Next Article in Journal
Identification of Circular RNAs in Hypothalamus of Gilts during the Onset of Puberty
Next Article in Special Issue
WNT-FRIZZLED-LRP5/6 Signaling Mediates Posterior Fate and Proliferation during Planarian Regeneration
Previous Article in Journal
Analysis of Transcriptional Changes in Different Brassica napus Synthetic Allopolyploids
Previous Article in Special Issue
The ADAR Family in Amphioxus: RNA Editing and Conserved Orthologous Site Predictions
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Genes
Volume 12
Issue 1
10.3390/genes12010083
genes-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Nuclear Receptors and Development of Marine Invertebrates

by
Angelica Miglioli
1,2,
Laura Canesi
2,
Isa D. L. Gomes
1,
Michael Schubert
1 and
Rémi Dumollard
1,*
1
Laboratoire de Biologie du Développement de Villefranche-sur-Mer (LBDV), Institut de la Mer de Villefranche, Sorbonne Université, CNRS, 181 Chemin du Lazaret, 06230 Villefranche-sur-Mer, France
2
Dipartimento di Scienze della Terra, dell’Ambiente e della Vita (DISTAV), Università degli Studi di Genova, Corso Europa 26, 16132 Genova, Italy
*
Author to whom correspondence should be addressed.
Genes 2021, 12(1), 83; https://0-doi-org.brum.beds.ac.uk/10.3390/genes12010083
Submission received: 14 December 2020 / Revised: 31 December 2020 / Accepted: 4 January 2021 / Published: 11 January 2021
(This article belongs to the Special Issue The Evolution of Invertebrate Animals)
Browse Figures
Review Reports Versions Notes

Abstract

:
Nuclear Receptors (NRs) are a superfamily of transcription factors specific to metazoans that have the unique ability to directly translate the message of a signaling molecule into a transcriptional response. In vertebrates, NRs are pivotal players in countless processes of both embryonic and adult physiology, with embryonic development being one of the most dynamic periods of NR activity. Accumulating evidence suggests that NR signaling is also a major regulator of development in marine invertebrates, although ligands and transactivation dynamics are not necessarily conserved with respect to vertebrates. The explosion of genome sequencing projects and the interpretation of the resulting data in a phylogenetic context allowed significant progress toward an understanding of NR superfamily evolution, both in terms of molecular activities and developmental functions. In this context, marine invertebrates have been crucial for characterizing the ancestral states of NR-ligand interactions, further strengthening the importance of these organisms in the field of evolutionary developmental biology.

