Relaxin and the Paraventricular Nucleus of the Hypothalamus

Megan Susan McGlashan

A Thesis Presented to The University of Guelph

In partial fulfillment of requirements for the degree of Master of Science in Biomedical sciences

Guelph, Ontario, Canada

RELAXIN AND THE PARAVENTRICULAR NUCLEUS OF THE HYPOTHALAMUS

Megan Susan McGlashan Advisor: University of Guelph, 2013 Professor A. J. S. Summerlee

The hormone relaxin regulates the release of the magnocellular hormones, oxytocin and vasopressin, from the central nervous system. Studies have yet to determine whether relaxin regulates magnocellular hormone release through the circumventricular organs alone, or whether relaxin can act on the brain regions containing the magnocellular neurons as well. The paraventricular nucleus of the hypothalamus was isolated from other brain regions and maintained in vitro, in order evaluate the effects of the relaxin and relaxin-3 on the somatodendritic release of oxytocin and vasopressin. At 50 nM concentrations, relaxin induced oxytocin release, while relaxin-3 inhibited oxytocin release. Neither relaxin nor relaxin-3 had an effect on the vasopressin release, however the RXFP3 specific agonist, R3/I5, induced vasopressin release. The effect of the relaxin peptides on the electrical activity of neurons in the paraventricular nucleus was also evaluated. Relaxin depolarized magnocellular neurons while relaxin-3 hyperpolarized the neurons. Relaxin and relaxin-3 appear to have differential effects on the magnocellular neurons of the paraventricular nucleus.

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ACKNOWLEDGEMENTS

When we first enter graduate school, we are very much children in the ways of research.

As the saying goes, it takes a village to raise a child, so it was for my master’s degree. I would like to thank my village. This thesis was made possible because of you.

I would like to thank my advisor, Alastair Summerlee. Thank you for your support, for your guidance, and for teaching me that research is as much about people as it is about bubbling buffer in a lab. Above all I would like to thank you for your continued confidence in my abilities.

Thank to my lab family. Jordan- thank you for seeing my potential. Roman- your motivational talks were pivotal in my success. Lindsay- thank you for your support and for inspiring me through your perseverance and success.

I would also like to thank the members of my advisory committee- John LaMarre,

Brian Wilson and Bettina Kalisch. Your advice was invaluable. Also, thank you to Craig

Bailey for being an enthusiastic and capable teacher; learning electrophysiology was far easier than it was reputed to be.

Finally, I would like to thank my incredible family. You are responsible for teaching me that education is not just steps between childhood and adulthood, rather it is a lifetime affair. This thesis is as much yours as it is mine. I could not have succeeded without you. Thank you.

DECLARATION OF WORK PERFORMED

I declare that all the work described in this thesis was carried out by the author alone in the laboratories of Dr. Summerlee and Dr. Bailey in the Department of Biomedical

Sciences at the University of Guelph, Guelph, Ontario, Canada.

TABLE OF CONTENTS

ACKNOWLEDGEMENTS...... iii

DECLARATION OF WORK PERFORMED...... iv

TABLE OF CONTENTS...... v

LIST OF FIGURES...... vii

LIST OF TABLES...... vii

LIST OF ABBREVIATIONS ...... viii

LITERATURE REVIEW ...... 1

Relaxin family of peptides ...... 1 Historical perspective ...... 1 Structure of relaxin peptides...... 3 Receptors...... 5 Signalling pathways ...... 7 Relaxin-3 ...... 9

Paraventricular Nucleus of the Hypothalamus ...... 12 Paraventricular Nucleus of the Hypothalamus ...... 12 The HNS as model of hormone release in the brain ...... 14 Heterogeneity within the parvocellular divisions ...... 15 Electrical properties of neuron types ...... 17 Relaxin and the hypothalamic-neurohypophyseal system ...... 18

INTRODUCTION AND PURPOSE OF EXPERIMENTS ...... 21

METHODS ...... 23

Animals ...... 23

Experiment 1...... 23 Development of the in vitro release protocol...... 23 Perfusion aCSF system ...... 24 Static aCSF system ...... 25 Static HEPES buffer system ...... 27

Experiment 2...... 28 vi

Patch-clamp recording...... 28

RESULTS...... 31

Experiment 1: The effect of relaxin-3 on the release of oxytocin and

vasopressin from isolated PVN...... 31 Perfusion aCSF system...... 32 Static aCSF system...... 32 Oxytocin...... 33 Vasopressin...... 34

Static HEPES buffer system...... 37 Oxytocin...... 37 Vasopressin ...... 38

Experiment 2: The effect of relaxin peptides on the intracellular electrical activity of isolated neurons from the PVN in vitro...... 42 The effect of relaxin-3 and relaxin-2 on the membrane potential of magnocellular neurons ...... 42 The effect of relaxin-3 and relaxin-2 on the membrane potential of parvocellular neurons...... 45

DISCUSSION...... 47

The regulation of somatodendritic release in the PVN...... 54 The function of relaxin-3 ...... 58 Technical considerations...... 59

CONCLUSION...... 63

REFERENCES ...... 64

vii

LIST OF FIGURES

Figure 1 Oxytocin release from PVN slices perfused with human relaxin-2...... 33

Figure 2 No effect of human relaxin-3 or R3/I5 on oxytocin release from the pvn...... 35

Figure 3 The effect of human relaxin-3 and R3/I5 on vasopressin release from the pvn...... 36

Figure 4 The effects of relaxin peptides on oxytocin release from PVN of young rats...... 39

Figure 5 The effects of relaxin peptides on vasopressin release from PVN of young rats...... 40

Figure 6 Identification of neuron types within the PVN by expression of low threshold spikes or outward rectification ...... 43

Figure 7 The effects of human relaxin-3 and human relaxin-2 on the membrane potential of magnocellular neurons in the paraventricular nucleus...... 44

Figure 8 The effects of human relaxin-3 and human relaxin-2 on the membrane potential of neuroendocrine parvocellular neurons

in the paraventricular nucleus...... 46

LIST OF TABLES

Table 1 Change in oxytocin and vasopressin release from isolated PVN when incubated in HEPES buffer with relaxin peptides...... 41

viii

LIST OF ABBREVIATIONS

Abbreviation Term

α-MSH Alpha-melanocyte stimulating hormone

ACTH Adrenocorticotropic hormone cAMP Adenosine 3’, 5’-cyclic monophosphate cGMP Guanosine 3’, 5’-cyclic monophosphate

DAG Diacyl glycerol

CRH Corticotropin-releasing hormone

GABA Gamma-aminobutyric acid

GRH Gonadotropin-releasing hormone

GPCR G-protein coupled receptor

HNS Hypothalamic-neurohypophysial system

IA current A-type potassium current

INSL Insulin-like

IP3 Inositol trisphosphate

LCDVs Large dense cored vesicles

LGR Leucine-rich repeat G-protein coupled receptor

LTS Lower threshold spikes

MAPK Mitogen-activated protein kinase

MMP Matrix metalloproteinase mRNA Messenger ribonucleic acid ix

NE Neuroendocrine

NF- kB Nuclear factor kappa B

NO Nitric oxide

NOS Nitric oxide synthase

PA Pre-autonomic

PI3K Phosphoinositide-3 kinase

PKA Protein Kinase A

PKC Protein Kinase C

PVN Paraventricular nucleus of the hypothalamus

RXFP Relaxin family peptide receptor

SON Supraoptic nucleus

TGF-β Transforming growth factor-β

TRH Thyroid-releasing hormone 1

LITERATURE REVIEW

1. Relaxin family of peptides

1.1 Historical perspective

Relaxin was first identified in 1926 when Fredrick Hisaw observed a loosening of the pelvic symphysis of female guinea pigs following an injection of ovarian extracts derived from pregnant guinea pigs (Hisaw 1926). The protein responsible for this relaxation was later isolated and named relaxin (Fevold et al. 1930). Relaxin was assumed to be a hormone of pregnancy but there was little progress made towards understanding its actions until a technique to extract and purify relaxin from ovarian extracts was developed in 1974 (Sherwood and O’Byrne 1974) and a sensitive radioimmunoassay was developed in 1979 (Sherwood and Crnekovic 1979). Once relaxin could be purified, researchers were able to determine its structure, sequence and binding sites (James et al. 1977, Schwabe et al. 1976; 1977). The relaxin-1 gene was cloned in 1983 (Hudson et al. 1983) followed by the identification of a second human relaxin gene, relaxin-2 (Hudson et al. 1984). Human relaxin-1 is found in primates only, while human relaxin-2 is equivalent to relaxin-1 (relaxin) in non-primates (Sherwood

2004). In fact, we now recognize seven members of the relaxin family: relaxin-1 (Hudson et al. 1983); relaxin-2 (Hudson et la. 1984); relaxin-3 (Bathgate et al. 2002a); and multiple insulin-like peptides (INSL), including INSL3 (Adham et al. 1993); INSL4 (Chassin et al. 1995); INSL5 (Conklin et al. 1999); and, INSL6 (Lok et al. 2000). It is commonly 2 accepted to refer to relaxin-2 simply as relaxin because it was the first relaxin to be identified.

Originally, relaxin was considered to be a reproductive hormone, but it has subsequently been shown to be produced in a variety of other tissues, and it is now known to have functions in a number of different systems including the brain, heart, cardiovascular system, skin, kidney, prostate etc. (See review: Sherwood 1994; Bathgate et al. 2013). Summerlee and colleagues (1984) were the first to report an action of relaxin in the brain. They reported an action for intravenously administered relaxin in suppressing reflex milk-ejection in anaesthetized rats, an action that implies a direct effect of relaxin on oxytocin release from the pituitary. Initially, there was considerable debate about whether or not relaxin was able to cross the blood-brain barrier and enter the nervous tissue to exert its effects, or whether relaxin was acting through a mediator outside the blood-brain barrier (Geddes and Summerlee 1995). It has since been shown that relaxin is produced in small quantities in the brain (Osheroff and Ho 1993,

Gunnersen et al. 1995) and that its cognate receptor, RXFP1, is present in brain tissue

(Burazin et al. 2005, Ma et al. 2006). Moreover, relaxin has been shown to affect drinking behaviour (Thornston and Fitzsimmons 1995, Summerlee and Robertson 1995), increase the frequency of action potentials of vasopressin and oxytocin neurons in the supraoptic nucleus (SON) of lactating rats (Dayanithi et al. 1987,Parry et al. 1990, Way and Leng 1992); suppress the electrically stimulated release of oxytocin in lactating rats

(O’Byrne et al. 1986); delay the timing of birth (Jones and Summerlee 1986, Summerlee et al. 1998a); and affect appetite in conscious rats (McGowan et al. 2005; 2006; 2007). 3

Relaxin-3 was the last of relaxin family peptides to be identified (Bathgate et al.

2002a). Relaxin-3 is highly conserved between species and is expressed almost exclusively in the brain where it appears to function as a neurotransmitter (Bathgate et al. 2013, Callander and Bathgate 2010). Its cognate receptor, RXFP3, is also found in the brain (Lui et al.2003a; Sutton et al. 2004; Ma et al. 2007), as is RXFP1, which relaxin-3 binds as well (Lui et al. 2003a). Although it has been suggested that relaxin-3 may have a significant role in the central nervous system, but to date, most of the research has focused on determining whether or not relaxin-3 has the same actions as relaxin in the brain.

1.2 Structure of relaxin peptides

The relaxin peptide family is classified as part of the insulin superfamily of peptides due to a number of structural similarities (James et al. 1977, Sherwood et al.

1974, Hudson et al. 1983). Relaxin/insulin peptides are synthesised as prohormones, consisting of a signal peptide and three chains in a B-C-A chain configuration (Hudson et al. 1983; 1984). The primary structure of relaxin/insulin peptides consists of the A and B chains connected by cysteine residues forming two inter-chain disulfide cross bridges

(Sherwood and O’Bryne 1974, Schwabe and MacDonald 1977, Rosengren et al. 2006a,

Rosengren et al. 2006b). The C-chain allows correct folding of the B and A chains into their final configurations (Hudson et al. 1984) but is cleaved in the mature form of the peptide. The A-chain consists of two parallel α-helices that are separated by a short ß strand, forming a loop (Eigenbrot et al. 1991, Rosengren et al. 2006a, Rosengren et al. 4

2006b). The A-chain loop is linked by two cysteine residues that form an intra-chain disulfide cross bridge (Schwabe and MacDonald 1977). The B-chain consists of an α- helix and a ß-strand (Eigenbrot et al. 1991, Rosengren et al. 2005a; 2006b). The A- and

B-chains are linked by two disulfide cross bridges formed by four cysteine residues

(Schwabe and MacDonald 1977).

