Distinct Stages of Myelination
Regulated by gamma-Secretase and Astrocytes in a Rapidly Myelinating CNS
Coculture System. Neuron 60:555-69
Windrem MS, Nunes MC, Rashbaum WK, Schwartz TH, Goodman RA, et al. 2004.
Fetal and adult human oligodendrocyte progenitor cell isolates myelinate the
congenitally dysmyelinated brain. Nat Med 10:93-7
Yamauchi J, Chan JR, Shooter EM. 2004. Neurotrophins regulate Schwann cell
migration by activating divergent signaling pathways dependent on Rho GTPases.
Proc Natl Acad Sci USA 101:8774-9
Yeh HJ, Ruit KG, Wang YX, Parks WC, Snider WD, Deuel TF. 1991. PDGF A-chain
gene is expressed by mammalian neurons during development and in maturity.
Cell 64:209-16
Yiu G, He Z. 2006. Glial inhibition of CNS axon regeneration. Nat Rev Neurosci 7:617-
27
 
 
101 
Zezula J, Casaccia-Bonnefil P, Ezhevsky SA, Osterhout DJ, Levine JM, et al. 2001.
p21cip1 is required for the differentiation of oligodendrocytes independently of
cell cycle withdrawal. EMBO Rep 2:27-34
Zhang H, Miller RH. 1996. Density-dependent feedback inhibition of oligodendrocyte
precursor expansion. J Neurosci 16:6886-95
Zhao C, Li WW, Franklin RJ. 2006. Differences in the early inflammatory responses to
toxin-induced demyelination are associated with the age-related decline in CNS
remyelination. Neurobiol Aging 27:1298-307
Zhou Q, Choi G, Anderson DJ. 2001. The bHLH transcription factor Olig2 promotes
oligodendrocyte differentiation in collaboration with Nkx2.2. Neuron 31:791-807

Abstract (if available)

Abstract As humans, a swift and highly accurate means of neuronal communication is required for both our basic survival, and our unique capacity for traits such as creativity and critical thinking. Communication between the nervous system and all other systems, as well as within the nervous system itself, depends on the transmission of neuronal action potentials, electrical signals required for the generation of functional outputs. The efficient transmission of the neuronal action potential is greatly enhanced by the insulating properties of the myelin sheath. Destruction of the myelin membrane, as a result of nerve injury or disease, significantly impairs the ability of the nervous system to communicate and can lead to a host of debilitating symptoms, as well as an ultimate loss of function. The development of novel strategies to promote remyelination is essential to limit the extent of sensory, motor and cognitive deficits that occur following demyelination. It is our belief that understanding how myelination occurs during development could provide insight into efforts to promote remyelination.

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Creator Rosenberg, Sheila Sara (author)

Core Title Exploring the role of cooperative and competitive interactions in oligodendrocyte differentiation and myelination

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Neuroscience

Publication Date 11/07/2010

Defense Date 03/29/2010

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag myelin,OAI-PMH Harvest,oligodendrocyte

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Chan, Jonah (committee chair), Chow, Robert HP. (committee member), Crump, Gage (committee member), Maxson, Robert E. (committee member), Tao, Huizhong (committee member)

Creator Email sheilasrosenberg@gmail.com,srosenbe@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m3046

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