Encyclopedia of Signaling Molecules

2018 Edition
| Editors: Sangdun Choi

Relaxin Family Peptide Receptors RXFP1 and RXFP2

  • Roger J. Summers
  • Michelle L. Halls
  • Ross A. D. Bathgate
Reference work entry
DOI: https://doi.org/10.1007/978-3-319-67199-4_362

Synonyms

 GREAT;  LGR7;  LGR8;  RXFP1;  RXFP2

Historical Background: Relaxin Family Peptides and Their Receptors

Relaxin was one of the first reproductive hormones to be identified, as a factor in the serum of pregnant guinea pigs that induced relaxation of the birth canal (Hisaw 1926). Until recently, relaxin was considered a hormone of pregnancy with little known of its roles in males and nonpregnant females. The isolation of relaxin from animal sources led to the determination of its structure, biological actions, and development of reliable bioassays (Schwabe and McDonald 1977; James et al. 1977; John et al. 1981), and this knowledge led to the use of recombinant DNA techniques to clone the rat (Hudson et al. 1981), pig (Haley et al. 1982), and human gene-1 (RLN1) (Hudson et al. 1983) and gene-2 relaxin (RLN2) genes (Hudson et al. 1984). The identification of other relaxin peptides and their cognate G protein-coupled receptors (GPCRs), more than 75 years after the identification of the peptide, stimulated a resurgence of interest in this pleiotropic hormone.

Relaxin family peptides are heterodimeric and closely related structurally to insulin. Relaxin, insulin-like peptide 3 (INSL3), relaxin-3, and insulin-like peptide 5 (INSL5) are the cognate ligands for the relaxin family peptide receptors RXFP1-4, respectively (Bathgate et al. 2006a, 2013; Halls et al. 2015). Humans and higher primates have two (RLN1 and RLN2) relaxin genes, whereas other mammals have only one (Rln1). The peptide encoded by RLN2 and mammalian Rln1 genes generates the peptide originally detected during pregnancy. The function of RLN1 in humans and higher primates is unknown. In humans, the relaxin family peptide genes have a similar structure and all synthesize prepropeptides (Hsu 2003) that are processed by convertases to the mature two-chain peptides: an A-chain linked to a B-chain by two disulfide bonds and an additional intrachain disulfide in the A-chain.

INSL3 (formerly Leydig insulin-like peptide) is found in the Leydig cells of the testis (Bathgate et al. 2006a, 2013; Halls et al. 2015; Adham et al. 1993) and has a critical role in testis descent with INSL3 knockout mice being cryptorchid and infertile (Nef and Parada 1999; Zimmermann et al. 1999). It plays an important role in gubernaculum development, which is involved in the first stage of testis descent, and also appears to have a role in the maintenance of ovarian function (Spanel-Borowski et al. 2001; Kawamura et al. 2004; Glister et al. 2013). INSL3 expression in other tissues occurs at much lower levels.

Thus, while relaxin and INSL3 resemble each other closely in structure, each is the cognate ligand for a specific G protein-coupled receptor (GPCR), and each possesses a wide variety of physiological functions. Relaxin has roles in reproduction, cardiovascular system, organ protection, metabolism, and as a neuropeptide in the brain; INSL3 although acting on a similar receptor has highly specialized roles in reproduction.

RXFP1, the cognate receptor for relaxin, has the typical seven transmembrane (TM) spanning regions of a GPCR, as well as a large extracellular domain comprising ten leucine-rich repeats (LRRs) and a unique N-terminal low-density lipoprotein receptor type A (LDLa) module (Hsu et al. 2002). RXFP1 mRNA and protein is found in reproductive tissues but also in the cardiovascular system as well as in a number of areas of the brain (for details see (Bathgate et al. 2013; Halls et al. 2015; Novak et al. 2006). Interaction of relaxin with RXFP1 involves at least three stages: high affinity binding between the Β-chain of relaxin and the RXFP1 LRR region, lower affinity binding to the TM extracellular loops (ECLs), and finally an essential interaction involving the LDLa module (Halls et al. 2005; Sudo et al. 2003). Although RXFP1 couples to numerous signal transduction pathways, early studies indicated that relaxin increased cAMP levels (Braddon 1978; Cheah and Sherwood 1980; Sanborn et al. 1980; Chen et al. 1988). The mode of cAMP production following stimulation of HEK293 cells expressing RXFP1 is complex and involves at least three G proteins (Halls et al. 2006, 2009a, b). RXFP1 also activates extracellular signal-regulated kinase 1/2 (ERK1/ERK2), tyrosine kinase(s), gene transcription, and nitric oxide (NO) signaling, and relaxin also interacts with the glucocorticoid receptor (GR) (Halls et al. 2015). The full implications of the pleiotropic effects of relaxin are still to be elucidated.

RXFP2 is the cognate receptor for INSL3 and is structurally similar to RXFP1 (Hsu et al. 2002) (Kumagai et al. 2002). It is primarily expressed in reproductive tissues (for details see Bathgate et al. 2013; Halls et al. 2015) and also influences bone metabolism (Ferlin et al. 2010). RXFP2 signaling also involves adenylyl cyclase (AC) activation and cAMP generation, utilizing a subset of the G proteins used by RXFP1 (Halls et al. 2006). Although in vitro cell systems that are often used to study RXFP2 show increases in cAMP levels, this is not always the case in endogenously expressing systems. Thus, in gubernacular cells (Kumagai et al. 2002) or osteoblasts (Ferlin et al. 2009, 2010), activation of RXFP2 increased cAMP, but in male germ cells and oocytes, decreased cAMP is observed (Kawamura et al. 2004), perhaps reflecting expression patterns of signaling proteins in different cells (Halls et al. 2009b). Although some species relaxins activate RXFP2 in vitro (Bathgate et al. 2006a, 2013; Halls et al. 2015; Hsu et al. 2002; Kumagai et al. 2002; Bathgate et al. 2006b; Scott et al. 2005a), there is no evidence that relaxin activates RXFP2 in vivo.

Thus, RXFP1 and RXFP2 are structurally similar receptors that utilize some common signaling mechanisms. However, current evidence suggests that the relaxin/RXFP1 system has a much wider range of distribution and functions than the INSL3/RXFP2 system.

Molecular Biology of RXFP1 and RXFP2

Human RXFP1 is located on chromosome 4q32.1, and RXFP2 is located on 13q13.1. The receptors share 60% amino acid sequence identity and 80% homology. Both RXFP1 and RXFP2 have multiple alternatively spliced isoforms (Bogatcheva et al. 2007; Bogzil et al. 2005; Bohm et al. 1994; Halls et al. 2007a, 2015). Thus far, 29 splice variants of RXFP1 and RXFP2 have been identified; four of these variants have been examined and show a wide range of tissue expression, but cannot bind either relaxin or INSL3 or increase cAMP accumulation. Although one isoform is highly expressed (RXFP2.1; deletion of exon 11, corresponding to leucine-rich repeat [LRR] 7), others were expressed either at very low levels (RXFP1.10; deletion of exon 3, flanking the low-density lipoprotein class a [LDLa] module), retained within the cell (RXFP1.2; deletion of exons 12 and 13, corresponding to LRRs 8 and 9), or, in the case of one variant (RXFP1.1; stop codon in exon 6, resulting in only the LDLa module and two LRRs), secreted, raising interesting questions regarding their role in endogenous regulation of full-length RXFP1 (Scott et al. 2006). Another even smaller secreted variant, RXFP1-truncate, has been identified in mouse, rat, and pig (Scott et al. 2005b) and consists of the receptor signal peptide, the LDLa module, 33 residues of the LRR flanking sequence, and a nonhomologous sequence of seven residues. When a construct encoding the truncate peptide is transiently expressed in HEK293T cells, the secreted protein inhibited relaxin-stimulated cAMP signaling mediated by the full-length receptor, acting as a functional “antagonist” of relaxin. Expression of RXFP1-truncate is increased during pregnancy in both mouse and rat, suggesting that it may functionally antagonize relaxin.

Structural Features and Functional Domains of RXFP1 and RXFP2

The similarity between RXFP1 and RXFP2 has facilitated the identification of the functional domains of the two receptors. Initial work with chimeric receptors and relaxin-3 revealed that the peptide interacts with both the LRR domain and ECL2 of the TM domain of RXFP1 to produce the full signaling profile (Sudo et al. 2003). This mechanism was later shown to be a feature of both receptors (Halls et al. 2005). Support for two binding sites also came from functional assays, with high-affinity LRR binding producing cAMP accumulation more effectively than the lower-affinity ECL site (Halls et al. 2005). In addition to the LRR region, the N-terminal LDLa module is essential for signaling but has no role in ligand binding (Scott et al. 2006; Hopkins et al. 2007; Kern et al. 2007) and likely interacts with other receptor domains (potentially the ECLs and TM domains) in a manner analogous to a tethered ligand.