Relaxin peptides have conserved characteristics which are important for the structure and activity of these peptides. All relaxin peptides have three glycine residues adjacent to the cysteine residues that form the disulfide cross-bridges (Büllesbach and

Schwabe 1995). The conserved glycines contribute flexibility to the peptide that allows for correct folding of the chains (Büllesbach and Schwabe 1994, Sherwood 1994).

Relaxin peptides are also characterised by containing the sequence Arg-X-X-X-Arg-X-X-

Ile/Val located on the B-chain α-helix, which is the receptor binding cassette (Büllesbach and Schwabe 2000).

Relaxin peptides also contain secondary binding sites. The secondary binding site that enables relaxin and relaxin-3 to bind RXFP1 is located in the C terminus region of the A chain of the peptides (Büllesbach and Schwabe 1994, Hossain et al. 2008, Hossain and Wade 2010). This secondary relaxin binding site sits opposite the primary receptor cassette on the B-chain and acts cooperatively with the primary binding site in forming a unique bond (Park et al. 2008).

Relaixn-3 also binds RXFP3; RXFP3 is bound by relaxin-3 alone (Lui et al. 2003a).

The B-chain of relaxin-3 is sufficient for the binding and activation of RXFP3 (Lui et al.

2003a, Hossain et al. 2008). The α-helix of the B-chain of relaxin-3 contains the RXFP3 5 binding region, while the c-terminus of the relaxin-3 B-chain is required for RXFP3 activation (Kuei et al. 2007). Taking advantage of this, the RXFP3 specific agonist, R3/I5, was created by substituting the A-chain of relaxin-3 with the A-chain of INSL5 that lacks the RXFP1 secondary binding site (Lui et al. 2005, Haugaard-Jonsson et al. 2008). This

RXFP3 agonist provides a tool for distinguishing between the presence of RXFP1 and 3 binding sites, which may be useful in distinguishing between the actions of relaxin-2 and relaxin-3.

1.3 Receptors

The relaxin family of peptide receptors were identified nearly 80 years after the discovery of relaxin (Hisaw 1926, Kumagai et al. 2002). The identity of the receptors was elusive simply because it was assumed that relaxin and insulin receptors would be similar: the cognate receptor for insulin is a tyrosine kinase receptor (Ebina et al. 1985,

Ullrich et al. 1985). We now know, however, that relaxin receptors are G protein- coupled receptors (GPCRs) (Hsu et al. 2002, Lui et al. 2003a).

The identification of the relaxin family peptide receptors followed two events.

First, genome searches identified a group of GPCRs with leucine-repeat-containing regions, classified as leucine-rich repeat GPCRs (LGR) (Hsu et al. 2000; 2002). Members of the LGR group include the follicle-stimulating hormone receptor, the luteinizing hormone receptor , the thyroid-stimulating hormone receptor , and two orphan receptors LGR7 and LGR8 (Hsu et al. 2000; 2002). INSL3 deficient mice were shown to exhibit bilateral cryptorchidism, a deficiency that was identical to a GPCR mutation in 6 mice (Zimmerman et al. 1999, Overbeek et al. 2001). The rat orthologue of this GPCR was determined to be LGR8 (Hsu et al. 2002) and INSL3 was found to bind LGR8 with high affinity, thereby identifying the LGR8 as the cognate receptor for INSL3 (Kumagai et al. 2002). LGR7 was then identified as the receptor for relaxin due to co-localized LGR7 mRNA expression with relaxin binding sites (Hsu et al. 2002). LGR7 deficient mice were found to exhibit the same impaired nipple development and parturition phenotype as relaxin deficient mice, thereby confirming LGR7 as the receptor for relaxin (Zhao et al.

1999, Krajnc-Franken et al. 2004). In order to establish consistent nomenclature for the relaxin family of receptors (RXFP), LGR 7 and LGR 8 were renamed RXFP1 and RXFP2 respectively (Bathgate et al. 2006).

Relaxin binding sites and high levels of RXFP1 mRNA have been identified in diverse brain structures, including in the SON, paraventricular nucleus of the hypothalamus (PVN), cerebral cortex, the circumventricular organs, amygdala, thalamus, hippocampus, midbrain pons, the pineal gland, the supermaxillary nucleus, and the arcuate nucleus (Osheroff and Phillips 1991, Burazin et al. 2005). The identification of relaxin in multiple brain regions suggests a role for relaxin in mediating diverse processes (Gundlach et al. 2009).

The distribution of RXFP2 is restricted compared with RXFP1 (Shen et al. 2005,

Sedaghat et al. 2008). RXFP2 mRNA has been detected in regions associated with sensimotor function, including thalamus nuclei and the motor, frontal and sensory layers of the cerebral cortex (Shen et al. 2005, Sedaghat et al. 2008). Surprisingly studies have not detected INSL3 expression in the brain, which researchers suggest is due to a 7 failure to detect it, rather than INSL3 not being expressed in the brain (Callander and

Bathgate 2010).

RXFP3, also known as somatostatin- and angiotensin- like peptide receptor, is unique among the relaxin family peptide receptors as it is bound by relaxin-3 alone (Lui et al.2003a). It is expressed extensively in the brain: RXFP3 mRNA is expressed in the rat at high levels in the olfactory bulbs, anterior olfactory nucleus, PVN, SON, thalamus, the amygdala, the dentate gyrus and CA3 field of the hippocampus, the interpeduncular nucleus, the superior colliculus and in the nucleus incertus (Sutton et al 2004, Ma et al.

2007). The distribution of relaxin-3 binding sites was found to correlate well with the expression of RXPF3 mRNA (Sutton et al. 2004). Studies of the mouse and primate have found complementary findings of RXFP3 expression and binding in the brain further illustrating a conservation of the relaxin-3-RXFP3 system across species (Smith et al.

2010, Ma et al. 2007; 2009).

1.4 Signalling pathways

RXFP1 is coupled to G-proteins that induce increases in adenosine 3’, 5’-cyclic monophosphate (cAMP) levels, while RXFP3 is coupled to other G-proteins that comparatively reduce cAMP levels (Bathgate et al. 2013). In the human embryonic kidney (HEK) 293 cells, RXFP1 induced cAMP accumulation is biphasic and modulated by multiple pathways (Nguyen et al. 2003, Halls et al. 2006). The initial phase (10-15 minutes) involves the coupling of Gαs and subsequent activation of adenylate cyclase that increases in cAMP levels, which is then inhibited by the coupling of GαoB (Halls et al. 8

2006). The second phase of RXFP1 induced cAMP accumulation is initiated by recruitment of Gαi3 protein and subsequent release of the Gαβγ subunits. The released subunits activate phosphoinositide-3 kinase (PI3K), which stimulates the translocation of protein kinase C (PKC)ζ to the cell membrane, where it activates adenylate cyclase to produce cAMP (Halls et al. 2006; 2009, Nguyen et al. 2003, Nguyen and Dessauer 2005).

RXFP1 activates theses pathways in several different cell types (Halls et al. 2009). RXFP1 also appears to increases cAMP levels through tyrosine kinase mediated inhibition of phosphodiesterase (Bartsch et al. 2001; 2004).

The RXFP1 receptor activates non-cAMP signalling pathways as well, including the mitogen-activated protein (MAPK) kinase pathway and nitric oxide (NO) pathway (Zhang et al 2002, Mookerjee et al. 2009). Activation of RXFP1 on renal fibroblasts induces the

MAPK- neuronal NO synthase (NOS)-NO-guanosine 3’, 5’ -cyclic monophosphate (cGMP) pathway to interfere with transforming growth factor-β (TGF-β1) signalling by inhibiting

Smad2 phosphorylation (Mookerjee et al. 2009) and induces matrix metalloproteinase

(MMP )-1,-2, -9 expression through inducible iNOS signalling (Chow et al. 2012). The activation of the RXFP1 receptor on fibrochondrocytes stimulates the expression of

MMP-9. As described above, ligand binding to RXFP1 activates PI3K which initiates a cascade of downstream signalling events including the activation of PKCζ, protein kinase

B (Akt) and MAPK resulting in enhanced of MMP-9 transcription; activation of PI3K-

PKCζ stimulates nuclear factor kappa B (NF- kB) while activation of all three pathways is necessary for AP-1 transcription factor activation (Ahmad et al. 2012). 9

In contrast to RXFP1, very little is known of the signalling pathways that mediate the actions of RXFP3. RXFP3 inhibits forskolin-stimulated cAMP accumulation in

HEK293T cells by coupling the GTPγS protein (Lui et. al 2003a; 2003b). RXFP3 also binds

GTPαS protein to activate PI3K and PKC, resulting in phosphorylation of extracellular signal-regulated kinase (ERK) 1/2 (Van der Westhuizen et al. 2007). Activation of RXFP3 on murine SN56 septal neurons also up-regulates the transcriptions of AP-1 and NF- kB, which are downstream of Gαi/o proteins (Van der Westhuizen et al. 2010).

1.5 Relaxin-3

Relaxin-3 was first identified in 2002 (Bathgate et al. 2002a), 76 years after Hisaw

(1926) discovered relaxin. Although relaxin-3 mRNA has been detected in reproductive tissue as well as in other tissues, the highest expression of relaxin-3 has been detected in the brain (Bathgate et al. 2002a, Burazin et al. 2002, Tanaka et al. 2005, Silvertown et al. 2010). Relaxin-3 is expressed centrally in the pontine raphe nucleus, the ventral and lateral periaqueductal gray, a region dorsal to the lateral substantia nigra, and the nucleus incertus (Bathgate et al. 2002a, Burazin et al. 2002, Tanaka et al. 2005). Relaxin-

3 expression in the nucleus incertus is highly conserved across species such that it is now an accepted identifying characteristic of the brain region (Bathgate et al. 2002a, Burazin et al. 2002, Tanaka et al. 2005, Donizetti et al. 2008, Silvertown et al. 2010).

The nucleus incertus is also known as the pars ventromedialis of the dorsal tegmental nucleus (Morest 1961), the nucleus recessus pontis medialis (Jennes et al. 10

1982), and nucleus ‘o’ (Messen and Olzewski 1949). It is located in of the midline floor of the fourth ventricle, adjacent to the ventromedial border of the caudal dorsal tegmental nucleus (Goto et al. 2001). The nucleus incertus has extensive efferent projections, many of which express relaxin-3 (Goto et al. 2001, Ma et al. 2007; 2009), suggesting that functions that can be regulated by the nucleus incertus might involve relaxin-3, and vice versa.

The nucleus incertus has been proposed to be involved in arousal (Goto et al.

2001, Olucha-Bordonau et al. 2003). The nucleus incertus shares bidirectional connections with the interpeduncular nucleus and the median raphe, which function in inhibiting behavioural activation, and it has been proposed that the nucleus incertus promotes processes mediating arousal through inhibitory inputs to these brain regions

(Goto et al. 2001). Relaxin-3 knockout mice have been found to exhibit hypo-activity during the arousal (dark) phase of their circadian cycle in comparison to wild-type mice

(Smith et al. 2012), which would be consistent with the view expressed above.

The nucleus incertus is involved in stress response (See review: Tanaka et al.

2010). The neurons of the nucleus incertus, including the majority of the neurons that produce relaxin-3 (Tanaka et al. 2005), express the corticotropin-releasing factor type 1 receptor (Bittencourt and Sawchenko 2000). There is an increase in c-fos immunoreactivity in the nucleus incertus following intracerebroventricular administration of corticotropin-releasing factor (Tanaka et al. 2005). Evidence that relaxin-3 is involved in stress response include the finding that relaxin-3 expression in the nucleus incertus increases following repeated forced swim stress, which is blunted 11 by prior administration of corticotrophin releasing factor 1 antagonist (Banerjee et al.

