THE EFFECTS OF EXCITOTOXICITY AND MICROGLIAL ACTIVATION ON OLIGODENDROCYTE SURVIVAL
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Brandon Andrew Miller, B.A.
*****
The Ohio State University 2006
Dissertation Committee: Approved by
Professor Phillip G. Popovich, Co-Advisor ______
Professor Michael S. Beattie, Co-Advisor ______
Professor Jacqueline C. Bresnahan, Co-Advisor ______
Professor Richard H. Fertel Co-Advisors, Neuroscience Graduate Studies Program Professor Chandan K. Sen
ABSTRACT
The nervous system transmits information over long distances by action potentials carried by neurons. In both the central and peripheral nervous system, neurons are insulated by myelin which improves the conduction of action potentials. In the peripheral nervous system, myelin is produced by Schwann cells. Oligodendrocytes (OLs) are the myelinating cells of the central nervous system (CNS). Both mature OLs and oligodendrocyte progenitor cells (OPCs) are lost in CNS disease and injury. Two contributing factors to OL and OPC cell death are inflammation and excitotoxicity. These studies were designed to better understand how OLs and OPCs respond to excitotoxicity and inflammatory stimuli. The glutamate receptor agonist kainic acid (KA) was used to induce excitotoxic cell death in OLs from optic nerve. We found that the pro- inflammatory cytokine tumor necrosis factor-alpha increased the susceptibility of optic nerve OLs to KA. Subsequent experiments tested the response of cortical
OLs at different developmental stages to KA and revealed that mature cortical
OLs were also vulnerable to KA-induced excitotoxicity. In the final set of experiments, we used combined culture of OLs or OPCs and microglia (MG) to study the effects of activated MG on OPC and OL survival. We found that MG activated with lipopolysaccharide induced OPC cell death, but MG, regardless of
ii activation state, protected mature OLs from apoptotic cell death. We also observed that MG cell death occurs as a consequence of activation. Taken together, these studies demonstrate that both excitotoxicity and inflammatory stimuli can induce OL and OPC death but that MG can also support the survival of OLs.
iii
Dedicated to Rhonda and Jeff Miller
iv
ACKNOWLEDGMENTS
I would like to acknowledge all of those who made this work possible:
Rochelle Diebert, Crystal Forrider, John Komon, Jeannine Crum, Tina Van
Meter, and especially Amy Tovar made invaluable technical and intellectual contributions to all of these projects. Thank you for helping me get things done right!
I would like to thank Dr. Michael Beattie and Dr. Jacqueline Bresnahan for their mentorship and support for the past six years. It has been a privilege to work with and learn from you.
Soon to be Dr. John Gensel – You’ve been a great friend and colleague for the past six years. It’s been an honor and a privilege to share an office with you and I wouldn’t trade you for a window any day! You will be an incredible mentor for young scientists one day soon.
Dr. Adam Ferguson – Thank you for all your help in experimental design, statistics, and the occasional coffee break. You are an awesome friend and mentor. It’s been great to live just down the street from you in SF!
Dr. Yvette Nout – Thank you for your advice and example for a young clinician-scientist. I know you’ll laugh (though you shouldn’t!), but one day I hope to emulate your success. Take good care of LR. v
Dr. Fang Sun – Thank you for your help with immunohistochemistry and all of our discussions about oligodendrocytes. I still think of your tenacity when I feel overwhelmed.
I would like to thank the members of my dissertation committee - Dr.
Phillip Popovich, Dr. Richard Fertel and Dr. Chandan Sen – for their mentorship, teaching and advice.
Dr. Allan Yates, Dr. Joanne Lynn, Dr. Charles Hitckcock, Dr. Mirza Baig,
Dr. Abhik Chaudhury, Dr. Louis Caragine, Dr. Michael Gong, Dr. David Feldman,
Dr. Howard Werman, Dr. Sandra Kostyk, Dr. David Bahner, Dr. Shirley Stiver, Dr.
Guy Rosenthal, and Dr. Geoff Manley - thank you for your mentorship as I’ve ventured outside the lab during my PhD.
Cheryl Yoder, Ashley Bertran, and Andrea Brown – thank you for your help with administrative issues - especially the copier!
Dr. Jeff Gidday – thank you introducing me to Neuroscience, and for your patience with a college student still learning so much. I hope we will have the opportunity to work together in the future.
My good friends who aren’t listed above– Nadine, Dan, Ronnie, Bethany,
BT, Gitis, Kristen, Sam, Kelly, Peter, Emma, TJ, Jason, Steve, Phillip, Michelle,
Amy, The Mikes, Liz, Chad, Eric and Jen – your support has made this possible and your friendship continues to make life great!
Most importantly, to my family: Rhonda and Jeff – thank you for dauntlessly supporting me all these years. Laura and Mike – thanks for so many
vi fun times in Delaware and Philly. My grandparents and great aunt – thank you for the value you’ve placed on education and hard work. Jared – your hard work and courage will always be an inspiration to me.
vii
VITA
12/17/1977 ...... Born – Cleveland, OH
5/8/2000 ...... B.A. Biology with College Honors Washington University in St. Louis
2000 – 2002 ...... Medical School Preclinical Curriculum The Ohio State University
2002 – 2006 ...... Graduate Research Associate The Ohio State University
2006-present ...... NRSA Dual Degree Fellow National Institute of Neurological Disorders and Stroke, and The Ohio State University
PUBLICATIONS
1. Miller BA, Baig M, Hayes J, Elton S. (2006) Injury Outcomes in Children Following Automobile, Motorcycle and All-Terrain Vehicle Accidents: an Institutional Review. The Journal of Neurosurgery: Pediatrics 105:182-186
2. Miller BA, Sun F, Christensen RN, Ferguson AR, Bresnahan JC, Beattie MS. (2005) A Sublethal Dose of TNF α Potentiates Kainate-Induced Excitotoxicity in Optic Nerve Oligodendrocytes. Neurochemical Research 30: 867-875
3. Miller BA, Perez RS, Shah AR, Gonzales ER, Park TS, Gidday JM. (2001) Cerebral Protection by Hypoxic Preconditioning in a Murine Model of Focal Ischemia-Reperfusion. NeuroReport 12:1663-1669
FIELDS OF STUDY
Major Field: Neuroscience
viii
TABLE OF CONTENTS
P a g e
Abstract...... ii
Dedication...... iv
Acknowledgments ...... v
Vita ...... viii
List of Figures ...... xi
Chapters:
Chapter 1: Introduction ...... 1
Chapter 2: A sublethal dose of TNF α Increases Excitotoxic Cell Death in Optic Nerve Oligodendrocytes ...... 18 Introduction ...... 18 Materials and Methods ...... 23 Results ...... 27 Discussion ...... 33
Chapter 3: Characterization of the Response of Cortical Oligodendrocytes to Kainic Acid Excitotoxicity ...... 40 Introduction ...... 40 Materials and Methods ...... 44 Results ...... 52 Discussion ...... 62
Chapter 4: The Effects of LPS-Activated Microglia on Oligodendrocyte Progenitor Cells and Mature Oligodendrocytes...... 73 Introduction ...... 73
ix
Materials and Methods ...... 75 Results ...... 78 Discussion ...... 90
Chapter 5: Microglial Cell Death in Response to LPS-Activation ...... 95 Introduction ...... 95 Materials and Methods ...... 97 Results ...... 98 Discussion ...... 102
Chapter 6: Summary, Conclusions and Future Directions ...... 107
References ...... 114
x
LIST OF FIGURES
Figure Page
1.1 Schematic of OPC and OL morphology ...... 3
1.2 Pathways of OL cell death after CNS trauma...... 15
2.1 Optic nerve OL immunocytochemistry ...... 28
2.2 Optic nerve OLs labeled for GluR4 ...... 29
2.3 Effect of short term KA exposure on optic nerve OL survival . . . 30
2.4 Effect of long term KA exposure on optic nerve OL survival . . . 32
2.5 Caspase activation in optic nerve OLs ...... 33
2.6 Effect of TNF α on KA excitotoxicity ...... 34
3.1 Cortical OL shakeoff procedure ...... 47
3.2 Comparison of live and dead OLs and OPCs ...... 51
3.3 Cortical OL response to KA ...... 54
3.4 Immunocytochemistry of OLs with and without TH ...... 55
3.5 Response of OLs with and without TH to KA ...... 56
3.6 OPC response to KA – cell counts ...... 57
3.7 OPC response to KA – LDH assay ...... 58
3.8 OL caspase labeling with caspase kit ...... 59
3.9 Caspase activation in response to time and KA ...... 60
3.10 OLs with and without MG ...... 62
3.11 Response of OLs with and without MG to KA ...... 63
xi
4.1 MG TNF α release in response to KA and LPS ...... 78
4.2 Immunocytochemistry of OPCs ...... 80
4.3 Immunocytochemistry of OPCs and OLs with MG ...... 81
4.4 OPC survival in response to MG and LPS ...... 82
4.5 OPCs labeled with BrdU ...... 83
4.6 OPC BrdU incorporation quantification ...... 84
4.7 OPCs labeled with caspase kit ...... 84
4.8 OPC apoptosis in response to MG and LPS ...... 85
4.9 Immunocytochemistry of OLs ...... 87
4.10 OL Survival in response to MG and LPS ...... 88
4.11 OL apoptosis in response to MG and LPS ...... 89
5.1 TNF α production by MG in different media ...... 99
5.2 MG cell death in the presence of OPCs and OLs ...... 100
5.3 MG cell death in the absence of OPCs and OLs ...... 101
5.4 LDH assay for MG cell death ...... 103
6.1 OL labeling with Carboxy-H2DCFDA ...... 109
6.2 Schematic of results in Chapters 4 and 5 ...... 113
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CHAPTER 1
INTRODUCTION
Oligodendrocytes in Health and Disease
Myelination allows neuronal axons to quickly conduct action potentials over long distances. Oligodendrocytes (OLs) are the myelin-producing cells of the central nervous system (CNS). Each OL contributes to the myelination of about 15 CNS axons, allowing neurons to efficiently conduct action potentials.
Though more prominent in the white matter of the brain and spinal cord, OLs are present in the grey matter as well. Mature OLs develop from a biopotential progenitor cell, often referred to as an oligodendrocyte progenitor cell (OPC), or oligodendrocyte – type 2 astrocyte cell (O2A) as it can mature into either an oligodendrocyte or astrocyte in vitro (Raff et al., 1983). Though more prevalent in the immature CNS, OPCs persist in the CNS of mature animals and humans
(Ffrench-Constant and Raff, 1986; Reyners et al., 1986; Wolswijk and Noble,
1989; Scolding et al., 1999), and have been shown to respond to CNS injury by proliferating and possibly taking the place of mature OLs that are lost during injury or disease (McTigue et al., 1998; McTigue et al., 2001; Woodruff et al.,
2004; Foote and Blakemore, 2005). However, there are physiologic differences
1 between OPCs from the mature CNS and OPCs from the immature CNS, as
OPCs from the adult CNS have been shown to have a different response to growth factors than their perinatal counterparts (Noble et al., 1989; Wolswijk et al., 1990).
The distinction between OPCs and mature OLs is often difficult to delineate in in vitro studies. In vitro OL development has been classically defined by morphology and expression certain antigens that appear sequentially as
OPCs develop into OLs. The proteoglycan NG2 is often used as a marker for immature, proliferating and motile bipolar OPCs (Rogister et al., 1999; Wilson et al., 2003). The antibody A2B5 is also used to label OPCs that are still dividing
(Temple and Raff, 1986). The proteoglycan galactocerebroside (GalC) has been used as a marker for more mature multiploar OLs that are no longer dividing.
Expression of myelin basic protein (MBP) is commonly used as evidence that
OLs have reached their terminally differentiated, myelinating stage (Dietrich et al., 2002). Unfortuneately, these markers for OL development have been described by others as “arbitrary and overlapping” (McDonald et al., 1998b), and we must agree with this assessment. Here, we rely on both antigenic markers and morphology when we apply the terms “OPC” and “OL,” and take care to clearly define the cells used in each experiment (Fig. 1.1). As we and others have shown, not only do OLs at different stages of development have different morphological characteristics, but OPCs and OLs vary greatly in their response to injury.
2
Figure 1.1 Simplified diagram of OPC and OL morphology in vitro . OPCs possess bipolar morphology and develop an increased number of processes as they mature. Mature OLs have multiple branched processes and elaborate myelin sheets. Solid lines represent the typical expression pattern of antigens used to define OL development in vitro.
Injury to and loss of OPCs and mature OLs occurs in many human
diseases. In periventricular leukomalacia (PVL), OPCs are lost as a
consequence of hypoxia, ischemia, or intrauterine infection (Blumenthal, 2004).
This loss of OPCs, and the resulting deficiency of mature OLs, is thought to be 3 the pathologic cause of spastic cerebral palsy (Haynes et al., 2003). This condition affects 5,700 children born each year in the United States (Volpe,
2001) and current treatment is focused on supportive measures rather than limiting OPC loss or promoting repair of damaged tissue. OPC loss in PVL is thought to be mediated by glutamate, cytokines, and free radicals. Glutamate is widely hypothesized to contribute to PVL (Follett et al., 2004; Jensen, 2005;
Back, 2006) and OPCs have been shown to be especially susceptible to glutamate-induced excitotoxicity (Jensen, 2002), which will be described in more detail later in this chapter. The cytokine tumor necrosis factor alpha (TNF α) is
found at elevated levels in the white matter of infants with PVL (Deguchi et al.,
1996; Yoon et al., 1997; Perlman, 1998) and OPCs have also been shown to
undergo cell death when exposed to TNF α in vitro (Pang et al., 2005). One source of TNF α in the CNS are microglia (MG), the resident immune cells of the
CNS. In addition to TNF α, MG can also produce free radicals which are though
to contribute to the pathology of PVL. Cerebrospinal fluid of infants with white
matter injury contains elevated lipid peroxidation products, which are generated
during free radical-mediated injury (Calabrese et al., 2000; Volpe, 2001). OPCs
themselves have been shown to be vulnerable to free radical-mediated damage
in vitro, possibly due to low levels of glutathione and superoxide dismutases
which protect cells free radical-mediated injury (Fragoso et al., 2004; Folkerth,
2006). Free radical-mediated damage to OPCs could be exacerbated by the
hemorrhage that occurs in PVL. Blood and blood breakdown products within the
4
CNS could increase CNS iron which catalyzes the reaction of hydrogen peroxide
(which is generated from the activity of superoxide dismutase) to the more reactive hydroxide free radical. Hydroxide free radicals can directly damage cells by reacting with and damaging DNA, proteins and lipids (Volpe, 2001; Madrigal et al., 2006).
Mature OLs are lost in multiple sclerosis (MS), the most common disease of the adult CNS, affecting over 250,000 people living in the US (Anderson et al.,
1992). Symptoms of MS can range from minor sensory or motor deficits, to severe cognitive impairment accompanied by total paralysis and eventually death. The etiology of MS is currently unknown, though it is has been hypothesized that MS develops due to a combination of environmental and genetic factors (Cook, 2004; Imitola et al., 2006). Many of the same factors that are thought to contribute to OPC loss in PVL also affect OLs in MS. Glutamate receptor activation is thought to contribute to OL loss in MS (Bolton and Paul,
2006; Matute et al., 2006) as are TNF α activity and free radical formation
(Gilgun-Sherki et al., 2004; Magnano et al., 2004).
OLs and OPCs also die after CNS trauma, such as brain and spinal cord
injury (Beattie et al., 2002b; Castejon, 1985). In spinal cord injury, it has been
shown that OLs undergo apoptosis (programmed cell death) for several weeks
after the initial injury (Shuman et al., 1997; Crowe et al., 1997). This delayed
death of OLs is thought to reduce the effectiveness of neural conduction in the
spared rim of white matter that often exists in injured spinal cords, and impair
5 optimal function of spared descending and ascending neural pathways. As in
PVL and MS, OL loss in trauma is thought to be mediated by excitotoxicity, inflammation and free radicals (Beattie, 2004).
Oligodendrocyte Cell Death via Excitotoxicity
Numerous experimental models have been used to study OL and OPC death, from in vitro preparations using purified cell populations to animal models
designed to closely mimic human disease. Across many models of different
diseases, glutamate-mediated excitotoxicity emerges as a primary cause of OL
and OPC death. Excitotoxicity was first described in neurons over 30 years ago
(Olney, 1969). In neurons, excess glutamate first activates non-N-methyl-D-
aspartate (NMDA) receptor ion channels of the α-amino-3-hydroxyl-5-methyl-4- isoxazolepropionate (AMPA) and kainate (or kainic acid, KA) families, leading to the expulsion of the magnesium ion from the pore of NMDA receptor-coupled ion channels (Michaelis, 1998). Simultaneously, glutamate induces NMDA channel opening and allows additional positive ions, most notably calcium, into the cell.
Excess calcium entry into neurons can initiate many intracellular events that mediate cell death (Arundine and Tymianski, 2004). Calcium can activate nitric oxide synthases (NOS) of the endothelial and neuronal isoforms (eNOS and nNOS) which produce the free radical nitric oxide (NO) (Nicotera and Lipton,
1999). As intracellular calcium rises, calcium is taken up into the mitochondria along its electrochemical gradient primarily by a uniporter – however the details of this process are poorly understood (Duchen, 2000; Camello-Almaraz et al.,
6
2006). The electrochemical gradient for calcium is maintained by a sodium calcium exchanger and by negative mitochondrial membrane potential relative to the intracellular space (Duchen, 2000). Excess calcium entry into mitochondria results in an increase in superoxide formation above normal levels produced during oxidative phosphorlyation (Camello-Almaraz et al., 2006). Superoxide can rapidly combine with NO to form the highly reactive free radical peroxynitrite
(ONOO -). Peroxynitrite is a potent oxidant and can directly damage DNA, protein and lipids. Superoxide can also be reduced to hydrogen peroxide by the enzyme superoxide dismutase. In the presence of iron, hydrogen peroxide can be converted to the hydroxyl free radical (OH -) which can also directly damage
cellular structures (Warner et al., 2004). Oxidative damage to mitochondria can
induce the release cytochrome c from the mitochondrial intermembrane space
(Petrosillo et al., 2003; Szeto, 2006). Once released into the extracellular space,
cytochrome c associates with Apaf-1 in the presence of ATP or dATP and this
“apoptosome” complex mediates the activation of caspase 9 (Cho and Choi,
2002). Caspase-9 subsequently activates caspase-3, which can also undergo
autolytic cleavage and activation. Once activated, caspase-3 induces many of
the cellular events associated with apoptosis such as DNA fragmentation and
cellular blebbing (Janicke et al., 1998). Apoptosis occurring through
mitochondrial cytochrome c release is often referred to as the intrinsic pathway of
apoptosis. Though caspase-3 is typically thought of as a downstream “effector”
caspase, it has been recently shown that caspase-3 also contributes to apoptotic
7 events that occur farther upstream such as mitochondrial depolarization and cytochrome c release (Lakhani et al., 2006). Intracellular calcium can also activate calpain enzymes that can subsequently activate caspase-3 or directly cleave structural proteins within cells, leading to apoptosis. Calpain and caspase activation have been shown to occur in OLs after spinal cord injury, where excitotoxic OL death also occurs (Ray et al., 2000; Ray et al., 2003; Park et al.,
2004a; McEwen and Springer, 2005). The linkage between excitotoxic and apoptotic OL death in vivo is of clinical importance because the multiple coordinated steps of apoptosis provide discrete therapeutic targets that may be amenable to pharmacologic intervention.
Though much evidence supports a link between excitotoxicity and apoptosis, excitotoxic cell death in neurons has been shown to be both apoptotic and necrotic in nature depending on the severity of the insult (Mergenthaler et al.,
2004). Additionally, it has been hypothesized that the determining factor as to whether neurons undergo apoptosis or necrosis in response to excitotoxicity is the amount of energy available to the cell. If cells have enough energy to execute active apoptotic cell death cascades, they may undergo apoptosis.
However, if cells do not have sufficient intracellular energy stores to activate apoptotic pathways, necrosis my occur by default (Nicotera and Lipton, 1999). In a model of glutamate-induced oxidative stress using a neuronal cell line, cells underwent a form of cell death that exhibited features of both apoptosis and necrosis together (Tan et al., 1998). While there is debate as to what determines
8 if glutamate elicits apoptotic or necrotic neuronal death, the detrimental effects of excess glutamate on neurons has been well established over time and is not currently under debate.
Like neurons, OLs are also vulnerable to excessive stimulation of glutamate receptors (McDonald et al., 1998a). Until recently, it was thought that
OLs lacked NMDA receptors, and only possessed glutamate receptors of the
AMPA and KA preferring varieties (McDonald et al., 1998a; Rosin et al., 2004).
Recently, it has been shown that OLs do possess NMDA receptors, and that these receptors are developmentally regulated and contribute to OL death in oxygen-glucose deprivation and in vivo white matter damage (Follett et al., 2000;
Karadottir et al., 2005; Salter and Fern, 2005).
It has been hypothesized that non-NMDA receptors normally play a role in stimulating OPC migration and reducing OPC proliferation (Gallo et al., 1996;
Gudz et al., 2006). One study showing a decrease in proliferation of OPCs in response to non-NMDA receptor activation demonstrated that this effect was specific to non-NMDA (as opposed to NMDA) receptors. Furthermore, the effects of non-NMDA receptor activation were not due to OPC death (Yuan et al.,
1998). It is possible that as the CNS develops and OPCs migrate and mature, that glutamate released from synapses serves as a signal to prevent OPCs from migrating into areas where they would interfere with synaptic activity.
Regardless of the normal physiologic roles of OL non-NMDA receptors, OLs and
OPCs in both in vitro and in vivo models of the aforementioned diseases have
9 been shown to die as a result of activation of AMPA and KA receptors (Follett et al., 2000; Smith et al., 2000b; Caruso et al., 2004; Soto et al., 2004; Leuchtmann et al., 2003; Pitt et al., 2000; Sanchez-Gomez et al., 2003; Wrathall et al., 1997).
The vulnerability of OLs to glutamate excitotoxicity is thought to be enhanced by the AMPA receptor subunit composition of OLs, which has been shown to lack the edited GluR2 subunit (Matute et al., 1997; Rosin et al., 2004), which is needed to block calcium entry through AMPA channels (Hollmann et al., 1991;
Tanaka et al., 2000).
Excitotoxic OL loss is thought be relevant to several human disease processes as elevated glutamate levels have been shown to occur in experimental SCI (McAdoo et al., 1999; Xu et al., 2004), in the cerebrospinal fluid of patients with active MS plaques (Sarchielli et al., 2003), and likely exist in PVL lesions (Volpe, 2001). It is possible that excitotoxicity is a final common mode of
OL and OPC cell death in many CNS diseases and trauma (Lipton and
Rosenberg, 1994). There are several potential sources of excess glutamate in
CNS disease, including neurons releasing excess neurotransmitter due to hyperexcitation or physical breakdown (Choi, 1988; Panter and Faden, 1992;
Springer et al., 1997), immune cells which produce glutamate (Werner et al.,
2000) and even OLs themselves, which have been shown to release glutamate after hypoxia (Li et al., 1999).
CNS Inflammation as a Mediator of Oligodendrocyte Death
10
There is also evidence that inflammation within the CNS contributes to OL loss in many diseases. Inflammation within the CNS is carried out in part by microglia (MG), the resident immune cells of the CNS. Resident MG in the CNS originate from bone marrow and then migrate into the CNS during early stages of development (Hess et al., 2004). MG display graded levels of activation in the
CNS, from resting ramified MG, to activated MG, to phagocytic macrophages
(Streit et al., 1988). MG are known to react quickly in response to central nervous system injury or disease (Kreutzberg, 1996), migrating into an injury site
(Carbonell et al., 2005) and secreting a wide array of molecules that are toxic to
OLs, including glutamate (Nakamura et al., 2003), free radicals (Benveniste,
1997) and tumor necrosis factor-alpha (TNF α) (Selmaj and Raine, 1988;
Kreutzberg, 1996; Nicholas et al., 2002; Dasgupta et al., 2003; Jana et al., 2003).
There are two different TNF α receptors – TNF-R1 (p55) and TNF-R2 (p75).
TNF-R1 is expressed on most cell types (including OLs) and mediates the majority of TNF α’s biological effects (Dopp et al., 2002; Shen and Pervaiz, 2006).
