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GPER/GPR30 : A TARGET FOR MODULATION

A Dissertation

Submitted to

The Temple University Graduate Board

In Partial Fulfillment

of the Requirements for the Degree of

Doctor of Philosophy

By

Elena Deliu

May, 2012

Examination Committee Members:

Dr. Nae J Dun (Advisor), Dept. of Pharmacology, Temple University

Dr. Mary E Abood, Dept. of Anatomy and Cell Biology, Temple University

Dr. Barrie Ashby, Dept. of Pharmacology, Temple University

Dr. Eugen Brailoiu, Dept. of Pharmacology, Temple University

Dr. Ellen M Unterwald, Dept. of Pharmacology, Temple University

Dr. Hreday Sapru, (External Examiner), Depts. of Neurosciences, Neurosurgery &

Pharmacology/Physiology, UMDNJ-NJMS

i ©

2012

By

Elena Deliu

All Rights Reserved

ii ABSTRACT

GPER/GPR30 : A TARGET FOR PAIN MODULATION

Elena Deliu

Doctor of Philosophy

Temple University, 2012

Doctoral Advisory Committee Chair: Nae J. Dun, Ph.D.

The G -coupled estrogen receptor GPER/GPER1, also known as GPR30, was originally cloned as an and later shown to be specifically activated by 17-

β-. This has led to its classification as an estrogen receptor and expanded the perspective on the mechanisms underlying the rapid estrogenic effects reported over the years. GPER is strongly expressed in the central nervous system and peripheral tissues and appears to be involved in a wide variety of physiological and pathological processes.

Estrogens are known to alter the processing of nociceptive sensory information and responses in the central nervous system. Both analgesic and pro-nociceptive effects of have been reported. Some pro-algesic estrogenic responses have a short latency, suggesting a non-genomic mechanism of action. Immunohistochemical studies in rodents prove the existence of GPER in pain-relevant areas of the nervous system such as dorsal root ganglia, superficial dorsal horn of the , (PAG), , trigeminal sensory nucleus and . In the periphery, activation of GPER results in pro-nociceptive effects. However, GPER involvement in pain processing at central levels is largely unexplored. Thus, the work

iii presented in this thesis was aimed at investigating whether GPER modulates nociception at spinal and supraspinal sites.

The behavioral response to GPER activation in the spinal cord and PAG was evaluated in an acute grooming test (scratching, biting and licking behavior) and in the hot plate test, respectively. Intrathecal challenge of mice with the GPER G-1 (0.1-

1 nmol) induced a dose-dependent increase in pain-related behaviors, that was abolished by pre-treatment with the GPER antagonist G15 (1-10 nmol), confirming GPER specificity of the response. Likewise, intra-PAG microinjection of G-1 (10-100 pmol) to rats reduced the nociceptive threshold in the hot plate test, an effect that was G15 sensitive. To obtain further insight on the mechanisms involved in the behavioral effects observed in whole animals, we tested the effect of GPER ligands on neuronal membrane

2+ potential, intracellular calcium concentration ([Ca ]i) and reactive oxygen species (ROS)

2+ accumulation. The membrane depolarization and the increases in [Ca ]i and ROS levels are markers of neuronal activation, underlying pain sensitization in the spinal cord and pain facilitation in the PAG. Electrophysiological recordings from superficial dorsal horn and lateral PAG neurons indicate neuronal depolarization upon G-1 application, an effect that was fully prevented by G15 pre-treatment. Both cultured spinal neurons and cultured

2+ PAG neurons responded to G-1 administration by elevating [Ca ]i and mitochondrial and cytosolic ROS levels. In the presence of G15, G-1 did not elicit the calcium and ROS responses.

Collectively, these results demonstrate that GPER modulates both the ascending and descending pain pathways to increase nociception via cytosolic calcium elevation and

ROS accumulation in spinal and PAG neurons, respectively. These findings broaden the

iv current knowledge on GPER involvement in physiology and pathophysiology, providing the first evidence of its pro-nociceptive effects at central levels and characterizing some of the mechanisms involved. Moreover, we show for the first time ROS accumulation downstream of GPER activation, extending the current understanding of GPER signaling.

v ACKNOWLEDGEMENTS

I have learned a lot as a graduate student in the Department of Pharmacology at

Temple University. It has been both a scientific and important life experience that contributed to who I am today. I owe my deepest gratitude to several people that helped me arrive at this moment.

I would particularly like to express my thankfulness to Dr. Dun for welcoming me in his lab and for his coordination throughout my research endeavors. I strongly appreciate that he encouraged me to study electrophysiology and also that he supported my research independence. I have benefited a lot from his flexibility, openness to questions, prompt replies and germane observations. I am really grateful for everything he has done for me.

It is difficult to overstate my gratitude for all the guidance, ideas and support Dr. E.

Brailoiu provided, as well as for all the trust he put in me despite the little research experience I had. He generously shared his expertise, knowledge and enthusiasm and got me involved in several projects. It’s been a tremendous opportunity to work with him and learn so much about the beauty of research.

I sincerely thank Dr. Ashby for giving me all the attention I needed at every stage.

Throughout these years, he was always available and took his time to answer questions, revise some of the drafts for previous examinations, as well as to attend the rehearsals of my presentations. His suggestions were extremely useful and I truly appreciate all the valuable observations and genuine support he provided.

I am very thankful to Dr. Unterwald and Dr. Abood for their insightful comments, very good questions regarding my project and for being so supportive.

vi I will not forget my first interaction with Dr. Unterwald and her patience in advising me how to adjust the protocol for the locomotor activity test so that it would fit the timings of other behavioral experiments I was performing at the time. Her attitude allowed me to have a bit more confidence in my reasoning, although I was at a very early stage in my development as a scientist. I am very grateful for that.

I also need to thank Dr. Abood for her enthusiasm and generous encouragement. I’m happy to have had the chance to interact with her and other people in her lab; I’ve learned research can often be rather enjoyable than stressful.

I would like to thank Dr. Sapru for taking time to serve as an external examiner on my Committee.

My special thanks go to Dr. Cristina Brailoiu for being a true prototype of hard work, dedication, generosity and cheerfulness. I have found in her a role model I so much longed to follow and I am truly blessed to have benefited from her friendship and scientific advice.

I am very thankful to Dr. Khalid Benamar for helping me complete the behavioral study of GPER involvement in supraspinal pain. He shared his time, chemicals, experimental settings and technical expertise and his contribution has been extremely valuable.

I am very appreciative of Dr. Lynn Kirby’s advice in electrophysiology. Her suggestions were priceless and helped me continue my experiments after stalling because of so many technical difficulties. I would also like to thank Dr. Alan Cowan for sharing his mice and observation boxes, which were helpful for learning the intrathecal injection procedure and performing behavioral tests, respectively.

vii I would like to thank Dr. J.B. Arterburn and Dr. T.I. Oprea for providing the GPER ligands.

I am very thankful to Mary Dun for sharing her immunohistochemical expertise and for so many discussions we’ve had in the lab, which greatly reduced my feeling of loneliness and created a more relaxed atmosphere.

Also, lots of thanks to Xin Gao and Xiaofang Huang for helping me with RT-PCR,

Western blotting and cell culture, as well as for being such dear presences in the lab and outside the lab. Among the graduate students Fan Yang and Neil Lamarre were my best friends; they have always been there to help and encourage me and I’m happy to express here my heartfelt gratitude towards them. I’ve also benefited from the help of, and the discussions with Nicole Enman, Bonnie Quattrochi, Bogdan Gurzu, Michael Jan and

Saadet Inan, and I thank them very much.

I am very thankful to the faculty in the Departments of Pharmacology, of Anatomy and Cell Biology, of Microbiology and Immunology and of Biochemistry, for sharing their knowledge and expertise during the courses and seminars at Temple University. I would also like to thank Liz Molina and Lor Hatch for their assistance and general positive attitude.

I cannot leave out so many other people that encouraged me to pursue a Ph.D. degree, former Professors, research assistants and colleagues at the “Carol Davila” University of

Medicine and Pharmacy in Bucharest and at the University of Iowa. Thank you very, very much!

This thesis is dedicated to those very dear ones that supported me wholeheartedly and wiped my tears along the way.

viii TABLE OF CONTENTS

PAGE

ABSTRACT ...... iii

ACKNOWLEDGEMENTS ...... vi

LIST OF TABLES ...... xii

LIST OF FIGURES ...... xiii

LIST OF EQUATIONS ...... xv

LIST OF ABBREVIATIONS ...... xvi

CHAPTER ONE: INTRODUCTION ...... 1

1.1. Estrogens ...... 1

1.1.1. Nuclear estrogen receptors ...... 1

1.1.2. Non-nuclear estrogen receptors ...... 3

1.1.3. The -coupled estrogen receptor (GPER) ...... 4

1.1.4. Tissue distribution of GPER ...... 6

1.1.5. Cellular localization of GPER ...... 7

1.1.6. GPER selective and non-selective ligands ...... 7

1.1.7. Estrogen involvement in health and disease ...... 9

1.1.7.1. Physiological effects of estrogen in reproductive organs ...... 10

1.1.7.2. Non-reproductive effects of estrogens in peripheral tissues ...... 10

1.1.7.3. Estrogenic effects in the central nervous system ...... 11

1.1.7.4. GPER involvement in health and disease ...... 12

1.2. Pain ...... 14

1.2.1. Spinal cord involvement in pain ...... 16

ix 1.2.2. Supraspinal sites of pain modulation ...... 18

1.2.3. Neuronal pathways in pain: calcium, depolarization, ROS ...... 20

1.2.4. Animal models of nociception ...... 23

1.2.4.1. Spinal pain: caudally directed biting and scratching in response to

spinal delivery of nociceptive molecules ...... 23

1.2.4.2. Supraspinal pain: hot plate test ...... 24

1.3. Estrogens and pain ...... 24

1.3.1. Estrogens and spinal pain ...... 25

1.3.2. Estrogens and supraspinal pain ...... 26

1.3.3. GPER and pain ...... 26

1.4. Specific aims ...... 28

CHAPTER TWO: EXPERIMENTAL PROCEDURES ...... 33

2.1. Drugs ...... 33

2.2. Experimental animals ...... 33

2.3. Mouse GPER mRNA analysis by RT-PCR ...... 35

2.4. Immunohistochemistry ...... 36

2.5. Intrathecal injection in mice ...... 37

2.6. Behavioral assay of spinal nociception ...... 37

2.7. Intra-PAG cannulation of rats ...... 38

2.8. Hot plate test ...... 38

2.9. Electrophysiology ...... 39

2.10. Neuronal culture ...... 41

2.11. Calcium imaging ...... 41

x 2.12. Detection of cytosolic ROS accumulation ...... 43

2.13. Detection of mitochondrial superoxide accumulation ...... 43

2.14. Statistical analysis ...... 44

2.15. Western blot for ERK1/2 – preliminary results ...... 44

CHAPTER THREE: RESULTS ...... 46

3.1. Spinal GPER activation is pro-nociceptive ...... 46

3.2. GPER is present in the mouse spinal cord at mRNA and protein level ...... 49

3.3. Supraspinal GPER activation is pro-nociceptive ...... 50

3.4.1. GPER activation with G-1 results in depolarization of spinal superficial dorsal

horn neurons ...... 53

3.4.2. G-1 depolarizes neurons located in the lateral PAG ...... 54

2+ 3.5.1. GPER activation elevates [Ca ]i in cultured spinal neurons ...... 57

2+ 3.5.2. GPER activation elevates [Ca ]i in cultured PAG neurons ...... 59

3.6. GPER activation results in cytosolic and mitochondrial ROS accumulation

...... 61

CHAPTER FOUR: DISCUSSION ...... 66

4.1. GPER and spinal pain ...... 66

4.2. GPER and supraspinal pain ...... 70

4.3. Summary and conclusions ...... 75

4.4. Future directions ...... 77

REFERENCES ...... 84

xi LIST OF TABLES

Table Page

Table 1 Time course of %MPE in the rat hot plate test ...... 51

xii LIST OF FIGURES

Figure 1 Domain structures of classical estrogen receptors α and β ...... 2

Figure 2 Simplified model of estrogen action in a target cell ...... 3

Figure 3 Structures of estrogen (17- β-estradiol, E2), GPER agonist G-1 and GPER antagonists G15 and G36 ...... 9

Figure 4 Pain pathways ...... 15

Figure 5 Hyperexcitability of dorsal horn neurons results in neuropathic pain ...... 17

Figure 6 Neuronal connections in the descending pain pathways from the PAG to the superficial laminae of the spinal cord ...... 19

Figure 7 Reactive oxygen and nitrogen species involvement in pain sensitization .....22

Figure 8 Effect of intrathecal G-1 on ERK1/2 phosphorylation ...... 29

Figure 9 Putative mechanisms of pro-nociceptive effects of GPER activation in the spinal cord and periaqueductal gray ...... 30

Figure 10 Spinal GPER activation results in nociceptive behavior in mice ...... 48

Figure 11 GPER expression in the mouse spinal cord ...... 49

Figure 12 Intra-PAG microinjection of G-1 is pro-nociceptive ...... 52

Figure 13 GPER stimulation results in depolarization of superficial dorsal horn neurons in the rat spinal cord ...... 54

Figure 14 Effects of GPER stimulation or blockade on membrane potential of lateral

PAG neurons ...... 56

Figure 15 G-1 application elevates cytosolic Ca 2+ in spinal neurons ...... 58

Figure 16 G-1 application elevates cytosolic Ca 2+ in cultured PAG neurons ...... 60 xiii Figure 17 GPER-mediated cytosolic and mitochondrial ROS accumulation in spinal neurons ...... 62

Figure 18 GPER-mediated cytosolic and mitochondrial ROS accumulation in PAG neurons ...... 63

Figure 19 Cytosolic ROS dye fluorescence in response to GPER activation in cultured spinal and PAG neurons ...... 64

Figure 20 Time-course of mitochondrial ROS accumulation ...... 65

Figure 21 Effects of selective receptor on G-1 induced pain-related behavior in mice ...... 78

Figure 22 Pre-treatment with selective opioid agonists prevents G-1 induced depolarization of superficial dorsal horn neurons ...... 79

xiv LIST OF EQUATIONS

Equation 1 Calculation of % maximal possible effect in the hot plate test ...... 39

Equation 2 Quantification of intracellular calcium concentration using Fura-2 fluorescent dye ...... 42

Equation 3 Quantification of ERK1/2 for preliminary studies ...... 45

xv LIST OF ABREVIATIONS

ACSF, artificial cerebrospinal fluid

AM, acetoxy methyl ester

AMP, adenosine monophosphate

AMPA, -amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid

ANOVA, analysis of variance apoE, apolipoprotein E

BK,

2+ [Ca ]i, intracellular calcium ions concentration cAMP, cyclic adenosine monophosphate cDNA, complementary DNA cGMP, cyclic guanosine monophosphate

CNS, central nervous system

COX-2, cyclooxygenase-2

CREB, cAMP response element-binding protein

Cys, cysteine cytoROS, cytosolic ROS

DBD, DNA binding domain

DCF, dichlorofluorescein df, degrees of freedom

DHPG, 3,5-dihydroxyphenylglycine (mGluR1/5 agonist)

DNA, deoxyribonucleic acid

xvi DPN, (ER β selective agonist)

E2, 17- β-estradiol

EAAC1, excitatory amino acid carrier 1 (the EAAT3 ortholog in rodents)

EAAT, excitatory amino acid transporter

ERs, nuclear estrogen receptors

ERE, estrogen response element

ERK1/2, extracellular signal regulated kinase 1/2

FITC, fluorescein isothiocyanate

GABA, gamma aminobutyric acid

GIRK, G protein-activated inwardly rectifying K + channels

GLAST, glutamate aspartate transporter (the EAAT-1 ortholog in rodents)

GLT-1, glutamate transporter 1 (the EAAT-2 ortholog in rodents)

GPCR, G protein-coupled receptor

GPER, G protein-coupled estrogen receptor

GPR30, G protein-coupled receptor 30

GS, glutamine synthetase

GSH, glutathione

H2-DCF, 2’,7’-dichlorodihydrofluorescein

H2-DCFDA, 2’,7’-dichlorodihydrofluorescein diacetate (cytosolic ROS indicator)

HBSS, Hanks balanced salt solution

HDL, high density lipoproteins

HEK, embryonic

HRE, hormone response elements xvii HSP, heat shock protein i.c.v., intracerebroventricular

IgG, immunoglobulin G i.p., intraperitoneal i.t., intrathecal

IUPHAR, International Union of Basic and Clinical Pharmacology

LBD, binding domain

LDL, low density lipoproteins

LTP, long term potentiation

MAPK, mitogen activated protein kinases mER, membrane estrogen receptor mGluRs, metabotropic glutamate receptors mitoROS, mitochondrial ROS

MPE, maximum possible effect mRNA, messenger ribonucleic acid

NADPH, adenine dinucleotide phosphate

NK1, neurokinin 1

NMDA, N-methyl D-aspartate

NMDAR, N-methyl D-aspartate receptor

NO, nitric oxide

NOS, nitric oxide synthase

NR,

NS, nociceptive specific

xviii PAG, periaqueductal gray

PBS, phosphate buffered saline pERK, phosphorylated ERK

PGE 2, prostaglandin E 2

PI 3K, phosphoinositide 3 kinase

PKA,

PKC, protein kinase C

PLC, phospholipase C

PPT, propylpyrazole-triol (ER α selective agonist)

RNA, ribonucleic acid

RNS, reactive nitrogen species

ROS, reactive oxygen species

RVM, rostroventral medulla s.c., subcutaneous

S.E.M., standard error of mean

SDH, superficial dorsal horn

SDS, sodium dodecyl sulphate

SOD, superoxide dismutase

TBS, Tris buffered saline tERK, total ERK

TRPV1, transient receptor potential vanilloid subclass 1 ion channel

UV, ultraviolet

WDR, wide dynamic range

xix CHAPTER ONE

INTRODUCTION

1.1. Estrogens

Naturally occurring estrogens include estradiol, and . The most active form of estrogen is 17 β-estradiol, E2, also referred here as “estrogen” or “estradiol”.