2010). The findings from work on relaxin-3 knock-out mice considering the effects of short-term or chronic stress have not been consistent: one study found significant differences between knockout and wild-type mice in measures of fear and anxiety

(Watanabe et al 2011); another study reported only trends (Smith et al. 2009); while another reported no effect at all (Smith et al. 2012). Given the propensity for knockout models to overcome functional deficits through developmental changes and/or compensatory systems, the findings of the relaxin-3 knockout studies can be seen as an indicator of a function for relaxin-3 in stress function, but whether relaxin-3 potentiates or attenuates stress response still requires investigation.

A key effector site of the stress response is the PVN. The PVN expresses mRNA for both RXFP1 and RXFP3 (Burazin et al. 2005, Tanaka et al. 2005, Silvertown et al. 2010).

A number of regions that project to the PVN also express RXFP3 mRNA, including the bed nucleus of the stria terminalis, dorsomedial and ventromedial nuclei, arcuate nucleus, suprachiasmatic nucleus, anterior and lateral hypothalamus, medial and lateral proptic area, supramammillary nucleus, raphe nuclei, nucleus of solitary tract (Ma et al.

2007). RXFP1 mRNA is also present in the arcuate nucleus, medial and lateral preoptic area, and supramammillary nucleus, as well as the circumventricular organs (Ma et al.

2006).

Relaxin-3 was shown to increase the c-fos immunoreactivity of corticotropin- releasing hormone (CRH) expressing neurons in the PVN, as well as increase plasma concentrations of adrenocorticotropic hormone (ACTH), which would suggest relaxin-3 12 is involved in mediating the neuroendocrine response to stress. Relaxin-3 also reduces the release of thyroid-releasing hormone (TRH), produced by neurons of the PVN, from hypothalamic explants (McGowan et al. 2006). Given that relaxin-3 induces increase in food intake (McGowan et al. 2005; 2006; 2007) when the central administration of CRH or TRH suppresses food intake (Arase et al. 1988, Suzuki et al. 1982), further study is needed to determine what role relaxin-3 has in stress response.

2.0 Paraventricular Nucleus of the Hypothalamus

The PVN is a paired nucleus located adjacent to the dorsal region of the third ventricle. Although it consists of only approximately one third of 1 mm3 of tissue, it is responsible for the integration and regulation of an array of neuroendocrine, autonomic and homeostatic functions. The PVN is an effector site for multiple central systems, including the circumventricular organs that releases angiotensin II to regulate plasma osmolarity (Johnson and Gross 1993), brainstem nuclei that releases serotonin, noradrenaline and adrenaline to regulate the hypothalamic-pituitary-adrenal axis during stress response (Pacak and Palkovits 2001), other brainstem nuclei that release dopamine to regulate sexual behaviour (Melis and Argiolas 1995), the arcuate nucleus that releases neuropeptide-Y, agouti-related peptide, and the precursor to alpha- melanocyte stimulating hormone (α-MSH) to regulate food intake (Schwartz et al. 2000), and the nucleus incertus that expresses relaxin-3 (Ryan et al. 2011)

The PVN consist of a multiplicity of neuron types in three magnocellular subdivisions and five parvocellular subdivisions (Swanson and Sawchenko 1983). The 13 magnocellular subdivisions (rostral, medial and caudal) consist of oxytocin and vasopressin neurons which, along with magnocellular neurons of the SON, are part of the hypothalamo-neurohypophysial system (HNS) that releases hormones from the neurohypophysis into the peripheral circulation (Swanson and Sawchenko 1983). The parvocellular subdivisions of the PVN (anterior, medial, dorsal, posterior and periventricular) contain neuroendocrine neurons, pre-autonomic neurons, and other centrally projecting neurons (Conrad and Pfaff 1976, Swanson and Sawchenko 1983,

Tasker and Dudek 1991).

The PVN is surrounded by GABAergic or glutamatergic interneurons located in what is known as the peri-PVN region (Roland and Sawcheko 1993, Boudaba et al. 1996;

1997). The GABAergic neurons of the peri-PVN region are located in the subparaventricular area which includes: the posterior bed nucleus of the stria terminalis

(BNST), dorsomedial hypothalamus and anterior hypothalamus, as well as the region that includes the suprachiasmatic nucleus (Roland and Sawcheko 1993, Boudaba et al.

1996). Glutamatergic neurons of the peri-PVN region are located in the perifornical area and dorsomedial hypothalamus (Boudaba et al. 1997). GABAergic and glutamatergic interneurons are also located within the PVN itself (Csáki et al. 2000) The majority of synaptic innervation of the PVN is GABAergic or glutamatergic (Deval and van den Pol

1992) implying a significant role for interneurons in controlling the activity of magnocellular and parvocellular neurons of the PVN (See review Herman et al. 2002,

Culliman et al. 2008).

2.1 The HNS as model of hormone release in the brain

The HNS is one the most extensively studied neuroendocrine systems and as such is a model of hormone release in the brain (See reviews: Landgraf and Neumann 2004,

Ludwig and Leng 2006). Magnocellular neurons, so-called because their soma are up to

20-30 µm in diameter, principally produce oxytocin and vasopressin (Swanson and

Sawchenko 1983). These peptides are packaged into neurosecretory granules, shipped down the long axons of the neurons and stored in swellings at the axon terminals.

Electrical activity of these neurosecretory neurons provokes the release of oxytocin and vasopressin into the circulation where they act on remote sites. A great deal is known about the pattern of electrical activity necessary to release these hormones (See review:

Leng et al. 1999, Murphy et al. 2012). Although each action potential travelling down the magnocellular neuron axons causes temporary depolarization of the neuron terminals which is the prerequisite for hormone release, the frequency of action potentials arriving at the terminals determines the amplitude of hormone release.

The electrical activity of oxytocin and vasopressin neurons is different, resulting in different patterns of oxytocin and vasopressin release from the neurons terminals

(Renaud and Bourque 1991). Vasopressin neurons have a characteristic phasic firing pattern and there is an almost constant release of hormone (Wakerley and Lincoln 1973,

Lincoln and Wakerley 1974). In contrast, oxytocin neurons generally have a slower pattern of firing but during reflex milk-ejection (Lincoln and Wakerley 1973, Lincoln and

Wakerley 1974, Wakerley et al. 1994) show a transient elevated firing rate that appears 15 to be coordinated across a significant number of oxytocinergic neurons that results in a bolus of oxytocin released into the blood stream.

Magnocellular neurons do not only release neuropeptides from their axons, they also release neuropeptides from their somas and dendrites (Pow and Morris 1989), and this can occur independent of axonal release (see review Ludwig and Leng 2006). This somatodendritic release is thought to affect the activity in adjacent somas (de Kock et al. 2003, Ludwig and Leng 1997) and the structure of adjacent astrocytes (See review:

Theodosis and Poulain 1993, Theodosis et al. 2008). Magnocellular neurons are tightly packed into discrete areas of the PVN (Swanson and Sawchenko 1983). Nevertheless, cell somas are normally separated by fine lamella-like processes of radial astrocytes

(Theodosis et al. 2008). These processes show a remarkable degree of plasticity. For example, in response to osmotic challenge (Tweedle and Hatton 1977), stress (Miyata et al. 1994), lactation (Tweedle and Hatton 1987), and parturition (Tweedle and Hatton

1982), the astrocytes in the PVN retract their processes. The mechanics underlying this retraction are not well understood but it appears to results in apposition of cell somas, allows the formation of gap junctions (Hatton 1997) and allows multiple pre-synaptic terminals to connect with single post-synaptic boutons (Theodosis et al. 1981, Tweedle and Hatton 1982).

2.2 Heterogeneity within the parvocellular divisions

Unlike the magnocellular subdivisions of the PVN, the parvocellular subdivisions contains a heterogeneity of neuron types that can be broadly classified into 16 neuroendocrine (NE) neurons, pre-autonomic neurons, and other central projecting neurons (Hoffman et al. 1991, Stern 2001).

The NE parvocellular neurons release factors into the hypophyseal portal system to induce the release of hormones from the anterior pituitary into peripheral circulation

(Swanson and Sawchenko 1983). The hypophyseal hormones produced by NE parvocellular neurons of the PVN include CRH, thyrotropin-releasing hormone (TRH) and gonadotropin-releasing hormone (GRH). Some NE parvocellular CRH neurons also co- express vasopressin; vasopressin induces the release of ACTH in the anterior pituitary by binding vasopressin V1b receptors, a vasopressin receptor isoform unique to the anterior pituitary and located on corticotropic cells (Antoni 1993).

The pre-autonomic parvocellular neurons project to the spinal cord, dorsal medulla, and dorsal vagal complex to regulate sympathetic activity, affecting heart rate, blood pressure and respiratory drive (Swanson and Kuypers 1980, Swanson et al. 1980;

Kc and Dick 2010). Specifically, anatomically distinct groups of PVN neurons project to the rostral ventrolateral medulla (Cootes et al. 1998), nucleus of the solitary tract, dorsal vagal complex (Van der Kooy et al. 1984), the pre-Botzinger complex (Kc et al. 2010), and the intermediolateral cell column of the spinal cord (Swanson and Kuypers 1980,

Swanson et al 1980, Zheng et al. 1995). In addition, parvocellular neurons also project to diverse brain regions such as the thalamus, hippocampus, amygdala, and regions in the hypothalamus (Conrad and Pfaff 1976). Like NE parvocellular neurons, centrally projecting parvocellular neurons express oxytocin, vasopressin and CRF (Conrad and

Pfaff 1976, Kc et al. 2002, Sawchenko 1987, Swanson and Sawchenko 1982). 17

2.3 Electrical properties of neuron types

Tasker and Dudek (1991) proposed that it was possible to distinguish between neuronal types in the PVN based on their electrical activity: PVN neurons were originally classified as type I, type II, and type III. Type I neurons express linear voltage-current relationships and a large A-type potassium (IA) current. Type II neurons exhibit lower threshold spiking (LTS). Finally, type III neurons express a small IA current and a lack of

+ LTS. IA current is a transient K current that is active at hyperpolarized membrane potentials so that it delays the rate of action potential generation (Connor and Stevens

1971). LTS are due to a low threshold T-type calcium current activated at membrane potentials below threshold and functions in neuronal oscillatory and bursting behaviour

(Huguenard 1996).

Type I neurons have subsequently been identified as magnocellular neurons

(Tasek and Dudek 1991, Hoffman et al. 1991, Luther and Tasker 2000, Luther et al.

2000). Type II neurons include the PA parvocellular neurons. These neurons exhibit retrograde staining from the spinal cord and medulla and express LTS (Stern 2001). Type

III neurons have been identified as the NE parvocellular neurons based on their uptake of peripherally administered fluoro-gold stain exclusive to neurons projecting to the pituitary (Luther et al. 2002).

2.4 Relaxin and the HNS

Investigations into the effects of relaxin in the HNS have focused on the possible role of relaxin in pregnancy (Sherwood 2004). Summerlee and colleagues (1984) were first to report that relaxin might have an action in the HNS. They showed that intravenously administered relaxin inhibited reflex milk-ejection in urethane anaesthetized rats, which could not be reinstated with exogenous oxytocin, evidence that relaxin acts centrally to inhibit pulsitile oxytocin release from the HNS (Summerlee et al. 1984). Relaxin was also shown to induce increases in blood pressure (Mumford et al. 1989, Parry et al. 1994), an effect that was attenuated by intravenous administration of a vasopressin (V1) receptor antagonist (Parry et al. 1991; Parry and Summerlee 1991), evidence that relaxin also acts centrally to induce vasopressin release. The direct evidence that relaxin regulates the activity of the HNS include that it induces increases in plasma levels of oxytocin and vasopressin in rats (Jones and Summerlee 1987,

Mumford et al. 1989, Parry et al. 1994, Geddes et al. 1994, Summerlee et al. 1998a), and that it increases the firing rate of magnocellular neurons in vivo (Way and Leng 1992).

The above mentioned studies, however, used urethane-anaesthetized rats and urethane, although it produces a stable level of anaesthesia, is known to elevate circulating levels of neurohypophyseal hormones (Spriggs and Stockham 1964). Rats anesthetised with a barbiturate also showed increase in blood pressure and plasma levels of vasopressin (Yang et al. 1995), which would suggest that the effect on vasopressin release is not an artefact of urethane anaesthesia. There is however contradictory evidence showing relaxin instead reduces blood pressure (Conrad and 19

Novak 2004), for example relaxin induces a fall in blood pressure in dogs anaesthetized with barbiturates (Miller and Kisley 1957), causes a relaxation of the mesocecum in rats

(Bigazzi et la. 1988), and provokes arteriolar dilation in skeletal muscle microvasculature

(Wilcox 2009).