1. Introduction

Nuclear receptors (NRs) are a superfamily of phylogenetically related transcriptional regulators and act as activators or repressors of gene transcription, either constitutively or depending on the binding of a ligand [1,2]. NRs are characterized by a conserved structural organization, comprising a variable N-terminal region (A/B domain), a DNA-binding domain (DBD, C domain), a hinge region (D domain), a ligand-binding domain (LBD, E domain) and a C-terminal domain (Figure 1) [3].
The LBD permits ligand binding, mostly through Van der Waals interactions and hydrogen bonds of specific amino acid residues in the ligand-binding pocket (LBP) [4]. The DBD is the region of the NR that mediates interactions with DNA and hence controls the specificity of the transcriptional response. NRs bind DNA at so-called response elements (REs), specific sequences located in the cis-regulatory regions of target genes [5]. REs consist of direct, inverted or palindromic repetitions of a core consensus motif separated by a variable number of nucleotides that are bound by NRs as monomers, homodimers or heterodimers [6]. Efficient dimeric complexes are formed in vivo through cooperative protein-protein and protein-DNA interactions [7]. While protein-protein dimerization interfaces are found in the DBD, the LBD and the hinge region of NRs, REs serve as NR dimerization sites on DNA [8,9,10].
NRs are divided into two main classes: NRs with known ligands and NRs, for which ligands do not exist or have yet to be identified, the so-called orphan receptors [11,12]. Recent phylogenetic analyses of DBD and LBD sequences defined nine NR subfamilies (Table 1), which have probably originated from a single ancestral NR [2,12,13,14,15,16].
NR genes have been found in all extant metazoan taxa, but not in fungi, plants or unicellular eukaryotes, suggesting that NRs originated at the base of the metazoans [1,2]. NR phylogenies have indicated that the ancestral receptor might have been a ligand-activated receptor, with fatty acids as possible ligands [15,16]. In the course of evolution, the NR superfamily has experienced a complex pattern of gene expansions that led to the current diversity of metazoan NRs (Figure 2) [17,18,19,20,21,22,23,24,27].
Genomes of placozoans contain members of NR subfamilies 2 and 3, suggesting that a first wave of NR gene expansion occurred relatively early in metazoan evolution [20,21,29,30]. The NR complement of cnidarians indicates that the diversification of the NR2 and the appearance of the NR6 and NR7/8 subfamilies predate the split of the cnidarian and bilaterian lineages (Figure 2) [20,27,30]. Another NR expansion occurred very early in bilaterian evolution, leading to the appearance of all extant NR subfamilies and an estimated complement of about 25 NR genes (Figure 2) [1,15,27]. The NR superfamily subsequently experienced lineage-specific expansions. The NR3C receptors, for example, arose at the base of the chordates and whole genome duplications further expanded the NR complement in the vertebrate lineage (Table 1, Figure 3) [31,32,33]. A specific lineage-specific duplication has further been reported in cephalochordates, whose genomes encode 10 NR1H receptors (Figure 3) [14]. Similarly, in nematodes, the orphan NR HNF4 experienced a lineage-specific burst of duplications (Figure 3) [34].
Gene losses have also been important for shaping the NR complements of extant metazoans. Ecdysozoans, for example, have lost a whole suite of NRs, including ER, THR, PPAR and NR7/8. RAR was further lost by arthropods and nematodes, and ERR by nematodes only (Figure 3) [24,27,35]. Similarly, within the chordates, NR0A genes have been lost during early diversification of the lineage, tunicates and vertebrates have lost NR7/8, and the tunicates have been subjected to additional NR losses, including, for example, ER, TLL/TLX and NR3C (Figure 3) [14,24,27,31,36].
Despite their presence in all metazoans, NR functions and signaling pathways have primarily been characterized in vertebrates where they were shown to be involved in a wide variety of biological processes, one of which is the regulation of embryonic and post-embryonic development [2,37]. NR names and classifications into orphan or ligand-activated receptors thus strictly reflect their functions and ligand binding properties in vertebrates, with limited relevance to other animal taxa [11,16,25]. In vertebrates, ligand-activated NRs are mainly found in the NR1 and NR3 subfamilies, and their endogenous ligands establish a diverse list of bioactive compounds (Table 1) [25,26]. Furthermore, NR1 and NR4 receptors generally exert their biological functions in heterodimeric complexes with RXR [38,39]. RXR is an orphan nuclear receptor of the NR2 subfamily, whose main biological function is to act as a permissive heterodimeric binding partner of NR1 and NR4 subfamily members, in a process commonly referred to as RXR subordination [39]. Conversely, NR2, NR3, NR5 and NR6 subfamily members mostly function in homodimeric complexes [6].
Given their ligand-dependent activity and their involvement in life threatening human pathologies, such as diabetes and cancer, NRs are major pharmacological targets in several different drug discovery programs [40]. Unfortunately, their ligand-dependent activity also makes them susceptible to a class of environmental pollutants defined as endocrine disrupting chemicals (EDCs), exogenous substances that alter the function of the endocrine system [41,42]. EDCs mimic the structure of endogenous NR ligands and can act as either agonists or antagonists of NR signaling pathways [42,43]. EDC exposure thus adversely affects the adult endocrine system and impacts developmental processes, leading to pathophysiological and pathological conditions, such as neurodevelopmental disorders and developmental dysfunction/arrest in all vertebrate taxa [42,43]. It was initially thought that invertebrates are not affected by EDCs as they were said to lack an endocrine system similar to that of vertebrates [44,45]. However, it is now known that invertebrates are extremely sensitive to EDC exposure, in particular during their embryonic and post-embryonic development [44,45,46]. For this reason, a specific set of NRs, including thyroid hormone (THR), retinoic acid (RAR), retinoid X (RXR) and estrogen (ER, ERR) receptors, is starting to be characterized in invertebrates in an effort to understand their functional traits in relation to those of their vertebrate orthologs. Even though it is still not clear to what extent the teratogenic effects of EDCs in invertebrates are mediated by NRs, it has been shown that NRs are involved in embryonic and post-embryonic development in these animals [47,48,49,50,51,52,53]. Yet, the mechanisms at play are not necessarily conserved with vertebrates, revealing lineage-specific adaptions in both invertebrates and vertebrates [1,10,17,47,48,49,50,51,52,53,54,55,56,57]. The aim of this review is to describe and discuss the developmental functions of NRs in invertebrates, with a special focus on marine organisms, highlighting the particular importance of comparative approaches using emerging marine invertebrate models for the field of evolutionary developmental biology.