LDLa module: RXFP1 and RXFP2 are the only known human GPCRs to contain an LDLa module (Bathgate et al. 2006a, 2013; Halls et al. 2015). The NMR solution structure of the RXFP1 LDLa module has been solved and reveals a fold generated by six cysteine residues and the incorporation of a calcium ion by a motif of acidic residues (Hopkins et al. 2007). Its role was first identified during characterization of a splice variant of RXFP2 that is missing the LDLa module. This receptor and a RXFP1 homologue are expressed at the cell surface and bind their cognate ligands normally but do not signal (Scott et al. 2006; Kong et al. 2013). A soluble recombinant form of the RXFP1 LDLa module acts as an RXFP1 antagonist when added to cells expressing RXFP1 (Scott et al. 2005b, 2006). These data suggest that the LDLa module acts as a tethered ligand at RXFP1. Further mutagenesis studies have provided evidence for an LDLa-RXFP1 interaction that drives receptor signaling. Mutants affecting folding of the LDLa module are unable to signal but maintain ligand binding profiles (Scott et al. 2006; Hopkins et al. 2007; Kern et al. 2007). Specifically, mutation of C47A and C53A involved in folding, or of D58E involved in calcium binding, produces mutants that do not increase cAMP accumulation (Kern et al. 2007). C27S and C40S mutations that affect calcium ligation and hence folding also abolish signaling (Hopkins et al. 2007) (Fig. 1). Similar studies at human RXFP2 with mutations of the calcium-ligating residue D70Y or conserved cysteine C71Y also resulted in a loss of signaling again highlighting the common mode of activation of the two receptors (Bogatcheva et al. 2007).
Relaxin Family Peptide Receptors RXFP1 and RXFP2, Fig. 1

Major signaling pathways observed following the activation of the cognate receptor for relaxin, RXFP1. Signalosomes are signaling complexes activated by sub-picomolar relaxin that are capable of responding to the amounts of relaxin commonly in the circulation. Canonical signaling in many cell types involves interaction of RXFP1 with at least three G proteins and the generation of cAMP and cGMP. The interactions are compartmentalized with RXFP1-Gαi3 interactions occurring in lipid-rich domains in the cell membrane. The small molecular weight agonist ML290 interacts at an allosteric site on RXFP1 to produce a biased signaling profile that preferentially activates cGMP over cAMP. There is also evidence that suggests that the anti-fibrotic effects of relaxin involve the formation of heterodimers between RXFP1 and the angiotensin AT2 receptor

Further evidence for an interaction between the LDLa module and RXFP1 comes from studies on chimeric and mutant RXFP1 (Hopkins et al. 2007; Kong et al. 2013). A chimera of the RXFP1 that contains a homologous LDLa module from the second ligand-binding domain of the LDL receptor produces a receptor (LB2-RXFP1) that is unable to signal to relaxin but binds ligand normally (Hopkins et al. 2007). Mutagenesis of conserved residues in the RXFP1 LDLa module highlighted potential roles for L29 and Y31 in receptor activation (Hopkins et al. 2007). Further studies with LB2-RXFP1 to define the potential “signaling surface” of the LDLa module confirmed key roles for L29 and Y31 and identified K39 as important residues in the RXFP1 LDLa module involved in hydrophobic interactions that drive the active receptor conformation (Kong et al. 2013). Studies with chimeric RXFP1 and RXFP2 with their LDLa modules swapped to the final cysteine residue of the module demonstrate that RXFP2 likely uses a similar but distinct mechanism to drive receptor activation. Importantly, chimeric receptors still bind ligand normally and signal, albeit with altered activity compared to wild-type receptors (Bruell et al. 2013). When the TM domains of the RXFP1 chimera with an RXFP2 LDLa (RXFP211) were swapped to match the LDLa module (RXFP212), the activity of the receptor approached wild type, suggesting a specific interaction between the LDLa module and the receptor TM domains.

The LDLa module may also influence receptor maturation and translocation to the cell surface. When expressed in HEK293 cells, a large fraction of RXFP1 remains in an immature form containing high mannose-type N-linked oligosaccharides within the endoplasmic reticulum (Kern et al. 2007). As for other glycoprotein hormone receptors (Ascoli et al. 2002; Tao et al. 2004; Pietila et al. 2005; Davis et al. 1995; Quintana et al. 1993), this may influence cell surface expression. In agreement with these findings, mutation of a conserved glycosylation site within the LDLa module results in a receptor with a reduced ability to generate cAMP, associated with a decrease in cell surface expression (Kern et al. 2007; Yan et al. 2008). However, there are inconsistencies, since an RXFP1 mutant lacking the LDLa module was expressed as the mature form only, as was a chimeric RXFP1 containing the LDLa module of RXFP2 (Kern et al. 2007), and studies on RXFP1 mutants with a disruption of the LDLa glycosylation site show only a minor (Yan et al. 2008) or no effect (Kong et al. 2013) on signaling and cell surface expression. Additionally, studies using LDLa-less RXFP1 and RXFP2 (Scott et al. 2006), LB2-RXFP1 mutants, RXFP1 LDLa loss of function mutants (Kong et al. 2013), or misfolded RXFP1 LDLa mutants (Hopkins et al. 2007) demonstrated no effect on cell surface expression. In RXFP2, mutations of amino acid residues that form the disulfide bond or coordinate calcium binding in the LDLa module (C70Y and D71Y) did reduce cell surface expression (Bogatcheva et al. 2007). Thus, the LDLa module plays an important and specific role in protein maturation, in cell surface expression, and in the activation of both RXFP1 and RXFP2.

LRR region: Glycosylation, a posttranslational modification common to many GPCRs, is important for receptor delivery to the cell surface, ligand binding, and signal transduction. In addition to sites within the LDLa module, the LRR region is also glycosylated at N105, N250, N303, and N346 that are important for receptor translocation and full signaling efficacy, but not ligand binding (Yan et al. 2008). The glycosylation sites of RXFP2 have yet to be studied in detail.

The LRR region is also essential as the primary high-affinity ligand-binding site for relaxin and INSL3 in RXFP1 and RXFP2 (Halls et al. 2005; Sudo et al. 2003). The B-chain residues of relaxin and INSL3 contribute most to binding affinity. Homology modeling of the LRRs and mutagenesis has been used to define the relaxin and INSL3 binding sites in RXFP1 and RXFP2. The RXFP1 LRRs were modeled on the crystal structure of the porcine ribonuclease inhibitor (a protein with LRRs) together with in silico peptide docking and receptor mutagenesis. Relaxin utilized the well-characterized RxxxRxxI binding motif within the peptide B-chain at 45° to five of the parallel LRRs (Bullesbach and Schwabe 2005a). The B-chain residues R13 and R17 were predicted to interact with acidic groups within the concave face of the LRRs (E277 and D279 in LRR8 and D231 and E233 in LRR6) (Bullesbach and Schwabe 2005a) (Fig. 1). The B-chain I20 was predicted to form a hydrophobic interaction with W180 and I182 within LRR4 and L204 and V206 within LRR5 (Bullesbach and Schwabe 2005a).

The interaction of INSL3 with RXFP2 utilizes different residues in the B-chain. Modeling of the INSL3 interaction with the RXFP2 LRRs utilized the NMR solution structure of INSL3 (Rosengren et al. 2006) and a molecular model of the RXFP2 LRR based upon the crystal structure of the Nogo receptor (Chen et al. 1988). Studies using a combination of peptide and LRR mutants in conjunction with in silico docking of the INSL3 B-chain to the LRR of RXFP2 predicted a B-chain residue R16 interaction with RXFP2 D227, H12 with RXFP2 W177, V19 with RXFP2 I179, R20 with RXFP2 E229 and D181, and W27 with RXFP2 F131 and Q133 (Scott et al. 2007). Although five of the RXFP2 residues that potentially interact with INSL3 are also found in RXFP1, the affinity of INSL3 for RXFP1 is very low (Halls et al. 2005; Sudo et al. 2003). Another study utilized the RXFP1 extracellular domain construct linked to a single CD8 TM domain, termed 7BP (Hsu et al. 2002) that has a high affinity for INSL3 (Halls et al. 2005) and was used as a template to further explore the interaction of INSL3 and the RXFP1 LRRs (Scott et al. 2012). Although only four amino acid changes are necessary to obtain the equivalent RXFP2 affinity for INSL3 in the RXFP1 LRRs, when these mutations were inserted into full-length RXFP1, they produced only a modest gain of function for INSL3. This confirmed that the TM domains have a role in modulating ligand binding. Molecular modeling studies demonstrated that relaxin and INSL3 have distinct orientations of binding to the LRRs. Taken together, the results suggest that there are critical differences both in the extracellular domain binding and the coordination of this binding with the TM binding site between RXFP1 and RXFP2 (Scott et al. 2012). Relaxin clearly binds to RXFP2 in a different manner from RXFP1 but also in a different manner from INSL3 binding to RXFP2.

TM domains and dimerization: The LRR binding site is the primary high-affinity binding site, but the studies with chimeric receptors and the selectivity of relaxin-3 for RXFP1 versus RXFP2 suggest an additional binding site in ECL2 (Sudo et al. 2003). This is supported by studies using the chimeric receptors with relaxin peptides that have selectivity for RXFP1 or RXFP2 (rat relaxin, RXFP1 > RXFP2; INSL3, RXFP2 > RXFP1) (Halls et al. 2005). Modeling studies suggest that A-chain residues in relaxin and INSL3 interact with this potential binding site within the TM domain (Hartley et al. 2009), but this has yet to be fully determined. It has been suggested that ligand binding to the LRRs (Halls et al. 2005; Bullesbach and Schwabe 2005a) and TM ECLs directs the LDLa to interact with the TM domain of a receptor homodimer partner to drive receptor activation (Bruell et al. 2013; Kong et al. 2010). Recent studies also identify the importance of the LRR-LDLa linker region in the ectodomain of RXFP1 such that binding to the latter region stabilizes and extends a helical conformation within the linker that acts as the critical switch for LDLa-mediated receptor activation (Sethi et al. 2016).

There is evidence that RXFP1 and RXFP2 form homo- and heterodimers and that this occurs in the absence of, and is independent of, ligand occupation of the receptor (Kern et al. 2008; Svendsen et al. 2008a, b). Heterodimers also form between the haloreceptor and a number of splice variants and are present at all stages of receptor translocation from the endoplasmic reticulum to the plasma membrane (Kern et al. 2008). Dimerization can also occur with a TM-only domain receptor (Svendsen et al. 2008a, b), suggesting that the TM domains drive dimerization, as with other GPCRs. The authors suggest that the receptor ectodomains may be required for stabilization of the dimer, due to the lower dimerization efficiency observed for the TM-only receptors. However, poor expression of the TM-only receptors could also explain these results (R.A.D. Bathgate unpublished).

Dimerization may also explain negative cooperativity (Svendsen et al. 2008a, b), whereby the affinity of unoccupied receptor binding sites progressively decreases as receptor occupancy increases. Two consequences of this are an increase in the functional concentration range of the ligand and a decrease in ligand residence time corresponding to an increase in free ligand concentration, potentially allowing selective activation of different signaling pathways (Shymko et al. 1997). It has also been suggested that this may provide an explanation for the observation that in many experimental and clinical situations, the concentration-response curves for relaxin acting at RXFP1 are bell shaped, although other mechanisms have been proposed (see section on “Signal Transduction Pathways of RXFP1 and RXFP2”).