They are biosynthesized from androgens by the aromatase, an irreversible process that occurs in both peripheral organs and central nervous system of males and females (Boon et al., 2010).

Estrogen is a member of the hormone family that also includes , , glucocorticoids and mineralocorticoids. Although considered to be the female steroid because of its traditional role in the female reproductive system, it is acknowledged today that estrogen is involved in a far wider range of actions, including metabolic regulation and neurological and behavioral effects in both male and female

(Boon et al., 2010).

1.1.1. Nuclear estrogen receptors

Classically, estrogen is known to activate nuclear receptors (ER α and ER β), which are sequence specific DNA-binding that recognize cis -acting hormone response elements (HRE) located typically in the promoter and enhancer regions of target .

This mechanism of action accounts for its long lasting effects that appear with a latency of hours to days. The two classical estrogen receptors described to date: ER α (Jensen and DeSombre, 1973) and ER β (Kuiper et al., 1996; Mosselman et al., 1996) are referred 1 to herein as ERs. ER α and ER β are highly homologous in their DNA- and ligand-binding domains (Fig. 1), however, they lack in their transcriptional and activation domains and present different tissue distribution (Kuiper et al., 1997;

Dechering et al., 2000), supporting the functional differences observed between the two receptor subtypes.

Fig. 1. Domain structures of classical estrogen receptors ααα and βββ.β... DBD, DNA binding domain; AF, activation domain (AF-1, AF-2), LBD, ligand binding domain. The percentages indicate the amino acid identity between domains of ER α and ER β (Edwards, 2005).

Both ERs are soluble proteins, that can shuttle between the cytoplasm and the nucleus, but are found predominantly (95%) in the nucleus. Upon ligand binding, ERs become activated and dissociate from protein chaperones, change their conformation, form homodimers and bind to HRE of target genes. DNA-bound receptors alter transcription rates of target genes via recruitment of coactivator or corepressor proteins, which are devoid of DNA binding activity and are recruited through protein-protein interaction, resulting in the formation either of an enzymatic protein complex capable of chromatin remodeling or of a “protein bridge” facilitating the assembly of RNA polymerase II initiation complex (Fig. 2). This ultimately leads to regulation of mRNA synthesis and thus, protein synthesis, resulting in cellular response.

2 Fig. 2. Simplified model of estrogen action in a target cell. NR, nuclear receptor; HSP, heat shock protein, HRE, hormone response element (http://human.freescience.org/htmx/nuclear_receptor.php )

Thus, the main feature of nuclear estrogen receptors is to control expression of specific sets of target genes; these effects are delayed in onset and prolonged in duration, as they depend on protein synthesis.

1.1.2. Non-nuclear estrogen receptors

A growing body of compelling evidence describes rapid estrogenic effects that occur on a scale of seconds to minutes and thus cannot be attributed to a genomic mechanism of action. Estradiol induces fast activation of protein kinases (such as MAPKs, PI 3K, PKC) phosphatases, release of cAMP, cGMP and calcium, in a variety of cell types (Losel and

3 Wehling, 2003; Levin, 2005). These rapid effects, much too fast to involve transcription, have been designated as “non-genomic” and there is increasing interest in the receptors involved in rapid, non-genomic estrogen signaling. Four types of receptors have been proposed. First, novel transmembrane receptors unrelated to nuclear hormone receptors, such as GPR30 (GPER) or a membrane estrogen receptor (mER) appear to mediate short-latency estrogen signaling. Second, there are transmembrane receptors for / hormones allosterically modulated by steroid hormones. Third, modified forms of the conventional steroid preferentially localized to the plasmalemma have also been suggested. Lastly, there are indications of a subpopulation of conventional steroid receptors that associates with sites of signaling complexes in the cytoplasm or the plasma cell membrane (Edwards, 2005).

1.1.3. The G protein coupled estrogen receptor (GPER)

In the late 1990s, multiple groups identified GPR30 as an orphan G-protein coupled receptor with 30% homology to the chemokine subfamily of GPCRs (Owman et al.,

1996; Carmeci et al., 1997; Takada et al., 1997; O'Dowd et al., 1998). A series of reports followed, which showed that GPR30 responded to E2 stimulation. In cells lacking nuclear ERs, but positive for GPR30 (SKBr3), E2 rapidly activates ERK1/2 (a MAPK) via G βγ -dependent transactivation of the epidermal growth factor receptor (Filardo et al.,

2000). Moreover, only when GPR30 is present in MDA-MB231 cells (ER α negative,

ER β positive), does E2 trigger a Gs-dependent activation of the -cAMP-

PKA pathway that results in Raf-1 inactivation and attenuation of ERK1/2 activity

(Filardo et al., 2002). In macrophages, E2 elicits a GPR30-dependent transcriptional 4 activation of nerve growth factor by inducing c-Fos expression via cAMP (Kanda and

Watanabe, 2003a). In human keratinocytes, E2 stimulates GPR30 to inhibit oxidative stress-induced apoptosis via an increase in Bcl-2 expression (Kanda and Watanabe,

2003b), as well as to elicit proliferation via cyclin D2 expression (Kanda and Watanabe,

2004); both responses occur in response to CREB phosphorylation by PKA, dependent on cAMP. Furthermore, in GPR30-expressing cells (SKBr3, GPR30- transfected MDA-MB231 and BT-20) E2 and two phytoestrogens ( and ) induce c-fos upregulation through the GPR30-dependent ERK1/2 activation

(Maggiolini et al., 2004).

In 2005, two groups independently demonstrated the existence of a GPR30 binding site specific for E2 (Revankar et al., 2005; Thomas et al., 2005). Several reports and reviews followed, supporting specific estrogenic activation of GPR30 and proposing it as a G protein-coupled estrogen receptor (Filardo and Thomas, 2005; Hasbi et al., 2005; Rae and Johnson, 2005; Funakoshi et al., 2006; Manavathi and Kumar, 2006; Pedram et al.,

2006; Thomas and Dong, 2006; Vivacqua et al., 2006; Albanito et al., 2007; Filardo et al., 2007; Haas et al., 2007; Hsieh et al., 2007; Prossnitz et al., 2007; Revankar et al.,

2007; Smith et al., 2007; Watson et al., 2007).

Consequently, IUPHAR classified GPR30 as a G-protein coupled estrogen receptor and initially named it GPER-1, then GPER.

( http://www.iuphar-db.org/DATABASE/ObjectDisplayForward?objectId=221 )

5 1.1.4. Tissue distribution of GPER

Expression of GPER has been characterized in the zebra finch (Acharya and

Veney, 2011) and Atlantic croaker ovaries (Pang et al., 2008), as well as in human tissues and tumors and in the rodent brain and peripheral organs. The original reports demonstrated high levels of GPER mRNA in the human , intestine, ovary and brain

(O'Dowd et al., 1998) and various levels of GPER transcripts in the placenta, lung and (Owman et al., 1996; Carmeci et al., 1997; Takada et al., 1997). Increased GPER protein expression has been reported in cancer cell lines and human primary malignancies of the breast (Carmeci et al., 1997; Filardo et al., 2000; Revankar et al., 2005; Albanito et al., 2008b), endometrium (Vivacqua et al., 2006; Leblanc et al., 2007; He et al., 2009), ovaries (Albanito et al., 2007; Albanito et al., 2008a; Henic et al., 2009), thyroid

(Vivacqua et al., 2006), lung (Siegfried et al., 2009), prostate (Chan et al., 2010) and testicular germ cells (Franco et al., 2011). In the rodent brain, GPER immunoreactivity was detected in the hypothalamic-pituitary axis, , brainstem autonomic nuclei and striatum (Brailoiu et al., 2007), as well as in the cortex, midbrain pontine nuclei, coeruleus and periaqueductal gray, trigeminal nuclei and cerebellum

Purkinje layer of the hindbrain (Hazell et al., 2009). GPER protein expression also occurs in the rat autonomic ganglia, dorsal root ganglia and superficial and deep laminae of the spinal cord (Dun et al., 2009; Takanami et al., 2009). In the rat and mouse periphery, adrenal medulla, renal pelvis and ovaries express GPER both at protein and mRNA levels

(Hazell et al., 2009).

6 1.1.5. Cellular localization of GPER

A consensus regarding the cellular distribution of GPER has not been reached yet.

Both plasma membrane and subcellular localizations have been reported. Apparently,

GPER transfected HEK 293 cells (Filardo et al., 2007), the human breast cancer cell line

SKBr3 (Thomas et al., 2005), as well as CA2 pyramidal neuronal cells and FLAG-GPER transfected HeLa cells (Funakoshi et al., 2006) express GPER on the plasma membrane.

However, when GPER is expressed in COS7 cells, it localizes to the (Revankar et al., 2005; Otto et al., 2008); the same subcellular localization has been reported for cells endogenously expressing GPER, such as HEC50 (Otto et al.,

2008). In neurons of the rat paraventricular and supraoptic nuclei, GPER is preferentially expressed in the Golgi apparatus and to a lesser extent in the endoplasmic reticulum, as shown by immunofluorescence and electron microscopy studies (Sakamoto et al., 2007). The same pattern of GPER distribution is reported for pyramidal cells of

CA1-3 and granule cells of the dentate gyrus in the rat hippocampal formation (Matsuda et al., 2008). In hypothalamic neurons, normal and malignant mammary and ovarian epithelia, GPER is similarly concentrated in organelles, while in the renal epithelial cells, it is localized to the basolateral membrane (Cheng et al., 2011). Functional studies in rat myometrial cells also support an intracellular localization for GPER (Tica et al., 2011).

1.1.6. GPER selective and non-selective ligands

In addition to 17- β-estradiol, GPER also interacts other ER ligands, such as:

• estriol - a GPER antagonist at high concentrations (Lappano et al., 2010),

7 • 4-hydroxytamoxifen - GPER agonist; also acts as ER agonist in some tissues and

ER antagonist in others, thus it is a selective estrogen (Filardo

et al., 2000; Jordan, 2007),

• /ICI 182,780 - ER antagonist, GPER agonist (Wakeling et al., 1991;

Filardo et al., 2002),

• phytoestrogens, such as genistein and quercetin (Maggiolini et al., 2004), as well

as , a non-steroidal equine estrogen that is a metabolite of the isoflavone

– GPER agonists (Rowlands et al., 2011),

• environmental estrogens such as – GPER agonists (Chevalier et al.,

2012),

• the herbicide – ER α and GPER agonist (Albanito et al., 2008a),

• , such as dichlorodiphenyltrichloroethane – GPER agonists (Thomas

and Dong, 2006)

In 2006 Bologa et al. identified G-1 as a GPER selective agonist, inactive against classical estrogen receptors (Bologa et al., 2006) and without binding activity on 25 other

G protein-coupled receptors (Blasko et al., 2009) or in Gper knockout mice (Haas et al.,

2009; Liu et al., 2009; Wang et al., 2009). Subsequently, GPER physiology was intensely studied in various systems, its activation resulting in calcium mobilization, PI 3K activation, ERK1/2 phosphorylation, stimulation of cyclic AMP – protein kinase A pathway, as well as activation of protein kinase C (Prossnitz et al., 2008; Prossnitz and

Maggiolini, 2009).

In 2009, G15 was identified as the first GPER-selective antagonist (Dennis et al.,

2009), followed by G36 in 2011, an antagonist with higher GPER-selectivity than G15 8 (Dennis et al., 2011). All G-1 effects tested, as well as many E2-dependent responses are inhibited by G15 (Dennis et al., 2009; Gingerich et al., 2010; Jenei-Lanzl et al., 2011;

Lindsey et al., 2011; Peyton and Thomas, 2011).

The structures of E2, G-1, G15 and G36 are shown in Fig. 3.

Fig. 3. Structures of estrogen (17- βββ-estradiol, E2), GPER agonist G-1 and GPER antagonists G15 and G36 (Dennis et al., 2011).

1.1.7. Estrogen involvement in health and disease

Estrogens affect a broad range of physiological and pathological processes in mammals, acting on reproductive organs and peripheral tissues, as well as in the central nervous system. Abnormal estrogen levels or dysfunctions of its receptors are involved in

9 the development or progression of various types of cancer (breast, cervical, endometrial, ovarian, colorectal, prostate), endometriosis, osteoporosis, cardiovascular disease, obesity, resistance, lupus erythematosus, neurodegenerative diseases etc (Deroo and Korach, 2006).

1.1.7.1. Physiological effects of estrogen in reproductive organs

In the female, estrogens stimulate the growth and maturation of reproductive organs and , the start of menstrual cycle and maintain the function of the reproductive system (Couse and Korach, 1999). In the male, some of the effects of testosterone are mediated through its aromatase-dependent conversion to estrogen. For instance, estrogen appears to be critical in the normal development of male gonadal functions and by controlling stem cell number and spermatid maturation in the seminiferous tubules (Hess, 2003).

1.1.7.2. Non-reproductive effects of estrogens in peripheral tissues

In the cardiovascular system estrogens induce angiogenesis, , protection against atherosclerosis, prevention of apoptosis in endothelial cells; thrombogenesis and blood clotting. In the liver upregulation of apolipoprotein E and of apoE/B LDL receptor in hepatocytes has been reported, which results in blood clearance of LDL and increase of circulatory HDL. In the , stimulation of epiphyseal closure and maintenance of normal bone mass (inhibition of bone resorption) occurs in response to estradiol. In the skin, estrogens are important for the maintenance of hydration and the stimulation of

10 collagen synthesis (Couse and Korach, 1999; Deroo and Korach, 2006; Boon et al.,

2010).