While both membrane-bound TNF α and soluble TNF α (which is cleaved from the
cell membranes by TNF α converting enzyme, or TACE) can activate TNF-R1,
TNF-R2 is predominantly activated by membrane-bound TNF α (Wajant et al.,
2003). TNF α binding causes receptors to trimerize and initiate several molecular cascades including the translocation of phosphatidylserine to the extracellular surface, a hallmark of apoptosis (Aktas et al., 2006). Like glutamate-mediated OL apoptosis, TNF α mediated apoptosis also induces
11 caspase activation, though through a different pathway (commonly referred to as the extrinsic apoptotic pathway). Trimerization of TNF-R1 subunits activates caspase-8 via adapter molecules that bind to the intracellular domain of the TNF-
R1 receptors. Caspase-8 subsequently activates caspase-3, which executes the final phases of apoptosis. The convergence of both excitotoxic and TNFα – induced apoptosis on caspase-3 creates potential for increased OL apoptosis in diseases where both excitotoxicity and MG activation / TNF α production occur.
Apart from activation of caspase-mediated apoptotic pathways, there are other signaling cascades induced by TNF α. Signaling through TNF-R1 can activate both the transcription factor Nuclear factor – kappa B (NF-kB) and c-Jun
NH2-terminal kinase (JNK, or stress-induced protein kinase). TNF α-induced OL
death has been shown to depend on JNK activation (Jurewicz et al., 2003) and it
has been shown that reactive oxygen species can serve as coactivators of JNK
in response to TNF α signaling. JNK activation also contributes to intracellular free radical production, leading to a possible positive feedback loop between JNK activation and ROS production (Shen and Pervaiz, 2006). Nf-kB activation can induce cellular antioxidant defenses such as manganese superoxide dismutase that can counteract JNK activation / free radical production (Pantano et al.,
2006). In contrast to TNF-R1, TNF-R2 activation preferentially induces Nf-kB activation and intracellular free radical defenses, but OLs in vitro do not express
TNF-R2 and therefore TNF-R1 signaling alone mediates their response to TNF α
(Dopp et al., 2002). The balance between JNK activation / free radical formation
12 and NF-kB / antioxidant defense induction may determine the fate of OLs responding to TNF α. In fact, it has been hypothesized that redox balace is a
central “switch” that determines the fate of OLs – with more reduced states
leading to OL proliferation and more oxidized states leading to OL differentiation
and increased susceptibility to cell death (Smith et al., 2000a; Noble et al., 2003).
A study examining the effects of TNF α and antioxidants in combination on OLs
lends some support to this hypothesis. Cammer found that antioxidants were
able to reverse the effects of TNF α on OLs, which supports the aforementioned
link between the effects of TNF α and ROS formation (Cammer, 2002). However, in this study, the dose of TNF α used did not induce OL death, rather only blocked
the maturation of OPCs to mature OLs. If the response of OLs to different levels
of oxidative stress operates along a simple continuum, starting with proliferation,
then progressing to differentiation and finally to cell death as OPCs/OLs become
more oxidized, it would be expected that if a sublethal dose of TNF α was
administered to OPCs/OLs it would actually promote differentiation. However this
effect of TNF slowing OL maturation has been shown in other studies as well
(Cammer and Zhang, 1999), therefore it is unlikely that TNF α, redox state and
OL maturation are related in such a simple linear fashion.
Not only can TNF α affect redox state, but the response of OPCs/OLs to
both TNF α and growth factors can be amplified or reduced depending on the
redox state of the cell, with more oxidized cellular states favoring a response to
TNF α and subsequent cell death. Therefore, any stimulus that can affect redox
13 state (i.e. –excitotoxic stimuli or oxygen changes) may determine how likely an
OPC/OL is to undergo TNF α-induced cell death (Noble et al., 2005). The
interplay between TNF α and free radical generation can lead to yet another positive feedback loop culminating in OPC/OL cell death. We hypothesize that both caspase activation and redox state are routes through which excitotoxic and inflammatory stimuli can synergize to increase OL death in several diseases where both MG activation and OL death occur.
It has been shown that activated MG contribute to OL death in models of
MS and PVL (Pang et al., 2000; Pang et al., 2003; Sriram and Rodriguez, 1997).
MG-induced OL damage may be further exacerbated by the fact that many factors that lead to OL death also lead to MG activation, such as glutamate
(Noda et al., 2000), kainic acid (Christensen et al., 2006), proinflammatory cytokines (Thery and Mallat, 1993), and physical trauma (Bellander et al., 2004).
The dual effects of these factors on both OLs and MG, and the ability of MG to release molecules that both damage OLs and activate MG has the potential to create a positive feedback loop resulting in increased OL cell death (Fig. 1.2).
MG may exert their effects on OLs and OPCs both at a distance and via direct cell-cell interaction. In vitro , MG have been shown to be capable of inducing
OPC death without being in direct contact with OPCs (Li et al., 2005). However, in vivo , MG have been observed in close proximity to dying OLs after spinal cord injury (Shuman et al., 1997). This proximity of microglia to OLs after CNS damage may increase the influence of MG on OL and OPC survival, as it has
14 been shown in vitro that physical contact between MG and OLs is a key factor in
mediating OL death via membrane bound microglial TNF α (Zajicek et al., 1992).
Figure 1.2 – Schematic of the effects of TNF α, free radicals and glutamate on MG and OPCs/OLs in CNS injury. All three molecules can both activate MG and damage OPCs/OLs resulting in a positive feedback loop increasing both MG activation and OPC/OL cell death.
Additionally, soluble factors produced by MG would have a greater effective
concentration if secreted into a small space between cells. Even the specific
region of an OL with which a MG interacts could affect the response of the OL.
NMDA receptors, which more prominently lead to demyelination when activated,
are located peripherally on OLs, whereas AMPA receptors which mediate cell
death are concentrated on OL soma (Matute, 2006). 15
The Possible Role of MG in Promoting Oligodendrocyte Survival
Though much of the literature cited thus far implies that MG play a deleterious role in CNS injury, there is evidence that MG and their secreted products may play a protective, or at least helpful role, in the injured CNS
(Filipovic and Zecevic, 2005; Shaked et al., 2005). In vitro, MG have been shown to be recruited by soluble factors released by stressed OLs, and to support OL survival via insulin-like growth factor 2 (Nicholas et al., 2002;
Nicholas et al., 2003). Additionally, cytokines produced by MG may aid in repair after injury, as mice lacking TNF α undergo delayed remyelination in a model of
toxic demyelination (Arnett et al., 2001). Even the observations of Shuman and
colleagues (1997), that MG phagocytose OLs after spinal cord injury, raises the
question – do MG destroy OLs that would otherwise survive after injury – or are
MG phagocytosing OLs already destroyed by other toxins in the damaged CNS?
It is also possible that microglia play a dual role in CNS injury, exacerbating
damage in some instances, and promoting repair or regeneration in others.
These studies were carried out to better understand the response of OPCs and
OLs to excitotoxicity and inflammation, and to provide insight into the complex
relationships between OPCs/OLs and MG in the injured CNS. The overarching
hypothesis for this work is that, due to their ability to both induce caspase
activation and increase intracellular free radical formation (which may have a
central role in determining OL survival), inflammatory and excitotoxic stimuli can
synergistically lead to OPC and OL cell death. As these studies progressed,
16 data from early experiments drove us to examine not only the effects of TNF α on
OL excitotoxicity, but the effects of activated MG on OPC and OL survival.
17
CHAPTER 2
A SUBLETHAL DOSE OF TNF α INCREASES EXCITOTOXIC CELL DEATH IN
OPTIC NERVE OLIGODENDROCYTES
Introduction
Excitotoxicity is a common mediator of cell death in central nervous system (CNS) disease and trauma. First described by Olney in 1969 (Olney,
1969), excitotoxicity classically refers to deadly overstimulation of neuronal ionotropic glutamate receptors. More recently, it has been shown that OLs can also be susceptible to cell death in response to elevated amounts of glutamate
(McDonald et al., 1998b; Yoshioka et al., 1995). Though OLs are not thought to conduct action potentials (for an alternate viewpoint see Lev-Ram and Grinvald,
1986, OL death in response to overstimulation of glutamate receptors is also referred to as excitotoxicity. In addition to receptor-dependent glutamate excitotoxicity, both neurons and OLs have been shown to be susceptible to receptor-independent glutamate toxicity, where glutamate interferes with cystine uptake into the cell, preventing synthesis of the antioxidant glutathione, ultimately leading to cell death as a result of oxidative stress (Rosin et al., 2004; Himi et al.,
2003; Li et al., 2003).
18
Neuronal excitotoxicity occurs in models of stroke (Ritz et al., 2004),
Huntington’s disease (Alberch et al., 2002), Parkinson’s disease (Beal, 1998),
CNS viral infection (Nargi-Aizenman et al., 2004), and amyotrophic lateral sclerosis (Wisman et al., 2003). Similarly, OL excitotoxicity is thought to be a factor in many CNS diseases and trauma, including multiple sclerosis (Werner et al., 2001) and spinal cord injury (Wrathall et al., 1997). Additionally, the loss of
OL progenitor cells (OPCs) in models of periventricular leukomalacia has been shown to involve excitotoxicity (Follett et al., 2000) and OL death induced by hypoxia in vitro can be reduced by blocking glutamate receptors (Tekkok and
Goldberg, 2001) .
Previously, it was thought that OLs lacked NMDA type glutamate receptors, only expressing only the AMPA- and KA-sensitive glutamate receptor subtypes (Matute et al., 1997). However, it has been reported recently that OLs have NMDA receptors as well, and that activation of these receptors can lead to
OL injury (Karadottir et al., 2005; Salter and Fern, 2005). Though recent work has examined the specific properties of NMDA receptors in OLs, more is known about non-NMDA (AMPA and KA) receptors in OLs.
AMPA and KA receptors are made up of different subunits, AMPA receptors being composed of subunits GluR 1-4, and KA receptors being composed of subunits GluR 5-7 and KA 1 and KA 2 (Tanaka et al., 2000;
Chittajallu et al., 1999). It has been shown that KA receptors on optic nerve OLs assemble into two different populations, with relatively high and low affinity for KA
19
(Sanchez-Gomez and Matute, 1999). High affinity KA receptors are composed of subunits KA1 and KA2, while low affinity KA receptors are composed of subunits GluR5-7 (Chittajallu et al., 1999). OLs have been shown to lack GluR2, the critical subunit that renders AMPA receptors impermeable to calcium
(Hollmann et al., 1991; Matute et al., 1997; Rosin et al., 2004). Thus, the binding of glutamate to AMPA and KA receptors on OLs, independent of NMDA receptor activation, is thought to initiate the same events that occur in neuronal excitotoxicity, where calcium influx culminates in cell death (Alberdi et al., 2002;
Sanchez-Gomez et al., 2003).
Recently, the rapid recycling of AMPA receptors that occurs in hippocampal neurons has gained attention as a mediator of synaptic plasticity and long term potentiation (Malinow and Malenka, 2002). This rapid recycling of
AMPA receptors is distinct from transcriptional and translational changes that occur over longer periods of time. Neurons retain an intracellular store of vesicles embedded with AMPA receptors which can be exocytosed rapidly
(within minutes) in response to environmental stimuli such as stimulation of glutamate receptors already present in the membrane. This upregulation of surface receptors leads to an increased sensitivity to further excitatory stimuli
(Park et al., 2004b; Man et al., 2000). Other than long term potentiation and depression, there are several physiologic and pathophysiologic conditions that may involve changes in AMPA receptor expression and/or recycling, including
CNS development (Petralia et al., 1999), neuropathic pain (Jones and Sorkin,
20
2004), psychiatric conditions (Du et al., 2004), and excitotoxic cell death (Jensen,
2002).
Like neurons, OLs might also exhibit dynamic glutamate receptor expression through transcription, translation or rapid recycling. Though extensive evidence exists, both in vitro and in vivo , that OLs die via excitotoxicity
(Matute et al., 2001), some studies have shown mature OLs to be resistant to excitotoxicity in vitro , especially when compared to OPCs (Rosenberg et al.,
2003; Wosik et al., 2004). There are many possible explanations for these differing experimental results, including cell source, time, and presence of other cell types in culture. Additionally, culture media has been demonstrated to have a profound effect on excitotoxicity in neurons (Ha et al., 2002), and the same could hold true for OLs. Another possible explanation for laboratories’ differing results could be that OLs, like neurons, have a dynamic glutamate receptor expression pattern in response to environmental stimuli. Indeed, work by Follett and colleagues has revealed a decrease in AMPA receptor expression and a corresponding decrease in susceptibility to glutamate toxicity as OPCs develop into mature cells (Follett et al., 2000). OPCs have also been shown to change their functional glutamate receptor expression in response to growth factors
(Chew et al., 1997), and over time in culture, leading to changes in AMPA- induced intracellular calcium accumulation (Itoh et al., 2002). Additionally, mature OLs in the spinal cord have been shown to change their glutamate receptor expression after injury (Park et al., 2003), though the exact mechanisms
21 of these changes are unknown. If mature OLs are able to rapidly change their
receptor expression in response to the altered CNS environment post injury, this
could have profound implications for their survival.
It has been shown that neurons in the spinal cord can be sensitized to
excitotoxicity by the proinflammatory cytokine tumor necrosis factor alpha (TNF α)
(Hermann et al., 2001). This potentiation of excitotoxicity may occur via rapid
receptor recycling, as hippocampal neurons rapidly increase the density of
functional AMPA receptors on the cell membrane in response to TNF α (Beattie et al., 2002a). This synergy of proinflammatory cytokines and excitatory amino acids may be of great significance in CNS injury, as TNF α is present at elevated levels in the injured CNS. TNF α has been shown to increase after both traumatic brain and spinal cord injury (Fan et al., 1996; Wang et al., 2002; Pan et al.,
2003). TNF α is also present in multiple sclerosis (MS) plaques (Hofman et al.,
1989) and is elevated in the brains of children with periventricular leukomalacia
(Kadhim et al., 2001). Additionally, TNF α is increased in serum of patients with chronic spinal cord lesions (Hayes et al., 2002) and the elevated TNF α levels in
MS lesions and MS patients’ cerebrospinal fluid have been shown to correlate with disease severity (Bitsch et al., 2000; Sharief and Hentges, 1991). There are many possible cellular sources of TNF α in the injured CNS: astrocytes (Zhao and
Brinton, 2004), microglia (Eskes et al., 2003), and cells of the peripheral immune system (Ito et al., 2001) have all been found to increase TNF α production in
response to CNS injury.
22
The experiments described in this study were designed to examine the effects of TNF α on excitotoxic death of OLs in vitro . Although TNF α alone has
been reported to be toxic to OLs (Selmaj and Raine, 1988), studies have shown
that this toxicity develops over periods longer than 24 hours (Louis et al., 1993;
McLaurin et al., 1995); and therefore does not exclude the potential for
additional, and more acute, effects of TNF α on OLs. We hypothesized that
because TNF α and excitotoxicity can activate the same cell death signaling
cascades, and TNF α has been shown to increase neurons’ vulnerability to excitotoxicity, TNF α would increase the susceptibility of OLs to excitotoxicity. In
the present experiments, TNF α was added in combination with the glutamate
receptor agonist KA, and we show that a sublethal dose of TNF α increases the magnitude of excitotoxic cell death in OLs. Results described here have been published previously in Miller et al., 2005.
Materials and Methods
OL Cultures
All experimental protocols in this dissertation were approved by the
Institutional Laboratory Animal Care and Use Committee of The Ohio State
University and are in agreement with NIH guidelines. All reagents were purchased from Sigma (St. Louis, MO) unless otherwise noted. OLs were prepared from postnatal day 7 (P7) Long Evans rat optic nerve in a similar manner as described by Raff and colleagues (Raff et al., 1983). Briefly, rats were sacrificed by rapid decapitation and optic nerves were removed and
23 cleaned of meninges while bathed in L-15 media (Invitrogen, Carlsbad, CA).
Tissue was then minced and incubated at 37 oC with 2 ml collagenase type 4
(13.3 mg/ml, Worthington Biochem, Lakewood, NJ ) in L-15 media for 15 minutes
and centrifuged at 300g for 5 minutes. Next, tissue was resuspended in 2.5 ml 3
% papain (Worthington Biochem) activated with N-acetyl cysteine in L-15 and
incubated at 37 oC for an additional 50 minutes. Enzymatic activity was stopped by addition of 2 ml Dulbecco’s Modified Eagle’s medium (DMEM, Invitrogen) with
0.53 mg/ml trypsin inhibitor, 0.4 mg/ml DNAse and 0.3 % bovine serum albumin
(BSA) for 10 minutes. Following centrifugation as above, cells were resuspended in DMEM containing Sato supplement (Bottenstein and Sato,
1979), and 80 ug/ml gentamicin, 0.33 ug/ml L-thyronine (T3) and 0.40 ug/ml L- thyroxine (T4) (hereafter referred to as “growth media”). Cells were further dissociated by sequential trituration through 18G and 25G needles. 2.75 to 4 x
10 3 trypan blue-excluding cells in 15 ul of growth media were initially plated in the
center of 13.3 ug/ml poly-l-lysine (mw 70,000-150,000) coated 12 mm glass
coverslips in 4-well plates (Nalge Nunc International, Rochester NY). After
waiting 5 minutes for cells to adhere, wells were brought to 500 ul total volume of
o growth media and placed at 37 C in a 5 % CO 2 and 95 % air humidified
incubator. All drug treatments occurred on day 3 in vitro . All test reagents were prepared at twice the final concentration in growth media and were added by replacing half the growth media in the wells with an equal amount of test reagent solution. Control wells were washed with growth media alone in the same
24 fashion. Cultures were fixed with 2% formaldehyde prepared from paraformaldehyde for 10 minutes, rinsed with Hanks Balanced Salt Solution
(HBSS), and stored in HBSS at 4 oC prior to immunocytochemistry.
Immunocytochemistry
For galactocerebroside (GalC, mature OLs) labeling, primary antibody,
(mouse anti-GalC hybridoma supernatant, generous gift of Dr. Mark Noble) was
applied at a dilution of 1:3 in HBSS with 5 % donor calf serum at room
temperature for 30 minutes, followed by goat anti-mouse IgG conjugated with
Alexa 488, (Molecular Probes, Eugene, OR) at 1:00 for 1 hour at room
temperature. Nuclear morphology was visualized via Hoechst labeling (10 ug/ml),
for 10 minutes prior to rinsing coverslips and mounting them onto uncoated glass
microscope slides with Immu-Mount (Thermo, Pittsburgh, PA).
For myelin basic protein labeling (MBP, mature myelinating OLs), cells
were fixed as above and permeabilized with 0.1 % triton in HBSS for 10 minutes.
Cells were then stained with an antibody to myelin basic protein (Chemicon,
Temecula, CA) at a dilution of 1:100 for 90 minutes, followed by goat anti-mouse
Alexa 594 (Molecular Probes) at 1:100 for 1 hour, rinsed, and stained with
Hoechst and mounted to slides as above.
For GluR4 labeling, cells were fixed with as above and incubated with
primary antibody to GluR4 (Chemicon) at 1:40 overnight, rinsed and then
incubated with fluorescently tagged secondary goat anti-rabbit secondary
25 antibody (Molecular Probes) antibody at 1:100 for 1.5 hours. Images of OLs labeled for GluR4 were taken on a Zeiss 510 Meta confocal microscope.
For cleaved caspase-3 and GalC double labeling, cells were rinsed with
PBS+ (0.1M PBS with 0.1 % Saponin and 0.02 % NaN 3) for 5 minutes three
times. Afterwards, cells were blocked with 1 % BSA in PBS+, and then
incubated with an antibody to cleaved caspase-3 at 1:100 (Cell Signaling
Technology, Beverley, MA) overnight in a humid chamber at room temperature.
The cells were then rinsed in PBS+ 3 times and incubated with goat anti-rabbit
Alexa 594 at 1:500 (Molecular Probes) for 2 hours at room temperature. Next,
cells were rinsed again in PBS+ and labeled with anti GalC antibody and Hoechst
as previously described.
Determination of Cell Death
For toxicity experiments, OLs were labeled with GalC and Hoechst as
described above. Cell counts were performed by a blinded observer using a
Zeiss Axioplan microscope (Zeiss, Germany) at 40x magnification. An area of
0.22 mm by 0.17 mm in the center of every second sequential field was
examined, with sampling performed over the entire coverslip. Using this method,
58 ± 23 (mean ± standard deviation) OLs per coverslip were counted. “n” values
correspond to the total number of coverslips analyzed for each condition over the
course of ten separate experiments. Each group was represented in at least two
separate experiments. OLs with an intact cell body and nucleus were scored as
“live.” OLs with an absent, fragmented, or condensed nucleus and either an
26 intact or fragmented cell body were scored as “dead.” Astrocytes were identified by their autofluorescence and morphology and counted. On average, cultures contained 21 ± 1 % astrocytes.
Statistical Analysis
Cell death data were analyzed by analysis of variance (ANOVA) followed by a post hoc Duncan’s Multiple Comparison test and are expressed as mean ±
SEM. Caspase-3 activation was analyzed by 2-way ANOVA with time and KA dose as between factors. For all analyses, statistical significance was set at p <
0.05. Statistical effect size was assessed using the standards established by
Cohen (Cohen, 1997) where partial eta squared ( η2) = 0.01 is considered a small, η2 = 0.06, a medium, and η2 = 0.14 a large effect size.
RESULTS
OLs Rapidly Exhibited a Mature Phenotype
By day three in vitro , OLs could be seen, under phase contrast
microscopy, to elaborate multiple branched processes. On day 4, OLs were
positive for both GalC (Fig. 2.1 A) and MBP (Fig. 2.1 B), indicating a mature
phenotype. Most OLs were observed to have intact, round and uniformly dense
nuclei as revealed by Hoechst labeling. A proportion (16 ± 2 %) of OLs in control
cultures exhibited fragmented, pyknotic or absent nuclei and were categorized as
dead at the time of fixation (Fig. 2.1 C, arrow).
27
Figure 2.1 Optic nerve OLs were positive for GalC (green, A) and MBP (red, B) at the time of fixation. Dead OLs were identified by fragmented nuclei (C, arrow). Apoptotic OLs were identified by GalC (D), Hoechst (E) and Caspase-3 (F) colocalization (G).
OLs Express the AMPA Receptor Subunit GluR4
To determine of OLs in our culture preparation expressed AMPA
receptors, we labeled OLs at day 4 in vitro with antibodies to GluR4, an AMPA receptor subunit that has been previously shown to be expressed on optic nerve
OLs (Matute et al., 1997). OLs labeled with primary antibody to GluR4 and florescent secondary antibody showed several small puncta indicating that GluR4 was present on the cell surface or within the cells (Fig 2.2). A negative control without primary antibody confirmed that actual receptor labeling rather than non- specific antobidy binding had occurred. Functionality of the receptors was shown in subsequent pharmacological experiments.
28
Figure 2.2 Optic nerve OLs labeled positively for the AMPA receptor subunit GluR4. Omission of the primary antibody resulted in negative staining.
OLs are Susceptible to KA Toxicity in a Dose-Dependent Manner
A preliminary experiment showed that KA-induced excitotoxicity was not evident after short periods of exposure, in contrast to data reported by others (Sanchez-
Gomez et al., 2003). OLs were exposed to KA for 4 hours, then the KA- containing media was replaced with fresh growth media and OL viability assessed 24 hours after the initial KA exposure (Fig. 2.3). There was no statistically significant elevation of cell death compared to control values. Control conditions, 100 µM, 300 µM and 1 mM KA resulted in 25 ± 1 %, 23 ± 4 %, 24 ± 1
%, and 23 ± 2 % cell death respectively. Therefore, in order to maximize our ability to detect cell death, experiments were carried out over 24 hours of continuous exposure to drug treatments. OLs were exposed to 100 µM, 200 µM,
29
300 µM and 1 mM KA for 24 hours (Fig. 2.4). 100 µM KA did not induce a significant increase in OL death over control conditions (20 ± 2% cell death versus 16 ± 2% for controls). However, 200 µM, 300 µM and 1 mM KA induced significant OL death in a dose-dependent manner (42 ± 2 %, 63± 4 %, and 88 ± 2
% cell death respectively). The non NMDA-receptor antagonist, 6-cyano-7- nitroquinoxaline-2,3-dione (CNQX) at a dose of 30 µM completely eliminated OL death due to 1mM KA (15 ± 2 % versus 88 ± 2 % cell death), demonstrating that
Figure 2.3 A 4 hour exposure to 100 µM, 300 µM and 1 mM KA did not induce cell death in optic nerve OLs. Cell death was assessed 24 hours after the initial exposure to KA.