Intracellular receptor domains: The ICL3 and the C-terminal tail have important roles in signaling. ICL3 couples RXFP1 to Gαs, essential for activation of AC and cAMP signaling. Fragments of the N-terminal region of ICL3 (615-629 and 619-629-Lys (Palm)) increase adenylyl cyclase (AC) activity (Shpakov et al. 2007). These peptides also functionally “antagonize” the cAMP response to relaxin in rat striatum and cardiac muscle (Shpakov et al. 2007). A peptide based on the C-terminus of Gαs (385-394) inhibited AC activity stimulated by relaxin or ICL3 peptides (Shpakov et al. 2007). In many GPCRs, ICL3 contains motifs that interact with G proteins (Kjelsberg et al. 1992; Ren et al. 1993; Herrick-Davis et al. 1997; Egan et al. 1998) and may also direct coupling to GαoB for both RXFP1 and RXFP2 although direct evidence is lacking. Truncation of the C-terminal tail of either receptor does not affect the Gαs or the GαOB components of cAMP signaling, suggesting a common interaction site.

The C-terminal tail of RXFP1 is necessary for cAMP accumulation through the Gαi3 pathway and essential for signalosome formation. RXFP1 but not RXFP2 increases cAMP accumulation by coupling to Gαi3 (Halls et al. 2006) followed by activation of AC5, utilizing a Gβγ-phosphatidylinositol 3-kinase (PI3K)-protein kinase C (PKC)ζ pathway. Activation of this pathway involves the final ten amino acids of the C-terminus and absolutely requires R752 (Halls et al. 2009a) (Fig. 1). The mechanisms suggested include direct Gαi3 coupling (a mechanism not observed for any other GPCR), Gαi3 coupling following receptor phosphorylation, or recruitment of scaffolding proteins for co-localization of the receptor with Gαi3.

The C-terminal tail of RXFP1 is also required for constitutive activity (Halls and Cooper 2010) where the receptor couples to AC2 involving helix 8 and A-kinase anchoring protein 79 (AKAP79). The signalosome facilitates Gαs and Gβγ-mediated stimulation of AC2 to cause cAMP accumulation in response to sub-picomolar concentrations of relaxin. The amount of cAMP generated by the signalosome is tightly controlled by protein kinase A (PKA)-stimulated phosphodiesterase (PDE) 4D3, which is scaffolded by β-arrestin 2 and S704 of RXFP1.

The RXFP1 C-terminal tail also contains consensus sequences for phosphorylation and protein-protein interactions and is the region most divergent between RXFP1 and RXFP2 (Halls et al. 2007a). It therefore represents an area of functional divergence between the two receptors that may relate to the more varied physiological roles of relaxin compared to INSL3.

RXFP1 and RXFP2 therefore have similarities in structure, binding sites, and signaling mechanisms. Both receptors possess a high-affinity binding site in the LRR region and a lower affinity site in ECL2 of the TM domain and require an intact LDLa module for signaling. RXFP1 has a more complex C-terminal region that contains residues essential for signaling involving Gαi3 and formation of signalosomes. Both receptors form homo- and heterodimers but the functional significance of this is not clear.

Ligands That Activate RXFP1

Relaxin family peptides: Although relaxin peptides show considerable variation across species, there are conserved regions. In addition to the cysteines necessary for the two-chain structure, there is a conserved RxxxRxxI/RxxxRxxV motif in the B-chain. The arginines are outward facing and located on the first and second loop of the α-helix (Eigenbrot et al. 1991) where together with an isoleucine or valine residue, they comprise the receptor binding site for relaxin (Schwabe and Bullesbach 1994; Tan et al. 2002). Replacement of arginine, isoleucine, or valine in this motif (RxxxRxxI/RxxxRxxV) markedly reduces or abolishes activity (Schwabe and Bullesbach 1994; Tan et al. 2002). Although the relaxin A-chain shows greater variation across species (Sherwood 1994), G14 is highly conserved and necessary for chain flexibility and structure (Bullesbach and Schwabe 1994). Structure activity studies have led to a potent and selective agonist for RXFP1, H2:A(4-24)(F23A), that has similar potency and biological activity to human relaxin but no significant activity at RXFP2 (Chan et al. 2012). Human relaxin binds to RXFP1 and RXFP2 by distinct mechanisms as illustrated by the species-specific nature of the structure activity relationships with mouse and rat relaxin or human relaxin-3 neither binding to nor activating RXFP2 (Bathgate et al. 2006b; Scott et al. 2005c). Peptides with a reduced B-chain (H2:(B7-24)) or reduced A- and B-chains (H2:(A4-24)(B7-24)) show some reduction in binding activity and ability to generate cAMP but have similar anti-fibrotic activity to human relaxin and less activity at RXFP2 (Hossain et al. 2011).

C1q-tumor necrosis factor-related protein 8 (CTRP8): Short linear peptides, derived from a naturally occurring protein containing a collagen-like repeat, appear to act at RXFP1 (Shemesh et al. 2009). The effects of the peptides CGEN-25009 and CGEN-25010 are extremely variable (Shemesh et al. 2008, 2009) but do suggest relaxin-like activity in THP-1 cells and in a fibrosis model (Pini et al. 2010). CGEN-25009 and human relaxin increased cAMP, cGMP, and nitrite, decreased collagen deposition, and increased matrix metalloproteinase (MMP-2) activity in human dermal fibroblasts (Pini et al. 2010). More recent studies with these peptides and the precursor protein C1q-tumor necrosis factor-related protein 8 (CTRP8) demonstrated activation of RXFP1 (Glogowska et al. 2013; Thanasupawat et al. 2015) with cAMP production and a PI3K-mediated pro-migratory phenotype in glioblastoma cell lines and primary cells. Co-immunoprecipitation studies suggest a direct interaction between human CTRP8 and RXFP1. Although these studies suggest that CTRP8 or peptide fragments activate RXFP1, it remains to be seen whether they are native ligands and/or whether activation of RXFP1 by CTRP8 occurs specifically in glioblastomas.

Small molecular weight allosteric agonists: An HTS strategy identified a series of 2-acetamido-N-phenylbenzamides active at the human RXFP1 (Xiao et al. 2012; Chen et al. 2013). Some compounds showed reasonable potency for cAMP generation in HEK293-RXFP1 and THP-1 cells and good specificity for RXFP1 versus RXFP2 (Xiao et al. 2013). Several compounds increased vascular endothelial growth factor (VEGF) gene expression in THP-1 cells with similar efficacy to relaxin, and compound 8 (ML290) increased cellular impedance in a label-free system with about 500-fold lower potency than relaxin (Xiao et al. 2013). Comparison of the properties of ML290 with that of relaxin suggests that it is an allosteric agonist at RXFP1 (Xiao et al. 2013). The interaction involves the seventh TM domain (TM7) and the third extracellular loop (ECL3) of RXFP1. Residues involved in agonist binding and receptor activation include a hydrophobic region in TM7 comprising W664, F668, and L670 (Hu et al. 2016). The G659/T660 motif in ECL3 is central to the species selectivity of the small molecular weight agonists. Other studies show that the D58E mutant of RXFP1 does not produce cAMP in response to relaxin, but the response to ML290 is unaffected (Xiao et al. 2013). Since the mouse RXFP1 does not respond to ML290, chimeras of human and mouse RXFP1 were used to identify the region that interacts with ML290 and revealed two pairs of amino acids divergent in ECL3. A mouse receptor with a double substitution of the residues, IL to VV and DS to GT, produced a mouse RXFP1 that responded to ML290 in a similar fashion to the human receptor (Xiao et al. 2013). More recent studies indicate that macaque and pig, in addition to human RXFP1, also respond to ML290 (Huang et al. 2015). It remains to be demonstrated how the profile of activity of ML290 and similar compounds compares with that of relaxin in a variety of cellular and animal disease models.

Ligands That Activate RXFP2

Relaxin family peptides: INSL3 is the cognate ligand for RXFP2 that interacts with RXFP1 only at high concentrations (Halls et al. 2005; Kumagai et al. 2002), suggesting that this interaction has no physiological role. In contrast, human relaxin potently activates RXFP2 although its mode of interaction differs from INSL3 that utilizes distinct amino acids in the B-chain to bind to and activate RXFP2 (Bullesbach and Schwabe 2004, 2005b) and individual substitutions only slightly reduce binding. However, combined mutation of H12A, R16A, V19A, R20A, and W27A abolishes INSL3 binding (Rosengren et al. 2006).

Truncation of the A-chain of human relaxin produces peptides that progressively lose the ability to bind to and activate both RXFP1 and RXFP2 (Hossain et al. 2008), whereas truncation of the A-chain of INSL3 completely abolishes activation of RXFP2 with no effect on binding (Bullesbach and Schwabe 2005b). Targeted point mutations within the A-chain selectively alter receptor binding and activation (Park et al. 2008). All studies highlight the importance of both A- and B-chains for INSL3 activation of RXFP2 and show that the interaction is different from that of relaxin-RXFP1 (Hossain et al. 2008).

Peptide antagonists: In contrast to RXFP1, there are numerous peptide antagonists of RXFP2. Deletion of ten residues from the N-terminus of the INSL3 A-chain, deletion of eight N-terminal residues of the B-chain, or disruption of the intra-A-chain disulfide bond (C10S/C15S or C10del/C15del) produce peptides that still bind to RXFP2, but no longer increase cAMP accumulation and are specific competitive inhibitors of INSL3 at RXFP2 (Bullesbach and Schwabe 2005b; Zhang et al. 2010). Substitution of eight A- or B-chain residues with alanine does not affect receptor signaling (Bullesbach and Schwabe 2005b). The INSL3 B-chain alone is a RXFP2 antagonist although it only has low affinity for the receptor (Del Borgo et al. 2006; Shabanpoor et al. 2007). Modifications of the single chain structure result in modest gains in affinity, but larger gains are obtained by linking two B-chains using the native cysteine residues (Shabanpoor et al. 2010) to produce antagonists with affinities only slightly lower than INSL3 (Shabanpoor et al. 2011).

