ACTIVIN IS CRITICAL for the DEVELOPMENT of PAIN HYPERSENSITIVITY AFTER INFLAMMATION by PIN XU Submitted in Partial Fulfillment

ACTIVIN IS CRITICAL FOR THE DEVELOPMENT OF PAIN HYPERSENSITIVITY

AFTER INFLAMMATION

PIN XU

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Thesis Adviser: Dr. Alison K. Hall

Department of Neurosciences

CASE WESTERN RESERVE UNIVERSITY

August, 2007

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the dissertation of

____Pin Xu______candidate for the Ph.D. degree *.

(signed)__ Gary Landreth______

(chair of the committee)

___Alison Hall______

___Jerry Silver______

___Susann Brady-kalnay______

______

(date) ____June 5, 2007______

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

contained therein.

DEDICATION

This thesis is dedicated to my parents, Hongfa Xu and Ruiyun Pan, and to my husband

Chen Liu.

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TABLE OF CONTENTS

Page

Title Page……………………..…………………………………………………….….…I

Typed ETD Sign-off Sheet……………………………………………………...... II

Dedication……………………………………………………………………………..…III

Table of Contents………………………………………………………..……..……..….IV

List of Tables……………………………………………………………………………VII

List of Figures…………………………………………………………………….....…VIII

Acknowledgements………………………………………………………………………X

Abstract…………………………………………………………………………...……...XI

Chapter I:

General Introduction……………………………………………………………………...1

Chapter II:

Activin induces tactile allodynia and increases CGRP upon peripheral inflammation…………………………………………………………………….………35

Abstract…………………………………………………………………………..36

Introduction………………………………………………………………………37

Methods and Materials…………………………………………………………...40

Results………………………………………………………………..…………..47

Summary…………………………………………………………………..……..54

Chapter III:

Activin acutely sensitizes DRG neurons and induces hyperalgesia via sensitize of

TRPV1………………………………………………………………………..…….……72

Abstract…………………………………………………………………….……..73

Introduction………………………………………………………………….……74

Methods and Materials…………………………………………………….……...76

Results………………………………………………………………..…………...80

Summary………………………………………………………………..…….…..83

Chapter IV:

Activin regulates CGRP expression in both embryonic and adult DRG neurons …………………………………………………………………………………..94

Abstract…………………………………………………………………….……..95

Introduction……………………………………………………………….………96

Methods and Materials…………………………………………………….……...98

Results………………………………………………………………..…….……101

Summary……………………………………………………………………..….110

Chapter V:

General discussion…………………………………………………….………………..131

Bibliography………………………………………………………….………………...155

LIST OF TABLES

Table No Page

Table 1.1. Family members, Activin A, BMPs and TGFβ signal through some shared receptors and signaling components………………………………………………….33

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LIST OF FIGURES

Figure No. Page

Figure 1.1. Sensory neurons respond to signals in skin for early differentiation, plasticity after wound and pain functions………………………………………………………....29

Figure 1.2. Activin and selected TGFβ family members, their receptors and signaling……………………………………………………………………….…..…....31

Figure 2.1. CFA induced inflammation. …………………………………………...…..58

Figure 2.2. Tracer labeled CGRP-IR neurons in the L4 DRG……………………..…...60

Figure 2.3. CGRP expression in different subpopulations of L4 DRG neurons………..62

Figure 2.4. The effect of inflammation on the size distribution of sensory neurons innervating ankle skin………………………………………………………………...... 64

Figure 2.5. Central projections of CGRP-IR fibers after localized inflammation………66

Figure 2.6. Activin A or NGF injection induces tactile allodynia…………………..…..68

Figure 2.7 Activin induces CGRP in the innervating DRG neurons………….……...... 70

Figure 3.1. Activin A acutely sensitizes the capsaicin induced current in the cultured

DRG neurons through the activin type I receptor AcRIB……………………….……....88

Figure 3.2. Activin A causes translocation of PKCδ to cell membrane of rat DRG neurons…………………………………………………………………………………..90

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Figure 3.3. Activin injection induces thermal hyperalgesia in wildtype, but not TRPV1 null mice……………………………………………………………………….………....92

Figure 4.1. Activin acts with NGF to induce CGRP mRNA in adult DRG cultures…...115

Figure 4.2. Activin effects on CGRP mRNA occur through its own receptor…………117

Figure 4.3. Activin signaling pathways in the adult sensory neurons……………….…119

Figure 4.4. Activin stimulates pSmad2 while NGF stimulates pERK2 in adult sensory neurons……………………………………………………………………….….…….. 122

Figure 4.5. NGF and activin signal through multiple MAPK pathways to regulate CGRP expression………………………………………………………………………….…...124

Figure 4.6. Activin increases CGRP mRNA in E14.5 DRG neuronal culture in a concentration dependent manner…………………………………….…………………126

Figure 4.7. Activin and NGF signaling pathways in embryonic sensory neurons…...... 128

ACKNOWLEGDEMENTS

I would like to thank my thesis advisor, Dr. Alison Hall. She taught me with great patience how to do science and design critical experiments. I have learned tremendously from her during my Ph.D. training. Also she set up an excellent model for me as a working mother and scientist. Alison always encourages me to pursue my personal goal and offers her suggestions whenever I meet problems. I am really grateful for her help.

I would also like to express my gratitude to members of my thesis committee, Drs.

Gary Landreth, Jerry Silver and Susann Brady-Kalnay for all of their supports and great advices on my thesis. I want to thank our collaborators, Dr. Liliana Berti-Mattera (CASE),

Dr. Andrew Russo (U. Iowa), Drs. Gerry Oxford and Weiguo Zhu (Stark Res. Inst.)

I appreciate the help from people in the Neurosciences department throughout my

Ph.D. training. Especially, I want to thank my current labmates, Shibani Mukerji and

Ekaterina Katsman, who make the lab life pleasant and I really enjoy our conversations about both scientific topics and personal life. I thank formal Hall lab residents, Bethany

Kiernan, Noah Haner and Rebecca Burke, who always offer their help.

Finally, I thank my family members for their endless love and support.

Activin is Critical for the Development of Pain Hypersensitivity after Inflammation

Abstract

PIN XU

Inflammatory pain is a major clinical challenge. The perception of pain is initiated by

specialized sensory neurons in the Dorsal Root Ganglia (DRG) called nociceptors. Upon injury, neuropeptides including Calcitonin Gene-Related Peptide (CGRP) are released from nociceptors and the neuropeptide contributes to the development of abnormal pain.

The studies in this thesis focus on the role of activin in regulating pain after skin

inflammation. In this study, experimental peripheral inflammation was accompanied by

increases in activin mRNA and activin immunoreactivity in both keratinocytes and

infiltrating immune cells. The number of CGRP containing DRG neurons increased after

inflammation, and this increase was among neurons that expressed the lectin IB4 or TrkA.

These data indicate that some adult sensory neurons remain plastic and can increase

CGRP expression after inflammation or activin administration. Strikingly, activin administration also induced thermal hyperalgesia and mechanical allodynia pain behaviors. I demonstrated that activin induced thermal hyperalgesia involved the sensitization of nociceptors through the Transient Receptor Potential Vallinoid subtype 1

(TRPV1). In addition to changing acute pain behaviors, activin also regulated neuropeptide gene transcription over chronic periods. Activin and nerve growth factor

(NGF) worked synergistically to increase CGRP mRNA in adult DRG cultures.

Biochemical and pharmacological studies indicate that this synergy was initiated by

independent stimulation of receptors and intracellular signals, and our data suggest that

activin and NGF effects converge on the CGRP promoter to regulate CGRP expression.

Taken together, these data demonstrate that increased activin expression after

inflammation results in both acute pain behaviors through sensitizing nociceptors and prolonged pain regulation by increasing CGRP expression.

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Chapter I

General Introduction

Inflammatory pain is a major medical challenge and can be difficult to manage clinically. Pain hypersensitivity is a key sign of inflammation and is not sufficiently controlled with commonly prescribed analgesics. Present studies demonstrate that activin

A (activin) regulates a group of specialized sensory neurons sensing nociceptive information. This thesis will focus on the functions of activin in regulating pain sensation.

In this section, a general introduction will review nociceptive sensation, neuropeptides

important for pain sensation, the regulation of neuropeptides by activin and NGF and current approach for pain management.

Perception is mediated by receptors, nervous pathways and specific areas in the

brain.

Our knowledge of the outside world starts from the senses which do not passively

record the surrounding world, but process information via brain activity leading to

perception. The sensory experience contains four basic types of information – modality,

location, intensity and timing, all of which are encoded in the nervous system by

specialized subgroups of neurons.

Modality is determined by the type of transmitted energy and the receptors specialized

to sense that energy. Sensory receptors transduce unique stimuli and convert them into

specific electrical signals, which are conducted to the cortex to produce different

perceptions. These sensory receptors include cutaneous mechanoreceptors, muscle and

joint receptors, cold and warm receptors and nociceptors in the somatosensory system.

Somatic sensibility has four major modalities: discriminative touch, proprioception,

nociception and temperature sense (Kandel, Schwartz et al. 2000).

All these modalities are conveyed by neurons in the Dorsal Root Ganglion (DRG),

which is located on the dorsal root of spinal nerves. One remarkable feature of DRG

neurons is their heterogeneity; they bear morphological distinct peripheral terminals and

contain different kinds of axons. Free nerve ending fibers are small-diameter; either

unmyelinated (C-fiber) or thinly myelinated (Aδ fiber). They transduce either pain or

thermal sensation, depending on the receptors expressed on the free nerve ending, at a

relatively lower speed (Croze, Duclaux et al. 1976; Ochoa and Torebjork 1989). Fibers

with encapsulated terminals are large or medium-diameter, myelinated (Aα, Aβ fiber) and

they transduce the sensation of touch and proprioception in a much faster way.

The location of perception is transferred by the spatial distribution of activated sensory receptors following a stimulus. The pattern of activated receptors conveys information about the site, size, shape and fine details of the stimulus. The intensity of a perception is coded by the firing rates of receptors and the timing is mediated by the duration of the firing, which is also partially determined by the adaptation rate of

receptors.

When a specific stimulus is applied to the body, morphologically specialized sensory

receptors are activated to generate receptor potential, which produce action potentials.

The burst of action potentials is conducted through first-order, and then higher-order

neurons and at each stage information is processed in relay nuclei, which determines

whether the information will be relayed to the cortex or not. When the information

reaches the cortex, it is finally translated into a specific sensation, such as touch, taste,

hearing, smell and vision.

How we sense pain

Pain is an unpleasant sensory and emotional experience associated with actual or

potential tissue damage (as defined by the International Association for the Study of

Pain). Pain is also important to protect the body by alerting and driving the behaviors that

remove or minimize the threat to avoid permanent tissue damage.

Our perception of pain is normally initiated in peripheral tissues such as skin that

contain axons of specialized sensory neurons whose development and function are

closely regulated. The pain sensors or nociceptors, including polymodal, thermal, and mechanical nociceptors, are activated by high intensity thermal, mechanical, or chemical noxious stimuli. This information is then conveyed from axons in the peripheral tissues,

like skin, muscle and joints, through neuronal cell bodies in the DRG to central synapses

within the spinal cord (Fig. 1.1) (Weddell 1955; Ignelzi and Atkinson 1980; Henry 1989).

The sensory afferent fibers predominantly project to the dorsal horn of the spinal cord, which contains multiple layers identified by the cytological feature of resident neurons.

Most nociceptive neurons project to the superficial layers of dorsal horn including the

marginal layer (lamina I) and substantia gelatinosa (lamina II). Aδ fibers project to

lamina I, V and the outer layer of lamina II to synapse with nociceptive-specific neurons

and wide dynamic-range neurons which respond to both noxious and nonnoxious

mechanical stimulation in a graded manner. C fibers mainly project to lamina II and

4 make connections with local interneurons which also connect to projection neurons in lamina I (Scott 1992). Aβ fibers terminate in Lamina III - VI and inner layer of lamina II and transmit nonnoxious information from low-threshold mechanoreceptor (Torebjork,

Vallbo et al. 1987). Wide dynamic-range neurons are the predominant neurons in lamina

V and they receive information conveyed by Aδ, Aβ and C fibers either through direct or indirect contacts via interneurons. Many neurons in lamina V also receive nociceptive information from the viscera.

In mammalian systems, nociceptors in the DRG utilize glutamate as a neurotransmitter to activate AMPA-type glutamate receptors and evoke fast excitatory postsynaptic potentials (EPSP). Further, they use one or more neuropeptides such as

CGRP and substance P that function as neuromodulators to evoke slow EPSP in second- order neurons in the dorsal horn of the spinal cord (De Biasi and Rustioni 1988). These spinal cord neurons project across the mid line and form the anterolateral tract. This ascending tract passes medulla, pons, midbrain and synapses with third-order neurons in the thalamus, which finally project to cortex. Although nociceptive information is conducted in such a specialized way, the perception of pain is not sensed until the higher brain centers process the passed-on information.

Pain is regulated by the central nervous system through modulating mechanisms. The first modulation happens in the spinal cord, also known as the gate control hypothesis

(Melzack and Wall 1965). This hypothesis suggests that the balance of activity in the nociceptive and nonnociceptive primary afferent fibers can modulate pain. The brain also uses descending inhibitory pathways to modulate pain. For example, the endogenous opioid peptides, including enkephalins, β-endorphine and dynorphin, are thought to

induce analgesia through descending inhibitory pathways (Morgan, Heinricher et al.

1992).

The perception of pain is very complicated but, practically, it can be measured using behavioral tests. Clinically, the commonly used method to measure acute pain is visual

analog scale (VAS) pain scores, which use a continuous spectrum of smiley faces to sad

faces to indicate from no pain, mild pain, and modest pain to very severe pain (Wewers

and Lowe 1990). In animal models, nociceptive behavior responses can be measured by

different behavioral assays, including the von Frey hair test and pin prick test for

mechanical nociceptive response and the hot-plate test, tail-flick test, Hargreaves test

(paw-flick test) for thermal nociceptive response (Crawley, Gerfen et al. 2006). In this

thesis studies some of these behavioral assays were used for detecting nociceptive

responses.

A subpopulation of dorsal root ganglion neurons responds to noxious stimuli

Our somatic sensations have four modalities, pain, touch, temperature and limb

proprioception. In the mature DRG, there are diverse subgroups of neurons responsible

for these modalities of sensation.

Different groups of sensory neurons are defined by their ability to effectively respond

to a particular excitatory stimulus such as nociceptors for pain, mechanoreceptors for

touch and proprioception and thermal receptors for temperature sensation.

Experimentally, they can also be identified by their morphological and

electrophysiological features. By Nissl staining, DRG neurons can be divided into two

major groups, large light (L) and small dark (SD) neurons (Andres 1961; Scott 1992).

Both L and SD neurons have a normal distribution of cell size, which overlap with each

other, with SD neurons being restricted to the lower end of the distribution (Lawson

1979). Experimentally, small neurons are defined arbitrarily by their cross-sectional area

containing nuclei up to 400µm2 and large neurons are those that have areas above

800µm2 (Fang, Djouhri et al. 2005). The nerve fibers of large and small neurons also

differ in conduction velocities. Myelinated cutaneous fibers include large (Aα), medium

(Aβ) and small (Aδ) fibers with Aα having the highest conduction velocity.

Unmyelinated cutaneous C fibers have smaller fiber diameter and slower conduction velocity. Generally, L neurons have A fibers while SD neurons bear C fibers (Lawson

and Waddell 1991). Moreover, functional different groups of DRG neurons also have

distinct firing properties. Most nociceptors are small diameter neurons with Aδ or C

fibers. Aδ fibers conduct information from thermal and mechanical nociceptors while C fibers transduce information from polymodal receptors. For example, when the thumb is hit by a hammer, the body first feels a sharp pain mediated by Aδ fibers and followed by

a more prolonged dull pain mediated by C fibers.

Nociceptors can also be further divided by their biochemical profile. In general, they

are divided into two almost equal groups, depending on the different neurotrophic sensitivity, NGF or glial cell line-derived neurotrophic factor (GDNF) responding nociceptors. Nociceptors responding to NGF express the tyrosine kinase receptor, TrkA.

Many of these neurons are also defined as “peptidergic” nociceptors, since they contain

peptides which are important for nociceptive sensation such as CGRP and substance P

(Verge, Grondin et al. 1992; Mu, Silos-Santiago et al. 1993; Averill, McMahon et al.

1995; Cao, Mantyh et al. 1998; Salmon, Damaj et al. 2001). CGRP-immunoreactive (IR)

neurons represent approximately 30% of rat DRG neurons and substance P is co-

expressed in a subset of CGRP-IR neurons (Gibbins, Furness et al. 1985; Lee, Takami et

al. 1985; Ju, Hokfelt et al. 1987; McCarthy and Lawson 1990). In this thesis, I mainly

focus on the CGRP-IR nociceptive neurons. By contrast, the remaining nociceptors

respond to GDNF and express c-Ret, and they can also be recognized by binding the

Griffonia simplicifolia isolectin B4 (IB4) (Molliver, Wright et al. 1997b; Bradbury,

Burnstock et al. 1998). These nociceptors generally do not express peptides and are defined as “nonpeptidergic” nociceptors. Other DRG neurons such as proprioceptors and low threshold mechanoreceptors can be identified by antibody RT97 binding to phosphorylated neurofilament protein (Lawson, Harper et al. 1984). Experimentally, these biochemical markers are widely used to discriminate between sensory neuron subtypes.

Role of neuropeptides in pain

Neuroactive peptides play many important roles in the nervous system, especially in sensory perception and emotions. These neuroactive peptides are made in the neuronal cell bodies and processed as secretory proteins. They are stored in large dense-core vesicles and moved to nerve terminals through fast axonal transport (McNeill, Coggeshall et al. 1988).

CGRP and substance P are key peptides that regulate the nociceptive response. CGRP was discovered in 1982 and is the most widely expressed neuropeptide in mammalian

sensory systems (Amara, Jonas et al. 1982). It is found in about 30% of vertebrate DRG

neurons among unmyelinated C and thinly myelinated Aδ fibers (Willis and Coggeshall

1991). Substance P is co-localized within a subset of CGRP-IR neurons. These neuropeptides represent not only operational markers for pain neurons, but their presence is also critical in pain perception, inflammation and wound healing (Brain 1997).

CGRP is a 37 amino acid peptide with α and β forms in rats, which correspond to

CGRP1 and CGRP 2 respectively in humans. In general, αCGRP and βCGRP co-express at variable ratios in the sensory nervous system. The protein ratio of αCGRP and βCGRP is 4:1 in the DRG and 3:1 in the spinal cord, and enteric neurons preferentially express more βCGRP (Mulderry, Ghatei et al. 1988). Alpha-CGRP and βCGRP are transcribed from two different genes and share >90% structural similarity (Amara, Arriza et al. 1985).

Alpha-CGRP is generated by RNA alternative splicing of αCGRP/calcitonin gene, which is transcribed in cells in both endocrine and nervous systems. After tissue-specific splicing, the mRNA encoding αCGRP is generated only in neuronal cells and the other product, calcitonin, is generated in the parafollicular cells of the thyroid gland which causes reduction in serum calcium (Amara, Jonas et al. 1982; Crenshaw, Russo et al.

1987; Leff, Evans et al. 1987). Beta-CGRP protein is not detected in the DRG and spinal cord in a αCGRP/calcitonin knock out mouse, suggesting that αCGRP might regulate the expression of βCGRP (Gangula, Zhao et al. 2000; Zhang, Hoff et al. 2001). However, this hypothesis is contradicted by a similar mouse model in which the coding region of

CGRP/calcitonin is replaced by a β-gal reporter. Although there is no detectable αCGRP expression in that mouse, the expression of βCGRP is normal at least in sensory neurons

(Schutz, Mauer et al. 2004). The discrepancy may be due to different genetic

backgrounds. Nevertheless, both of the genetic null mice exhibit reduced hypersensitivity

after inflammation which suggest that αCGRP and βCGRP have similar functions in

respect to nociceptive behavior (Salmon, Damaj et al. 2001; Zhang, Hoff et al. 2001).

CGRP receptors have two subtypes, CGRP1 and CGRP2 receptors, both of which are

G-protein coupled receptors (Quirion, Van Rossum et al. 1992; Waugh, Bockman et al.

1999). CGRP1 receptor mediates the majority of CGRP functions and is widely

expressed throughout the vasculature and nervous systems (Wimalawansa 1996;

Edvinsson 2004). The CGRP1 receptor can be specifically inhibited by an antagonist,

CGRP8-37, a truncated peptide that binds to CGRP1 receptor without initiating any

function and is widely used for testing the role of CGRP and its receptor (Chiba,

Yamaguchi et al. 1989; Dennis, Fournier et al. 1990). On the other hand, a second, less

prevalent receptor, CGRP2, is found in the vas deferens where it function to inhibit the

twitch responses and is less sensitive to agonist CGRP8-37 (Wisskirchen, Burt et al. 1998;

Poyner, Sexton et al. 2002).

CGRP has been shown to contribute to an increasing number of biological activities

(Brain and Cambridge 1996). CGRP was first found as a potent vasodilator affecting

blood flow and blood pressure (Brain, Williams et al. 1985; Brain, Tippins et al. 1986;

Marshall, Al-Kazwini et al. 1988; Gangula, Zhao et al. 2000). Indeed, CGRP-IR sensory

fibers innervate skin, muscle, joint, viscera, and most of the time they are in close vicinity to blood vessels. Further, in peripheral skin, CGRP1 receptor immunoreactivity is present in arteriolar smooth muscle, venular endothelium and capillary endothelium, which is

consistent with its function in blood vasodilation (Hagner, Haberberger et al. 2002).

CGRP participates in the nociceptive response. In the spinal cord, CGRP-IR sensory fibers terminate in the superficial layers of the dorsal horn in the spinal cord. CGRP receptors are expressed in the neurons of the superficial dorsal horn (Ye, Wimalawansa et al. 1999). These expression patterns suggest that CGRP signaling is involved in mediating nociceptive information. In the context of skin injury, CGRP is released both centrally to convey pain information, and peripherally from free nerve endings to cause peripheral vasodilation and release serum proteins in the injury area contributing to recovery (Brain, Tippins et al. 1986).

CGRP plays only a limited role in normal baseline pain responses. Two different kinds of αCGRP null mice all show normal basal nociceptive responses to mechanical and thermal stimuli (Salmon, Damaj et al. 2001; Zhang, Hoff et al. 2001). This idea is further supported by the data that intrathecal injection of the agonist, CGRP8-37, does not affect the basal mechanical response in the rats and subcutaneous injection CGRP into the skin does not induce nociceptive responses in rat or human (Nakamura-Craig and Gill

1991; Pedersen-Bjergaard, Nielsen et al. 1991; Sun, Lawand et al. 2003). However, there might be a basal release of CGRP in the spinal cord, which is not affected by innocuous mechanical stimuli. On the other hand, electrical stimulation which is sufficient to excite

C fibers or noxious thermal and mechanical stimuli can significantly increase this basal release of CGRP (Morton and Hutchison 1989; Schaible, Freudenberger et al. 1994).

CGRP release in the spinal cord plays a key role in inflammation-induced pain hypersensitivity (see below and (Sun, Lawand et al. 2003)), which suggests the amount of released CGRP needs to reach a threshold to mediate nociceptive response and this threshold is reached after inflammation (Seybold, Galeazza et al. 1995). For example, C

fibers fire repetitively under severe injury and the response of the projection neurons in

dorsal horn increases progressively, a phenomenon called “wind up”, which is an

example of central sensitization (Herrero, Laird et al. 2000). It is conceivable that

repeated firing of C fibers releases CGRP which builds up quickly to reach the threshold

and this process contributes to projection neuron hypersensitivity.