1.1.7.3. Estrogenic effects in the central nervous system

In addition to regulation of the and other brain regions related to reproduction, estrogens have been found to have widespread influence throughout the brain, such as the basal forebrain cholinergic system, the hippocampus and , the caudate-putamen, midbrain raphe and brainstem , and the spinal cord. These systems are involved in a variety of estrogen actions on mood, locomotor activity, pain sensitivity, vulnerability to epilepsy, as well as attentional mechanisms and cognition (McEwen and Alves, 1999); e.g.:

• In the ventromedial hypothalamus, estrogens regulate female sexual behavior via

modulation of , induction of oxytocin receptors,

progesterone receptor and cyclic synaptogenesis.

• In the basal forebrain cholinergic system, there is upregulation of cholinergic

markers and nerve growth factor receptors in response to estrogen, which

promotes neural survival. In this central nervous system site, sex differences are

programmed during early development.

• In the midbrain serotonergic system, E2 modulates tryptophan hydroxylase,

serotonin transporters and certain serotonin receptor subtypes. At this level, sex

differences are apparent in progestin receptor expression and in serotonin

turnover.

11 • In the midbrain and hypothalamic dopamine system and projections, estrogens

elicit developmentally programmed sex differences in incertohypothalamic

dopamine neuron number and function. In addition, studies in rats show that

facilitation of - or -stimulated dopamine release and

locomotor activity occurs in response to estrogen.

• In the brainstem catecholaminergic system, there is estrogen-dependent regulation

of tyrosine hydroxylase gene and immediate early gene expression.

• In the hippocampus, E2 triggers de novo synapse formation on pyramidal neurons

via modulation of NMDA receptors.

• In the spinal cord, there are sex differences in estrogen modulation of nociception

in and animals.

• In glial cells, estrogens influence the normal- and lesion-induced-synaptic

plasticity via regulation of glial specific genes, such as glial fibrillary acid protein

and apolipoprotein E within astrocytes and microglia.

1.1.7.4. GPER involvement in health and disease

Investigations in the past ten years concluded that GPER/GPR30 is an important player in mammalian physiology and pathology (Prossnitz and Barton, 2011). GPER activation promotes neuronal survival following brain ischemia (Lebesgue et al., 2009;

Lebesgue et al., 2010) and reduces depressive-like behaviors in the mice (Dennis et al.,

2009). In addition, protection against functional and histological manifestations of experimental autoimmune encephalomyelitis (mouse model of multiple sclerosis) has been reported in response to GPER stimulation (Blasko et al., 2009; Wang et al., 2009). 12 GPER mediates the reduction of ischemia-reperfusion injury after myocardial infarction, and of the infarct size, improving the contractile function (Deschamps and Murphy, 2009;

Patel et al., 2009; Bopassa et al., 2010; Deschamps et al., 2011). Moreover, GPER activation causes vasodilatation in human, porcine and rodent arteries (Haas et al., 2009;

Broughton et al., 2010; Lindsey et al., 2011; Meyer et al., 2011), lowering in normotensive (Haas et al., 2009) and hypertensive rats (Lindsey et al., 2009; Lindsey et al., 2011). GPER is implicated in proteinuria reduction and creatinine clearance improvement in hypertensive rats despite continued hypertension (Gilliam-Davis et al.,

2010). Furthermore, GPER activation results in anti-inflammatory properties in pancreatic islets (Balhuizen et al., 2010) and protection of pancreatic islet survival (Liu and Mauvais-Jarvis, 2009); mice deficient in GPER develop insulin resistance and obesity in a sex-dependent manner (Haas et al., 2009; Martensson et al., 2009); moreover, estrogenic protection on survival of pancreatic β-cells is absent in GPER-deficient animals (Liu et al., 2009).

Nevertheless, detrimental effects of GPER activation have also been reported, such as cancer cell proliferation, with poor outcome of GPER-positive breast cancer, ovarian carcinoma, endometrial cancer and uterine carcinosarcoma (Prossnitz and Barton 2011).

Moreover, peripheral pain sensitization in response to GPER stimulation has also been reported (Kuhn et al., 2008; Lu et al., 2009).

13 1.2. Pain

According to the International Association for the Study of Pain, pain is an unpleasant sensory and emotional experience associated with actual or potential tissue damage, or described in terms of such damage (Merskey and Watson, 1979). Pain can manifest itself in three ways:

• as a physiological protective mechanism – an alarm system that notifies the body

of the presence of noxious stimuli in the environment;

• as a repair promoter – once tissue damage has occurred, the hypersensitivity of

the injured part (inflammatory pain) leads to minimization of contact or

movement until healing;

• as neuropathic pain – disturbances in neural and non-neural cells lead to

maladaptive changes in neurons of the sensory system, resulting in spontaneous,

persistent pain and severe hypersensitivity (Basbaum and Woolf, 1999).

14 Fig. 4. Pain pathways. A, Ascending pathways for pain, including receptors and primary afferent neurons, ascending spinothalamic tract and thalamocortical pathways to somatosensory cortex. B, Descending pain modulatory pathways – this system originates in the cerebral cortical areas, including amygdala, and projects to central gray. The latter, in turn, projects to cells in the rostroventral medulla, which in turn project to the dorsal horn; both inhibitory and facilitatory effects are exerted at the dorsal horn level (i.e. the control of nociceptive transmission is bi-directional).

Painful peripheral stimuli activate nociceptive neurons (), initiating a train of action potentials that propagate along the axons of primary afferent nerve fibers to nerve terminals embedded in laminae I and II of the spinal cord dorsal horn. These terminals release pro-nociceptive neurotransmitters (e.g. glutamate, , calcitonin gene related peptide), which further stimulate postsynaptic receptors on the 15 superficial dorsal horn (SDH) neurons. SDH neurons are part of the spinothalamic tract and project to the thalamus; thus, pain is perceived (Krarup, 2003). However, there are also descending pathways from the cortex to the spinal cord that modulate pain responses

(Fig. 4) (Porreca et al., 2002).

1.2.1. Spinal cord involvement in pain

The spinal cord is the central concourse along which all pain messages travel to and from the brain. The dorsal horn is the major site of termination for primary afferent neurons, which are distributed according to fiber size and sensory modality. Nociceptors are small-diameter afferents, with either finely myelinated (A δ, carrying fast, localized pain sensation) or unmyelinated (C, conveying slow, poorly localized pain) fibers and usually synapse with secondary neurons and interneurons in laminae I and II of the SDH.

Neurons in these laminae further project to several brain sites, including the thalamus, periaqueductal gray matter (PAG), lateral parabrachial area, nucleus tractus solitarius and medullary reticular formation; these projections form the ascending pain pathway. In addition, these neurons also interact with GABAergic and glycinergic interneurons in laminae II and III, acting as local modulators to inhibit excessive nociceptive signals

(Todd, 2002; Thomas Cheng, 2010).

Although nociceptors are rather accessible as a target for analgesic therapy, some clinical persistent pain conditions, triggered by intense tissue or nerve injury, result in a reorganized spinal cord dorsal horn; in this situation, pain cannot be treated by blocking messages from the primary sensory neurons.

16 Fig. 5. Hyperexcitability of dorsal horn neurons results in neuropathic pain. Modified from (Inoue et al., 2007).

Changes in pain transmission (i.e. “central sensitization”) include induction of new genes, sprouting of primary sensory neurons, upregulation of neurotransmitters and their receptors and second messenger systems – these cause a prolonged increase in the excitability of dorsal horn neurons (Basbaum and Woolf, 1999). Central sensitization manifests itself as allodynia (pain in response to previously non-painful stimuli) or hyperalgesia (exacerbated response to normally painful stimuli). The plastic changes at dorsal horn synapses that result in central sensitization are comparable to those

17 underlying learning and memory-related long term potentiation (LTP) of hippocampal synapses (Sharif-Naeini and Basbaum, 2011).

1.2.2. Supraspinal sites of pain modulation

It has been known since 1954 that stimulation of the reticular formation, the cerebellum or the cerebral cortex has a controlling influence on the flow of nociceptive impulses up the anterolateral tract (Hagbarth and Kerr, 1954). Further investigations revealed that the firing of dorsal horn neurons in response to noxious stimuli can be inhibited by electrical stimulation of a series of brain regions such as the PAG and lateral reticular formation in the midbrain, the raphe nuclei, the locus coeruleus and various regions of the medullary reticular formation, as well as sites in the hypothalamus, septum, and sensorimotor cortex (Gebhart et al., 1984; Zimmermann, 1984).

The chemical mediators of the descending inhibitory pathways from the brain to the

SDH neurons include endogenous , norepinephrine and serotonin, which are expressed in cell bodies located in the PAG, locus coeruleus and nucleus raphe magnus, respectively (Thomas Cheng, 2010).

In patch clamp experiments, analgesic substances, such as opioids, produce an outward current or hyperpolarization in a majority of caudal PAG output neurons

(Osborne et al., 1996) and opioid-induced analgesia has been supported by findings that opioids directly hyperpolarize subpopulations of neurons predominantly located in the lateral PAG (Behbehani et al., 1990a; Chieng and Christie, 1994).

18 Fig. 6. Neuronal connections in the descending pain pathways from the PAG to the superficial laminae of the spinal cord. (-), inhibitory connection; (+), excitatory connection; PAG, periaqueductal gray; RVM, rostroventral medulla.

Moreover, GABA A receptor antagonists, such as bicuculline methiodide, induce membrane depolarization, increase firing frequency and increase frequency of excitatory postsynaptic potentials of neurons located in the PAG, in patch clamp experiments performed on acute midbrain slices (Behbehani et al., 1990b); this is particularly important since opioid-containing terminals in the PAG are hypothesized to make contact with GABAergic inhibitory interneurons that tonically inhibit descending projection neurons from the PAG to the nucleus raphe magnus (Reichling et al., 1988), and reduces GABA release in the lateral PAG of the rat (Renno et al., 1992). Thus, 19 opioids inhibit lateral PAG GABAergic neurons, allowing excitatory ventrolateral PAG neurons to excite “OFF-cells” and inhibit “ON-cells” located in the rostral ventral medulla (Fig. 6), leading to attenuation of pain (Fields et al., 1991; Morgan et al., 1992;

Osborne et al., 1996).

1.2.3. Neuronal pathways in pain: calcium, depolarization, ROS

Upon activation of nociceptors, their central terminals release glutamate or substance

P; glutamate is an activator of ionotropic (NMDA and AMPA), as well as metabotropic

(mGluRs) receptors, whereas substance P binds the phospholipase C (PLC)-coupled neurokinin 1 (NK1) receptor, thus transmitting inflammatory signals and pain. Continued activation of primary afferent pain fibers elicits an increase in intracellular Ca 2+ , which is the trigger for central sensitization. The rise in cytosolic Ca 2+ can occur through the opening of NMDA receptor-channels by depolarization-induced removal of their magnesium block; nevertheless, Ca 2+ influx may result via voltage-dependent calcium channels, secondary to the depolarization that occurs after inhibition of a potassium current and/or activation of a nonselective cation current (Sharif-Naeini and Basbaum,

2011).

2+ 2+ Thus, the increase in intracellular Ca concentration, [Ca ]i, is a common signature of pain mediators and a key step which leads to the activation of several calcium- dependent kinases (e.g. calcium-calmodulin kinase II α - CamKII α), as well as cyclooxygenase-2 (COX-2) and the NO synthase (NOS) family, that are responsible for the production of pain-sensitizing molecules (Kuner, 2010). Additionally, at supraspinal sites, such as thalamo-cortical neurons, concurrent regulation of T-type and L-type

20 calcium currents results in reduced visceral pain-responses, proving their importance in the gating of incoming nociceptive inputs (Cheong et al., 2008).

Reactive oxygen species (ROS) contribute to neuropathic pain by reducing spinal

GABA release (Yowtak et al., 2011) and are involved in the potentiation of synaptic excitability in the spinal cord dorsal horn (Fig. 7A, B) (Lee et al., 2009), that is a mechanism for central nociceptive sensitization (Ji and Woolf, 2001). In addition, the increase in cytosolic free calcium is followed by downstream mitochondrial calcium uptake, with subsequent production of ROS (Fig. 7C), resulting in underlying chronic pain (Kim et al., 2011). Furthermore, at the supraspinal level, reactive oxygen and nitrogen species are incriminated for descending facilitation of nociception

(Salvemini et al., 2011) and antinociceptive tolerance (Doyle et al., 2010). In amygdala neurons, increased excitability is mediated by mitochondrial reactive oxygen species, and results in pain behavior (Li et al., 2011).

21 Fig.7. Reactive oxygen and nitrogen species involvement in pain sensitization. A, NADPH oxidase- and mitochondrial-derived superoxide oxidizes NO to peroxynitrite (ONOO -), which further enhances its own production via modulation of NADPH oxidase and protein kinase C – feedforward mechanisms involved in central sensitization. B, Peroxynitrite enhances glutamatergic signaling through: (i) nitration and activation of the NMDARs and the protein kinases involved in NMDARs activation; (ii) nitration and inactivation of the glutamate transporters (GLT-1, GLAST, EAAC1) and of the glutamine synthetase (GS). Thus, toxic levels of glutamate accumulate in the synapse. Moreover, compromised cysteine (Cys) transport due to EAAC1 inhibition results in low levels of glutathione (GSH) and increased neuronal nitroxidative stress (Salvemini et al., 2011). C, Upon cytosolic Ca 2+ increase, most of the Ca 2+ is sequestered by mitochondria; this consequently increases ROS production, leading to activation of intracellular signaling cascades (i.e. PKC, PKA, ERK), which in turn results in synaptic plasticity of the dorsal horn neurons (Kim et al., 2011).

22 1.2.4. Animal models of nociception

Rather than biochemical indicators or electrophysiological parameters, the most reliable signs of pain, in both humans and animals, are behavioral reactions to algogenic stimuli; however, these are not always very specific either (Le Bars et al., 2001). In animals, pain may be defined as an aversive sensory experience caused by actual or potential injury that elicits progressive motor and vegetative reactions, results in learned avoidance behavior and may modify species specific behavior, including social behavior

(Zimmermann, 1986). Depending on the animal pain model, the nociceptive response can be evaluated from the following parameters: (i) the pain perception threshold – the point at which the stimulus is perceived by animals as painful; (ii) the latency – the maximum duration of painful stimulus that can be endured without manifestation of pain-related behavior; (iii) the total number of (or time spent engaged in) pain-related behaviors after application of an algogenic stimulus (Besson and Besson, 1997; Le Bars et al., 2001).

1.2.4.1. Spinal pain: caudally directed biting and scratching in response to spinal delivery of nociceptive molecules

Intrathecal administration of pro-nociceptive mediators, such as: excitatory amino acids, substance P, , spermine, D-cycloserine or sulfur containing amino-acids results in a specific behavior consisting of scratching, biting and licking directed caudally, to the tail and hind limbs (Hylden and Wilcox, 1981; Aanonsen and Wilcox,

1986; Aanonsen and Wilcox, 1987; Sakurada et al., 1999; Sakurada et al., 2000; Tan-No et al., 2000; Osborne and Coderre, 2003; Tan-No et al., 2007). This type of behavior is

23 reduced or fully prevented by systemic, intrathecal or i.c.v. morphine, thus, it has been accepted as an indicator of nociception.

1.2.4.2. Supraspinal pain: hot plate test

In the hot plate test, rats placed on a plate heated to a constant temperature present two behavioral components: paw licking and jumping; both are considered to be supraspinally integrated responses and are measured in terms of their reaction times (Le

Bars et al., 2001).

1.3. Estrogens and pain

Sex-hormone dependent differences in pain and analgesia have been documented both in human and animal studies, with females presenting more severe, frequent, anatomically diffuse and longer lasting pain than males (Unruh, 1996; Cairns and

Gazerani, 2009), suggesting an emerging role for estrogens and androgens in this process

(Craft et al., 2004; Greenspan et al., 2007; Fillingim et al., 2009). Estrogen modulation of pain is very complex, with either pro- or antinociceptive effects depending on the type of pain disorders and on the system of the body via which they exert their action (Craft,

2007).