There is also the question of how relaxin, which circulates outside the brain during pregnancy and is too large to cross the blood brain barrier, gains access to the HNS.

Dayanithi et al. (1987) suggested that relaxin was acting on the neurohypophysis to affect the release of oxytocin and vasopressin from the neural lobe stores, however subsequent studies failed to detect relaxin binding or RXFP1 mRNA expression in the pituitary (Osheroff et al. 1990, Osheroff and Phillips 1991, Burazin et al. 2005, Ma et al.

2006). It is possible that circulating relaxin acts through the circumventricular organs to affect hormone release from the magnocellular nuclei (Mumford et al. 1989, Parry et al.

1991; 1994, Geddes and Summerlee 1995). Lesioning the subfornical organ, one of the principal components of the circumventricular organs, blocks the effects of relaxin on reflex milk-ejection (Summerlee et al. 1987) and blood pressure (Mumford et al. 1989).

The effects of relaxin in the HNS appears to be mediated through angiotensinergic projections to the magnocellular nuclei as blocking central angiotensin with an angiotensin II receptor blocker partly negates the effects of relaxin on magnocellular hormone release (Geddes et al. 1994).

Since the time of these original controversies, relaxin binding sites and relaxin receptors have been identified inside the blood-brain barrier in rats (Burazin et al. 2005,

Osheroff and Phillips 1991), mice (Piccenna et al. 2005) and primates (Silvertown et al. 20

2010) and relaxin mRNA has been detected in the rostral olfactory nucleus, tenia tecta, pyriform cortex, neocortex, hippocampus, and orbitofrontal and anterior cingulate cortices, and arcuate nuclei (Osteroff and Ho 1993, Ma et al. 2006). These findings suggest that relaxin is produced and active in the brain and substantiates the concept proposed by Geddes and Summerlee (1995) that perhaps there is a central relaxin system within the blood-brain barrier which can be influenced by relaxin circulating in the periphery if the blood levels of the hormone reach a critical level to “spill over” into the brain (Geddes and Summerlee 1995). Finally, the discovery of relaxin-3, which is almost exclusively found in the brain (Burazin et al. 2002, Tanaka et al. 2005, Silvertown et al. 2010) and interacts with RXFP-1, suggests that relaxin-3 may be a predominant form of relaxin in the brain.

INTRODUCTION AND PURPOSE OF EXPERIMENTS

The hormone relaxin regulates the activity of the HNS: relaxin inhibits the pulsatile release of oxytocin driving lactation, while inducing increases in the basal release of oxytocin and vasopressin from the central nervous system (Summerlee et al.

1998a). Peripherally circulating relaxin acts through the circumventricular organs to regulate HNS activity (Summerlee et al. 1987, Mumford et al. 1989, Sunn et al. 2002).

Relaxin and relaxin-3 are produced in the brain (Osheroff and Phillips 1993, Bathgate et al. 2002, Burazin et al. 2005). Studies have yet to determine whether central relaxin regulates HNS activity through the circumventricular organs alone, or whether relaxin can also act directly in the HNS.

The HNS consists of two brain regions: magnocellular regions of the SON and the

PVN (Swanson and Sawchenko 1983). Magnocellular neurons release their hormones from their soma and dendrites (in the brain), as well as from their terminals (from the brain), which can be regulated independently (Ludwig and Leng 2006). The PVN has particularly dense expression of the relaxin peptide receptors RXFP1, bound by relaxin and relaxin-3, and RXFP3, bound by relaxin-3 alone (Bathgate et al. 2002, Burazin et al.

2005, Tanaka et al. 2005, Silvertown et al. 2010). The PVN functions in central processes which have been suggested as involving relaxin-3, including the stress response (Tanaka et al. 2005, 2010, Banerjee et al. 2010, Watanabe et al. 2011), food intake (McGowan et al. 2005, 2007), and reproduction (McGowan et al. 2008), and as such has been a focus of studies investigating the neuroendocrine role of relaxin-3. 22

In order to investigate the actions of relaxin and relaxin-3 in the PVN, the following experiments were conducted:

1. An in vitro release protocol was developed, in which the PVNs of rats were

isolated from their afferent and efferent projections, in order to evaluate the

effect of relaxin and relaxin-3 on the somatodendritic release of oxytocin and

vasopressin in the PVN

2. In order to evaluate the effect of relaxin and relaxin-3 on electrical activity of

neuroendocrine neurons in the PVN, intracellular electrophysiology was

conducted on rat brain slices

METHODS

Animals

Male Wistar rats (Charles Rivers, Saint-Coustant, QC, Canada) were housed at the

Central Animal Facility of the University of Guelph, Ontario. All animals were given food and water ad libitum and maintained on a 12-h light, 12-h dark lighting regimen. All work was done in accordance with an animal usage protocol approved by the Animal

Care Committee at the University of Guelph.

Experiment 1: The effect of relaxin on the somatodendritic release of oxytocin and vasopressin from the PVN

Development of the in vitro release protocol

Three different approaches were used during the development of the in vitro release protocol. In all three approaches, rats were placed under deep anaesthesia using sodium pentobarbital (50mg/Kg IP) and decapitated. Brains were removed in less than two minutes and immersed in ice cold incubation buffer for a further two minutes.

Brains were placed in a brain matrix (Zivic instruments, Pittsburgh, PA) and the region containing the hypothalamus was isolated by slicing in the horizontal plane at the optic chiasma, and 2 mm caudal to the optic chiasma.

The three approaches to the in vitro release protocol differed by the age of rats, buffer, and method of tissue incubation used. The initial approach (experiment 1.1) 24 utilized PVN slices from older rats (greater than 50 days old) perfused with artificial cerebral spinal fluid (aCSF). The aCSF was composed of 124 mM NaCl, 1.2 mM MgSO4,

1.2 mM KH2PO4, 3.25 mM KCl, 26 mM NaHCO3, 10 mM D-glucose and 1.0 mM CaCl2, adjusted to a pH of 7.4 with NaOH and saturated with a gas mixture of 5% CO2 and 95%

O2. The other approaches employed a static system in which the PVN was isolated from its afferent projections and incubated in buffer that was exchanged every 5 minutes.

The PVN was harvested from older rats (greater than 50 days old) and incubated in aCSF

(experiment 1.2), or harvested from young rats (less than 28 days old) and incubated in

HEPES buffer (experiment 1.3). The HEPES buffer was composed of 140 NaCl mM, 5 KCl mM, 10 HEPES mM, 10 glucose mM, 1.2 KH2PO4 mM, 1.2 MgSO4 mM, and 1.8 CaCl2 mM, adjusted to pH 7.4 with NaOH, and saturated with 100% O2 .

Experiment 1.1: Perfusion aCSF system

The PVN was harvested as described above and the tissue block containing the

PVN was mounted to the stage of the vibratome buffer bath, filled with ice cold aCSF, using cyanoacrylate glue (Instant Krazy glue, Borden Company, Willowdale, Ontario,

Canada). The tissue was cut into 300 μm coronal slices with a vibratome (VT1200, Leica

Microsystems, Wetzlar, Germany). The coronal slices were further dissected to separate the left from the right nucleus.

The tissue slices were then transferred to a humid brain slice chambers

(Scientific Systems Design Inc, Mississauga, ON), that were perfused with aCSF (4 mL/minute) and maintained at a temperature of 36°C. Following a 30 minute recovery 25 period, brain slices containing the right and left PVN nuclei were placed in separate slice chambers. The rate of perfusion of aCSF through the slice chambers was reduced to 1.5 mL/5 minutes and the tissue was allowed to equilibrate for a further 30 minutes.

Following a 20 minute interval to establish basal hormone release, the tissue was treated with 50 nM human relaxin-2 in aCSF or aCSF alone for 20 minutes. The incubation media was collected in 10 minute intervals and frozen at -20°C, until further testing using an enzyme-linked assay kits (EIA) (Enzo Life Sciences, Plymouth Meeting,

PA, oxytocin) to determine the oxytocin (catalogue no. ADI-900-153) and vasopressin

(catalogue no. ADI-900-017) concentrations of the samples. The sensitivity of the oxytocin and vasopressin ELISAs were 11.7 pg/mL (range 15.6 - 1,000 pg/mL) and 3.39 pg/mL (range 4.10 -1,000 pg/mL) respectively. The response to relaxin treatment was evaluated by determining the percentage change in hormone concentrations during treatment, in 10 minute intervals, from average basal hormone concentration. The hormone concentrations in the basal and treatment intervals were compared using a two-way repeated measures ANOVA, evaluating the effect of treatment over time.

Experiment 1.2: Static aCSF system

After the tissue containing the PVN was harvested as described above, it was trimmed to remove the hypothalamic tissue surrounding the PVN using the rostral commissure, the third ventricle and the fornices as indicators of the PVN’s location

(-0.80 to -2.30 mm bregma, anterior to posterior (Paxinos and Watson, 1996). The left and right nuclei were separated. Each isolated nucleus was then transferred to a humid 26 brain slice chamber (Scientific Systems Design Inc, Mississauga, ON), perfused with aCSF

(4 mL/minute) and maintained at a temperature of 36°C.

Following 30 minute recovery period, the left and right nuclei were placed in separate wells of a 24 well cell culture plate, which was floated in a heated water bath to maintain the buffer temperature at 36°C. The wells contained 750 µL of aCSF which was replaced every 5 minutes. At the end of a 30 minute equilibrium period, the nuclei were incubated in aCSF alone for three 5 minute intervals to determine basal levels of hormone release, followed by four 5 minute intervals in aCSF containing 50 nM or 500 nM of human relaxin-3, 50 nM or 500 nM of R3/I5 (RXFP-3 agonist), or aCSF alone

(control). The samples of buffer were then frozen at -20°C until the oxytocin and vasopressin concentrations were determined using the EIA kits (described above).

In order to determine the response to treatment, the hormone concentrations were standardized across trials by calculating hormone concentrations as a percentage difference from average basal period before treatment. The hormone concentrations in the basal interval and treatment intervals were compared using a two-way repeated measures ANOVA, evaluating the effect of treatment over time. If the hormone concentrations failed to meet the assumptions of normality and equal variance, or if there was significant interaction between time and treatment, then comparisons were made within treatment groups using a non-parametric alternative, the Friedman’s test.

Post-hoc comparisons were then made using the Dunn’s method. The results of the statistical tests were taken as significant if p<0.05.

Experiment 1.3: Static HEPES buffer system

The PVN nuclei were isolated and allowed to recover as described above

(Experiment 1.2). The nuclei from two brains were then placed in 750 µL of HEPES buffer in one well of a 24 well cell culture plate floated on a heated water bath to maintain the buffer temperature at 36°C. The buffer was replaced every 5 minutes.

At the end of a 30 minute equilibrium period the buffer was replaced by the following: (1) HEPES buffer alone (3 x 5 minutes) followed by (2) high potassium HEPES buffer (2x5 minutes) in order to evaluate tissue viability at the beginning of the trial, then (3) HEPES buffer alone (4 x 5 minutes) to allow the tissue to recover, followed by

(4) HEPES buffer alone (3 x 5 minutes) then (5) 50 nM of relaxin peptide in HEPES (6x5 minutes) to evaluate the change in hormone release with treatment, and finally (6)

HEPES buffer alone (6 x 5 minutes) followed by (6) high potassium HEPES (2x5 minutes) in order to evaluate tissue viability at the end of the trial. The high potassium HEPES buffer consisted of 90 NaCl mM, 70 KCl mM, 10 HEPES mM, 10 glucose mM, 1.2 KH2PO4 mM, 1.2 MgSO4 mM, and 1.8 CaCl2 mM, adjusted to pH 7.4 with NaOH. During the treatment period (period 5), the HEPES buffer contained one of the following: 50 nM human relaxin-2, 50 nM human relaxin-3, 50 nM RXFP3 agonist R3/I5 or HEPES buffer alone (control). The buffer samples were frozen at -20°C.