Signal Transduction Pathways of RXFP1 and RXFP2

RXFP1

Canonical signaling pathways: RXFP1 signals to a variety of pathways including cAMP, MAP kinases, tyrosine kinases, and NO. The importance of cAMP as a signaling pathway for relaxin was confirmed during RXFP1 deorphanization (Hsu et al. 2000, 2002). RXFP1 couples to Gαs to increase cAMP (Hsu et al. 2000, 2002, Halls et al. 2006), an effect modulated by coupling to GαOB (Halls et al. 2006), but also couples to Gαi3 to cause a delayed surge of cAMP accumulation utilizing a Gβγ-PI3K-PKCζ pathway that activates AC5 (Halls et al. 2006, 2009a; Nguyen et al. 2003; Nguyen and Dessauer 2005). The delayed pathway was first identified in THP-1 cells endogenously expressing RXFP1 where relaxin causes a biphasic increase in cAMP accumulation, with the later phase partially blocked by the PI3K inhibitors LY294002 and Wortmannin. Relaxin also increased PI3K activity (Nguyen et al. 2003) that was shown to be essential for translocation of PKCζ to the cell membrane and for the late cAMP response (Nguyen and Dessauer 2005). This pathway also occurs in HEK293 cells transiently or stably expressing RXFP1 (Halls et al. 2006, 2009a) and is downstream of Gα and Gβγ subunits. Transfection of cells with Gαi/o mutants that are insensitive to pertussis toxin (PTX) shows coupling of RXFP1 to Gαi3 as the mediator of the Gβγ-PI3K-PKCζ-AC5 pathway. This also occurs in rat left atria where relaxin increases cAMP accumulation that is reduced by pertussis toxin (PTX) pretreatment, which also reduced the inotropic and chronotropic responses to relaxin (Kompa et al. 2002) and in the failing human heart where there is increased Gαi/o expression (Bohm et al. 1994; Eschenhagen et al. 1992) and where the positive inotropic effects of relaxin in humans are preserved (Dschietzig et al. 2011). The final ten amino acids of the RXFP1 C-terminus, particularly R752, are essential for activation of the Gαi3 pathway (Halls et al. 2009a) (Fig. 1) as is the presence of lipid-rich membrane domains, suggesting compartmentalization of the cAMP response (Halls et al. 2009a). GTPγS-immunoprecipitation studies show that Gαi3 is activated immediately after RXFP1 stimulation, suggesting that the delay occurs downstream of the G protein and is due to the translocation of PKCζ (Halls et al. 2009a). In HEK293 cells, activation of Gαs- and GαOB-dependent cAMP pathways increases CRE-mediated gene transcription, whereas Gαi3-mediated signaling selectively regulates NFκB-dependent gene transcription (Halls et al. 2007b), again suggesting compartmentalization of signaling events, and inferring that distinct physiological outcomes can be anticipated downstream of different cAMP signaling branches (Halls et al. 2007b).

cAMP accumulation in response to relaxin may also occur by G protein-independent mechanisms such as via activation of tyrosine kinases. In THP-1 cells and some primary cell cultures, cAMP accumulation in response to porcine relaxin is blocked by tyrosine kinase inhibition (Kuznetsova et al. 1999; Bartsch et al. 2001; Anand-Ivell et al. 2007; Heng et al. 2008) and potentiated by phosphotyrosine phosphatase inhibitors (bpV(phen) and mpV(pic)) that mimic tyrosine kinase activation (Bartsch et al. 2001). In human lower uterine segment fibroblasts, relaxin caused tyrosine phosphorylation with no effect upon cAMP accumulation (Palejwala et al. 1998). The tyrosine kinase-dependent increase in cAMP accumulation may occur by PDE inhibition. However, tyrosine kinase inhibitors do not affect relaxin-stimulated cAMP accumulation in HEK293 cells expressing RXFP1 (Anand-Ivell et al. 2007), emphasizing differences in cellular responses between different cell types. In cells utilizing the tyrosine kinase pathway, there was some degree of cAMP inhibition by the PI3K inhibitor, LY294002 (Anand-Ivell et al. 2007; Heng et al. 2008), and evidence of a negative feedback loop involving PKA (Anand-Ivell et al. 2007).

NO production, cGMP generation, and PKG activation are other pathways by which relaxin exerts its effects. Relaxin has been shown to activate endothelial nitric oxide synthase (eNOS) (Dschietzig et al. 2012; Baccari et al. 2007) and neuronal NOS (nNOS) (Baccari et al. 2004; Mookerjee et al. 2009) or stimulate inducible NOS (iNOS) expression (Bani et al. 1998a; Alexiou et al. 2013). In rat lung, relaxin-mediated iNOS upregulation is stimulated by ERK1/ERK2 activation and inhibited by PI3K stimulation (Alexiou et al. 2013).

Many cells also respond to relaxin by activation of ERK1/ERK2. In normal human endometrial cells (NHE cells), relaxin causes a rapid, transient increase of pERK1/pERK2 (Zhang et al. 2002). A similar pERK1/pERK2 response is also observed in THP-1 cells and in human coronary artery and pulmonary artery smooth muscle cells and is associated with increased transcription of VEGF (Zhang et al. 2002). In HeLa cells and human umbilical vein endothelial cells (HUVECs), the pERK1/pERK2 response is more prolonged (Dschietzig et al. 2003). In HeLa, EAhy926, HT-29, and primary fibrochondrocyte cells, the pERK1/pERK2 and pAkt responses are maximal at 30 min (Dschietzig et al. 2009a; Ahmad et al. 2012). In primary fibrochondrocytes, relaxin also activates PI3K, PKCζ, NFκB, c-fos, and Elk-1, all of which influence the expression of MMP-9 (Ahmad et al. 2012). Relaxin also produces a rapid peak followed by a sustained increase in pERK1/pERK2 in rat renal myofibroblasts, with the peak potentiated by inhibition of Gαi/o by PTX (Mookerjee et al. 2009). In contrast, in human vascular smooth muscle cells (hVSMC), relaxin stimulation did not increase pERK1/pERK2 but did increase phosphorylation of p38 MAPK (Dschietzig et al. 2003). In rat pulmonary arterial endothelial and smooth muscle cells, relaxin increased iNOS expression and activity dependent on a balance of pERK1/pERK2 and PI3K pathway activation (Alexiou et al. 2013). Thus, relaxin produces a variety of effects on phosphorylation of a number of kinases in multiple cell types.

Relaxin has well-established anti-fibrotic actions due to inhibition of TGF-β1 signaling that involves activation of PI3K and NOS-NO-cGMP (Ahmad et al. 2012; Chow et al. 2012). Relaxin also influences differentiation of neonatal fibroblasts into myofibroblasts by preventing TGF-β1-mediated inhibition of Notch-1 signaling which controls cell differentiation and fate and is involved in fibrosis (Fan et al. 2011). Downregulation of Notch-1, which is induced by TGF-β1, is a necessary step in the differentiation of rat cardiac fibroblasts into myofibroblasts (Fan et al. 2011).

Noncanonical signaling pathways: Relaxin activates the glucocorticoid receptor (GR), a nuclear receptor that acts as a ligand-dependent transcription factor (Dschietzig et al. 2004), an action that may account for the effects of relaxin upon gene transcription. Relaxin treatment of THP-1 cells differentiated into macrophages blunts the production of cytokines with the effect being abolished by the GR antagonists RU-486 and D06 (Dschietzig et al. 2004, 2009a). Relaxin co-immunoprecipitates with the GR and levels of nuclear GR increase after 30 min stimulation with relaxin. Relaxin also displaces classic glucocorticoids from the GR (Dschietzig et al. 2004, 2009a), but the region of relaxin that binds to the GR differs from that involved in relaxin binding to RXFP1 (Dschietzig et al. 2009a). Relaxin interaction with the GR autoregulates its own expression by binding to half sites of glucocorticoid response elements upstream of transcription start at the human RLN2 promoter (Dschietzig et al. 2009b). Actions at the GR may also be involved in the vasodilator effects of relaxin since in rat aortic rings, relaxation to acetylcholine is impaired by TNF-α and the effect is reversed by relaxin (Dschietzig et al. 2012). The effects of relaxin are blocked by the PI3K inhibitor Wortmannin or by the GR and progesterone receptor antagonist RU-486. In rat primary aortic endothelial cells, TNF-α treatment increased endothelin-1 (ET-1) and arginase II expression, decreased superoxide dismutase (SOD)-1 expression, and stimulated superoxide and nitrotyrosine formation (Dschietzig et al. 2012). All of these effects were reversed or attenuated by relaxin acting at the GR (Dschietzig et al. 2012). Thus, relaxin protects the endothelium by activating the canonical PI3K-Akt-eNOS pathway but also the GR, with SOD-1 upregulation being dependent on relaxin-GR-c/EBP-β signaling (Dschietzig et al. 2012). Other reports that support a relaxin-GR interaction include relaxin-mediated protection in a rodent model of severe acute pancreatitis (an action that is attenuated by treatment with RU-486 (Cosen-Binker et al. 2006)), relaxin-induced GR activation in reporter gene experiments (Halls et al. 2007b), and blockade of relaxin-mediated decreases in GM-CSF and IL-8 secretion from primary decidual macrophages by RU-486 (Horton et al. 2011).

The RXFP1 signalosome: Relaxin activates canonical signaling pathways in the nanomolar concentration range (Bathgate et al. 2013), concentrations higher than those normally in the circulation (Sherwood 2004). However, an RXFP1-dependent cAMP response has been identified using FRET-based cAMP biosensors in cells expressing RXFP1 (Halls and Cooper 2010) that depends upon a protein complex, or signalosome, linked to the relaxin receptor and sensitive to attomolar concentrations of relaxin (Fig. 1). This mechanism may provide the basis for physiological responses to relaxin at levels present in the circulation. The signalosome comprises RXFP1 scaffolded to AC2 by AKAP79, facilitating efficient activation of the AC by Gαs and Gβγ subunits. cAMP production is tightly regulated by PKA-activated PDE4D3 itself, scaffolded to the receptor C-terminus (at S704) by β-arrestin 2 (Halls and Cooper 2010). The signalosome thus possesses both stimulatory (AKAP79 and AC2) and regulatory (β-arrestin 2, PKA, and PDE4D3) arms that are both spatially and functionally distinct. There is no effect of inhibitors of classical pathway-specific proteins (including Gαi/o, PI3K, and PKC) on cAMP generated in response to sub-picomolar concentrations of relaxin and no effect of inhibition of signalosome-specific proteins (including AC2, AKAP79, and β-arrestin 2) upon classical relaxin cAMP signaling. The pathways also generate cAMP in quite distinct regions of the cell. Thus, signalosome-specific AC2 is excluded from lipid-rich domains (Willoughby et al. 2007) that are essential for the activation of the Gαi3 pathway with nanomolar relaxin (Halls et al. 2009a). Since AC2 expression occurs predominantly in the brain, lung, skeletal muscle, heart, and uterine myometrium (Willoughby et al. 2007; Defer et al. 2000; Sadana and Dessauer 2009), it is likely that some tissues display RXFP1 signalosome signaling, whereas others do not, which may help to determine its physiological role.