2. Development of Marine Invertebrates and Associated NR Cohorts

In this section, we will correlate NR expression with the development of marine invertebrates, focusing on representatives from three phyla: cnidarians, mollusks and chordates. Marine invertebrate development can generally be classified into four stages: embryonic development, embryo to larva transition (EtL), larval development and metamorphosis. Embryonic development is characterized by cell proliferation and germ layer specification [47]. EtL consists of a series of morphogenetic processes resulting in the formation of a primitive larva. These processes include axial and body patterning, initiation of neurogenesis and organogenesis [58]. During larval development, additional structures for feeding, light sensing and swimming/crawling are formed [59]. Most of these structures will subsequently be lost during metamorphosis, which results in the emergence of the adult body plan [60]. This typical life cycle can be completed by an asexual reproduction phase. Juvenile polyps of Medusozoan cnidarians, for example, can undergo strobilation, leading to segmentation of the polyp into so-called ephyras, which will subsequently grow into adults [61]. Furthermore, several tunicate lineages independently gained the capacity for asexual reproduction using different budding strategies [62,63].
As detailed above, NR complements can vary greatly between different marine invertebrate phyla. Cnidarians, for instance, possess only a limited number of NRs [20,30,61,64]. The genome of the cnidarian Nematostella vectensis, an anthozoan, was thus estimated to encode 17 NRs, including orthologs of vertebrate COUP-TF, TLX/PNR, HNF4, TR2/4 and GCNF [30]. The genomes of medusozoan cnidarians, such as Aurelia aurita, include additional NRs, such as orthologs of vertebrate RXR and an ER-like NR, called NR3E (Figure 2; Figure 3) (Table 1) [20,64,65]. In comparison, mollusks have larger NR complements with representatives of each NR subfamily. In the genomes of Crassostrea gigas, Biomphalaria glabrata and Lottia gigantea, for example, 43, 39 and 33 genes encoding NRs were respectively identified, with most of them being orthologous to vertebrate NRs [19,23,24]. While the stereotypical NR complement of chordates is similar to that of mollusks, tunicates are characterized by significantly reduced NR complements. The ascidian tunicate Ciona intestinalis, for example, only has 17 NR genes [36].
It is intriguing to speculate that differences in NR complements are correlated with the diversity of life cycles, developmental strategies and reproductive adaptations observed in marine invertebrates [66]. NRs are essential regulators of vertebrate development [67,68]. Given that a certain number of developmental processes are conserved in metazoans (or at least bilaterians), it is conceivable that NRs are also pivotal regulators of embryonic and post-embryonic development in invertebrates [48,69]. However, developmental expression of NRs has only been established in a limited number of marine invertebrates, and their developmental functions in marine invertebrates are chiefly unknown [30,47,48,53,65].
In N. vectensis (Figure 4), 13 of the 17 NRs are dynamically expressed during development, with distinctive temporal patterns during embryogenesis, EtL, larval development and metamorphosis. While homologs of COUP-TF and TLX/PNR are highly expressed during EtL, HNF4, TR2/4, COUP-TF and NR7/8 expression is characteristic for larval development. Intriguingly, although GCNF is detectable throughout development, its expression increases during late development and marks the metamorphic stage [30]. Furthermore, in A. aurita, RXR expression peaks during strobilation (Figure 4) [64,65]. In the bivalve mollusk C. gigas, 34 of the 43 NRs are characterized by dynamic expression patterns during development (Figure 4). Orthologs of several NRs, including 2DBD-NRγ, HNF4 and NR7/8, are strongly expressed during embryonic development, with transcripts being almost exclusively of maternal origin [24,47]. While EtL is characterized by a number of NR transcripts including THR, RAR, PPAR, RXR, Rev-ErbA, TLL/TLX and PNR, larval development is marked by a general downregulation of NR expression [24,47,70]. Metamorphosis, in turn, is characterized by NR0B, 2DBD-NRδ, EcR, RXR, COUP-TF/SVP, TLL/TLX, ER and ERR expression [47]. Intriguingly, various members of the C. gigas-specific NR1P subgroup are dynamically expressed during development [17,47]. In the ascidian tunicates C. intestinalis and Phallusia mammillata, a principal component analysis of NR transcripts has not yet been performed. However, the expression profiles of ascidian NRs available in the Aniseed database (https://www.aniseed.cnrs.fr/) allow at least some level of developmental clustering (Figure 4). Embryonic development is thus characterized by expression of TR2/4, RXR, PPAR and LXR, all of which are of maternal origin. EtL is associated with RAR, HNF4, GCNF and ROR expression, while peaks in the expression of THR, Rev-Erb, ERR, COUP-TF and PPAR occur during larval development [53]. Altogether, these results demonstrate that the development of marine invertebrates is characterized by the stage-specific expression of NR subsets, suggesting that NR activity is correlated with distinctive developmental processes at different stages of the life cycle.

3. Development of Marine Invertebrates and NR Diversification

Despite the gene duplications and losses that accompanied NR diversification, a basic set of NRs is implicated in the development of distantly related metazoans. This basic set of NRs comprises COUP-TF/SVP, TLX/PNR, HNF4 and RXR (Figure 5, Table 1). Information regarding their developmental functions in marine invertebrates is still extremely scarce and is largely derived from the biological activity of their vertebrate orthologs.

3.1. Chicken Ovalbumin Upstream Promoter Transcription Factor, COUP-TF

In vertebrates, COUP-TF generally acts as a transcriptional repressor and regulates the development of muscles and heart as well as the differentiation of hindbrain and photoreceptors [71,72]. COUP-TF orthologs are expressed in early and late larval stages of N. vectensis and in late pre-metamorphic stages of both C. gigas and C. intestinalis (Figure 4). As this receptor is expressed in nematoblasts and subsets of neural cells in the cnidarian Hydra vulgaris as well as in the posterior photoreceptive ocellus of P. mammillata, COUP-TF is considered as a conserved neural marker involved in the formation of photoreceptive organs [53,73].