In order to determine the response to treatment, the hormone concentrations were standardized across trials by dividing hormone concentrations by the change in hormone release during the initial potassium challenge (period 2/period 1). The 28 hormone concentrations during treatment (period 5) were compared as a percentage difference from the average hormone concentration in the interval preceding treatment

(period 4) using a one-way repeated measures ANOVA. If the hormone concentrations failed to meet the assumptions of the normality and equal variance, or if there was significant interaction between time and treatment, then comparisons were made within treatment groups using a non-parametric alternative, the Friedman’s test. Post- hoc comparisons were then made using the Tukey’s test. The results of the statistical tests were taken as significant if p<0.05.

Experiment 2: The effect of relaxin on the intracellular electrical activity of isolated neurons from the PVN in vitro

Patch-clamp recording

Coronal brain slices containing the PVN were prepared as described above in experiment 1.1. After at least a one hour recovery period, slices were placed in a recording chamber mounted under a Zeiss Axioskop 2 FS plus microscope (Carl Zeiss,

Gottingen, Germany). The microscope was fitted with infrared differential interference contrast optics. The slices were visualized using a Dage IR-1000 infrared sensitive charge-coupled device camera (Dage-MTI, Michigan City, IN). Slices were submerged in aCSF, perfused at a rate of 1-2 mL/minute and heated to a temperature of 36 °C with a

C-20 In-line Heater and CL-100 Temperature Controller (Warner Instrument, Hamden,

CT). Recording electrodes were pulled from borosilicate glass pipettes (1.0 mm OD, 29

Sutter Instrument Co., Novato, CA) using a Sutter p-97 pipette puller (Sutter Instrument

Co., Novato, CA), producing a resistance of 5-8 MΩ. The recording electrode was filled with an intracellular solution composed of 120 mm K-gluconate, 5 mm KCl, 2 mm MgCl,

4 mm K2-ATP, 400 μm Na2-GTP, 10 mm Na2-phosphocreatine, and 10 mm HEPES buffer, adjusted to pH 7.3 with KOH. The recording electrode was connected to the recording circuit using a potassium chloride plated silver wire. The recording signal was digitized using a Digidata 1440A and amplified with a Multiclamp 700B (Molecular Devices,

Sunnyvale, CA). The signal was visualized and manipulated using Multiclamp 700B and

Clampex 10.1 software (Molecular Devices, Sunnyvale, CA).

The recording electrode was advanced into the tissue using a MPC-200 micromanipulator (Sutter Instrument Co., Novato, CA). Before entering the tissue, the difference in potential between the electrode solution and the aCSF was corrected and a positive pressure was applied to the electrode to prevent contamination of the tip by debris once it entered the tissue. Once a live cell was located, the electrode was moved onto the cell and the electrode’s positive pressure applied to the electrode was reversed to form a GΩ seal between the cell membrane and electrode. In voltage clamp mode, the holding voltage was dropped to -70 mV and a negative pressure quickly applied to the electrode solution in order to facilitate the electrode tip gaining access to the internal cell environment. Upon entering the cell, the series resistance dropped to MΩ levels.

Neurons were then distinguished as magnocellular, PA parvocellular or NE parvocellular neurons by their distinct electrophysiological characteristics (Tasker and 30

Dudek 1991, Hoffman et al. 1991, Luther et al. 2002). A current step protocol was applied to patch-clamped neurons in which they were alternatively injected with a negative pA pulse for 250 ms, to drop the membrane potential to approximately -80 mV, with a series of progressively more depolarizing pulses, increased in 10 pA increments, from 0 pA to 100 pA. Neurons were identified as magnocellular if they displayed a

+ damping of the membrane charging curve indicative of an A-type K (IA) current (Tasker and Dudek 1991, Luther et al. 2000). Neurons were identified as PA parvocellular if they displayed action potentials with the 0-30 pA depolarizing pulses indicative of a type T

Ca2+ current. Finally, neurons were classified as NE parvocellular if they displayed neither a damping of the membrane charging curve or low-threshold spikes (Hoffman et al. 1991, Luther et al. 2002).

Following a 5 minute recovery period, the brain slices containing the identified magnocellular or parvocellular neurons were perfused with aCSF containing 50 nM of human relaxin-2 or 50 nM of human relaxin-3 for 5 minutes. Recordings were analyzed by determining the average change in membrane resting potential during treatment, in

30 second intervals, from the average in the 30 second period preceding treatment.

Neurons were classified as responding to treatment if the average change in resting membrane potential during treatment was at least twice the standard deviation of the average in the 30 seconds preceding treatment. The average changes in membrane potential were compared between responding and non-responding neurons using the student’s t-test.

RESULTS

Experiment 1: The effect of relaxin-3 on the release of oxytocin and vasopressin from isolated PVN

Three different approaches were considered in the development of the in vitro release protocol. The protocol was optimized by considering the following for each approach: the concentration of hormone in effluent sample, whether the release of the hormones was consistent throughout trials, and finally whether tissue was capable of responding to treatment. The average concentrations of oxytocin in control buffer samples were 7.81+3.29 pg/mL, 44.68+4.83pg/mL and 39.98+2.03pg/mL for the perfusion aCSF system, static aCSF system and static HEPES buffer system (samples collected during period 3 and 4) respectively. All means are reported + standard error of the mean. The intra- and inter- oxytocin assay variations were 13.54 % and 15.10 % respectively.

Vasopressin concentrations were not evaluated for the perfusion aCSF system as the pilot study evaluating whether this preparation could sustain tissue revealed that oxytocin concentrations were low and declined significantly over time (see figure 1). The average concentrations of vasopressin in control buffer samples were 22.81+2.81pg/mL

(n=53) and 13.21+1.62 pg/mL (n=36) for the static aCSF system and HEPES buffer system respectively. The intra- and inter-vasopressin assay variations were 11.91 % and 12.95 % respectively.

Perfusion aCSF system

A pilot study was conducted to determine the validity of using PVN slices perfused with aCSF to measure the response of the PVN neurons to relaxin peptides.

The oxytocin concentrations of the effluent were standardized by calculating the hormone concentrations as a percentage difference from basal concentrations (3x10 minutes). The oxytocin concentrations were compared between slices treated with 50 nM human relaxin-2 or aCSF alone (3x10 minutes) using a two-way repeated measures

ANOVA. The test determined that oxytocin concentrations decreased over time in effluent collected from tissue treated with either 50 nM of human relaxin-2 or aCSF alone (figure 1, time p< 0.05). There was no effect of treatment (50 nM human relaxin-2 or control) on oxytocin concentrations (treatment p= 0.899, treatment x time p= 0.239).

Given the low concentrations of oxytocin in effluent samples (kit range of 15.6 – 1000 pg/mL, sensitivity of 11.7 pg/mL), the decrease of oxytocin concentrations over time, and an inability to maintain a consistent rate of perfusion throughout trials, the perfusion system was abandoned for a static system.

Static aCSF system

The second approach used to measure hormone release consisted of incubating whole PVN nuclei in aCSF. After a basal interval (3x5 minutes), nuclei were treated with human-3, R3/I5, at a concentration of 50 nM or 500 nM, or aCSF alone (4x5 minutes).

Nuclei from individual brains underwent treatment at the same concentration. The hormone concentrations of the incubations buffer were compared as percentage 33 difference from average basal concentration using a two-way repeated measure

ANOVA.

Figure 1 Oxytocin release from PVN slices perfused with human relaxin-2 PVN slices were perfused with aCSF (1.5 mL/minute) alone for 15 minutes before treatment with 50 nM human relaxin-2 (n=4, triangles) or aCSF alone (n=4, circles) for 20 minutes (bar on x-axis). The average oxytocin concentrations during the control and treatment trials were 5.73 + 2.05 pg/mL and 5.75 + 3.12 pg/mL respectively. The release of oxytocin decreased significantly over time irrespective of treatment (treatment p=0.899, time p<0.05, treatment x time p= 0.239). The results shown are means + SEM.

Oxytocin

Figure 2 shows the effect of human relaxin-3 and R3/I5 on the release of oxytocin from the PVN of rats. Neither human relaxin-3 nor R3/I5 had an effect on oxytocin release, at either 50 nM (figure 2A, n=3 for each treatment group, treatment p= 0.472, time p= 0.177, treatment x time p=0.675) or 500 nM treatment concentrations

(figure 2B, n=6 for each treatment group, treatment p=0.620, time p< 0.05, treatment x 34 time p=0.878). There was some fluctuation in oxytocin release at both treatment concentrations, however there was only a significant decrease in oxytocin release across the 500 nM concentration trials (time p< 0.05).

Vasopressin

Figure 3 shows the effect of human relaxin-3 and R3/I5 on the release of vasopressin from the PVN of rats. As was found for oxytocin release, neither human relaxin-3 nor R3/I5 had an effect on the release of vasopressin at 500 nM treatment concentrations (figure 3B, relaxin-3: n=3, R3/I5: n=3, control: n=4, treatment p= 0.829, time p=0.673, treatment x time p=0.736).

Unlike what was found for oxytocin release, R3/I5 at a concentration of 50 nM had a significant effect on the release of vasopressin from the rat PVN. Vasopressin concentrations in the 50 nM trials failed the assumption of normality so the changes in concentration across time (for both basal and treatment intervals) were evaluated separately by treatment group using the Friedman’s test.

R3/I5 induced a significant increase in the release of vasopressin at 20 minutes of treatment (figure 3A, n=4, p<0.05). Although 50 nM of human relaxin-3 appeared to reduce the release of vasopressin, this response was not significant (figure 3A, n=3, p=

0.249). There was some fluctuation in vasopressin release, as was seen for oxytocin release, however there was no change in the release of vasopressin in control trials over time (figure 3A, n=4, p=0.297). 35

Figure 2 No effect of human relaxin-3 or R3/I5 on oxytocin release from the PVN The nuclei of the PVN were incubated in aCSF that was exchanged every 5 minutes. After 15 minutes, the nuclei were incubated with the following (bar on x-axis): human relaxin-3 (closed circles), R3/I5 (closed diamonds), at a concentration of (A) 50 nM (n=3) or (B) 500 nM (n=6), or in aCSF alone (open circles). At 50 nM treatment concentrations, the relaxin peptides had no effect on oxytocin release (treatment p= 0.472). Similarly, at 500 nM treatment concentrations, the relaxin peptides had no effect on oxytocin release (treatment p= 0.620), but there was a significant decrease in the release of oxytocin concentrations over time for all the treatment groups including for the control (P<0.05). The results shown are the means + SEM. 36

Figure 3 The effect of human relaxin-3 and R3/I5 on vasopressin release from the pvn The nuclei of the PVN were incubated in aCSF that was exchanged every 5 minutes. After 15 minutes, the nuclei were incubated with the following (bar on x-axis): human relaxin-3 (closed circles), R3/I5 (closed diamonds), at a concentration of (A) 50 nM (n=3, 4) or (B) 500 nM (n=3, 3), or in aCSF alone (n=4, 4; open circles). Although human relaxin-3 appeared to reduce the release of vasopressin at a 50 nM treatment concentration, this trend was not significant (p= 0.249). R3/I5, on the other hand, significantly increased the release of vasopressin with 20 minutes of treatment (P<0.05). Neither human relaxin-3 nor R3/I5 had an effect on vasopressin release at a 500 nM treatment concentration (p=0.829). The results shown are the means + SEM.

Static HEPES buffer system

The third and final approach used to measure the release of the hormones involved incubating the tissue from younger rats in HEPES buffer. In this system, PVN were challenged with high potassium concentrations before and after treatment in order to evaluate the viability of the tissue, as well as to confirm the tissue’s responsiveness. As is seen by the peaks in release in figures 4 and 5, high potassium concentrations induce the release of oxytocin and vasopressin. In order for a trial to be included in analysis, the isolated PVN had to respond to these potassium challenges with increases in hormone release.

The treatment concentrations were compared to hormone concentration buffer collected in the 15 minutes prior to treatment (period 4), the basal concentrations. The effect of treatment on hormone release was determined by statistically comparing the treatment concentrations either to all three basal concentrations or to the average of the basal concentrations. As there was no difference in the outcomes of the methods of comparison, the former analysis is presented in the remainder of the thesis.