30 the effects of KA were indeed mediated by the activity of AMPA and/or KA receptors (Fig. 2.4). The morphology of many of the dead OLs was consistent with an apoptotic mechanism of cell death, with nuclear fragmentation and condensation often observed (Fig. 2.1 C). A separate experiment was conducted to confirm that OL death had an apoptotic component.
KA-Induced OL Death Leads to Caspase-3 Activation
In a separate experiment, OLs were fixed after 6, 14, 18 and 24 hours of continuous exposure to 300 µM KA, and evaluated for cleaved caspase-3 expression at each time point. Cleaved caspase-3 immunoreactivity could be observed in some, but not all, cells with fragmented nuclei (Fig. 2.1 D-G). The number of cells expressing cleaved caspase-3 increased over time in this experiment and revealed a significant main effect of 300 µM KA on caspase-3 activation (p < 0.001, Fig. 2.5). There was no main effect of time on caspase activation, though the effect approached statistical significance (p = 0.06).
TNF α alone does not induce OL death, but potentiates KA-induced toxicity
in OLs
In a preliminary experiment, OLs were initially exposed to 10 ng/ml, 50 ng/ml and
100 ng/ml TNF α for 24 hours, none of which induced in an increase in OL death
(data not shown). Further experiments were conducted using 100 ng/ml TNF α
which again was not observed to have an effect on OL cell death over 24 hours
(15 ± 1 % cell death versus 16 ± 2 % for control; Fig. 2.6). Next, OLs were
exposed to 100 ng/ml TNF α and two doses of KA simultaneously for 24 hours.
31
Figure 2.4 OLs exhibit a dose-dependent response to KA, which is entirely blocked by the AMPA antagonist CNQX.
Cell death in OLs exposed to 100 µM KA in combination with 100 ng/ml TNF α was 19 ± 2 % as compared to 20 ± 2 % cell death with 100 µM KA alone, showing no effect of TNF α on a dose of KA that did not induce OL death by itself
(Fig. 2.6). However, 200 µM KA in combination with 100 ng/ml TNF α resulted in
49 ± 3 % OL death compared to 42 ± 2 % OL death when 200 µM KA was applied alone (Fig. 2.6). An ANOVA followed by Duncan’s post hoc revealed that this was a statistically significant increase in cell death due to TNF α (p < 0.05), indicating that a sublethal dose TNF α at 24 hours potentiated an intermediate 32
Figure 2.5 Optic Nerve OLs exhibit graded caspase activation in response to 300 µM KA. There was a main effect of KA on caspase activation over time.
dose of KA. Although the absolute magnitude of this difference appears to be
small, the partial eta squared effect size statistic revealed that the impact of
TNF α represented a very large effect ( η2 = 0.179).
DISCUSSION
Here we show that the glutamate receptor agonist KA is toxic to optic nerve OLs in a dose-dependent fashion, and that this toxicity can be blocked by the AMPA and KA receptor antagonist CNQX, as demonstrated previously by others (Matute et al., 1997). The toxic effect of KA is most likely through the activation of AMPA receptors and low affinity kainate receptors, as the doses
33
Figure 2.6 OL death in response to 200 µM KA is increased by 100 ng/ml TNF α. Data from groups with KA alone is taken from Fig. 2.3. Data from groups with TNF α was collected in the same experiments shown in Fig. 2.3 but shown separately here for clarity. The dashed line represents the levels of OL cell death under control conditions.
used in this study were well above those shown to mediate OL death
through high affinity KA receptors (Sanchez-Gomez and Matute, 1999). Many
dead OLs in our study exhibited an apoptotic morphology, with cell death
occurring over time and dead cells having fragmented and condensed nuclei.
Previously, OL death in response to excitotoxins has been described as both
apoptotic and necrotic in nature (Benjamins et al., 2003; Leuchtmann et al.,
2003). OLs in our laboratory exposed to 300 µM KA exhibited increasing
caspase-3 activation over 24 hours, confirming that there is an apoptotic
34 component to excitotoxic death of OLs. However, this finding does not exclude a necrotic component to OL excitotoxicity, especially at higher doses of KA.
We have shown for the first time that excitotoxic OL death can be directly potentiated by the pro-inflammatory cytokine TNF α. Though this effect is small in
magnitude, it was consistently observed. Moreover, the statistical effect size was
very large, indicating that the phenomenon was robust. The biological
significance of this effect may be important, as both TNF α and glutamate are
increased after CNS injury (McAdoo et al., 1999; Wang et al., 2002). Data from
other laboratories suggest that TNF α can affect OL response to excitotoxic stimuli. Pitt and colleagues (2003) have shown that TNF α can reduce the reuptake of glutamate by OLs, which could result in a increased vulnerability to excitotoxic stimuli. Additional in vitro data has shown that TNF α and interleukin-
1B can induce OL death via excitotoxicity in a mixed glial environment, indicating
a role for other cell types in mediating OL death in the injured CNS (Takahashi et
al., 2003). In vitro studies have shown that stressed OLs may attract microglia via a soluble mediator and that activated microglia may induce contact- dependent OL cell death (Nicholas et al., 2003). These data are supported by previous studies from our laboratory which have shown activated microglia in close proximity to apoptotic OLs after spinal cord injury (Shuman et al., 1997); however, whether dying OLs lead to microglial activation or microglial activation initiates OL death in vivo has yet to be resolved (Beattie et al., 2002b).
Additionally, data from other laboratories have shown that microglia play a role in
35
OL loss in models of periventricular leukomalacia (Haynes et al., 2003) and multiple sclerosis (Sriram and Rodriguez, 1997). The importance of cell-cell interactions in the damaged CNS raises the possibility that the effects of soluble
TNF α on OLs in culture do not accurately model the cell-cell interactions and
other events that occur in vivo (Peterson et al., 2002). It is possible that as
microglia become activated in CNS injury and disease, and migrate close to OLs,
that the local concentration of glutamate and cytokines in the space between OLs
and microglia is much higher than doses used in experimental applications.
Therefore, it is possible that current in vitro dosing strategies may actually
underestimate the effect of TNF α on OLs in vivo . It has been shown in in vitro experiments with mixed oligodendrocyte-microglia combined cultures, that membrane bound TNF α present on microglia is more important than soluble
TNF α in mediating cytotoxic effects on OLs (Zajicek et al., 1992). This could be
the case for other effects of TNF α as well, such as the effect we have shown here. We believe that there needs to be an increased focus on understanding the role of microglia in OL death and survival, as studies have demonstrated both potentially beneficial (Shaked et al., 2005; Nicholas et al., 2002) and harmful
(Arevalo-Martin et al., 2003; Benveniste, 1997) roles of these cells after CNS injury.
Our results reported here may seem inconsistent with a recent report from
Wosik and colleagues suggesting that TNF α does not affect the resistance of
human OLs to excitotoxicity (Wosik et al., 2004). In this study, human OLs were
36 found to be insensitive to excitotoxicity, and the addition of TNF α did not change this lack of response to TNF α. It should be emphasized that our study has
shown a subthreshold dose of KA (100 µM KA) did not become toxic in the
presence of TNF α, which is consistent with the observation of Wosik and colleagues. However, when we applied a suprathreshold dose of KA (200 µM
KA) in combination with TNF α to our rat optic nerve OL cultures, we did observe
potentiation of excitotoxic OL death by TNF α.
There are several mechanisms by which TNF α could potentiate OL excitotoxicity. It is possible that TNF α could initiate increased synthesis of calcium permeable glutamate receptors, leading to greater vulnerability of OLs to
KA. This could be accomplished by either a net increase in calcium-permeable glutamate receptors, or a change in subunit composition favoring more GluR2- lacking, and therefore calcium permeable, receptors without an overall change in receptor number, as seen in hippocampal neurons (Stellwagen et al., 2005). An increase in functional glutamate receptors in OLs could also occur via rapid membrane insertion of already-synthesized glutamate receptors as occurs in response to TNF α in neurons (Beattie et al., 2002a). Another possible
mechanism for TNF α’s potentiation of KA toxicity could be due to the intersection
of intracellular signaling cascades known to be common to TNF α-induced and excitotoxic cell death as described in Chapter 1. Both KA and TNF α have been
shown to induce caspase activation in OLs (Sanchez-Gomez et al., 2003; Ye and
D'Ercole, 1999) and it is possible that TNF α and KA binding to their respective
37 receptors induce separate signaling events that converge at one or more points in the apoptotic pathway, such as caspase-3 activation, which we have shown to occur in excitotoxic OL death. Both TNF α and excitotoxicity also induce oxidative stress in OLs. It may be that a sublethal dose of TNF α induces a level of oxidative stress that OLs can survive, but that when excitotoxicity has already overwhelmed OL antioxidant defenses OLs may become vulnerable to lower levels of TNF α-induced oxidative stress. Further work will be needed to determine the underlying mechanism of the interaction of TNF α and KA on OLs, and may lead to new therapeutic strategies to protect OLs after CNS injury.
Experiments studying the effects of TNF α and KA in combination on OL
precursor cells and on OLs in vivo would also generate insight into the biological
significance of this interaction.
Though TNF α has been shown in many studies, including this one, to
have a detrimental effect on OLs, the array of cytokines and other molecules OLs
are exposed to in CNS injury in vivo , is much more extensive than TNF α alone.
After CNS injury, microglia become activated and produce TNF α (Jana et al.,
2003), IL1-B (Zhao et al., 2006), and free radicals (Zhao et al., 2006; Li et al.,
2005; Dasgupta et al., 2002; Boje and Arora, 1992). Since microglia may be the
first immune cells to affect OLs after CNS injury, the cumulative effect of the
molecules they produce on OLs may be a determining factor as to whether OLs
survive or die as a result of the injury. Studies described in the following
chapters were undertaken to determine if microglia themselves would have an
38 effect on OL survival both under control conditions and in response to excitotoxicity.
39
CHAPTER 3
CHARACTERIZATION OF THE RESPONSE OF CORTICAL
OLIGODENDROCYTES TO KAINIC ACID EXCITOTOXICITY
Introduction
One of the barriers to understanding the response of OLs to excitotoxic injury has been the variety of cell culture systems used by different laboratories.
While excitotoxicity has been studied extensively in optic nerve OLs (Matute et al., 1997; Sanchez-Gomez and Matute, 1999; Alberdi et al., 2002; Sanchez-
Gomez et al., 2003; Soto et al., 2004; Miller et al., 2005; Ibarretxe et al., 2006),
OLs from cerebral cortex and corpus callosum have also been shown to be vulnerable to excitotoxicity (Yoshioka et al., 1995; McDonald et al., 1998a;
McDonald et al., 1998b). However, there are also reports showing cortical OLs to be resistant to excitotoxicity (Rosenberg et al., 2003; Wosik et al., 2004). This resistance may be due to cell source, growth media and supplements used, or time in culture. Often, individual laboratories focus on a single culture system without comparative studies between OLs from different sources. Additionally,
OLs within one laboratory are often grown in a single media formulation, which
40 may confound results, as a single growth factor can dramatically alter OL response to injury (Louis et al., 1993). The experiments described below were undertaken to characterize the response of cortical OLs to excitotoxicity under conditions similar to experiments conducted using optic nerve OLs. We hypothesized that OLs from cerebral cortex would display similar responses to
KA as our optic nerve OLs, based on the fact that OLs from different regions of the CNS are susceptible to excitotoxicity in vivo (Xu et al., 2001; Soto et al.,
2004; Xu et al., 2004). However, a difference in the response of cortical OLs versus optic nerve OLs is possible, and would emphasize regional differences in
OLs, as has been shown in developmental and toxilogical studies (Power et al.,
2002; Du et al., 2003; Stidworthy et al., 2003). Experiments were also carried out to determine if OLs at an earlier developmental stage in vitro and oligodendrocyte progenitor cells (OPCs) have a different excitotoxic threshold than more mature OLs, as has been reported by other laboratories (Itoh et al.,
2002; Jensen, 2002).
Another goal for conducting experiments with cortical OLs was to develop an OL culture system that would produce higher yields of cells that our previously described optic nerve culture system. The optic nerve culture system described in Chapter 2 generated less than 1x10 6 cells per dissection, precluding the use of molecular biology assays that require higher cell numbers. A higher yield of OLs would also allow more experimental groups to be analyzed in parallel, increasing efficiency and allowing for more replications per experimental preparation.
41
Lastly, we sought to develop a culture system in which OLs and microglia (MG) from the same source could be combined for experiments in this chapter and
Chapters 4 and 5.
While the previous study demonstrated that TNF α alone could make OLs more susceptible to excitotoxicity (Miller et al., 2005), the studies in this chapter were designed to address the question: what role do MG play in OLs response to excitotoxicity? MG not only secrete TNF α when activated (Jana et al., 2003) but can also secrete IL-1B (Zhao et al., 2006), which, along with TNF α, has been shown to decrease glutamate uptake by OLs (Pitt et al., 2003) and induce OL cell death that can be blocked by glutamate antagonists (Takahashi et al., 2003).
Additionally, microglia are able to produce free radicals, which can be directly toxic to OLs (Baud et al., 2004). This toxicity can occur via direct reaction between free radicals and intracellular components, leading to membrane peroxidation (Bernardo et al., 2003), DNA damage, or damage to proteins within the cell (Virag et al., 2003). However, free radicals do not need to directly damage cellular structures in order to be toxic to cells. Lower concentrations of free radicals can induce cell death by serving as intracellular messengers that induce caspase activation and apoptosis (Fragoso et al., 2004). The overall balance between free radicals and antioxidants may be a key regulator of cell survival as in OLs, the overall redox state of the cell can determine if a cell goes on to proliferate, differentiate, or die (Noble et al., 2003). Excitotoxic OL cell death has been shown to involve free radical production (Ibarretxe et al., 2006;
42
Sanchez-Gomez et al., 2003) and TNF α shifts the redox state of OLs towards a more oxidized state (Mayer and Noble, 1994), which favors cell death.
Therefore, it is possible that redox imbalance is a common pathway through which the effects of excitotoxicity, TNF α, and free radicals converge to mediate
OL cell death. Since MG can be activated by glutamate to secrete TNF α (Hagino et al., 2004), we hypothesized that OLs cultured in the presence of MG would be more susceptible to excitotoxicity.
The mixed cortical culture system first described by McCarthy and de
Vellis (1980) provides both OLs and MG from the same animals, thereby reducing the potential for results based on different genotypes of cell types within one study. Though other groups have used OL-MG combined cultures (Li et al.,
2005), the system in the present study was novel in that both OLs and MG were grown on the same coverslips to allow maximum opportunity for cell-cell interactions. This is important because membrane-bound TNF α produced by MG
preferentially effects TNF-R1, while soluble TNF α effects both TNF-R1 and TNF-
R2. These different receptors have different effects, with TNF-R1 activation
favoring cell death (see Chapter 1). We added MG to OL cultures 1 day after OL
plating (when OLs were already seen to have multiple processes) in order to
minimize any effects MG might have on OL development, as MG, astrocytes and
neurons have all been shown to affect OL development in vitro (Dutly and
Schwab, 1991; Nicholas et al., 2001).
43
We hypothesized that cortical OLs would be susceptible to KA-induced excitotoxicity similar to optic nerve OLs and that cortical OLs at less mature developmental stages would be more susceptible to excitotoxicity. Finally, we hypothesized that the presence of MG in culture with OLs would increase the vulnerability of OLs to excitotoxicity. Our results demonstrate that cortical oligodendrocytes are susceptible to similar doses of KA as optic nerve OLs. We found that media composition has a profound effect on OL response to excitotoxicity, as omission of thyroid hormone from the growth media increased
OL response to KA. Surprisingly, in our culture system, a dose of KA that induced cell death in more mature cortical OLs did not induce cell death in immature oligodendrocyte progenitor cells (OPCs). We also found that while the presence of MG in OL cultures did not alter OL response to excitotoxicity, the presence of MG decreased OL cell death overall. This finding was further investigated and data are reported in Chapter 4.
Materials and Methods
Cortical OL and OPC Cultures
Cortical OLs were isolated by a protocol similar to that described by
McCarthy and de Vellis (1980) with some modifications. Postnatal day 2 Long-
Evans rat pups were sacrificed by rapid decapitation, cleaned with 70 % ethanol and placed under a dissecting microscope with fiber optic illumination. Skin was removed by making a single midline cut with scissors followed by dissection with forceps. The skull was then cut along the sagital suture and the brain was lifted
44 from the skull base, cerebellum and olfactory bulbs were removed and meninges were dissected away with forceps. Brains were placed in media containing 7.1 mg/ml NaCl, 0.36 mg/ml KCl, 0.166 mg/ml KH 2PO 4, 257 mg/ml d-Glucose, 2.4
mg/ml NaHCO 3, 0.01 mg/ml phenol red, 0.9 mg/ml BSA Fraction V, and 0.33 mg/ml MgSO 4, (hereafter referred to as “dissection media”) and minced with a sterile scalpel blade. The solution was then spun at 300 rcf for 5 minutes in a sterile poly-propylene tube, the supernatant was aspirated and the tissue was resuspended with 10 mls of dissection media with 0.280 ng/ml trypsin (Sigma).
The tube was then placed vertically in an orbital shaker at 200 rpm for 13-15
o minutes, at 37 C. Following shaking, 10 mls of dissection media with 0.0128 mg/ml DNAse (Sigma) and 0.0832 mg/ml soy bean trypsin inhibitor (Sigma) was added and the tube was briefly shaken and centrifuged at 690 rcf for 5 minutes.
The supernatant was then removed and tissue resuspended in 3 ml of dissection media with 0.080 mg/ml DNAse and 0.520 mg/ml SBTI added. The solution was triturated 6 times with a 5 ml pipette and then triturated 15 times with a fire- polished glass pipette to obtain a single cell suspension. After trituration, 4 ml of dissection medium with 0.114 mg/ml MgCl 2 and 0.011 mg/ml CaCl 2 was added to
the single cell suspension, and the tube was spun for 5 minutes at 200 rcf. After
the supernatant was aspirated, cells were resuspended in DMEM with 10 % fetal
bovine serum containing 0.292 mg/ml L-glutamine (Sigma, G-5763), 1 mM
sodium pyruvate (Gibco, 11360-070), 0.04 mg/ml gentamicin (Gibco, 15750-060)
at a volume of approximately 5 ml per brain. The suspension was then plated
45 into T-75 flasks at 10 mls per flask. Flasks were kept in an incubator at 37 o C
and 5 % CO 2. Media was replaced after 24 hours in order to clear debris and dead cells that did not adhere to the bottom of the flask.
Once in culture, astrocytes began to proliferate across the bottom of the flasks with OPCs and MG growing on top of the astrocyte bed (Fig. 3.1). Cells were allowed to proliferate until confluent, at which time OPCs were isolated from the mixed cortical cultures of astrocytes, OPCs and MG, corresponding to day 9-
13 post dissection. In preparation for OPC isolation, MG were first removed by placing flasks on an orbital shaker at 200 rpm for 30 minutes to 2 hours. The orbital shaker was maintained at 37 oC and flasks were sealed with parafilm to
prevent gas exchange. After shaking, the supernatant was removed and
replaced with fresh DMEM with 10 % fetal bovine serum as described above and
returned to the incubator for 2 hours to allow the new media to achieve proper
temperature and pH. After equilibration, flasks were resealed and returned to the
shaker overnight (approximately 18 hours). At the completion of the overnight
shake, supernatant was removed and spun in a 50 ml polypropylene tube at 300
rcf for 5 minutes. The supernatant was then aspirated and cells were
resuspended in 4 ml of OL growth medium as defined in Chapter 2 (Miller et al.,
2005) unless otherwise stated, and triturated 15 times with a fire-polished glass
pipette. The cell suspension was then plated onto a sterile petri dish with a
radius of 2.5 mm and returned to the incubator for 45 minutes.
After 45 minutes, petri dishes were gently rinsed with growth media in
46
Figure 3.1 Shakeoff procedure for obtaining MG and OPCs/OLs from the same primary culture. Mixed cortical cultures were obtained and grown for 9-13 days and then shaken for 2 hours to remove MG. After adding fresh media, and allowing the mixed cortical culture to equilibrate in the incubator for 2 hours, the flask was shaken overnight which yielded OPCs that remained in the progenitor stage or differentiated into OLs depending on the media formulation used. After the overnight shake, the flask consisted of mostly astrocytes, which were not utilized for any of the experiments described here.
47 order to remove OPCs, but leave more tightly adherent MG behind. The resulting cell suspension was adjusted to a density of 20,000 cells per ml and plated on 12 mm glass coverslips in 4-well plates as described previously. For experiments utilizing OPCs, T3 and T4 were not present in the medium, and basic fibroblast growth factor (bFGF) and platelet-derived growth factor (PDGF) were included in the growth media, both at a concentration of 10 ng/ml. This growth factor combination has been shown previously to maintain OPC in an immature stage in vitro (Bogler et al., 1990; Engel and Wolswijk, 1996).
MG Isolation and Addition to OL Cultures
In selected experiments, MG were added to OLs on day 1 in vitro . MG
were harvested from flasks derived from the same primary dissections as the
OPCs or OLs with which they were combined. To remove MG from flasks
without removing OPCs, T75 flasks were tapped on the lab bench and the
supernatant removed. MG were resuspended in the same medium as the OLs
with which they were combined. When plated alone, the resuspended cells were
shown to be > 95 % pure MG. MG were plated onto pre-existing OL cultures at a
ratio of 1:1 by initially resuspending MG at twice the desired concentration,
removing 250 ul of media from the OPC or OL cultures and adding 250 ul of MG
suspension to OPC or OL cultures. OL cultures not receiving MG had half of their
media replaced to control for possible effects of fresh media.
Immunocytochemistry
At the conclusion of all experiments, cells were fixed with 2 %
48 formaldehyde prepared from paraformaldehyde. All primary antibodies were purchased from Chemicon (Temecula, CA) unless otherwise noted. All secondary antibodies were purchased from Molecular Probes (Carlsbad, CA).
Antibodies were diluted in Hank’s Buffered Salt Solution with 5 % donor calf serum and coverslips were rinsed twice between all antibody applications. For
NG2 labeling (a marker for immature and dividing OPCs), cells were first permeabilized with 0.1 % Triton for 10 minutes, and then incubated with anti-NG2 antibody at a concentration of 1:400 for 1.5 hours, followed by goat anti-rabbit
Alexa 594 at 1:100 for 1 hour. For A2B5 labeling (a marker for OPCs), cells were incubated with an antibody to A2B5 conjugated to Alexa 488 (Chemicon) at
1:100 for 1.5 hours. For GalC labeling (a marker of more mature OLs), cells were incubated with an antibody to GalC at 1:400 for 1 hour followed by goat anti-mouse Alexa 488 at 1:200 for 1 hour. For myelin basic protein (MBP) labeling (a marker for mature, myelinating OLs), cells were first permeabilized with 0.1 % Triton for 10 minutes and incubated with an antibody against MBP at
1:200 for 1 hour followed by goat anti-rabbit Alexa 488 at 1:100 for 1 hour. For
IB4 labeling (used to label MG), which was completed following labeling for GalC, cells were incubated with Alexa 594-conjugated IB4 (Molecular Probes) at 1:100 for 30 minutes in a solution of HBSS with 1 mM calcium and no serum. For all experiments, cells were incubated with 10 ug/ml Hoechst for 5 minutes, rinsed with H 2O and mounted on glass slides with Immu-Mount (Thermo, Pittsburgh,
PA) after antibody applications.
49
Cell counts were performed blind to experimental conditions, using a Zeiss
Axioplan 2 microscope (Carl Zeiss, Germany). Counts were conducted by evaluating all cells within the microscope reticule over the entire diameter of each coverslip. For mature OLs, both live and dead cells were visible, with live cells having intact nuclei and cell bodies, and dead cells having absent or fragmented nuclei and fragmented cell bodies (Fig. 3.2 A&B). Counts of both live and dead
OLs were performed and values are expressed as percentages of live cells relative to control. Live OPCs were defined as A2B5 positive cells with two or more intact processes and intact nuclei. It was not possible reliably quantify dead OPCs, as cell bodies were often absent and it could not be determined if processes left behind were part of the same of different cells (Fig 3.2 C&D).