Signaling aspects of RXFP1 oligomers: There is evidence that RXFP1 receptors form homo- and heterodimers (Kern et al. 2008; Svendsen et al. 2008a, b). It has been suggested that dimer formation is necessary for signal transduction with ligand binding occurring at one dimer partner, followed by interaction of the bound ligand with the ECL2 of the second partner and initiation of signaling (Kong et al. 2010). However, there are no experiments with two inactive mutant receptor dimer partners that show complementation and rescue of function to support this concept. Negative cooperativity at RXFP1 has been suggested by the finding that the presence of unlabeled relaxin modestly increases the rate of dissociation (Svendsen et al. 2008a, b) although such behavior does not necessarily reflect dimer formation (Chabre et al. 2009). Studies are required that utilize receptors that retain function but are unable to form dimers or on receptors expressed in model phospholipid bilayers that allow examination of the functional characteristics of the monomer (Velez-Ruiz and Sunahara 2011; Whorton et al. 2007).

One intriguing aspect of relaxin-RXFP1 pharmacology is the phenomenon of bell-shaped concentration-response curves that are observed in many bioassays from studies in recombinant and primary cell systems (Halls et al. 2006; Sarwar et al. 2015), animal studies (Danielson and Conrad 2003; Debrah et al. 2005), and clinical trials (Teerlink et al. 2009). Negative cooperativity has been suggested as a potential explanation, but currently available information does not support this mechanism. While both RXFP1 and RXFP2 display negative cooperativity, bell-shaped curves for cAMP accumulation are only observed in response to three or 30 min activation of RXFP1 or 30 min activation of RXFP2, but not 3 min activation of RXFP2 (Halls et al. 2006). It could be argued that the RXFP2 system at 3 min is not in equilibrium and that increasing receptor occupation balances out reduced receptor affinity, but this does not convincingly explain the difference between RXFP1 and RXFP2 at the same time point. In human vascular cells that endogenously express RXFP1, venous endothelial and smooth muscle cells displayed pronounced bell-shaped concentration-response curves, yet arterial smooth muscle cells showed conventional sigmoidal concentration-response curves (Sarwar et al. 2015). This would appear to be incompatible with an explanation based on RXFP1 homodimer formation and negative cooperativity. The shape of the concentration-response curves could also be altered from bell shaped to sigmoid by selective inhibition of particular G proteins (Sarwar et al. 2015).

In HUASMC, cAMP and cGMP concentration-response relationships were sigmoidal, suggesting that responses involved Gαs and Gαi3 but not GαOB as inhibition of Gαs with NF449 or Gαi with NF023 desensitized the response but did not change the shape of the curves. Blockade of both G proteins completely abrogated cAMP and cGMP responses (Sarwar et al. 2015). In contrast, in HUVSMC, cAMP and cGMP concentration-response curves were bell shaped, suggesting that the responses involved not only Gαs and Gαi3 but also GαOB and that the latter coupled less efficiently to RXFP1. Inhibition of Gαs desensitized the responses but did not change the shape of the curves, whereas inhibition of Gαi both desensitized and changed the shape of the curve to sigmoid (Sarwar et al. 2015). Again, blockade of Gαs, Gαi3, and GαOB completely abrogated responses. In both arterial and venous cells, disruption of lipid rafts with filipin III had the same effect as inhibition of Gi/Go proteins (Sarwar et al. 2015). The results suggest that both Gαs and Gαi3 contribute to the increases in cAMP (Halls et al. 2006, 2009a) and cGMP observed in both HUASMC and HUVSMC and that GαOB inhibits these responses but only operates in HUVSMC and interacts with RXFP1 with lower affinity than the other G proteins.

There is also evidence for dimer formation between the haloreceptor and splice variants (encoding only the LDLa module and up to eight LRRs), and dimers are present at all stages of receptor translocation from the endoplasmic reticulum to the plasma membrane (Kern et al. 2008). Dimers were formed less effectively between the haloreceptor and a TM-only domain receptor (Svendsen et al. 2008a, b). Examination of three of the splice variants cloned from human fetal membranes showed that they did not signal (Kern et al. 2008), but co-expression with RXFP1 produced a parallel shift to the right of the cAMP concentration-response curve. Since the splice variants also markedly reduced cell surface expression of RXFP1, the reduced responses are likely due to a dominant negative effect (Kern et al. 2008).

Recent studies also suggest heterodimer formation between RXFP1 and other GPCRs. The anti-fibrotic actions of relaxin are mediated through a RXFP1-pERK1/pERK2-nNOS-NO-cGMP pathway leading to regulation of collagen-degrading MMPs (Chow et al. 2012). The effect of relaxin was completely absent in angiotensin type 2 receptor (AT2R)-/y mice or in mice treated with the AT2R antagonist PD123319 suggesting that the AT2R was necessary for the anti-fibrotic actions of relaxin (Fig. 1). BRET studies showed that RXFP1 and AT2R form constitutive heterodimers suggesting that the receptor complex is responsible for the novel pharmacology. There is no direct binding of relaxin to the AT2R, and the peptide does not affect the BRET signal from RXFP1-AT2R complexes (Chow et al. 2014). Importantly, this also explains why the anti-fibrotic actions of relaxin are only observed in pathological states: under normal physiological conditions, AT2Rs are expressed at low levels but are dramatically increased with injury or disease (Jones et al. 2008; Carey 2005; Siragy and Carey 1997; Matsubara 1998; Savoia et al. 2006).

Allosteric agonist signaling: The small molecular weight allosteric agonist ML290 (Xiao et al. 2013) does not require the LDLa module for signaling (Xiao et al. 2013). ML290 interacts with ECL3 of RXFP1 (Fig. 1), specifically requiring G659 and T660 within ECL3 for activity (Xiao et al. 2013). ML290 does not interact with the classical binding sites on RXFP1 utilized by relaxin and does not compete for 125I relaxin binding.

RXFP2

Canonical signaling pathways: Signaling pathways activated by stimulation of RXFP2 are simpler than those observed for RXFP1. INSL3 (or relaxin) stimulation of HEK293-RXFP2 cells causes Gαs-mediated increases in cAMP accumulation, which are negatively modulated by GαoB (Halls et al. 2006; Kumagai et al. 2002). This stage of signaling closely resembles the first stage of RXFP1 signaling (Fig. 1) (Halls et al. 2006). The contribution of GαoB to the response is seen by the increase in cAMP following the removal of Gβγ using βARK-ct or inhibition of Gαi/o with PTX (Halls et al. 2006). RXFP2 does not display constitutive activity or a high sensitivity response to INSL3 as seen with the RXFP1 signalosome (Halls and Cooper 2010); neither is there activation of the Gαi3-cAMP signaling pathway (Halls et al. 2006, 2009a).

s and GαoB are also involved in cAMP accumulation in cells that endogenously express RXFP2. In rat gubernacular cells (Kumagai et al. 2002), in a human osteoblast cell line (MG-63) (Ferlin et al. 2008) and in mouse primary Leydig cells (Pathirana et al. 2012), INSL3 stimulation of RXFP2 increases cAMP accumulation, probably involving Gαs. However, primary cultures of testicular germ cells and oocytes respond to INSL3 activation of RXFP2 with a PTX-sensitive inhibition of cAMP accumulation (Kawamura et al. 2004), consistent with RXFP2 coupling to GαoB. Thus, as for RXFP1, the net signaling outcome of RXFP2 stimulation will depend upon the signaling components (especially G protein isoforms) that are expressed in a particular cell type.

Localization of RXFP1 and RXFP2 Receptors

RXFP1 is present in a variety of tissues (reviewed in Bathgate et al. 2006a; Bathgate et al. 2013; Halls et al. 2007a, 2015; van der Westhuizen et al. 2008). Northern blots of human tissue identified relaxin receptor mRNA in the ovary, uterus, placenta, testis, prostate, brain, kidney, heart, lung, liver, adrenal gland, thyroid gland, salivary glands, muscle, peripheral blood cells, bone marrow, and skin. An additional shorter length receptor mRNA was also identified in the oviduct, uterus, colon, and brain. Human relaxin receptor protein expression has been identified by immunohistochemical analysis in the uterus, cervix, vagina, nipple, and breast. Studies using Northern blots of rat tissue have additionally identified receptor mRNA in the small intestine and oviduct, and LacZ reporter expression was also identified in oviduct in a relaxin receptor knockout mouse model.

INSL3 receptor mRNA expression in human (reviewed in (Bathgate et al. 2006a, 2013; Halls et al. 2007a, 2015; van der Westhuizen et al. 2008; Ivell and Anand-Ivell 2009) occurs in the uterus, testis, brain, pituitary, kidney, thyroid, muscle, peripheral blood cells, and bone marrow as determined by reverse transcriptase-polymerase chain reaction (RT-PCR). Additional expression was identified in the ovary and gubernaculum in the mouse as determined by RT-PCR and specifically in the mesenchymal and cremaster muscle of the gubernaculum using immunohistochemistry, in the rat gubernaculum using RT-PCR and Northern blot analysis, and in the rat ovary using RT-PCR, Northern blot analysis, and in situ hybridization.

Physiological Roles of Relaxin/RXFP1 and INSL3/RXFP2

Relaxin and RXFP1

Reproduction: Initially, the role of relaxin produced by the corpus luteum and/or placenta, as described in pregnant guinea pigs, was thought to be preparation of the birth canal for parturition (Hisaw 1926). Although this special endocrine function is found in species such as rodents and some other mammals, it is now apparent that it is of less importance in higher species. Currently it is established that in women, maximum circulating relaxin levels occur during the first trimester of pregnancy, suggesting that it is likely to be associated with first trimester events such as embryo implantation (Bathgate et al. 2006c). The peptide also causes a number of additional changes associated with pregnancy, including uterine growth and development, myometrial contractility, central control of plasma osmolality, and cardiovascular adaptations (reviewed in Bathgate et al. 2006a).