One report suggests that CGRP may affect the basal thermal response in a special

mouse strain. This report shows that there are different basal CGRP expression levels among different mice strains and mice that express lower levels of CGRP, such as AKR mice, show increased thermal nociceptive response after exogenous CGRP administration.

Other mice, like C57BL/6 mice, which have high endogenous levels of CGPP show no response to the same amount of exogenous CGRP. Furthermore, this difference is only seen in the thermal nociceptive response but not the mechanical nociceptive response

(Mogil, Miermeister et al. 2005). However, it is not known what regulates this CGPR sensitivity in AKR strain. In conclusion, CGRP is generally not required for normal baseline pain responses for rats and mice.

CGRP plays a key role in plastic changes occurring in the development of abnormal pain, including allodynia and hyperalgesia (Seybold, Galeazza et al. 1995; Brain 1997;

Zhang, Hoff et al. 2001; Jang, Nam et al. 2004; Ruiz and Banos 2005)(and see below).

Allodynia is defined as “pain due to a stimulus which does not normally provoke pain” and hyperalgesia is defined as “increased response to a stimulus which is normally painful” (IASP, 1994). According to the location of the hyperalgesia, it is further classified into primary hyperalgesia where the pain hypersensitivity occurs at the injury sites and secondary hyperalgesia where the pain hypersensitivity occurs at a distance

12 away from the injury sites. The CGRP antagonist, CGRP8-37 alleviates both mechanical and thermal allodynia in a chronic neuropathic pain model induced by spinal hemisection suggesting that CGRP signaling is required for the development of neuropathic pain

(Bennett, Chastain et al. 2000). Further the antagonist retards the initiation and diminishes secondary mechanical allodynia and hyperalgesia induced by capsaicin caused inflammation (Sun, Lawand et al. 2003). Anti-CGRP antibody delivered intrathecally also suppresses the carrageenan-induced hyperalgesia (Satoh, Kuraishi et al.

1992). These data strongly argue that CGRP is required for the development of abnormal pain in different pathologies.

CGRP functions in abnormal pain are further revealed by the studies performed in alpha-CGRP null mice. These mice present an attenuated abnormal pain response caused by chemical insults which induce inflammation, such as capsaicin, formalin, acetic acid and complete Freud’s adjuvant (CFA) (Salmon, Damaj et al. 2001; Zhang, Hoff et al.

2001). These data further support that CGRP is required for the development of abnormal pain. The central mechanism why CGRP is important for inflammation-induced allodynia and hyperalgesia may include direct sensitization of dorsal horn neurons, like wide dynamic range neurons and substantia gelatinosa neurons (Neugebauer, Rumenapp et al.

1996; Yoshinaga, Inoue et al. 2004; Bird, Han et al. 2006). In this thesis, CGRP regulation will be studied extensively.

CGRP also has functions in immune responses such as modulating antigen presentation in Langerhans cells in the skin (Hosoi, Murphy et al. 1993; Torii, Tamaki et al. 1998) and degranulating dural mast cells to release cytokines and inflammatory agents which contribute to migraine pathogenesis (Theoharides, Donelan et al. 2005).

Substance P was the first neuropeptide discovered and was recognized as a sensory neurotransmitter by Lembeck in 1953 (US and Gaddum 1931; Lembeck 1953). It is a member of tachykinin family whose best known members are substance P, neurokinin A and neurokinin B (Satake and Kawada 2006). Members of the tachykinin family are widely expressed in the mammalian peripheral and central nervous system (Pennefather,

Lecci et al. 2004). Substance P is exclusively found in small-diameter neurons in the

DRG and is released from sensory nerve endings after nociceptive stimulation (Hokfelt,

Kellerth et al. 1975; Sicuteri, Fanciullacci et al. 1990; Tiseo, Adler et al. 1990).

Tachykinins function through three types of receptors, NK1, NK2, NK3, which are all G protein-coupled receptors. Substance P preferentially acts through NK1, which is widely expressed in neurons, vascular endothelial cells, muscle and immune cells and superficial layers of the dorsal horn in the spinal cord (Pennefather, Lecci et al. 2004). Their expression patterns in the spinal cord suggest possible role in nociceptive information transmission.

Substance P is a less potent vasodilator compared to CGRP (Brain and Cox 2006) but causes plasma extravasation and degranulates mast cells during neurogenic inflammation

(Foreman, Jordan et al. 1983; Skofitsch, Donnerer et al. 1983). In contrast to CGRP, substance P is required for the basal pain response, as all substance P null mice show deficits in response to moderate and intense mechanical and thermal stimuli (Cao,

Mantyh et al. 1998; Zimmer, Zimmer et al. 1998). Whether substance P contributes to abnormal pain is unclear. Studies in these null mice have shown no deficits in hyperalgesia in CFA model and a neuropathic pain model (Cao, Mantyh et al. 1998;

Zimmer, Zimmer et al. 1998). In conclusion, although both CGRP and substance P are

14 required for central sensitization (see below), they perform different functions in different situations, and CGRP plays a more important role in inflammation-induced hyperalgesia.

Both CGRP and substance P play important roles in neurogenic inflammation.

Stimulation of nerves in target skin causes nerve firing to transmit nociceptive information to the spinal cord in the orthodromic direction. Further it also elicits an antidromic signal transmitted towards the peripheral nerve endings which releases vesicles into the innervated skin, a phenomenon called “axon reflex” (Schmelz and

Petersen 2001). The released vesicles contain neuropeptides including CGRP and substance P. When they are released into the skin by the axon reflex, a classical ‘triple response’ results in the skin (Lembeck and Holzer 1979). This response includes heat, redness (both due to CGRP induced vasodilation), and swelling (due to substance P induced plasma extravasation), accompanied by nociceptive hypersensitivity, all of which are classical signs of inflammation. In this example, inflammation is initiated by the axon reflex and thus represents neurogenic inflammation, defined as an inflammation caused by substances released from primary sensory nerve terminals (Richardson and Vasko

2002). It is believed that neurogenic inflammation has evolved as part of the natural defense to accelerate tissue repair (Brain 1997), but it may also contribute to chronic inflammatory pain in some diseases, like arthritis, migraine and asthma (Kidd, Photiou et al. 2004; Durham 2006; Nassenstein, Kutschker et al. 2006).

Nociceptive neuron phenotype is regulated during development

One hallmark characteristic of nociceptive neurons is the expression of neuropeptides

such as CGRP and substance P which are important for their physiological functions.

How nociceptors acquire their biochemical phenotype during development may include

signals from target tissues and intrinsic signaling mechanisms (Patel, Jackman et al. 2000;

Hall, Burke et al. 2002; Chen, Broom et al. 2006).

Nociceptive neurons require target derived factors for full phenotype differentiation.

Neurotrophins such as NGF, brain derived neurotrophic factor (BDNF), neurotrophin 3

(NT 3) and neurotrophin 4 (NT4) play key roles in sensory neuron development (Snider

1994; Snider and Silos-Santiago 1996). In particular, NGF and NT3 regulate sensory neuroblast proliferation in early neurogenesis and later supports selected neuronal

survival (Memberg and Hall 1995). Interestingly, in animals that lack the proapoptotic

BCL-2 homolog BAX, as well as NGF, natural cell apoptosis is inhibited and many sensory neurons survive. These neurons extend axons, but fail to make appropriate superficial cutaneous projections and lack biochemical markers such as CGRP that are characteristic of nociceptive neurons. These studies suggest that peripheral target contact is required for the phenotypic differentiation of sensory neurons (Patel, Jackman et al.

2000). During development, CGRP expression can be detected at E18-E19 in rats, which

is after the nerve innervating target tissues, and all nociceptive cells have been born by

then (Lawson and Biscoe 1979; Kitao, Robertson et al. 1996). These studies provide

further evidence support target-derived factors regulating CGRP expression in

nociceptors.

Sensory neuron phenotype is also regulated by intrinsic mechanisms. Postnatally, half

of early NGF responding neurons downregulate TrkA and upregulate GDNF receptor, c-

Ret (Silos-Santiago, Molliver et al. 1995; Molliver and Snider 1997; Molliver, Wright et

16 al. 1997). This transition is regulated by a Runt domain transcription factor, Runx1 (Chen,

Broom et al. 2006). Runx1 is initially expressed in 88% TrkA-IR neurons and as development continues, Runx1 expression decreases in TrkA positive neurons which also express neuropeptides including CGRP and is maintained in GDNF responding nociceptors. In these GDNF-responding neurons, Runx1 has been suggested to suppress

CGRP, the opioid receptor MOR, the proton gated channel DRASIC expression and promote the expression of TRPs (TRPA1, TRPM8, TRPC3, TRPV1, and TRPV2), P2X3, and Nav1.9. How Runx1 expression decreases in the TrkA-IR neurons and is sustained in c-Ret-IR neurons is not known. In addition, Runx3 has been shown to promote the differentiation of proprioceptors through decreasing TrkB expression in prospective TrkC sensory neurons (Kramer, Sigrist et al. 2006).

In summary, sensory neuron phenotype is determined by both intrinsic and target- derived signals. The intrinsic signals determine which cells are capable to respond to target-derived signals and the availability of target-derived factors finally resolve the ultimate size of neuronal population to obtain a certain sensory phenotype.

The role of activin in sensory development

The Transforming Growth Factor (TGF)-β family member activin has recently been implicated in neural development. During sensory neuron development, activin from peripheral target tissues can regulate pain neuropeptide expression.

Activin is initially identified as a regulator of pituitary follicle-stimulating hormone in the reproductive axis (Vale, Rivier et al. 1986) and has later been implicated to have

many roles in the nervous system. For example, studies suggest activin is a survival

factor for rodent CNS hippocampal and cortical neurons (Schubert, Kimura et al. 1990;

Iwahori, Saito et al. 1997; Mukerji, Katsman et al. 2006). More recently, activin

biological activity has emerged in another surprising arena: the specification and

differentiation of neurons.

Studies on developing rodent sensory neurons point to activin as a skin-derived factor

that induces de novo CGRP expression (Ai, Cappuzzello et al. 1999; Hall, Dinsio et al.

2001; Hall, Burke et al. 2002). Early embryonic (E14) rat DRG contains no detectable

CGRP mRNA or protein, although after peripheral connections are functional neuropeptides become apparent and increase (E18-19) (Kessler and Black 1981; Senba,

Shiosaka et al. 1982; Marti, Gibson et al. 1987; Hall, Ai et al. 1997). These data point to

target-derived factors in the development of subsets of neuropeptide-containing sensory

nociceptors. Not only is the appearance of neuropeptides coincident with target contact,

but the co-culture of embryonic DRG neurons with an embryonic skin cell line or

treatment with skin conditioned medium from a skin cell line induces de novo CGRP expression (Hall, Ai et al. 1997; Hall, Dinsio et al. 2001). Application of recombinant activin specifically induces neuropeptide expression in naïve embryonic rat sensory neurons in a concentration dependent manner (Ai, Cappuzzello et al. 1999). Furthermore, anti-activin antibody blocked the ability of skin-conditioned medium to induce CGRP expression in embryonic neurons suggesting activin is the active ligand in the skin (Hall,

Dinsio et al. 2001). Indeed, embryonic skin contains activin protein suggesting activin in

the target tissue may affect CGRP expression in the ganglion. In this thesis, the signaling

mechanisms involved in activin inducing CGRP expression in the embryonic sensory

neuron culture were studied.

NGF and activin signaling in the nervous system

NGF signals through TrkA and p75NTR, a member of the tumour-necrosis factor

receptor superfamily (Frade and Barde 1998). When bound to NGF dimers, TrkA dimers

get activated through phosphorylation and initiate multiple parallel downstream signaling

pathways including mitogen-activated protein kinase (MAPK), phosphatidylinositol 3-

kinase (PI3K) and phospholipase C-γ (PLCγ) pathways. There are four distinct groups of

MAPKs that have been characterized in mammals: extracellular signal-regulated kinases

(ERKs), c-Jun N-terminal kinases (JNKs), p38 isoforms and ERK/big MAP kinase 1

(BMK1) (Zarubin and Han 2005). In neurons, MAPK pathways are involved in synaptic

plasticity, long-term potentiation, neurite outgrowth and survival (Grewal, York et al.

1999). The PI3K pathway is involved in cell survival (Dudek, Datta et al. 1997; Crowder

and Freeman 1998). The PLCγ pathway regulates Ca2+, which, as a second messenger,

has multiple roles in the nervous system, such as regulation of axonal growth, growth-

cone guidance, synapse formation and neurotransmitter release (Cockcroft 2006). NGF

plays a key role in pain and temperature perception. In a large family from northern

Sweden, individuals lack deep pain and temperature perception, a mutation is found in the coding region of the NGFβ gene (Einarsdottir, Carlsson et al. 2004).

Like other members of the TGF-β superfamily, activins are dimeric proteins

consisting of disulphide-linked βA and βB subunits forming homodimeric activin A (βA-

βA), activin B (βB-βB) and heterodimeric activin AB (βA-βB) (Massague 2000). Activin

A is the focus of this thesis. Activins signal through type I receptor (called activin

receptor like kinases or ALK4/ActRIB) and type II receptor (activin receptor IIA and

ActRIIB), both of which are Ser/Thr kinases. Activated type II receptor further stimulate downstream Smad pathways which include receptor-regulated Smad (R-Smad,

Smad1,2,3,5 or 8), the co-mediator Smad (co-Smad, Smad4) and at least two inhibitory

Smads (I-Smad, Smad6 and 7; Fig 1.2) (Massague 2000). When activin binds to ActRII,

ActRIB is recruited to form the 'six-chain complex' due to the dimeric nature of the ligands (Greenwald, Vega et al. 2004; Lin, Lerch et al. 2006). After the complex is formed, the constitutively active ActRII trans-phosphorylates and activates ActRIB,

which further phosphorylates Smad2 or Smad3 which are recruited to the receptor by

Smad Anchor for Receptor Activation (SARA) (Tsukazaki, Chiang et al. 1998;

Panopoulou, Gillooly et al. 2002). Phosphorylated Smad2 or Smad3 then binds to Smad4

to form a heteromeric complex, which is translocated into the nucleus and, in conjunction

with other nuclear binding proteins, regulates the transcription of target genes (Chen,

Lebrun et al. 1996; Feng and Derynck 2005; Massague, Seoane et al. 2005).

Activin signaling can be regulated at multiple levels, including ligands, receptors,

intracellular signaling molecules, nucleus translocation and negative feedback

mechanisms (Massague and Chen 2000; Massague, Seoane et al. 2005; Massague and

Gomis 2006). In this thesis, both NGF and activin signals were studied to understand the

regulation of CGRP expression in the sensory neurons.

Nociceptor sensitization includes NGF sensitizing the TRPV1 channel

Inflammation after skin wounds or infections is a clinical challenge characterized by cell infiltration, release of mediators by inflammatory and damaged cells and increased pain sensitivity. Multiple inflammatory mediators increase the transduction sensitivity of nociceptors resulting in abnormal pain (Dray 1995; Katz and Gold 2006). In particular,

NGF is well known to cause both acute and prolonged mechanical and thermal hyperalgesia (Lewin, Ritter et al. 1993; Andreev, Dimitrieva et al. 1995; Amann,

Schuligoi et al. 1996; Bennett, al-Rashed et al. 1998; Gould, Gould et al. 2000). NGF increases after inflammation and stimulates neuropeptide synthesis in DRG neurons, which makes it possible that more neuropeptides will be released to facilitate afferent transmission to cause abnormal pain (Lindsay and Harmar 1989; Urban, Thompson et al.

1994).

Recent studies suggest that NGF can sensitize transient receptor potential vanilloid receptor subtype 1 (TRPV1) to reduce the threshold of nociceptors involving in temperature sensation. TRPV1 is a member of TRP family of ion channels. It is a nonselective cation channel gated by capsaicin (a main pungent ingredient in "hot" chili peppers), protons (extracellular pH<6) and heat (>43°C) (Tominaga, Caterina et al. 1998), all of which are enough to excite nociceptors and evoke pain in vivo (Steen, Reeh et al.

1992; Wall and Melzack 1999). In this way TRPV1 functions as an integrator of both physical and chemical stimuli. TRPV1 is selectively expressed by small to medium diameter neurons in DRG, either peptidergic or nonpeptidergic nociceptors. Also, strong

TRPV1 immunoreactivity is observed in the terminals of afferent fibers projecting to the superficial layers of the dorsal horn (Caterina, Schumacher et al. 1997; Tominaga,

Caterina et al. 1998). The TRPV1 expression profile suggests that it may function in

mediating nociception. Indeed, the burning sensation we feel after having spicy food is

mediated by capsaicin which activates TRPV1 and then triggers action potential firing of

sensory neurons, releasing glutamate and CGRP (Holzer 1991; Mogil, Miermeister et al.

2005). TRPV1 null mice show normal responses to noxious mechanical stimuli and

normal conduction velocities of C and Aδ fibers, but lose capsaicin response and thermal

hyperalgesia after inflammation (Caterina, Leffler et al. 2000; Davis, Gray et al. 2000).

TRPV1 is a nonselective cation channel which is predicted to have six transmembrane

domains and a short pore-forming hydrophobic region between the fifth and sixth

transmembrane domain. When TRPV1 gets activated, cells exhibit a time- and Ca2+ dependent outward current followed by a long-lasting refractory state, during which the channel is desensitized and does not respond to further stimuli (Tominaga, Caterina et al.

1998). This slow desensitization provides a potential mechanism for regulating peripheral nociceptors under different pathological conditions. Indeed, a variety of proinflammatory factors released from injured tissue after inflammation such as bradykinin, NGF, prostaglandins, serotonin, histamine, TNFα, ATP, and cations sensitize TRPV1(Cesare and McNaughton 1996; Nicol, Lopshire et al. 1997; Lopshire and Nicol 1998; Shu and

Mendell 1999; Hu, Bhave et al. 2002; Kim, Lee et al. 2004; Sugiuar, Bielefeldt et al.

2004; Ahern, Brooks et al. 2005). Sensitization reduces the temperature threshold to activate TRPV1, and this sensitization makes normal body temperature possibly capable of activating nociceptive neurons (Tominaga, Caterina et al. 1998; Vyklicky, Vlachova et al. 1999; Liang, Haake et al. 2001).

NGF has been shown to act through TrkA to rapidly sensitize TRPV1 in sensory

neurons (Shu and Mendell 2001; Bonnington and McNaughton 2003). In this thesis, whether activin required TRPV1 to cause thermal hyperalgesia was studied.

Inflammation increases CGRP, changes neuronal phenotype and pain responses

Under certain pathological circumstances, the normal arrangement of the information delivered by DRG neurons changes. A well-known case is after inflammation, when more sensory neurons show CGRP and substance P immunoreactivity accompanied by decreased nociceptive threshold and increased response to a nociceptive stimulus

(Bessou and Perl 1969; Willis and Cornelsen 1973; Kniffki, Mense et al. 1978; Sluka and

Westlund 1993).

The inflammatory response is part of the innate immune system and it includes the activities of vascular and immune systems. During inflammation, phagocytes remove bacteria, as well as damaged cells and tissues to promote healing. Without inflammation, the wound and infection might never heal, jeopardizing the survival of the organism. On the other hand, the inflammation response itself has the potential to cause harm, such as sepsis.

There are two types of inflammation, acute and chronic inflammation. Acute inflammation is usually a short lived process lasting from minutes up to days and has classic signs including heat, redness, swelling, pain and loss of function. When tissue is injured, cytokines and other chemicals are released from the blood vessels and

neutrophils migrate into the injury area to clear the initial injurious agent. When the

harmful stimuli are removed, the inflammation stops. Acute inflammation is a protective

mechanism but under certain pathological conditions, it can become a prolonged procedure lasting for weeks, months or even years and itself becomes a threat, also known as chronic inflammation. Chronic inflammation does not necessarily present the classic signs of inflammation, but it features in the infiltration of lymphocytes and

macrophages (Cotran, Robbins et al. 1998). Chronic inflammation commonly arises from

persistent infections, prolonged exposure to potential toxic agents and autoimmune

diseases. Normally acute pain comes with the acute inflammation and chronic pain

accompanies the chronic inflammation.

Pain can be similarly classified into two kinds depending on its time span, acute pain

and chronic pain. Acute pain is a protective mechanism, which disappears when the

injury is eliminated and chronic pain is the disease of pain which is persistent. Acute pain

can be both normal and abnormal; however, chronic pain is abnormal. Abnormal pain has

two kinds, allodynia and hyperalgesia, both of which can rise from chronic inflammation

and nerve injury and are resistant to most medical treatments. Although these

phenomenon have long been observed, the mechanism that accounts for pain

hypersensitivity is not clear (Hardy, Wolff et al. 1950).

Both allodynia and hyperalgesia are key signs of inflammation. These abnormal pain

phenomena have both peripheral and central origins. In the periphery, following

inflammation, substances such as potassium, serotonin, bradykinin, prostaglandins and

others from damaged cells or infiltrating immune cells can activate or sensitize

24 nociceptive neurons which decrease the threshold for activating nociceptive neurons, i.e. a benign stimulus become painful (Katz and Gold 2006). The central mechanism of abnormal pain involves the hyperexcitability of dorsal horn neurons, such as “wind up” which can be attenuated by blocking N-methyl-D-aspartate (NMDA)-type glutamate receptor, neurokinin (NK) receptors and CGRP receptor in the spinal cord under different pain pathology (Malmberg and Yaksh 1992; Ren, Williams et al. 1992; Thompson, Dray et al. 1994; Barbieri and Nistri 2001; Davis and Inturrisi 2001; Yashpal, Fisher et al. 2001;

Sun, Tu et al. 2004).

In tactile allodynia, a mechanism involving Aβ fibers has also been proposed

(Campbell, Raja et al. 1988; Ochoa and Yarnitsky 1993; Neumann, Doubell et al. 1996).

It is suggested that Aβ fibers which normally process information coming from low threshold mechanoreceptors change their modality to transmit nociceptive information during inflammation. Evidence shows that inflammation increases the excitability of Aβ neurons and more Aβ afferents express substance P after inflammation (Dubner and

Ruda 1992; Neumann, Doubell et al. 1996; Djouhri and Lawson 1999; Xu and Zhao

2001). However, whether the increased excitability of Aβ neurons or increase of substance P in Aβ afferents enhances nociception is not known. New synaptogenesis has also been suggested to partially contribute to the sensory modality change seen after inflammation. For example the reorganization of synaptic connections of low-threshold mechanoreceptors in the spinal cord can cause touch information transmitted by Aβ to be processed as nociception due to the altered synaptic connectivity (Maihofner, Neundorfer et al. 2003; Lewin and Moshourab 2004). Among these mechanisms, new synaptogenesis is much more likely to be the key one for the development of tactile allodynia.

After inflammation, DRG neurons undergo substantial changes in neuropeptide expression. Inflammation increases CGRP mRNA and protein levels and release in the inflamed skin, sciatic nerve, innervating DRG and spinal cord (Smith, Harmar et al. 1992;

Donnerer, R.Schuligoi et al. 1993; Nahin and Byers 1994; Schaible, Freudenberger et al.

1994; Galeazza, Garry et al. 1995; Seybold, Galeazza et al. 1995; Neumann, Doubell et al.