Therefore, for OPCs, values were expressed raw number of live cells counted.
Experiments were analyzed via ANOVA followed by Tukey’s post hoc test unless otherwise stated and values are expressed as percent of control ± SEM.
Statistical significance was set at p < 0.05. For all experiments evaluating
OPC/OL survival, cells were assessed at 3 days in vitro .
Caspase Assay A commercially available caspase assay kit (FLICA kit,
Immunohistochemistry Technologies, Bloomington, Minnesota) that utilizes a
fluorescently-tagged poly-caspase inhibitor was used in accordance with the
manufacturer’s recommendations. Caspase positive OLs were defined as cells
50
Figure 3.2 Comparison of live and dead cortical OLs and OPCs. (A) Live OLs are seen to have multiple highly branched processes and intact Hoechst-labeled nuclei. (B) Dead OLs are seen to have fragmented cell bodies and fragmented or absent nuclei. (C) Live OPCs are seen to have two or more intact processes and intact nuclei. (D) Dead OPCs are seen as fragments of processes without clearly visible cell bodies, preventing accurate quantification of dead cells.
51 with fragmented nuclei that also labeled brightly with the caspase detection kit, so that the cell body and processes were visible to confirm OL morphology.
Cells counts were performed as described above, and the number of caspase positive OLs was recorded.
Lactate Dehydrogenase Release Assay for Assessment of OPC Death A commercially available lactate dehydrogenase (LDH) assay kit (Roche,
Indianapolis, IN) was used to quantify LDH release according to the manufacturer’s instructions. For positive controls, OPCs were treated with 1 % triton for 10 minutes prior to LDH release quantification. Cell death release was expressed as experimental values minus control values as a percent of positive control values minus negative control values (% cell death = ( experimental value
– neg. control ) / ( pos. control – neg. control )).
Results
Cortical OLs are Susceptible to KA in a Dose-Dependent Manner
Cortical OLs exhibited a dose-dependent response to KA (Fig 3.3), with
100 µM KA having no significant effect on OL death, and 150 µM and 200 µM significantly increasing OL cell death to 55 ± 4 % and 72 ± 5 % respectively.
1mM KA induced 91 ± 3 % OL death, which was completely blocked by 30 µM
CNQX, demonstrating that KA toxicity in cortical OLs is receptor dependent as in optic nerve OLs.
Cortical OLs Grown Without Thyroid Hormone do not Exhibit a Different
Phenotype but are More Vulnerable to KA Excitotoxicity
52
It has been shown that OLs at immature stages of development are more susceptible to excitotoxicity than OLs at mature development stages (Rosenberg et al., 2003). This in vitro data is supported by in vivo data that showing
developing OLs to have a greater number of glutamate receptors and are more
susceptible to excitotoxicity than mature OLs (Jensen, 2002) . Thyroid hormone acts on OPCs and OLs in the developing nervous system to induce myelination, and levels of T3 peak during the period in which brain myelination occurs
(Ahlgren et al., 1997; Rodriguez-Pena, 1999). Experiments were performed to determine if OLs grown in media without T3 and T4 (TH), which speed OPC development to mature OLs (Baas et al., 1997; Gao et al., 1998), would be more sensitive to KA toxicity than mature OLs.
OLs were isolated as described above, except for the substitution of TH- free growth media for the initial growth media for all cells. Two hours after plating, TH-free media was replaced with growth media containing TH for selected wells, and all other wells had TH-free growth media replaced with fresh
TH-free growth media to control for possible effects of fresh media. Addition of drugs to TH-treated and TH-free OLs was carried out simultaneously at day 2 in vitro . OLs in both media conditions were fixed at day 3 for labeling with antibodies to NG2, A2B5, GalC and MBP to asses developmental stage.
OLs cultured with and without TH were seen to rapidly acquire a highly branched phenotype characteristic of mature OLs in vitro . Upon fixation and immunocytochemical analysis on day 3 in vitro , both OLs grown with and without
53
Figure 3.3 Cortical OLs exhibit a dose-dependent response to KA excitotoxicity. 150 µM, 200 µM and 1 mM KA all induced significant cell death compared to controls. The effect of 1 mM KA was completely blocked by 30 µM CNQX.
TH exhibited the same profile of antigen labeling. In both conditions, cells were completely negative for NG2 and there was faint A2B5 labeling. In both conditions, all cells labeled strongly for GalC and MBP throughout their processes (Fig. 3.4).
OLs cultured without TH did not differ significantly from OLs cultured with
TH in amount of cell death under control conditions (Fig. 3.5, 39 ± 3 % cell death for OLs with TH, 45 ± 8 % for OLs without TH). However, OLs without TH were significantly more susceptible to KA toxicity than OLs with TH, undergoing 80 ± 2 54
% cell death after exposure to 100 µM KA and 95 ± 2 % cell death in response to
200 µM KA. In contrast, OLs cultured with TH were not susceptible to 100 µM
KA (43 ± 4 % cell death, not significant from control), and exhibited 72 ± 6 % cell death in response to 200 µM KA. Factorial ANOVA confirmed an interaction of
KA and TH (p = 0.016) indicating that TH affected OL response to KA.
Figure 3.4 OLs grown with and without thyroid hormone exhibit similar morphology at 3 days in vitro . OLs with and without TH labeled for GalC and MBP but did not label for A2B5.
OL Progenitor Cells are Less Susceptible to KA-induced Excitotoxicity than
Mature OLs
55
Experiments were conducted on OPCs to determine if these cells were more vulnerable to excitotoxicity than mature OLs, as has been shown by other laboratories (Volpe, 2001; Rosenberg et al., 2003). OPCs were prepared as described above, and exposed to 200 µM KA from day 2 to day 3 in vitro. In the presence of 200 µM KA, OPC viability was reduced from an average of 87 ± 16 cells per coverslip, to an average of 56 ± 16 cells per coverslip, a 36 % reduction
Figure 3.5 OLs with and without TH have different susceptibilities to KA toxicity. OLs with and without TH exhibited similar levels of cell death in control conditions. In response to 100 µM KA, OLs with TH did not undergo cell death, however without TH OLs underwent a significant amount of cell death. In response to 200 µM KA, OLs with and without TH both underwent cell death, however OLs without TH underwent cell death to a significantly higher degree (* = p < 0.05, Tukey’s post hoc).
56
in cell number. However, this difference did not reach statistical significance
(Fig. 3.6).
To confirm this null finding, a LDH assay was conducted. Cells with disrupted membranes release LDH which can be quantified and compared to positive controls to estimate the percentage of dead cells as described in materials and methods . 200 µM KA resulted in 41 ± 17 % cell death, however,
this was not significantly different from the control condition (Fig. 3.7). Though
both of these experiments showed a trend towards OPC death after exposure to
200 µM KA, no additional experiments were performed. In experiments with
Figure 3.6 Counts were used to determine the number of OPCs after treatment with 200 µM KA. The difference between controls and OPCs treated with 200 µM KA did not reach statistical significance (n=4 for each group).
57 mature cortical OLs, the effects of 200 µM KA on cell death attained statistical significance with n=4 in each group. In addition, both OPC count data and LDH data supported the same conclusion (and showed similar magnitudes of OPC death), therefore, we chose to conduct further experiments studying excitotoxicity only on mature OLs.
Time, but not KA, Induces Caspase Activation in OLs.
Several studies have shown excitotoxicity in OLs to have an apoptotic component that is mediated by caspase enzymes (Benjamins et al., 2003;
Sanchez-Gomez et al., 2003; Miller et al., 2005). To determine if
Figure 3.7 LDH release as percent of positive controls. There was no effect of 200 µM KA on OPC LDH release (n=5 for each group).
58 excitotoxicity induces apoptosis in mature cortical OLs, caspase activation was evaluated over time after exposing OLs to 200 µM KA. Caspase activation was evaluated using a commercially evaluated caspase detection kit as described in materials and methods. Caspase positive OLs were defined as cells that were labeled with the caspase indicator brightly to identify their typical OL morphology, and also exhibited fragmented nuclei (Fig. 3.8). KA was administered at day 2 in vitro as in previous experiments. Caspase activation was quantified at 30 minutes, 6 hours and 30 minutes, and 24 hours after KA addition, corresponding to 2 days in vitro (DIV), 2.25 DIV, and 3 DIV (Fig. 3.9). At 2 DIV, 15 ± 2 caspase positive OLs were observed in the control condition, and 19 ± 1 caspase positive
OLs were observed after treatment with 200 µM KA. At 2.25 DIV, 23 ± 3
Figure 3.8 A typical example of a caspase positive OL. Caspase positive OLs were defined as cells which had both fragmented nuclei and were labeled sufficiently brightly with the FLICA kit for OL morphology to be confirmed.
59
Figure 3.9 Time, but not 200 µM KA, induces caspase activation in OLs. OLs were assayed for caspase activation on day 2, day 2 + 6 hours, and day 3 in vitro . There was a significant main effect of time (p < 0.005), but not KA on caspase activation.
caspase positive OLs were observed in the control condition and 26 ± 2 caspase positive OLs were observed after treatment with 200 µM KA. At 3 DIV, 35 ± 4 caspase positive OLs were observed in the control condition and 32 ± 8 caspase positive OLs were observed after treatment with 200 µM KA. Factorial ANOVA revealed a main effect of time (p = 0.004), but not KA on caspase activation in
OLs.
Microglia do not Alter OL Response to Excitotoxicity, but Reduce OL Cell
Death Overall in vitro .
60
MG have been hypothesized to play a detrimental role in CNS trauma, although studies have shown that activated MG have the ability to play a protective as well as detrimental role in OL survival depending on the activating stimulus (Butovsky et al., 2006). It has been shown that MG respond quickly to
KA, by reorganizing actin filaments and becoming motile (Christensen et al.,
2006) and that MG can release glutamate when activated (Barger and Basile,
2001). We hypothesized that OLs cultured with MG would be more susceptible to excitotoxicity than OLs alone due to MG activation and release of molecules such as glutamate, nitric oxide and TNF α which have all been shown to be toxic to OLs in vitro (Selmaj and Raine, 1988; Pang et al., 2000; Ibarretxe et al., 2006) and can all induce common intracellular death pathways as described in Chapter
1. Though MG have been directly shown to produce TNF α in response to glutamate receptor activation (Hagino et al., 2004), it is not known if MG produce other inflammatory cytokines and free radicals after glutamate receptor activation
(Farber and Kettenmann, 2005).
MG were added to OL cultures as described in methods . Successful MG transfer to OL cultures was verified using IB4 labeling (Fig. 3.10). 200 µM KA was administered to OLs alone and to OLs that were grown in combination with
MG (Fig. 3.11). As in previous experiments, 200 µM KA induced a significant amount of cell death in mature OLs (Fig. 3.11). 35 ± 1 % of OLs were scored as dead in control conditions and 25 ± 2 % of OLs were scored as dead in the presence of MG alone. 200 µM KA induced 72 ± 5 % OL death without MG
61 added and 61 ± 3 % OL cell death in the presence of MG. Contrary to our hypothesis, there was no interaction between MG and KA on OL cell death.
However, there was a significant main effect of MG increasing OL survival, as
revealed by factorial ANOVA (p = 0.002). It was hypothesized that MG protect
OLs by reducing caspase activation, as OL caspase activation occurs over time
in vitro but not in response to KA, and MG protected OLs from cell death overall,
Figure 3.10 (A) Cortical OLs grown without MG labeled with GalC on day 3 in vitro . (B) Cortical OLs with MG that were added on day 2 in vitro labeled with IB4 (red).
but not from KA. This hypothesis was directly tested in experiments described in
the Chapter 4.
Discussion
Numerous reports have shown OLs to be susceptible to excitotoxicity in
vitro (McDonald et al., 1998b; Miller et al., 2005; Ibarretxe et al., 2006). This
62 excitotoxic cell death has been shown in response to glutamate itself, and the glutamate receptor agonist KA. However, not all laboratories have shown OLs to be susceptible to excitotoxicity (Rosenberg et al., 2003; Wosik et al., 2004). This could be due to different culture conditions used across laboratories, different cell source within the CNS, time in culture, or different animal source of OLs. These
Figure 3.11 MG do not alter OL response to excitotoxicity but do reduce OL cell death overall (p = 0.002, factorial ANOVA).
experiments were conduced to verify that under our culture conditions, cells from
different regions of the CNS would be vulnerable to excitotoxicity. Another
reason for these experiments was to develop a culture system of cortical OLs 63 and MG from the same source in order to conduct the experiments examining the interactions between these two cell types. The cortical OL and MG culture system described here holds several advantages over the optic nerve OL culture system described in Chapter 2 and Miller et al., 2005. The larger volume of tissue available from the initial dissection and the proliferation of the initial cultures in T75 flasks before OL isolation results in a much higher yield of cells, allowing for more groups per experiment. Higher yields of cells also allowed the use of a LDH assay which preliminary experiments showed to be more sensitive when OLs were plated at higher densities (data not shown).
In our culture system, cortical OLs exhibited a similar response to excitotoxicity as optic nerve OLs described in the Chapter 2. Both cortical and optic nerve OLs were responsive to KA at doses of 20 µM and above, and the effect of 1 mM KA was able to be completely blocked by 30 µM CNQX, as has been shown in other studies of OL excitotoxicity (Matute et al., 1997).
Media composition may contribute to differing results in OL excitotoxicity studies. DMEM Sato is a commonly used medium for growing OLs, though some researchers use formulations such as F12 (Liu et al., 2002) and media containing serum (Wosik et al., 2004). Thyroid hormone is often present in growth media in order to drive OLs quickly to maturation. However, it has been documented that OLs will mature in vitro if OLs are grown in serum free media without growth factors (Raff et al., 1983). Sera has been shown to alter the development of OLs (Raff et al., 1983), and a preliminary experiment we
64 conduced verified this to be the case, as OLs cultured with even 1 % sera had a morphology resembling type 2 astrocytes (data not shown), as described by Raff and colleagues (1983), and were insensitive to excitotoxicity, as has been shown for astrocytes (Yamaya et al., 2002).
We conducted an experiment testing the response of OLs grown without
TH to KA excitotoxicity with the hypothesis that OLs cultured without TH would mature slower and be more susceptible to excitotoxicity. Though many studies have shown OLs at earlier developmental stages to be more vulnerable to excitotoxicity, no study to our knowledge has directly tested the effect of TH alone on OL excitotoxicity. We expected the omission of thyroid hormone from the growth media to be an effective way to compare OLs at different developmental stages without extending time in culture (which, as we demonstrated, can increase caspase activation). We expected OLs cultured without TH to mature at a slower rate in vitro than OLs with TH, based on prior studies reporting TH to increase the rate of OL maturation (Baas et al., 1997).
Cortical OLs in our experiments were observed to take on the multipolar morphology of mature OLs at day 1 in vitro , (more rapidly than optic nerve OLs which typically appeared to be in an OPC developmental stage until day 2 in vitro ), and we expected to observe a delay in OL maturation in the absence of
TH. Surprisingly, we did not observe a qualitative difference in morphology or
antigenicity of OLs grown with or without thyroid hormone. Though we assayed
for several development markers of OLs, it is possible that there were
65 developmental differences to which our labeling paradigm was not sensitive. The system of antigen labeling to describe OL developmental stage has been described as “arbitrary and overlapping,” (McDonald et al., 1998b) an opinion with which we agree, as our own experience has shown OL developmental markers to overlap in cells with clearly different OPC/OL phenotypes (see
Chapter 4). There are other characteristics used to define OL maturation stage, such as specificity to the antibodies O1 and O4 (Sommer and Schachner, 1981;
Bansal et al., 1989; Reynolds and Hardy, 1997), expression of the transcription factors Olig1, Olig2, and the recently described cytoskeletal protein, Ermin
(Brockschnieder et al., 2006). Though OLs grown with and without TH exhibited the same morphology and immunolabeling profile, there are more subtle changes that occur as OLs mature and these may have contributed to the higher excitotoxic threshold of TH-cultured OLs. As OLs mature, they decrease the expression of AMPA receptors that mediate excitotoxicity (Rosenberg et al.,
2003). It is possible that the TH treated cells, while not grossly more mature than cells without TH treatment, had less AMPA receptors and therefore a higher excitotoxic threshold. Since excitotoxicity has been shown to involve free radical formation in both OLs and neurons, it is also possible that the TH-treated OLs had developed more glutathione stores, which is a characteristic of more mature
OLs, along with increased resistance against oxidative stress (Back et al., 1998;
Baud et al., 2004). It is also possible that TH has pro-survival effects on OLs independent of what have been described as developmental changes. One
66 study has demonstrated that TH can reduce caspase activation in OLs that occurs over time in culture (Jones et al., 2003). Though our results showed no main effect of excitotoxicity on caspase activation, it is possible that a decreased basal level of caspase activation could help protect OLs from excitotoxicity.
Further investigation into the protective effects of TH would be useful to determine if TH has potential for use in treatment of white matter diseases, as has been shown with other hormones, including estrogen and progesterone
(Thomas et al., 1999; Cantarella et al., 2004; De Nicola et al., 2006).
After detecting a pharmacologic, but not morphologic difference in OLs grown with and with out TH, we investigated the response of OPCs to KA excitotoxicity. As stated previously, many reports have indicated OPCs to be more susceptible than OLs to a variety of insults. Our two experiments testing the effects of 200 µM KA on OPCs (a dose that induced cell death in both optic nerve and cortical OLs) yielded surprising results. There was no statistically significant effect of 200 µM KA on OPCs, in contrast to our results showing mature OLs to be vulnerable to this dose of KA. Though several reports have shown OLs at earlier developmental stages in vitro and in vivo to be more
sensitive to excitotoxicity than mature OLs, this may be an oversimplified view.
One report (Itoh et al., 2002) has described a more dynamic change in glutamate
receptor expression in OLs in vitro . Itoh and colleagues (2002) observed a peak in glutamate receptor expression early in OL development. However, OPCs at the earliest stages examined actually had lower functional glutamate receptor
67 levels than mature OLs. This has also been described in vivo , with a temporary
peak in glutamate receptor expression corresponding with increased excitotoxic
vulnerability (Jensen, 2002). It is possible that the OPCs in our culture system
were of this earliest developmental stage and had not undergone this transient
receptor peak. OPCs in our experiments were NG2 positive (see Chapter 4),
which is a marker of the most immature stage of OPCs, which Itoh and
colleagues found to have low levels of calcium-permeable AMPA receptors.
Further study of our OPCs would be needed to determine the mechanism of their
relative resistance to excitotoxicity.
While typically thought of as more fragile than OLs, OPCs have been shown to better tolerate damage induced by complement proteins (Scolding et al., 1990). Recently, it has been shown that sublethal doses of glutamate can sensitize OLs to complement-mediated cell death (Alberdi et al., 2006). This effect was mediated through oxidative stress, as it could be blocked by the antioxidant Trolox, and could be replicated by inducing oxidative stress without glutamate receptor activation. This provides further evidence that oxidative stress could be a cellular “switch” that predisposes OLs to cell death in response to several different stimuli.
After CNS injury, OLs and OPCs are exposed to a wide range of injurious stimuli including excitotoxins, proinflammatory cytokines, free radicals, activated complement proteins and perhaps autoantibodies (Rebhun and Botvin, 1980;
McAdoo et al., 1999; Hayes et al., 2002; Ankeny et al., 2006). After spinal cord
68 injury, there are distinct time courses of both OL death and OPC proliferation. In rats, OL apoptosis distant from the lesion center peaks 8 days after injury, and is sustained at 3 weeks post injury (Crowe et al., 1997). OPC proliferation is elevated from 1 to 4 weeks post SCI (McTigue et al., 2001) and is also seen from
1 to 2 weeks after ischemic injury in the rat brain (Tanaka et al., 2003).
Protecting proliferating OPCs from cell death may allow the CNS to better repair itself after injury by generating new OLs from this expanding OPC population, which has been shown to occur in vivo after demyelination (Watanabe et al.,
2002). In order to both minimize OL cell death and maximize OPC replacement
of lost OLs, it may be necessary to separately target specific biochemical
pathways in order to protect these distinct cell types when they are each most
vulnerable. Our results indicate that blocking excitotoxicity may confer benefit
during time periods where OLs are dying, but not when OPCs are proliferating. A
better understanding of what factors contribute to OPC and OL loss at distinct
time periods would help in the development of time-specific treatments for SCI
and other neurological injuries.
Our results showing OL caspase activation increasing over time are not
surprising. In vitro isolation is stressful for cells and the serum free media used in our experiments is not designed for prolonged time in culture. Serum supplementation has been shown to increase the lifespan of OLs in vitro (Louis et al., 1993), however we chose to exclude sera from experiments due to the unknown milieu of growth factors, complement proteins and other substances
69 that may be present. Others have shown OLs to undergo cell death after isolation from mixed cortical cultures, and that cell death varied by OL plating density and media composition (Yang et al., 2005b). However, a pilot experiment in our laboratory did not reveal a difference in OL survival when OLs were plated at 5,000 or 20,000 cells per coverslip (data not shown). Others have described OL death in vitro at levels greater than 50% in control conditions
(Nicholas et al., 2002), so it was not surprising that OLs in our system underwent apoptosis over time in culture. Unfortunately, many reports in the literature do not disclose the net number of dead cells in control conditions, further complicating the comparison of culture conditions across laboratories.
We were surprised not to observe caspase activation in response to KA- induced excitotoxicity. Excitotoxic cell death has been shown to be both necrotic and apoptotic in nature, depending on the strength of the stimuli (Cheung et al.,
1998). 200 µM KA may be a relatively high dose of KA for our cortical OLs, inducing 72 ± 5 % cell death and therefore may have induced necrosis rather than apoptosis. Perhaps a lower dose of KA would have induced caspase activation as in optic nerve OLs (See Chapter 2). However, the fact that 200 µM
KA did not induce OL caspase activation, gave us an important tool to contrast caspase-dependent and caspase-independent OL death. Using 200 µl KA as a model of necrotic OL cell death and time in culture as a model of apoptotic OL death gave us insight into the mechanism of MG-mediated OL survival. We had initially hypothesized that the presence of MG in combined culture with OLs
70 would make OLs more susceptible to excitotoxic cell death. However, our experiment revealed that MG had no effect on the response of OLs to 200 µM KA and in fact reduced OL cell death overall. This suggests that while MG do not alter the response of OLs to a necrotic insult, MG are able to protect OLs from a more gradual apoptotic insult. We directly tested this hypothesis in the Chapter
4.
In summary, OL cell death occurs in vivo in a number of neurological diseases and trauma. Numerous in vitro models have been developed to further
understand the mechanism of OL cell death and to develop protective strategies
that can be translated into clinical therapies. In order for in vitro studies to have
relevance in vivo , there needs to be convergent evidence from studies that use a
variety of conditions and experimental paradigms. We have developed a cell
culture system of cortical OLs and MG that allows for the isolation of both cell
types alone and in combination. OLs in this system are vulnerable to KA-induced
excitotoxicity under similar conditions as optic nerve OLs. There is an effect of
media composition and cell developmental stage on OL/OPC response to KA,
with thyroid hormone reducing OL susceptibility to KA and OPCs being
comparatively resistant to a 200 µM dose of KA. This indicates that therapeutic
strategies that may protect mature OLs that die in SCI may not have the same
beneficial effects on proliferating OPCs after injury. We observed a beneficial
effect of MG on OL survival, possibly by reducing apoptosis that occurs over time
in culture. This finding suggests that MG may have the ability to promote OL
71 survival in conditions in which OLs undergo apoptosis and that MG due not play an exclusively negative role in CNS injury. Our combined OL-MG culture system was utilized to further study interactions between activated MG and OPCs/OLs as described in Chapter 4.