Relaxin has trophic effects on the mammary gland and/or nipple in several species. Nipple development is completely blocked in relaxin knockout mice with limited effects on the mammary gland (Zhao et al. 1999) which causes pups born to relaxin knockout mice to die within 24 h unless cross-fostered to wild-type mothers (Zhao et al. 1999). The same phenotype is displayed by RXFP1 knockout mice (Krajnc-Franken et al. 2004) and is not rescued by transgenic overexpression of INSL3 (Kamat et al. 2004). Relaxin binding sites are present in the mammary glands of pigs, rats, and humans where RXFP1 receptors are localized to the nipple, epithelial cells (Kohsaka et al. 1998), and stromal tissue (Ivell et al. 2003).

Relaxin is also found in the male reproductive tract in most mammals in human seminal plasma, appears to be produced in the prostate, and is identical to luteal relaxin (Winslow et al. 1992; Sokol et al. 1989; Yki et al. 1983). RXFP1 is expressed in sperm in both mice (Krajnc-Franken et al. 2004) and humans (Carrell et al. 1995; Ferlin et al. 2012) and facilitates sperm motility and penetration into oocytes (Weiss 1989). Seminal relaxin may act on the female reproductive tract to prepare the endometrium for implantation (Unemori et al. 1999; Telgmann and Gellersen 1998).

Central nervous system: In mammals, RXFP1 in the subfornical organ (SFO) and organum vasculosum of the lamina terminalis (OVLT) is activated by relaxin to cause a reduction in plasma osmolality (Sunn et al. 2002). In humans, this effect may not be completely attributable to relaxin as women that become pregnant following ovum donation (and lack circulating relaxin) display attenuated (Smith et al. 2006a) or no decrease in plasma osmolality (Johnson et al. 1991, 1996). However, the apparent lack of relaxin in these studies may have to be reassessed given the discovery of responses to fM concentrations of relaxin (Halls and Cooper 2010).

Activation of RXFP1 in the circumventricular organs and hypothalamic nuclei may also have a role in the timing of parturition which is disrupted by central administration of a relaxin monoclonal antibody (Summerlee et al. 1998). RXFP1 is also highly expressed in the basolateral amygdala, and administration of relaxin to this region impairs fear-related memory consolidation in rats (Ma et al. 2005). Although RXFP1 is highly expressed in other regions associated with memory formation such as the neocortex, thalamic nuclei, hippocampus, and supramammillary nucleus, there are no studies to date that examine effects on memory.

Blood vessels: Relaxin influences many of the adaptive cardiovascular changes that occur in pregnancy (Conrad 2011) such as increases in cardiac output, heart rate, and a decrease in vascular resistance (Debrah et al. 2005, 2006; Conrad and Novak 2004; Conrad et al. 2004). In rats, chronic relaxin administration increases renal plasma flow and glomerular filtration rate (Danielson et al. 1999), and a few studies suggest similar effects in humans (Conrad and Shroff 2011) (Erikson and Unemori 2001; Teichman et al. 2009; Dschietzig et al. 2009c).

Vasodilation in arterioles, capillaries, and venules to relaxin occurs in many tissues including reproductive tissues (Bani et al. 1988; Lee et al. 1992; Vasilenko et al. 1986), the heart (Bani et al. 1998b; Masini et al. 1997; Bani-Sacchi et al. 1995), the liver (Bani et al. 2001), and the cecum (Bigazzi et al. 1986). Relaxin is a potent vasodilator in some but not all arteries (Conrad 2010; McGuane et al. 2011) and is a physiological antagonist of vasoconstrictors (Bani et al. 1998a; Massicotte et al. 1989; St Louis and Massicotte 1985; Longo et al. 2003). The vasodilator effects in guinea pig and rat coronary arteries and in bovine-cultured smooth muscle cells are associated with increased NO synthesis (Bani et al. 1998a; Bani-Sacchi et al. 1995). In humans, there are vasodilator effects in gluteal resistance or subcutaneous arteries but little or no effect in pulmonary, myometrial, or placental vessels (McGuane et al. 2011; Fisher et al. 2002; Petersen et al. 1991). In gluteal arteries, the vasodilator responses likely involve NO and interestingly were influenced by the medication being taken by patients. Arteries from patients on angiotensin-converting enzyme (ACE) inhibitors showed marked attenuation of the vasodilator response to relaxin, effects that were further enhanced by inhibition of cyclooxygenase (Fisher et al. 2002).

Relaxin is the major renal vasodilator responsible for increases in renal plasma flow and glomerular filtration rate (GFR) during rat pregnancy (Novak et al. 2001), but relaxin also has similar effects in nonpregnant female rats and male rats (Danielson and Conrad 2003; Danielson et al. 1999; Bogzil et al. 2005). In humans, the evidence is less compelling although women who are pregnant through egg donation (relaxin deficient) display lower GFR than in normal pregnancies (Smith et al. 2006a). In human volunteers, relaxin (5 h) increased renal plasma flow, by ∼75%, and fractional sodium excretion, by ∼25% but did not affect GFR (Smith et al. 2006b). However, during chronic relaxin infusion in the scleroderma trials, the GFR rose significantly (Seibold et al. 2000; Khanna et al. 2009). It should be noted that the vasodilator effects of relaxin may also influence creatinine handling in the gut and that this is often upregulated in scleroderma or CHF. The renal effects of relaxin in humans therefore remain to be confirmed.

The vasodilator mechanisms suggested for relaxin in mammals involve activation of NOS (Conrad and Novak 2004; Nistri and Bani 2003), VEGF, placental growth factor (PGF), matrix metalloproteinases, and ETB receptors (Dschietzig et al. 2003; Novak et al. 2002) and modification of the extracellular matrix of the vessel walls (McGuane et al. 2011; Jeyabalan et al. 2003; Lekgabe et al. 2005; Xu et al. 2010). These responses have a distinct temporal hierarchy with acute (minutes) responses being endothelium dependent and blocked by NOS inhibitors, by the PI3K inhibitors Wortmannin and LY294002, and by PTX pretreatment but not by the VEGF receptor antagonist SU5416 (McGuane et al. 2011). The effects are consistent with Gβγ activation of PI3K, Akt, and eNOS.

Longer-term (hours) vasodilator responses to relaxin involve endothelial ETB receptors that release NO (Dschietzig et al. 2003; Jeyabalan et al. 2003). Two mechanisms involving the relaxin-ETB receptor pathway have been found. In rodent and human vessels of ∼100–300 μm diameter, relaxin activates MMP-9 and later also MMP-2 to generate ET-1(1-32), rather than ET-1(1-21), from big ET-1 (Jeyabalan et al. 2003). ET-1(1-32) then preferentially binds to the endothelial ETB receptor causing NO release. In blood vessels from pregnant or relaxin-treated nonpregnant rats, pro-MMP-2 and MMP-2 activity and pro-MMP-2 protein and mRNA are increased (Jeyabalan et al. 2003, 2006). These findings support those obtained with the ETB antagonist RES-701-1 that blocks renal hemodynamic changes produced by relaxin (Danielson et al. 2000) in rats and antagonizes inhibition of renal artery smooth muscle produced by relaxin or in pregnancy (Novak et al. 2002). Similar effects were produced in vitro by the ET receptor antagonist SB209670 but not by the ETA selective BQ123 (Gandley et al. 2001). An alternative mechanism, which occurs mainly in larger vessels and in the pulmonary circulation (Dschietzig et al. 2001a, b, 2003), involves relaxin-induced upregulation of endothelial ETB receptor (Dschietzig et al. 2003). Thus, relaxin acts as a functional ET-1 antagonist because the endothelial ETB receptor mediates NO production and, equally importantly, ET-1 clearance (Dschietzig et al. 2001b, 2003).

Evidence also exists for a role of VEGF in the longer-term vasodilator actions of relaxin. In human endometrial cells, relaxin increases cAMP levels and VEGF expression, and the effects are prevented by AC inhibition and mimicked by forskolin or a PDE inhibitor (Unemori et al. 1999). Relaxin promotes angiogenesis (Unemori et al. 1999, 2000) by inducing VEGF and basic fibroblast growth factor synthesis. These effects are likely important in wound healing (Bitto et al. 2013) and also in heavy, irregular or prolonged menstrual bleeding that accompanies relaxin administration (Unemori et al. 1999). Relaxin also increases arterial compliance by its effects on geometric remodeling (increases in unstressed wall area and wall-to-lumen area ratio) and compositional remodeling (decrease in collagen-to-total protein ratio) in certain arteries (Debrah et al. 2011; Chan and Cipolla 2011; Gooi et al. 2013). These properties expedite tissue and organ perfusion in the long term.

Compared with arteries, remarkably little is known of the effects of relaxin on veins. It has been reported that the vasodilator potency of relaxin is lower in veins than arteries (Li et al. 2005). However, it is becoming clear that the actions of relaxin on both endothelium and smooth muscle can vary considerably between vascular beds. Human umbilical arterial endothelial cells (HUAEC) do not contain detectable RXFP1 and do not respond to relaxin, whereas venous endothelial (HUVEC), smooth muscle (HUVSMC), and arterial smooth muscle cells (HUASMC) contain the receptor and respond to relaxin with increases in cGMP and cAMP (Sarwar et al. 2015). Interestingly in cocultures of endothelial and smooth muscle cells, particularly large cGMP responses are observed when HUVECs are paired with HUVSMC suggesting that venodilation may be a significant vascular response to relaxin. In addition it was established that human coronary artery endothelial cells (HCAEC) but not HUVEC not only release NO to activate cGMP but secrete prostanoids to increase cAMP in cocultured SMCs (Sarwar et al. 2016). Other variations include the observation that rat mesenteric veins, in contrast to arteries, are not remodeled by relaxin (Jelinic et al. 2014). It is also important to recognize that venous tone is significantly more dependent on sympatho-adrenergic innervation and pacemaker cells (Cajal cells) than arterial tone (Gelman 2008). Given the interest in the utility of vasodilators in the treatment of cardiac failure (Bhushan et al. 2014), venodilator responses to relaxin promises to be an important future area of research.