1996; Mulder, Zhang et al. 1997; Calza, Pozza et al. 2000; Bulling, Kelly et al. 2001).

When CGRP is released in the periphery, it causes vasodilation to promote healing.

When it is released centrally, it is involved in the development of abnormal pain (also see above). In this thesis, the mechanisms of abnormal pain that developed after inflammation are studied.

Current pain relief drugs

The commonly prescribed analgesics include opioids, steroids, and nonsteroidal anti- inflammatory drugs (NSAIDS).

Opioid receptors located in the descending inhibitory pathways which are used by brain to modify pain and opioids function through these pathways to induce analgesia.

Opioid analgesics (narcotics), such as morphine, diamorphine and codeine are widely used to relieve acute pain such as post operative pain and chronic pain such as cancer pain (Caimi and Cymet 2006; McCleane and Smith 2007; Myles and Power 2007).

However, they produce significant side effects, such as respiratory depression, physical dependence and sedation. Furthermore, their utilization in neuropathic pain is limited because high dosages are required to relieve pain, probably due to reduced opioid receptor expression in the DRG after peripheral nerve injury (Rashid, Inoue et al. 2004).

Therefore, although opioids are the most powerful analgesics, they are not universally useful for all kinds of pain and their usage is strictly controlled.

During inflammation, two mediators causing pain are prostaglandins and bradykinin.

Prostaglandin production can be inhibited by NSAIDS. Prostaglandins are metabolism

products of phospholipids. Cell membrane phospholipids are catalyzed by phospholipases

to form arachidonic acid (AA), which can be further metabolized through two different

pathways. One is cyclooxygenase (COX) pathway which AA is catalyzed by COX to

produce prostaglandins and thromboxane and the other is lipoxygenase pathway which

AA is catalyzed by 5-lipoxygenase to produce leukotrienes.

Most NSAIDS are COX blockers such as aspirin and ibuprofen and thus inhibit all

prostaglandins and thromboxane synthesis. There are two major forms of COX, one is

COX-1 which is expressed in gastric mucosa where prostaglandins prevent acid-induced

damage. The other is COX-2 which is primarily expressed at sites of inflammation and

produces prostaglandins involved in pain (Adelizzi 1999). The traditional NSAIDS, such

as aspirin, ibuprofen (Advil®, Motrin®) and naproxen sodium (Aleve®) target both

COX-1 and COX-2, and have side effects such as peptic ulcers and stomach bleeding.

New types of NSAIDS which only target COX-2 have been developed in order to avoid these side effects, such as Celecoxib (Celebrex®), Valdecoxib (Bextra®) and Rofecoxib

(Vioxx®). Although they did show reduced side effects on peptic ulcers, all these COX-2 specific NSAIDS also showed increased risk for heart attack and Vioxx and Bextra were eventually withdrawn from the American market (Silverstein, Faich et al. 2000; Juni,

Nartey et al. 2004). Furthermore, because the COX pathway also produces thromboxane which is required for the clotting by platelets, all NSAIDS may cause side effects such as

bleeding and not recommended for treating post-operative pain. In particular, aspirin,

even at low doses increases the risk of major bleeding by approximately 70% compared to the patients taking placebo (McQuaid and Laine 2006). Although NSAIDS are widely used to treat mild to moderate inflammatory pain, the dosage and time span of the treatment need to be considered to balance the benefits versus harm.

Other commonly used analgesics include paracetamol or acetaminophen (Tylenol®)

and steroids. Tylenol is often used to relieve fever and mild pain. It may function through

indirect activation of cannabinoid CB(1) receptors and does not have similar side effects

caused by opioids and NSAIDS (Hogestatt, Jonsson et al. 2005; Ottani, Leone et al.

2006). However, it is not as effective as opioids and NSAIDS and usually cannot control

moderate to severe pain. Steroids such as glucocorticoids can have rapid analgesic effects,

though their functions are nonspecific as they suppress all kinds of immune activities.

Besides, they also cause a lot of side effects including weight gain and osteoporosis.

Although targeted delivery and use of different combinations of analgesics improve

the management of pain, the insufficient relief and considerable side effects are still

significant remaining problems, which make developing new pain relievers necessary.

In this thesis, I performed experiments to determine which populations of adult

sensory neurons still remain plastic and increase CGRP after inflammation. I tested the

hypothesis that activin is a primary factor in the pathology of abnormal pain resulting

from inflammation. I also examined the hypothesis whether activin is sufficient to

regulation CGRP expression and explored the underling molecular mechanisms.

Figure 1.1. Sensory neurons respond to signals in skin for early differentiation,

plasticity after wound and pain functions.

Sensory neuron endings in skin convey information from free endings in the skin past the

neuronal cell bodies in the DRG and into superficial layers of the spinal cord. During

development, and after injury in the adult, sensory nerve endings in skin receive activin

and NGF signals that alter neuropeptides including CGRP. After injury and inflammation

in the adult, both activin and NGF from the injured region can increase the amount of

CGRP. When released into the inflamed area, CGRP functions to increase vasodilation

and release serum proteins in the wound area, and when released into the spinal cord, increases the perception of pain.

Figure 1.2. Activin and selected TGFβ family members, their receptors and signaling.

Activin A binds to Type-II receptors, which recruit and activate the Type-I receptors.

These type I receptors are serine/threonine kinases that activate Smad2/3, which dissociate from the anchor protein SARA to form a heteromeric complex with the common Smad4. This smad complex translocates into the nucleus and regulates target gene expression with other co-factors.

Table 1.1. Family members, Activin A, BMPs and TGFβ signal through some shared receptors and signaling components.

Type II Ligands Type I receptor R-Smad C-Smad I-Smad receptor

Activin A ALK4 (ActRIB) ActRII(A) Smad2, 3 Smad4 Smad7

ActRIIB

ALK2 (ActRI) ActRII(A) BMP-2, 4, 7 ALK3 (BMPRIA) ActRIIB Smad1, 5, 8 Smad4 Smad6, 7

ALK6 (BMPRIB) BMPRII

TGF-β1, β2, β3 ALK5 (TGFβRI) TGFβRII Smad2, 3 Smad4 Smad7

ALK1 (TSR1)

Chapter II

Activin induces tactile allodynia and increases CGRP upon peripheral inflammation

A portion of this Chapter has been published

(Xu, et al., 2005)

ABSTRACT:

Calcitonin gene-related peptide (CGRP) is a sensory neuropeptide important in inflammatory pain that conveys pain information centrally and dilates blood vessels

peripherally. Previous studies indicate activin increases CGRP-immunoreactive (IR)

sensory neurons in vitro, and following wound, activin protein increases in skin and more

neurons have detectable CGRP expression in the innervating dorsal root ganglion (DRG).

These data suggest some adult sensory neurons respond to activin or other target-derived

factors with increased neuropeptide expression.

This study was undertaken to test if activin contributes to inflammatory pain and

increased CGRP, and to learn which neurons retained plasticity. After adjuvant-induced

inflammation, activin mRNA, but not NGF or GDNF, increased in skin. To examine

which DRG neurons increased CGRP immunoreactivity, retrograde tracer-labeled

cutaneous neurons were characterized after inflammation. The proportion and size of

tracer-labeled DRG neurons with detectable CGRP increased after inflammation. One-

third of CGRP-IR neurons that appear after inflammation also had IB4 binding. By

contrast, the increased proportion of CGRP-IR neurons did not appear to come from

RT97-IR neurons. To learn if central projections were altered after inflammation, CGRP

immunoreactivity in the PKCγ-IR lamina IIi was quantified and found to increase.

Injection of activin protein alone caused robust tactile allodynia and increased CGRP in

the DRG. Together, these data support the hypothesis that inflammation and skin changes

involving activin cause some sensory neurons to increase CGRP expression and pain

responses.

INTRODUCTION

The sensory neuropeptide CGRP is essential for pain following inflammation but the factors that regulate its expression are not well understood. Following noxious

stimulation, neuropeptides are released from sensory nerves to transmit pain information

to the spinal cord, and to promote vasodilation of the skin (Wallengren and Hakanson

1987; Holzer 1998). Inflammation increases CGRP in the innervating DRG, sciatic nerve

and inflamed skin, suggesting that sensory neurons increase CGRP and the peptide is

transported to the inflamed area (Smith, Harmar et al. 1992; Nahin and Byers 1994;

Neumann, Doubell et al. 1996; Bulling, Kelly et al. 2001). However, the factors from

inflamed skin that increase CGRP are not clear.

The TGF-β family member, activin, is a strong candidate to mediate changes in CGRP expression. Activin induces CGRP expression in cultured embryonic and adult DRG neurons in a dose dependent manner (Ai, Cappuzzello et al. 1999; Hall, Dinsio et al. 2001;

Cruise, Xu et al. 2004). After skin wounds, activin is upregulated and CGRP increases in the innervating DRG (Hübner, Hu et al. 1996a; Cruise, Xu et al. 2004). Activin may be a common component in inflammatory injury (Hubner, Brauchle et al. 1997; Jones,

Brauman et al. 2000; Phillips, Jones et al. 2001; Jones, Kretser et al. 2004).

The sensory neurons that acquire CGRP and presumably nociceptive functions in the

DRG after injury have been difficult to identify. Distinctive neurons process specialized sensory information. Some 30-40% of DRG neurons are CGRP-IR but neuropeptides are present in 50% of the cutaneous neurons (Bennett, Dmietrieva et al. 1996a). Forty percent of neurons express the NGF receptor TrkA, and most of these neurons also express

CGRP (Verge, Grondin et al. 1992; Mu, Silos-Santiago et al. 1993; Averill, McMahon et

al. 1995). By contrast, other nociceptors derived from late-developing sensory neurons

are GDNF sensitive, express the c-Ret receptor and bind the Griffonia simplicifolia

isolectin B4 (IB4) (Molliver, Wright et al. 1997b; Bradbury, Burnstock et al. 1998).

Roughly 30% of sensory neurons express IB4 (Silverman and Kruger 1990). Although

the reports are variable, it is generally assumed that there is very little colocalization

between TrkA and IB4 (Averill, McMahon et al. 1995; Kashiba, Uchida et al. 2001).

Proprioceptive afferent neurons are recognized by antibody RT97 (Lawson, Harper et al.

1984) and about half the lumbar DRG neurons are RT97-IR (Scott 1992) and 18% TrkA-

IR neurons express RT97 (Averill, McMahon et al. 1995). Physiological characteristics

also distinguish the fibers delivering pain information. All Aδ fibers and half of the C-

fiber nociceptors are NGF dependent and the remaining C fiber nociceptors are GDNF dependent during development (Priestley, Michael et al. 2002). In rat lumbar DRG, 40% of the C-fiber cells, 33% of the Aδ-fiber cells and 17% of the fast A-fiber cells are

CGRP-IR (Scott 1992).

This study was undertaken to test if activin contributes to inflammatory pain, and to learn which DRG neurons retain phenotypic plasticity. Retrograde tracer labeled cutaneous neurons and inflammation were induced. We show that inflammation increases activin mRNA in skin and CGRP-IR neurons in the DRG. The increased population of

CGRP-IR neurons after inflammation is derived from neurons that also bind IB4 -IR, and others that are TrkA-IR. We show for the first time that activin induces tactile allodynia and CGRP expression in the DRG. These data suggest that inflammatory wounds involve

38 activin upregulation in the skin and increased neuronal expression of neuropeptides essential for pain.

MATERIALS AND METHODS:

Animals. Sixty-five adult female Sprague-Dawley rats (8-10 weeks old, 200-250g,

Zivic Miller, Pittsburgh, PA) were used in the experiments. All the rats were housed in cages and maintained on a 12-h light; 12-h dark cycle and the experimental protocols with animals were reviewed and carried out in accordance with the Institutional Animal

Care and Use Committee at Case Western Reserve University.

Inflammation induction and retrograde label of cutaneous neurons innervating ankle area. Fourteen control and fourteen experimental adult rats were anesthetized by intraperitoneal injection (150µl) of a cocktail of 100mg/ml Ketamine (Fort Dodge, Iowa),

20mg/ml Xylazine (Phoenix Scientific, St. Joseph, MO), and 10mg/ml Acepromazine

Maleate (Boehringer Ingelheim Vetmedica, St.Joseph, MO). Hair was shaved from both of rear ankles and 10µl total volume of a saline solution containing 2.5%

Hydroxystilbamidine Methanesulfonate (HM; similar to FluoroGoldTM; Molecular Probe,

Eugene, OR) was injected subcutaneously with a 20μl Hamilton syringe with 27G1/2

needle at 4 sites around the left ankle to label cutaneous sensory neurons innervating this

area. After 4 days, four injections of 80µl total volume of CFA (Sigma, St Louis, MO)

were injected into the same area to induce inflammation. The circumference of the ankle

was measured before tracer injection, before CFA injection and 2 days after CFA

injection. The experiments were done 3 times independently, with 10 rats in each of two experiments (5 control/5 CFA) and 8 rats in the third (4 control/4 CFA).

Activin A, NGF ankle injection: Rats were anesthetized by ether and both ankles were shaved. A dose of 1 µg carrier free human recombinant activin /20 µl (R&D systems,

Minneapolis, MN), or 300ng NGF/20µl (Sigma, St Louis, MO) in sterile saline (Sigma,

St Louis, MO) was injected in the outer lateral side of the left ankle skin. Saline injection and 20µgBSA/20µl saline injection were used as negative control. In particular, 1µg

BSA/20µl saline injection was used as the negative control for 1µg activin injection at 48 hr.

Tactile allodynia. Responses to mechanical stimulation were assessed using calibrated

von Frey filaments. A series of filaments with logarithmically incremental stiffness from

1.4 to 15 g were used to determine the force required to elicit 50% withdrawal response

using the up-down method of Dixon (Chaplan, Bach et al. 1994). Briefly, rats were

placed singly in a wire cage and allowed to acclimate for 15 min. Filaments were applied

perpendicularly onto the lateral skin surface of either left or right ankles with a pressure

that caused the filament to buckle. Testing was initiated with the filament that possesses a

buckling weight of 2 g followed by consecutive stimuli (ascending or descending).

Licking or withdrawal of the paw was recorded as positive responses, and the next

lightest filament was chosen for the next measurement. Absence of response after 5

seconds prompted the use of the next heavier filament. The test was complete when four measurements were made after the initial change in behavior, or after five consecutive

negative or four positive responses had occurred. The resulting sequence of positive and

negative responses was used to interpolate the 50% response thresholds as described by

(Chaplan, Bach et al. 1994). The effects of activin, NGF or saline injections on tactile

allodynia were assessed by an investigator blind to the treatment at 1h, 6h, 12h, 24h and

48h after injection. The effect of 20µg BSA/20µl saline injection was assessed at 1h and

6h. The effect of 1µg BSA/20µl saline injection was assessed at 48h. Control animals

routinely displayed paw withdrawal thresholds between 13-15g. Any animals with

allodynic responses (< than 8 g) in the uninjected right ankle were not used for further

analysis.

Tissue collection. For mRNA isolation, rats were anesthetized by ether and the left

ankle skin was rapidly collected, and fresh frozen at –80°C. For immunohistochemistry,

rats were then perfused intracardially with 4% paraformaldehyde in 0.1M phosphate

buffer and ankle skin, the fourth and fifth lumbar DRGs (L4, L5 DRG), and L4 spinal

cord segments (identified by the level of dorsal root entry) were collected, post-fixed,

frozen, and sections collected on a cryostat at 10µm (DRG) or 20µm (skin, spinal cord)

thickness. The sections were collected on gelatin-subbed slides and kept at –20°C until

use.

Skin RNA isolation and cDNA synthesis. Fresh frozen skin biopsy material was placed

in 1ml ice-cold Trizol reagent (Invitrogen, Carlsbad, CA), homogenized, and extracted in

chloroform. RNA was precipitated in isopropanol at room temperature for 45 min. RNA

was washed with 75% ice-cold ethanol twice and resuspended in 50μl DEPC water. RNA

quantity was determined using 260nm absorbance. Extracted RNA was treated with

DNase to remove genomic DNA contamination using the DNA-freeTM kit (Ambion,

Austin, TX) according to the manufacturer’s instructions, and confirmed by testing in

real-time PCR reactions with all sets of the primers. No valuable threshold cycle (Ct)

value was detected. For the first-strand cDNA synthesis, 2μg RNA was then reverse- transcribed (RT) using the SuperscriptTM III RNaseH- reverse transcriptase and random

primers (Invitrogen, Carlsbad, CA) in a 20 μl reaction volume according to the manufacturer’s instructions. Any contamination of the reagents was tested by omitting

RNA in the RT reactions followed by real-time PCR with GAPDH primers. No valuable

Ct was detected.

Quantitative real time PCR. Two-step SYBR green PCR reaction was performed using an iCycler (Bio-Rad laboratories, Hercules, CA). One microliter RT reaction from

total 20µl reaction was added to 20 μl PCR reaction mixture based on SYBR Green PCR buffer. The final concentration of each reagent was: 3mM MgCl2, 0.2mM each of dATP,

dCTP, dGPT, 0.4mM dUTP, 25U/ml iTaq DNA polymerase, SYBR Green I, ROX

reference dye, 200nM primers. The amplification conditions are: step 1: 95°C, 3min to

activate the iTaq DNA polymerase; step 2: 95°C, 30s for denaturation, 61°C 30s for

annealing and extension, 40 cycles. After amplification, a melting curve protocol was run

to detect primer dimers and to ensure only one product was amplified. Product specificity

of the PCR products was confirmed by 2% agarose gel electrophoresis. All samples were

run in triplicates for each experiment and each sample was amplified in two or three

independent experiments. PCR reaction without cDNA template was used as negative

control. Standard curves were generated for each gene and all the primers were found to

have excellent PCR amplification efficiency (90-100%) as determined by the slope of the

standard curves. Generally, the Ct of each gene from each sample was normalized against

that of GAPDH and fold changes of RNA levels were calculated by 2-ΔΔCt method (Livak

and Schmittgen 2001), in which the relative changes of genes of interest in the experimental group was calculated as the ratio of normalized data over control group.

The following PCR primers were used. Rat Activin βA subunit: forward, 5’-

tgtgaacagtgccaggaga-3’; reverse, 5’- agaaagacggaagtgacgga-3’ (Becker, Hertel et al.

2003) with an expected 102bp fragment. Rat NGF: forward, 5'- catggtacaatctccttcaac-3’;

reverse, 5’-ccaacccacacactgacactg-3’ (modified from (Koh, Armugam et al. 2004) with

an expected 110bp fragment. Rat GDNF: forward, 5’-acgaaaccaaggaggaactga-3’; reverse,

5’-tttgtcgtacattgtctcggc-3’ (Yamagata K, Tagami M et al. 2002) with an expected 74 bp

fragment. Rat GAPDH: forward, 5’-tcaaggctgagaatgggaag-3’; reverse, 5’-

tactcagcaccagcatcacc-3’ (Becker, Hertel et al. 2003) with an expected 103bp fragment.

GAPDH was used as the internal control.

Immunohistochemistry: Sections were washed in PBS and blocked for 1 hour at 25°C

in 0.4% Triton X-100, 3% BSA in PBS (8%BSA for TrkA staining). Sections were then

incubated in primary antibody diluted in dilution buffer (0.4% Triton X-100, 3% BSA,

and 10% donkey serum (Jackson ImmunoResearch, West Grove, PA) in PBS) overnight

at 4°C. For skin staining, skin sections were fixed with acetone at -20°C for 20mins and

air dry for 20mins at room temperature then incubate with primary antibody, goat-anti-

human Activin (1:50) diluted in the dilution buffer and amplified with donkey anti-goat

biotin (1:300, Jackson ImmunoResearch) and streptavidin Cy2 (1:750, Jackson

ImmunoResearch). For CGRP/IB4 and CGRP/RT97 double staining, follow antibodies

were used: rabbit anti-CGRP (1:4000, Sigma), IB4-Alexa 486 (1:100, Molecular Probes)

and mono-RT97 (1:20, Chemicon). After rinses with PBS, sections were incubated in

secondary antibody in 0.4% Triton X-100, 3% BSA, and 10% donkey serum in PBS for

1.5hours. Donkey anti-Rabbit Cy3 was use for rabbit anti-CGRP, donkey anti-mouse Cy2 was used for mono-RT97 (1:100, Jackson ImmunoResearch). For TrkA and CGRP double staining, additional antibodies were used: Guinea pig anti-CGRP (1:200,

Peninsula, San Carlos, CA), followed by donkey anti-guinea pig Cy3 (1:100, Jackson

ImmunoResearch); Rabbit anti-TrkA (1:200, Upstate, Lake Placid, NY), amplified with

goat anti-rabbit biotin (1:300, Chemicon) and streptavidin Cy2 (1:750, Jackson

ImmunoResearch). For PKCγ and CGRP double labeling, rabbit anti-CGRP (1:4000,

Sigma) followed by donkey anti-rabbit Cy3, mono-PKCγ (1:50, BD transduction

laboratories, San Jose, CA) was amplified with donkey anti-mouse biotin (1:300) and streptavidin Cy2 (1:750, Jackson ImmunoResearch). Sections were mounted in ProLong®

Antifade kit according to the manufacturer’s protocol (Molecular Probes).

Neuronal profile counting and area measurement: All the slides were blinded as to

animal treatment by another investigator in the laboratory. Neuronal images were

collected using a Leica DMR microscope and a Hamamatsu, digital CCD camera, C4742-

95-12G04. Leica filter cube A was used to collect tracer-labeled neuronal profile, filter cube +L5 was used to collect Cy2 fluorescence labeled neuronal profile and filter cube

N2.1 was used to collect Cy3 fluorescence labeled neuronal profile. Only neurons that contained a visible nucleus were counted and at least 400 tracer labeled L4 neurons from each rat were counted in the double labeling experiments and at least 400 tracer labeled

L4 and L5 neurons from each rat were counted in 48h activin or BSA injected rats. For neuron size measurement, each neuron with a visible nucleus was drawn with a computer mouse and the soma area was calculated by Openlab 3.1.5. (Improvision, Lexington,

MA). Small neurons < 400 µm2, medium neurons: 400-800 µm2, large neurons > 800

µm2 (Fang, Djouhri et al. 2002). For spinal cord measurement, the spinal cord sections

(38 sections from 3 control rats and 60 sections from 5 CFA rats) were double stained with PKCγ (green) and CGRP (red). Both PKCγ and CGRP images of the same section

were taken unsaturated in either green or red channel. The PKCγ-IR area in the spinal

45 cord was detected depending on PKCγ staining (green) on each image. Both PKCγ-IR area and total fluorescence were collected under green channel. The CGRP fluorescence

(red) in the PKCγ area was collected under red channel with Openlab 3.1.5.

Statistical analyses utilized the unpaired t test for two groups’ comparison and one- way ANOVA followed by Bonferroni/Dunn’s test for multiple groups’ comparison. The

Kolmogorov Smirnov test was used to determine whether neuronal size –frequency distributions differed between populations. ***p<0.0001, **p<0.001, *p<0.05 (Statview

4.1 software, Abacus Concepts, Inc., Berkeley, CA). Data are present as mean±SEM.