Oxytocin

Figure 4 shows the effect of the relaxin peptides on the release of oxytocin from the PVN of young rats. Human-relaxin-3 significantly reduced the release of oxytocin from the PVN (figure 4A, n=3, p<0.01). Although human relaxin-3 appeared to reduce oxytocin concentration with 15 minutes of treatment, oxytocin concentrations were only significantly different from basal concentrations after 25 and 30 minutes of 38 treatment (p<0.05). The mean percentage decrease from average basal concentration was 44.77 + 1.64 % at 25 minutes and 34.48 + 5.50 % at 30 minutes of human relaxin-3 treatment (table 1).

In order to evaluate whether relaxin-3 mediates this effect through RXFP1 or

RXFP3, PVN were also treated with human-relaxin-2 and R3/I5. R3/I5 significantly reduced the release oxytocin from PVN of young rats (figure 4B, n=3, p<0.05), while comparatively human-relaxin-2 induced a significant increase in oxytocin release from the PVN (figure 4B, n=3, p<0.05). R3/I5 appeared to reduce oxytocin release between 10 to 15 minutes of treatment, however Post hoc comparisons did not reveal any significant differences in oxytocin concentrations between basal and R3/I5 treated samples. Human relaxin-2 induced an increase in oxytocin release after 5 minutes of treatment, with an average increase of 96.31 + 15.62 % from average basal concentrations (table 1). There was no change in oxytocin concentrations across time in control trials (figure 4A, n=6, p=421).

Vasopressin

Figure 4 shows the effect of the relaxin peptides on the release of oxytocin from the PVN of young rats. Although there was some variation in release of vasopressin in the control trials, these changes in vasopressin concentration were not significant

(figure 5A, n=4, p=0.625). There was no apparent effect of human-relaxin-3 on the release of vasopressin from the PVN (figure 5A, n=3, p=0.394), however, as was found in the static aCSF study, there was a significant increase in the release of vasopressin from isolated PVN after long term treatment with R3/I5 (figure 5B, n=3, p<0.05). Vasopressin

Figure 4 The effects of relaxin peptides on oxytocin release from the PVN of young rats After the pvn of young rats were challenged with high potassium concentrations (grey bars) to evaluate tissue viability, they were incubated in HEPES buffer alone (A, n=5, open circles), or in 50 nM of human relaxin-3 (A, n=3, filled circles), R3/I5 (B, n=4, filled diamonds), or human relaxin-2 (B, open diamonds, n=3) for 30 minutes (black bar). Both human relaxin-3 and R3/I5 significantly reduced oxytocin release (p<0.05), while human relaxin-2 significantly increased oxytocin release (p<0.05). There was no change in oxytocin release from PVN incubated in HEPES buffer alone (p=0.425). The results shown are the means + 40

SEM.

Figure 5 The effects of relaxin peptides on vasopressin release from the PVN of young rats After, the pvn of young rats were challenged with high potassium concentrations (grey bars) to evaluate tissue viability, they were incubated with HEPES buffer alone (A, n=4, open circles) or in 50 nM of human relaxin-3 (A, n=3, filled circles), R3/I5 (B, n=3, filled diamonds), or human relaxin-2 (B, n=3, open diamonds) for 30 minutes (black bar). Neither human relaxin-3 nor human relaxin-2 had an effect on vasopressin release (p=0.394, 0.659 respectively). R3/I5, however, significantly induced vasopressin release following 30 minutes of treatment (p<0.05). The results shown are the means + SEM. 41

Table 1 Change in oxytocin and vasopressin release from isolated PVN incubated in Hepes buffer with relaxin peptides After an initial potassium challenge to evaluate viability, isolated PVN were allowed to recover before treatment with hepes buffer alone, 50 nM human relaxin-3, 50 nM R3/I5, or 50 nM human relaxin-2 for 30 minutes. Hormone concentrations were standardized across trials by dividing by the change in hormone concentration during the initial potassium challenge (period 2/period1). The hormone concentrations in each treatment group were compared as a percentage change from average hormone concentration in the 15 minute interval preceding treatment (period 4). * indicates p<0.05. The results reported are the means + SEM. Change From Average Basal Concentration (%) Average Basal Treatment Time (min) Concentration Treatment (pg/mL) 5 10 15 20 25 30 Oxytocin

Control 17.73 + 0.97 5.78 + 10.30 40.31 + 43.11 24.74 + 22.59 - 11.56 + 12.19 26.96 + 18.51 24.80 + 22.36 (n=5) 15.63 + 1.71 15.63 + 1.17 24.17 + 8.46 -6.18 + 14.97 - 44.77 + 1.64 * -34.48 + 5.50 * -13.57 + 20.99 Human Relaxin-3 (n=3) 25.34 + 6.12 14.27 + 20.81 -26.31 + 15.92 -43.30 + 10.18 -25.02 + 10.18 -7.82 + 25.35 16.73 + 25.94 R3/I5 (n=3) 17.37 + 2.00 96.31 + 15.62 * 27.13 + 19.07 55.43 + 27.42 22.36 + 16.12 4.99 + 5.43 9.96 + 23.50 Human Relaxin-2 (n=3) Vasopressin

Control 5.80 + 1.78 15.36 + 26.79 49.19 + 50.79 49.38 + 48.96 25.15 + 18.54 40.37 + 28.99 98.39 + 70.84 (n=4)

Human Relaxin-3 10.36 + 4.61 -0.80 + 3.94 24.25 + 3.99 16.84 + 14.65 35.96 + 32.93 8.01 + 15.64 -16.54 + 55.70 (n=3)

R3/I5 9.72 + 1.73 18.32 + 20.24 -21.98 + 16.86 -22.59 + 17.55 -17.14 + 21.68 -12.94 + 23.06 240.25 + 147.63 * (n=3)

Human Relaxin-2 12.40 + 0.48 -0.990 + 6.16 19.13 + 18.48 3.90 + 20.51 -8.79 + 18.14 -22.42 + 16.43 -12.67 + 13.58 (n=3) 42

concentrations increased by an average of 291.27 + 189.61 % at 30 minutes of R3/I5

treatment (table 1). Human relaxin-2, on the other hand, had no effect on vasopressin

release from the PVN of young rats (figure 5B, n=3, p=0.659).

Experiment 2: The effect of relaxin peptides on the intracellular electrical activity of isolated neurons from the PVN in vitro

The effect of relaxin and relaxin-3 on the membrane potential of magnocellular neurons

Magnocellular neurons (n=15) were identified by their large size (approximately

30 µm), location in the PVN (magnocellular subdivisions), and by their expression of a

prominent transient potassium current (IA), visualized as an outward rectification

following injection with a hyperpolarizing current (see figure 6A). The magnocellular

neurons had a mean membrane potential of -53.85±5.12 mV, mean action potential amplitude of 77.56+3.71 mV, and mean input resistance of 368.18 ± 39.40 mV. Six

magnocellular neurons were treated with 50 nM human relaxin-2 in order to evaluate

the electrical response of the neurons to relaxin. The treatment concentration was

chosen in order to maintain consistency with the in vitro release experiment; relaxin has

been shown to have biphasic response with concentration in other systems (Sherwood

2004). Neurons were classified, according to the criteria used by similar studies (Hoyda et al. 2009, Price et al. 2009), as responding to treatment if the average change in membrane potential during the 5 minute treatment period was at least 2 times the standard deviation of the average membrane potential in 30 second period preceding treatment. 43

-62 mV

-61 mV

-78 mV

20 mV 100ms

Figure 6 Identification of neuron types within the PVN by the expression of low threshold spikes or outward rectification Neuron types were identified by their response to injections of positive current following hyperpolarization. Magnocellular neurons exhibited transient outward rectification visualized as delay to action potential (top; see arrow). Pre-autonomic parvocellular neurons exhibited low threshold spikes with current injection from 0 to 30 mV (middle). NE parvocellular neurons exhibited neither a delay to action potential or low threshold spikes (bottom).

A A B

Figure 7 The effects of human relaxin-2 and human relaxin-3 on the membrane potential of magnocellular neurons in the paraventricular nucleus Patch-clamped magnocellular neurons treated with 50 nM human-relaxin-2 and human relaxin-3 for 5 minutes showed heterogeneous responses. (A) Magnocellular neurons responded to human relaxin-2 by depolarizing (black points), with changes in membrane potential that were significantly (p<0.05) greater from neurons that showed no response to human relaxin-2 (grey points). Magnocellular neurons responded to human relaxin-3 by hyperpolarizing (white points), with changes in membrane potential that were significantly (p<0.05) greater from neurons that showed no response to human relaxin-3 (grey points). (B) Examples of depolarization (top) and hyperpolarisation (bottom) of single magnocellular neurons induced by human relaxin-2 and human relaxin-3 respectively over the 5 minute treatment period (overhead bar).

Magnocellular neurons responded to 50 nM human relaxin-2 with either increases in membrane potential (n=3) or with no response (n=3) (figure 7). The average increase in membrane potential with human relaxin-2 treatment was

3.23+1.65 mV, which was significantly larger than the average of the non-responding neurons, 0.04+0.52 mV (p<0.05).

The electrical response of magnocellular neurons to human relaxin-3 was also evaluated. Of the 8 magnocellular neuron treated with 50 nM human-relaxin-3, five responded with decreases in membrane potential and three showed no response (figure 45

7). The average decrease in membrane potential with human relaxin-3 treatment was -

6.08+2.52 mV, which was significantly smaller than the average of the non-responding

neurons, 0.22+1.20 mV (p<0.05).

The effect of relaxin and relaxin-3 on the membrane potential of NE parvocellular neurons

Parvocellular neurons (n=18) were identified by their location (parvocellular divisions) as well as by their lack of low threshold or delayed action potentials following injection of hyperpolarizing current (figure 6c). The parvocellular neurons had a mean membrane potential of -62.69 ± 7.35 mV, mean action potential amplitude of 70.86 ±

5.55 mV, and mean input resistance of -272.46 ± 41.03 mV.

The response of the NE parvocellular neurons to relaxin and relaxin-3 was

evaluated. Of the eight NE parvocellular neurons treated with 50 nM human relaxin-2,

three responded with increases in membrane potential while five showed no response

(figure 8). The average increase in membrane potential with human relaxin-2 treatment

was 7.63 + 2.19 mV, which was significantly larger than the average of the non-

responding neurons, 0.57 + 1.47 mV (p<0.01). Of the ten NE parvocellular neurons

treated with 50 nM human relaxin-3, three responded with increases in membrane

potential, three with decreases in membrane potential and four showed no response

(figure 8). The average increase in membrane potential with human relaxin-3 treatment

was 5.95 + 2.65 mV, which was significantly larger than the average of the non-

responding neurons, -0.59 + 1.26 mV (p<0.01). The average decrease in membrane 46 potential with human relaxin-3 treatment was -3.95 + 2.18 mV which was significantly smaller than the average of the non-responding neurons (p<0.05).

A B

Figure 8 The effect s of human relaxin-2 and human relaxin-3 on the membrane potential of neuroendocrine parvocellular neurons in the paraventricular nucleus Patch-clamped NE parvocellular neurons treated with 50 nM human-relaxin-2 and human relaxin- 3 for 5 minutes also showed heterogeneous responses. (A) NE parvocellular neurons responded to 50 nM human relaxin-2 by depolarizing (p<0.001; black points), with changes in membrane potential that were greater than those of neurons that did not respond to human relaxin-2 (grey points). Human-relaxin-3 had a heterogeneous effect on NE parvocellular neurons. NE parvocellular neurons responded to human relaxin-3 by depolarizing (black points) and hyperpolarizing (white points); the changes in membrane potential of responding neurons were significantly different than those neurons that showed no response to human relaxin-3(p<0.01, p<0.001 respectively). (B) Example of depolarization of a NE parvocellular neuron by human relaxin-2 (top) as well as examples of depolarization (middle) and hyperpolarization (bottom) of NE parvocellular neurons by human relaxin-3 over the 5 minute treatment period (overhead bar).