Heart: Rat atria contain relaxin binding sites (Osheroff et al. 1992; Osheroff and Ho 1993), and relaxin was shown subsequently to be a potent inotrope in left and chronotrope in right atria (Bani-Sacchi et al. 1995; Kakouris et al. 1992; Coulson et al. 1996; Thomas and Vandlen 1993; Toth et al. 1996; Mathieu et al. 2001; Tan et al. 1998; Wade et al. 1994; Ward et al. 1992). Chronotropic responses to relaxin are accompanied by the secretion of atrial natriuretic peptide in rat isolated perfused hearts (Toth et al. 1996). In rat atrial myocytes, relaxin inhibits outward potassium currents, increases action potential duration, and enhances calcium entry to produce the positive inotropic effect (Piedras-Renteria et al. 1997a, b). The inotropic effects of relaxin also occur in human atria (but not in ventricular myocardium) (Dschietzig et al. 2011) and are preserved in failing hearts and involve PKA, outward potassium currents, and PI3K.

Relaxin protects against myocardial injury caused by ischemia and reperfusion in rats (Bani et al. 1998b). These findings were confirmed in an in vivo pig model of myocardial infarction in which relaxin attenuated leukocyte recruitment and oxidative damage and improved contractile recovery (Perna et al. 2005). In a mouse infarction model, relaxin improves post-infarction remodeling by suppressing reactive fibrosis in vital myocardium, without affecting reparative fibrosis (scarring) within the infarcted area (Samuel et al. 2011). Relaxin attenuates hypertrophy in rat neonatal cardiomyocytes by inhibiting myofibroblast activation and the subsequent paracrine release of growth factors (Moore et al. 2007), confirming an earlier study in spontaneously hypertensive rats and suggesting that endogenous relaxin has anti-hypertrophic actions (Dschietzig et al. 2005). Atrial fibrillation, a common arrhythmia that develops during aging-related fibrosis, hypertension, or heart failure, is suppressed by relaxin in spontaneously hypertensive rats, an established model of end-organ damage (Parikh et al. 2013).

Relaxin has cardioprotective properties in chronic heart failure where the expression of relaxin is increased in both atria and ventricles to levels that correlate with the degree of failure (Dschietzig et al. 2001b; Fisher et al. 2003). Increased relaxin gene expression in chronic heart failure was also observed in two independent studies utilizing a rat infarction model (Kompa et al. 2002; Zhang et al. 2005). Relaxin has since been used to treat human heart failure, and in a hemodynamic pilot study in patients with stable chronic heart failure (Dschietzig et al. 2009c), a 24-h i.v. infusion of recombinant human relaxin markedly elevated cardiac index without affecting heart rate and decreased pulmonary wedge pressure, systemic, and pulmonary vascular resistance, without affecting systolic or mean arterial blood pressure or central venous pressure. Hemodynamic changes occurred within 45–60 min, and relaxin infusion improved creatinine and blood urea nitrogen clearance and was free of relevant side effects. The use of relaxin to treat acute heart failure (defined as new onset or, more frequently, worsening of known heart failure) is promising. In the Pre-RELAX (phase II) and RELAX (phase III) trials (Teerlink et al. 2009, 2013), relaxin moderately improved dyspnea (the primary endpoint), was exceptionally safe, improved renal function, and lowered all-cause as well as cardiovascular mortality at day 180. Since both trials were not powered statistically to assess mortality, another extensive adequately powered mortality trial in acute heart failure is in progress. If successful, relaxin would become the first evidence-based remedy for this highly challenging clinical syndrome.

Fibrosis: The anti-fibrotic properties of relaxin were the first biological effects to be recorded (Hisaw 1926), and attempts have been made to exploit this function therapeutically. Relaxin was safe, well tolerated, and effective in some patients in a phase II trial (Seibold et al. 2000) but failed to show clinical efficacy in a phase III trial (Erikson and Unemori 2001). The discrepancy between the effectiveness of relaxin in preclinical anti-fibrosis models and its failure in the phase III human scleroderma trial may stem from an incomplete understanding of the pathophysiology of scleroderma and by the enrolment of patients in the terminal stages of the disease. It is noteworthy that dermal fibrosis in the relaxin knockout mouse is rescued by exogenous relaxin at 6 but not 12 months of age (Samuel et al. 2005). There are many studies in animals that demonstrate that relaxin influences collagen turnover. In male relaxin knockout mice, there is an increased tissue fibrosis with age that is prevented (Samuel et al. 2007) or reversed in the lung (Samuel et al. 2003), kidney (Samuel et al. 2004a), and heart (Samuel et al. 2004b) by relaxin.

In the lung and in cardiac fibroblasts, relaxin reduces the expression of collagens I and III, increases MMP levels, and reduces fibrosis (Samuel et al. 2004b; Unemori et al. 1996). In renal fibroblasts, relaxin inhibits pro-fibrotic changes induced by TGF-β by activating the NO/guanylyl cyclase pathway and decreasing Smad2 phosphorylation and nuclear localization (Mookerjee et al. 2009). Relaxin has anti-fibrotic effects in rat renal fibrosis models (Garber et al. 2001; McDonald et al. 2003) and in spontaneously hypertensive rats (Lekgabe et al. 2005). In cardiac fibrosis induced by isoprenaline (Zhang et al. 2005) or in transgenic mice overexpressing cardiac β2-adrenoceptors (Bathgate et al. 2008), relaxin reduced cardiac fibrosis. In streptozotocin-treated mRen-2 rats (a model for diabetic cardiomyopathy), relaxin reduced left ventricular collagen, myocardial stiffness, and diastolic dysfunction (Samuel et al. 2008), accompanied by decreased TIMP1 expression and increased extracellular matrix-degrading MMP-13 (Samuel et al. 2008). Relaxin also has anti-fibrotic actions in the liver (Bennett 2009). However, in a particularly severe chronic pressure overload model in mice, relaxin was ineffective (Xu et al. 2008). Relaxin increases expression of MMP-9 and MMP-13 mRNA (Ahmad et al. 2012) in primary fibrochondrocytes that is associated with activation of PI3K, Akt, PKCζ, and ERK1/ERK2. Inhibition of these signaling pathways blocked MMP-9 induction and the anti-fibrotic effects. Increases in MMP-9 expression were also blocked by transfection of a dominant negative Akt or by siRNA knockdown of ERK1/ERK2, PKCζ, Elk-1, c-fos, and to a lesser extent NFκB (Ahmad et al. 2012). This study provides insights for the translation of the anti-fibrotic effects of relaxin into a clinical setting.

The anti-fibrotic properties of relaxin are clearly seen in disease conditions associated with excessive collagen deposition and are not simply a result of activation of RXFP1. In rat kidney myofibroblasts, the anti-fibrotic actions of relaxin are completely abolished by the AT2R antagonist PD123319 (Chow et al. 2014), and in an in vivo fibrosis model in mice, the protective effects of relaxin are lost when the AT2R is either absent (in AT2R-/y mice) or blocked by PD123319, confirming that the AT2R is obligatory for the anti-fibrotic actions. However, relaxin does not bind directly to the AT2R although RXFP1 does form constitutive heterodimers with AT2R that may mediate the downstream anti-fibrotic signaling pathways originally attributed to relaxin (Mookerjee et al. 2009; Chow et al. 2012; Heeg et al. 2005), which include inhibition of the TGF-β1/pSmad2 axis and reduced TGF-β1-induced collagen deposition. As AT2Rs are normally expressed at low levels in tissues (Jones et al. 2008) (Carey 2005; Matsubara 1998) and fibroblasts, but are dramatically increased in pathological conditions (Siragy and Carey 1997; Savoia et al. 2006), this may create an environment where RXFP1-AT2R heterodimerization is more likely to take place and explain why relaxin displays more pronounced anti-fibrotic effects under pathological conditions.

Relaxin also has beneficial effects in wound healing (Casten and Boucek 1958) that may involve vasodilator effects but also the synthesis of new blood vessels by enhancing the local production of VEGF (Unemori et al. 2000).

Organ protection: Relaxin protects from ischemia-reperfusion (IR) injury in the heart (see above), but also in rat liver (Boehnert et al. 2005, 2008, 2009) and kidney (Yoshida et al. 2013). In a similar model, relaxin decreases inflammatory cytokines, counteracts oxidative damage by increasing SOD-1 and SOD-2 expression, and ameliorates neutrophil-related injury as assessed by myeloperoxidase levels (Collino et al. 2013). In rat isolated lungs (Alexiou et al. 2010, 2013), relaxin causes inhibition of leukocytes and oxidative surge, as well as prevention of ET-1 stimulation resulting in less edema formation and lower pulmonary vascular pressure.

NO appears to be critically involved in organ protection (Alexiou et al. 2013; Masini et al. 1997; Collino et al. 2013; Alexiou et al. 2010). In rat lung, relaxin increases iNOS (Alexiou et al. 2013), whereas in rat kidney (Collino et al. 2013), it upregulates iNOS and activates eNOS via PI3K.

Formation and spread of tumors: Relaxin is expressed in endometrial (Kamat et al. 2006), mammary (Tashima et al. 1994), thyroid (Hombach-Klonisch et al. 2006), and prostate tumors (Feng et al. 2007; Thompson et al. 2006) and has been associated with breast cancer (Silvertown et al. 2003; Bani 1997) where relaxin treatment of breast cancer cells implanted into nude mice increases their invasive potential (Binder et al. 2002). In contrast, longer-term application of relaxin (up to 8 days) reduces mammary xenograft growth in mice (Radestock et al. 2008). The elevated serum relaxin levels in breast cancer patients are associated with metastatic disease (Binder et al. 2004). Relaxin is also associated with prostate cancer progression in the mouse xenograft model (Silvertown et al. 2006), where blocking relaxin or its receptor decreases cancer growth (Feng et al. 2010). In patients with prostate cancer, elevated relaxin levels are linked to cancer progression, metastasis, and androgen independence (Thompson et al. 2006). Likewise, relaxin heightens the collagenolytic potency of thyroid cancer cells by upregulating MMP-2 that facilitates greater in vitro invasiveness (Bialek et al. 2011). In human osteosarcoma cells, relaxin promotes faster in vitro growth, invasion, and angiogenesis via the Akt and VEGF pathways, and siRNA-mediated knockdown of relaxin mitigates these effects (Ma et al. 2013a, b). A clinical study in 108 patients with hepatocellular carcinoma identified expression of relaxin in tumors as associated with a poor prognosis (Pan et al. 2013). Similarly, high plasma relaxin levels were associated with a poor prognosis in 146 patients with esophageal squamous cell carcinoma (Ren et al. 2013). A recent study also suggests that C1q-tumor necrosis factor-related protein (CTRP8) interacts with RXFP1 to facilitate cell migration in brain cancer, dependent on activation of PI3K and PKC (Glogowska et al. 2013). In summary, relaxin does not appear to initiate cancer, but, like other growth and angiogenic hormones, it promotes cancer growth and/or spread by its actions on enhanced matrix degradation and angiogenesis. To date, this is clearly clinically relevant at least for mammary, thyroid, and prostate cancer.