RESULTS

CFA injection causes ankle inflammation, tactile allodynia and activin increase

Retrograde tracer label was injected intradermally around the ankle to identify cutaneous neurons innervating this area (Bennett, Dmietrieva et al. 1996a). The neurons

were labeled four days before subsequent experimentation to allow full retrograde

accumulation of the dye in DRG neurons. Injections of CFA in the ankle (Grubb, Stiller

et al. 1993; Hanesch and Schaible 1995) resulted in dramatic increases in ankle diameter

as well as increased nociceptive perception. The circumference of the CFA injected

ankles increased significantly as expected (Fig. 2.1A; p<0.0001). Dye injection alone did

not increase ankle circumference, and the contralateral right ankles did not show swelling.

To test if inflammation caused physiological changes, measurements with von Frey

filaments were performed on the ankles (Grubb, Stiller et al. 1993) and as expected, only

CFA injected ankles were accompanied by pronounced tactile allodynia (Fig.2.1B;

p<0.0001). These data indicate that localized CFA injection caused ipsilateral swelling

and tactile allodynia, but did not induce a systemic inflammation response that affected

the uninjected ankle.

Molecular changes in the skin were observed following inflammation. To begin to understand which changes in skin may affect CGRP expression by sensory afferents after inflammation, the mRNA levels of several growth factors implicated in neuropeptide regulation including activin, NGF and GDNF were assayed by quantitative real-time PCR

in both the inflamed ankle skin and control animals at two days. Both NGF and activin

mRNA levels have been previously shown to increase in skin after excisional wound or

inflammation (Hübner, Hu et al. 1996a; Manni, Lundeberg et al. 2002) and these and

other skin ligands including GDNF can alter neuropeptide levels (Lindsay and Harmar

1989; Ramer, EJ et al. 2003). Activin mRNA levels increased more than 2 fold in inflamed ankle skin compared with control ankle skin, GDNF mRNA levels decreased and no obvious change in NGF mRNA level was found after 2 days inflammation

(Fig.2.1C).

Keratinocytes and immune cells secret proinflammatory cytokines and growth factors after inflammation and actually activin expression was found in the keratinocytes

(Hubner and Werner 1996; Cruise, Xu et al. 2004). We sought to find which cells expressed activin after two days of CFA-induced inflammation. Keratinocytes presented activin immunoreactivity in both control and inflamed skin (Fig.2.1D, E, F) and distinctly, there was a group of infiltrating cells showed up in inflamed skin with obvious activin immunoreactivity (Fig.2.1F, G).

Ankle inflammation increases the proportion of CGRP-IR cutaneous neurons in innervating DRG

Cutaneous sensory neurons in the L4 dermatome in the ankle area were labeled by injection of retrograde tracer that showed bright yellow fluorescence in the innervating

DRG neurons (Fig.2. 2A). Half of the animals received a CFA injection in the left ankle and all ipsilateral L4 DRG were harvested at two days, sectioned and blinded for immunohistochemical analysis. In control animals, 47.8±1.3% of the tracer-labeled

neurons were CGRP-IR, while in CFA-injected animals, 59.5±1.4% of the tracer-labeled

neurons were CGRP-IR (Fig. 2.2B; p<0.0001, 5110 and 4502 tracer labeled neurons were

counted in 10 control and 10 CFA animals). Because the tracer was injected 4 days

before the experiment, the same cutaneous neuronal populations were pre-labeled in each

animal, and the increase was unlikely to be due to altered retrograde transport of dye.

Thus, among tracer-labeled neurons, there was a twelve percent increase in the neurons

with detectable CGRP after inflammation.

To test which subpopulation of adult DRG neurons contributed to the increased CGRP

expression upon inflammation, neuronal size and antigenic expression were assayed.

Generally, neurons delivering nociceptive information have small-medium soma size and

express either CGRP immunoreactivity or IB4 binding (Fig. 2.3A-D), while neurons delivering proprioceptive information have larger soma size and can be identified by bright RT97 antibody reactivity (Fig. 2.3E, F). Overall, the mean size of tracer-labeled sensory neurons increased after inflammation, from 741.9±8.3μm2 to 777.6±8.6μm2 (Fig.

2.4A; p<0.01, 2871 and 2565 L4 DRG neurons were measured in 9 control and 9 CFA

rats). When CGRP-IR, tracer-labeled neurons were measured, the majority of these neurons were small with some medium and large cells and the mean size increased after inflammation, from 648.1±8.7μm2 to 710.6±8.8μm2 (Fig. 2.4B; p<0.0001, 2068 and 2102

neurons were measured in 9 control and 9 CFA rats). By contrast, RT97-IR neurons

showed a broad size spectrum, including small, medium and large cells and the mean size

of RT97-IR tracer labeled sensory neurons did not change after inflammation, remaining

around 1090μm2 (Fig. 2.4C; p>0.9999, 876 and 764 neurons were measured in 5 control

49 and 5 CFA rats). Thus, two days after inflammation, the CGRP-IR, tracer labeled neurons had a larger soma size, while RT97-IR neurons remained unchanged.

To test which population of DRG neurons accounted for the increased CGRP-IR neurons observed with inflammation, double label immunohistochemistry of the tracer- labeled neurons was performed. Both in the control and following inflammation, approximately 34% of tracer labeled neurons in the L4 DRG showed IB4 binding which suggests that IB4 binding was not affected by inflammation and this marker was used as a stable marker. CGRP-IR and IB4 populations in the DRG were generally two distinct populations with only a minor overlap (Fig. 2.3A, B). In the control L4 DRG, 14.9±0.7% tracer labeled neurons expressed both IB4 and CGRP, while in the CFA L4 DRG,

19.1±1.1% tracer labeled neurons expressed both IB4 and CGRP (p<0.05, 2356 and 1952 tracer labeled neurons from 5 control and 5 CFA rats were counted). These data suggest that more IB4 neurons have detectable CGRP expression after inflammation. Indeed, the increase in CGRP-IR/IB4 double labeled neurons accounts for a third of the increase in

CGRP-IR neurons seen after inflammation.

In our experiments, TrkA-IR and CGRP-IR also showed considerable co-expression

(Fig.2.3C, D) and TrkA-IR neurons represented a stable 59.0% of tracer labeled L4 DRG neurons both in control and after inflammation. TrkA is the high affinity neurotrophin receptor for NGF, and previous studies illustrated that about half of the L3 cutaneous

DRG neurons were TrkA-IR (McMahon, Armanini et al. 1994) and 92% of TrkA-IR neurons express CGRP (Averill, McMahon et al. 1995). When the proportion of CGRP-

IR, TrkA-IR tracer labeled neurons was quantified, this group increased from 41.7±3.4% to 48.6±4.9% after inflammation (p<0.05, 2829 and 2522 tracer labeled neurons from 5

50 control and 5 CFA rats were counted). Thus, these data suggest that around 7% more

CGRP/TrkA neurons are detected after inflammation, and more TrkA neurons have detectable CGRP expression after inflammation. By contrast, RT97-IR neurons and

CGRP-IR neurons are two distinct populations with small overlap (Fig, 2.3E, F) and around 46% of tracer labeled neurons both in the control and CFA L4 DRG showed

RT97-IR. While these markers are occasionally present in the same sensory neuron, there was no significant difference found in the CGRP/ RT97 doubly -IR tracer labeled population after inflammation (1914 and 1596 tracer labeled neurons from 5 control and

5 CFA rats were counted). These data suggest that the increase in CGRP-IR neurons after inflammation did not arise from an RT97 -IR pool. In summary, the increase in CGRP-IR neurons observed after inflammation is accounted for by summing the changes in

CGRP/IB4 and CGRP/TrkA neurons.

Central projection of sensory afferents may change after inflammation

To test if the increase in CGRP-IR neurons after inflammation was associated with changes in their central projections in the superficial laminae of the dorsal horn, CGRP immunoreactivity in the spinal cord was assayed. Nociceptive afferent fibers terminate predominantly in lamina I and II of the dorsal horn of the spinal cord (Scott 1992). In the control spinal cord section, CGRP staining occupied laminae I-IIo (outer) as a curved zone (Gibson, Polak et al. 1984) (Fig. 2.5A, B). IB4 staining was seen throughout laminae I and II, but primarily in lamina IIi (inner) (Bennett, Michael et al. 1998) and some overlap between CGRP and IB4 staining could be found (Fig. 2.5B). Since the total proportion of IB4-labeled neurons did not change after inflammation in the DRG, it is

reasonable to believe their central projections would also remain stable. Compared to the

control spinal cord, the CGRP-IR zone in the CFA spinal cord became broader and more

CGRP-IR fibers projected deeply into the IB4-binding zone (Fig. 2.5C). The strong

CGRP fluorescence in the IB4-binding zone and their broad overlap made fluorescence

measurements very difficult to accomplish. For this reason, we used another more

restricted lamina IIi marker, Protein Kinase Cγ. PKCγ staining occupied the inner layer of

laminae II (Mori, Kose et al. 1990; Malmberg, Chen et al. 1997) and only a minor

overlap between CGRP and PKCγ staining could be found in control spinal cord (Fig.

2.5D-F). To measure CGRP-IR fluorescence, sections of L4 spinal cord were doubly

labeled with PKCγ and CGRP. In each case, the slides were processed together and were

blinded as to treatment. The PKCγ area is around 5.0x104µm2 and total fluorescence of

PKCγ is around 2.4x107 units in both control and experimental CFA spinal cord (p>0.05,

38 sections from 3 control rats and 60 sections from 5 CFA rats were included). We then

identified the area of PKCγ reactivity and quantified the second marker, CGRP, in the same domain on the same sections. Compared to control spinal cord, there was a significant increase of CGRP immunofluorescence in the inner lamina II PKCγ domain from 1.46±0.04 x107 to 1.57±0.03x107 after inflammation (p<0.05).

Activin injection results in tactile allodynia and increases the proportion of CGRP--

IR cutaneous neurons in the innervating DRG

Systemic application of NGF (Lewin, Ritter et al. 1993) as well as intraplantar NGF

injections (Woolf, Safieh-Garabedian et al. 1994) have been shown to produce thermal

and mechanical hyperalgesia. Following a similar approach, rats were injected with

300ng NGF or 200ng-1 µg activin in the ankle and assessed for tactile allodynia

responses. In each case, some animals did not respond behaviorally to injection

(“failures”; Fig. 2.6A). At 300ng NGF, approximately 60% of the rats developed

allodynia after 1 hour with a mean response 4.4±3.0g and this response was still present

at 24 hours with a mean response of 7.9g. At low doses (200ng) of activin, 60% of the

animals responded with mean response 9.0g at 6 hour and 80% rats responded with a

mean response 10g at 24 hours (n=4, data not shown). When the dose of activin was

increased to 1µg, animal responses were more reliable, such that 43% rats responded at 1

hour with a 2.7±1.3g mean response. All rats responded 12 hours after 1 µg activin

injection with a 5.2±1.6g mean response. This allodynic response following activin injection lasted at least 48 hours (Fig. 2.6B). By contrast, control injections of 20µl or

1µg BSA and saline did not induce any response (Fig. 2.6A). These data indicate that

activin injection alone is sufficient to cause tactile allodynia.

To test whether injected activin can induce CGRP expression, cutaneous sensory

neurons innervating outer lateral left ankle were labeled by injection of retrograde tracer.

Activin or BSA injection was performed and immunohistochemical analyses performed

at 48 hr. In BSA injected animals, 57.8±1.8% of the tracer-labeled neurons were CGRP-

IR, while in activin-injected animals, 65.9±1.4% of the tracer-labeled neurons were

CGRP-IR (Fig.2.7A, B, C, *p<0.05, 3255 and 7259 tracer labeled neurons in L4 and L5

DRG were counted in 5 BSA and 8 activin injected animals and one BSA injected rat was

not included due to failure to show any tracer labeling).

SUMMARY

Sensory neuropeptides such as CGRP are markers of nociceptive neurons and essential mediators of pain; thus, the regulation of neuropeptides following injury is important for our understanding of pain. CGRP expression is regulated upon injury or inflammation, but the factors that increase CGRP expression are not clear. This study highlights activin as a candidate to mediate changes in CGRP expression and pain responses after injury.

Inflammation increases sensory neuropeptides and pain

Localized inflammation increases CGRP expression in innervating DRG neurons

(Smith, Harmar et al. 1992; Nahin and Byers 1994; Hanesch and Schaible 1995;

Neumann, Doubell et al. 1996; Bulling, Kelly et al. 2001). In this study, a retrogradely transported tracer was used to label cutaneous neurons in the ankle. This approach allowed us to study the same population of DRG neurons with and without peripheral inflammation. The proportion of tracer labeled, CGRP-IR DRG neurons increased after inflammation, suggesting that additional neurons increased CGRP expression. We acknowledge several potential difficulties with this interpretation. These studies utilized antibody detection of CGRP and neurons were subjectively scored as “positive” or

“negative” in slides blinded as to experimental variable. Antibody specificity has been established (Hall, Ai et al. 1997). It remains possible that the present results reflect an increase in CGRP to detectable levels, rather than de novo induction in neurons. Despite

54 this concern, the proportion of cutaneous CGRP-IR neurons in this report is similar to that seen by others (Kuraishi, Nanayama et al. 1989; Donaldson, Harmar et al. 1992).

Increased CGRP -IR neurons and Deeper Central Projections

Size was used as the first criteria to distinguish among different sensory neurons after inflammation. Indeed, the size of tracer labeled neurons increased after inflammation, but these changes occurred only among the RT97-negative neurons. Two possibilities may underlie this observation: RT97-negative, tracer labeled neurons may hypertrophy due to growth factor availability after inflammation (Albers, Wright et al. 1994; Bradbury,

Burnstock et al. 1998) or newly CGRP-IR neurons may derive from larger neurons.

Markers of DRG subtypes were then used to learn which neurons gave rise to the increased CGRP-IR neurons after inflammation. Three populations were tested, IB4+

TrkA- neurons that represent nonpeptidergic nociceptors, TrkA-IR neurons that are mostly peptidergic nociceptors, and RT97-IR neurons that are largely proprioceptors and low-threshold mechanoreceptors. Most newly detectable CGRP-IR neurons after inflammation were derived from TrkA-IR neurons and surprisingly, one third of the newly CGRP-IR neurons also bound IB4, which represents nonpeptidergic nociceptors.

By contrast, there was no increase in CGRP+RT97+ tracer-labeled neurons after inflammation, suggesting that proprioceptors and other large neurons were not responsible for the increase. An earlier report (Neumann, Doubell et al. 1996) suggested that some low-threshold mechanoreceptors which contain Aβ fibers contribute to inflammatory hypersensitivity by expressing SP. However, RT97 is a marker for all A fibers and we did not see any change in CGRP+RT97+ tracer-labeled neurons.

Furthermore, it is not clear if this phenotypic change is permanent or reversible after the inflammatory response has abated.

CGRP immunoreactivity increased in deeper spinal cord laminae after inflammation, suggesting neurons in addition to primary nociceptors had become CGRP-IR. The expression of PKCγ is quite restricted (Mori, Kose et al. 1990; Malmberg, Chen et al.

1997) and serves as a marker for lamina IIi. Neurons in lamina IIi respond preferentially to non-noxious inputs (Light, Trevino et al. 1979; Woolf and Fitzgerald 1983; Malmberg,

Chen et al. 1997) which may be important for neuropathic pain. It is not clear from this assay, however, if new axons express detectable CGRP or whether the increased fluorescence reflects new branches deeper in the spinal cord.

Activin induces tactile allodynia and increases CGRP-IR neurons

In this report, activin or NGF was injected into ankle skin and tactile allodynia was detected over 48 hours. The timing of CGRP and pain responses following inflammation or activin injection suggests particular cellular mechanisms. Allodynia observed 24-48hr after injections is consistent with new CGRP synthesis by neurons (and see this role for

NGF in (Lewin and Mendell 1994)). Our data demonstrating an early onset of allodynia also suggests a peripheral mechanism, in which activin may directly alter the sensitivity of sensory neurons.

These data demonstrate that activin is sufficient to induce behavioral changes and increase CGRP-IR neurons. Thus, activin administration initiated a pathway that eventually regulated CGRP expression in neuronal cell bodies. While activin is sufficient

56 to initiate CGRP expression, the increased CGRP-IR neurons with activin injection were fewer than those observed with inflammation. It is not clear if sufficient activin was injected, or if additional molecules present with inflammation also raise CGRP levels. It will be interesting to learn if activin inhibition during inflammation can abrogate the increase in CGRP-IR neurons, but because inflammation is complex, inhibition of one factor is likely to give only a partial result that may be difficult to resolve.

In summary, these data suggest that inflammation involves activin upregulation in the skin resulting in increased sensory neuron expression of neuropeptides essential for pain.

Further, activin itself confers behavioral correlates of pain in addition to CGRP increases.

Figure 2.1. CFA induced inflammation.

A, Circumference of ipsilateral (dark bar) and contralateral (light bar) ankles two days after subcutaneous injection of tracer or CFA, compared with uninjected (No Inj) ankles.

Only ankles injected with CFA swelled (***p<0.0001, n=14 for each group). B.

Withdrawal threshold two days after CFA injection. CFA injection reduced leg withdrawal threshold (mean±SEM, n=10 for each group, ***p<0.0001). C. mRNA changes in skin with inflammation. Quantitative real-time PCR reveals mean±SEM of fold changes in activin, NGF and GDNF in inflamed skin relative to control ankle skin.

GAPDH was used as the housekeeping gene (n=14 for control and CFA groups). D, E.

Activin immunoreactivity in control skin and inflamed skin (F, G). Higher magnification of the squared area was shown as E and G.

Figure 2.2. Tracer labeled CGRP-IR neurons in the L4 DRG.

A. Labeled sensory neurons in DRG after injection of tracer in ankle skin. B, CGRP-IR neurons in the same DRG section. Some tracer labeled neurons were CGRP-IR (arrow) while others were not (arrowhead).

Figure 2.3. CGRP expression in different subpopulations of L4 DRG neurons.

A, C, E, L4 sensory neurons that innervated the ankle skin contained tracer. B. IB4 binding (green) and CGRP-immunoreactivity (red) were sometimes co-expressed in small-sized neurons. D. TrkA (green) and CGRP (red) immunoreactivities were often co- expressed in neurons. F, RT97 (green) and CGRP (red) immunoreactivities were sometimes co-expressed, especially in medium- to large-sized neurons. Where red and green labels were co-expressed, the cells appear yellow in the merged image. Arrows represent double labeled neurons and arrowheads represent neurons with a single label but not CGRP. Scale bar, 100μm.

Figure 2.4. The effect of inflammation on the size distribution of sensory neurons innervating ankle skin.

A, CFA increased the size of the total FG labeled neurons. (2871 and 2565 L4 DRG neurons were measured in 9 control and 9 CFA rats). B, CFA increased the size of

CGRP-IR tracer labeled neurons (2068 and 2102 neurons were measured in 9 control and

9 CFA rats). C, The size of RT97-IR neurons was not changed after CFA (876 and 764 neurons were measured in 5 control and 5 CFA rats). All the population comparisons were done by Kolmogorov Smirnov test. ***p<0.0001, **p<0.001. Open bar: Control L4

DRG neurons. Filled bar: CFA L4 DRG neurons.

Figure 2.5. Central projections of CGRP-IR fibers after localized inflammation.

A, Schematic figure of lumbar spinal cord dorsal horn lamina. B, L4 spinal cord section

from the control rats was stained to show IB4 (green) and CGRP (red). The zone of

CGRP labeling is in the lamina I and the outer layer of lamina II. The zone of IB4

staining is in the lamina II. C, L4 spinal cord section from CFA rats was stained to show

IB4 and CGRP. The zone of CGRP labeling in the dorsal horn is broader and project deeply into IB4 zone. D. L4 spinal cord section reacted with PKCγ antibody. PKCγ staining was in lamina IIi. E, The same section reacted with CGRP antibodies. CGRP immunoreactivity is usually found in lamina I and the outer layer of lamina II. The white dotted line in D and E indicated the PKCγ-IR area, and was used for CGRP fluorescence measurement in each section. The overlapping of D and E is shown in F. Scale bar:

100um.

Figure 2.6. Activin A or NGF injection induces tactile allodynia.

Rats received a 20µl injection of activin A, NGF, saline or BSA solution under the skin of the left ankle. Leg withdrawal was assessed with von Frey filaments at 1-48h after injection and the number of animals with a response scored in the table (A). The withdrawal threshold among responding animals was averaged, and plotted in the lower graph (B) One microgram activin A induced tactile allodynia at 1 hr that was still present at 48 hr (*) and 300ng NGF induced allodynia and lasted at least 12 hours after injection

(#). Data from 8 activin A, 6 NGF, 6 saline or 6 BSA animals were analyzed by ANOVA followed by Fisher’s PLSD. *, #, p<0.05 denotes the significance level.

Figure 2.7 Activin induces CGRP in the innervating DRG neurons

Activin A injection (1 µg) increased cutaneous CGRP-IR neurons in the DRG. A.Tracer labeled (green) CGRP-IR (red) neurons in control DRG section and activin injected DRG section (B). Some tracer labeled neurons were CGRP-IR (arrow) while others were not

(arrowhead). Scale bar, 100 µm. C. Activin A injection (1 µg) increased cutaneous

CGRP-IR neurons in the DRG (65.9 ± 1.4%) compared to BSA injection (57.8 ± 1.8%);

3255 and 7259 cutaneous tracer-labeled neurons in L4 and L5 DRG were counted in five

BSA and eight activin A-injected animals, respectively. *P<0.05 was defined with unpaired t test.

Chapter III

Activin acutely sensitizes DRG neurons and induces hyperalgesia via sensitizing of

TRPV1

A portion of this work has been submitted for publication. Part of this work was done by

collaborators Dr.Gary Oxford and Dr.Weiguo Zhu and is indicated in the legends.

ABSTRACT

Previous studies have shown that activin increases following peripheral injury and is sufficient to induce acute nociceptive behavior and increase pain peptides in sensory ganglia. This study was designed to test the possibility that the enhanced nociceptive responsiveness associated with activin involved sensitization of TRPV1 in primary sensory neurons. Whole cell patch clamp physiology performed by our collaborators showed that activin acutely sensitized capsaicin responses, and depended on activin receptor kinase activity. Furthermore, activin administration caused acute thermal hyperalgesia in wild type mice, but not in TRPV1 null mice. These data suggest that activin signals through its own receptor and probably PKCε signaling to sensitize the

TRPV1 channel and contributes to acute thermal hyperalgesia.

INTRODUCTION

A well recognized adjunct to tissue damage and repair is the somatosensory tenderness around a wound that triggers nocifensive behaviors (e.g. withdrawal) that serve to guard against additional damage during the healing process. Recent evidence suggests that activin, a member of the transforming growth factor β (TGFβ) superfamily, plays an important role in this response to tissue damage.

Activin has been shown to cause both acute and prolonged tactile allodynia in vivo

(Chapter II). The prolonged nociceptive behavior may be due to increased pain peptides.

Activin has been shown to increase CGRP expression (Ai, Cappuzzello et al. 1999;

Cruise, Xu et al. 2004), which makes it possible that more neuropeptides will be released to facilitate afferent transmission to cause abnormal pain (Urban, Thompson et al. 1994).

As sensitization of nociception by activin occurs within 60 minutes of its administration, the underlying mechanisms of this acute response are unlikely to involve transcriptional regulation of gene expression; rather it is more likely that post- translational events underlie the phenomenon. Increasing evidence has linked sensitization of the capsaicin receptor-ion channel, TRPV1, to a decrease in the threshold of the nociceptor invoked by many inflammatory mediators (Bhave and Gereau 2004).