While systemic estrogens appear to decrease acute inflammatory pain in rat animal models (Multon et al., 2005; Mannino et al., 2007), central (i.c.v.) administration of estradiol augments formalin-induced nociception (Aloisi and Ceccarelli, 2000; Ceccarelli et al., 2004). Moreover, estradiol has been reported to affect neuronal excitability leading to amplification of nociceptive responses to excitotoxic spinal cord injury in male rats

24 (Gorman et al., 2001) and also to enhance stress-induced facilitation of pain (Bradesi et al., 2003). Using a radiant heat source, Bradshaw and colleagues (2000) observed an increase in thermal hyperalgesia during proestrus in female rats when compared to males, ovariectomized females or females in diestrus or estrus. As estrogen levels reach a peak during proestrus, this study further suggests a pro-nociceptive effect (Bradshaw et al.,

2000).

The mechanisms by which estrogens regulate nociception are complex, including direct action on the central and peripheral nervous system, as well as indirect actions via their modulation of skeletal and immune system (Craft, 2007).

1.3.1. Estrogens and spinal pain

In the mammalian spinal dorsal horn, ER α and/or ER β are located mainly in the lamina II and to some extent I, III, V and X (Evrard, 2006), while GPER is present in laminae I, II and V (Dun et al., 2009; Liu et al., 2011). E2 can reach central levels upon uptake from circulation, but is also synthesized locally by spinal aromatase, which converts testosterone to estradiol; aromatase is present in spinal cord laminae I and II, areas involved in nociception and opioid antinociception (Evrard and Balthazart, 2004;

Evrard, 2006; Liu et al., 2011). Spinal delivery of E2 or testosterone rapidly decreases foot withdrawal latency in male quail hot water immersion test; treatment with an aromatase inhibitor abolishes the effect of testosterone, but not of E2, supporting E2 as a pro-nociceptive mediator in the spinal cord (Evrard and Balthazart, 2004). Moreover, high E2 concentrations are able to inhibit glycine receptor-mediated current in spinal neurons, thus increasing their excitability (Jiang et al., 2009). Furthermore, intrathecal

25 administration of E2 increases colorectal distension-induced nociceptive responses in the rat (Ji et al., 2011).

1.3.2. Estrogens and supraspinal pain

Interactions of estrogens and androgens with the supraspinal opioid system have been reported in several studies, suggesting a pro-nociceptive estrogenic modulation at the supraspinal level. There is decreased antinociception in females versus males to intra-

PAG morphine, as well as decreased analgesia in high-estrogen state females versus low- estrogen state females (Bernal et al., 2007). Moreover, male rodents display greater analgesia following µ-opioids administration into the lateral ventricle, the periaqueductal gray (PAG) or the rostro-ventral medulla than female rodents (Kepler et al., 1989; Kepler et al., 1991; Boyer et al., 1998; Krzanowska and Bodnar, 1999; Krzanowska and Bodnar,

2000; Loyd and Murphy, 2006). Estrogen decreases the potency of systemic and ventricular morphine analgesia on visceral pain tests in ovariectomized female rats (Ji et al., 2007) and increases pain responses to visceral stimulation following implantation on the amygdala in the rats (Myers et al., 2011).

1.3.3. GPER and pain

In the spinal cord, immunohistochemical analysis revealed GPER positively labeled cells in the superficial layers of the dorsal horn of both male and female rats (Dun et al.,

2009). The outer layer of the spinal dorsal horn had a dense accumulation of GPER-ir which was distributed throughout the cervical to sacral levels in both sexes (Takanami et al., 2009). These findings suggest that GPER is a possible site of estrogen mediated

26 central nociceptive processing observed during the high estrogen phase of the estrous cycle (Okamoto et al., 2003).

Moreover, trigeminal ganglion cultures derived from female rats express functional

GPER within the nociceptive, TRPV1 positive subpopulation of sensory neurons

(Fehrenbacher et al., 2009). In ovariectomized females, the population of GPER positive neurons extending small, unmyelinated axons, involved in the transmission of painful stimuli, was double that of GPER neurons with myelinated fibers (Liverman et al., 2009).

This further supports a putative role of GPER in estrogenic modulation of pain.

The receptor expressed in these systems is functional and activated by G-1 in vitro . In trigeminal ganglion cultures derived from female rats, G-1 rapidly and significantly triggers increases in intracellular calcium concentration (Fehrenbacher et al., 2009), supporting the hypothesis that GPER activation increases the activity of peripheral nociceptors. Furthermore, brief (15 min) pretreatment with estradiol of trigeminal ganglion neurons derived from ovariectomized female rats resulted in a potentiation of the cellular responses to the inflammatory mediators bradykinin (BK) and prostaglandin

E2 (PGE 2); these effects were reinstated with membrane impermeable 17- β-estradiol conjugated to bovine serum albumin, suggesting a non-genomic mechanism of action

(Fehrenbacher et al., 2009).

The first in vivo study describing estrogenic activation of GPER to induce mechanical hyperalgesia was published by Kuhn et al. (2008). The group had previously reported rapid stimulatory effects of estrogen on nociceptive neurons, as well as induction of hyperalgesia (Hucho et al., 2006). Kuhn et al. (2008) found that in male rats, estradiol, and the selective GPER agonist G-1 (Bologa et al., 2006) and ICI 182,780 (fulvestrant) –

27 that acts as GPER agonist and ER α and ER β antagonist (Wakeling et al., 1991; Filardo et al., 2002) – rapidly induce protein kinase C ε (PKC ε) activation and translocation to the membrane in vitro , mainly in isolectin B4 positive nonpeptidergic neurons in the dorsal root ganglia. In vivo , local GPER activation with G-1 or ICI 182,780 injected in the hind paw elicits PKC ε - dependent mechanical hyperalgesia (Kuhn et al., 2008). This sensitization is reversed by pretreatment with εV1-2, a PKC ε inhibitor (Kuhn et al.,

2008). Downstream of GPER, PKC ε phosphorylates the ion channel transient receptor potential vanilloid subclass 1 (TRPV1), which hinders tubulin binding to TRPV1, leading to microtubule destabilization and retraction of microtubules from filopodial structures, thus to a microtubule-dependent pain sensitization (Goswami et al., 2011).

Additionally, another in vivo study reported pronociceptive effects of GPER activation in a rat model of visceral hypersensitivity (Lu et al., 2009). While overiectomy resulted in a loss of visceral hypersensitivity to serotonin during colorectal distension, estradiol, G-1 and estradiol + ICI 182,780, but not PPT (ER α selective agonist) or DPN

(ER β selective agonist) treatment restored the visceral motor response. Moreover, knockdown of GPER using specific antisense oligodinucleotides suppressed serotonin- induced visceral hypersensitivity, supporting the hypothesis that some pro-nociceptive estrogenic effects result from GPER activation.

1.4. Specific aims

Immunohistochemical studies in rodents support the existence of GPER in pain- relevant areas of the nervous system such as dorsal root ganglia, spinal cord superficial laminae, periaqueductal gray, amygdala, trigeminal sensory nucleus and thalamus (Dun 28 et al., 2009; Hazell et al., 2009; Takanami et al., 2009). Several studies suggest that peripheral GPER activation is pro-nociceptive (Kuhn et al., 2008; Lu et al., 2009) and

2+ mechanisms may involve an increase in [Ca ]i (Fehrenbacher et al., 2009) or PKC ε translocation to membrane and activation of TRPV1 channel (Kuhn et al., 2008;

Goswami et al., 2011). Nevertheless, GPER involvement in pain processing at central levels has not been explored. Preliminary results show that intrathecal challenge of mice with GPER agonist G-1 increases spinal phosphorylated ERK1/2 levels (Fig. 8), which is a sign of nociceptive activation (Ji and Woolf, 2001).

Fig. 8. Effect of intrathecal G-1 on ERK1/2 phosphorylation. A, Representative blots illustrating expression of phosphorylated (p) and total (t) ERK1/2 (42/44 kDa) in the mouse lumbar spinal cord upon intrathecal administration of either vehicle (negative control), G-1 or DHPG (mGluR1/5 agonist, positive control, (Hu et al., 2007)). B, Similar to DHPG, intrathecal G-1 significantly increased pERK1/2 levels in the mouse lumbar spinal cord (n = 3 for each treatment group, *P < 0.05). Statistical analysis for the G-1 treatment: df = 7, F(1,7) = 9.68, P = 0.017; for the DHPG treatment: df = 5, F(1,5) = 16.33, P = 0.0099.

Therefore, the major goal of this research is to investigate whether GPER is involved in pain modulation at spinal and supraspinal sites; an additional aim is to explore the underlying mechanisms. The working hypothesis is illustrated in Fig. 9: GPER activation 29 2+ produces an increase in neuronal [Ca ]i and cytosolic and mitochondrial ROS levels via disparate or sequential mechanisms, which results in membrane depolarization underlying pain facilitation in the PAG or pain sensitization in spinal cord. Neuronal

2+ membrane depolarization may further enhance the rise in [Ca ]i via voltage-gated calcium channels, providing a feed-forward mechanism for the pro-nociceptive effect.

Fig. 9. Putative mechanisms of pro-nociceptive effects of GPER activation in the spinal cord and periaqueductal gray (PAG).

The proposed research uses behavioral assays, reverse transcription polymerase chain reaction, immunohistochemistry, electrophysiology and optical imaging with fluorescent dyes. The specific aims are as follows:

30 Specific aim 1

GPER activation at the spinal and supraspinal level results in pro-nociceptive behaviors

1.1. To evaluate whether intrathecal administration of the GPER selective agonist G-1

(Bologa et al., 2006) elicits pain-related behaviors consisting of caudally directed biting, scratching and licking in mice and test whether the GPER antagonist G15 (Dennis et al.,

2009) prevents this effect.

1.2. To assess that the behavior elicited by intrathecal injection of GPER agonists is pain- related: evaluate whether morphine inhibits caudally directed biting and scratching elicited by G-1.

1.3. To test whether intra-periaqueductal gray delivery of G-1 decreases paw-withdrawal latency in the rat using the hot plate test and whether GPER antagonism with G-15 diminishes or abolishes this effect.

Sub-aim 1: GPER is present in the mouse spinal cord a. To identify GPER mRNA in various segments of the mouse spinal cord. b. To identify GPER-immunoreactive neurons in the dorsal horn of the spinal cord.

31 Specific aim 2

Mechanisms of GPER-mediated pro-nociceptive effects at spinal and supraspinal sites

2.1. To determine the effects of GPER activation or blockade on the membrane potential of the neurons located in the superficial dorsal horn of the spinal cord and of the PAG neurons using whole cell patch clamp recording in rat lumbar spinal cord and midbrain slices.

2.1.1. To test whether G-1 depolarizes superficial dorsal horn neurons and whether

G15 can antagonize the effect of G-1.

2.1.2. To test the effect of G-1 on PAG neurons in the midbrain slice and how this response is altered by G15.

2.2. To examine the effects of GPER activation or blockade on intracellular Ca 2+ concentration and cytosolic and mitochondrial ROS accumulation in cultured spinal and

PAG neurons.

2.2.1 . To determine the calcium response of cultured spinal neurons and of cultured

PAG neurons upon G-1 application and the effect of G-15 on this response.

2.2.2. To evaluate how G-1 or G-1 + G15 application affects the cytosolic and mitochondrial ROS levels in cultured spinal and PAG neurons.

32 CHAPTER TWO

EXPERIMENTAL PROCEDURES

2.1. Drugs

The GPER agonist G-1 was purchased from Calbiochem, EMD Biosciences

(Rockland, MA), morphine was obtained from the National Institute of Drug Abuse and the GPER antagonist G15 was synthesized and provided by Dr. Jeffrey B. Arterburn. For behavioral studies G-1 and G15 stock solutions of 1 mg/mL in ethanol were diluted to the desired concentration using a vehicle containing 0.9% NaCl, 20% ethanol and 2% Tween

20. Morphine was dissolved in saline (0.9% NaCl) solution.

2.2. Experimental animals

Spinal nociception, immunohistochemistry and RT-PCR : Experiments were performed on male ICR mice (Ace Animal Inc., Boyertown, PA) weighing 20-25 g.

Animals were maintained at room temperature (22-25 ºC), with free access to tap water and standard food, under a 12/12-h light/dark cycle (lights on at 8:00 h) and housed 5 animals/cage.

Hot plate test : Experiments were performed on Sprague-Dawley male rats (Ace

Animals, Inc., Boyertown, PA), weighing 300-350 g. Animals were housed in groups of

2-3 for at least one week in an animal room maintained at room temperature (22-25 ºC) and constant humidity (55 ± 5%), on a 12/12-h light/dark cycle (lights on at 7:00), with free access to food and water. All the experiments were conducted between 9:00 am and

3:00 pm.

33 Whole-cell patch clamp experiments : Sprague-Dawley rats 5-12 days old were used.

Neuronal culture : Sprague-Dawley rats 0-2 days old were used.

Comment on the use of male animals in behavioral studies: In vivo studies of GPER physiology have been carried out in both male and female animals suggesting its importance in both sexes. The levels of estrogen activating GPR30 are in the nanomolar

(3 – 7 nM) range (Revankar et al., 2005; Thomas et al., 2005), whereas plasma estrogen levels are typically in the picomolar range. Thus, GPER is activated when estrogen synthesis/secretion is increased locally. All of our studies are performed in male animals, raising the question whether this is relevant at all for GPER physiology. GPER expression levels appear similar in male and female animals (Brailoiu et al., 2007; Dun et al., 2009) and estrogen has important physiological functions in the male (de Ronde et al.,

2003). In addition, aromatase enzyme converting testosterone into estrogen is present in the male, its knockout resulting in reduced fertility, osteoporosis and increased intra- abdominal adiposity (Murata et al., 2002). Thus, estrogen can regulate similar functions in male and female. Moreover, it has been suggested that high local estradiol concentrations can be rapidly obtained in the male spinal cord due to large available pool of testosterone and this has been shown to be relevant for estrogenic regulation of nociception (Evrard and Balthazart, 2004; Evrard, 2006). In conclusion, the result of this study will provide new insight relative to the physiological role of GPER in pain modulation.

34 2.3. Mouse GPER mRNA analysis by RT-PCR

Adult male ICR mice weighing 20-25 g were rapidly decapitated and the spinal cord removed surgically and sectioned in segments according to location (cervical, thoracic, lumbar and sacral); all tissues were placed in ice-cold Trizol® reagent and immediately frozen (-70 ºC).

Total RNA from various tissues and brain regions were isolated using Trizol® reagent according to the manufacturer’s protocol (Invitrogen). The purity and concentration of RNA was assessed by spectrophotometrical analysis. First-strand cDNA was synthesized from 2 µg of total RNA with random hexamer primers and the

SuperScript III reverse transcriptase according to the manufacturer’s protocol

(Invitrogen). Two µL of 20 µL cDNA product were amplified with the following specific intron-spanning primers based on GPER (NM_029771) and β-actin (NM_007393) sequences: mouse GPER: 5’-TCA GCA GTA CGT GAT TGC CCT CTT-3’ (upstream),

5’-ACT GCT CTG TGC TGT CTG GAA TGA-3’ (downstream); mouse β-actin: 5’-TCG

TAC CAC AGG CAT TGT GAT GGA-3’ (upstream), 5’-ACT CCT GCT TGC TGA

TCC ACA TCT-3’ (downstream). Amplifications were performed using Platinum Taq

DNA polymerase (Invitrogen) as follows: GPER: 32 cycles of 94 ºC for 60 s, 57 º C for 60 s, 72 ºC for 60 s; β-actin: 29 cycles of 94 ºC for 30 s, 55 ºC for 45 s and 72 ºC for 60 s.