3.2. Tailless/Photoreceptor Cell-Specific Nuclear Receptor, TLX/PNR

In vertebrates and fruit flies, TLL/TLX is an orphan receptor involved in eye and forebrain development as well as in anteroposterior patterning of the embryo, suggesting some level of functional conservation between vertebrates and invertebrates [48,74]. In both N. vectensis and C. gigas, TLX/PNR orthologs are highly expressed in EtL stages (Figure 4). Their predominant expression in neural tissues is indicative of functions in neurogenesis and in the development of photoreceptive organs [30,47]. While TLX/PNR receptors were secondarily lost in tunicates, the TLL/TLX ortholog of the cephalochordate amphioxus is dynamically expressed in EtL stages. In neurulae, the amphioxus TLL/TLX gene marks developing sensory neurons, and in larvae, the gene is expressed in the central nervous system and anterior notochord [75]. It is thus likely that TLX/PNR genes are also involved in neurogenic processes in developing invertebrate chordates.

3.3. Hepatocyte Nuclear Factor 4, HNF4

In humans, HNF4 binds endogenous fatty acids as ligands and regulates hepatocyte differentiation, energy metabolism, xenobiotic detoxification and stem cell maintenance in the germ line [76,77]. HNF4 also participates in primary endoderm development in frogs, regulates expression of transcription factors necessary for endoderm specification in mice and is required for gut formation in insects [78,79,80]. HNF4 is highly expressed during larval development in N. vectensis, is a maternal transcript in C. gigas early embryos and expressed at EtL stages in C. intestinalis (Figure 4) [30,47]. In P. mammillata, HNF4 is expressed in endoderm cells of the trunk [53]. Developmental expression of HNF4 could thus play a role in endoderm specification and the formation of endodermal organs [30,47,53].

3.4. Retinoid X Receptor, RXR

In vertebrates, RXR is commonly the silent heterodimeric partner of NR1 and NR4 subfamily members and is involved in a variety of developmental processes in subordination to its heterodimeric binding partners. In the medusozoan cnidarian A. aurita, RXR is dynamically expressed during early strobilation, suggesting a potential role for RXR in this asexual reproduction process (Figure 4) [65]. RXR is highly expressed in C. gigas EtL stages, together with the NR1 subfamily members THR, RAR and PPAR [47,70]. In addition, there is a second peak of RXR expression prior to metamorphosis, which is paralleled by the NR1 subfamily member EcR [47]. Conversely, RXR in C. Intestinalis is expressed mainly during embryonic development, together with PPAR, and less strongly during EtL and larval stages, which are, respectively, characterized by the expression of the NR1 subfamily members RAR and ROR and THR, Rev-erb and PPAR [53]. While the developmental clustering of RXR does not strictly follow that of the NR1 subfamily members, there is nonetheless a tendency for regrouping RXR expression with that of the representatives of the NR1 subfamily. The notable absence of NR4 subfamily members from the developmental clusters in cnidarians, mollusk and tunicates suggests that heterodimers of RXR and NR1 receptors might play much more important roles during invertebrate development than heterodimers of RXR and NR4 receptors.
The receptors defining the basic set of NRs acting during development of marine invertebrates are all members of the NR2 subfamily. This subfamily has appeared very early in the metazoan lineage and has diversified before the cnidarian-bilaterian split [27,30]. Of all NRs, the NR2 subfamily member HNF4 is actually considered the extant NR that most closely resembles the ancestral NR that originated in the last common ancestor of all metazoans [15,16]. Based on the observations detailed above, it thus seems likely that the early diversification of the NR2 subfamily was accompanied by the elaboration of a NR2-dependent gene regulatory network involved in different aspects of animal development [81]. If this hypothesis is correct, at least some elements of this core gene regulatory network should still exist in extant animals and control conserved developmental functions in different metazoans. The basic set of NRs identified here and their potential involvement in invertebrate development could serve as a starting point for future studies aimed at identifying these ancestral NR-dependent features of animal development.