For example, NGF, prostaglandins, protons, bradykinin, and serotonin are among the factors known to sensitize TRPV1 through both common and distinct signaling pathways

(Cesare and McNaughton 1996; Nicol, Lopshire et al. 1997; Lopshire and Nicol 1998;

Shu and Mendell 2001; Hu, Bhave et al. 2002; Bonnington and McNaughton 2003;

Sugiuar, Bielefeldt et al. 2004; Zhuang, Xu et al. 2004; Ahern, Brooks et al. 2005; Zhang,

Huang et al. 2005; Stein, Ufret-Vincenty et al. 2006; Zhu and Oxford 2007).

We sought to examine the possibility that the enhanced pain responsiveness associated with activin administration or elevation during injury might involve direct sensitization of

TRPV1 in primary sensory neurons. Using electrophysiological and behavioral approaches in collaboration with Dr. Gary Oxford and Dr. Weiguo Zhu, we have confirmed such a link.

MATERIALS AND METHODS

Reagents. Activin A was purchased from R&D systems (Minneapolis, MN) and

reconstituted in sterile 0.1% BSA saline at 10 ng/µl as stock solution. The activin

receptor like kinase 4 (ActRIB) inhibitor SB431542 was purchased from Tocris

(Ellisville, MO) and dissolved in dimethylsulfoxide (DMSO) at 50 mM stock solution.

Adult dorsal root ganglion (DRG) cell culture. Sprague Dawley (SD) rats (Harlan

Sprague Dawley, Indianapolis, IN) were maintained at the laboratory animal resource

center (LARC) of Indiana University School of Medicine. Animals were anesthetized

with isoflurane and then decapitated, which is consistent with the recommendations of

the Panel on Euthanasia of the American Veterinary Medical Association. DRG neurons

from young adult SD rats (150-200 gm) were dissociated and cultured as described

previously (Koplas et al., 1997; Shu and Mendell, 1999). Dissociated cells were plated on

poly-D-lysine-coated glass coverslips, and maintained in Dulbecco’s modified Eagle’s

medium (DMEM) (Gibco, Invitrogen, Grand Island, NY, USA) supplemented with 10%

fetal bovine serum (FBS) (Hyclone, Logan, UT, USA) and 100 units/ml penicillin and

100mg/ml streptomycin for 16-18 hours at 37°C under 5% CO2.

Electrophysiology. Currents were recorded under voltage clamp by whole cell patch

recording methods. In all experiments the holding potential was -60mV. Electrodes

(N51A borosilicate glass) exhibited resistances of 2-5 MΩ. The standard external

solution (SES) contained (in mM; all from Sigma): 145 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2,

10 HEPES, and 10 glucose, pH 7.3. The internal solution consisted of (mM): 130 K- gluconate, 10 EGTA, 1 MgCl2, 1 CaCl2, 10 HEPES, and 2 Mg-ATP, pH 7.4. Data were

collected using an Axopatch 200B patch clamp amplifier, Digidata 1200 interface, and

pClamp 7.0 software (Axon Instruments, Foster City, CA), filtered at 1 kHz, and sampled

at 5 kHz.

Drug treatment. Two capsaicin challenges (50 nM, 40 sec) were delivered to cells from a

quartz capillary (Polymicro Technologies, Phoenix AZ) with a 10 min interval between

challenges. External solutions with 0.1% BSA saline solution (control), 10 ng/ml activin

(experiment) or 10 ng/ml activin plus various pharmacological reagents (interference)

were superfused during the 10 min interval. For experiments assessing pharmacological

interference with the activin effect, a 10 min pretreatment with the relevant inhibiting reagent preceded recording.

PKCε inmmunocytochemical staining. Adult DRG neurons were planted on the PLL and laminin coated glass coverslips in the defined neurobasal medium (Gibco-BRL) with B27 medium supplement (Gibco-BRL), penicillin– streptomycin (1:200, Gibco-BRL), 3 mM glutamine for 4 hours and then treated with activin 20ng/ml for 5mins. Neurons were fixed in 4% paraformaldehyde-PBS for 30mins, washed in PBS, premeabilized in 0.4%

Triton X-100 in PBS for 5mins and blot in 0.4% Triton X-100, 3% BSA, 10% donkey serum (Jackson ImmunoResearch, West Grove, P), 10% goat serum (Gibco-BRL) for

10mins. Neurons were then incubated in Rabbit anti-PKCε (1:100, Santa Cruz), diluted in

0.4% Triton X-100, 3% BSA, 10% goat serum in PBS overnight at 4°C. After rinses with

PBS, cover slips were incubated in goat anti-Rabbit biotin (1:250, Chemicon) in 0.4%

Triton X-100, 3% BSA, 10% goat serum in PBS for 1.5hours and followed with streptavidin Cy2 (1:750, Jackson ImmunoResearch). Neurons were protected with 2% n- propyl gallate in 50% glycerol-PBS for anti-fading.

Behavioral assays of thermal hyperalgesia in wild type or TRPV1 null mice. C57BL/6J and TRPV1 null mice (B6.129X1-Trpv1tm1Jul/J, backcrossed to C57BL/6J for 10 generations, Jackson Laboratories) of 8 weeks age were used for behavioral studies and animal protocols were approved by the Institutional Animal Care and Use Committee at

Case Western Reserve University School of Medicine. Mice were habituated to plastic chambers for an hour per time, twice a day and 3 continuous days. Before each test, the mice were habituated for one hour. In all experiments, the experimenter was blinded to the genotypes of the mice being analyzed. Mice were anesthetized with 3% isoflurane inhalation anesthetic, both legs were shaved and 5 µl saline or 250ng/5μl carrier free activin A (R&D Systems) was injected to the lateral ankle skin with a 26-gauge needle connected to a 25μl Hamilton syringe. Basal nociceptive responses were collected at 1h,

24hrs and 48hrs. Thermal hyperalgesia was detected by a Hargreave test (Hargreaves,

Dubner et al. 1988) (Ugo Basile Plantar™ Analgesia Instrument, Ugo Basile) at 1h, 24hrs and 48hrs. A mobile radiant heat source was located under the glass table and focused onto the plantar area of the hind paw. Paw withdrawal latencies (PWL) were recorded automatically. The intensity of the radiant heat gave a basal latency of 4-5 s on an intact normal mouse. Each trial was repeated 5 times at 4 min intervals and the PWL is the average of the five responses.

Data analysis. A one-way repeated measure ANOVA was used to analyze the majority of data. When p<0.05 a Newman-Keul’s post hoc test was also utilized to make individual comparisons between groups. A student’s t-test was also used for data analysis when only two conditions are being compared.

RESULTS

Activin acutely sensitizes capsaicin currents in cultured DRG neurons

Whole cell currents were recorded from cultured DRG neurons under voltage clamp

conditions (-60 mV). Each of two 50 nM capsaicin administrations separated by a 10

minute exposure to SES containing 0.1% BSA induced inward currents, the second being

consistently smaller than the first due to calcium-dependent desensitization (Docherty,

Yeats et al. 1996; Koplas, Rosenberg et al. 1997; Zhu and Oxford 2007) (Fig. 3.2A). In

contrast, when 10 ng/ml activin was perfused during the 10 minute interval, the second

current response was augmented (Fig. 3.1A). Activin increased the ratio of the second

capsaicin response compared to the first response by 5.17±1.75 fold (n=12), whereas the ratio declined to 0.47±0.07 (n=10) for the vehicle control (Fig. 3.1B, p<0.05). Pre- treatment with ActRIB inhibitor SB431542 (20 µM) for 10 minutes and exposure to both activin and SB431542 during the interval between capsaicin challenges prevented the augmentation (Fig. 3.1A). Under this condition the second-to-first response ratio was not

different from that observed in vehicle controls (0.43±0.09, n=13) (Fig. 3.1B) indicating

that activin A sensitizes capsaicin current through its receptor ActRIB.

Activin causes translocation of PKCε to the DRG neuron plasma membrane

Multiple mechanisms have been proposed to sensitize nociceptive neurons, including

phosphatidylinositol-3-kinase (PI3K), protein kinase C (PKC), calcium-calmodulin-

80 dependent protein kinase II (CaMK II) and others. Several PKC isoforms (βI, βII, δ, ε, ζ) are present in rat DRG neurons (Cesare, Dekker et al. 1999) that might mediate TRPV sensitization. Activation of several of these isoforms requires their translocation from the cytosol to the plasma membrane for interaction with membrane phospholipids. In particular, activation of PKCε translocation is believed to underlie the sensitization of

TRPV1 by bradykinin and NGF (Cesare and McNaughton 1996; Cesare, Dekker et al.

1999; Bonnington and McNaughton 2003). To examine the possible involvement of such translocation in activin sensitization of TRPV1, PKCε translocation was assayed in adult

DRG neurons. Under vehicle control conditions, immunoreactivity of PKCε was evenly distributed in the cytoplasm of adult rat DRG neurons (Fig. 3.2A), whereas activin

(10ng/ml) treatment for 5 minutes caused rapid translocation of PKCε immunoreactivity to the plasma membrane in most DRG neurons (Fig. 3.2 B).

Activin induced thermal hyperalgesia in mice requires the TRPV1 channel

Since our electrophysiological studies reveal a strong sensitization of TRPV1 by activin and TRPV1 has been implicated by many labs as a critical molecule for the expression of acute inflammatory hyperalgesia, we hypothesized that TRPV1 may be required for hyperalgesia associated with elevated levels of activin. To test this hypothesis the effect of activin administration on thermal pain responses was evaluated in

TRPV1 null and wild-type (WT) mice. Activin or saline was injected subcutaneously into one ankle and thermal hyperalgesia assayed over time in operator-blinded studies. WT mice injected with saline demonstrated thermal pain-induced paw withdrawal within

about 5 seconds. Activin administration in WT mice produced more rapid paw

withdrawal as expected since TRPV1 channels were intact in these animals. This

behavioral thermal hyperalgesia was significant when assessed 1 hour following activin

injection, but was absent 24 and 48 hours post-injection (Fig. 3.3). In contrast to WT

mice, activin injection into TRPV1-/- null mice produced withdrawal latencies

comparable to saline injection and these latencies were not different from saline

injections at any time examined. These studies indicate that TRPV1 is required for the induction of thermal hyperalgesia by activin.

SUMMARY

Activin has functions in skin wound healing, such as facilitating the repair process and

modulating scar formation (Munz, Hubner et al. 1999; Munz, Smola et al. 1999; Wankell,

Werner et al. 2003), and enhancing CGRP expression in the innervating DRG neurons

and spinal cord (Chapter II). Activin injection directly into the skin of a rat produces

tactile allodynia (chapter II), but how activin enhances pain sensation in unknown. In the present study we have extended these observation to reveal a fundamental mechanism underlying activin-induced hyperalgesia supported by two novel findings: (1) activin potently sensitizes TRPV1 through its own receptor complex in adult DRG neurons and probably through PKC ε pathway (2) thermal hyperalgesia induced by activin in vivo

requires TRPV1.

Activin sensitizes TRPV1 to mediate acute thermal hyperalgesia

TRPV1 is a multimodal transducer of noxious stimuli including chemicals such as

capsaicin and protons and physical changes such as heat. These responses were initially

taken as evidence for a role in normal nociception (Tominaga, Caterina et al. 1998).

Recently, however, evidence from physiological and behavioral assessments of TRPV1

null mice has indicated that TRPV1 is not required for detecting noxious heat (Woodbury,

Zwick et al. 2004), but plays a significant role in pathological nociception (e.g.

hyperalgesia) associated with tissue injury and inflammation (Caterina, Leffler et al. 2000;

Davis, Gray et al. 2000; Bolcskei, Helyes et al. 2005; Pogatzki-Zahn, Shimizu et al.

2005). This is believed to occur through post-translational changes in the sensitivity and membrane density of TRPV1 as well as transcriptional changes in TRPV1 expression in

response to a variety of molecules released during injury or inflammation including NGF,

bradykinin, and PGE2 (Lopshire and Nicol 1998; Shu and Mendell 2001; Bonnington

and McNaughton 2003; Zhuang, Xu et al. 2004; Zhang, Huang et al. 2005; Stein, Ufret-

Vincenty et al. 2006; Zhu and Oxford 2007). Activin can also sensitize TRPV1 within

minutes of its application consistent with its ability to induce hyperalgesia upon injection

into the skin. Furthermore, hyperalgesia in response to activin injection is absent in

TRPV1 null mice suggesting that sensitization of TRPV1 is both necessary and sufficient

for activin induced acute pain behavior.

TRPV1 knockout mice were used to assay activin mediated hyperalgesia. Most

importantly, the TRPV1-/- sensory ganglia are normal and the expression of IB4 and

substance P, which recognize two major populations of sensory neurons are normal

(Caterina, Leffler et al. 2000). In other reports, this knockout mouse showed normal

thermal sensation at 43°C, the TRPV1 gating temperature, but lost thermal hyperalgesia

after CFA and was widely used to detect TRPV1 dependent nociceptive events (Amadesi,

Nie et al. 2004; Sugiuar, Bielefeldt et al. 2004; Pogatzki-Zahn, Shimizu et al. 2005;

Szabo, Helyes et al. 2005; Jin and Gereau 2006; Negri, Lattanzi et al. 2006; Helyes,

Elekes et al. 2007).

Activin induced thermal hyperalgesia was tested through the Hargreave test, in which

a ramped heat stimulus is delivered, such that a longer latency may reflect a difference in

heat threshold required for response. Mice injected with activin did show thermal

hyperalgesia as early as 1 hour but not at 24, 48 hours. There are several ways to explain why prolonged thermal hyperalgesia was not present. First, in the tactile allodynia experiments, activin was injected into the ankle skin and allodynia was measured by

placing Von Frey filaments in situ. In the thermal hyperalgesia experiments, restricted by the protocol and instruments setting, activin was injected into the ankle skin and thermal

hyperalgesia was measured at the plantar surface of the hindpaw. Any effect seen in the

paw is from diffused activin from the ankle. Although activin is pretty soluble (Jones,

Armes et al. 1996), only a small amount of activin may diffuse into the paw which is

enough to function at the beginning. Activin may degrade overtime and its effect drop

below the detectable level at later time points. Secondly, rats were used for tactile

allodynia and mice were used for thermal hyperalgesia test and there may be a species

difference.

In the TRPV1 null mice, significant higher basal withdrawal latencies were detected

compared to wild-type mice. Through a skin nerve preparation, Woodbury and his

colleagues clearly showed that there was no difference in gating 43°C threshold of the

cutaneous sensory neurons prepared from wild-type and TRPV1 null mice (Woodbury,

Zwick et al. 2004). These null mice also showed normal thermal responses to moderate

temperature (45°C, 48°C) and only showed less sensitive to higher heat (Caterina, Leffler

et al. 2000). These data suggest that TRPV1 channel is not the only thermal sensors responsible for gating 43°C body temperature.

Activin sensitizes TRPV1 through its own receptors and PKC signaling pathway

Activin binds to either the constitutively active Ser/Thr kinase ActRII or IIB to mediate its biological effects. This binding recruits and transphosphorylates ActRIB which enables subsequent phosphorylation of downstream substrates (Massagué 1998; de

Caestecker 2004). Activin receptors are expressed in embryonic sensory neurons (Ai,

Cappuzzello et al. 1999) where they are likely to play a role in phenotypic differentiation.

In adult DRG neurons, activin sensitization of TRPV1 is blocked by a selective ActRIB

receptor antagonist, suggesting that this receptor kinase is required for activin-induced

sensitization of TRPV1 in DRG neurons.

Both canonical and non-canonical signaling pathways are activated by these receptors.

Canonical signaling involves activation of the Smad family of transcription factors by phosphorylation and translocation to the nucleus to activate target gene transcription.

More recently, several non-canonical pathways, including activation of ERK, JNK and p38 MAP kinases, PI3K, PKC and PKA have been reported in different cell types

(Moustakas and Heldin 2005). These signals may be required to mediate and/or augment maximal Smad-dependent responses, or they may act on distinct downstream effectors in different cell types. The precise mechanisms bridging these non-canonical signaling pathways with the activated receptor complexes are not fully understood. Given that both

the activin-induced hyperalgesia and TRPV1 sensitization are rapid responses, these

latter signaling pathways, not involving transcriptional changes are strongly implicated.

There are extensive studies on the signaling pathways linking inflammatory agents to

the acute sensitization of TRPV1. While still controversial, the links between activation

of the NGF receptor TrkA and TRPV1 are the most comprehensively studied and

consensus supports the involvement of both PI3K and src-kinase, and to a lesser extent

p42/p44 Erk phosphorylation in this coupling (Zhuang, Xu et al. 2004; Zhang, Huang et

al. 2005; Stein, Ufret-Vincenty et al. 2006; Zhu and Oxford 2007). However, none of

these signaling is required for activin sensitization TRPV1 (personal communication with

Dr. Gary S. Oxford and Dr.Weiguo Zhu). It is suggested that PKC signaling sensitizes heat responses and potentiates CGRP and substance P release from cultured DRG neurons (Cesare and McNaughton 1996). Also, it is shown that PKC signaling sensitizes nociceptive neurons to both thermal and mechanical stimuli in intact peripheral nerve preparations (Schepelmann, Messlinger et al. 1993; Leng, Mizumura et al. 1996).

Moreover, it has also been reported that the phosphorylated PKC, probably PKCα or

PKCε, sensitizes the TRPV1 (Bhave, Hu et al. 2003). Based on these reports, we focused

on PKCε activation and found that activin did activate PKCε. Further, inhibiting PKCε

activation suppressed activin sensitizing TRPV1 (personal communication with Dr. Gary

S. Oxford and Dr.Weiguo Zhu). These data provide necessary evidence supporting that

activin sensitize TRPV1 through PKCε pathway. However, whether the activations of

other isoforms of PKC, or CaMK II are involved in activin sensitizing TRPV1 needs

further study. Also, it will be interesting to learn the molecular mechanism of how activin activates PKCε in such a short time.

In summary, this study reveals activin mediated sensitization of TRPV1 to be an important component of injury-induced hyperalgesia.

Figure 3.1. Activin A acutely sensitizes the capsaicin induced current in the cultured DRG neurons through the activin type I receptor ActRIB.

A. Representative current traces from 3 DRG neurons challenged by two 50nM capsaicin administrations separated by a 10 minute treatment with either vehicle, 10ng/ml activin A, or 10ng/ml activin plus 20μM SB431542 (additional 10 minutes pretreatment), respectively. B. Summary data (mean±SEM) of normalized capsaicin responses (second peak current/first peak current) indicate that activin through ActRIB can greatly sensitize capsaicin currents in DRG neurons. Open bar is the vehicle control group (0.47 ± 0.07, n=10), black bar is the activin A group (5.17±1.75, n=12, p<0.05 compared to control group), grey bar is the activin A plus SB431542 group (0.43±0.09, n=13, p<0.05 compared to activin A group and p>0.05 compared to control group).

Figure 3.2. Activin A causes translocation of PKCε to cell membrane of rat DRG neurons.

A. PKCε immunoreactivity is uniform distributed in the cytoplasm under control conditions (arrow). B. Activin treatment (20ng/ml) for 5 minutes cause membrane staining of PKCε (arrow ahead).

Figure 3.3. Activin injection induces thermal hyperalgesia in wild-type mice, but

not TRPV1 null mice.

Activin (250ng/5ul) injection induces thermal hyperalgesia in wild-type mice (gray bars, n=19) but not TRPV1 null mice (black bars, n=16). Thermal hyperalgesia was measured by paw withdrawal latency (s) with plantar heating at 1-48h after injection. Data groupings represent assessments done at the indicated times following activin injection.

Wild-type mice show 4-5s withdrawal latency in the absence of activin (white bars), while TRPV1 null mice exhibit longer latencies (hatched bar, n=8). * p<0.05 relative to wild-type responses in activin.

Chapter IV

Activin regulates CGRP expression in both embryonic and adult DRG neurons

A portion of this work has been submitted for publication.

ABSTRACT

NGF and activin can regulate CGRP mRNA and protein in sensory neurons following inflammation and in the embryonic development both NGF and activin are required for

CGRP expression. This study was designed to investigate how neurons integrate these two signals to regulate the neuropeptide important for inflammatory pain. In embryonic rat sensory neuron cultures, NGF supported neuronal survival but did not increase CGRP mRNA, while activin addition induced CGRP mRNA in a concentration-dependent manner. In the adult DRG, NGF is not required for survival, but NGF or activin increases

CGRP-IR neurons in the innervating DRG. While activin alone had no effect on CGRP mRNA induction in adult DRG neurons in vitro, in the presence of NGF, CGRP mRNA was increased by activin. In both embryonic and adult DRG cultures, activin activated a phosphoSmad2 (pSmad2) signaling pathway and NGF activated a phoshpoERK (pERK) signaling pathway and any cross talk is limited. In adult DRG neurons, studies with activin receptor or TrkA receptor kinase inhibitors suggested that each ligand when presented together utilized its cognate receptor. Kinase inhibitor studies demonstrated that activin synergy required several NGF intracellular signals to be activated. Because activin did not further stimulate, but did require NGF intracellular signals, it appears that activin and NGF converge not in cytoplasmic signals, but through transcriptional mechanisms to regulate CGRP in sensory neurons.

INTRODUCTION

CGRP expression can be stimulated by growth factors such as NGF or the cytokine activin but whether or how these factors cooperate in regulating the neuropeptide is not known (Lindsay and Harmar 1989; Amann, Sirinathsinghji et al. 1996; Ai, Cappuzzello et al. 1999; Hall, Dinsio et al. 2001).

Following inflammation, both NGF and activin increase in skin (Woolf, Safieh-

Garabedian et al. 1994; Amann, Peskar et al. 2001; Xu, Van Slambrouck et al. 2005).

Exogenous NGF increases CGRP expression both in adult DRG cultures and normal rats

(Lindsay and Harmar 1989; Amann, Sirinathsinghji et al. 1996; Winston, Toma et al.

2001; Cruise, Xu et al. 2004). Similarly, activin also increases CGRP-IR neurons in DRG cultures and injection of activin into adult rodent skin increases CGRP-IR sensory neurons in the innervating DRG (Cruise, Xu et al. 2004; Xu, Van Slambrouck et al. 2005).

NGF/TrkA signaling is required for CGRP expression in sensory neurons during development, and can increase CGRP expression in adult sensory neurons (Lindsay and

Harmar 1989; Patel, Jackman et al. 2000; Winston, Toma et al. 2001; Cruise, Xu et al.

2004). However, NGF does not induce CGRP expression in embryonic neurons by itself in a serum free culture medium (Ai, Cappuzzello et al. 1999). Application of another target-derived factor, activin, with NGF specifically induces CGRP expression in naïve embryonic rat sensory neurons in a concentration-dependent manner (Ai, Cappuzzello et al. 1999). Both NGF and activin are required for CGRP expression, but how sensory neurons respond to these two signals and regulate CGRP expression is unknown.

Activin and NGF can induce transcriptional changes in sensory neurons, but are

thought to utilize discrete intracellular signals (Durham and Russo 2003; Cruise, Xu et al.

2004). Traditionally, activin is thought to bind the activin receptor complex and activate

Smad2/3 phosphorylation that ultimately forms a heteromeric complex with smad4 that in conjunction with other nuclear binding proteins to regulate the transcription of target genes (Attisano, Wrana et al. 1996; Heldin, Miyazono et al. 1997; Massague and Gomis

2006). By contrast, NGF has been reported to regulate CGRP promoter activity through

ERK-MAP kinase pathways in transfected PC12 (Freeland, Liu et al. 2000) and trigeminal neurons (Durham and Russo 2003). However, it is possible that these factors

converge on intracellular signals as in some other studies (Kretzschmar, Doody et al.