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CHAPTER 4
THE EFFECT OF LPS-ACTIVATED MG ON OLIGODENDROCYTE
PROGENITOR CELLS AND MATURE OLIGODENDROCYTES
Introduction
Though the data described above show that MG do not affect excitotoxicity in mature OLs, there are numerous reports of activated MG having deleterious effects on OPCs (Pang et al., 2000; Li et al., 2005). When activated,
MG can produce a wide array of cytokines, free radicals, glutamate and other molecules that are toxic to OL lineage cells. It has been shown that OPCs are more sensitive to many insults than more mature OLs, including excitotoxicity
(Rosenberg et al., 2003), glutathione depletion (Back et al., 1998), TNF α (Pang et al., 2005), and radiation injury (Fukuda et al., 2005). Therefore, it is possible that while MG can protect OLs from cell death, they may induce cell death in
OPCs.
In the previous study, we expected to observe an increase in OL cell death in the presence of MG. Previous data from our laboratory had shown MG to be rapidly activated by KA and to assume a phenotype typical of activated MG in vivo (Christensen et al., 2006) . However, we did not observe any changes in
73
OL cell death in response to a mid-range dose of KA in the presence of MG.
Since MG have been hypothesized to contribute to OL and OPC death in vivo ,
we further explored the effects of activated MG on OL/OPC survival in vitro in
order to better study the interactions of these two cell types.
Previous studies have been conducted with OPCs in culture with MG, and
have demonstrated that activated MG lead to OPC death that is mediated by free
radical production and dependent on toll-like receptor signaling in MG (Lehnardt
et al., 2002). In these studies, lipopolysaccharide (LPS) was used as an
activator of MG. LPS is isolated from the cell wall of gram-negative bacteria, and
has been shown in vivo and in vitro to be a potent activator of macrophages and microglia, inducing TNF α and free radical production (Zhao et al., 2006). We
used a TNF α ELISA to confirm that LPS induced TNF α production by MG in our culture system, and also to determine if the same was true for KA, which we hypothesized would also induce MG TNF α production. Though there are very
few studies examining the intracellular signaling cascades induced by glutamate
and glutamate agonists on MG (Hagino et al., 2004; Farber and Kettenmann,
2005), it has been shown that glutamate agonists can induce the phosphorylation
and activation of the transcription factor CREB in OLs (Deng et al., 2003) and
that CREB activation is associated with TNF α production by MG (Min et al.,
2003).
After confirming that LPS induced TNF α production by MG in our culture
system, we proceeded to study the effect of LPS-activated MG on OPCs and
74
OLs. We hypothesized that LPS-activated, TNF α producing MG would induce cell death in OPCs, as has been shown by others (Lehnardt et al., 2002). We further hypothesized that LPS-activated MG would also induce cell death in mature OLs, which has not been studied previously. MG have been implicated in inducing both OPC and mature OL death in CNS injury, but it is not clear if mature OLs respond to activated MG in the same manner as OPCs. The present study demonstrates that LPS, but not KA, induces rapid TNF α production by MG, and that LPS-activated MG reduce survival of OPCs. We also show that both
MG alone and LPS-activated MG decrease caspase activation and increase survival of mature OLs. This surprising observation indicates that MG can have divergent effects on OL lineage cells depending on OL developmental stage. It also emphasizes that while activated MG can have deleterious effects on some cells within the CNS, MG activation does not necessarily eliminate all beneficial effects of MG on other CNS cells. Taken together with Chapter 3, these data show that while mature OLs are vulnerable to excitotoxicity, OPCs are vulnerable to MG-mediated injury and different therapeutic strategies may be needed to protect OPCs and mature OLs after CNS injury.
Materials and Methods
OPC and Mature OL Culture
OPCs were isolated from mixed cortical cultures in the same manner as
described in Chapter 3. To keep OPCs in an immature stage, bFGF and PDGF
were both present in the media at concentrations of 10 ng/ml throughout the
75 experiment as described above. To induce maturation of OPCs into mature OLs, thyroid hormone was used instead of bFGF/PDGF. MG were added to OPC/OL cultures one day after OPC/OL plating as described in Chapter 3.
Immunocytochemistry for OL Lineage Markers
Immunocytochemistry for GalC (mature OLs) and A2B5 (OPCs) was performed as described above. For NG2 labeling (OPCs), cells were first permeabilized with 0.1 % Triton for 10 minutes, and then incubated with anti-NG2 antibody at a concentration of 1:400 for 1.5 hours, followed by goat anti-rabbit
Alexa 594 at 1:100 for 1 hour. For MBP labeling (mature OLs), cells were permeabilized with 0.1 % Triton for 10 minutes and incubated with MBP antibody at 1:200 for 1 hour followed by goat anti-rabbit Alexa 488 at 1:100 for 1 hour.
Cell counts for OPC Survival
OPC survival was assessed as described in Chapter 3 (only live OPCs were counted) and is reported here as OPC survival relative to the control condition (OPCs alone).
Cell Counts for OL Survival
OL survival was assessed as described in Chapter 3 (live and dead OLs were counted), and is reported here as OL survival relative to the control condition (OLs alone).
Cell Proliferation Assay A BrdU incorporation assay, similar to that employed by Kondo and Raff
(2000) was used to asses OPC and OL proliferation. Cells were exposed to 20
µM BrdU (Sigma) for six hours prior to fixation, and then fixed in 2 % 76 formaldehyde as previously described. Cells were post fixed in 100 % methanol for 10 minutes at -20˚C then treated followed by DNA denaturation with 2 M hydrochloric acid for 30 minutes. Acid was then neutralized with 0.1 M sodium borate at pH of 8.5 for 10 minutes. Cells were then permeabilized with 0.1 %
Triton for 10 minutes and incubated with primary antibody against BrdU
(hybridoma monoclonal antibody developed by Stephen J. Kaufman,
Developmental Studies Hybridoma Bank, The University of Iowa, Department of
Biological Sciences, Iowa City, IA) at 1:5 for 1 hour followed by goat anti-mouse secondary antibody conjugated to Alexa 594 at 1:100 for 1 hour. This labeling was done in combination with A2B5 labeling for OPCs and followed by Hoechst labeling of nuclei. BrdU labeling was evaluated by examining five random fields per coverslip and the percentage of OPCs or OLs incorporating BrdU was compared to that of the control condition.
Caspase Assay A commercially available caspase assay kit (FLICA kit, Immunochemistry
Technologies, Bloomington, Minnesota) that utilizes a fluorescently-tagged poly caspase inhibitor was used in accordance with the manufacturer’s recommendations. Caspase positive OPCs/OLs were defined as cells with fragmented nuclei that also labeled brightly with the caspase detection kit, so that the cell body and processes were visible to confirm OPC/OL morphology. Cells counts were performed as described above, and caspase activation values were expressed as percent of control.
Statistical Analysis 77
Experiments were analyzed via factorial ANOVA or ANOVA followed by
Tukey’s post hoc test unless otherwise stated and values are expressed as percent of control ± SEM. Significance was set at p < 0.05.
Results
LPS, but not KA, Induces TNF α Production by MG
To confirm that LPS induces MG to produce TNF α, as has been shown by
others (Strohmeyer et al., 2005; Qin et al., 2005), we conducted a TNF α ELISA assay. We also tested if KA induced TNF α production, as we have previously
observed KA to induce morphological changes in MG consistent with an
activated phenotype (Christensen et al., 2006). At 4 hours, 1 ng/ml LPS, but not
Figure 4.1 After 4 hours, MG produce TNF α in response to LPS but not KA. 1 ng/ml LPS produced 386 ± 76 pg/ml TNF α which was significantly different from control (p < 0.05, Tukey’s post hoc).
78
1 mM KA, induced TNF α production (Fig. 4.1). This confirms that MG in our culture system respond to LPS, but that KA may not induce the same type or extent of MG activation that is induced by LPS. A previous study demonstrating
MG TNF α release in response to 1 mM KA for 3 hours showed a significant
release of TNF α on the order of 25 pg/ml (Hagino et al., 2004). This result is very similar to our result of 35 pg/ml after 4 hours exposure to 1mM KA.
Our results did not show this level to be statistically significant because our TNF α
ELISA was designed to detect the higher levels of TNF α that have been shown to be induced by LPS exposure (Wang et al., 2005).
OPCs Cultured with bFGF and PDGF Express Markers of both Immature and Mature OL Lineage Cells
OPCs grown continuously with PDGF and bFGF were observed to have a bipolar morphology typical of OPCs by day 1 in vitro and were labeled with
antibodies to both A2B5 and NG2 in their somata and processes on day 3 in
vitro . OPCs also labeled positive for GalC or MBP, which are typically
considered markers of mature OL lineage cells (Fig. 4.2).
OPC Viability is Reduced by LPS-Activated MG
For these experiments, > 95 % pure OPC cultures and OPC cultures with
MG added at a ratio of 1:1 were utilized (Fig. 4.3 A). There was no effect of MG
addition on OPC number; additionally, there was no effect of LPS addition on
OPC number when OPCs were cultured alone. In OPC-MG combined cultures,
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Figure 4.2 OPCs grown in bFGF/PDGF labeled for markers of both immature and mature OLs on day 3 in vitro . OPCs were positive for NG2 and A2B5, which are typically considered markers of immature OLs as well as GalC and MBP, which are considered markers of mature OLs.
80
Figure 4.3 MG labeled with IB4 in combined culture with OPCs (A) and mature OLs (B).
there was a significant reduction in OPC number in the presence of 10 ng/ml LPS
(Fig. 4.4). The number of live OPCs in the presence of MG plus 10 ng/ml LPS was reduced to 34 ± 5 % of control values (p = 0.002), indicating that the presence of MG and LPS in combination decreased OPC survival. This was confirmed via factorial ANOVA, revealing a significant (p = 0.036) interaction between MG and LPS and a significant effect of MG (p = 0.001).
OPC Proliferation is not altered by MG or LPS-Activated MG
OPCs were observed to incorporate BrdU in all experimental conditions, with an average 6 % of OPCs incorporating BrdU in the control conditions (Fig.
81
4.5). There was no significant effect or interaction between LPS or MG on OPC proliferation (Fig. 4.6).
Figure 4.4 LPS addition and MG addition had no effect on OPC survival, but MG and LPS in combination resulted in a significant decrease in survival of OPCs (p = 0.002, Tukey’s post hoc).
Both MG and LPS-Activated MG Reduce Caspase Activation in OPCs
To determine if OPC loss in response to activated MG was mediated by caspase
activation; caspase activation was assayed at 3 days in vitro (Fig. 4.7). As
in the previous experiments, OPCs alone, OPCs plus 10 ng/ml LPS, OPCs plus
MG and OPCs plus MG and LPS were analyzed. Surprisingly, there was a
significant main effect of MG in reducing caspase activation of OPCs (p = 0.023,
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Figure 4.5. Example of an OPC labeled with BrdU. OPCs incubated with BrdU prior to fixation were labeled with Hoechst, antibody to BrdU and A2B5. BrdU labeling was observed to colocalize with nuclear Hoechst labeling.
83
Figure 4.6 BrdU incorporation in OPCs was not altered by MG or LPS, and there was no interaction of MG and LPS on altering BrdU incorporation. However, it should be noted that the main effect of MG approached statistical significance (p = 0.056).
Figure 4.7 Composite of an apoptotic OPC labeled with Hoechst and a commercially available caspase detection kit (FLICA). Apoptotic OPCs were defined as cells that had fragmented nuclei and cell bodies labeled brightly with the caspase detection kit.
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Fig. 4.8). However, the absolute number of OPCs expressing active caspase enzymes was very small, averaging 2.4 ± 0.2 caspase positive OPCs per coverslip in the control condition, representing less than 2 % of live OPCs
counted per coverslip. This indicates that although MG reduced apoptosis at the
timepoint we examined, this effect only involves a small subpopulation of OPCs
and therefore is not powerful enough to affect overall OPC number.
Figure 4.8 MG reduce caspase activation in OPCs. Though the addition of 10 ng/ml LPS and MG alone and in combination did not significantly change OPC caspase activation, there was a main effect of MG on reducing caspase activation in OPCs (p = 0.023).
OLs Cultured with Thyroid Hormone Only Express Markers of Mature OLs
85
OLs grown continuously with thyroid hormone were seen to have multiple
processes by day 1 in vitro and were labeled with antibodies to GalC and MBP on day 3 in vitro . OLs grown in thyroid hormone did not label positively for NG2 or A2B5 (Fig. 4.9).
Non-Activated as well as LPS-Activated MG Increase Survival of Mature
OLs
For these experiments, pure OL cultures and OL cultures with MG added were utilized (Fig. 4.3 B). As with OPCs, there was no effect of LPS on OLs in culture alone. However, there was an effect of MG on OL survival (Fig. 4.10). In the presence of MG, OL survival was significantly increased by 53 ± 11 % (p =
0.013 from control), and in the presence of MG and LPS, OL survival was significantly increased 63 ± 12 % (p = 0.004 from control). Factorial ANOVA confirmed a main effect of MG (p < 0.001) but no main effect of LPS or interaction between MG and LPS.
To exclude a possible influence of thyroid hormone on the pro-survival effect of MG, this experiment was repeated with thyroid hormone-containing media being replaced with thyroid hormone-free media on day 1 in vitro (at the
same time as MG were added to OLs). Under these conditions, the significant
protective effect of MG was retained (p = 0.043, data not shown).
86
Figure 4.9 OLs grown in media containing thyroid hormone labeled for oligodendrocyte markers at day 3 in vitro . OLs did not label positively for NG2 or A2B5, but did label positive for GalC and MBP.
87
Figure 4.10 MG, and MG and LPS in combination significantly increased OL survival (p = 0.013 and p = 0.014, respectively), and there was a significant main effect of MG in increasing OL survival (p < 0.001).
Both MG and LPS-Activated MG Reduce Caspase Activation in Mature OLs
To further investigate the mechanism of MG-enhanced OL survival, we quantified caspase activation in OLs cultured with and without MG and LPS.
Based on previous experiments (see Chapter 3) we knew that OLs in our culture system underwent caspase activation that increased over time. Experiments here examined caspase activation in OLs at 3 DIV. There was a significant main
88 effect of MG in reducing caspase activation in OLs (Fig 4.11). In the presence of
MG, caspase activation was significantly decreased 66 ± 9 % from controls (p =
0.015, Tukey’s post hoc). In the presence of MG and LPS, caspase activation
Figure 4.11 In mature OLs, there was a significant main effect of MG in reducing caspase activation (Factorial ANOVA, p < 0.001).
was decreased 58 ± 9 % from controls, which did not achieve statistical significance. Factorial ANOVA demonstrated a significant main effect of MG (p <
0.001) but no main effect of LPS or interaction between MG and LPS.
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Mature OLs do not Incorporate BrdU
BrdU incorporation was used to evaluate cell division in mature OLs.
Since TH has been shown to block OL proliferation (Baas et al., 1997), it was hypothesized that BrdU incorporation would not be observed in mature OLs. As expected, we did not detect any OLs in the control condition that had incorporated BrdU, and there was no effect of MG or LPS on BrdU incorporation in mature OLs (data not shown).
Discussion
Here we demonstrate that the maturation state of OLs has a profound effect on their response to MG and MG activation. While reducing the survival of
OPCs, LPS-activated MG reduce caspase activation and increase survival of mature OLs. LPS is often used as a potent activator of MG and is generally thought to induce a pathological phenotype of MG. Several previous studies have shown that LPS-activated MG are toxic to OPCs both in vitro and in vivo
(Lehnardt et al., 2002; Pang et al., 2003; Pang et al., 2000) and that this effect is mediated in part by oxidative stress (Li et al., 2005). Previous studies have also demonstrated that OPCs are more vulnerable than mature OLs to a variety of insults, including TNF α (Pang et al., 2005), interferon gamma (Baerwald and
Popko, 1998), excitotoxicity (Rosenberg et al., 2003; Wosik et al., 2004), radiation injury (Fukuda et al., 2005), and free radical mediated injury (Bernardo et al., 2003). As all of these stimuli can cause an increase in intracellular oxidative stress (Spitz et al., 2004), it is possible the vulnerability of OPCs to
90 many types of injury is due to a relative deficiency in antioxidant enzymes and other factors (such as high intracellular iron) that predispose OPCs to oxidative stress (Thorburne and Juurlink, 1996). Recent data from our laboratory suggest that OPCs proliferate after spinal cord injury but do not overcome the net loss of
OLs that occurs (Sun et al., 2006). It is possible that proliferating OPCs in vivo ,
as in our study, are unable to withstand toxins produced by activated. This OPC
proliferation followed by apoptosis may represent a failed attempt at
remyelination after spinal cord injury due to the intrinsic vulnerability of OPCs to
inflammation. Therapies that help boost OPCs antioxidant defenses may have
application in many diverse diseases such as PVL and spinal cord injury where
rescue of proliferating OPCs may lead to better functional outcomes for patients.
There are many developmental changes that occur in OLs that may make
mature cells more resistant to injury, including a decrease in ionotropic glutamate
receptor expression (Itoh et al., 2002), an increase in metabotropic glutamate
receptor expression (Luyt et al., 2006), changes in pro and anti-apoptotic gene
expression (Itoh et al., 2003), or changes antioxidant enzyme levels (Folkerth et
al., 2004). It is possible that OLs in our study resisted MG-mediated injury due to
increased glutathione stores as reported by Fragoso and colleagues (Fragoso et
al., 2004), which would render them less susceptible to oxidative injury mediated
by MG, as well as TNF α and excitotoxicity which also induce oxidative stress.
Since LPS-activated MG in our study and others’ were able to induce OPC loss,
it is should be asked: do LPS-activated MG confer a benefit for mature OLs, or
91 do OLs obtain protection only from the initial presence of MG before LPS- activation and then simply tolerate the negative effects of MG after LPS activation? We hypothesize the latter - that LPS activation is actually pathological for CNS cells in general, but that mature OLs can withstand this activation while still benefiting from the presence of MG before the addition of
LPS. This question could be addressed by activating MG with LPS immediately after adding them to OL cultures. If this procedure did not increase OL survival, it would indicate that OL survival in the presence of LPS-activated MG is really due to the presence of MG without LPS for the 24 hours. This is a reasonable hypothesis considering the fact that LPS activation actually reduces MG cell number (see Chapter 5). If MG secrete growth factors that promote OL survival, it would be expected that as MG number decreases, OL survival would decrease as well. Since the magnitude of OL survival and reduction in apoptosis was similar in the MG and MG + LPS conditions, even as MG number dramatically decreased, it would seem that even having MG present for a short time in culture with OLs modified the culture environment to provide extended benefit to OLs, even if MG did not survive for extended periods.
Though activated MG reduce OPC survival, our findings and others’ demonstrate that MG have the ability to protect mature OLs from apoptotic cell death. Pro-survival factors secreted by MG, such as insulin-like growth factor-2, may allow OLs to resist TNF α and other molecules also produced by activated
MG (Nicholas et al., 2002). However, interferon-gamma has been shown to
92 reverse MG phenotype from protective to toxic for OLs when the two cell types are in direct contact (Nicholas et al., 2003). The fact that in our experiments,
LPS-activation did not change the anti-apoptotic effect of MG, shows that increased activation does not always cause MG to lose their beneficial effects. It also indicates that different activating stimuli may have induce profoundly different types of MG activation. Others have shown MG to have divergent effects on OPC genesis depending on the cytokine profile that was used to activate MG (Butovsky et al., 2006). In the injured CNS, multiple inflammatory cytokines are produced (Yang et al., 2005a) and the combination of cytokines may change over time, potentially resulting in a dynamic MG phenotype after injury. Subsequent experiments will need to determine which factors produced by activated MG mediate OPC death and OL survival, in order to develop strategies to block the former and promote the latter. Pharmacologic therapies that inhibit MG-mediated OPC cell death but sustain the production of pro- survival factors could be of benefit in several diseases that involve MG activation and OPC/OL cell death.
An important factor to consider in light of the divergent effects of MG on
OPCs and OLs is that OPCs and OLs themselves are in very different states in vitro. OPCs are continually proliferating (as evidenced by BrdU incorporation) while mature OLs are gradually undergoing apoptosis (as described in Chapter
3). The fact that the effects of MG and activated MG are able to overcome the
“natural” states of OPCs and OLs in vitro emphasizes the power of MG to
93 dramatically affect OPC/OL survival. As MG are involved in many CNS disease processes, and can have such dramatic effects on OPC and mature OL survival, further study of MG activation is needed. Chapter 5 begins to address this subject, examining MG TNF α production and survival after LPS activation under
different culture conditions.
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CHAPTER 5
MICROGLIAL CELL DEATH IN RESPONSE TO LPS ACTIVATION
Introduction
Microglia are known to react quickly in response to central nervous system injury (Kreutzberg, 1996), migrating into an injury site and secreting a wide array of molecules, including TNF α (Kreutzberg, 1996; Dasgupta et al.,
2003), glutamate (Nakamura et al., 2003), free radicals (Boje and Arora, 1992)
and other inflammatory and anti-inflammatory mediators that affect cell survival
(Barron, 1995). It has been shown that activated MG can lead to the death of
CNS neurons (Thery and Mallat, 1993; Shin et al., 2004) and contribute to OL
death in models of MS (Benveniste, 1997; Sriram and Rodriguez, 1997; Arevalo-
Martin et al., 2003). In PVL, MG surrounding necrotic foci express TNF α
(Deguchi et al., 1996), and MG have also been shown to produce TNF α in an
experimental model of multiple sclerosis (Renno et al., 1995). As previously
mentioned, MG have also been found in close apposition to apoptotic OLs in a
spinal cord injury (Shuman et al., 1997) and it is possible that these MG are
triggering OL death, as activated MG in vitro exert their cytotoxic effect on OLs
primarily through membrane-bound TNF α (Zajicek et al., 1992). Another in vitro
95 study of MG and OLs showed that while MG-conditioned media enhanced OL maturation, conditioned media of LPS-stimulated MG was toxic to OL precursors
(Pang et al., 2000).
Many of the factors that influence OL survival also lead to MG activation, such as glutamate (Noda et al., 2000), and proinflammatory cytokines (Thery and
Mallat, 1993; Kreutzberg, 1996). LPS, the stimulus used to activate MG in our experiments (see Chapter 4), is commonly used to induce MG-mediated neuronal and OPC death. LPS acts on MG through Toll-like receptor 4 (TLR4), activating the transcription factor Nuclear factor kappa-B that initiates production of pro-inflammatory cytokines (Lee and Lee, 2002). The TLR4 pathway is central to MG-induced OPC damage, as TLR4 is not present on OPCs but MG-mediated
OPC death is dependent on TLR4 function (Lehnardt et al., 2002). Interestingly,
MG themselves have also been shown to undergo cell death in response to LPS activation (Liu et al., 2001). However, MG cell death in response to activation does not preclude MG from inducing cell death in other CNS cells (Lehnardt et al., 2003). This MG cell death in response to LPS stimulation in vitro is likely not an experimental artifact, as activated MG have been shown to undergo cell death in parallel with activation in vivo (Shuman et al., 1997).
These experiments were performed to determine if MG survival in our experiments was affected by prolonged activation by LPS. The first set of data reported here was gathered from OPC/OL-MG combination cultures that were described in Chapter 4. Subsequent experiments were carried out in cultures of
96
MG alone. We hypothesized that as MG were activated by LPS, they would undergo cell death, as MG have been reported to undergo apoptosis during CNS inflammation in vivo (White et al., 1998). We also performed these experiments to determine if the different media formulations necessary to generate both OPCs and mature OLs would have different effects on MG TNF α production and
survival after exposure to LPS. We did not observe MG TNF α production to be
affected by media composition; however, we did observe that different media
formulations affected MG cell death. Thus, growth factors that modulate OL
development may also play a role in MG survival and could have multiple effects
in CNS diseases and trauma where both OPC/OL death and MG activation
occur.
Materials and Methods
MG Isolation
For experiments examining MG activation in the presence of OLs, MG
were isolated as described in Chapters 3 and 4. For experiments involving MG
alone, MG were plated onto coverslips in an identical fashion as in experiments
involving both MG and OLs.
TNF α ELISA
An enzyme-linked immunosorbant assay for TNF α was performed as described in Chapter 4.
MG Immunocytochemistry MG were labeled with IB4 as described in Chapters 3 and 4.
MG Cell Counts 97
IB4 labeled microglia were counted across the entire diameter of the coverslip. Only MG with intact nuclei were scored as “live;” however, MG with fragmented or absent nuclei were rarely observed, therefore MG counts were expressed as percent of live MG in control conditions.