Diabetes: There are studies that suggest that relaxin influences carbohydrate metabolism. Hypoglycemic episodes are not uncommon in insulin-dependent diabetic women during the first trimester of pregnancy, when circulating levels of relaxin are higher, and increased insulin may be required in the third trimester, when relaxin concentration are lowest (O’Byrne et al. 1978). Experimentally, intraperitoneal relaxin inhibits ad libitum feeding in male rats when given during the early dark phase of the circadian cycle (McGowan et al. 2010).

Relaxin concentrations in the serum of diabetic women are higher than those in nondiabetic women during each trimester of pregnancy (Steinetz et al. 1992; Whittaker et al. 2003). This may be a physiological response to the insulin resistance that occurs during pregnancy. More circumstantial findings show that circulating relaxin levels correlate positively with insulin sensitivity and inversely with β-cell function in women with type 2 diabetes mellitus (T2DM) (Szepietowska et al. 2008) and increased plasma relaxin after successful anti-diabetic therapy in men with T2DM (Schondorf et al. 2007). The cellular mechanism may involve a relaxin-mediated increase in insulin binding to its receptors on adipocytes (Olefsky et al. 1982; Jarrett et al. 1984). In rat cardiac fibroblasts, high glucose increases the expression of relaxin (Wang et al. 2009). In human amniotic epithelial cells, relaxin enhances expression of insulin-like growth factor-2 (Millar et al. 2003). In insulin-resistant lean mice following a high-fat diet, relaxin increases glucose uptake into skeletal muscle (Bonner et al. 2013), and in a genetic mouse model of T2DM, relaxin given over 12 days lowered blood glucose levels (Bitto et al. 2013). Relaxin has also been shown to activate PPARγ and enhance the actions of rosiglitazone in cells expressing RXFP1 (Singh and Bennett 2010). Thus, relaxin treatment would be expected to influence insulin sensitivity.

INSL3 and RXFP2

Reproduction: INSL3 is primarily a reproductive hormone, first cloned from testicular cDNA libraries (Adham et al. 1993; Pusch et al. 1996) and shown to be highly expressed in testicular Leydig cells (Adham et al. 1993; Pusch et al. 1996). Male INSL3 knockout mice (Nef and Parada 1999; Zimmermann et al. 1999) or RXFP2 knockout mice (Overbeek et al. 2001; Gorlov et al. 2002) are infertile and bilaterally cryptorchid. The INSL3/RXFP2 system is therefore essential for the development of the gubernaculum during embryogenesis and for normal transabdominal testicular descent.

INSL3 is highly expressed in testicular Leydig cells in adult mammals and is an important marker of Leydig cell function (Ivell and Anand-Ivell 2009; Ivell et al. 2014). INSL3 is a circulating hormone in males with levels rising in puberty and being maintained in adults (Foresta et al. 2004). The function of INSL3 in adult males is unknown; however, it has been reported to have potential endocrine functions on bone metabolism (see below) and roles in germ cell function. RXFP2 receptors are expressed on germ cells, and studies in male rats suggest that it is a paracrine factor affecting male germ cell survival (Kawamura et al. 2004). RXFP2 receptors are also expressed on Leydig cells, and recent studies suggest that INSL3 influences steroid production by activating RXFP2 on Leydig cells (Kawamura et al. 2004; Anand-Ivell et al. 2006; Filonzi et al. 2007) (Pathirana et al. 2012).

In females, INSL3 is produced in the thecal cells of the ovary, and in the corpus luteum (Ivell and Bathgate 2002) and in mice, Insl3 expression is higher in the follicular phase of the cycle (Zimmermann et al. 1997). Female INSL3 knockout mice have longer estrous cycle length, impaired fertility, smaller litter sizes, accelerated follicular atresia and luteolysis, and premature loss of corpora lutea (Nef and Parada 1999; Spanel-Borowski et al. 2001). In women, INSL3 is a circulating hormone, and levels are indicative of the number of ovarian antral follicles recruited and growing within a follicular wave (Anand-Ivell et al. 2013). As INSL3 is a product of the thecal cells, plasma levels are elevated in polycystic ovary syndrome (PCOS) (Anand-Ivell et al. 2013; Szydlarska et al. 2012; Gambineri et al. 2007). A recent study in cows demonstrated that INSL3 has a positive autoregulatory role in maintaining the thecal androgen production essential for normal ovarian follicle development (Glister et al. 2013).

Central nervous system: RXFP2 expression occurs in human brain as shown by RT-PCR (Hsu et al. 2002) and in the rat parafascicular nucleus, dorsolateral, ventrolateral, and posterior thalamic nuclei and medial habenula by in situ hybridization histochemistry (Shen et al. 2005) (Sedaghat et al. 2008). These sites map with 125I-INSL3 binding (Sedaghat et al. 2008). Insl3 expression has been demonstrated in the rat brain and in the bovine hypothalamus (Bathgate et al. 1996, 1999). The effects of INSL3 administration to rat brain suggest that RXFP2 influences sensorimotor function (Sedaghat et al. 2008).

Bone and other tissues: Physiological functions of INSL3 in other tissues postulated on INSL3 and RXFP2 expression patterns include cryptorchid hypogonadism (T222P mutation of RXFP2), where 64% had reduced bone density but normal plasma testosterone levels (Ferlin et al. 2008). RXFP2 mRNA and protein occur in human osteoblasts, and an osteoblast cell line responded to INSL3 with a concentration-dependent increase in cAMP. Mouse osteoblasts also express RXFP2, but not Insl3, and RXFP2 knockout mice are osteopenic and have functional osteoblast impairment (Ferlin et al. 2008). INSL3 regulates expression of genes required for proliferation and differentiation, matrix deposition, and osteoclastogenesis in cultured human osteoblasts (Pepe et al. 2009). Deficient INSL3/RXFP2 signaling is correlated with reduced bone mass (Ferlin et al. 2008, 2009) suggesting that INSL3 has a role in bone physiology and that RXFP2 mutations may be linked with osteoporosis in men (Ferlin et al. 2013). Recent studies indicate that RXFP2 is associated with horn growth in sheep (Johnston et al. 2013) and cows (Wiedemar et al. 2014), suggesting actions of INSL3 on the bone are common features in male mammals.

Tumors show altered expression of INSL3, and a splice variant has been described in human hyperplastic thyroid adenoma and thyroid cancer (Hombach-Klonisch et al. 2003). INSL3 is also present in prostate carcinomas and increases motility in the human androgen-insensitive prostate carcinoma cell line PC-3 (Klonisch et al. 2005). All human thyroid adenomas, three types of human thyroid carcinomas, and mouse noncancerous follicular epithelial cells of the thyroid express RXFP2 mRNA (Hombach-Klonisch et al. 2010). Thyroid cancer cells expressing INSL3 demonstrate enhanced motility and increased colony formation in vitro and enhanced tumor growth in vivo. Finally, INSL3 increases levels of the calcium-binding protein S100A4, which increases cancer cell mobility and enhances tumor tissue vascularization, suggesting that S100A4 is a downstream target of RXFP2 signaling in thyroid cells (Hombach-Klonisch et al. 2010).

In the kidney, RXFP2 is expressed in mesangial cells in mature glomeruli and inhibits proliferation of cultured primary glomerular cells, suggesting that INSL3 and RXFP2 influence the genesis or maturation of renal glomeruli and regulate mesangial cell density (Fu et al. 2006). Pod1 is a transcription factor involved in glomerulogenesis (Quaggin et al. 1999), and an E-box consensus sequence capable of binding Pod1 and other helix loop helix transcription factors is located upstream of the gene for RXFP2 (Funato et al. 2003). RXFP2 expression levels in the embryonic kidney were significantly greater in Pod1 knockout mice than in heterozygous or wild-type controls, indicating that RXFP2 is downstream of Pod1 and Pod1 negatively regulates expression of RXFP2 in the glomeruli (Familari et al. 2009).

Although the INSL3 and RXFP2 system has primarily been studied for its specialized roles in both male and female reproductive physiology, there is emerging evidence to suggest that it may have roles in the CNS, cancer metastasis, bone physiology, and the kidney.

Summary

RXFP1 is coupled to cAMP and many other signaling pathways in different cell types. Linking these effectors of RXFP1 activation to specific physiological end points should allow the design and development of targeted therapies. RXFP1 has therapeutic potential for treatment of fibrosis and cancer metastasis and is currently in extended phase III clinical trials for heart failure. Although not as extensively studied, RXFP2 appears to have a simpler physiological role and fewer downstream effectors compared to RXFP1. RXFP2 has possible therapeutic potential for the treatment of some types of cryptorchidism and could be used to control fertility; however, more extensive research is required to assess its true therapeutic potential.

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Copyright information

© Springer International Publishing AG 2018

Authors and Affiliations

  • Roger J. Summers
    • 1
  • Michelle L. Halls
    • 1
  • Ross A. D. Bathgate
    • 2
  1. 1.Drug Discovery Biology, Monash Institute of Pharmaceutical SciencesMonash UniversityParkvilleAustralia
  2. 2.Neuropeptides Division, Florey Institute of Neuroscience and Mental Health and Department of Biochemistry and Molecular BiologyUniversity of MelbourneParkvilleAustralia