1997; Lutz, Krieglstein et al. 2004; Bao, Tsuchida et al. 2005; Imamichi, Waidmann et al.

2005; Zhang, Deng et al. 2005). Therefore, the aim of this study was to obtain a

molecular understanding about how activin and NGF act together to increase CGRP in

embryonic sensory neurons during development and adult sensory neurons after

inflammation.

MATERIALS AND METHODS

Primary neuron culture. Primary cultures of adult DRG lumbar neurons were prepared from 8-10 week old Sprague Dawley rats (Charles River, Wilmington, MA) (Cruise, Xu et al. 2004). For CGRP mRNA induction assays, cells were plated at 2.1x103cell/cm2 and allowed to attach overnight in defined neurobasal medium (Gibco-BRL, Gaithersburg,

MD; with B27 medium supplement, penicillin– streptomycin 1:200, 3 mM glutamine).

Defined medium was chosen as adult DRG neurons are neurotrophin-independent and do not require NGF for survival (Lindsay 1988), and the absence of serum restricts glial proliferation. Other studies demonstrate robust neuronal survival and differentiation under these conditions (Cruise 2004). Reagents were added the following day (day1) and included human recombinant activin A (Activin, R&D Systems, Minneapolis, MN) and nerve growth factor (NGF, Austral Biologicals, San Ramon, CA). For pharmacological experiments, cultures were pretreated with SB431542 (Sigma, St Louis, MO), K-252a,

U0126, SB203580 or SP600125 (Calbiochem, La Jolla, CA) for one hour, followed by activin, NGF or combination treatment. All drugs were dissolved in dimethyl sulfoxide

(DMSO), such that the final concentration of DMSO ranged from 0.02% (K252a) to

0.08% (U0126) and control wells or those with ligand alone each contained 0.08%

DMSO vehicle. DMSO at these concentrations has no effect on CGRP expression (Data not shown). The culture medium was changed every other day, except for cultures used for pharmacological experiments that were treated daily, and collected on day 5. For

Western blot assay, cells were plated at 5.2x103cell/cm2 in neurobasal medium for four hours, before specific ligands were added for 0.5-60 minutes. Four hours plating was

98 chosen to reduce any cell proliferation, and to identify cell signals that were not modified by extended in vitro cultures.

RNA isolation, cDNA synthesis and Quantitative real time PCR. RNA isolation of DRG cultures were performed with RNeasy Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. RNA quantity was determined, first-strand cDNA synthesis was generated and two-step SYBR green PCR reaction was performed as described in

Chapter II. The following PCR primers were used. Rat alpha-type CGRP: forward, 5-

’aaccttagaaagcagcccaggcatg-3’; reverse, 5’-gtgggcacaaagttgtccttcacca-3’ with an expected 246bp fragment. (a generous gift from Dr.Andy Russo, University of Iowa); Rat

GAPDH, forward, 5’-tcaaggctgagaatgggaag-3’; reverse, 5’-tactcagcaccagcatcacc-3’

(Becker JC, Hertel M et al. 2003) with an expected 103bp fragment. GAPDH was used as the internal control. In general, each condition was run twice, and each experiment performed at least in triplicate.

Western Blotting. Immunoblotting was performed according to protocol described in

(Cruise, Xu et al. 2004). Proteins were denatured in 4x NuPAGE® LDS Sample buffer

(Invitrogen, Carlsbad, CA) containing 0.5% BME at 70°C for 10 minutes and 5 µg total protein was separated on the precast 4% stacking, 10% separating SDS-PAGE gel (Bio-

Rad) and transferred to nitrocellulose membranes. The blots were blotted in 5% skim milk for 1 hr and incubated overnight at 4°C in appropriate primary antibody diluted in

5% BSA-TBST solution: rabbit anti-phospho Smad2 (1:250, Cell Signaling), mouse anti-

Smad2, rabbit anti-phospho ERK1/2, rabbit anti-phosphoP38, rabbit anti-P38, rabbit anti- phosphoJNK, rabbit anti-JNK (1:1000; Cell Signaling), rabbit anti-ERK2 (1:20,000,

Santa Cruz Biotechnology). Secondary antibodies used are: goat anti-rabbit IgG or goat

99 anti-mouse IgG (1:2500, Jackson Immunoresearch). Primary antibody was omitted in control studies, and specific signal was absent. Western immunoblots well below saturation were quantified on a Versadoc scanner in arbitrary units, and data presented from at least duplicate experiments.

Statistical analyses. The relative gene expressions were calculated with Gene Expression

Macro supplied by Bio-Rad laboratories. Generally, the Ct of CGRP was normalized against that of GAPDH in each sample and fold changes of RNA levels was calculated by

2-ΔΔCt method (Livak and Schmittgen 2001), in which the relative changes of genes of interest in the experimental group was calculated as the ratio of normalized data over control group. This analysis was performed for all variables in a study, and at least three independent experiments were compared. The unpaired t test was used for two group comparisons and one-way ANOVA followed by Bonferroni/Dunn’s test was used for multiple group comparisons. Data are presented as mean ± SEM and confidence indicated with asterisks. ***p<0.0001, **p<0.001, *p<0.05 (Statview 4.1 software,

Abacus Concepts, Inc., Berkeley, CA).

100

RESULTS

Inflammation produced by CFA increased activin mRNA in skin, produced local

edema, and increased CGRP in the sensory neurons that project to the inflamed region

(Xu, Van Slambrouck et al. 2005). Our previous studies indicated that activin addition to

embryonic or adult DRG cultures increased CGRP (Ai et al., 1999; Cruise et al., 2004)

and I have shown that direct activin injection increased CGRP-IR neurons in the

innervating DRG (Xu, Van Slambrouck et al. 2005). Similarly, NGF mRNA and protein

increases have been reported in skin following CFA injection (Ma and Woolf 1997;

Amaya, Shimosato et al. 2004; Malin, Molliver et al. 2006), and NGF administration in

vitro or in vivo can increase CGRP (Lindsay and Harmar 1989; Amann, Sirinathsinghji et

al. 1996; Gangula, Zhao et al. 2000; Patel, Jackman et al. 2000; Winston, Toma et al.

2001; Ruiz and Banos 2005). Thus, activin and NGF increase locally following

inflammation, and can increase CGRP in sensory neurons, but how their signals might be

summed by neurons is not known.

Activin acted synergistically with NGF to increase CGRP mRNA in adult DRG

neurons.

To test if activin and NGF ligands acted in parallel or together, each ligand was added

alone and in combination, and CGRP mRNA was assayed. The addition of high levels of

NGF (100ng/ml) did not increase CGRP mRNA at 2 or 6 hours (data not shown) but

increases were detected at 24 hours and CGRP mRNA levels continued to increase each day in this NGF concentration (Fig. 4.1A gray bars). While CGRP mRNA levels appear

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to decline with time without added NGF (Doughty, Atkinson et al. 1991; Shadiack, Sun

et al. 2001) neuronal cell survival is known to remain high in these cultures (Omri and

Meiri 1990; Cruise 2004), and indeed, GAPDH mRNA levels detected by real-time PCR

are similar (data not shown). From these data, we chose to examine CGRP regulation at 4

days in vitro to obtain robust and reproducible CGRP mRNA increases. To learn more

about NGF actions, neurons were then treated with different concentrations of NGF and

CGRP mRNA was quantified (Fig. 4.1B). Maximal CGRP mRNA induction of 8- fold

occurred at 10 ng/ml NGF, and 2.5 ng/ml NGF produced half maximal neuropeptide

induction. By contrast, neurons treated with 1-50ng/ml activin alone had no CGRP

mRNA increase (Fig 4.1A black bars; Fig 4.1C). To learn if activin can modulate NGF-

induced CGRP mRNA synthesis, adult DRG cultures were then treated with activin in the

presence of NGF concentrations that produced half-maximal stimulation of CGRP. When

added together, activin at each concentration tested increased CGRP mRNA to levels

above that induced by NGF or activin alone (Fig 4.1D). These data suggest that the ligands or their downstream signals cooperate in some fashion to increase CGRP.

Activin effects with NGF on CGRP induction required the activin receptor.

Because activin actions occurred only in the presence of NGF, it seemed possible that activin was acting on the NGF receptor itself. To test if activin effects required functional activin or NGF receptors, pharmacological inhibitors were added to specifically block the activation of each receptor. Each inhibitor was tested initially after a single ligand was added. NGF (2.5 ng/ml) induction of CGRP mRNA was largely blocked with the

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addition of K252a (200nM), as expected as this drug is a potent inhibitor of the NGF receptor TrkA kinase (Koizumi, Contreras et al. 1988). Some residual CGRP induction remained, but higher concentrations of K252a (400nM) could not be used as they were toxic (data not shown). By contrast, the addition of SB431542 (10 µM) had no effect on

NGF-induced CGRP mRNA, as it is a competitive ATP binding site inhibitor of ActRIB

(Laping et al., 2002; data not shown). To address which receptors were required to

mediate the synergistic effects of activin in combination with NGF, drugs were added in

the presence of 20 ng/ml activin plus 2.5 ng/ml NGF that produces robust CGRP

expression. The addition of K252a partially blocked activin plus NGF effects, but some

CGRP induction remained after drug action (Fig. 4.2). The activin receptor inhibitor

SB431542 also partially blocked CGRP induction by the ligand combination, but less

effectively. Increased levels of SB431542 (20 µM) did not further decrease activin plus

NGF effects (data not shown) suggesting that the dose administered was maximally

effective. These data suggest that each cognate receptor contributed to the CGRP mRNA

induction when these ligands were presented at the same time.

Activin does not augment NGF mediated intracellular signals or vice versa

The synergistic stimulation of CGRP mRNA by activin and NGF resulted from

intracellular signals initiated by the ligands and receptors. Other reports indicate that

activin receptor binding phosphorylates Smad2/3, while NGF/ TrkA receptor stimulation

regulates CGRP promoter activity through ERK-MAP kinase pathways. To test which

intracellular signals were stimulated by each ligand alone or in combination, adult DRG

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were acutely isolated for four hours before stimulation with ligand, and phospho-specific

antibody reagents were used to assay intracellular signals over minutes. In these neuronal

cell culture, most of the adult DRG neuron showed translocation of Smad2 from the

cytoplasm to the nucleus (Fig. 4.3 A, B) or phospho-Smad2 (pSmad2) nucleus staining after one hour of activin treatment (Fig. 4.3 C, D). Nonneuronal cells in these cultures did

not show significant Smad2 immunoreactivity, which suggests that neurons are the major

responding cells to activin in these cultures. To learn which concentration of activin

maximally stimulated intracellular signals, activated phospho-Smad2 (pSmad2) was

assayed at one hour. Smad2 activation increased with added activin, and maximal

stimulation was reached at 6ng/ml activin (Fig. 4.3 E). This pSmad2 reagent detects

Smad2 only when dually phosphorylated at serine 465 and serine 467, and may detect

phosphorylated Smad3 at its equivalent site. The present studies revealed one band on the

western blot consistent with Smad2 phosphorylation. To learn more about the temporal

stimulation of Smad2 by activin, 5ng/ml activin was added to neurons, and

phosphorylated Smad2 was evaluated at subsequent times. Activin resulted in maximal

stimulation of pSmad2 at one hour, and sustained levels of pSmad2 were still present at 2

hour (Fig. 4.3 G). By contrast, similar activin concentration and temporal stimulation

studies showed no effect on phospho-ERK (pERK) stimulation (Fig. 4.3 F, H). These

observations indicate that activin stimulates pSmad2 at nanomolar concentrations and

that stimulation by activin results in a long-lived pSmad2 signal.

The activity of major protein kinases after addition of a single ligand or the

combination of NGF and activin was assayed. As expected, activin (20ng/ml) addition for

5 or 15 min stimulated pSmad2 while NGF (50ng/ml) did not (Fig. 4.4 A). By contrast,

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NGF addition for 5 or 15 min stimulated pERK2 activation but activin had no effect on

this kinase (Fig. 4.4B). In addition, NGF addition resulted in modest activation of p38 but

not JNK (Fig. 4.4C, D) in adult sensory neurons, but activin had no effect on these

kinases. These data support the activation of canonical intracellular pathways by each

ligand, when added alone.

To test if activin increased NGF-mediated intracellular signals, neurons were treated

with both 20ng/ml activin plus 50ng/ml NGF for 15mins. The two ligands added together

stimulated pSmad2 to levels achieved by activin alone, and pERK2 to levels stimulated

by NGF alone (Fig. 4.4 A, B). Further, phospho-CREB was also not activated by activin

(data not shown), which is one transcriptional factor activated by NGF to activate the

CGRP promoter (Freeland, Liu et al. 2000). Thus, the synergistic stimulation of CGRP

mRNA seen when activin and NGF were added together was not mediated by additional

stimulation of these kinase pathways. In addition, several studies were designed to test if presentation of one ligand first altered responses to the second. For example, neurons were treated with one ligand for 5mins or 4hours and followed with 15min stimulation by the second ligand. Such “priming” by activin or NGF did not demonstrate any effect on the later treatment (data not shown). Thus, in combination, NGF and activin appear to increase the classical signaling pathways associated with these ligands in adult DRG neurons, and no obvious cross talk between these two signaling pathways was observed.

In a second approach to test if NGF-initiated signals were necessary for the effects of activin addition, pharmacological inhibitors were used to learn which signaling pathways were required for CGRP induction. Initial tests were designed to determine which signaling pathways were required for NGF alone to induce CGRP mRNA. The potent and

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specific inhibitor of MEK1 and MEK2 that activate the ERK1/2 pathway, U0126 (Favata,

Horiuchi et al. 1998; Portis and Longnecker 2004) decreased NGF mediated CGRP mRNA induction by less than half (Fig. 5.5 A). Higher concentrations of U0126 (10, 20 and 40 µM) did not further reduce NGF mediated induction, and 60 µM U0126 caused cell death (data not shown). In other studies, tumor necrosis factor-alpha (TNFα) is

shown to regulates CGRP expression in trigeminal neurons through JNK and p38

pathways (Bowen, Schmidt et al. 2006). To test the role of these kinases, SB203580

(20µM), a highly specific inhibitor of p38 MAP kinase (Lee, Laydon et al. 1994) was

used, but this reagent only slightly decreased NGF induced CGRP mRNA (Fig. 4.5 A).

Similarly, the addition of SP600125 (10µM), a potent inhibitor of JNK (Bennett, Sasaki

et al. 2001; Clerk, Kemp et al. 2002), again only slightly decreased NGF induced CGRP

mRNA (Fig. 4.5 A). These data suggest that each pathway may contribute to NGF

mediated signals that increase CGRP. Cells were then treated with different combinations

of inhibitors. While the ERK pathway appeared to play the largest role in CGRP

induction, the addition of inhibitors to p38 and JNK showed reductions that suggested

these pathways also contributed to mRNA induction. The maximum inhibition of CGRP

induction was obtained only when both ERK and p38 or JNK pathways were blocked or

all three pathways were blocked together (Fig. 4.5 A). These data suggest that ERK is the

predominant signaling pathway used by NGF to regulate CGRP mRNA but that all three

MAPK pathways can combine to regulate CGRP expression by NGF.

The signaling pathways used by activin in combination with NGF to stimulate CGRP

expression are similar to those used by NGF alone. Again, inhibition of the MAPK

pathway by U0126 partially decreased CGRP induction by the ligand combination, but

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the most robust inhibition was obtained only by inhibiting ERK plus p38 or JNK pathways (Fig. 4.5 B). In combination, the intracellular signal activation data indicate that activin addition does not further stimulate NGF intracellular kinases, and the pharmacological inhibitor data demonstrates that NGF-stimulated kinase functions are required for both ligands to act together. These observations suggest that the induction of

CGRP mRNA by activin in combination with NGF include cooperative transcriptional mechanisms in the nucleus.

Activin increases CGRP mRNA in embryonic DRG culture

In previous studies, activin increased the number of CGRP-IR neurons in serum free embryonic DRG culture that contained NGF (Ai, Cappuzzello et al. 1999). In this case, approximately 4% of neurons had detectable CGRP-IR in control wells and 60% neurons express CGRP after activin was added. To determine if activin similarly induced CGRP mRNA, embryonic DRG cultures were treated with 1ng/ml activin for 1-4 days and RNA was evaluated by quantitative real time PCR. In “control” cultures containing 25 ng/ml

NGF to support neuronal survival, there was no increase in CGRP mRNA after 4 days

(Fig. 4.6), nor was CGRP increased by 125 ng/ml NGF (data not shown). Because embryonic neurons depend upon NGF for survival, it was not possible to test activin effects on CGRP mRNA induction alone. By contrast, activin addition (1ng/ml) in the presence of NGF (25 ng/ml), increased CGRP mRNA at 24 hours and maximum induction was achieved at day 3 (Fig. 4.6). These data demonstrate that NGF alone does

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not alter CGRP mRNA in embryonic DRG neurons, but activin in the presence of NGF

increases CGRP mRNA.

Activin and NGF signal independently

To test if activin and NGF effects might converge via common intracellular signaling,

the signals initiated by these factors were tested in embryonic DRG cultures. In order to

get clear ligand responses, embryonic DRG cells were dissociated and plated in serum free NB medium without NGF for 4 hours and followed with different ligand treatments.

Brief activin stimulation resulted in robust pSmad2 activation in embryonic culture, as

expected. In embryonic DRG cultures, activin increased pSmad2 at 15 minutes treatment

and with maximal effect at 6ng/ml activin (Fig. 4.7A). Maximal pSmad2 activation was

observed at one hour, and sustained levels of pSmad2 were still present at 4 hours (Fig.

4.8B). These data suggest that either receptor activation or pSmad2 remains for hours in

these embryonic cultures. No pERK activation was detected after activin treatment. By

contrast, 25ng/ml NGF transiently increased pERK level at 5mins and the response

almost disappeared at 4 hours. No significant pSmad2 activation was detected after NGF

treatment (Fig. 4.7C). Embryonic neurons treated with 0.5ng/ml activin plus 25ng/ml

NGF for 15mins showed similar pSmad2 activation compared to neurons treated with

activin alone and similar pERK activation level compared to neurons treated with NGF

alone (Fig. 4.7C,D), which suggests there is no cross talk between the signals for these

ligands. Studies in which neurons were briefly treated with NGF or activin for 5mins or 4

hours to occupy receptors, followed by 15mins activin/NGF treatment did not show any

108 effect on the later treatment (data not shown), which suggest occupying one receptor did not affect signaling activated by the other ligand.

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SUMMARY

Both NGF and activin are required for CGRP expression in sensory neurons during

development and both ligands increase after skin inflammation and can induce CGRP,

but how activin and NGF might work together or separately to regulate the neuropeptide was not known. The present study identifies for the first time that activin in combination with NGF synergistically stimulates CGRP induction, that each cognate receptor is stimulated and parallel intracellular signals are initiated, but that activin synergistic action requires the function of an intact NGF signaling cascade. These data rule out various cytoplasmic cross-talk mechanisms and strongly suggest that activin stimulatory actions with NGF include cooperative mechanisms in the nucleus.

We and others have demonstrated that NGF increases CGRP mRNA and peptide in adult DRG cultures (Lindsay and Harmar 1989; Winston, Toma et al. 2001). In previous studies, subcutaneous activin increased CGRP-IR neurons in the innervating DRG neurons at 2 days (Xu, Van Slambrouck et al. 2005), but the present data suggest that activin alone cannot regulate CGRP mRNA in isolated neurons. In combination, this suggests either that NGF or factors with similar permissive activity are available in vivo, or possibly that activin also regulates CGRP at the translational level. It likely that existing NGF levels are sufficient to allow activin effects to be apparent, as both NGF mRNA and protein are present in adult smooth muscle, hair follicle sheath cells, keratinocytes, and hypodermal fibroblasts (Hasan, Zhang et al. 2000).

This study confirms that NGF regulates CGRP expression in adult DRG cultures, and begins to untangle the signaling cascades that underlie its effects in primary neurons. In

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previous reports, NGF activated a 1250bp rat CGRP promoter that contains proximal

cylicAMP- and Ras-responsive regions and a distal enhancer (Durham and Russo 2003).

In promoter-reporter studies with transfected trigeminal neurons or PC12 cells, NGF

mediated activation was totally blocked by PD98059 or U0126, which suggests that NGF

signals through ERK to drive the CGRP promoter (Freeland, Liu et al. 2000; Durham and

Russo 2003). By contrast, in the present report, U0126 did not totally block NGF effects

on CGRP induction in adult dorsal root ganglion neurons. Instead, U0126 blocked perhaps half the NGF effect, and further inhibition was achieved only by blocking either p38 or JNK in addition to ERK. These data suggest that ERK is the predominant but not the only pathway used by NGF to regulate CGRP expression in adult sensory neurons, and when the ERK pathway is inhibited, NGF signals can be mediated through p38 or

JNK.

There are at least three differences between the previous CGRP promoter/reporter assays and the present study. An obvious difference is that the promoter assay was performed within 2 hours but our experiments lasted 4 days. Thus, it was possible that

U0126 degraded in our culture over time. To address this potential concern, pERK activation was tested in fresh DRG cells that were treated with medium from 4 day

U0126 treated cultures, and very low pERK levels were detected, indicating the drug remained active (data not shown). A second difference is that the promoter assay tested the activation of a 1250bp rat CGRP promoter but the present study utilized the entire

CGRP mRNA. It is possible that the 1250bp rat CGRP promoter contains elements responding to ERK but not additional elements required to respond to p38 or JNK signaling pathways. However, TNF-alpha has been shown to activate the 1250bp rat

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CGRP promoter through p38 and JNK signaling pathways in transfected trigeminal

neurons, which suggest this mini CGRP promoter does contain elements needed to

respond to p38 or JNK signaling pathways (Bowen, Schmidt et al. 2006). It is also possible that our western blots were not sensitive enough to show that activation. The last difference is the promoter assay was done in trigeminal cultures and we used DRG cell cultures, and a cell type difference may account for the signaling observed.

Occupying one receptor with its corresponding ligand and treating with the other ligand did not cause any facilitation or inhibition on the second signal pathway, which largely ruled out the possible receptor level interaction between NGF and activin.

However, the effects of activin in combination with NGF on CGRP mRNA was abolished by the specific pharmacological blocker SB431542 that inhibits activin receptor like kinases (Laping, Grygielko et al. 2002), suggesting that activin requires its own receptor complex to effect CGRP expression. Activin receptors phosphorylate

Smad2/3 (Attisano, Wrana et al. 1996; Heldin, Miyazono et al. 1997) that interacts with common Smad4 and translocates to the nucleus to alter transcription. In addition to this classical pathway, TGF-beta family members have been shown to activate alternative pathways in different cell systems including ERK, JNK and p38 MAP kinases

(Kretzschmar, Doody et al. 1997; Bao, Tsuchida et al. 2005; Imamichi, Waidmann et al.

2005; Zhang, Deng et al. 2005). Further, phospho-CREB was also not stimulated by activin in adult sensory neurons (Chapter V), which is one transcriptional factor activated by NGF to drive CGRP promoter (Freeland, Liu et al. 2000). No NGF signaling pathway tested was activated by activin and one target transcriptional factor for NGF activating

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CGRP promoter was also not phosphorylated by activin, suggesting that activin is

unlikely to affect CGRP through augmenting NGF intracellular signals.