LDH Assay for Assessment of MG Cell Death A commercially available LDH assay kit (Roche, Indianapolis, IN) was used to quantify LDH release in experiments with MG grown alone. In 3 independent replications, MG were isolated as described above, and plated at densities varying from 50,000 to 150,000 cells/ml into a 96 well plate coated with poly-L-lysine. 24 hours after plating, 10 ng/ml LPS was administered for 24 hours and LDH release was assayed according to the manufacturer’s instructions. For positive controls, MG were treated with 1 % triton for 10 minutes prior to LDH release quantification. Cell death was expressed as experimental values minus control values as a percent of positive control values minus negative control values (% cell death = ( experimental value – neg. control )
/ ( pos. control – neg. control )).
Results MG TNF α production is not Altered by Media Composition
A TNF α ELISA was used to confirm MG activation and to test whether MG would produce TNF α regardless of media formulation. Addition of 10 ng/ml LPS to MG cultures induced significant TNF α production. MG cultured in media
containing thyroid hormone (TH, which was used for experiments of combined
OLs and MG) produced an average of 416 ± 37 pg/ml TNF α over 24 hours and
98
MG in media containing bFGF/PDGF (which was used for OPCs and MG) produced 341 ± 25 pg/ml TNF α (Fig. 5.1), but there was no significant effect of media composition on TNF α production. In experiments using combined MG-
OPC/OL cultures, there was no detectable TNF α production by OLs or OPCs
alone treated with LPS or MG cultures that were not treated with LPS (data not
shown).
MG undergo Cell Death in Response to LPS Activation, and LPS-Induced MG Cell Death is More Pronounced in Media Containing TH than Media Containing bFGF/PDGF In experiments described in Chapter 4, counts of MG were also
conducted, and revealed significantly lower numbers of MG in cultures treated
Figure 5.1 There is no statistically significant difference in TNF α production by MG treated with 10 ng/ml LPS in media containing TH or bFGF/PDGF (Student’s T-Test).
99
with LPS. In experiments conducted with MG in combined culture with OPCs, where media contained bFGF and PDGF, MG survival was significantly decreased 42 ± 8 % (p < 0.001 from control, Student’s T- test) after treatment with 10 ng/ml LPS for 24 hours (Fig. 5.2 A). In experiments conducted with MG in combined culture with OLs in media containing TH, MG numbers were significantly decreased 86 ± 2 % (p < 0.001 from control, Student’s T-test) after treatment with 10 ng/ml LPS for 24 hours (Fig. 5.2 B).
To directly compare the magnitude of MG loss in both media conditions, a separate experiment was conducted culturing MG alone in media containing either bFGF and PDGF or TH (Fig. 5.3). In these experiments, MG numbers
Figure 5.2 In separate experiments, 10 ng/ml LPS treatment reduced the number of surviving MG when MG were cultured with OLs and with OPCs (*= p < 0.05).
100 were also decreased following LPS treatment, with 54 ± 8 % of MG grown in bFGF and PDGF containing media being lost and 76 ± 4 % of MG grown in TH- containing media being lost. Factorial ANOVA revealed a significant interaction between media composition and LPS treatment (p = 0.003), but no effect of media alone. This indicates that media composition affected MG loss only in response to LPS. It was confirmed that media composition had no effect on MG number in the absence of LPS by comparing the raw number of MG in
TH-containing media to bFGF/PDGF-containing media, which did not differ (data not shown).
Figure 5.3 10 ng/ml LPS significantly reduced MG number in both bFGF/PDGF- and TH-containing media. There was a significant interaction between media composition and LPS, indicating the media composition altered MG survival in response to LPS (p = 0.003, factorial ANOVA).
101
To confirm that the lower numbers of MG counted were due to MG death and not simply MG detachment from coverslips before fixation, a LDH assay was conducted. 10 ng/ml LPS induced significant LDH release from MG as compared to controls, and as in the previous experiment, there was a significant difference in MG death in response to LPS in different media compositions (Fig.
5.4). MG cultured in media containing bFGF/PDGF underwent 32 ± 3 % cell death as measured by LDH release, and MG cultured in media containing TH underwent 63 ± 1 % cell death, again demonstrating an effect of media composition on MG survival after LPS activation (p < 0.008, Student’s T-Test).
Discussion
MG cell death in response to activation has been shown to occur both in vitro and in vivo (Ryu et al., 2003). As with OPC/OL cell death, MG cell death both in vitro and in vivo is associated with free radical production and caspase activation (Lee et al., 2001b; Ryu et al., 2003). MG cell death has been described in response to mitochondrial toxins (Ryu et al., 2003), proteins in
Alzheimer’s disease plaques (Kingham et al., 1999), and polyamines (Takano et al., 2003). MG also undergo apoptosis along with OLs after spinal cord injury and peripheral nerve injury (Gehrmann and Banati, 1995; Shuman et al., 1997).
Others have shown that MG activated with LPS undergo cell death in a dose-responsive manner (Liu et al., 2001), though not all reports show MG death associated with LPS activation (Chock and Giffard, 2005). This MG “autocrine”
102
Figure 5.4 LDH assay confirmed that MG exposed to 10 ng/ml LPS for 24 hours underwent cell death. MG cultured in TH-containing media underwent a significantly higher level of cell death compared to MG in bFGF/PDGF-containing media (p < 0.001, Student’s T-Test).
death is mediated by free radicals, but not TNF α (Lee et al., 2001b). Though
LPS can induce caspase activation in MG (Lee et al., 2001a), LPS exposure may also cause MG to be more likely to undergo necrotic cell death under some conditions (Nagano et al., 2006). In our experiments assessing caspase activation in OLs and OPCs cultured with MG, we did not observe caspase activation in MG. It may have been that the timepoint we examined was too late to detect caspase activation that would precede a net loss of MG or that MG death was necrotic in nature. Regardless of the nature of MG cell death, it is important to note that the effects of MG on OLs and OPCs described in Chapter
103
4 occurred even in the presence of activation-induced MG cell death. This has been described previously, in a study showing that MG are also lost during MG- mediated neuronal death (Lehnardt et al., 2003). It is possible that sufficient MG- derived toxins or survival factors for OPCs/OLs were produced before MG were lost due to activation. Alternately, the effects of MG may only require a small number of MG, and the MG that persist in the presence of LPS-activation may be sufficient to support OL survival or OPC death. This question could be answered by better resolving the exact timepoints at which OPC and MG cell death occur.
If MG cell death precedes OPC death, it would show that only a few MG are needed to carry out the effects on OPCs. Culturing MG at different densities with
OLs could also answer this question by revealing the minimum density of MG needed to protect OLs from gradual apoptosis. In future experiments, it would be interesting to determine if the effect of MG on OLs can be modulated by the number of MG present at the onset of LPS addition, and if lower doses of LPS that may not induce MG cell death would lead to greater changes in OL/OPC survival.
MG cell death occurs during neuroinflammation in vivo (White et al., 1998)
and may be a form of autoregulation in order to limit the MG response to injury
and prevent excessive damage to other cells in the CNS. It is possible that MG
exhibit a graded activation that can culminate in cell death if the activation
stimulus is extreme enough. This suggestion is supported by data showing that
MG cultured for less time in vitro produce greater amounts of TNF α and undergo
104 cell death to a greater extent than MG cultured for longer periods of time (Chock and Giffard, 2005), demonstrating a possible correlation between MG activation intensity and susceptibility to cell death. In one study examining MG cell death in response to LPS-activation, it was shown that higher LPS concentrations induced more MG cell death and consequently less total TNF α production by MG. This
indicates that MG over-activation and cell death has the potential to reduce the
presence of inflammatory molecules and may be a way to protect other CNS
cells from inflammatory mediators produced by MG (Liu et al., 2001). In
preliminary experiments we conducted examining a range of LPS doses on MG
TNF α production and MG cell death, we observed this to be the case as well,
with the magnitude of TNF α production and MG cell death increasing with LPS dose (data not shown).
The difference in the survival of activated MG in different media conditions observed in the present study could be due to the effects of bFGF on MG. MG have been shown increase transcription of mRNA for fibroblast growth factor receptor 1 (the receptor for bFGF) during experimental autoimmune encephalitis
(Liu et al., 1998). Since bFGF reduces human MG iNOS expression and nitrite production (Colasanti et al., 1995) and LPS-induced MG cell death has been shown to depend on free radical production (Lee et al., 2001b) this may be a mechanism by which bFGF reduces activation-induced MG cell death. It is possible that the bFGF present in our media, while not sufficient to reduce MG
105 activation enough to protect OPCs, did reduce MG activation so that less MG were lost to free radicals and other inflammatory mediators.
In summary, we have demonstrated that MG undergo cell death after
activation by LPS. This MG cell death does not preclude MG-induced OPC loss
or OL survival. There is also an effect of media composition on the magnitude of
MG cell death in response to LPS activation, which may be related to the
presence of bFGF which has been shown to decrease free radical production by
MG. MG cell death due to overactivation may be a regulatory mechanism that
helps protect other CNS cells from the inflammation that occurs after CNS injury,
as a higher density of MG has been shown to correlate with more extensive CNS
damage in response to LPS injection (Kim et al., 2000). Based on data reported
in Chapter 4, it is likely that this activation induced MG cell death would be of
benefit in reducing MG number and possible bystander damage to other CNS
cells. MG cell death has not been studied to the extent of OPC/OL and neuronal
cell death. Further research is needed to determine what factors contribute to
MG survival and cell death in CNS disease and if MG survival can be modulated
to improve outcomes in CNS injury and disease.
106
CHAPTER 6
SUMMARY, CONCLUSIONS AND FUTURE DIRECTIONS
These studies, taken together, show that OLs and OPCs are vulnerable to
cell death from both excitotoxic and inflammatory stimuli. Optic nerve OLs were
shown to be susceptible to KA-induced excitotoxicity which could be increased
by TNF α. Cortical OLs were also susceptible KA-induced excitotoxicity in a dose-dependent manner.
Several studies were conducted to understand more about the mechanism of KA toxicity in OLs, and to explore possible therapies for excitotoxic OL loss.
Though OL cell death in several in vivo experimental models can be blocked by glutamate receptor antagonists (Kanwar et al., 2004; Follett et al., 2000) these compounds have been shown to have side effects when used clinically (Muir and
Lees, 1995) and therefore effort has been made to develop new and clinically applicable pharmacotherapies that will interfere with excitotoxic cell death pathways. One therapy that has been shown to be efficacious in animal models, and can be tolerated by humans is topiramate (Sfaello et al., 2005; Guerrini and
Parmeggiani, 2006). Topiramate acts in part by antagonizing glutamate receptors (Walker and Sander, 1996). We tested if the simultaneous application
107 of topiramate with KA would reduce excitotoxic cell death in cortical OLs, but a preliminary experiment did not yield promising results (data not shown). This negative result, which examined only a single dose of topiramate, dose of KA and timepoint must be viewed as inclusive. Further experiments will be needed to test the effects of different doses of topiramate and other therapeutics on OL excitotoxicity.
In vivo data from our laboratory has shown that oxidative changes precede OL death after spinal cord injury (Sun et al., 2006), and we sought to determine if oxidative stress preceded excitotoxic OL death in our experimental preparations, as has been shown by others (Ibarretxe et al., 2006). We conducted several experiments using the reactive oxygen species (ROS) sensitive dye Carboxy-H2DCFDA at several timepoints after administration of KA
to OLs. This dye is originally present in a nonfluorescent form and becomes
fluorescent when reduced by intracellular ROS. While preliminary experiments
showed Carboxy-H2DCFDA to be sensitive to oxidative changes in our positive controls (Fig. 6.1), we were not able to obtain reliable data demonstrating oxidative changes in response to KA. Further experiments using Carboxy-
H2DCFDA to examine excitotoxicity and MG-induced OPC death are planned. In additional experiments designed to determine if oxidative changes affected OL excitotoxicity, we exposed OLs to KA at 5 % (rather than 20 %) oxygen. We hypothesized that lower oxygen levels in vitro may better simulate the in vivo condition and help cells resist excitotoxicity due to lower basal levels of oxidative
108 stress in cells. A previous study has shown that culturing neurons at lower oxygen tensions that more closely approximate the actual CNS environment resulted in enhanced survival in reduction to (Studer et al., 2000). However, we did not observe an effect of different oxygen levels on excitotoxic OL death (data not shown). We also tested the antioxidant tocotrionol in our OL excitotoxicity paradigm, but did not observe an effect of tocotrionol on excitotoxic OL death
Figure 6.1 Use of Carboxy-H2DCFDA to detect reactive oxygen species in cortical OLs. Positive controls induced to produce reactive oxygen species emitted qualitatively higher signals than negative controls. Experiments examining ROS formation during KA excitotoxicity were inconclusive.
(data not shown). Again, this negative result does not exclude the possibility that oxidative stress plays a role in excitotoxic OL cell death and additional experiments are planned to assess the efficacy of antioxidants in reducing excitotoxic OL cell death.
109
Continued interest in the synergy of inflammatory cytokines and excitotoxicity in inducing OL cell death led to experiments utilizing cultures of cortical OLs in combination with MG, which have been shown to produce TNF α in response to excitotoxic stimuli (Noda et al., 2000). Driven by in vivo
observations made previously in our laboratory (Shuman et al., 1997), we hoped
to develop a tissue culture model that would allow us to study MG-OL
interactions and test specific hypotheses about these cells’ roles and fates in
CNS injury. Our first hypothesis was that MG, which respond to KA stimulation
(Christensen et al., 2006), would increase OL response to KA when the two cell
types were cultured together. We hypothesized that this effect would be due to
TNF α release by MG, which we had observed to increase OL response to KA
(see Chapter 2). Contrary to our hypothesis, there was no interaction between
MG and OL response to KA, however MG did reduce OL cell death overall in this
system, leading to our next set of experiments.
We combined cortical OPCs with MG, and were able to obtain similar
results as others (Lehnardt et al., 2002), observing OPC cell death in response to
MG activated with LPS. Others have shown that MG-induced OPC death is
dependent on free radical formation (Haynes et al., 2005) and we plan to
determine if OPC death in our system is amenable to treatment with anti-
oxidants. We also observed MG to undergo cell death in this preparation, which
seems to parallel MG death that occurs in vivo after neurological injury (Shuman
et al., 1997). One question that remains unanswered is the time course of this
110
OPC and MG cell death. Do MG begin undergoing cell death before OPCs, or is
MG cell death a relatively late event that may occur after all OPC death has occurred? If the later is true, then the hypothesized autoregulatory role of activation-induced MG cell death may not be an important event. However, if
MG numbers decline as OPC death is occurring, then MG loss would likely reduce the loss of additional OPCs. One way to resolve this question would be the use of live cell imaging studies, where culture preparations could be monitored continuously and both OPC and MG viability could be tracked on a continuous basis. We are currently developing the technology to carry out these experiments, measuring caspase activation and free radical production at timepoints that precede the onset of cell death observed in live cell experiments.
Surprisingly, we found that MG, regardless of activation state, reduced apoptotic cell death of mature OLs. Further experiments will be conducted with specific caspase probes, such as caspase-8 and -9, to determine if OL apoptosis is occurring through the extrinsic or intrinsic pathway. Though other studies have demonstrated MG-mediated OL protection (Nicholas et al., 2001), we are the first to show that even MG activated to an extent that induces OPC death can provide support for OLs in vitro (Fig. 6.2). We hypothesize that this protection is due to
MG production of IGF2, which has been shown to mediate the protective effect of
MG on OLs in similar experiments (Nicholas et al., 2002). Future experiments, such as determining if MG-mediated OL protection can be blocked antibodies to
IGF2 or other growth factors, will be needed to determine the mechanism of this
111 effect. Additionally, varying the relative proportion of MG in our cultures would help determine if MG cell death is in fact a glial-protective effect. If MG at higher densities can reduce OL survival when activated, it could be inferred that MG cell in CNS disease is a autoregulatory and protective event in CNS disease.
Finally, we hope that further research will be conducted to better define the term “microglial activation,” as this term has been sometimes carelessly applied to in vivo and in vitro conditions that may not be synonymous (Hurley et al., 1999). MG in culture may be constitutively activated to a certain degree, as rod-like MG, which are defined as resting in vivo are rarely observed in vitro .
Though LPS and other activating stimuli clearly alter MG phenotype in vitro,
thorough profiling of all cytokines and growth factors produced in response to
different activating stimuli has not been conducted. MG expression of certain
receptors, such as cannabinoid receptor type-2 (CB2), is differentially induced by
different activating stimuli (Carlisle et al., 2002). This differential receptor
expression in response to different activating stimuli may be of great importance,
as activation of the CB2 receptor has been shown to reduce MG-mediated
neuronal death in vitro (Ramirez et al., 2005). As scientists develop a better understanding of the role of activated MG in CNS injury and recovery, future therapies that suppress injurious effects and promote beneficial effects of MG may be developed.
112
Figure 6.2 Schematic of combined results from Chapters 4 and 5. LPS activated MG induce OPC cell death and undergo cell death themselves. MG reduce cell death of mature OLs regardless of activation state and cell death.
113
REFERENCES
1. Ahlgren SC, Wallace H, Bishop J, Neophytou C, Raff MC (1997) Effects of thyroid hormone on embryonic oligodendrocyte precursor cell development in vivo and in vitro. Mol Cell Neurosci 9: 420-432.
2. Aktas O, Prozorovski T, Zipp F (2006) Death ligands and autoimmune demyelination. Neuroscientist 12: 305-316.
3. Alberch J, Perez-Navarro E, Canals JM (2002) Neuroprotection by neurotrophins and GDNF family members in the excitotoxic model of Huntington's disease. Brain Res Bull 57: 817-822.
4. Alberdi E, Sanchez-Gomez MV, Marino A, Matute C (2002) Ca(2+) influx through AMPA or kainate receptors alone is sufficient to initiate excitotoxicity in cultured oligodendrocytes. Neurobiol Dis 9: 234-243.
5. Alberdi E, Sanchez-Gomez MV, Torre I, Domercq M, Perez-Samartin A, Perez-Cerda F, Matute C (2006) Activation of kainate receptors sensitizes oligodendrocytes to complement attack. J Neurosci 26: 3220-3228.
6. Anderson DW, Ellenberg JH, Leventhal CM, Reingold SC, Rodriguez M, Silberberg DH (1992) Revised estimate of the prevalence of multiple sclerosis in the United States. Ann Neurol 31: 333-336.
7. Ankeny DP, Lucin KM, Sanders VM, McGaughy VM, Popovich PG (2006) Spinal cord injury triggers systemic autoimmunity: evidence for chronic B lymphocyte activation and lupus-like autoantibody synthesis. J Neurochem 99: 1073-1087.
8. Arevalo-Martin A, Vela JM, Molina-Holgado E, Borrell J, Guaza C (2003) Therapeutic action of cannabinoids in a murine model of multiple sclerosis. J Neurosci 23: 2511-2516.
9. Arnett HA, Mason J, Marino M, Suzuki K, Matsushima GK, Ting JP (2001) TNF alpha promotes proliferation of oligodendrocyte progenitors and remyelination. Nat Neurosci 4: 1116-1122.
114
10. Arundine M, Tymianski M (2004) Molecular mechanisms of glutamate- dependent neurodegeneration in ischemia and traumatic brain injury. Cell Mol Life Sci 61: 657-668.
11. Baas D, Bourbeau D, Sarlieve LL, Ittel ME, Dussault JH, Puymirat J (1997) Oligodendrocyte maturation and progenitor cell proliferation are independently regulated by thyroid hormone. Glia 19: 324-332.
12. Back SA (2006) Perinatal white matter injury: the changing spectrum of pathology and emerging insights into pathogenetic mechanisms. Ment Retard Dev Disabil Res Rev 12: 129-140.
13. Back SA, Gan X, Li Y, Rosenberg PA, Volpe JJ (1998) Maturation- dependent vulnerability of oligodendrocytes to oxidative stress-induced death caused by glutathione depletion. J Neurosci 18: 6241-6253.
14. Baerwald KD, Popko B (1998) Developing and mature oligodendrocytes respond differently to the immune cytokine interferon-gamma. J Neurosci Res 52: 230-239.
15. Bansal R, Warrington AE, Gard AL, Ranscht B, Pfeiffer SE (1989) Multiple and novel specificities of monoclonal antibodies O1, O4, and R-mAb used in the analysis of oligodendrocyte development. J Neurosci Res 24: 548- 557.
16. Barger SW, Basile AS (2001) Activation of microglia by secreted amyloid precursor protein evokes release of glutamate by cystine exchange and attenuates synaptic function. J Neurochem 76: 846-854.
17. Barron KD (1995) The microglial cell. A historical review. J Neurol Sci 134 Suppl: 57-68.
18. Baud O, Haynes RF, Wang H, Folkerth RD, Li J, Volpe JJ, Rosenberg PA (2004) Developmental up-regulation of MnSOD in rat oligodendrocytes confers protection against oxidative injury. Eur J Neurosci 20: 29-40.
19. Beal MF (1998) Excitotoxicity and nitric oxide in Parkinson's disease pathogenesis. Ann Neurol 44: S110-S114.
20. Beattie EC, Stellwagen D, Morishita W, Bresnahan JC, Ha BK, Von Zastrow M, Beattie MS, Malenka RC (2002a) Control of synaptic strength by glial TNFalpha. Science 295: 2282-2285.
21. Beattie MS (2004) Inflammation and apoptosis: linked therapeutic targets in spinal cord injury. Trends Mol Med 10: 580-583.
115
22. Beattie MS, Hermann GE, Rogers RC, Bresnahan JC (2002b) Cell death in models of spinal cord injury. Prog Brain Res 137: 37-47.
23. Bellander BM, Bendel O, Von Euler G, Ohlsson M, Svensson M (2004) Activation of microglial cells and complement following traumatic injury in rat entorhinal-hippocampal slice cultures. J Neurotrauma 21: 605-615.
24. Benjamins JA, Nedelkoska L, George EB (2003) Protection of mature oligodendrocytes by inhibitors of caspases and calpains. Neurochem Res 28: 143-152.
25. Benveniste EN (1997) Role of macrophages/microglia in multiple sclerosis and experimental allergic encephalomyelitis. J Mol Med 75: 165-173.
26. Bernardo A, Greco A, Levi G, Minghetti L (2003) Differential lipid peroxidation, Mn superoxide, and bcl-2 expression contribute to the maturation-dependent vulnerability of oligodendrocytes to oxidative stress. J Neuropathol Exp Neurol 62: 509-519.
27. Bitsch A, Kuhlmann T, Da Costa C, Bunkowski S, Polak T, Bruck W (2000) Tumour necrosis factor alpha mRNA expression in early multiple sclerosis lesions: correlation with demyelinating activity and oligodendrocyte pathology. Glia 29: 366-375.
28. Blumenthal I (2004) Periventricular leucomalacia: a review. Eur J Pediatr 163: 435-442.
29. Bogler O, Wren D, Barnett SC, Land H, Noble M (1990) Cooperation between two growth factors promotes extended self-renewal and inhibits differentiation of oligodendrocyte-type-2 astrocyte (O-2A) progenitor cells. Proc Natl Acad Sci U S A 87: 6368-6372.
30. Boje KM, Arora PK (1992) Microglial-produced nitric oxide and reactive nitrogen oxides mediate neuronal cell death. Brain Res 587: 250-256.
31. Bolton C, Paul C (2006) Glutamate receptors in neuroinflammatory demyelinating disease. Mediators Inflamm 2006: 93684.
32. Bottenstein JE, Sato GH (1979) Growth of a rat neuroblastoma cell line in serum-free supplemented medium. Proc Natl Acad Sci U S A 76: 514-517.
33. Brockschnieder D, Sabanay H, Riethmacher D, Peles E (2006) Ermin, a myelinating oligodendrocyte-specific protein that regulates cell morphology. J Neurosci 26: 757-762.
116
34. Butovsky O, Landa G, Kunis G, Ziv Y, Avidan H, Greenberg N, Schwartz A, Smirnov I, Pollack A, Jung S, Schwartz M (2006) Induction and blockage of oligodendrogenesis by differently activated microglia in an animal model of multiple sclerosis. J Clin Invest 116: 905-915.
35. Calabrese V, Bates TE, Stella AM (2000) NO synthase and NO- dependent signal pathways in brain aging and neurodegenerative disorders: the role of oxidant/antioxidant balance. Neurochem Res 25: 1315-1341.