4. Thyroid Hormone Receptor (THR) Signaling Regulates Developmental Transitions in Marine Invertebrates

In vertebrates, THR is a ligand-activated transcription factor and its ligands are generally referred to as thyroid hormones (THs), which are either synthesized endogenously or taken up from the environment (Table 1) [82,83]. The main THs of vertebrates are triiodothyronine (T3) and tetraiodothyronine (T4), with T3 being the biologically active TH. THs are key regulators of vertebrate development and homeostasis and are involved, for example, in animal growth and metabolism as well as in the regulation of metamorphosis [84]. THR is dynamically expressed during development of both mollusks and tunicates, which is suggestive of a possibly conserved function in bilaterian development (Figure 5). THR genes originated at the base of bilaterians and have already been identified in the genomes of a wide variety of protostomes and deuterostomes [10,52,85]. However, the THR gene was lost in ecdysozoans and lineage-specific THR duplications occurred, for example, in platyhelminths, likely by independent duplication events in trematodes and turbellarians (Figure 3) (Table 2) [27,85].
Ascidian tunicates possess a THR ortholog and can endogenously produce T4 [86,87,88]. T4 is present in pre-metamorphic stages, and experimental evidence suggests that it could be a regulator of metamorphosis [87,88]. However, a direct involvement of THR in this process remains elusive [86,87]. In the cephalochordate amphioxus, several elements of the THR signaling system are shared with vertebrates, but the biologically active TH is triiodothyroacetic acid (TRIAC), rather than T3 [89,90]. TRIAC binds and strongly activates the amphioxus THR, and TH-dependent signaling plays a pivotal role in the regulation of metamorphosis [89,90]. In echinoderms, a THR gene has been cloned from sea urchins, but it has been shown that this receptor is not activated by THs or their metabolites [52,83]. Yet, T3 and T4 are actively accumulated during echinoderm development, and an exogenous supply of T4 can accelerate development, skeletogenesis and metamorphosis in a variety of echinoderm species [52,91].
Ligand-controlled THR signaling has further been suggested to regulate development and metamorphosis in different protostomes. However, in the absence of convincing evidence of endogenous TH synthesis in protostomes, it is currently believed that protostomes have to take up THs, or its precursors, from external sources [52,54,92]. In annelids, the THR of Platynereis dumerilii is activated in the presence of T3 or TRIAC [10,54]. Treatments with exogenous T3 or TRIAC induce an acceleration of the morphological switch from the trochophore to the crawling larva. This morphological switch is further characterized by a peak of THR expression [10,54]. The THR of the annelid P. dumerilii might thus be a ligand-activated receptor with endogenous ligands similar to T3 or TRIAC that mediates developmental transitions between larval stages [54,93].
In mollusks, experimental evidence also points to a morphogenetic function of THR and THs. While the transcriptional activity of C. gigas THR is not stimulated at relevant physiological concentrations of T4, T3 or TRIAC in vitro, both T4 and T3 are present in vivo in embryos and larvae, with their concentrations increasing significantly between the gastrula and the feeding larva [94]. In addition, the THR protein is detectable from blastula to trochophore stages in C. gigas, suggesting that the THR and the TH signaling system might be involved in the regulation of the embryo to larva transition [94]. Similarly, although platyhelminth THRs are not activated by THs, exogenous THs accelerate the development of parasitic platyhelminth lineages [85,95,96]. Taken together, it seems likely that THs have an ancestral role in lophotrochozoan development and that THRs are involved in mediating these roles. However, the endogenous ligands of lophotrochozoans THRs seem to be different from those of vertebrate THRs.
THs were further shown to function in developmental transitions in animals that lack a THR, such as cnidarians where THs regulate metamorphosis and strobilation [52,83]. A role for TH signaling in the control of developmental transitions might thus predate the evolutionary origin of THRs [52,54,83]. It has been proposed that the first TH signaling systems to evolve used TH precursors obtained from algal sources and that the ancestral THR evolved as a sensor for iodinated tyrosine and indicator of food availability [10,52,83,92]. The appearance of THR in bilaterian animals thus allowed for the elaboration of a ligand-dependent control mechanism for development and growth, whose activity is directly coupled to environmental cues favoring larval survival [10,52,83,92,97].