NGF supports embryonic sensory neuron survival (Crowley, Spencer et al. 1994;

Smeyne, Klein et al. 1994; Silos-Santiago, Greenlund et al. 1995), but its role in

regulating neuropeptide expression is not clear. Previous data showed that cultured E14.5

DRG in 25ng/ml NGF only contained 4-7% CGRP-IR neurons after 7 days culture (Ai,

Cappuzzello et al. 1999) and in the present report, 25-125ng/ml NGF treatment did not

increase CGRP mRNA in these cultures over several days. By contrast, activin induces

concentration dependent CGRP-IR neuron induction without affecting neuron survival,

which suggests activin but not NGF alone can modulate CGRP expression in embryonic development. Because embryonic neurons required trophic support from NGF, these

studies were performed in the presence of 25 ng/ml NGF and thus it is hard to tell

whether NGF plays a role in CGRP expression other than survival.

In an attempt to support neuronal survival in the absence of NGF, Patel and his

colleagues used Bax and NGF double deficient transgenic animals. In these studies, few

DRG neurons contained CGRP in vivo, and many cutaneous fibers had unusual projections that did not extend into the skin, suggesting NGF is required for skin innervation and skin factors might be important for CGRP induction. However, this study

also examined explanted E13.5 DRG from Bax-/- NGF-/- mice, and found that 41% of

cultured neurons were CGRP-IR after 3.5 days in 50ng/ml NGF and heat-inactivated

horse serum medium but had only a few CGRP-IR neurons without NGF. These studies

were interpreted to indicate that NGF /TrkA signaling is required for the full phenotypic

differentiation of sensory neurons (Patel, Jackman et al. 2000). It is important to

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recognize that serum contains activin (Krummen, Woodruff et al. 1993; Torii, Hanai et al.

1993; Woodruff, Sluss et al. 1997; Ai, MacPhedran et al. 1998; Baccarelli, Morpurgo et

al. 2001; Eldar-Geva, Spitz et al. 2001; Morpurgo, Beck-Peccoz et al. 2002), and thus,

both factors may have been present in this study. Over all, these data suggest that NGF is

not only a survival factor but also plays a permissive function for CGRP expression, and

activin is the key inducer for CGRP expression in development.

In summary, embryonic sensory neurons turn on CGRP expression in the presence of both NGF and activin and this two molecules signal through their canonical signaling pathways to regulate CGRP expression. In the adult, activin in combination with NGF stimulates CGRP mRNA expression and this effect requires an intact NGF receptor and intracellular signaling, but independently stimulates the activin receptor and Smad signals.

We interpret these results to indicate that this activin synergy with NGF is unlikely to occur at the receptor or cytoplasm, but is more likely to modulate promoter function. It will be important and interesting to identify any changes in the transcriptional complex that regulates CGRP after inflammation.

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Figure 4.1. Activin acts with NGF to induce CGRP mRNA in adult DRG cultures.

Adult sensory neurons were maintained in serum free medium without NGF overnight followed by A. NGF (100ng/ml, labeled N) or Activin (50ng/ml, labeled A) treatment for

1-4 days. After NGF treatment, CGRP mRNA increased each day in culture (one-way

ANOVA *p<0.05, ***p<0.0001, values compared to D1 NGF). B. Increased concentrations of NGF for 4 days stimulated CGRP induction with maximal stimulation at 10 ng/ml (all values significantly increased above 1 ng/ml NGF, p<0.0001). C.

Different concentrations of Activin did not increase CGRP mRNA after 4 days. D. In the presence of half-maximal concentration of NGF at 2.5 ng/ml, activin (5-50 ng/ml) synergistically increased CGRP mRNA compared to NGF alone ***p<0.0001, *p<0.05.

In each case, quantitative real-time PCR Ct values for CGRP were normalized to Ct values of the housekeeping gene GAPDH as in (Livak and Schmittgen 2001), to reflect mean±SEM of fold changes of CGRP mRNA. Three independent experiments were compared.

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Figure 4.2. Activin effects on CGRP mRNA occur through its own receptor.

Primary adult DRG cultures were plated in serum-free medium overnight and pretreated

with drugs K252a (200nM, 0.02% DMSO) or SB431542 (10µM; 0.04% DMSO) for 1

hour, followed by activin (20ng/ml) or NGF (2.5ng/ml) or both for 4 days. DMSO

control wells, activin or NGF or both treated wells all contained 0.08% DMSO. Inhibition

of trkA kinase with K252a reduced NGF stimulation (***p<0.0001). When activin and

NGF were added at the same time, synergistic stimulation of CGRP occurred (p<0001,

also as shown in Fig 1D) and this stimulation was reduced by inhibition of either trkA

kinase with K252a or activin receptor kinase with SB431542 (### p<0.0001, values

compared to 2.5ng/ml NGF plus 20ng/ml activin). In each case, quantitative real-time

PCR Ct values for CGRP were normalized to Ct values of the housekeeping gene

GAPDH as in (Livak and Schmittgen 2001), to reflect mean±SEM of fold changes of

CGRP mRNA. Three-eight independent experiments were compared.

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Figure 4.3. Activin signaling pathways in the adult sensory neurons.

A. Intracellular Smad2 phosphorylation and translocation in neurons following activin stimulation. Dissociated adult DRGs were cultured for 3 days in basal medium, and then stimulated for 1 h with 20 ng/ml activin. Cultures were stained with antibody to Smad2

(A and B) or phosphospecific Smad2 (C and D). Following one hour of activin (20ng/ml)

stimulation, most neurons showed Smad2 translocation into the nucleus (arrowheads, A

and B), and phospho-specific Smad2-immunoreactivity was localized in neuronal nuclei

(C and D, arrowheads). Phospho-Smad2 was present in 20.7 ± 5% neurons in control

cultures and 91 ± 5% neurons after activin stimulation (200 cells counted in each of six

experiments). E. For immunoblotting assay, adult sensory neurons were plated in serum-

free neurobasal medium for 4 hours and followed with different concentrations of activin

for an hour. Representative blots of pSmad2 and a reprobe for Smad2 in the same

samples. F. Activin does not activate pERK. Representative blots of pERK2 and ERK2

reprobe. (n=2) G. Activin (5 ng/ml) stimulates long-lasting pSmad2 activation,

representative blots of pSmad2 and Smad2. H. Activin (5ng/ml) does not activate pERK,

representative blots of pERK2 and ERK2.

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Figure 4.4. Activin stimulates pSmad2 while NGF stimulates pERK2 in adult sensory neurons.

Activin (20ng/ml) or NGF (50ng/ml) or both were added to adult DRG neurons for 5 or

15 minutes to evaluate stimulation of intracellular signaling pathways A. Activin activates pSmad2 while B. NGF increases pERK2 but not vice versa, and activin and

NGF in combination did not further increase phosphorylation of either pathway (n>3).

Representative blots of pSmad2 and a reprobe for Smad2 are shown. Likewise, representative blot for pERK2 and a reprobe for ERK2 in the same samples are shown. C.

In similar studies, the p38 pathway was slightly activated by NGF but not activin. D. The

JNK pathway was not activated after NGF or activin treatment. (n=2) Representative blots of p-p38 and a reprobe for p38, and a second blot for pJNK and a reprobe for JNK in the same samples are shown.

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Figure 4.5. NGF and activin signal through multiple MAPK pathways to regulate

CGRP expression.

A. NGF alone or B. Activin plus NGF. Primary adult DRG cultures were plated in serum- free medium overnight and pretreated with drugs U0126 (40µM) to inhibit ERK,

SB203580 (20µM) to inhibit p38, or SP600125 (10µM) to inhibit JNK for 1 hour, followed by activin (20ng/ml) or NGF (2.5ng/ml) or both for 4 days (vehicle contained

0.08% DMSO, see Methods). Data represent quantitative real-time PCR, corrected for

GAPDH expression and reflects the mean±SEM of fold changes of CGRP mRNA in 3 experiments. ***p<0.0001, **p<0.001, *p<0.05 was defined with one-way ANOVA followed by Bonferroni/Dunn’s test with comparison to 2.5ng/ml NGF and ### p<0.0001,

#p<0.05 was obtained from comparing to 2.5ng/ml NGF plus 20ng/ml Activin. (n>=3 for each condition).

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Figure 4.6. Activin increases CGRP mRNA in E14.5 DRG neuronal culture in a concentration dependent manner

Relative CGRP mRNA changes in E14.5 DRG cultures in control conditions containing

25 ng/ml NGF (C), or after activin (1ng/ml) (A) treatment for 1-4 days (D1-D4).

Quantitative real-time PCR reveals mean±SEM of fold changes of CGRP mRNA relative to control neurons (N) only treated with NGF. GAPDH was used as the housekeeping gene. (n=3 for each condition). * p<0.05 is obtained by compared to activin treated for one day with one way ANOVA followed by Bonferroni/Dunn’s test.

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Figure 4.7. Activin and NGF signaling pathways in embryonic sensory neurons.

A. Activin dose response in embryonic sensory neuron culture for 15mins. Embryonic sensory neurons were plated in serum-free neurobasal medium without NGF for 4 hours, followed by addition of different concentrations of activin treatment for 15mins.

Representative blots of pSmad2 and Smad2 for activin dose response experiments. (n=2)

B. Activin (5ng/ml) time course in embryonic sensory neuron culture. Embryonic sensory neurons were plated in serum-free neurobasal medium without NGF for 4 hours and followed with 5ng/ml activin treatments for different times. Western blot revealed the ratio of the phospho-Smad2 level over Smad2 level at each time points compared to the control. C, D. Activin and NGF signal independently. Embryonic sensory neurons were plated in serum-free neurobasal medium without NGF for 4 hours and followed with different ligand treatments. (C) Activin (0.5ng/ml) activates pSmad2 pathway and (D)

NGF (25ng/ml) activates pERK pathway but not vice versa, and activin and NGF in combination did not further stimulate signals by either pathway. (n>3)

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(Activin)

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Chapter VI

General Discussion

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Inflammatory pain is caused by mediators including growth factors, cytokines, plasma proteases and other chemicals which change the excitability of the nociceptive neurons.

An incomplete understanding of these inflammatory mediators leads to insufficient therapeutic strategies to manage inflammatory pain. In this discussion, the functions of activin in nociceptive response as an inflammatory cytokine are discussed.

The studies in this thesis provide evidences that some adult DRG neurons retain plasticity to respond to target-derived factors including activin and activin increases

CGRP expression after inflammation. The studies also provide both in vitro and in vivo data to reveal activin functions in hyperalgesia. Acutely, activin sensitized the TRPV1 channel to decrease the threshold of nociceptors and produce hypersensitivity. In the long term, activin acted with NGF to increase CGRP. These data suggest that increased activin in the inflamed skin contributes to inflammatory pain and inhibiting activin may be a valuable therapeutic strategy to relieve inflammatory pain.

I have focused my thesis on how sensory neurons respond to inflammation using a

CFA induced skin inflammation model and in vitro models including a serum free, growth factor free rodent DRG neuronal culture system. Specifically, I focused on two questions. First, which population of DRG neurons still remain plastic to turn on neuropeptides including CGRP after inflammation in adult animals; second, how activin induces both acute and prolonged hyperalgesia in animals.

I chose to use a skin inflammation paradigm in order to understand the function of activin in inflammation. Skin contains limited amount of activin and skin damage causes a dramatic mRNA and protein elevations in activin expression (Hubner, Hu et al. 1996;

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Cruise, Xu et al. 2004). Transgenic overexpression of activin or of the activin binding protein, follistatin, in mice keratinocytes results in accelerated or retarded wound healing accompanied by increased or decreased scar formation, respectively, suggesting activin is involved in injury (Munz, Smola et al. 1999; Wankell, Munz et al. 2001). These observations suggest that activin released during tissue damage can promote the repair process. Increases in activin are also found following systemic inflammation and arthropathies and may represent a common component in inflammatory injury (Hubner,

Brauchle et al. 1997; Munz, Hübner et al. 1999; Jones, Brauman et al. 2000; Phillips,

Jones et al. 2001; Jones, Kretser et al. 2004; Werner and Alzheimer 2006).

Adult rat sensory neuronal culture was used to understand the molecular signals induced by activin. This is a widely used in vitro system for studying sensory neurons.

The adult sensory neuronal culture is often used to study gene regulation and underlying signaling pathways (Lindsay and Harmar 1989; Vedder, Affolter et al. 1993; Ghassemi,

Dib-Hajj et al. 2001; Ramer, Bradbury et al. 2001; Winston, Toma et al. 2001; Skoff,

Resta et al. 2003). Defined culture conditions without serum were used for the study of

CGRP regulation both in the embryonic and adult sensory neuronal cultures in this thesis.

Serum contains multiple growth factors of which activin is a prominent component and has been reported to induce CGRP expression (Krummen, Woodruff et al. 1993; Torii,

Hanai et al. 1993; Hall, Ai et al. 1997; Woodruff, Sluss et al. 1997; Ai, MacPhedran et al.

1998; Ai, Cappuzzello et al. 1999; Baccarelli, Morpurgo et al. 2001; Eldar-Geva, Spitz et al. 2001; Morpurgo, Beck-Peccoz et al. 2002). In serum free cultures, adult and embryonic sensory neurons survival well and glial cells proliferate slowly (Ai,

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MacPhedran et al. 1998; Ai, Cappuzzello et al. 1999). These serum free cultures provide

a clean system to study activin function on CGRP expression.

In the following sections, activin functions are discussed in the context of

inflammation.

Sensory neuron phenotypes are determined during development but some remain

plasticity in the adult

Limited availability of activin may regulate embryonic CGRP induction. During

development, most sensory neurons express TrkA and activin receptors including ActRI,

ActRII and ActRIIB (Ai, Cappuzzello et al. 1999; Huang and Reichardt 2001). With

NGF, fewer than 5% neurons are CGRP positive in embryonic DRG cultures, and low concentrations of activin (0.5 ng/ml ) results in 60% of DRG neurons to turn on CGRP

(Ai, Cappuzzello et al. 1999). On the other hand, in the adult, about 30% of DRG neurons are CGRP-IR (Scott 1992) and inflammation increases another 12% CGRP-IR neurons.

These data suggest there is a larger group of neurons capable of expressing CGRP in the

embryo than the adult. Since CGRP expression is turned on after E18-E19 and gradually

increases and maintains in around 30% of total adult DRG neurons, it is possible that

limited amount of target-derived factors determine the percentage of sensory neurons that turn on CGRP. Indeed, NGF mRNA peaks at E12-13 in mouse skin and gradually declines to levels seen in the adult by E18 (Ernfors, Hallbook et al. 1988; Buchman,

Sporn et al. 1994). ActivinβA mRNA is first detected in the dermal layer of the rat skin at

E13 and disappears after E17 (Roberts, Sawchenko et al. 1991). Although it’s not known

134 how long the proteins are sustained after mRNA decreases, both NGF and activin have narrow expression windows in development which may limit their availability for competent sensory neurons to express CGRP.

Activin is the key inducer of CGRP in embryonic neurons. Studies from the Bax, NGF double null mice suggest that NGF is required for CGRP expression in the presence of serum (Patel, Jackman et al. 2000). However, different concentrations of NGF alone failed to induce CGRP expression in a serum free embryonic sensory neuron culture

(Chapter V). On the other hand, activin from skin induces CGRP expression in naive embryonic sensory neurons cultured with NGF in a concentration dependent manner (Ai,

Cappuzzello et al. 1999). These studies suggest that NGF provides a permissive condition for neurons to respond to activin and activin is the key inducer of CGRP.

The sensory neuron phenotype is also regulated by intrinsic mechanisms. Runx1 is responsive for the transition of half of the early NGF responding neurons to downregulate

TrkA and upregulate GDNF receptor, c-ret, postnatally (Silos-Santiago, Molliver et al.

1995; Molliver and Snider 1997; Molliver, Wright et al. 1997). Runx1 null mice have more TrkA-IR neurons and CGRP-IR neurons in the DRG. Also, these mice lose nociceptive ion channels commonly seen in nonpeptidergic nociceptors. These data suggest that Runx1 inhibits CGRP expression and promotes nociceptive ion channels expression in nonpeptidergic nociceptors. However, during development, some CGRP-IR neurons express a low level of Runx1. These data suggest that either this low level of

Runx1 is not sufficient for inhibiting CGRP expression or other factors such as activin inhibiting Runx1 function. The second hypothesis points to that activin may de-repress

CGRP expression through stopping Runx1 function. This hypothesis can be tested by in

135 vitro culturing E14.5 Runx1 null sensory neurons with NGF. If the hypothesis is true, they should turn on CGRP expression since the inhibition from Runx1 is not present.

Further addition of exogenous activin to these cultures should not affect CGRP expression. Moreover, overexpressing Runx1 in the sensory neurons should block activin function on CGRP induction.

Adult sensory neurons appear to respond to target-derived factors that are re-expressed after inflammation. Mature skin, unlike that in the embryo only has low levels of activin, however, skin wound and inflammation rapidly increase activin mRNA and protein

(Chapter II and Hubner, Hu et al. 1996a; Cruise, Xu et al. 2004). The cellular source of activin following inflammation is not yet clear and probably involves both infiltrating immune cells and keratinocytes (Chapter II). Similarly, after wound or inflammation,

NGF is upregulated in skin. The cellular source of NGF in this case may include mast cells, keratinocytes, fibroblasts and immune cells in the wounded area (Di Marco,

Marchisio et al. 1991; Markenson 1996; Pincelli and Marconi 2000). Because both NGF and activin are upregulated after inflammation, these re-occurring target-derived factors are likely to drive the induction of CGRP (Woolf, Safieh-Garabedian et al. 1994; Amann,

Peskar et al. 2001),.

Target-derived factors can regulate neuropeptides expression. Systemic administration of anti-NGF antibody during experimental inflammation can partially reduce CGRP increases (Woolf, Safieh-Garabedian et al. 1994). Moreover, direct injection of NGF or activin in vivo increased CGRP expression in the innervating sensory neurons (Chapter II and Woolf, Safieh-Garabedian et al. 1994). These studies suggest that some sensory neurons can respond to target-derived factors from inflamed skin to increase

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neuropeptide expression and these neurons still maintain phenotypic plasticity even in

adulthood. Indeed, after skin inflammation, de novo CGRP-IR neurons were found in the

innervating DRG and 60% of these newly CGRP-IR neurons came from TrkA-IR population. In each case, it is not clear whether the change of neuropeptide expression present in adult sensory neurons after injury reflects a permanent conversion or just a transient modulation.

One third of the de novo CGRP-IR neurons in adult DRG after CFA induced inflammation came from the IB4 binding, nonpeptidergic nociceptive population.

Although both peptidergic (express TrkA and respond to NGF) and nonpeptidergic (bind

IB4 and respond to GDNF) nociceptors transduce nociceptive signals, they have been suggested to contribute to inflammatory pain or neuropathic pain respectively (Snider and

McMahon 1998). Biochemically, these two groups of nociceptors respond to different trophic factors in the adult (Boucher, Okuse et al. 2000) and selectively express different peptides, ion channels and receptors (Scott 1992; Bradbury, Burnstock et al. 1998; Dong,

Han et al. 2001; Potrebic, Ahn et al. 2003; Amaya, Shimosato et al. 2004). Anatomically, their central projections are also different. Peptidergic fibers mainly project to lamina I and the outer layer of lamina II of the dorsal horn of the spinal cord while nonpeptidergic nociceptive fibers project to the inner layer of lamina II (Malmberg, Chen et al. 1997).

These biochemical and anatomical features support the idea that these two types of nociceptors may have distinct functions (Braz, Nassar et al. 2005). Indeed, GDNF family of neurotrophins has a therapeutic effect on neuropathic pain (Malmberg, Chen et al.

1997; Boucher, Okuse et al. 2000; Gardell, Wang et al. 2003; Wang, Guo et al. 2003;

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McMahon and Jones 2004). Runx1 null mice which do not have nonpeptidergic

nociceptors also do not develop neuropathic pain (Chen, Broom et al. 2006).

Nonpeptidergic nociceptors also express Ret and possibly Runx1 in the adult. Runx1

is believed to inhibit CGRP expression in IB4 neurons postnatally and it is not known

whether the inhibition mechanism is still valid in the adult. It would be interesting to know whether the de novo expression of CGRP in IB4 neurons after inflammation requires the down-regulation of Runx1 or inhibiting Runx1 signaling. One can speculate that increased target-derived factors such as activin may function to inhibit Runx1. These data suggest that although two populations of nociceptors may function in different pathological conditions, under CFA induced inflammation some IB4 nonpeptidergic nociceptors can respond to target-derived factors to express CGRP.

There are several hypotheses to explain how allodynia develops after inflammation.

Allodynia is believed to have a peripheral origin, in which chemicals such as bradykinin,

prostaglandins, serotonin, histamine, NGF or as I suggested activin released near the

peripheral nerve terminals sensitize or activate ion channels expressed on the free nerve

endings of nociceptors to decrease the firing threshold. In this way, a benign stimulus

becomes harsh enough to initiate action potentials in the nociceptors to deliver

nociceptive information.

It is also proposed that there is a central component for allodynia that mainly involves

projection neuron hypersensitivity (“wind up”), low mechanical Aβ fiber phenotypic

“switch” and circuits rewinding in the spinal cord. The phenomenon “wind up” happens

under noxious constant stimulation and is a frequency-dependent increase in the

138 excitability of spinal cord neurons. It can be evoked by repeated C fiber firing, probably due to the constant buildup of glutamate, CGRP and substance P which facilitate the firing of projection neurons (Herrero, Laird et al. 2000). The “wind up” is similar to long term potentiation in learning and memory. CGRP can facilitate projection neurons firing

(Bird, Han et al. 2006) and indeed there is more CGRP-IR in the PKCγ-IR layer (IIi) after inflammation. These data suggest that the pain hypersensitivity developed after inflammation may be due to “wind up” (Chapter II).

It is also suggested that low threshold mechanoreceptors can “switch” their phenotype after inflammation by expressing substance P to resemble pain fibers. As a consequence, even light touch can release substance P to sensitize dorsal horn neurons in the spinal cord and cause increased central responses to innocuous stimuli (Neumann, Doubell et al.

1996; Xu and Zhao 2001). However, Allen et al. found that the release of substance P increased after activating Aδ and C fibers but failed to detect substance P release from Aβ fiber after inflammation (Allen, Li et al. 1999). In my studies, the number of RT-97 (label all A fibers (Lawson, Harper et al. 1984)) and CGRP-IR double positive sensory neurons remained unchanged after inflammation. These data confirm that CGRP expression is not changed in Aβ neurons after inflammation.

Furthermore, new synaptogenesis is proposed to rewire local circuits in the spinal cord and cause allodynia. For example, it is suggested that Aβ fibers may re-organize their synaptic connections by nerve sprouting in the spinal cord so that information of touch transmitted by Aβ to be processed as nociceptive information (Lewin and Moshourab

2004; Hoseini, Hoseini et al. 2006). In the spinal cord, there was more CGRP-IR in the

PKCγ-IR layer (IIi) after two days inflammaiton, however it can reflect either more axons

139 expressing detectable CGRP or more branchings of existing CGRP expressing axons into deeper layers in the dorsal horn. However, this data supports new synaptogesis as an explanation as newly branched out CGRP-IR fibers are very likely to make new synapse to nociceptive projection neurons after inflammation. To further test this possibility, one can make a transgenic mouse which uses CGRP promoter to drive the expression of transneuronal tracer, wheat germ agglutinin (WGA). In this way, all CGRP neurons and their postsynaptic projection neurons will be labeled by WGA. One can account the number of WGA labeled and CGRP-IR projection neurons in the animals with or without inflammation. If there are more WGA labeled and CGRP-IR projection neurons after inflammation, new synaptogenesis occurs.