The PCR products were separated using agarose gel electrophoresis on 1% agarose gels and visualized by an ethidium bromide staining under UV light.

35 2.4. Immunohistochemistry

Adult male ICR mice, weighing 20-25 g were used. Mice were anesthetized with urethane (1.2 g/kg, i.p.) and intracardially perfused with 0.1 M phosphate buffered saline

(PBS), followed by 4% paraformaldehyde/0.2% picric acid in PBS. Lumbar spinal cord was removed, post fixed for 2h and stored in 30% sucrose/PBS solution overnight. Spinal cord was sectioned to 50 µm using a Vibratome. Tissues were processed for GPER- immunoreactivity by the avidin-biotin complex procedure or fluorescent method

(Brailoiu et al., 2007). Tissues were first treated with 3% hydrogen peroxide to quench endogenous peroxidase, washed several times, blocked with 10% normal goat serum and incubated in GPER antiserum. After thorough rinsing, sections were incubated in biotinylated anti-rabbit IgG (1:150 dilution, Vector Laboratories, Burlingame, CA, USA) for 2 h, rinsed with PBS and incubated in avidin-biotin complex solution for 1 h (1:100 dilution, Vector Laboratories). Following washes in Tris-buffered saline (TBS), sections were incubated in 0.05% diaminobenzidine/0.001% H 2O2 solution and washed for at least

2 h with TBS. Sections were mounted on slides with 0.25% gel , air-dried, dehydrated with absolute alcohol followed by xylene, and coverslipped with Permount.

Tissues processed with pre-immune serum instead of GPER antiserum served as a negative control (Brailoiu et al., 2007; Dun et al., 2009). For fluorescent immunohistochemistry, incubation with GPER antiserum was followed by several washes with PBS, then by incubation with anti-rabbit IgG (1:50, Vector Laboratories), washes with PBS and incubation with avidin-fluorescein isothiocyanate (FITC). After rinsing with PBS, spinal cord sections were mounted in Citifluor, coverslipped and

36 examined under a confocal scanning laser microscope (Leica TCS SL) with excitation/emission wavelengths set to 488/520 for FITC.

2.5. Intrathecal injection in mice

The i.t. injection procedure was according to the method described by Fairbanks

(Fairbanks, 2003): a 30-gauge stainless steel needle attached to a 50 µL Hamilton microsyringe is inserted between L5 and L6 of an unanaesthetized mouse and drugs given in a volume of 5 µL over the course of 1-2 s. Puncture of the dura was indicated by a reflexive lateral flick of the tail or the formation of an S shape by the tail. Following the injection, the syringe was rotated slightly and removed.

2.6. Behavioral assay of spinal nociception

Mice were acclimated to the laboratory for at least two days before testing and used only once throughout experiments. Approximately 60 min before injections, the mice were adapted to an individual box which was also used as the observation chamber after the intrathecal (challenging) injection.

Each mouse received a subcutaneous injection of either vehicle (saline with 20% ethanol and 2% Tween 20) or chemical (G-15, morphine), followed by an intrathecal injection of G-1 30 min later. Immediately after the i.t. injection, the mouse was placed in the transparent observation box and the accumulated response time of hind limb scratching directed toward the flank, biting and/or licking of the hind paw and the tail was measured for 10 min. 6-10 mice were assigned for each treatment group.

37 2.7. Intra-PAG cannulation of rats

Rats were anesthetized with a mixture of hydrochloride (100-150 mg/kg) and acepromazine maleate (0.2 mg/kg). A sterilized stainless steel C313G guide cannula

(22 gauge, Plastics One Inc., Roanoke, VA) was implanted in the PAG and fixed with dental cement. The stereotaxic coordinates were as follows: 7.8 mm posterior to bregma,

0.5 mm from midline and 5 mm ventral to the dura mater (Paxinos and Watson, 1998). A

C313DC cannula dummy (Plastics One Inc., Roanoke, VA) of identical length was inserted into the guide tube to prevent its occlusion and the animals were housed individually after surgery. Experiments began one week postoperatively. Each rat was used only once. At the end of the behavioral experiments, each rat was injected with 0.5

µl of cresyl violet, anesthetized and perfused transcardially with 0.9% isotonic saline followed by 4% paraformaldehyde in PBS (pH 7.4). The brain was removed, stored in the same fixative for 4 h, then in 20% sucrose overnight, and cut into 20 µm sections on a freezing microtome. Each coronal section was mounted according to standard histological procedures (Benamar et al., 2004), and the site of the injection was verified by locating the dye.

2.8. Hot plate test

A 52 ºC hot plate (Ugo Basile, Varese, Italy) was used, according to the method of

Dewey et al. (Dewey et al., 1970). The baseline response latency was obtained for each animal after two conditioning runs. Each rat was retested on the hot plate at +10 min and thereafter at 10 min intervals up to +30 min by using jumping or hind-paw licking as the

38 nociceptive endpoint and 30 s as the cutoff point. 4-7 rats were assigned for each treatment group.

The percentage of maximal possible effect (%MPE) for each animal at each time point was calculated using the formula:

test latency - baseline latency %MPE = x 100 30 - baseline latency

Equation 1. Calculation of % maximal possible effect in the hot plate test.

2.9. Electrophysiology

Visualized whole-cell patch clamp was performed on spinal cord and midbrain slices from Sprague-Dawley rats 5-12 days old. Animals were decapitated and the spinal cord or midbrain rapidly removed surgically and placed in an ice-cold high-sucrose artificial cerebrospinal fluid (ACSF) containing (in mM): 248 sucrose, 11 glucose, 23 NaHCO 3, 2

KCl, 1.25 KH 2PO 4, 2 CaCl 2, 2 MgSO 4, saturated and continuously bubbled with 95% O 2

– 5% CO 2 (Breton et al., 2009). Pia membranes were carefully removed with a pair of fine forceps and the lumbar segment of the spinal cord or the midbrain fixed onto an agar block with cyanoacrylic glue. Transverse slices (250-300 µm thick) were cut in the slicing chamber of a Vibratome (series 1000), using a sapphire blade and stored at room temperature (21 ± 0.5 ºC) into regular ACSF containing (in mM): 126 NaCl, 26 NaHCO 3,

1.25 NaH 2PO 4, 2.5 KCl, 2 CaCl 2, 2 MgCl 2, 10 glucose, pH 7.35 (adjusted with NaOH).

Spinal cord or midbrain slices were incubated at least one hour before the start of experiments. One slice was transferred to the recording chamber (RC-22, Harvard

39 Apparatus), immobilized by the grid of a fine nylon mesh and superfused continuously with oxygenated ACSF at a mean flow rate of 3 mL per minute. Recording experiments were performed at room temperature (spinal cord) or at 32-34 ºC (midbrain) (Chen et al.,

2011).

Patch pipettes were pulled from thin-walled fiber-filled borosilicate glass (WPI, OD

2.0 mm), filled with an intracellular recording solution containing (in mM): 130 K + gluconate, 1 MgCl 2, 2 CaCl 2, 4 ATP, 10 EGTA, 10 HEPES, pH 7.3 (adjusted with KOH) and only the ones with appropriate resistance (3-7 MΩ) were used. Cells were visualized using a Nikon (Eclipse FN1) upright microscope and the ones located in the superficial laminae (spinal cord dorsal horn) or PAG (midbrain) were approached by the patch electrode under positive pressure. Once the patch electrode was in contact with the cell, the positive pressure was released and a giga-ohm seal established by applying light suction to the patch electrode. Recordings were made using a CV-4 headstage (Molecular

Devices) attached to an Axopatch 1D amplifier (Axon Instruments/Molecular Devices).

After the seal, the cell was voltage clamped at -60 mV and the membrane ruptured under negative pressure, resulting in a whole-cell configuration of the spinal superficial dorsal horn neuron or PAG neuron. The patched cell was studied under current clamp mode (I =

0 pA). Pharmacological compounds were administered by bath superfusion at a rate of 3 mL/min and the data was digitized and stored for later analysis using a Digidata 1320A analog-digital converter (Axon Instruments/Molecular Devices) and analyzed using the pClamp 10.2 software (Molecular Devices).

40 2.10. Neuronal culture

Spinal cord or periaqueductal gray neurons were dissociated from neonatal (1-2 day old) Sprague Dawley rats as previously described (Brailoiu et al., 2005; Dun et al., 2009).

Newborn rats were decapitated and the spinal cord and quickly removed surgically and immersed in ice-cold Hanks balanced salt solution (HBSS) without calcium and magnesium (Mediatech, Herndon, VA). The PAG was identified (Paxinos and Watson

1998) and carefully removed with a forceps. Tissues were minced and subjected to enzymatic with papain for 3-4 min at 37 ºC, followed by mechanical trituration in total medium to stop enzymatic activity. The total medium was composed of:

Neurobasal-A (Invitrogen, Carlsbad, CA) containing 1% GlutaMax (Invitrogen), 2% penicillin-streptomycin-amphotericin B solution (Mediatech, Manassas, VA) and 10% fetal bovine serum. After centrifugation at 500 g, fractions enriched in neurons were collected and resuspended in total medium. Cells were plated on round 25 mm glass coverslips coated with poly-D-lysine (Sigma-Aldrich, St Louis, MO) in 6-well plates.

Cultures were maintained at 37 ºC in a humidified atmosphere with 5% CO 2. The mitotic inhibitor cytosine β-arabino furanoside (1 µM) (Sigma) was added to the culture the third day to inhibit glial cell proliferation (Schoniger et al., 2001). The media was changed every 2-3 days and cells used after 5 days in culture.

2.11. Calcium imaging

2+ Measurements of [Ca ]i were performed as previously done in our lab and described in the literature (Brailoiu et al., 2005; Brailoiu et al., 2011; Deliu et al., 2011b). Cells were incubated with 5 µM fura-2 AM (Invitrogen) in Hanks’ balanced salt solution

41 (HBSS) at room temperature for 45 min, in the dark, washed three times with dye-free

HBSS, and then incubated for another 45 min to allow for complete de-esterification of the dye. Coverslips (25 mm diameter) were subsequently mounted in an open bath chamber (RP-40LP, Warner Instruments, Hamden, CT) on the stage of an inverted microscope Nikon Eclipse TiE (Optical Apparatus Co., Ardmore, PA). The microscope was equipped with a Perfect Focus System and a Photometrics CoolSnap HQ2 CCD camera (Roper Scientific, Optical Apparatus Co.). During the experiments the Perfect

Focus System was activated. Fura-2 AM fluorescence (emission = 510 nm), following alternate excitation at 340 and 380 nm, was acquired at a frequency of 0.25 Hz. Images were acquired and analyzed using NIS-Elements AR 3.1 software (Nikon/Optical

Apparatus Co.). The ratio of the fluorescence signals (340/380 nm) was converted to

Ca 2+ concentrations using the following formula:

R - Rmin [Ca 2+ ] = Kd x Q x Rmax - R

Equation 2. Quantification of intracellular calcium concentration using Fura-2 fluorescent dye (Grynkiewicz et al., 1985). Kd, Ca 2+ dissociation constant of Fura-2; Q, ratio Fmin/Fmax at 380 nm; R, fluorescence intensity ratio F(340 nm)/F(380 nm) for the ion-bound and ion-free indicator, respectively, in the experimental conditions. Rmin, Fmin and Rmax, Fmax correspond to completely ion-free and ion-saturated indicator, respectively and are determined during calibration, along with Kd. In our experimental conditions Kd = 220 nM, Q = 6, Rmin = 0.2, Rmax = 7.5.

Cultured cells with neurites at least three times longer than the cell bodies, which

2+ respond to 20 mM KCl solution by an increase in [Ca ]i were considered neurons

(Brailoiu et al., 2005).

42 2.12. Detection of cytosolic ROS accumulation

Assessment of cytosolic ROS levels was achieved using the carboxy-H2-DCFDA (5-

(and-6)-carboxy-2’,7’-dichlorodihydrofluorescein diacetate) dye (Invitrogen). This assay is based on the principle that the nonpolar, nonionic H 2-DCFDA crosses the cell membrane and is enzymatically hydrolyzed by intracellular esterases to nonfluorescent

H2-DCF. In the presence of ROS, H 2-DCF is rapidly oxidized to become highly fluorescent DCF. Cells were incubated with 1 µM H2-DCFDA and corresponding treatment in HBSS at room temperature for 15 min, in the dark and washed with dye-free

HBSS. Coverslips (25 mm diameter) were subsequently mounted in an open bath chamber (RP-40LP, Warner Instruments) on the stage of an inverted microscope Nikon

Eclipse TiE (Optical Apparatus Co.). The intensity of green fluorescence following excitation at 495 nm was acquired at a frequency of 0.25 Hz and evaluated as a measure of cytosolic ROS accumulation.

2.13. Detection of mitochondrial superoxide accumulation

Measurement of mitochondrial ROS levels was achieved using the MitoSOX Red superoxide indicator (Invitrogen). MitoSOX Red reagent permeates live cells, where it selectively targets the mitochondria. At the mitochondrial level, it is rapidly oxidized by superoxide, but not by other ROS and reactive nitrogen species (RNS); the oxidized product is highly fluorescent upon binding to nucleic acid. Cells were incubated with 3

µM MitoSOX Red in HBSS at room temperature for 25 min (out of which 15 min were with the corresponding treatment), in the dark and washed with dye-free HBSS.

Coverslips (25 mm diameter) were subsequently mounted in an open bath chamber (RP- 43 40LP, Warner Instruments) on the stage of an inverted microscope Nikon Eclipse TiE

(Optical Apparatus Co.). The intensity of red fluorescence after excitation at 510 nm was acquired at a frequency of 0.25 Hz and evaluated as a measure of mitochondrial superoxide accumulation.

2.14. Statistical analysis

The data were expressed as the mean ± standard error of mean (S.E.M.). Statistical analysis of difference between treatment groups was assessed with one-way or two-way analysis of variance (ANOVA) using Origin 7 (OriginLab Corporation, Northampton,

MA). ID 50 values for G15 were determined by linear regression analysis using Origin 7 software. Statistical significance was set at P < 0.05.

2.15. Western blot for ERK1/2 – preliminary results

Five minutes after intrathecal injection of either vehicle, G-1 (0.5 nmol) or DHPG (1

µmol), mice were decapitated and the lumbar portion of the spinal cord removed quickly and immersed in 300 µL ice-cold lysis buffer containing 10 mM Tris, 0.2 M NaCl, 1 mM

EDTA, 1% TritonX (Sigma), 1 mM sodium orthovanadate (AlfaAesar), 100 mM NaF

(Sigma) and protease inhibitor cocktail (Roche), pH 7.4. Samples were homogenized using an electric sonicator and further maintained under constant agitation for 2 h at 4 ºC, then centrifuged for 20 min at 12,000 rpm at 4 ºC in a microcentrifuge. The protein concentration of each sample was determined spectrophotometrically using the BCA assay kit (Pierce Biotechnology, Inc) and bovine serum albumin as a protein standard.