36. Camello-Almaraz C, Gomez-Pinilla PJ, Pozo MJ, Camello PJ (2006) Mitochondrial reactive oxygen species and Ca2+ signaling. Am J Physiol Cell Physiol 291: C1082-C1088.
37. Cammer W (2002) Protection of cultured oligodendrocytes against tumor necrosis factor-alpha by the antioxidants coenzyme Q(10) and N-acetyl cysteine. Brain Res Bull 58: 587-592.
38. Cammer W, Zhang H (1999) Maturation of oligodendrocytes is more sensitive to TNF alpha than is survival of precursors and immature oligodendrocytes. J Neuroimmunol 97: 37-42.
39. Cantarella G, Risuglia N, Lombardo G, Lempereur L, Nicoletti F, Memo M, Bernardini R (2004) Protective effects of estradiol on TRAIL-induced apoptosis in a human oligodendrocytic cell line: evidence for multiple sites of interactions. Cell Death Differ 11: 503-511.
40. Carbonell WS, Murase S, Horwitz AF, Mandell JW (2005) Migration of perilesional microglia after focal brain injury and modulation by CC chemokine receptor 5: an in situ time-lapse confocal imaging study. J Neurosci 25: 7040-7047.
41. Carlisle SJ, Marciano-Cabral F, Staab A, Ludwick C, Cabral GA (2002) Differential expression of the CB2 cannabinoid receptor by rodent macrophages and macrophage-like cells in relation to cell activation. Int Immunopharmacol 2: 69-82.
42. Caruso A, Di GG, V, Castiglione M, Marinelli F, Tomassini V, Pozzilli C, Caricasole A, Bruno V, Caciagli F, Moretti A, Nicoletti F, Melchiorri D (2004) Testosterone amplifies excitotoxic damage of cultured oligodendrocytes. J Neurochem 88: 1179-1185.
43. Castejon OJ (1985) Electron microscopic study of central axons degeneration in traumatic human brain edema. J Submicrosc Cytol 17: 703-718.
117
44. Cheung NS, Pascoe CJ, Giardina SF, John CA, Beart PM (1998) Micromolar L-glutamate induces extensive apoptosis in an apoptotic- necrotic continuum of insult-dependent, excitotoxic injury in cultured cortical neurones. Neuropharmacology 37: 1419-1429.
45. Chew LJ, Fleck MW, Wright P, Scherer SE, Mayer ML, Gallo V (1997) Growth factor-induced transcription of GluR1 increases functional AMPA receptor density in glial progenitor cells. J Neurosci 17: 227-240.
46. Chittajallu R, Braithwaite SP, Clarke VR, Henley JM (1999) Kainate receptors: subunits, synaptic localization and function. Trends Pharmacol Sci 20: 26-35.
47. Cho SG, Choi EJ (2002) Apoptotic signaling pathways: caspases and stress-activated protein kinases. J Biochem Mol Biol 35: 24-27.
48. Chock VY, Giffard RG (2005) Development of neonatal murine microglia in vitro: changes in response to lipopolysaccharide and ischemia-like injury. Pediatr Res 57: 475-480.
49. Choi DW (1988) Glutamate neurotoxicity and diseases of the nervous system. Neuron 1: 623-634.
50. Christensen RN, Ha BK, Sun F, Bresnahan JC, Beattie MS (2006) Kainate induces rapid redistribution of the actin cytoskeleton in ameboid microglia. J Neurosci Res 84: 170-181.
51. Cohen J (1997) Statistical power analysis for the behavioral sciences. New York: Academic Press.
52. Colasanti M, Di Pucchio T, Persichini T, Sogos V, Presta M, Lauro GM (1995) Inhibition of inducible nitric oxide synthase mRNA expression by basic fibroblast growth factor in human microglial cells. Neurosci Lett 195: 45-48.
53. Cook SD (2004) Does epstein-barr virus cause multiple sclerosis? Rev Neurol Dis 1: 115-123.
54. Crowe MJ, Bresnahan JC, Shuman SL, Masters JN, Beattie MS (1997) Apoptosis and delayed degeneration after spinal cord injury in rats and monkeys. Nat Med 3: 73-76.
55. Dasgupta S, Jana M, Liu X, Pahan K (2002) Myelin basic protein-primed T cells induce nitric oxide synthase in microglial cells. Implications for multiple sclerosis. J Biol Chem 277: 39327-39333.
118
56. Dasgupta S, Jana M, Liu X, Pahan K (2003) Role of very-late antigen-4 (VLA-4) in myelin basic protein-primed T cell contact-induced expression of proinflammatory cytokines in microglial cells. J Biol Chem 278: 22424- 22431.
57. De Nicola AF, Gonzalez SL, Labombarda F, Deniselle MC, Garay L, Guennoun R, Schumacher M (2006) Progesterone treatment of spinal cord injury: Effects on receptors, neurotrophins, and myelination. J Mol Neurosci 28: 3-15.
58. Deguchi K, Mizuguchi M, Takashima S (1996) Immunohistochemical expression of tumor necrosis factor alpha in neonatal leukomalacia. Pediatr Neurol 14: 13-16.
59. Deng W, Rosenberg PA, Volpe JJ, Jensen FE (2003) Calcium-permeable AMPA/kainate receptors mediate toxicity and preconditioning by oxygen- glucose deprivation in oligodendrocyte precursors. Proc Natl Acad Sci U S A 100: 6801-6806.
60. Dietrich J, Noble M, Mayer-Proschel M (2002) Characterization of A2B5+ glial precursor cells from cryopreserved human fetal brain progenitor cells. Glia 40: 65-77.
61. Dopp JM, Sarafian TA, Spinella FM, Kahn MA, Shau H, de Vellis J (2002) Expression of the p75 TNF receptor is linked to TNF-induced NFkappaB translocation and oxyradical neutralization in glial cells. Neurochem Res 27: 1535-1542.
62. Du J, Gray NA, Falke CA, Chen W, Yuan P, Szabo ST, Einat H, Manji HK (2004) Modulation of synaptic plasticity by antimanic agents: the role of AMPA glutamate receptor subunit 1 synaptic expression. J Neurosci 24: 6578-6589.
63. Du Y, Fischer TZ, Lee LN, Lercher LD, Dreyfus CF (2003) Regionally specific effects of BDNF on oligodendrocytes. Dev Neurosci 25: 116-126.
64. Duchen MR (2000) Mitochondria and calcium: from cell signalling to cell death. J Physiol 529 Pt 1: 57-68.
65. Dutly F, Schwab ME (1991) Neurons and astrocytes influence the development of purified O-2A progenitor cells. Glia 4: 559-571.
66. Engel U, Wolswijk G (1996) Oligodendrocyte-type-2 astrocyte (O-2A) progenitor cells derived from adult rat spinal cord: in vitro characteristics and response to PDGF, bFGF and NT-3. Glia 16: 16-26.
119
67. Eskes C, Juillerat-Jeanneret L, Leuba G, Honegger P, Monnet-Tschudi F (2003) Involvement of microglia-neuron interactions in the tumor necrosis factor-alpha release, microglial activation, and neurodegeneration induced by trimethyltin. J Neurosci Res 71: 583-590.
68. Fan L, Young PR, Barone FC, Feuerstein GZ, Smith DH, McIntosh TK (1996) Experimental brain injury induces differential expression of tumor necrosis factor-alpha mRNA in the CNS. Brain Res Mol Brain Res 36: 287-291.
69. Farber K, Kettenmann H (2005) Physiology of microglial cells. Brain Res Brain Res Rev 48: 133-143.
70. Ffrench-Constant C, Raff MC (1986) Proliferating bipotential glial progenitor cells in adult rat optic nerve. Nature 319: 499-502.
71. Filipovic R, Zecevic N (2005) Interaction between Microglia and Oligodendrocyte Cell Progenitors Involves Golli Proteins. Ann N Y Acad Sci 1048: 166-174.
72. Folkerth RD (2006) Periventricular leukomalacia: overview and recent findings. Pediatr Dev Pathol 9: 3-13.
73. Folkerth RD, Haynes RL, Borenstein NS, Belliveau RA, Trachtenberg F, Rosenberg PA, Volpe JJ, Kinney HC (2004) Developmental lag in superoxide dismutases relative to other antioxidant enzymes in premyelinated human telencephalic white matter. J Neuropathol Exp Neurol 63: 990-999.
74. Follett PL, Deng W, Dai W, Talos DM, Massillon LJ, Rosenberg PA, Volpe JJ, Jensen FE (2004) Glutamate receptor-mediated oligodendrocyte toxicity in periventricular leukomalacia: a protective role for topiramate. J Neurosci 24: 4412-4420.
75. Follett PL, Rosenberg PA, Volpe JJ, Jensen FE (2000) NBQX attenuates excitotoxic injury in developing white matter. J Neurosci 20: 9235-9241.
76. Foote AK, Blakemore WF (2005) Repopulation of oligodendrocyte progenitor cell depleted tissue in a model of chronic demyelination. Neuropathol Appl Neurobiol 31: 105-114.
77. Fragoso G, Martinez-Bermudez AK, Liu HN, Khorchid A, Chemtob S, Mushynski WE, Almazan G (2004) Developmental differences in HO- induced oligodendrocyte cell death: role of glutathione, mitogen-activated protein kinases and caspase 3. J Neurochem 90: 392-404.
120
78. Fukuda A, Fukuda H, Swanpalmer J, Hertzman S, Lannering B, Marky I, Bjork-Eriksson T, Blomgren K (2005) Age-dependent sensitivity of the developing brain to irradiation is correlated with the number and vulnerability of progenitor cells. J Neurochem 92: 569-584.
79. Gallo V, Zhou JM, McBain CJ, Wright P, Knutson PL, Armstrong RC (1996) Oligodendrocyte progenitor cell proliferation and lineage progression are regulated by glutamate receptor-mediated K+ channel block. J Neurosci 16: 2659-2670.
80. 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.
81. Gehrmann J, Banati RB (1995) Microglial turnover in the injured CNS: activated microglia undergo delayed DNA fragmentation following peripheral nerve injury. J Neuropathol Exp Neurol 54: 680-688.
82. Gilgun-Sherki Y, Melamed E, Offen D (2004) The role of oxidative stress in the pathogenesis of multiple sclerosis: the need for effective antioxidant therapy. J Neurol 251: 261-268.
83. Gudz TI, Komuro H, Macklin WB (2006) Glutamate stimulates oligodendrocyte progenitor migration mediated via an alphav integrin/myelin proteolipid protein complex. J Neurosci 26: 2458-2466.
84. Guerrini R, Parmeggiani L (2006) Topiramate and its clinical applications in epilepsy. Expert Opin Pharmacother 7: 811-823.
85. Ha BK, Vicini S, Rogers RC, Bresnahan JC, Burry RW, Beattie MS (2002) Kainate-induced excitotoxicity is dependent upon extracellular potassium concentrations that regulate the activity of AMPA/KA type glutamate receptors. J Neurochem 83: 934-945.
86. Hagino Y, Kariura Y, Manago Y, Amano T, Wang B, Sekiguchi M, Nishikawa K, Aoki S, Wada K, Noda M (2004) Heterogeneity and potentiation of AMPA type of glutamate receptors in rat cultured microglia. Glia 47: 68-77.
87. Hayes KC, Hull TC, Delaney GA, Potter PJ, Sequeira KA, Campbell K, Popovich PG (2002) Elevated serum titers of proinflammatory cytokines and CNS autoantibodies in patients with chronic spinal cord injury. J Neurotrauma 19: 753-761.
121
88. Haynes RL, Baud O, Li J, Kinney HC, Volpe JJ, Folkerth DR (2005) Oxidative and nitrative injury in periventricular leukomalacia: a review. Brain Pathol 15: 225-233.
89. Haynes RL, Folkerth RD, Keefe RJ, Sung I, Swzeda LI, Rosenberg PA, Volpe JJ, Kinney HC (2003) Nitrosative and oxidative injury to premyelinating oligodendrocytes in periventricular leukomalacia. J Neuropathol Exp Neurol 62: 441-450.
90. Hermann GE, Rogers RC, Bresnahan JC, Beattie MS (2001) Tumor necrosis factor-alpha induces cFOS and strongly potentiates glutamate- mediated cell death in the rat spinal cord. Neurobiol Dis 8: 590-599.
91. Hess DC, Abe T, Hill WD, Studdard AM, Carothers J, Masuya M, Fleming PA, Drake CJ, Ogawa M (2004) Hematopoietic origin of microglial and perivascular cells in brain. Exp Neurol 186: 134-144.
92. Himi T, Ikeda M, Yasuhara T, Murota SI (2003) Oxidative neuronal death caused by glutamate uptake inhibition in cultured hippocampal neurons. J Neurosci Res 71: 679-688.
93. Hofman FM, Hinton DR, Johnson K, Merrill JE (1989) Tumor necrosis factor identified in multiple sclerosis brain. J Exp Med 170: 607-612.
94. Hollmann M, Hartley M, Heinemann S (1991) Ca2+ permeability of KA- AMPA--gated glutamate receptor channels depends on subunit composition. Science 252: 851-853.
95. Hurley SD, Walter SA, Semple-Rowland SL, Streit WJ (1999) Cytokine transcripts expressed by microglia in vitro are not expressed by ameboid microglia of the developing rat central nervous system. Glia 25: 304-309.
96. Ibarretxe G, Sanchez-Gomez MV, Campos-Esparza MR, Alberdi E, Matute C (2006) Differential oxidative stress in oligodendrocytes and neurons after excitotoxic insults and protection by natural polyphenols. Glia 53: 201-211.
97. Imitola J, Chitnis T, Khoury SJ (2006) Insights into the molecular pathogenesis of progression in multiple sclerosis: potential implications for future therapies. Arch Neurol 63: 25-33.
98. Ito A, Bebo BF, Jr., Matejuk A, Zamora A, Silverman M, Fyfe-Johnson A, Offner H (2001) Estrogen treatment down-regulates TNF-alpha production and reduces the severity of experimental autoimmune encephalomyelitis in cytokine knockout mice. J Immunol 167: 542-552.
122
99. Itoh T, Beesley J, Itoh A, Cohen AS, Kavanaugh B, Coulter DA, Grinspan JB, Pleasure D (2002) AMPA glutamate receptor-mediated calcium signaling is transiently enhanced during development of oligodendrocytes. J Neurochem 81: 390-402.
100. Itoh T, Itoh A, Pleasure D (2003) Bcl-2-related protein family gene expression during oligodendroglial differentiation. J Neurochem 85: 1500- 1512.
101. Jana M, Dasgupta S, Saha RN, Liu X, Pahan K (2003) Induction of tumor necrosis factor-alpha (TNF-alpha) by interleukin-12 p40 monomer and homodimer in microglia and macrophages. J Neurochem 86: 519-528.
102. Janicke RU, Sprengart ML, Wati MR, Porter AG (1998) Caspase-3 is required for DNA fragmentation and morphological changes associated with apoptosis. J Biol Chem 273: 9357-9360.
103. Jensen FE (2002) The role of glutamate receptor maturation in perinatal seizures and brain injury. Int J Dev Neurosci 20: 339-347.
104. Jensen FE (2005) Role of glutamate receptors in periventricular leukomalacia. J Child Neurol 20: 950-959.
105. Jones SA, Jolson DM, Cuta KK, Mariash CN, Anderson GW (2003) Triiodothyronine is a survival factor for developing oligodendrocytes. Mol Cell Endocrinol 199: 49-60.
106. Jones TL, Sorkin LS (2004) Calcium-permeable alpha-amino-3-hydroxy-5- methyl-4-isoxazolepropionic acid/kainate receptors mediate development, but not maintenance, of secondary allodynia evoked by first-degree burn in the rat. J Pharmacol Exp Ther 310: 223-229.
107. Jurewicz A, Matysiak M, Tybor K, Selmaj K (2003) TNF-induced death of adult human oligodendrocytes is mediated by c-jun NH2-terminal kinase- 3. Brain 126: 1358-1370.
108. Kadhim H, Tabarki B, Verellen G, De Prez C, Rona AM, Sebire G (2001) Inflammatory cytokines in the pathogenesis of periventricular leukomalacia. Neurology 56: 1278-1284.
109. Kanwar JR, Kanwar RK, Krissansen GW (2004) Simultaneous neuroprotection and blockade of inflammation reverses autoimmune encephalomyelitis. Brain 127: 1313-1331.
123
110. Karadottir R, Cavelier P, Bergersen LH, Attwell D (2005) NMDA receptors are expressed in oligodendrocytes and activated in ischaemia. Nature 438: 1162-1166.
111. Kim WG, Mohney RP, Wilson B, Jeohn GH, Liu B, Hong JS (2000) Regional difference in susceptibility to lipopolysaccharide-induced neurotoxicity in the rat brain: role of microglia. J Neurosci 20: 6309-6316.
112. Kingham PJ, Cuzner ML, Pocock JM (1999) Apoptotic pathways mobilized in microglia and neurones as a consequence of chromogranin A-induced microglial activation. J Neurochem 73: 538-547.
113. Kondo T, Raff M (2000) The Id4 HLH protein and the timing of oligodendrocyte differentiation. EMBO J 19: 1998-2007.
114. Kreutzberg GW (1996) Microglia: a sensor for pathological events in the CNS. Trends Neurosci 19: 312-318.
115. Lakhani SA, Masud A, Kuida K, Porter GA, Jr., Booth CJ, Mehal WZ, Inayat I, Flavell RA (2006) Caspases 3 and 7: key mediators of mitochondrial events of apoptosis. Science 311: 847-851.
116. Lee J, Hur J, Lee P, Kim JY, Cho N, Kim SY, Kim H, Lee MS, Suk K (2001a) Dual role of inflammatory stimuli in activation-induced cell death of mouse microglial cells. Initiation of two separate apoptotic pathways via induction of interferon regulatory factor-1 and caspase-11. J Biol Chem 276: 32956-32965.
117. Lee P, Lee J, Kim S, Lee MS, Yagita H, Kim SY, Kim H, Suk K (2001b) NO as an autocrine mediator in the apoptosis of activated microglial cells: correlation between activation and apoptosis of microglial cells. Brain Res 892: 380-385.
118. Lee SJ, Lee S (2002) Toll-like receptors and inflammation in the CNS. Curr Drug Targets Inflamm Allergy 1: 181-191.
119. Lehnardt S, Lachance C, Patrizi S, Lefebvre S, Follett PL, Jensen FE, Rosenberg PA, Volpe JJ, Vartanian T (2002) The toll-like receptor TLR4 is necessary for lipopolysaccharide-induced oligodendrocyte injury in the CNS. J Neurosci 22: 2478-2486.
120. Lehnardt S, Massillon L, Follett P, Jensen FE, Ratan R, Rosenberg PA, Volpe JJ, Vartanian T (2003) Activation of innate immunity in the CNS triggers neurodegeneration through a Toll-like receptor 4-dependent pathway. Proc Natl Acad Sci U S A 100: 8514-8519.
124
121. Leuchtmann EA, Ratner AE, Vijitruth R, Qu Y, McDonald JW (2003) AMPA receptors are the major mediators of excitotoxic death in mature oligodendrocytes. Neurobiol Dis 14: 336-348.
122. Lev-Ram V, Grinvald A (1986) Ca2+- and K+-dependent communication between central nervous system myelinated axons and oligodendrocytes revealed by voltage-sensitive dyes. Proc Natl Acad Sci U S A 83: 6651- 6655.
123. Li J, Baud O, Vartanian T, Volpe JJ, Rosenberg PA (2005) Peroxynitrite generated by inducible nitric oxide synthase and NADPH oxidase mediates microglial toxicity to oligodendrocytes. Proc Natl Acad Sci U S A 102: 9936-9941.
124. Li J, Lin JC, Wang H, Peterson JW, Furie BC, Furie B, Booth SL, Volpe JJ, Rosenberg PA (2003) Novel role of vitamin k in preventing oxidative injury to developing oligodendrocytes and neurons. J Neurosci 23: 5816- 5826.
125. Li S, Mealing GA, Morley P, Stys PK (1999) Novel injury mechanism in anoxia and trauma of spinal cord white matter: glutamate release via reverse Na+-dependent glutamate transport. J Neurosci 19: RC16.
126. Lipton SA, Rosenberg PA (1994) Excitatory amino acids as a final common pathway for neurologic disorders. N Engl J Med 330: 613-622.
127. Liu B, Wang K, Gao HM, Mandavilli B, Wang JY, Hong JS (2001) Molecular consequences of activated microglia in the brain: overactivation induces apoptosis. J Neurochem 77: 182-189.
128. Liu HN, Giasson BI, Mushynski WE, Almazan G (2002) AMPA receptor- mediated toxicity in oligodendrocyte progenitors involves free radical generation and activation of JNK, calpain and caspase 3. J Neurochem 82: 398-409.
129. Liu X, Mashour GA, Webster HF, Kurtz A (1998) Basic FGF and FGF receptor 1 are expressed in microglia during experimental autoimmune encephalomyelitis: temporally distinct expression of midkine and pleiotrophin. Glia 24: 390-397.
130. Louis JC, Magal E, Takayama S, Varon S (1993) CNTF protection of oligodendrocytes against natural and tumor necrosis factor-induced death. Science 259: 689-692.
125
131. Luyt K, Varadi A, Durant CF, Molnar E (2006) Oligodendroglial metabotropic glutamate receptors are developmentally regulated and involved in the prevention of apoptosis. J Neurochem.
132. Madrigal JL, Garcia-Bueno B, Caso JR, Perez-Nievas BG, Leza JC (2006) Stress-induced oxidative changes in brain. CNS Neurol Disord Drug Targets 5: 561-568.
133. Magnano MD, Robinson WH, Genovese MC (2004) Demyelination and inhibition of tumor necrosis factor (TNF). Clin Exp Rheumatol 22: S134- S140.
134. Malinow R, Malenka RC (2002) AMPA receptor trafficking and synaptic plasticity. Annu Rev Neurosci 25: 103-126.
135. Man HY, Ju W, Ahmadian G, Wang YT (2000) Intracellular trafficking of AMPA receptors in synaptic plasticity. Cell Mol Life Sci 57: 1526-1534.
136. Matute C (2006) Oligodendrocyte NMDA receptors: a novel therapeutic target. Trends Mol Med 12: 289-292.
137. Matute C, Alberdi E, Domercq M, Perez-Cerda F, Perez-Samartin A, Sanchez-Gomez MV (2001) The link between excitotoxic oligodendroglial death and demyelinating diseases. Trends Neurosci 24: 224-230.
138. Matute C, Domercq M, Sanchez-Gomez MV (2006) Glutamate-mediated glial injury: mechanisms and clinical importance. Glia 53: 212-224.
139. Matute C, Sanchez-Gomez MV, Martinez-Millan L, Miledi R (1997) Glutamate receptor-mediated toxicity in optic nerve oligodendrocytes. Proc Natl Acad Sci U S A 94: 8830-8835.
140. Mayer M, Noble M (1994) N-acetyl-L-cysteine is a pluripotent protector against cell death and enhancer of trophic factor-mediated cell survival in vitro. Proc Natl Acad Sci U S A 91: 7496-7500.
141. McAdoo DJ, Xu GY, Robak G, Hughes MG (1999) Changes in amino acid concentrations over time and space around an impact injury and their diffusion through the rat spinal cord. Exp Neurol 159: 538-544.
142. McCarthy KD, de Vellis J (1980) Preparation of separate astroglial and oligodendroglial cell cultures from rat cerebral tissue. J Cell Biol 85: 890- 902.
126
143. McDonald JW, Althomsons SP, Hyrc KL, Choi DW, Goldberg MP (1998a) Oligodendrocytes from forebrain are highly vulnerable to AMPA/kainate receptor-mediated excitotoxicity. Nat Med 4: 291-297.
144. McDonald JW, Levine JM, Qu Y (1998b) Multiple classes of the oligodendrocyte lineage are highly vulnerable to excitotoxicity. Neuroreport 9: 2757-2762.
145. McEwen ML, Springer JE (2005) A mapping study of caspase-3 activation following acute spinal cord contusion in rats. J Histochem Cytochem 53: 809-819.
146. McLaurin J, D'Souza S, Stewart J, Blain M, Beaudet A, Nalbantoglu J, Antel JP (1995) Effect of tumor necrosis factor alpha and beta on human oligodendrocytes and neurons in culture. Int J Dev Neurosci 13: 369-381.