5. Retinoic Acid Receptor (RAR)-Dependent Signaling Is Required for Neurogenesis in Marine Invertebrates

As in the case of THR, RAR is also dynamically expressed during larval morphogenesis in invertebrates (Figure 5). It is thus reasonable to assume that this NR is also involved in the regulation of developmental functions in marine invertebrates. Vertebrate RARs are ligand-activated transcription factors that act as constitutive repressors in the absence of a ligand [98]. RARs bind different isomers of the small, lipophilic molecule retinoic acid (RA), such as all-trans-RA, 9-cis-RA or 13-cis-RA, with all-trans-RA being the main biologically active RA isomer [99]. RA acts as a morphogen, whose functions are mediated by RARs. During vertebrate development, RA signaling is, for example, required for axial patterning, nervous system development and organogenesis, with HOX genes being amongst the major targets of this signaling pathway [99,100]. Most bilaterian genomes contain a single RAR gene [17,27,36,49,50,101,102,103]. However, RARs have been lost in most ecdysozoans as well as in appendicularian (larvacean) tunicates (Figure 3) (Table 3) [27,101,103].
Many elements of a vertebrate-like RA signaling system are present in invertebrate deuterostomes, including the genes encoding the receptors and the enzymes required for the synthesis and degradation of endogenous RA [99]. In tunicates that have not secondarily lost the RAR gene, RAR-dependent RA signaling is implicated in neurogenesis as well as in tissue regeneration and bud development of budding tunicates [104,105]. The cephalochordate amphioxus is characterized by the most vertebrate-like RA signaling system of all invertebrates [99,106]. It has been demonstrated that the amphioxus RAR/RXR heterodimer can be activated by all-trans-RA, that HOX genes are directly regulated by RAR/RXR and that HOX-mediated RA signaling is essential for neurogenesis and axial patterning [107]. Conversely, little is known about RAR and RA signaling in ambulacrarians. In echinoderms, the sea urchin RAR can bind RA in vitro, although with low affinity, and, while RA treatments might either disrupt or delay sea urchin development, the molecular mechanisms underlying these effects remain to be established [49,99].
In lophotrochozoans, extensive functional characterizations of RAR orthologs and RA signaling have been carried out in both annelids and mollusks [49,50,51,55,102]. The annelid P. dumerilii possesses a vertebrate-like RA signaling machinery composed of a ligand-activated RAR as well as enzymes for RA synthesis and degradation [49,101]. P. dumerilii RAR is activated by RA binding and regulates transcription in a heterodimer with RXR [49]. However, the P. dumerilii RAR has a lower affinity for RA than its vertebrate orthologs, and the conformation of the ligand within the LBP is not conserved between the annelid and vertebrate receptors (Figure 1) [49]. When exposed to all-trans-RA or 13-cis-RA, P. dumerilii embryos experience neuroblast depletion leading to reduced numbers of differentiating motor neurons and suggesting a direct effect of RA on dividing neural stem cells [49]. Moreover, knockdown of P. dumerilii RAR or RXR causes severe malformations of the developing larval nervous system [49]. RAR-dependent RA signaling is thus required for neurogenesis in annelids [49].
Contrasting the situation in annelids, mollusk RARs have lost the ability to bind RA. It has been suggested that mollusk RARs secondarily lost the capacity to bind RA by accumulating independent single mutations in different mollusk lineages [51,56]. Accordingly, mollusk RARs likely function as constitutive transcriptional repressors [50,51,102]. It has recently been proposed that this repressive activity of RAR within the RAR/RXR heterodimer could be modified by RXR-dependent ligand binding [50,102]. It thus remains elusive how RA controls development and neurogenesis in mollusks [50,55,56,99,106]. Similarly, although the genomes of most ecdysozoans do not encode a RAR gene, there are indications for active roles of RA signaling in ecdysozoans, for example, during nervous system regeneration and in tissue repair of insects [108,109]. As a matter of fact, RAR has been lost very early during ecdysozoan evolution, after the split of the priapulid lineage [103]. Notably, the RAR of the priapulid Priapulus caudatus binds all-trans-RA and 9-cis-RA with affinities similar to those of the P. dumerilii RAR [49,103]. In cnidarians, whose genomes do not encode RARs, RA has been implicated in the regulation of metamorphosis and neural development [55,64,99,110]. Independent of the presence of RAR, RA can thus be considered a potent morphogen involved in metazoan neurogenesis. Future work will have to address how the RA signal is mediated in the absence of RAR, and several scenarios, including the involvement of a liganded RXR, have already been proposed for the evolutionary origin of RA signaling at the base of metazoans [50,99]. RARs only arose later, at the base of bilaterians, as ligand-activated RA sensors that significantly facilitated the regulatory control of RA signals [49].

6. Retinoid X Receptor (RXR) Functions during Marine Invertebrate Development

In vertebrates, RXRs act as heterodimeric binding partners of NR1 and NR4 subfamily receptors. Their activity is thus subordinated to those of the heterodimeric binding partners, which participate in a wide variety of developmental processes [39,111]. However, under very specific circumstances, RXRs may also function as ligand-activated receptors. In vitro, RXRs are activated by a number of different compounds, including 9-cis-RA and fatty acids, such as docosahexaenoic acids (DHAs) [112,113]. Orthologs of vertebrate RXRs have been identified in most metazoan taxa, including sponges, placozoans and cnidarians, suggesting that RXRs originated at the base of the animal tree of life [57,114]. Lineage-specific duplications of RXR genes are rare and have so far only been reported in two lophotrochozoans: bryozoans and platyhelminths (Figure 3) [57]. RXR is widely expressed during development of most marine invertebrate taxa, including cnidarians, mollusks and tunicates. Compared to other NRs, RXR does not show a clear pattern of developmental clustering. Its expression profiles tend to follow the ones of its heterodimeric binding partners (Figure 4).
Marine invertebrates provide excellent examples for the importance of RXRs as regulators of development and point to a possible ligand-activated function of these receptors (Table 3). In the sea urchin Strongylocentrotus nudus, for example, an ambulacrarian deuterostome, knockdown of RXR induces abnormal early embryonic development and leads to a complete arrest of embryonic development at the early gastrula stage, suggesting that RXR is crucially required for sea urchin development [115]. Furthermore, a study of the RXR from the sea urchin Paracentrotus lividus suggests that exposure to an endogenous gonadal fatty acid mixture stimulates the activity of the PPAR/RXR heterodimer in cellulo and that this stimulation is mediated by ligand binding to RXR. In this particular context, the transcriptional activity of the PPAR/RXR heterodimer might thus be regulated by RXR [116]. In ecdysozoan and lophotrochozoans protostomes, RXR mainly acts as a heterodimeric partner of other NRs, in accordance with the process of RXR subordination [117,118]. However, a ligand-activated function of RXRs has been proposed in mollusks, where it was shown, in C. gigas, Nucella lapillus and Acanthochitona crinita, that RXR-specific ligands abrogate the repressive state of RAR/RXR heterodimers [50]. Given that mollusk RARs do not seem to bind RA, the activity of RAR/RXR during mollusk development might thus be regulated by ligand binding to RXR. Further studies need to address this hypothesis in vivo and evaluate the biological relevance of RXR ligands during development.
While the genomes of non-bilaterians encode RXR genes, their heterodimeric binding partners, i.e., members of the NR1 and NR4 subfamilies, are absent. The RXR of medusozoan cnidarians binds 9-cis-RA with high affinity and is required for strobilation [64]. Cnidarian RXRs might further be involved in the RA-dependent regulation of nervous system patterning and neurogenesis [65,99,110]. It is thus tempting to speculate that RXR is mediating the developmental roles of RA in cnidarians, potentially as a homodimer [119].