DISCUSSION

This study was undertaken to evaluate the actions of the relaxin peptides, relaxin

and relaxin-3, in regulating the somatodendritic release of oxytocin and vasopressin

within the PVN, and in regulating the electrical activity of neuroendocrine neurons of the PVN. In its role as a hormone of reproduction, relaxin that originates in the peripheral circulation acts centrally to regulate the release of oxytocin and vasopressin from the brain (Geddes and Summerlee 1995; Summerlee et al. 1998a). It has yet to be established if relaxin can also act directly on the magnocellular nuclei to regulate oxytocin and vasopressin release. This study investigated the effect of relaxin and relaxin-3 on the activity of magnocellular nuclei in the PVN, and demonstrated that relaxin and relaxin-3 regulate the somatodendtritic release of oxytocin and vasopressin in this brain region in rats. Relaxin and relaxin-3 appear to regulate the somatodendtritic release of oxytocin and vasopressin by acting on receptors located within the PVN, as the relaxin peptides regulated the release of the hormones when the PVN was isolated from its afferent projections. Relaxin and relaxin-3 were also demonstrated to regulate the electrical activity of neuroendocrine neurons in the PVN in vitro, including magnocellular neurons.

Peripheral relaxin has a dual effect on the release of oxytocin from the brain, inhibiting reflex milk-ejection by inhibiting the pulsitile release of oxytocin (Summerlee et al. 1984, Summerlee et al. 1998a), while also increasing the basal release of oxytocin as well as vasopressin in lactating, early pregnant and non-pregnant rats (Way and Leng

1992, Parry et al. 1994, Summerlee et al. 1998a). Evidence suggests peripheral relaxin 48

regulates the release of oxytocin and vasopressin by acting through the

circumventricular organs that are exempt from the blood-brain barrier (Mumford et al.

1989; Parry et al. 1991; 1994; Geddes and Summerlee 1995).

Relaxin is also expressed in the brain, as is relaxin’s receptor, RXFP1 (Osheroff

and Ho 1993, Burazin et al. 2005, Piccenna et al. 2005, Ma et al. 2006). Relaxin-3 is

produced centrally as well, and binds RXFP1 (Bathgate et al. 2002a, Sutton et al. 2004,

Ma et al. 2007; 2009, Smith et al. 2010, Silvertown et al. 2010). Presumably centrally

produced relaxin can also act through the circumventricular organs to regulate oxytocin

and vasopressin release, given that the subfornical organ lies adjacent to regions that

express relaxin mRNA, including the paraventricular thalamic area and dendrate gyrus

(Ma et al. 2006). A study by Summerlee and colleagues (1987) found that administering relaxin centrally inhibited reflex milk-ejection, which was negated by lesions to the

subfornical organ.

The PVN and SON express RXFP1 mRNA and receive projections from the arcuate

nucleus that produces relaxin (Burazin et al. 2005, Tanaka et al. 2005, Ma et al. 2006),

suggesting relaxin could also regulate oxytocin and vasopressin release by acting on receptors in these brain regions. The PVN and the SON also express RXFP3 mRNA,

relaxin-3’s other receptor, and receive projections from the nucleus incertus that

produces relaxin-3 (Bathgate et al. 2002a, Sutton et al. 2004, Ma et al. 2007), suggesting relaxin-3 could also regulate oxytocin and vasopressin release by acting at one or both of the receptors in the PVN and SON. 49

The PVN has been a focus of studies investigating the neuroendocrine role of

relaxin-3 (Watanabe et al. 2011). The challenge of examining the role of relaxin-3 and

relaxin in the PVN is isolating the actions of the peptides acting in the PVN from the actions of the peptides acting through the diverse brains regions that project to the PVN that express RXFP1 (arcuate nucleus, subfornical organ, lateral hypothalamus), RXFP3

(bed nucleus of the stria terminalis, dorsal medial nucleus of the hypothalamus, dorsal raphe nuclei, suprachiasmatic nucleus, nucleus incertus), or both receptors (nucleus of the solitary tract, medial preoptic area) (Sutton et al. 2004, Burazin et al. 2005, Ma et al.

2006; 2007; 2009, Silvertown et al. 2010, Smith et al. 2010). Therefore, the objective of the present study was to develop in vitro models in which the direct effects of relaxin and relaxin-3 in the PVN could be examined.

Two techniques were employed to examine the effect of centrally acting relaxin and relaxin-3 on PVN activity: (1) The PVN was dissected free from its efferent projections, and maintained in vitro to evaluate the effects of the relaxin and relaxin-3

acting in the PVN on the somatodendritic release of oxytocin and vasopressin, and (2)

intracellular electrophysiology was employed using brain slices in which neuroendocrine neurons, identified by their response to hyperpolarizing current (Tasek and Dudek 1991;

2000, Hoffman et al. 1991, Luther et al. 2000; 2002, Stern 2001), were treated with relaxin and relaxin-3 to evaluate their electrical response to the relaxin peptides. The treatment concentration used in both studies was 50 nM, a concentration chosen because prior studies have found that 10 nM concentrations of relaxin-3 can affect the release of hormones in the PVN (McGowan et al. 2006; 2008). 50

Unlike the in vitro release study, the electrophysiology study is an indirect measure of the effect of the relaxin peptides on the release of oxytocin and vasopressin.

The frequency of action potentials arriving at the terminals of magnocellular terminals determines the amplitude of oxytocin and vasopressin release from the posterior pituitary (Dayanithi et al. 2012, Leng et al. 1999). The electrophysiology study was therefore an indirect method to evaluate the effect of the relaxin peptides on the release of oxytocin and vasopressin from the terminals of magnocellular neurons in the

PVN.

When the PVN was dissected free of its afferent projections and maintained in

vitro, relaxin appeared to induce the release of oxytocin from isolated PVN. In contrast,

relaxin-3 at the same concentration appeared to reduce the release of oxytocin in the

PVN, indicating a novel action of relaxin-3. The RXFP3 agonist, R3/I5, also reduced the

release of oxytocin from the PVN, indicating relaxin-3 may act at the RXFP3 receptor to

mediate its apparent effects on somatodendritic oxytocin release in the PVN.

Relaxin-3 appeared to have no effect on the release of vasopressin from isolated

PVN, and neither did relaxin. However, the RXFP3 agonist, R3/I5, appeared to induce

the release of vasopressin from isolated PVN. R3/I5 is a hybrid peptide, consisting of the

B-chain of the relaxin-3, necessary to bind and activate RXFP3 (Lui et al. 2003, Hossain et

al. 2008), and the A-chain of INSL5, without the secondary binding site necessary to

activate RXFP1 (Lui et al. 2003). As R3/I5, but not relaxin-3, induced the release of

vasopressin, RXFP3 activation appears to induce the release of vasopressin in the 51

absence of RXFP1 binding, suggesting RXFP1 interferes with RXFP3 signalling on

vasopressin neurons.

Given that human relaxin-3 binds RXFP1 with lower affinity than RXFP3 in rats

(Lui et al. 2003a), if RXFP1 inhibits RXFP3 activity on vasopressin neurons, then relaxin-

3’s function in the PVN may depend on the magnitude of its release, as well as the availability of each of the relaxin receptors on vasopressin neurons. RXFP1 and RXFP3 appear to have opposing effects on oxytocin release in the PVN, creating a similar situation in which the function of relaxin-3 in the PVN could depend on the concentration of relaxin-3 and availability of the receptors. Although this may seem paradoxical, a bell shaped response has been reported for relaxin in its function of collagen remodelling: human dermal and lung fibroblasts have been found to exhibit a dose-dependant increase in the expression of metalloprotinease-1 in response to relaxin concentrations only up to 10 and 100 ng/mL concentrations respectively, with a decrease in expression at higher concentrations in vitro (Unemori and Amento 1990,

Unemori et al. 1996).

As hormone release from the soma and dendrites of magnocellular neurons can occur independent from their terminal release (see Ludwig and Leng 2006 for review), the present study also investigated the electrical response of neuroendocrine neurons of the PVN to relaxin and relaxin-3. Way and Leng (1992) examined the effect of peripheral relaxin on the firing rate of magnocellular neurons of the SON in vivo using extracellular electrophysiology, and found that intravenously administered relaxin increased the frequency of action potentials of oxytocin and vasopressin neurons, an 52 effect that is presumably mediated by relaxin acting through the circumventricular organs (Geddes and Summerlee 1995). By using an in vitro electrophysiology approach in the present study, the effect of the peptides acting through the circumventricular organs was eliminated through dissection. Any effect of relaxin and relaxin-3 on the electrical activity of neuroendocrine neurons would be due to the peptides acting in brain regions that were maintained in the brain slices. The brain slices included the peri-

PVN and the PVN itself, as well as hypothalamic nuclei in the immediate vicinity of the

PVN, such as the arcuate nucleus, suprachiasmatic nucleus, the lateral hypothalamus and the medial preoptic area. The neuroendocrine neurons of the PVN were identified by their responses to a hyperpolarization, with magnocellular neurons responding to injections of current following hyperpolarization with delayed action potentials, and neuroendocrine parvocellular neurons, the neurons that project to anterior pituitary, responding to hyperpolarization with neither a delay to action potential nor low threshold spikes that are characteristic of non-neuroendocrine parvocellular neurons

(Tasek and Dudek 1991; 2000, Hoffman et al. 1991, Luther et al. 2000; 2002, Luther and

Tasker 2000, Stern 2001).

Relaxin depolarized magnocellular neurons, which suggests relaxin may increase the firing rate of these neurons. Relaxin has been indicated as having an effect on the terminal release of oxytocin and vasopressin release from isolated neural lobes of rats in vitro, in other words, from magnocellular neurons terminals dissected free of their neurons (Dayanthni et al. 1987). Under these conditions, relaxin inhibited the basal release of oxytocin and vasopressin while potentiating electrically stimulated release. A 53

similar study using neural lobes from lactating rats, however, found no effect of relaxin

on the electrically stimulated release of the magnocellular hormones (O’Bryne et al.

1986), which is possibly due to a reduction in response of magnocellular terminals to

relaxin during lactation (Geddes and Summerlee 1995). Studies have yet to detect

relaxin binding or RXFP1 mRNA in the pituitary of rats (Bathgate et al. 2013), so the

mechanism by which relaxin would induce the release of the hormones from the pituitary remains unknown. The finding that relaxin can depolarize magnocellular

neurons suggests, however, that relaxin may also be able to act in the brain to regulate the peripheral release of magnocellular hormones by affecting the frequency of action potentials arriving at the magnocellular neuron terminals in the posterior pituitary. In contrast to relaxin, relaxin-3 hyperpolarized magnocellular neurons, which would

suggest relaxin-3 decreases the firing rate of magnocellular neurons. As activation of

RXFP1 depolarized magnocellular neurons, and as relaxin-3 binds to and activates both

RXFP1 and RXFP3 (Lui et al. 2003a), it is surprising that relaxin-3 hyperpolarized

magnocellular neurons. This effect, along with the potential interaction between RXFP3

and RXFP1 in the regulation of vasopressin release, if any, requires further investigation.

The effects of the relaxin peptides on the electrical activity of NE parvocellular

neurons in the PVN were also investigated. The role of relaxin-3 in the PVN has been a

focus of research investigating the central function of relaxin-3. It has been shown to

increase c-fos immunoreactivity of CRH and vasopressin neurons in the parvocellular

regions of the PVN (Watanabe et al. 2011). Relaxin-3 has also been shown to increase

plasma release of luteinizing hormone (McGowan et al. 2008) and reduce the release of 54

plasma thyroid stimulating hormone (McGowan et al. 2006). Relaxin-3 both depolarized

and hyperpolarized NE parvocellular neurons in this study. Relaxin, on the other hand,

depolarized NE parvocellular neuron, which would suggest relaxin has neuroendocrine

functions in the PVN other than regulating oxytocin and vasopressin release.

The regulation of somatodendritic release in the PVN

The mechanisms of somatodendritic release in the HNS are not well understood

(Dayanithi et al. 2012). At the axon terminals of magnocellular neurons, exocytosis of

the large dense cored vesicles (LDCVs) containing oxytocin and vasopressin is initiated

by the entrance of extracellular calcium through voltage gated channels opened by the

arrival of action potentials (Leng et al. 1999). In contrast, somatodendritic release

appears to be dependent on the mobilization of intracellular calcium sources (Ludwig and Leng 2006).

Both oxytocin and vasopressin induce their own somatodendritic release by liberating internal calcium stores (Lambert et al. 1994, Ludwig et al. 2002, Ludwig et al.