Once protein concentrations were equalized, 2X Laemmli buffer (4% SDS, 10% 2- 44 mercaptoethanol, 20% glycerol, 0.004% bromophenol blue, 0.125 M Tris HCl, pH 6.8) was added to each sample and the samples boiled for 5 min at 95-100 ºC to reduce and denature the proteins. Protein samples were loaded onto 10 % Tris-Glycine gels and further run at 200 V using PowerPAC HC source (BioRad) in a buffer containing 25 mM

Tris base, 250 mM glycine and 0.1% SDS. Proteins from the gel were transferred on nitrocellulose membranes (LI-COR) at 100V using a buffer containing 39 mM glycine,

48 mM Tris base, 20% methanol and 0.037% SDS. After the transfer, membranes were blocked in Tris buffered saline (TBS, 15 mM NaCl, 5 mM Tris base, pH 7.5) with 5% nonfat milk and incubated at 4 ºC, overnight, in primary antibodies (rabbit anti-pERK1/2, rabbit anti-tERK1/2 or mouse anti- β-actin, Cell Signaling) diluted in Li-Cor blocking solution. After three washes with TBST (TBS containing 0.1% Tween 20), 15 minutes each, blots were incubated at room temperature for 1 h in fluorescent secondary antibodies (goat anti-rabbit, goat anti-mouse, LI-COR) diluted in Li-Cor blocking solution. Following another 2 washes with TBST (15 min, 10 min) and one with TBS (5 min), blots were visualized using LI-COR Odyssey. Each membrane was incubated with anti- β-actin antibody and either anti-pERK1/2 or anti-tERK1/2 antibody. Phosphorylated

ERK1/2 quantification was done using Odyssey V 3.0 software and the following equation:

pERK pERK βββactin(P) = tERK tERK βββactin(T)

Equation 3. Quantification of ERK1/2 for preliminary studies. 45 CHAPTER THREE

RESULTS

3.1. Spinal GPER activation is pronociceptive

Intrathecal (i.t.) administration of the GPER agonist G-1 produced a characteristic behavioral response consisting of scratching, biting and licking directed caudally, to the tail and hind limbs, along with a slight hind limb scratching directed toward the flank.

The total time the mice engaged in this specific behavior upon G-1 (0.1 nmol, 0.5 nmol, 1 nmol/mouse) treatment was 45 ± 13.4 s/10min (n = 8), 60.6 ± 5.6 s/10min (n = 10), 127.3

± 19 s/10min (n = 7). The responses were statistically significant (F (3, 28) = 19.43, *P <

0.0001) for all the doses tested.

The vehicle had no effect (0 ± 0 s/10min, n = 7) (Fig.10A). This behavior was apparent 3-5 min after i.t. delivery of G-1 and almost disappeared at 8 min after injection.

Thus, mice were observed for 10 min after injection in all behavioral experiments.

To determine whether the response elicited with G-1 is pain-related, the effect of morphine on G-1-induced behavior was assessed. Morphine pre-treatment (1, 3, and 10 mg/kg, s.c.) completely abolished the effect of GPER activation at all the analgesic doses tested (1.75 ± 0.8 s/10min, n = 8,; 0 ± 0 s/10min, n = 7; 0 ± 0 s/10min, n = 7, respectively) (Fig. 10B); F (3, 28) = 79.61. This suggests that the behavioral response elicited with G-1 is related to nociception.

To further verify GPER specificity of the response, the effect of the GPER antagonist

G15 on G-1-induced pain-related behavior was examined. G15 (0.1, 1 and 10 nmol/mouse, s.c.) caused a dose-dependent inhibition (34.8 ± 8 s/10min, n = 7; 20 ± 4.2*

46 s/10min; 4.5 ± 2* s/10min, n = 6, respectively; F (3, 21) = 10.97, *P < 0.001) of caudally directed biting and scratching produced by i.t. G-1 with an ID 50 value of 3.95 ± 1.35 nmol/mouse (Fig. 10C). Comparisons with the G-1 alone group were done using one-way

ANOVA and the statistc information for the groups pre-treated with G15 0.1 nmol/mouse, 1 nmol/mouse and 10 nmol/mouse was: df = 11, F(1,11) = 1.17, P = 0.3

(not significant); df = 10, F(1,10) = 27.93, P = 0.0035; df = 10, F(1,10) = 202.65, P = 0, respectively.

47 Fig. 10. Spinal GPER activation results in nociceptive behavior in mice. A , Effects of varying doses (0.1-1 nmol/mouse) of GPER agonist G-1 administered i.t. in mice. The duration of caudally directed scratching, biting and licking induced by G-1 was determined over a 10-min period starting immediately after injection. This type of behavior was not observed in vehicle-treated mice. *P < 0.01 when compared to vehicle. B, Effect of morphine on G-1-induced scratching, biting and licking response in mice. Morphine was administered s.c. 30 min prior to i.t. (challenging) injection of G-1. **P < 0.0001 when compared to G-1 alone. C, Effect of GPER antagonist G15 on G-1-induced nociceptive behavior in mice. G15 was given s.c. 30 min prior to i.t. injection of G-1. *P < 0.001 when compared to G-1 alone. A-C , Data are given as the means and S.E.M. for groups of 6-10 mice. (Deliu et al., 2010; Deliu et al., 2011a)

48 3.2 GPER is present in the mouse spinal cord at mRNA and protein level

Presence of GPER in pain-specific areas of the spinal cord is well documented in the rat, but not in the mouse.

Fig. 11. GPER expression in the mouse spinal cord. A-C , GPER immunoreactivity (irGPER) is observed in the superficial dorsal horn in the mouse lumbar spinal cord sections labeled with GPER antiserum using avidin-byotin complex ( A, B ) or FITC procedure ( C). B, Higher magnification of the area outlined in A. irGPER is detected in terminals and neurons of small diameter (arrows) in the superficial dorsal horn ( B, C ). Scale bar: A, 250 µm; B, C 50 µm. D, RT-PCR results of GPER mRNA in different segments of the mouse spinal cord, C-cervical, T-thoracic, L-lumbar, S-sacral. Housekeeping gene β-actin was used as an internal control. The product size of mouse GPER was 944 bp and that of β-actin was 647 bp. Results are representative of 3 experiments. (Deliu et al., 2010) 49 Since the behavioral studies evaluating the role of GPER in the modulation of spinal pain were carried out in mice, presence of GPER needed to be established in the mouse.

Examination of tissue sections prepared from the spinal cord of six mice showed that

GPER immunoreactivity was present in the superficial layers of the mouse spinal dorsal horn (Fig. 11A-C). At mRNA levels, GPER appeared to be enriched only in the lumbar section of the mouse spinal cord (Fig. 11D).

3.3. Supraspinal GPER activation is pro-nociceptive

To test the sensory performance at the supraspinal level, we measured the latency to responses from the hot plate set at 52 ºC. The cutoff latency was set at 30 s.

Baseline latency was evaluated at three time points before administration of compounds. The latency of the nocifensive behavior was measured 10, 20 and 30 min after PAG microinjection. The time-course of the %MPE is presented in Table 1.

Intra-PAG delivery of G-1 (10, 50, 100 pmol) dose-dependently decreased nociceptive threshold in the hot plate test (%MPE was -11.3 ± 8%, -25.4 ± 8%, -47.7 ±

15%, respectively, Fig. 12). The G-1-induced effect waned over time, but there was a significant reduction in the nociceptive threshold 10 min after administration, in the rats treated with 50 and 100 pmol G-1 (F (3, 18) = 3.95, P < 0.05, Fig. 12B). Pre-treatment with G15 (500 pmol) prevented G-1 (50 pmol) from eliciting its hyperalgesic effect

(%MPE brought to -1 ± 1%, F (1, 10) = 6.12, P < 0.05, Fig. 12A, B).

50 Treatment %MPE %MPE %MPE

Group 10 min 20 min 30 min

Vehicle 4.3 ± 9 10.7 ± 17 -4.7 ± 3

G-1 10 pmol -11.3 ± 8 -19.2 ± 10 -16.1 ± 13

G-1 50 pmol -25.4 ± 8* -20 ± 6 -24.5 ± 8

G-1 100 pmol -47.7 ± 15* -38.7 ± 15 -32.3 ± 12

G15 500 pmol + G-1 50 -1 ± 1** -0.3 ± 1 -0.5 ± 3

pmol

Table 1. Time course of %MPE in the rat hot plate test. P < 0.05 compared to vehicle (*) or to G-1 50 pmol (**).

51 Fig. 12. Intra-PAG microinjection of G-1 is pro-nociceptive. A, Time-courses of the %MPE in rats treated with vehicle, G-1 (10, 50 and 100 pmol) or G-1 (50 pmol) 10 minutes after G15 (500 pmol). B, %MPE 10 minutes after treatment (same treatment groups as in A). Data are given as the means and S.E.M. for groups of 4-7 rats. P < 0.05 compared to control (*) or to50 pmol G-1 (**).

52 3.4.1. GPER activation with G-1 results in depolarization of spinal superficial dorsal horn neurons

Whole-cell patch clamp recordings in current-clamp mode were made from neurons located in the superficial laminae of acute lumbar spinal cord slices, which had a mean resting membrane potential of -58 ± 1 mV, mean input resistance of 301 ± 24 M Ω and a mean capacitance of 19.6 ± 0.8 pF (n = 31). The response of a laminae II spinal neuron is shown in Fig. 13A: G-1 (100 nM) depolarized the neuron from a baseline level of -50.5 mV to – 41 mV. The G-1-induced depolarization returned to a baseline level of -50.4 mV following drug washout. G-1 depolarized 61% (19/31) of the superficial dorsal horn neurons from an average baseline of -58.3 ± 1.6 mV to -53 ± 1.7 mV (n=19), producing a mean depolarization of 5.3 ± 0.4 mV (Fig. 13C, D). Depolarizations in response to G-1 were apparent 1-3 min after drug application. 39% (12/31) of the neurons tested did not change membrane potential upon G-1 treatment (Vm = -57.8 ± 1.6 mV). G15 application alone (1 µM) did not change the average membrane potential from baseline levels (Vm =

-57 ± 2.4 mV, n=15, Fig. 2C). Nevertheless, G15 pre-treatment blocked G-1 induced depolarization (n=15) in all superficial dorsal horn neurons tested (F (1, 32) = 121, P <

0.00001, Fig. 13C). Fig. 13B presents a representative recording from a neuron pre- treated with G15, where G-1 failed to induce depolarization of the membrane.

53 Fig. 13. GPER stimulation results in depolarization of superficial dorsal horn neurons (SDH) in the rat spinal cord. A, Representative continuous membrane potential recording of a SDH neuron depolarized by bath-applied GPER agonist G-1 (100 nM). B, Pre-treatment of SDH neuron with GPER antagonist G15 (1 µM) prevents G-1- induced depolarization (representative trace). C, G-1 (100 nM) depolarizes SDH neurons with a mean value of 5.3 ± 0.4 mV (n=19). G15 (1 µM) has no effect on resting membrane potential, but pre-treatment with G15 blocks G-1-induced effect (n=15). *P < 0.00001 when compared to G-1 alone. D, Depolarizations induced by G-1 (100 nM) in individual SDH neurons (3-9 mV).

3.4.2. G-1 depolarizes neurons located in the lateral PAG

Whole-cell patch clamp recordings in current-clamp mode were made from neurons located in the lateral PAG region of the midbrain slice, which had a mean resting

54 membrane potential of -50.4 ± 2.6 mV, mean input resistance of 163 ± 15 M Ω and a mean capacitance of 24 ± 2 pF (n = 12). A representative recording of a neuron that depolarized in response to G-1 is shown in Fig. 14A. Upon G-1 (100 nM) application, the membrane potential changed from a baseline of -59.4 mV to -53.2 mV; after drug washout, the membrane potential returned to a baseline level of -58 mV. G-1 depolarized all the PAG neurons analyzed (6/6) from an average baseline of -54.2 ± 2.9 mV to -49 ±

3.2 mV, producing a mean depolarization of 4.4 ± 0.9 mV (Fig. 14C, D). Conversely,

G15 (1 µM) treatment resulted in hyperpolarization of 83% (5/6) of the tested PAG neurons, from baseline levels of -52.2 ± 2.3 mV to -58.9 ± 2.5 mV; the mean hyperpolarization was -5.7 ± 1.2 mV (Fig. 14C, D) and lasted for at least 30 min in all responsive cells. A representative recording is shown in Fig. 14B. One cell was followed until its membrane potential returned to baseline, which occurred 45 min after drug washout (Fig. 14E). G15 completely prevented G-1 from eliciting its depolarizing effect; a representative recording from a cell hyperpolarized with G15 is shown in Fig. 14B. In one cell G15 had no effect on its own; nevertheless, G-1 application after G15 did not elicit a response either (Fig. 14F). It is noteworthy that the changes in membrane potential induced by GPER ligands were relatively slow in onset, apparent 2-4 min after drug application.

55 Fig. 14. Effects of GPER stimulation or blockade on membrane potential of lateral PAG neurons. A, Bath application of G-1 (100 nM) produces membrane depolarization in a lateral PAG neuron (representative trace). B, Representative current-clamp recording of a lateral PAG neuron hyperpolarized by GPER antagonist G15 (1 µM), in which G-1 failed to produce its depolarizing effect. C, G-1 (100 nM) depolarizes lateral PAG neurons with a medium value of 4.4 ± 0.9 mV (n = 6), whereas G15 (1 µM) produces a medium hyperpolarization of -5.7 ± 1.2 mV, which is not reversed by G-1. D, Changes in membrane potential induced by G-1 (100 nM, 3.5 to 7 mV) or G15 (1 µM, -4.5 to -8 mV). E, Membrane potential returns to baseline 45 min after G15 washout. F, Recording from a cell that does not modify its membrane potential in response to either G15 or G-1 application. 56 2+ 3.5.1. GPER activation elevates [Ca ]i in cultured spinal neurons

2+ 2+ The mean basal cytosolic Ca concentration, [Ca ]i, of cultured spinal neurons was

124 ± 3.2 nM (n = 151 cells), which is in agreement with the concentration (30 – 200 nM) reported in mammalian central neurons (Connor, 1986). Bath application of G-1 (10

2+ nM, 100 nM, 1000 nM) induced a concentration-dependent rise in [Ca ]i, with a mean amplitude of 114 ± 2.6 nM (n = 37 cells), 617 ± 6.1 nM (n = 43 cells) and 1040 ± 11.2

2+ nM (n = 29 cells), respectively (Fig. 15C). [Ca ]i elevation elicited with G-1 occurred slowly, beginning 1-3 min after G-1 administration and reached a plateau after 5-7 min; representative traces are shown in Fig. 15B. Pre-treatment with GPER antagonist G15 (1

2+ µM, 15 min) largely abolished (15 ± 1 nM, n = 42 cells, P < 0.05) the increase in [Ca ]i induced by G-1 (100 nM) (Fig. 15A, C); a representative recording from a cell pre-treated with G15 for 15 min and then stimulated with G-1 is shown in blue in Fig. 15B.

57 Fig. 15. G-1 application elevates cytosolic Ca 2+ in spinal neurons . A, Fura-2 AM fluorescence ratio (340/380 nm) before (left) and after (right) administration of G-1 (100 nM) in the absence (top panels) and presence (bottom panels) of G15 (1 µM); the fluorescence ratio scale (0-2) is magnified in top left panel. B, Representative recordings 2+ 2+ of cytosolic Ca concentration, [Ca ]i, in response to different concentrations of G-1 (10-1000 nM) and G-1 (100 nM) after pre-treatment with G15 (1 µM, 15min). C, 2+ Comparison of increases in [Ca ]i produced by G-1 (10-1000 nM) and G-1 (100 nM) after pre-treatment with G15 (1 µM, 15min); G-1 induced a concentration-dependent rise 2+ in [Ca ]i; the effect was abolished by G15. P < 0.05 compared to control (*) or to G-1 alone (**). (Deliu et al., 2011a)

58 2+ 3.5.2. GPER activation elevates [Ca ]i in cultured PAG neurons

2+ 2+ The mean basal cytosolic Ca concentration, [Ca ]i, of cultured PAG neurons was 164

2+ ± 2.6 nM (n = 136 cells). Bath application of G-1 (100 nM) elevated [Ca ]i, with a mean amplitude of 795 ± 8.3 nM (n = 79 cells) (Fig. 16C). Similar to the effect observed in

2+ spinal cord neurons, [Ca ]i elevation elicited with G-1 occurred slowly, beginning 3 min after G-1 application and reached a maximum after 7 min; a representative trace is shown in black in Fig. 16B. Pre-treatment with GPER antagonist G15 (1 µM, 15 min) largely

2+ reduced (56 ± 3 nM, n = 57 cells, P < 0.05) the rise in [Ca ]i elicited with G-1 (100 nM)

(Fig. 16A, C); a representative recording from a cell pre-treated with G15 for 15 min and then stimulated with G-1 is shown in red in Fig. 16B.