7. Estrogen Receptor (ER), Estrogen-Related Receptor (ERR) and the Development of Marine Invertebrates

In vertebrates, ERs are activated by estradiol (E2) and function during the formation of the nervous system, during development of secondary sexual characteristics, in the regulation of the immune system, in the maintenance of bone density and in the control of social behavior [120,121,122]. ERRs form a group of orphan receptors that are closely related to ERs and that are critical for the regulation of neurogenesis and metabolism as well as for cell proliferation and cell movements [123,124,125]. While the endogenous ligand of ERRs still remains elusive, their activity can be modulated by synthetic ligands, some of which also activate or inhibit ERs, such as, respectively, diethylstilbestrol (DES) or 4-hydroxytamoxifen (4-HT) [126]. ER and ERR are members of the NR3 subfamily, which originated early in metazoan evolution, likely after the split of the sponge lineage, and experienced diversification in early bilaterians and subsequently also in early chordates (Figure 2; Figure 3) [20,21,127]. Phylogenetically, non-chordate ERs thus group at the base of chordate ERs and SRs [20,127,128]. Single orthologs of ER and ERR have been identified in most protostomes, with ecdysozoans having lost ER and the nematodes, which are ecdysozoans, having additionally lost ERR (Figure 3) [27]. The genomes of invertebrate deuterostomes also encode single copies of ER and ERR and those of cephalochordates additionally a single SR [14,31]. Within chordates, the tunicates have secondarily lost both ER and SR (Figure 3) [27,36,127]. The lineage-specific loss of an ER-like gene has further been reported in anthozoan cnidarians (Figure 3) [20,30]. Even though their expression profiles are indicative of a function during invertebrate development (Figure 5), the role played by these NRs outside vertebrates remains virtually unknown.

7.1. ERR Might Play a Role during Development of Marine Invertebrates

Ascidian tunicates possess a single ERR gene [36,53], whose expression, albeit dynamically modulated during late larval development, is exclusively localized in the brain (Figure 4) (Table 4) [53,129]. DES and 4-HT both affect brain formation and trunk elongation in developing P. mammilata larvae, suggesting that ERR acts during nervous system development and body extension of tunicates [53,129]. In the cephalochordate amphioxus, ERR is expressed in the hindbrain homolog and in the developing musculature [130]. The segmented patterns of ERR expression in hindbrain and muscles suggest that this NR is involved in establishing neuromuscular contacts in developing amphioxus [130]. In protostomes, ERR functions have been established in ecdysozoans, where it regulates metabolic processes supporting larval growth and cell proliferation [131,132]. ERR is further suspected to be a pivotal player during metamorphosis [131]. In contrast, there is only limited information available on ERR expression and functions in lophotrochozoans. In C. gigas, ERR is dynamically expressed during development with a peak at pre-metamorphic and metamorphic stages [47]. Accordingly, it is parsimonious to assume that ERR plays specific functional roles during development of mollusks, and more generally, of lophotrochozoans.

7.2. Developmental Functions of ER in Marine Invertebrates Remain Largely Elusive

In the cephalochordate amphioxus, estrogens have been proposed to play important roles in reproductive functions, such as spawning [133]. However, it is currently unknown if ER and SR are required to mediate these functions (Table 4). In contrast, experimental evidence from marine protostomes, such as annelids and rotifers, supports the hypothesis that ERs are involved in reproductive processes of at least some invertebrates [134,135,136,137]. However, studies addressing the developmental roles of ER-like receptors in protostomes remain extremely rare, with the only description of developmental functions of ER-like receptors in protostomes coming from annelids. In fact, estrogens are endogenously synthesized in annelids and are required for the formation and proliferation of primordial germ cells [136,137,138]. This regulation is dependent on ER, which binds the endogenous estrogens and hence directly controls this process [136,138]. Estrogen also induces proliferation of primitive germ cells in vertebrates, suggesting that germ cell regulation represents an ancestral trait of ER signaling in bilaterians [138]. In contrast, estrogen binding and ligand-dependent activation were lost in mollusk ERs [128,139]. Mollusk ERs are constitutively active and retained the ability to regulate their own gene transcription, but do not bind estrogens or other steroids, as their LBPs underwent vestigialization [128,139,140]. Nevertheless, in the bivalve mollusks C. gigas and Mytilus galloprovincialis, dynamic ER expression was detected at larval stages, which is suggestive of a role for ER in larval development [47,141]. Outside bilaterians, ER-like receptors have been identified in placozoans and cnidarians [20,21]. While their expression and function have yet to be assessed, it has been shown that the H. vulgaris ER-like receptor can bind paraestrol A, an ancestral estrogen [20,134]. Notably, a compound isolated from the cnidarian Dendronephthya studeri is structurally similar to paraestrol A, suggesting that ER-like receptors might act as ligand-activated transcription factors, at least in cnidarians [134,142].