2005, Ludwig and Leng 1997). Somatodendritic release of these hormones is also induced by thapsigargin, a calcium-ATPase inhibitor that discharges calcium from

endoplasmic reticulum stores (Ludwig et al. 2002, Ludwig et al. 2005, Ludwig and Leng

1997). Somatodendritic oxytocin release appears to occur independently of the

electrical status of oxytocin neurons (Ludwig and Leng 2006). Voltage-operated calcium

channels can induce somatodendritic oxytocin release if the release is primed (Ludwig

and Leng 2006). Priming involves increasing the availability of releasable LDCVs through 55

mechanisms that are not well understood but appear to involve increases in intracellular

calcium concentration (Ludwig et al. 2002, Ludwig et al. 2005, Ludwig and Leng 2006).

Unlike oxytocin, somatodendritic release of vasopressin is partially dependent on voltage-operated calcium channels (Sabateir et al. 1997, Ludwig et al. 2005). In the in vitro somatodendtritic release preparation, relaxin failed to affect vasopressin release from PVN. The PVN was isolated from its efferent connections in the preparation, therefore the possibility cannot be excluded that relaxin can act through other brain regions to change the electrical activity of vasopressin neurons, and thereby somatodendritic release of vasopressin, in the PVN.

Oxytocin induces increases in intracellular calcium of oxycocin neurons by activating phospholiase C, which increases levels of inositol trisphosphate (IP3) and diacyl glycerol (DAG). This then liberates intracellular calcium from endoplasmic reticulum stores (Dayanithi et al. 2000). Vasopressin induces intracellular calcium release in the same manner as oxytocin, but also by increasing cAMP concentrations to activate protein kinase A (Dayanithi et al. 2000).

Treatment of dissociated hypothalamic neurons with INSL5, another relaxin

peptide family member, has been shown to induce thapsigargin-sensitive increases in

intracellular calcium in calcium free media, with RXFP4 coupling to Gα16 in order to

induce mobilization of intracellular calcium (Dun et al. 2006). So far there is no evidence

RXFP3 can also couple to Gα16 (Bathgate et al. 2013). Given the finding that relaxin-3, as

well as R3/I5, reduced the somatodendritic oxytocin release in the PVN, activation of

RXFP3 on oxytocin neurons may instead inhibit calcium signalling pathways. 56

Alternatively, RXFP3 signalling may inhibit somatodendritic oxytocin release by disrupting actin remodelling. The plasma membranes of magnocellular neurons are lined by an internal F-actin framework, which is disrupted during potassium stimulated hormone release (Tobin and Ludwig 2007). Unlike release at terminal axons, somatodendritic release does not occur at particular active zones of the membrane

(Pow and Morris 1989), rather F-actin is seen as acting to scaffold exocytosis (Tobin et al. 2012). The role of F-actin in regulating oxytocin neuron activity is dynamic (Wang and

Hatton 2007): when magnocellular neurons of the SON were treated with oxytocin, they exhibited an initial sub-cortical actin ring formation (by 5 min), which was followed by a decrease in actin expression upon bursting behaviour in vitro and milk-ejection in vivo (by 30 min); these changes in actin expression coincided with increases and decreases in the phosphorylation of ERK 1/2 respectively. Tobin and Ludwig (2007) found that treating the SON with agents to depolymerise or polymerize F-actin acted to inhibit thapsigargin induced priming of oxytocin release, indicating a role for the f-actin remodelling in regulating priming (Tobin and Ludwig 2007). Studies have yet to examine the role of pERK 1/2 in the regulation of somatodendritic release (Tobin et al. 2012).

Given that activation of RXFP3 increases the phosphorylation of ERK ½, relaxin-3 may prevent actin remodelling by maintaining pERK ½ levels when they would otherwise decrease and resulting in the inhibition of oxytocin autoregulatory activity. There is some evidence relaxin-3 can affect actin remodelling in astrocytes; astrocytes treated with relaxin, relaxin-3 or R3/I5 in vitro exhibited changes in their F-actin arrangement

(Willcox 2013). 57

Another cell type on which oxytocin acts to increase intracellular calcium

concentrations is myometrial cells (Anwer et al. 1989). As is seen in oxytocin neurons of

the HNS, oxytocin acts on its receptor on myometrial cells to increase IP3 and DAG

production, liberating calcium from intracellular stores which initiates contraction

(Anwer et al. 1989). Relaxin inhibits oxytocin induced myometrial cell contraction by inhibiting IP3 and DAG production through PKA activation (Anwer et al. 1990).

Somatodendritic release of oxytocin is essential for the coordinated burst of

oxytocin neuron activity that drives reflex milk-ejection (Lambert et al. 1993). Centrally

administered relaxin inhibits reflex milk-ejection (Summerlee et al. 1998a), which

suggests that relaxin would inhibit the somatodendritic release of oxytocin in the PVN.

The findings of this study, however, indicate that relaxin may instead induce

somatodendritic oxytocin release in the PVN. Relaxin may act through pre-synaptic

connections to regulate oxytocin neuron electrical activity, however the findings of this

study are that relaxin instead depolarizes magnocellular neurons in the PVN. The effect

of relaxin in other systems is biphasic; RXFP1 activates different signalling pathways at

high (nanomolar) and low (picomolar) relaxin concentrations, with the liberation of PKA

only occurring at high concentrations (Bathgate et al. 2013). Relaxin could have a

biphasic effect on the somatodendritic release of oxytocin in the PVN, inhibiting the

release of oxytocin at concentrations higher than was considered in the present study.

The function of Relaxin-3

Relaxin-3 has been indicated in an array of central functions including appetitive behaviour (McGowan et al. 2005; 2007; 2008) and stress response (Tanaka et al. 2005;

2010, Banerjee et al. 2010, Watanabe et al. 2011), and these functions appear to be affected in part by relaxin-3 actions on or in the PVN. The PVN is an effector site for regulating appetive behaviour (Schwartz et al. 2000), producing neuropeptides which suppress food intake when released centrally, including TRH, CRH and oxytocin

(Schwartz et al. 2000, Arase et al. 1988, Suzuki et al. 1982). Relaxin-3 induces increases c-fos expression of CRH neurons in the PVN (Watanabe et al. 2011), but the effect of relaxin-3 on the central release of CRH has yet to be investigated. It inhibits the central release of thyroid-releasing hormone (McGowan et al. 2006). Relaxin-3 may also regulate appetitive behaviour by inhibiting somatodendritic oxytocin release in the PVN.

Relaxin-3 is involved in stress response, though its role in it remains unclear

(Bathgate et al. 2013). The nucleus incertus has been suggest as being involved in controllable (escapable) rather than uncontrollable (inescapable) stress, due to the finding that rats undergoing forced swim exhibited more neural activation in the nucleus incertus if prior trials included an escape platform (Goto et al. 2001). Relaxin-3 immunoreactivity increases following forced swim stress (Tanaka et al. 2010). A recent study examined the role of relaxin-3 in regulating behaviour during forced swim, finding that a RXFP3 agonist decreased the time rats spent immobile if they had undergone prior stress tests (Ryan et al. 2013). Forced swim stress induces oxytocin release within 59

the PVN and SON (Wotjak et al. 1998; 2001). Though it needs to be re-examined in a

forced swim model, the findings of the present study support this model as relaxin-3

reduced the release of oxytocin in the PVN. Also, as central administration of an

oxytocin antagonist into the PVN increases plasma ACTH concentrations in rats during

forced swim stress (Neumann et al. 2000), relaxin-3 may also regulate plasma ACTH

release by inhibiting oxytocin release in the PVN.

Technical considerations

The in vitro somatodendritic release protocol used in this study was developed by the investigator. The development of the protocol was beset with some technical difficulties such that three versions of the protocol were developed. When the in vitro

hormone release protocol was originally developed, it consisted of a modified

electrophysiology protocol in which tissue was continuously perfused with aCSF. The

challenge with adapting an electrophysiology protocol to investigating hormone release

is maintaining viability when the perfusion rate is decreased to allow for detectable

concentrations of the released hormones. This is particularly difficult for quantifying

oxytocin and vasopressin since they are released from the PVN at low picograms levels

in the absence of electrical or ionic stimulation. A pilot experiment was conducted to evaluate the efficacy of the perfusion protocol as a measure of hormone release in the

PVN. The average oxytocin concentrations of the perfusant, which were low to begin with, decreased significantly over the evaluation period, indicating that there were problems with maintaining tissue viability. 60

Similar studies used a static incubation system to evaluate the effect of relaxin-3

on the release of hormones in the PVN (McGowan et al. 2006; 2008). The protocol used

of the McGowan studies was adopted, but modified by changing the aCSF every 5

minutes, instead of every 20 minutes as reported in the McGowan studies, in order to prevent confounding oxytocin and vasopressin autoregulation (Ludwig et al. 2006).

With this static protocol, oxytocin and vasopressin were released at quantifiable concentrations from PVN nuclei. The hormones concentrations detected were reasonably stable over time in control trials. This suggested efficacy of the protocol in measuring induced changes in the hormone release. Surprisingly, there were no apparent changes in oxytocin and vasopressin release in response to relaxin-3 treatment

when evaluated with the static aCSF protocol. This was contradicted by subsequent

work.

Though the tissue containing oxytocin and vasopressin neurons may be viable in

terms of survival, the conditions may not be optimal for neurons to be capable of

responding to stimulation in vitro. As indicated by a study conducted by Widmer and

colleagues (2003), the ability of oxytocin neurons in the SON to respond to high potassium concentrations decreases with age in rats. In light of that study, a further static incubation experiment was conducted in which brains were harvested from younger rats. The static incubation protocol was further modified by using HEPES buffer,

which is a more pH resilient buffer, as well as including potassium challenges at the

beginning and end of trials as a control. The smaller brain size of the younger rats made 61 it necessary that two PVN be used per trial for the hormone concentrations to be detectable.

Other technical considerations of this study include the small samples numbers used. This may have reduced the ability of the present study to detect the effects the relaxin peptides on the release of hormone and electrical activity in the PVN. Relaxin and relaxin-3 appeared to have no effect on the release of vasopressin in the PVN, however the power of the statistical tests used may have been too small to detect a change in vasopressin release with treatment due to the small sample numbers used.

Similarly, the difference in response of magnocellular neurons to relaxin and relaxin-3 could have been a consequence of small sample numbers. The phenotype of the neurons was not specifically identified. Some PVN neurons of each classification showed no change in membrane potential in response to the relaxin peptides, a common result for a study investigating the effect of peptides using intracellular electrophysiology (see for example: Hoyda et al. 2009, Price et al. 2009, Sirzen-Zelenskaya et al. 2011). It is possible that the magnocellular neurons that responded to relaxin or relaxin-3 may be oxytocinergic and the magnocellular neurons that failed to respond to the relaxin peptides may be vasopressinergic, or visa versa. Without identifying the phenotype of the neurons in these experiments, the usefulness of the findings of this study as an evaluation of the role of relaxin peptides in the PVN is limited.

Similarly, the expression of the relaxin receptors by the different neuronal populations in the PVN the relaxin receptors was not investigated. It has been shown in a prior study using in situ hybridization that both magnocellular and parvocellular 62 regions of the PVN express RXFP3 mRNA (Tanaka et al. 2005), and relaxin has been found to bind in both PVN regions as well (Osheroff et al. 1993). Without evaluating what types of neurons in the PVN express each of the relaxin receptors, as well as whether the receptors are expressed on the same or different neurons, the interpretation of the findings of the present study is limited.

CONCLUSION

The studies presented in this thesis were conducted in order to expand the understanding of the neuroendocrine functions of the relaxin family peptides, relaxin and relaxin-3. The findings suggest relaxin can act in the brain to induce the somatodendritic release of oxytocin and increase magnocellular neuron activity within the PVN. In contrast to relaxin, relaxin-3 appears to inhibit the somatodendritic release of oxytocin and decrease the activity of magnocellular neurons within the PVN. Given that relaxin-3 binds relaxin’s receptor, RXFP1, with lower affinity than its other receptor,

RXFP3, this dichotomy of magnocellular neuron response to the relaxin peptides

suggests the neuroendocrine function of relaxin-3 in the magnocellular regions of the

PVN could be dependant on the degree of relaxin-3 release and the availability of each

of its receptors on the magnocellular neurons.

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