59 Fig. 16. G-1 application elevates cytosolic Ca 2+ in cultured PAG neurons . A, Fura-2 AM fluorescence ratio (340/380 nm) before (left) and after (right) administration of G-1 (100 nM) in the absence (top panels) and presence (bottom panels) of G15 (1 µM); the fluorescence ratio scale (0-2) is magnified in top left panel. B, Representative recordings 2+ 2+ of cytosolic Ca concentration, [Ca ]i, in response to G-1 (100 nM) in absence or 2+ presence of G15 (1 µM, 15min pre-treatment). C, Comparison of increases in [Ca ]i produced by G-1 (100 nM) alone or after pre-treatment with G15 (1 µM, 15min); P < 0.05 compared to control (*) or to G-1 alone (**).

60 3.6. GPER activation results in cytosolic and mitochondrial ROS accumulation

Stimulation of H2-DCFDA loaded spinal neurons for 15 min with G-1 (100 nM) lead to a 3.3 times (231%) increase in relative fluorescence (2495 ± 205, n = 23), which is statistically significant (F (1, 39) = 46.2, P < 0.00001) when compared to untreated cells

(752 ± 121), suggesting oxidation of the dye in response to cytosolic ROS production

(Fig. 17A). G15 (1 µM, 5 min) pre-treatment prevented G-1 from inducing cytosolic

ROS accumulation, maintaining relative fluorescence at basal levels (643 ± 91, n = 17, F

(1, 38) = 53.69, P < 0.00001) (Fig. 17A).

Mitochondrial superoxide production was significantly elevated (F (1, 23) = 27.84, P

< 0.00005) in spinal neurons upon G-1 (100 nM) incubation as suggested by the 2.1 times

(110%) increase of MitoSOX Red fluorescence (142 ± 15, n = 12 in G-1 treated vs. 59 ±

4, n = 13 in untreated neurons) (Fig. 17B). The G-1 effect was largely diminished after

G15 (1 µM) pre-treatment (78 ± 7, n = 10, F (1, 20) = 11.86, P < 0.005) (Fig 17B).

61 Fig. 17. GPER-mediated cytosolic and mitochondrial ROS accumulation in spinal neurons. A, GPER agonist G-1 (100 nM) increases cytosolic ROS levels in cultured spinal cord neurons, whereas pre-treatment with GPER antagonist G15 (1 µM) prevents G-1-induced response, maintaining ROS at basal levels. B, G-1 (100 nM) elevates mitochondrial ROS; pre-treatment with G15 blocks G-1 effect. P < 0.05 compared to control (*) or to G-1 alone (#).

In cultured PAG neurons, basal H2-DCFDA fluorescence was 465 ± 97 (n = 13 cells).

G-1 application produced a significant increase (800%) in cytosolic ROS dye fluorescence compared to untreated cells (4195 ± 291, n = 19 cells, F (1, 30) = 105.21, P

< 0.00001, Fig. 18A). Pre-treatment with G15 (1 µM) largely diminished the effect of G-

1 (Fig. 18A), bringing H2-DCFDA relative fluorescence to 809 ± 135 (n = 16 cells, F (1,

33) = 97.84, P < 0.00001).

62 Fig. 18. GPER-mediated cytosolic and mitochondrial ROS accumulation in PAG neurons. A, GPER agonist G-1 (100 nM) increases cytosolic ROS levels in cultured PAG neurons, whereas pre-treatment with GPER antagonist G15 (1 µM) prevents G-1- induced response. B, G-1 (100 nM) elevates mitochondrial ROS; pre-treatment with G15 blocks G-1 effect. P < 0.05 compared to control (*) or to G-1 alone (#).

Additionally, mitochondrial ROS levels were increased in response to G-1 (100 nM) in cultured PAG neurons (MitoSOX Red fluorescence was 401 ± 32, n = 17 G-1 treated cells vs. 177 ± 14, n = 15 untreated cells; F (1, 30) = 37.69, P < 0.00001, Fig. 18B). G15

(1 µM) pretreatment maintained MitoSOX Red fluorescence at basal levels (173 ± 13, n

= 16 cells, F(1, 31) = 41.86, P < 0.00001), despite further G-1 application (Fig. 18B).

Modifications in dye fluorescence are proportional to ROS accumulation.

Representative pictures illustrating the H2-DCFDA fluorescence in cultured spinal and

PAG neurons in response to different treatments are shown in Fig. 19.

63 Fig. 19. Cytosolic ROS dye fluorescence in response to GPER activation in cultured spinal and PAG neurons. H2-DCFDA fluorescence in representative spinal (top) and PAG (bottom) neurons upon control vehicle (left), G-1 100 nM (middle) or G15 1 µM + G-1 100 nM treatment (right).

Both H2-DCFDA and MitoSOX Red are cumulative dyes, their fluorescence increases with exposure time; loading time with a particular ROS dye was according to the manufacturer’s protocol and our preliminary studies and differed between mitochondrial and cytosolic ROS experiments; however, the incubation time with a particular treatment was identical in all ROS experiments. In Fig. 20, representative time courses of ROS fluorescence upon application of various treatments is shown. According to the GPER ligands kinetics, we chose 15 min as the most appropriate incubation time.

64 Fig. 20. Time-course of mitochondrial ROS accumulation. Representative recordings of MitoSOX Red fluorescence modification in response to control vehicle (black) or G-1 treatment in a spinal (red) and in a PAG neuron (green).

65 CHAPTER FOUR

DISCUSSION

4.1. GPER and spinal pain

The first round of experiments was conducted to evaluate the ability of previously characterized GPER ligands (Bologa et al., 2006; Dennis et al., 2009) to affect nociceptive behavior initiated at the spinal level in vivo . The data presented here clearly shows that intrathecal challenge of mice with the GPER agonist G-1 results in a behavioral syndrome indicative of nociception, consisting of caudally directed biting, scratching and licking. This type of behavior is characteristic for nociceptive molecules and considered reflective of pain if it is reduced or prevented by morphine. Accordingly, in our experiments, subcutaneous delivery of analgesic doses of morphine completely abolished the G-1-induced nociceptive response. The GPER antagonist G15 dose- dependently diminished the effect of G-1, confirming the specificity of GPER involvement in nociception at the spinal level.

Activation of GPER in vivo has been reported to produce pain sensitization. When injected in the hind paw of male rats, G-1 (0.025-2.5 nmol) decreases mechanical nociceptive threshold, pointing to a hyperalgesic effect of peripheral GPER stimulation

(Kuhn et al., 2008). Moreover, 2.5-125 nmol/kg G-1, s.c., increases visceral hypersensitivity to 5-hydroxytryptophan in ovariectomized female rats (Lu et al., 2009).

Both studies are indicative of peripheral nociceptive sensitization, although it is possible that G-1 crossed the blood-brain barrier to enhance hyperalgesic response via central mechanisms. The lowest pro-algesic dose of G-1 evaluated in the present study appears to

66 be 0.1 nmol (~0.4 nmol/kg), while the doses reported by Lu et al. (2009) are considerable higher. In addition, we have reported effects of 1 nmol (4 nmol/kg) G-1 upon i.p. delivery to mice (Dennis et al., 2009), indicating that G-1 has the ability to accumulate and elicit responses centrally. Thus, even low systemic G-1 doses, such as those used by Kuhn et al. (2008), are likely to have produced additional pain sensitization upon reaching central levels.

Expression of GPER has been intensely studied in the rat central nervous system

(Brailoiu et al., 2007; Dun et al., 2009; Hazell et al., 2009; Takanami et al., 2009), whereas reports regarding its expression in mice are rather scarce and restricted to the brain (Hazell et al., 2009). Since some behavioral experiments were carried out in mice, we needed to confirm GPER presence in the mouse spinal cord. Similar to what was observed in the rat (Dun et al., 2009), GPER immunoreactivity was detected in cells and cell processes of the superficial dorsal horn, which are known to be involved in pain transmission. GPER mRNA was enriched in the lumbar spinal segment, which is particularly important since i.t. injections are performed between L5 and L6 and the nociceptive behavior observed in the current study involves the tail and the hind limbs.

There are no reports regarding the possible differential distribution of GPER mRNA in various segments of the rat or mouse spinal cord. Nevertheless, using in situ hybridization, Takanami et al. (2009) showed very low levels of GPER mRNA in the rat lumbar spinal cord, although the lumbar dorsal root ganglia (DRG) presented very high levels; moreover, upon selective spinal dorsal rhizotomy of the lumbar spinal cord, GPER immunoreactivity in the lumbar superficial dorsal horn was significantly reduced on the ipsilateral side, suggesting that GPER protein was synthesized in the DRG and then

67 transported down the dorsal root to the terminals of nociceptive afferents in laminae I and

II of the spinal dorsal horn. Thus, in the mouse, it is likely for this phenomenon to occur mostly in the cervical, thoracic and sacral spinal regions, while in the lumbar segment

GPER is also synthesized locally.

Further insight was sought on the mechanism underlying spinal nociceptive effects in response to GPER activation. Neurons in the superficial laminae of the spinal dorsal horn function as the main output source to supraspinal sites, playing a crucial role in the integration of peripheral nociceptive messages (Bishop, 1980; Craig, 2000; Furue et al.,

2004). Depolarization of superficial dorsal horn neurons is involved in central nociceptive sensitization (Ji and Woolf, 2001), while analgesic molecules, such as opioids, are known to hyperpolarize neurons in the laminae II of the spinal cord (Eckert and Light, 2002). Consistent with a pro-algesic effect of GPER stimulation at the spinal level, we found that bath application of G-1 depolarized 61% of the neurons analyzed, while G15 prevented this response. Membrane properties such as resting membrane potential, membrane resistance and time constant are correlated with the neuronal response patterns (Stebbing et al., 1999; Abdulla and Smith, 2001; Weng and Dougherty,

2002). The depolarizations observed with G-1 were only partial, never large enough to produce a depolarization block. However, a partial depolarization turns a neuron with nociceptive specific (NS) functional profile into a neuron with wide dynamic range

(WDR) functional profile (Weng and Dougherty, 2002). NS neurons have passive membrane properties that would require higher amounts of synaptic current to drive the cells toward spike threshold as compared to WDR neurons. A modest change in membrane potential makes a NS neuron respond to a low intensity stimulus that would

68 typically affect only the activity of a WDR cell. Therefore, the partial membrane depolarization elicited by GPER activation in postsynaptic neurons is likely to contribute to the development of hypersensitivity to peripheral stimuli in intact animals after G-1 administration.

2+ The increase in [Ca ]i is important for release (Berridge et al., 2000;

Brailoiu et al., 2003) and further transmission of nociceptive information. Moreover,

2+ 2+ membrane depolarization elicits a rapid [Ca ]i rise via voltage-gated Ca channels, a mechanism involved in pain transmission in superficial dorsal horn (SDH) neurons

2+ (Heinke et al., 2004). We found that G-1 both depolarizes and increases [Ca ]i in spinal cord neurons, with both effects being subject to GPER blockade. This further supports a pro-nociceptive effect of GPER at the spinal level.

2+ The increase in [Ca ]i in spinal neurons is followed by downstream mitochondrial

Ca 2+ uptake, with subsequent production and release of ROS, resulting in synaptic plasticity underlying chronic pain (Kim et al., 2011). Indeed, recent research suggests that reactive oxygen and nitrogen species are involved in central sensitization associated with acute and chronic inflammatory and noninflammatory neuropathic pain via modulation of protein kinases, alterations in glutamatergic transmission, neuroinflammation and modulation of ion channels such as TRPV1 (Westlund et al., 2010; Salvemini et al.,

2011). These alterations take place in the periphery, in the spinal cord and at supraspinal sites. At the spinal level, ROS contribute to neuropathic pain by reduction of GABA release (Yowtak et al., 2011), sensitization of WDR neurons in the dorsal horn (Lee et al.,

2007) and potentiation of synaptic excitability in spinal superficial laminae (Lee et al.,

2009). We found that GPER activation results in cytosolic and mitochondrial ROS

69 accumulation, indicating a potential mechanism for GPER-mediated spinal nociceptive sensitization.

4.2. GPER and supraspinal pain

Supraspinal sites involved in pain processing – such as the amygdala, periaqueductal gray (PAG), trigeminal sensory nucleus, dorsal raphe or central medial nucleus of the hypothalamus – express GPER immunoreactivity (Hazell et al., 2009). In the hot plate test, hind paw licking and jumping are considered to be the result of supraspinal sensory integration (Caggiula et al., 1995; Rubinstein et al., 1996). Thus, the reduction of the hot plate nociceptive threshold upon PAG G-1 microinjection indicates specific involvement of GPER in supraspinal pain facilitation. This finding is further supported by the ability of G15 pre-treatment to prevent the acute hyperalgesic effect of G-1. Accordingly, the supraspinally mediated decrease in opioid analgesia in response to estrogen (Bernal et al.,

2007; Ji et al., 2007), as well as estrogenic pro-nociceptive effects upon amygdala implantation (Myers et al., 2011) are at least partially dependent on GPER.

The contribution of other estrogen receptor types cannot be excluded, especially since

ER α is also localized on PAG neurons projecting to rostroventral medulla in the descending pain modulatory pathway (Loyd and Murphy, 2008). Nevertheless, the PAG is a midbrain region involved in the regulation of a variety of functions and behaviors, as shown by both anatomical and physiological studies. Thus, it modulates the autonomic nervous system controlling blood pressure, heart rate and regional blood flow, all of which have shown to be influenced by estrogen (Alper and Schmitz, 1996; Morgan and

Pfaff, 2001). Moreover, defensive or aggressive behaviors are initiated in the PAG

70 (Bandler et al., 1985; Bandler and Carrive, 1988), both of which are under gonadal hormone modulation (Albert et al., 1991; Scordalakes and Rissman, 2004). Sexual behavior is also under PAG regulation, as PAG stimulation facilitates lordosis behavior in female rats (Sakuma and Pfaff, 1979a; McCarthy et al., 1991), while PAG lesions suppress this behavior (Sakuma and Pfaff, 1979b; Lonstein and Stern, 1998). Therefore, the presence of both GPER and ER α in the PAG may have additional functional implications, which are worthy of further exploration.

In order to unravel cellular mechanisms underlying GPER-dependent pain facilitating effect observed in vivo , an electrophysiological approach was undertaken. GABAergic interneurons located in the lateral PAG are under inhibitory control of endogenous opioid system; they further synapse on excitatory PAG neurons projecting to OFF cells in the rostroventral medulla (RVM) that mediate pain inhibition in the spinal cord; additionally, they decrease the activity of RVM ON cells involved in pain facilitation (Fig. 6). Thus, we examined the effect of GPER activation in the lateral PAG neurons, which undergo hyperpolarization in response to opioids, a mechanism known as “disinhibition”

(Osborne et al., 1996). In agreement with the pro-nociceptive effect observed upon in vivo delivery into the PAG, G-1 depolarized all the lateral PAG neurons analyzed. Thus, the pain facilitation in response to G-1 is likely a result of a PAG-mediated overall inhibitory effect on the OFF cells located in the RVM (Fig. 6). Interestingly, GPER antagonist G15 hyperpolarized 83% of the tested neurons, which may indicate constitutive activity of GPER in the PAG, unlike in the spinal cord neurons. Although not explored in the mammals, expression of aromatase in cells projecting to the PAG has been reported in the Japanese quail (Absil et al., 2001); thus, estrogen may be synthesized

71 and released locally to account for the GPER constitutive activity we observed in the

PAG.