Overall, these proposed hypotheses are not mutually exclusive and all point to the hyperexcitability of nociceptors or projection neurons after inflammation (Herrero, Laird et al. 2000). The peripheral mechanism involves quick sensitization which accounts for acute inflammatory pain and most of the central mechanisms require hours or days to increase peptide expression, upregulate receptors, sprout nerves or rewire circuits. Each of these may explain why chronic inflammatory pain develops gradually over a rather prolonged period of time.

Activin causes thermal hyperalgesia requires TRPV1

Pain hypersensitivity is a main sign of tissue damage, but how molecules from peripheral tissues affect sensory neuron physiology is not fully understood. Activin has been suggested to have functions in several tissue repair and neuronal protective

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processes. However, the present report is the first time that activin has been shown to cause thermal hyperalgesia. Inflammatory mediators released during inflammation sensitize nociceptors to cause hypersensitivity (Julius and Basbaum 2001). Nociceptor

sensitization after inflammation may involve TRPV1 (Caterina, Leffler et al. 2000; Davis,

Gray et al. 2000). Activation of TRPV1 facilitates the firing of nociceptors and increase

the release of neuropeptides (Petho, Izydorczyk et al. 2004). In fact, TRPV1 antagonists

are currently under clinical trial for treating acute and chronic pain (Rami and Gunthorpe

2004; Appendino, De Petrocellis et al. 2005).

TRPV1 can be regulated at both transcriptional and post-translational levels. Indeed, it

has been shown that inflammation mediators modulate TRPV1 to sensitize the channel to

cause thermal hyperalgesia. For example, NGF increases TRPV1 activity through

increasing its expression and membrane insertion and inhibiting its desensitization in

multiple pathways including p38, src-kinase, PI3K and ERK pathways (Ji, Samad et al.

2002; Zhuang, Xu et al. 2004; Zhang, Huang et al. 2005; Stein, Ufret-Vincenty et al.

2006; Zhang and McNaughton 2006; Zhu and Oxford 2007). Bradykinin, ATP and

inflammatory proteases have been reported to function through their corresponding

receptors to sensitize TRPV1 through the PKC pathway (Tominaga, Wada et al. 2001;

Sugiura, Tominaga et al. 2002; Amadesi, Nie et al. 2004). Serotonin has been suggested

to signal through PKA to sensitize TRPV1 (Sugiuar, Bielefeldt et al. 2004). Although

activin challenges to DRG neurons failed to increase either phosphorylation of ERK or

Akt as indicators of stimulation of ERK and PI3K pathways respectively, activin does

cause PKCε translocation, indicating its activation in the adult DRG neurons.

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Others mechanisms of hyperalgesia exist in addition to TRPV1 (Caterina, Leffler et

al. 2000). For example, other thermosensitive TRP family members are also expressed in

sensory neurons. TRPV2 is activated by heat >52°C; TRPV3 is activated by heat > 32-

39°C; TRPV4 is activated by heat >27-35°C; TRPM8 is activated by temperature below

25-28°C and TRPA1 is activated by temperature below 17°C (Numazaki and Tominaga

2004). They function in normal innocuous and noxious thermal sensation or thermal

hyperalgesia or both. Among these thermal sensitive channels, gene ablation studies have shown that both TRPV4 and TRPA1 are generally not required for gating body temperature but instead are involved in inflammation or nerve injury induced hyperalgesia (Todaka, Taniguchi et al. 2004; Obata, Katsura et al. 2005; Bautista, Jordt et al. 2006). It will be interesting to see whether activin can sensitize these thermal sensors and modulate thermal sensation in different modality to cause hyperalgesia, such as cold.

Skin keratinocytes have been proposed to participate in thermal sensation. It was

thought that only DRG neurons sense temperature (Hensel 1981). Recently, TRPV3 and

TRPV4 are all found expressed in keratinocytes. Activation of these thermosensors on

skin keratinocytes are suggested to affect temperature sensitivity and produce inflammatory pain (Guler, Lee et al. 2002; Inoue, Koizumi et al. 2002; Peier, Reeve et al.

2002; Southall, Li et al. 2003). For example, TRPV3 is only expressed in keratinocytes and TRPV3 null mice have deficits in both innocuous and noxious thermal stimuli

(Moqrich, Hwang et al. 2005). The mechanism about how keratinocytes participate in thermal sensation is largely unknown. It may involve nonsynaptical communication between keratinocytes and nerve fibers. Activin immunoreactivity is found in the keratinocytes in both normal skin and inflamed skin (Seishima, Nojiri et al. 1999; Werner,

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Beer et al. 2001). It is reasonable to speculate that activation of thermosensors on the

keratinocytes will release activin. Released activin can function as a paracrine factor to

affect TRPs on the free nerve ending and as an autocrine regulator to sensitize TRPs

channels on keratinocytes.

Activin causes acute and prolonged mechanical tactile allodynia

Activin induced acute mechanical tactile allodynia has a different mechanism than acute thermal hyperalgesia. Activin induces mechanical tactile allodynia as quickly as one hour after administration and the symptoms last at least 48 hours (Chapter II). The acute phase of tactile allodynia probably involves a post-translational mechanism, but

TRPV1 is probably not required for mechanical allodynia. In TRPV1 null mice, normal mechanical response is similar compared to wild-type mice. These animals do not develop thermal hyperalgesia; however, still develop mechanical hyperalgesia after inflammation (Caterina, Leffler et al. 2000; Jin and Gereau 2006). These data strongly support the idea that TRPV1 is not required for mechanical allodynia. Nevertheless, there is some evidence suggesting that TRPV1 may contribute to mechanical allodynia in certain experimental models and animal species. For example, in an experimental arthritis model, TRPV1 null mice showed some decreased mechanical hypersensitivity compared to wild-type mice (Barton, McQueen et al. 2006). In guinea pig but not rats and mice, capsazepine, a TRPV1 antagonist, partially blocked CFA and neuropathic injury-induced mechanical allodynia (Walker, Urban et al. 2003). After all, these data suggest that there are other mechanisms for mechanical hyperalgesia.

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It seems likely that other TPR channels, sodium channels account for activin effects on tactile allodynia. TRPV4 plays a crucial role in mechanical allodynia through cAMP/PKA, PKCε or PKD pathways (Alessandri-Haber, Dina et al. 2006; Grant,

Cottrell et al. 2007). Since activin directly cause tactile allodynia and induce PKCε translocation, it is reasonable to speculate that activin might function through TRPV4 to induce mechanical allodynia (Chapter III and personal communication with Drs. Zhu and

Oxford).

Tetrodotoxin (TTX)-resistant sodium channels are well documented as critical players in modulating mechanical hypersensitivity under inflammatory and neuropathic pain conditions (Wood, Abrahamsen et al. 2004; Wood, Boorman et al. 2004). The voltage gated sodium channels, Nav1.8 and Nav1.9 are predominantly expressed in nociceptive neurons and both channels are coexpressed with TRPV1 (Fjell, Cummins et al. 1999).

TTX-resistant sodium currents are suggested to contribute to the generation of action potential, repetitive firing, and setting membrane potential in nociceptive neurons, which make it a candidate modulator of peripheral sensitization (Bhave and Gereau 2004).

Indeed, Nav1.8 null mice show a pronounced analgesia to noxious mechanical stimuli

(Akopian, Souslova et al. 1999). Temporally decreasing Nav1.8 leads to a decrease in prostaglandin-induced mechanical hyperalgesia (Khasar, Gold et al. 1998). Nav1.9 null mice show normal basal thermal and mechanical pain sensitivity but decreased mechanical hypersensitivity after administration of bradykinin, prostaglandin or capsaicin

(Amaya, Wang et al. 2006). Furthermore, all inflammatory mediators known to sensitize nociceptors also potentiate TTX-resistant sodium currents, such as PGE2, adenosine, serotonin and TNF α (England, Bevan et al. 1996; Gold, Reichling et al. 1996; Cardenas,

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Del Mar et al. 1997; Gold, Levine et al. 1998; Fitzgerald, Okuse et al. 1999; Gold 1999;

Jin and Gereau 2006). In addition to the TTX-resistant sodium currents, both voltage gated potassium and calcium channels can be regulated at post-translational level to contribute to peripheral sensitization (Bhave and Gereau 2004). It will be interesting to

know whether activin can also modulate these channels to provoke mechanical allodynia.

By contrast, the prolonged tactile allodynia may involve gene regulation. In my hands,

activin-induced mechanical tactile allodynia is present at 48 hours after treatment. The

mechanism underlying long term tactile allodynia caused by activin may include new

protein synthesis, such as CGRP. I have shown that activin can induce CGRP expression

both in vivo and in vitro. Although I did not test whether activin challenge could cause

CGRP release from the nerve terminals, it seems likely since TRPV1 activation usually

leads to vesicle release (Amadesi, Nie et al. 2004). My working model is that when

activin is applied to the nerve terminal, it sensitizes TRPV1 and perhaps sodium channels

to decrease the threshold of the nociceptors to fire action potentials, which leads to acute

hypersensitivity. Meanwhile skin-derived activin, retrogradely transported back to the

neuronal cell bodies in the DRG, acts with NGF to increases CGRP content which

contributes to long term tactile allodynia.

Activin regulates CGRP expression

CGRP protein can be regulated by NGF and activin in both embryonic and adult DRG

cultures (Lindsay and Harmar 1989; Ai, Cappuzzello et al. 1999; Patel, Jackman et al.

2000; Cruise, Xu et al. 2004). In this study, I focused on CGRP regulation at the mRNA

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level and sought the possible signaling pathways to explain how both NGF and activin

regulate CGRP.

Activin appears to potentiate NGF effects on CGRP expression through CGRP promoter regulation. NGF has been suggested to signal through ERK to activate the

CGRP promoter in transfected PC12 cells and trigeminal neurons (Freeland, Liu et al.

2000; Durham and Russo 2003). The canonical signaling for activin is through phosphorylated Smad2/3 that interacts with common Smad4 and then translocates to the nucleus to alter gene transcription (Attisano, Wrana et al. 1996; Heldin, Miyazono et al.

1997). Activin did activate phosphorylated Smad2 in the adult sensory neurons and no major NGF intracellular signaling was activated by activin (Chapter V), suggesting it is unlikely that activin promotes NGF effect on CGRP through augmenting NGF intracellular signals. These data suggest that the interaction may occur at the promoter level. Others have shown that NGF activation of the CGRP promoter requires a cAMP response element located at bases -103 to -109 and a cell-specific enhancer that contains a basic helix-loop-helix site and an adjacent octamer-binding motif (Peleg, Abruzzese et al. 1990; Klein, Lamballe et al. 1992; Tverberg and Russo 1993; Watson, Ensor et al.

1995; Lanigan and Russo 1997). Whether activin modifies NGF recruited transcriptional factors or recruits its own transcriptional factors to potentiate NGF effects on CGRP is unclear. It is interesting to know whether activin signals modify NGF recruited transcriptional factors or it recruits its own transcriptional factors to potentiate NGF effect on CGRP.

Transcriptional factors reported to drive CGRP promoter includes a heterodimer of the bHLH-Zip protein USF-1 and -2 and a winged-helix transcription factor Foxa2 (Lanigan

146 and Russo 1997; Viney, Schmidt et al. 2004). USF-1 and 2 are ubiquitously expressed proteins whereas Foxa2 expression is cell-specific and found in several neuronal-like cell lines such as CA77 thyroid C cell line, P19 cells as well as in rat trigeminal ganglia where CGRP is expressed (Tverberg and Russo 1993; Jacob, Budhiraja et al. 1997; Viney,

Schmidt et al. 2004). One should look at the expression of Foxa2 in the DRG neurons.

Should it be expressed, it would be interesting to know whether activin signaling affects the activity of these transcription factors, through which it potentiates NGF effect on

CGRP expression. Several experiments can be used to test this hypothesis. First, activin signaling may positively regulate the expression of both factors; therefore, the levels of these factors may change upon activin treatment. Second, as phosphorylation is a must for the transcriptional activities of both factors, it is reasonable to test whether activin treatment would affect the phosphorylation status of both factors. Third, evidence suggests that other co-factors may interact with both USFs and Foxa2 to stabilize the enhancersome (Viney, Schmidt et al., 2004), therefore activin treatment might alter the transcription complex which can be revealed by either immunoprecipitation or EMSA assays.

Among those factors that can regulate CGRP expression, many function through distinct kinase pathways. In the present study, activin signals through Smads, however, no substantially augmented kinase pathways were observed in embryonic or adult sensory neurons. Although PKCε translocation was detected, it is a transient signal that returns to control level one hour after activin treatment (Personal communication with

Dr.Weiguo Zhu and Dr. Gary Oxford). This abbreviated activity makes it unlikely that activin uses PKCε pathway to modify gene expression. These data suggest that activin

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activated Smads probably function as transcription factors to regulate CGRP expression.

Using the rVista search engine on the rat 2kb CGRP promoter only, no Smad4 binding

sites are found except that one potential Smad3 site. However, the predicted Smad3 site may not be relevant in my neuronal cultures as I showed Smad2 was the dominant activated Smads. Although there are more Smad4 binding sites can be found in further

upstream regions in human, rat and mice CGRP gene, their functional relevance remains

to be tested.

Alternatively, activin may regulate the CGRP promoter through recruiting

transcription factors other than Smads. The dimer of Smad2 and Smad4 binds to DNA

sequence weakly and the single binding of the dimer is usually not sufficient to drive

gene expression (Zawel, Dai et al. 1998; Massague and Wotton 2000). Indeed, other

DNA binding partners are required to form TGFβ/activin response factor (TRF) which

binds to activin response element (ARE) to drive target gene expression. Two forkhead

domain proteins have been found to bind ARE, one is FAST-1 which regulates

homeobox gene Mix.2 in early Xenopus tissue and the other is the mammalian homolog

FAST-2 which regulates the goosecoid (gsc) promoter (Huang, Murtaugh et al. 1995;

Labbe, Silvestri et al. 1998). Thus, it will be interesting to see whether activin can

activate the CGRP promoter in DRG neurons and which DNA binding partner is required.

I did not find ARE for FAST-2 in the 2kb rat promoter, but it is still possible that it exists

in even further upstream region. One approach might be to transfect sensory neurons with

FAST-2 and test CGRP promoter reporter activity after activin treatment. If FAST-2 does

play a role in CGRP mRNA regulation by activin, CGRP promoter activity will increase.

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Overall, activin functions through its own receptor to modify NGF effects on CGRP

and the regulation probably happens at the promoter level. Actually, quite similar activin synergy with NGF was also seen on substance P (Katsmen.E and Hall AK, unpublished data). This observation raises the interesting question about whether this is a common pattern used for activin to potentiate NGF effects on gene regulation during pain sensation. NGF is known to regulate many genes involved in nociception and abnormal pain, such as brain-derived neurotrophic factor (BDNF), purinergic receptor of ATP

(P2X3), ASIC3, µ-opiate receptor, bradykinin B2 receptor, TRPV1,voltage gated ion channels including sodium channels, potassium channels and calcium channels (Hilborn,

Vaillancourt et al. 1998; Fjell, Cummins et al. 1999; Mamet, Baron et al. 2002; Priestley,

Michael et al. 2002; Park, Choi et al. 2003; Amaya, Shimosato et al. 2004; Pezet and

McMahon 2006). NGF mRNA increases after four days CFA induced inflammation, peaks at seven days by two folds (Malin, Molliver et al. 2006) and then decreases, which suggests that NGF increase is slow and modest. Such a response is unlikely to be the critical component in chronic inflammatory pain. If activin does potentiate NGF effect on these genes after inflammation, it may explain why NGF dependent gene regulation is seen. This hypothesis can be tested by injecting both CFA and anti-activin antibody and checking the expression of NGF regulated genes such as TRPV1. If the hypothesis is right, TRPV1 expression will be decreased compared to CFA injected only animals. If this is true, activin may be the fundamental component in chronic inflammatory pain and just by inhibiting activin we can relieve pain.

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Interaction between nervous system and immune system during inflammation

Inflammatory pain is not just a sensation due to the activity of nervous system but is also strongly modulated by the immune system (Gold, Archelos et al. 1999; Machelska and Stein 2000; Watkins and Maier 2000; DeLeo and Yezierski 2001; Elenkov, Iezzoni et al. 2005; Marchand, Perretti et al. 2005; Moalem and Tracey 2006). In CFA induced skin inflammation, the immune system is activated first to recruit phagocytes to clean the bacteria. During this process, proinflammatory cytokines, growth factors and other chemicals are released. Some inflammatory mediators increase vascular permeability while others activate nociceptors to transduce nociceptive information, and further

release neuropeptides such as CGRP and substance P. These peptides also induce

vasodilation and extravasation and results in neurogenic inflammation (Levine, Clark et

al. 1984; Lundberg, Franco-Cereceda et al. 1985). It is intriguing to know whether activin induced hyperalgesia is due to the activity of any of the immune cells. For example, neutrophils depleted or mast cells predegranulated animals can be used to see whether activin induced pain hypersensitivity depends on neutrophils or mast cells function

(Lewin, Rueff et al. 1994; Bennett, al-Rashed et al. 1998).

It is worth mentioning that NGF is also involved in the inflammatory response and is probably regulated in a similar manner. After inflammation, immune cells release NGF which not only directly sensitizes nociceptors but also indirectly does so via activating immune cells, for example, NGF can cause mast cell degranulation to release amines, cytokines or proteases (Mazurek, Weskamp et al. 1986; Lewin, Rueff et al. 1994; Woolf,

Ma et al. 1996; Lambiase, Micera et al. 2004). Furthermore, NGF may be chemotactic for neutrophils (Gee, Boyle et al. 1983; Boyle, Lawman et al. 1985) which partially

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contributes to NGF induced thermal hyperalgesia (Bennett, al-Rashed et al. 1998; Foster,

Wicks et al. 2002).

Inflammation causes abnormal pain through the release of inflammatory mediators, cytokines and growth factors, which include bradykinin, prostaglandins, histamine, serotonin, NGF, activin, interleukin-1β (IL-1β) and TNFα. Most of these factors directly sensitize or activate nociceptors. However, it is still a puzzle whether these factors work synergistically together or in a sequential order to cause pain. Both mechanisms have been proposed. For example, carrageenan induced mechanical hyperalgesia is initiated by bradykinin leaked from the dilated blood vessel, which facilitates the release of TNFα.

TNFα then triggers the release of IL-6/ IL-1β which further liberates prostaglandins

(Cunha, Lorenzetti et al. 1991; Cunha, Poole et al. 1992). By contrast, in the CFA inflammation model, NGF up-regulation has been reported to be secondary to an earlier increase of IL-1β and later a sequential release cascade stemming from TNFα to IL-1β and finally NGF is proposed, although IL-1β is not the only cytokine that can induce

NGF in this inflammation model (Safieh-Garabedian, Poole et al. 1995; Woolf, Allchorne et al. 1997). On the other hand, in a mouse writhing model induced by acetic acid, the writhing response was mediated by TNFα, IL-1β, and IL-8 in a concomitant and synergistic fashion (Ribeiro, Vale et al. 2000). Activin can induce the expression of

TNFα, IL-1β, prostanoid in rat bone marrow-derived macrophages (Nusing and Barsig

1999) and TNFα, IL-1β can increase the expression of activin in keratinocytes and fibroblasts (Hubner and Werner 1996). In an acute systemic inflammation model induced by lipopolysaccharide (LPS), activin increase within an hour after injection, earlier than

TNFα and IL-6 (Jones, Brauman et al. 2000). These data suggest that inhibiting activin

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may inhibit the cytokine cascade and largely stop inflammatory responses. Then anti-

activin medicine will provide a very promising new strategy to manage inflammatory

pain.

Activin may be a candidate target for development of novel pain therapeutics

The management of pain is still a major challenge since most common pain relievers

have suboptimal therapeutic efficacies and considerable side effects.

In this thesis, I have shown that increased activin in the skin after inflammation can induce acute nociceptive responses through a TRPV1 dependent mechanism. I have also shown that activin contribute to prolonged pain response probably through regulating neuropeptides important for pain including CGRP. These changes caused by activin are unlikely to be permanent since the mechanism found so far involves sensitizing TRPV1 channels and increased CGRP expression. The phenomenon that activin sensitizes

TRPV1 channel can probably be reversed by dephosphorylation or downregulate TRPV1.

CGRP expression largely depends on the target-derived factors, since the nerve axotomy decreases CGRP expression (Doughty, Atkinson et al. 1991). It is reasonable to speculate

that when activin level decreases in the skin, the CGRP expression will also drop in the

DRG. It seems unlikely that activin causes change in nerve circuits in the central nervous system. Therefore, inhibiting activin may relieve both acute and chronic inflammatory pain and further, stopping activin function at the early onset of the injury may prevent the development of chronic pain.

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Systemic inhibition of activin is probably not a good idea; however, activin has a wide

range of different functions. In the reproduction organs, it induces the release of follicle

stimulating hormone, which stimulates the maturation of germ cells (Bilezikjian, Blount et al. 2006). During pregnancy, activin level increases in the serum in the third trimester

and may be involved in normal labor (Woodruff, Sluss et al. 1997). Interesting, pregnant

women are more sensitive to radiant heat during the last two weeks of pregnancy

(Goolkasian and Rimer 1984), which might be due to the increased activin. However,

activin is also important for multiple organ development and inhibiting activin may

jeopardize the development of the fetus. Activin also protects tissues such as kidney and

brain against ischemic injuries (Tretter, Hertel et al. 2000; Maeshima, Zhang et al. 2001;

Mukerji, Katsman et al. 2006). Furthermore, activin has been shown to promote skin

wound healing by increasing healing speed but also increases scar formation (Munz,

Smola et al. 1999). On the other hand, the function of activin in nerve injury needs further study. These data suggest drugs like an anti-activin antibody might be suitable to control some types of pain such as inflammatory pain but not others, such as postoperative pain, and needs to be delivered locally to avoid side effects.

Recently, activin has been suggested to be involved in multiple inflammatory diseases, such as gastrointestinal inflammation, arthropathy and systemic inflammation (Werner and Alzheimer 2006). Local delivery of anti-activin antibody as a drug is a very promising treatment for this inflammatory pain. In particular, activin is secreted by macrophages and synoviocytes and elevated in joint synovia in rheumatoid arthritis and inhibiting activin in the joints may provide good pain relief (Yu, Dolter et al. 1998). A recent clinical trial has shown that local delivery of an anti-NGF antibody reduces

153

osteoarthritis pain (Genentech). Maybe in the near future, Activo (commercial name for

anti-activin antibody) will be used for treat rheumatoid arthritis. The pathogenesis of

migraine includes increased CGRP and blocking CGRP signals shows valuable therapeutic function for treating migraine (Edvinsson 2004; Theoharides, Donelan et al.

2005; Durham 2006; Durham, Niemann et al. 2006). Given that activin can regulate

CGRP expression, it is promising that Activo will also relieve migraine. In this thesis, I

applied both in vitro and in vivo experiments to understand the molecular mechanism of

activin function in the inflammatory pain. I present evidence to support that activin

contributes to both acute inflammatory pain through sensitizing TRPV1 channel and

prolonged inflammatory pain by increased CGRP expression. These data provide the

mechanisms and promise for the new inflammatory pain reliever Activo.

154

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