147. McTigue DM, Horner PJ, Stokes BT, Gage FH (1998) Neurotrophin-3 and brain-derived neurotrophic factor induce oligodendrocyte proliferation and myelination of regenerating axons in the contused adult rat spinal cord. J Neurosci 18: 5354-5365.
148. McTigue DM, Wei P, Stokes BT (2001) Proliferation of NG2-positive cells and altered oligodendrocyte numbers in the contused rat spinal cord. J Neurosci 21: 3392-3400.
149. Mergenthaler P, Dirnagl U, Meisel A (2004) Pathophysiology of stroke: lessons from animal models. Metab Brain Dis 19: 151-167.
150. Michaelis EK (1998) Molecular biology of glutamate receptors in the central nervous system and their role in excitotoxicity, oxidative stress and aging. Prog Neurobiol 54: 369-415.
151. Miller BA, Sun F, Christensen RN, Ferguson AR, Bresnahan JC, Beattie MS (2005) A sublethal dose of TNFalpha potentiates kainate-induced excitotoxicity in optic nerve oligodendrocytes. Neurochem Res 30: 867- 875.
152. Min KJ, Jou I, Joe E (2003) Plasminogen-induced IL-1beta and TNF-alpha production in microglia is regulated by reactive oxygen species. Biochem Biophys Res Commun 312: 969-974.
153. Muir KW, Lees KR (1995) Clinical experience with excitatory amino acid antagonist drugs. Stroke 26: 503-513.
127
154. Nagano T, Kimura SH, Takai E, Matsuda T, Takemura M (2006) Lipopolysaccharide sensitizes microglia toward Ca(2+)-induced cell death: mode of cell death shifts from apoptosis to necrosis. Glia 53: 67-73.
155. Nakamura Y, Ohmaki M, Murakami K, Yoneda Y (2003) Involvement of protein kinase C in glutamate release from cultured microglia. Brain Res 962: 122-128.
156. Nargi-Aizenman JL, Havert MB, Zhang M, Irani DN, Rothstein JD, Griffin DE (2004) Glutamate receptor antagonists protect from virus-induced neural degeneration. Ann Neurol 55: 541-549.
157. Nicholas R, Stevens S, Wing M, Compston A (2003) Oligodendroglial- derived stress signals recruit microglia in vitro. Neuroreport 14: 1001- 1005.
158. Nicholas RS, Stevens S, Wing MG, Compston DA (2002) Microglia- derived IGF-2 prevents TNFalpha induced death of mature oligodendrocytes in vitro. J Neuroimmunol 124: 36-44.
159. Nicholas RS, Wing MG, Compston A (2001) Nonactivated microglia promote oligodendrocyte precursor survival and maturation through the transcription factor NF-kappa B. Eur J Neurosci 13: 959-967.
160. Nicotera P, Lipton SA (1999) Excitotoxins in neuronal apoptosis and necrosis. J Cereb Blood Flow Metab 19: 583-591.
161. Noble M, Mayer-Proschel M, Proschel C (2005) Redox regulation of precursor cell function: insights and paradoxes. Antioxid Redox Signal 7: 1456-1467.
162. Noble M, Smith J, Power J, Mayer-Proschel M (2003) Redox state as a central modulator of precursor cell function. Ann N Y Acad Sci 991: 251- 271.
163. Noble M, Wolswijk G, Wren D (1989) The complex relationship between cell division and the control of differentiation in oligodendrocyte-type-2 astrocyte progenitor cells isolated from perinatal and adult rat optic nerves. Prog Growth Factor Res 1: 179-194.
164. Noda M, Nakanishi H, Nabekura J, Akaike N (2000) AMPA-kainate subtypes of glutamate receptor in rat cerebral microglia. J Neurosci 20: 251-258.
165. Olney JW (1969) Brain lesions, obesity, and other disturbances in mice treated with monosodium glutamate. Science 164: 719-721. 128
166. Pan W, Csernus B, Kastin AJ (2003) Upregulation of p55 and p75 receptors mediating TNF-alpha transport across the injured blood-spinal cord barrier. J Mol Neurosci 21: 173-184.
167. Pang Y, Cai Z, Rhodes PG (2000) Effects of lipopolysaccharide on oligodendrocyte progenitor cells are mediated by astrocytes and microglia. J Neurosci Res 62: 510-520.
168. Pang Y, Cai Z, Rhodes PG (2003) Disturbance of oligodendrocyte development, hypomyelination and white matter injury in the neonatal rat brain after intracerebral injection of lipopolysaccharide. Brain Res Dev Brain Res 140: 205-214.
169. Pang Y, Cai Z, Rhodes PG (2005) Effect of tumor necrosis factor-alpha on developing optic nerve oligodendrocytes in culture. J Neurosci Res 80: 226-234.
170. Pantano C, Reynaert NL, van d, V, Janssen-Heininger YM (2006) Redox- sensitive kinases of the nuclear factor-kappaB signaling pathway. Antioxid Redox Signal 8: 1791-1806.
171. Panter SS, Faden AI (1992) Pretreatment with NMDA antagonists limits release of excitatory amino acids following traumatic brain injury. Neurosci Lett 136: 165-168.
172. Park E, Liu Y, Fehlings MG (2003) Changes in glial cell white matter AMPA receptor expression after spinal cord injury and relationship to apoptotic cell death. Exp Neurol 182: 35-48.
173. Park E, Velumian AA, Fehlings MG (2004a) The role of excitotoxicity in secondary mechanisms of spinal cord injury: a review with an emphasis on the implications for white matter degeneration. J Neurotrauma 21: 754- 774.
174. Park M, Penick EC, Edwards JG, Kauer JA, Ehlers MD (2004b) Recycling endosomes supply AMPA receptors for LTP. Science 305: 1972-1975.
175. Perlman JM (1998) White matter injury in the preterm infant: an important determination of abnormal neurodevelopment outcome. Early Hum Dev 53: 99-120.
176. Peterson JW, Bo L, Mork S, Chang A, Ransohoff RM, Trapp BD (2002) VCAM-1-positive microglia target oligodendrocytes at the border of multiple sclerosis lesions. J Neuropathol Exp Neurol 61: 539-546.
129
177. Petralia RS, Esteban JA, Wang YX, Partridge JG, Zhao HM, Wenthold RJ, Malinow R (1999) Selective acquisition of AMPA receptors over postnatal development suggests a molecular basis for silent synapses. Nat Neurosci 2: 31-36.
178. Petrosillo G, Ruggiero FM, Paradies G (2003) Role of reactive oxygen species and cardiolipin in the release of cytochrome c from mitochondria. FASEB J 17: 2202-2208.
179. Pitt D, Nagelmeier IE, Wilson HC, Raine CS (2003) Glutamate uptake by oligodendrocytes: Implications for excitotoxicity in multiple sclerosis. Neurology 61: 1113-1120.
180. Pitt D, Werner P, Raine CS (2000) Glutamate excitotoxicity in a model of multiple sclerosis. Nat Med 6: 67-70.
181. Power J, Mayer-Proschel M, Smith J, Noble M (2002) Oligodendrocyte precursor cells from different brain regions express divergent properties consistent with the differing time courses of myelination in these regions. Dev Biol 245: 362-375.
182. Qin L, Li G, Qian X, Liu Y, Wu X, Liu B, Hong JS, Block ML (2005) Interactive role of the toll-like receptor 4 and reactive oxygen species in LPS-induced microglia activation. Glia 52: 78-84.
183. Raff MC, Miller RH, Noble M (1983) A glial progenitor cell that develops in vitro into an astrocyte or an oligodendrocyte depending on culture medium. Nature 303: 390-396.
184. Ramirez BG, Blazquez C, Gomez dP, Guzman M, de Ceballos ML (2005) Prevention of Alzheimer's disease pathology by cannabinoids: neuroprotection mediated by blockade of microglial activation. J Neurosci 25: 1904-1913.
185. Ray SK, Hogan EL, Banik NL (2003) Calpain in the pathophysiology of spinal cord injury: neuroprotection with calpain inhibitors. Brain Res Brain Res Rev 42: 169-185.
186. Ray SK, Matzelle DD, Wilford GG, Hogan EL, Banik NL (2000) Increased calpain expression is associated with apoptosis in rat spinal cord injury: calpain inhibitor provides neuroprotection. Neurochem Res 25: 1191- 1198.
187. Rebhun J, Botvin J (1980) Complement elevation in spinal cord injury. Ann Allergy 44: 287-288.
130
188. Renno T, Krakowski M, Piccirillo C, Lin JY, Owens T (1995) TNF-alpha expression by resident microglia and infiltrating leukocytes in the central nervous system of mice with experimental allergic encephalomyelitis. Regulation by Th1 cytokines. J Immunol 154: 944-953.
189. Reyners H, Gianfelici dR, Regniers L, Maisin JR (1986) A glial progenitor cell in the cerebral cortex of the adult rat. J Neurocytol 15: 53-61.
190. Reynolds R, Hardy R (1997) Oligodendroglial progenitors labeled with the O4 antibody persist in the adult rat cerebral cortex in vivo. J Neurosci Res 47: 455-470.
191. Ritz MF, Schmidt P, Mendelowitsch A (2004) Acute effects of 17beta- estradiol on the extracellular concentration of excitatory amino acids and energy metabolites during transient cerebral ischemia in male rats. Brain Res 1022: 157-163.
192. Rodriguez-Pena A (1999) Oligodendrocyte development and thyroid hormone. J Neurobiol 40: 497-512.
193. Rogister B, Ben Hur T, Dubois-Dalcq M (1999) From neural stem cells to myelinating oligodendrocytes. Mol Cell Neurosci 14: 287-300.
194. Rosenberg PA, Dai W, Gan XD, Ali S, Fu J, Back SA, Sanchez RM, Segal MM, Follett PL, Jensen FE, Volpe JJ (2003) Mature myelin basic protein- expressing oligodendrocytes are insensitive to kainate toxicity. J Neurosci Res 71: 237-245.
195. Rosin C, Bates TE, Skaper SD (2004) Excitatory amino acid induced oligodendrocyte cell death in vitro: receptor-dependent and -independent mechanisms. J Neurochem 90: 1173-1185.
196. Ryu JK, Nagai A, Kim J, Lee MC, McLarnon JG, Kim SU (2003) Microglial activation and cell death induced by the mitochondrial toxin 3- nitropropionic acid: in vitro and in vivo studies. Neurobiol Dis 12: 121-132.
197. Salter MG, Fern R (2005) NMDA receptors are expressed in developing oligodendrocyte processes and mediate injury. Nature 438: 1167-1171.
198. Sanchez-Gomez MV, Alberdi E, Ibarretxe G, Torre I, Matute C (2003) Caspase-dependent and caspase-independent oligodendrocyte death mediated by AMPA and kainate receptors. J Neurosci 23: 9519-9528.
199. Sanchez-Gomez MV, Matute C (1999) AMPA and kainate receptors each mediate excitotoxicity in oligodendroglial cultures. Neurobiol Dis 6: 475- 485. 131
200. Sarchielli P, Greco L, Floridi A, Floridi A, Gallai V (2003) Excitatory amino acids and multiple sclerosis: evidence from cerebrospinal fluid. Arch Neurol 60: 1082-1088.
201. Scolding NJ, Morgan BP, Campbell AK, Compston DA (1990) Complement mediated serum cytotoxicity against oligodendrocytes: a comparison with other cells of the oligodendrocyte-type 2 astrocyte lineage. J Neurol Sci 97: 155-162.
202. Scolding NJ, Rayner PJ, Compston DA (1999) Identification of A2B5- positive putative oligodendrocyte progenitor cells and A2B5-positive astrocytes in adult human white matter. Neuroscience 89: 1-4.
203. Selmaj KW, Raine CS (1988) Tumor necrosis factor mediates myelin and oligodendrocyte damage in vitro. Ann Neurol 23: 339-346.
204. Sfaello I, Baud O, Arzimanoglou A, Gressens P (2005) Topiramate prevents excitotoxic damage in the newborn rodent brain. Neurobiol Dis 20: 837-848.
205. Shaked I, Tchoresh D, Gersner R, Meiri G, Mordechai S, Xiao X, Hart RP, Schwartz M (2005) Protective autoimmunity: interferon-gamma enables microglia to remove glutamate without evoking inflammatory mediators. J Neurochem 92: 997-1009.
206. Sharief MK, Hentges R (1991) Association between tumor necrosis factor- alpha and disease progression in patients with multiple sclerosis. N Engl J Med 325: 467-472.
207. Shen HM, Pervaiz S (2006) TNF receptor superfamily-induced cell death: redox-dependent execution. FASEB J 20: 1589-1598.
208. Shin WH, Lee DY, Park KW, Kim SU, Yang MS, Joe EH, Jin BK (2004) Microglia expressing interleukin-13 undergo cell death and contribute to neuronal survival in vivo. Glia 46: 142-152.
209. Shuman SL, Bresnahan JC, Beattie MS (1997) Apoptosis of microglia and oligodendrocytes after spinal cord contusion in rats. J Neurosci Res 50: 798-808.
210. Smith J, Ladi E, Mayer-Proschel M, Noble M (2000a) Redox state is a central modulator of the balance between self-renewal and differentiation in a dividing glial precursor cell. Proc Natl Acad Sci U S A 97: 10032- 10037.
132
211. Smith T, Groom A, Zhu B, Turski L (2000b) Autoimmune encephalomyelitis ameliorated by AMPA antagonists. Nat Med 6: 62-66.
212. Sommer I, Schachner M (1981) Monoclonal antibodies (O1 to O4) to oligodendrocyte cell surfaces: an immunocytological study in the central nervous system. Dev Biol 83: 311-327.
213. Soto A, Perez-Samartin AL, Etxebarria E, Matute C (2004) Excitotoxic insults to the optic nerve alter visual evoked potentials. Neuroscience 123: 441-449.
214. Spitz DR, Azzam EI, Li JJ, Gius D (2004) Metabolic oxidation/reduction reactions and cellular responses to ionizing radiation: a unifying concept in stress response biology. Cancer Metastasis Rev 23: 311-322.
215. Springer JE, Azbill RD, Kennedy SE, George J, Geddes JW (1997) Rapid calpain I activation and cytoskeletal protein degradation following traumatic spinal cord injury: attenuation with riluzole pretreatment. J Neurochem 69: 1592-1600.
216. Sriram S, Rodriguez M (1997) Indictment of the microglia as the villain in multiple sclerosis. Neurology 48: 464-470.
217. Stellwagen D, Beattie EC, Seo JY, Malenka RC (2005) Differential regulation of AMPA receptor and GABA receptor trafficking by tumor necrosis factor-alpha. J Neurosci 25: 3219-3228.
218. Stidworthy MF, Genoud S, Suter U, Mantei N, Franklin RJ (2003) Quantifying the early stages of remyelination following cuprizone-induced demyelination. Brain Pathol 13: 329-339.
219. Streit WJ, Graeber MB, Kreutzberg GW (1988) Functional plasticity of microglia: a review. Glia 1: 301-307.
220. Strohmeyer R, Kovelowski CJ, Mastroeni D, Leonard B, Grover A, Rogers J (2005) Microglial responses to amyloid beta peptide opsonization and indomethacin treatment. J Neuroinflammation 2: 18.
221. Studer L, Csete M, Lee SH, Kabbani N, Walikonis J, Wold B, McKay R (2000) Enhanced proliferation, survival, and dopaminergic differentiation of CNS precursors in lowered oxygen. J Neurosci 20: 7377-7383.
222. Sun F, Lin C, McTigue DM, Shan X, Tovar C.A., Bresnahan J.C., Beattie MS (2006) Dorsal rhizotomy induces oligodendrocyte progenitor proliferation and oligodendrocyte genesis, while spinal contusion induces oligodendrocyte death. J Neurosci in revision . 133
223. Szeto HH (2006) Mitochondria-targeted peptide antioxidants: novel neuroprotective agents. AAPS J 8: E521-E531.
224. Takahashi JL, Giuliani F, Power C, Imai Y, Yong VW (2003) Interleukin- 1beta promotes oligodendrocyte death through glutamate excitotoxicity. Ann Neurol 53: 588-595.
225. Takano K, Nakamura Y, Yoneda Y (2003) Microglial cell death induced by a low concentration of polyamines. Neuroscience 120: 961-967.
226. Tan S, Wood M, Maher P (1998) Oxidative stress induces a form of programmed cell death with characteristics of both apoptosis and necrosis in neuronal cells. J Neurochem 71: 95-105.
227. Tanaka H, Grooms SY, Bennett MV, Zukin RS (2000) The AMPAR subunit GluR2: still front and center-stage. Brain Res 886: 190-207.
228. Tanaka K, Nogawa S, Suzuki S, Dembo T, Kosakai A (2003) Upregulation of oligodendrocyte progenitor cells associated with restoration of mature oligodendrocytes and myelination in peri-infarct area in the rat brain. Brain Res 989: 172-179.
229. Tekkok SB, Goldberg MP (2001) Ampa/kainate receptor activation mediates hypoxic oligodendrocyte death and axonal injury in cerebral white matter. J Neurosci 21: 4237-4248.
230. Temple S, Raff MC (1986) Clonal analysis of oligodendrocyte development in culture: evidence for a developmental clock that counts cell divisions. Cell 44: 773-779.
231. Thery C, Mallat M (1993) Influence of interleukin-1 and tumor necrosis factor alpha on the growth of microglial cells in primary cultures of mouse cerebral cortex: involvement of colony-stimulating factor 1. Neurosci Lett 150: 195-199.
232. Thomas AJ, Nockels RP, Pan HQ, Shaffrey CI, Chopp M (1999) Progesterone is neuroprotective after acute experimental spinal cord trauma in rats. Spine 24: 2134-2138.
233. Thorburne SK, Juurlink BH (1996) Low glutathione and high iron govern the susceptibility of oligodendroglial precursors to oxidative stress. J Neurochem 67: 1014-1022.
234. Virag L, Szabo E, Gergely P, Szabo C (2003) Peroxynitrite-induced cytotoxicity: mechanism and opportunities for intervention. Toxicol Lett 140-141: 113-124. 134
235. Volpe JJ (2001) Neurobiology of periventricular leukomalacia in the premature infant. Pediatr Res 50: 553-562.
236. Wajant H, Pfizenmaier K, Scheurich P (2003) Tumor necrosis factor signaling. Cell Death Differ 10: 45-65.
237. Walker MC, Sander JW (1996) Topiramate: a new antiepileptic drug for refractory epilepsy. Seizure 5: 199-203.
238. Wang AL, Yu AC, Lau LT, Lee C, Wu lM, Zhu X, Tso MO (2005) Minocycline inhibits LPS-induced retinal microglia activation. Neurochem Int 47: 152-158.
239. Wang CX, Reece C, Wrathall JR, Shuaib A, Olschowka JA, Hao C (2002) Expression of tumor necrosis factor alpha and its mRNA in the spinal cord following a weight-drop injury. Neuroreport 13: 1391-1393.
240. Warner DS, Sheng H, Batinic-Haberle I (2004) Oxidants, antioxidants and the ischemic brain. J Exp Biol 207: 3221-3231.
241. Watanabe M, Toyama Y, Nishiyama A (2002) Differentiation of proliferated NG2-positive glial progenitor cells in a remyelinating lesion. J Neurosci Res 69: 826-836.
242. Werner P, Pitt D, Raine CS (2000) Glutamate excitotoxicity--a mechanism for axonal damage and oligodendrocyte death in Multiple Sclerosis? J Neural Transm Suppl 375-385.
243. Werner P, Pitt D, Raine CS (2001) Multiple sclerosis: altered glutamate homeostasis in lesions correlates with oligodendrocyte and axonal damage. Ann Neurol 50: 169-180.
244. White CA, McCombe PA, Pender MP (1998) Microglia are more susceptible than macrophages to apoptosis in the central nervous system in experimental autoimmune encephalomyelitis through a mechanism not involving Fas (CD95). Int Immunol 10: 935-941.
245. Wilson HC, Onischke C, Raine CS (2003) Human oligodendrocyte precursor cells in vitro: phenotypic analysis and differential response to growth factors. Glia 44: 153-165.
246. Wisman LA, van Muiswinkel FL, de Graan PN, Hol EM, Bar PR (2003) Cells over-expressing EAAT2 protect motoneurons from excitotoxic death in vitro. Neuroreport 14: 1967-1970.
135
247. Wolswijk G, Noble M (1989) Identification of an adult-specific glial progenitor cell. Development 105: 387-400.
248. Wolswijk G, Riddle PN, Noble M (1990) Coexistence of perinatal and adult forms of a glial progenitor cell during development of the rat optic nerve. Development 109: 691-698.
249. Woodruff RH, Fruttiger M, Richardson WD, Franklin RJ (2004) Platelet- derived growth factor regulates oligodendrocyte progenitor numbers in adult CNS and their response following CNS demyelination. Mol Cell Neurosci 25: 252-262.
250. Wosik K, Ruffini F, Almazan G, Olivier A, Nalbantoglu J, Antel JP (2004) Resistance of human adult oligodendrocytes to AMPA/kainate receptor- mediated glutamate injury. Brain 127: 2636-2648.
251. Wrathall JR, Teng YD, Marriott R (1997) Delayed antagonism of AMPA/kainate receptors reduces long-term functional deficits resulting from spinal cord trauma. Exp Neurol 145: 565-573.
252. Xu GY, Hughes MG, Ye Z, Hulsebosch CE, McAdoo DJ (2004) Concentrations of glutamate released following spinal cord injury kill oligodendrocytes in the spinal cord. Exp Neurol 187: 329-336.
253. Xu H, Barks JD, Liu YQ, Silverstein FS (2001) AMPA-Induced suppression of oligodendroglial gene expression in neonatal rat brain. Brain Res Dev Brain Res 132: 175-178.
254. Yamaya Y, Yoshioka A, Saiki S, Yuki N, Hirose G, Pleasure D (2002) Type-2 astrocyte-like cells are more resistant than oligodendrocyte-like cells against non-N-methyl-D-aspartate glutamate receptor-mediated excitotoxicity. J Neurosci Res 70: 588-598.
255. Yang L, Jones NR, Blumbergs PC, Van Den HC, Moore EJ, Manavis J, Sarvestani GT, Ghabriel MN (2005a) Severity-dependent expression of pro-inflammatory cytokines in traumatic spinal cord injury in the rat. J Clin Neurosci 12: 276-284.
256. Yang Z, Watanabe M, Nishiyama A (2005b) Optimization of oligodendrocyte progenitor cell culture method for enhanced survival. J Neurosci Methods 149: 50-56.
257. Ye P, D'Ercole AJ (1999) Insulin-like growth factor I protects oligodendrocytes from tumor necrosis factor-alpha-induced injury. Endocrinology 140: 3063-3072.
136
258. Yoon BH, Romero R, Kim CJ, Koo JN, Choe G, Syn HC, Chi JG (1997) High expression of tumor necrosis factor-alpha and interleukin-6 in periventricular leukomalacia. Am J Obstet Gynecol 177: 406-411.
259. Yoshioka A, Hardy M, Younkin DP, Grinspan JB, Stern JL, Pleasure D (1995) Alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA) receptors mediate excitotoxicity in the oligodendroglial lineage. J Neurochem 64: 2442-2448.
260. Yuan X, Eisen AM, McBain CJ, Gallo V (1998) A role for glutamate and its receptors in the regulation of oligodendrocyte development in cerebellar tissue slices. Development 125: 2901-2914.
261. Zajicek JP, Wing M, Scolding NJ, Compston DA (1992) Interactions between oligodendrocytes and microglia. A major role for complement and tumour necrosis factor in oligodendrocyte adherence and killing. Brain 115 ( Pt 6): 1611-1631.
262. Zhao J, Brooks DM, Lurie DI (2006) Lipopolysaccharide-activated SHP-1- deficient motheaten microglia release increased nitric oxide, TNF-alpha, and IL-1beta. Glia 53: 304-312.
263. Zhao L, Brinton RD (2004) Suppression of proinflammatory cytokines interleukin-1beta and tumor necrosis factor-alpha in astrocytes by a V1 vasopressin receptor agonist: a cAMP response element-binding protein- dependent mechanism. J Neurosci 24: 2226-2235.
137