A noteworthy observation is that G-1 application did not modify the hyperpolarization induced by G15. The 1:10 concentration ratio chosen for G-1 (100 nM) and G15 (1 µM) was according to their binding affinity to the GPER (Bologa et al., 2006;

Dennis et al., 2009) and proven to be perfectly antagonistic in the optical imaging experiments employed in this study, as well as in other published reports (Dennis et al.,

2009; Tica et al., 2011). However, a recent study suggests that G15 may function as a partial agonist of transcription mediated by ERs. Apparently, 10 µM G15 presents ~25% activity at the estrogen response element (ERE) reporter compared to the maximal E2 response; in addition, 10 µM G15 inhibits the E2-induced ERE effect by ~30%, with no significant response at 1 µM (Dennis et al., 2011). The partial agonist activity observed with G15 applies only to transcriptional events through the classical ERs and not to its rapid effects (Dennis et al., 2009; Dennis et al., 2011). It is debatable whether or not 1

µM G15 might have acted also through a genomic mechanism to induce the G-1 insensitive prolonged hyperpolarization in the PAG. While rapid and delayed effects of are easily distinguishable from each other at the extremes in terms of mechanisms, actions that have onset times of minutes are difficult to classify as genomic or nongenomic. For example, glucocorticoids have genomic actions on lymphocytes with onset latencies of 20 min (McEwen et al., 1978). Thus, there is a possibility that G15 initiated its effect via GPER antagonism (rapidly) and continued it via ERE modulation

(after 10-15 min); however, at this point it is rather a speculation as no experiments have been employed to test this hypothesis. Moreover, although tested for GPER specificity 72 and shown not to interact with at least 25 other GPCRs (Bologa et al., 2006; Blasko et al.,

2009; Dennis et al., 2009; Dennis et al., 2011), GPER ligands may still have heretofore uncharacterized GPER-independent activities. These off target effects are hard to exclude when using pharmacological approaches, therefore, future studies should focus on genetic manipulations in order to clarify the additional mechanisms involved in G15-induced response.

Intriguingly, although G15 hyperpolarized PAG neurons in acute midbrain slices and prevented G-1 from modulating its response, in whole animals G15 did not elicit antinociception, while perfectly antagonizing G-1-induced hyperalgesia. This may be explained by an increased G15 metabolism in vivo .

Similar to the responses observed in cultured spinal cord neurons, both elevation of

2+ [Ca ]i, and increases in mitochondrial and cytosolic ROS occurred also in the PAG neurons, upon application of GPER agonist G-1; all these effects were sensitive to GPER blockade by G15. At both spinal and supraspinal sites, ROS accumulation was reported to be involved in the development of morphine antinociceptive tolerance (Doyle et al.,

2009; Doyle et al., 2010). Moreover, at the supraspinal level, ROS are mediators linking activation of metabotropic glutamate receptors type I (mGluR1/5) to an increase in neuronal excitability and pain behavior (Li et al., 2011). This confirms our hypothesis that ROS accumulation in the periaqueductal gray following GPER activation is one of the mechanisms underlying its hyperalgesic effects.

The sources of ROS in the CNS are various, including monoamine oxidase, cyclooxygenase, NADPH oxidase, nitric oxide synthase, as well as mitochondrial oxidative phosphorylation (Kukreja et al., 1986; Pou et al., 1992; Chetkovich et al.,

73 1993). As shown in this study, GPER activation results in cytosolic Ca 2+ increase. Thus, the major source of ROS is likely the mitochondria, as mitochondrial oxidative phosphorylation increases during enhanced Ca 2+ signaling (Dykens, 1994; Dugan et al.,

1995; Reynolds and Hastings, 1995); moreover, studies in isolated mitochondria indicate that Ca 2+ accumulation causes leakage of electrons from the respiratory chain and increases the production of superoxide (Castilho et al., 1995). This mechanism likely occurs similarly in the two types of cells tested, as G-1 induces a ~2 fold increase in mitochondrial ROS both in spinal and PAG neurons. However, cytosolic ROS elevation in response to GPER activation is ~8 fold in the PAG and only ~3.5 fold in the spinal neurons. The significance of this difference, as well as the underlying mechanism necessitates further exploration. One hypothesis worth testing is the differential regulation of NADPH oxidase, a superoxide-generating enzyme system involved in the development of central sensitization associated with inflammatory hyperalgesia (Ibi et al.,

2008) and peripheral nerve injury-induced neuropathic pain (Kim et al., 2010). NAPDH oxidase is regulated by protein kinase C (PKC) either directly or via activation of mitogen-activated protein kinase (MAPK) pathways (Sharma et al., 1991; El Benna et al.,

1996). Both kinases are downstream targets of GPER activation in various cell types, including neurons (Kuhn et al., 2008; Prossnitz et al., 2008; Goswami et al., 2011). It is worth testing whether the expression and activation of different NADPH isoforms in spinal and PAG neurons may account for the disparate cytosolic ROS responses observed upon GPER activation (Salvemini et al., 2011).

74 4.3. Summary and conclusions

The studies presented in this thesis evaluated the involvement of G protein-coupled estrogen receptor GPER in central nociceptive processing, namely in the spinal superficial dorsal horn and in the periaqueductal gray.

Behavioral experiments carried out in mice indicate pro-algesic dose-dependent effects of GPER agonist G-1 upon spinal injection, which was sensitive to GPER antagonism with G15 (systemic pre-treatment). Similarly, intra-PAG microinjection of

G-1 produced a concentration-dependent decrease in nociceptive threshold in the hot plate test; this response was also G15 sensitive. Thus, both at spinal and supraspinal sites

GPER activation in vivo is pro-nociceptive.

The underlying mechanisms hypothesized and tested in our studies are generally similar, although some discrepancies have also been noted. In acute spinal cord slice preparations, G-1 application depolarizes superficial dorsal horn neurons, indicating involvement of GPER in spinal nociceptive sensitization. Likewise, recordings from lateral PAG neurons located in acute midbrain slices show depolarization in response to

G-1 applied by superfusion; this underscores GPER involvement in pain facilitation at supraspinal levels. Antagonism with G15 prevents the responses induced by G-1 in both paradigms; however, G15 induces a G-1-insensitive hyperpolarization of PAG neurons by a mechanism that remains to be further examined.

2+ In cultured spinal and PAG neurons, G-1 administration results in [Ca ]i elevation that is G15-sensitive. Similarly, cytosolic and mitochondrial ROS accumulation is a common mechanism underlying pro-nociceptive effects of GPER activation in both types

75 of cells. Nevertheless, in the PAG, there is increased contribution of cytosolic ROS compared to spinal neurons.

Corroborating all these findings, the present thesis supports the involvement of GPER in spinal nociceptive sensitization and PAG-dependent pain facilitation. As depicted in

Fig. 9, GPER activation is followed by increases in intracellular calcium concentration, as well as cytosolic and mitochondrial ROS levels, all of which are correlated with nociceptive processing via depolarization of specific neurons located in the superficial dorsal horn or in the PAG. These results are in agreement with previous studies indicative of algogenic effects of GPER activation, implicating GPER in some of the sensitizing effects of estrogen on nociception. Accordingly, GPER may represent a new target for pain management (Kuhn et al., 2008), and its specific modulation may prove beneficial in the treatment of visceral and trigeminal pain disorders, which are more prevalent in high estrogenic states (Craft, 2007). Moreover, there is an overall agreement between studies evaluating the detrimental proliferative role of GPER in breast, endometrial and ovarian (Prossnitz and Barton, 2009; Prossnitz and Barton, 2011), conditions highly correlated with pain. Targeting these cancers with GPER antagonists to reduce proliferation may prove beneficial in alleviating the associated pain.

76 4.4. Future directions

The work presented in this thesis gives limited insight regarding the involvement of

GPER in nociceptive processing at central sites and some of the putative underlying mechanisms. Additional research should be carried out to further clarify the role of GPER in pain physiology and pathology, as well as the therapeutic implications of these findings.

i) Putative interactions of GPER with the opioid system via modulation of K + channels

Although not presented in the Results section of this thesis, the work we conducted lead to preliminary results that support the proposal of additional experiments to further our understanding of the mechanism of GPER-induced spinal nociception.

As shown in Fig. 21, the pain-related behaviors induced by G-1 intrathecal administration to mice is reduced or abolished not only by morphine, but also by κ

(U50,488H) and δ (SNC80) selective agonists. Moreover, very low doses of morphine, U50,488H or SNC80, that do not significantly hyperpolarize the membrane of superficial dorsal horn neurons (Eckert and Light, 2002), completely prevent the depolarization induced by G-1 (Fig. 22).

77 Fig. 21. Effects of selective opioid receptor agonists on G-1 induced pain-related behavior in mice. The effect of G-1 is reduced or completely abolished by analgesic doses of A, morphine (MOR, 1, 3 and 10 mg/kg); B, U50,488H (U, 1, 3 and 10 mg/kg); C, SNC80 (SNC, 0.5, 1.25 and 5 mg/kg). **P < 0.0001; +P < 0.01 compared to G-1 alone. Data are representative for groups of 6-10 mice. (Deliu et al., 2010; Deliu et al., 2011a)

78 Fig. 22. Pre-treatment with selective opioid agonists prevents G-1 induced depolarization of superficial dorsal horn neurons. Representative current-clamp recordings indicating that pre-application of A, morphine (100 nM), B, U50,488H (100 nM) or C, SNC80 (100 nM) abolishes the G-1 induced membrane potential modification. (Deliu et al., 2011a)

It is generally acknowledged that opioid receptors couple predominantly to Gi/Go proteins to stimulate G protein-activated inwardly rectifying K + channels (GIRKs)

(Waldhoer et al., 2004). In addition, the increase of membrane conductance mediated by the activation of GIRKs in the postsynaptic substantia gelatinosa (lamina II) neurons leading to the subsequent hyperpolarization is one of the mechanisms underlying µ− , κ− and δ− opioid receptor induced spinal analgesia (Grudt and Williams, 1993; Grudt and

Williams, 1994; Grudt and Williams, 1995; Eckert and Light, 2002).

Therefore, future experiments employing electrophysiology in the superficial laminae neurons of the dorsal spinal cord should include evaluation of the ionic currents

79 underlying G-1 induced depolarization, with particular emphasis on testing the hypothesis that GPER activation results in inhibition of K + channels.

Redox regulation of ion channel activity has been reported for NMDA receptors expressing the NR1 subunit (Sullivan et al., 1994), Ca 2+ -activated K + channel hslo

(DiChiara and Reinhart, 1997), the voltage-gated K + channel Kv1.4 (Ruppersberg et al.,

+ 1991), the inwardly rectifying K channel IRK 1 (Ruppersberg and Fakler, 1996), as well as GIRKs (Zeidner et al., 2001). Additionally, inhibition of various types of K + channels in response to ROS has been reported in rat (Muller and Bittner, 2002) and mouse hippocampal neurons (Takeuchi and Yoshii, 2008), as well as in cultured catecholaminergic CATH.a neurons (Yin et al., 2010). Thus, it is worth investigating the correlation between ROS elevation and ion channel modulation in response to GPER activation. This mechanism should be evaluated also in PAG neurons; however, given the distinct electrophysiological responses reported in this thesis, activation or blockade of

GPER in the PAG may involve differential regulation of ion channels compared to that in the superficial dorsal horn.

Furthermore, it is worth considering that our preliminary results showed an increase in phosphorylated ERK 1/2 in the spinal cord upon intrathecal injection of G-1 (Fig. 7); activated ERK 1/2 acts on A-type potassium channels (Adams et al., 2000; Schrader et al., 2006) in the dorsal horn neurons to inhibit their activity and potentiate arriving excitatory inputs (Hu and Gereau, 2003; Hu et al., 2003; Hu et al., 2006; Hu et al., 2007).

Thus, a new hypothesis would be that the G-1 induced depolarization is a result of A-type

K+ channels inhibition downstream of ERK 1/2 and should be prevented by GPER antagonists and MEK inhibitors. Moreover, if this proves to be true, GPER activation

80 should result in increased excitability of the SDH neurons; therefore, the effect of G-1 on action potentials evoked by current injection, i.e. on first spike latency and spike frequency, should be evaluated.

ii) involved in the ROS responses to GPER activation

Another study of great interest should focus on obtaining insights regarding the differences and similarities observed between the GPER-mediated effects on cytosolic and mitochondrial ROS in the spinal cord and in the PAG. In this respect, we propose studying the GPER-dependent regulation of mitochondrial superoxide dismutase (SOD) and of various NADPH oxidase enzyme complexes in both types of neurons.

Experimental approaches should include electrophysiological and imaging studies with

SOD and NADPH oxidase inhibitors, as well as immunoblotting of the enzyme proteins or protein subunits upon treatment with GPER ligands.

iii) Genetic approaches to confirm GPER specificity of the nociceptive effects

GPER-specificity of the results reported in this thesis has been confirmed using pharmacological approaches. However, to further strengthen this conclusion, the studies should also be performed in GPER-deficient animals and neuronal cells.

iv) GPER contribution to trigeminal and cancer pain

Future studies in whole animals should focus on the contribution of GPER to modulation of trigeminal and cancer pain, using wild-type and GPER-deficient animals of both sexes; intact females should be separated into groups according to their estrous

81 cycle state and ovariectomized females supplemented with both low and high estrogen doses.

Animal models of trigeminal neuralgia are generated by producing various lesions to the trigeminal nerve, such as infraorbital nerve loose ligation (Vos et al., 1994), application of complete Freund adjuvant to the trigeminal root (Benoliel et al., 2002), removal of tooth pulp (Gobel and Binck, 1977) or implanting the trigeminal root with chromic suture (Burchiel, 1980). However, these animal models present shortcomings related to the lack of reproducibility in terms of the observed sensory, motor and molecular changes; therefore, a more accurate model of trigeminal neuropathic pain should be used, such as application of Cobra venom to the animal’s infraorbital nerve trunk; the procedure is easy to perform and is likely to be relevant for human studies (An et al., 2011). These animals should be further treated intrathecally or intra-PAG with

GPER ligands and the degree of mechanical allodynia examined using von Frey hairs.

Modeling cancer has been difficult; however, in the past three decades, several animal models of cancer pain have been developed, that can be divided into five categories: bone cancer pain models, non-bone cancer pain models, cancer invasion pain models, cancer chemotherapeutic-induced peripheral neuropathy models and spontaneous occurring cancer pain models (Pacharinsak and Beitz, 2008). Of particular interest would be studying the involvement of GPER in a metastatic bone cancer pain model in which MDA-MB231 human breast cancer cells (5 x 10 5 per mL) are injected into the femoral arteries of nude rats (Bauerle et al., 2005). Breast and prostate cancers often lead to skeletal metastasis (Coleman, 2000; Blouin et al., 2005) and their clinical outcome is negatively correlated with GPER expression (Prossnitz and Barton,

82 2011). Thus, this model is likely to be relevant for studying the involvement of GPER in cancer pain, especially if both GPER-negative and GPER-overexpressing MDA-MB231 cancer cells will be used for the animal model generation. Animals will be examined for mechanical hyperalgesia and allodynia using weight-bearing tests and von-Frey monofilaments, respectively.

Moreover, electrophysiological recordings of MDA-MB231 injected animals in the presence or absence of GPER ligand treatments in the superficial spinal cord neurons and

PAG neurons should be performed. Apparently, in animals injected with carcinoma cells, the receptive field size for superficial dorsal horn neurons is enlarged and nociceptive- specific neurons, which are excited only by noxious stimuli, are responsive to non- noxious stimuli (Urch et al., 2003). In addition, the responses of superficial wide dynamic range neurons (excited by both painful and non-painful stimuli) are significantly increased, while deeper WDR neurons are minimally affected, suggesting involvement of both ascending and descending facilitation pain pathways (Urch et al., 2003; Urch et al.,

2005).

v) Involvement of the neurosteroid E2 to GPER pro-nociceptive responses at central sites

To test for constitutive GPER activation at spinal and PAG levels, the effect of the neurosteroid E2 should be evaluated by using aromatase inhibitors, both in vivo , ex viv o and in vitro . This paradigm should be evaluated using both pharmacological and genetic approaches.

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