8. Are NRs a Primary Target of EDCs in Marine Invertebrates?

Invertebrate protostomes and deuterostomes were initially thought not to be affected by EDCs, as they supposedly lack an elaborate endocrine system [44]. This assumption was shown to be incorrect, following the collapse of gastropod and bivalve mollusk populations in the Arcachon Bay in France. It was later shown that the mollusks were affected by EDCs in marine antifouling paints, already known to modulate the activity of PPAR/RXR heterodimers in vertebrates [45]. Today, it is well established that marine invertebrates are sensitive to EDC pollution and clearly manifest their adverse effects, with embryonic stages being particularly vulnerable. The scientific community has thus invested greatly in the study of EDC exposure and endocrine systems of marine invertebrates, turning their embryos into valuable alternative models for EDC and chemical testing [143]. However, with the exception of the RXR-dependent imposex phenotype in gastropod mollusks [144], evidence for NRs as primary targets of EDCs in marine invertebrates is still extremely circumstantial and mainly based on in vitro studies that do not necessarily reflect the molecular mechanisms in vivo [44,45,46,145,146]. This does not mean that NR-mediated endocrine disruption does not occur on a large scale in marine invertebrates. In fact, the chemicals currently classified as EDCs have chiefly been identified and characterized in vertebrates, and most studies into marine invertebrates have simply used these compounds to assess whether they also affect their endocrine system [46]. This vertebrate-centric view has thus heavily biased our current understanding of invertebrate endocrine systems.
Based on the work reviewed here, it is not surprising that high affinity EDCs of vertebrate NRs do not necessarily affect NRs of marine invertebrates (and vice versa) [44,45,145]. In the course of their evolution and diversification, NRs have been subjected to significant alterations of their sequences and structures, which explains why orthologous NRs are not necessarily characterized by conserved ligand binding affinities, downstream targets and biological functions [1,2,13,15,16,17,25]. Accordingly, there are probably numerous substances that can act as NR-mediated EDCs in marine invertebrates, and their modes of action are likely to be very different from those known in vertebrates [46,146]. Furthermore, it is currently impossible to estimate to what extent the endocrine systems of marine invertebrates are comparable to those of vertebrates, including the involvement of NRs [147,148,149]. There are thus a number of different points that need to be addressed in order to establish the endocrine disrupting potential of a particular set of chemicals in marine invertebrate taxa. These include the characterization of the roles of NRs in marine invertebrate physiology and endocrine systems, the assessment of their functions in embryonic and post-embryonic development and the detailed definition of the physiological and developmental outcomes of NR-dependent endocrine disruption.

Author Contributions

Writing—original draft preparation, A.M., M.S., R.D.; writing—review and editing, A.M., L.C., I.D.L.G., M.S., R.D.; funding acquisition, L.C., M.S., R.D. All authors have read and agreed to the published version of the manuscript.

Funding

This work is financed by an ANR grant to Rémi Dumollard (Marine-EmbryoTox, ANR-14-OHRI-0009-01-1). Michael Schubert is supported by the CNRS.

Acknowledgments

The authors are indebted to Emma Bokobza and Manon Brunet, to the members of the Ascidian BioCell and EvoInSiDe teams at the Laboratoire de Biologie du Développement de Villefranche-sur-Mer as well as to the members of the Canesi team of Environmental Physiology at the Università Degli Studi Di Genova for fruitful discussions. The authors would like to thank Laurent Gilletta and Régis Lasbleiz of the Centre de Ressources Biologiques Marines of the Institut de la Mer de Villefranche that is supported by EMBRC-France and funded by the Agence Nationale de la Recherche (ANR) (ANR-10-INBS-02).

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Escriva, H.; Delaunay, F.; Laudet, V. Ligand binding and nuclear receptor evolution. BioEssays 2000, 22, 717–727. [Google Scholar] [CrossRef]
  2. Escriva, H.; Bertrand, S.; Laudet, V. The evolution of the nuclear receptor superfamily. Essays Biochem. 2004, 40, 11–26. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. Bain, D.L.; Heneghan, A.F.; Connaghan-Jones, K.D.; Miura, M.T. Nuclear receptor structure: Implications for function. Annu. Rev. Physiol. 2007, 69, 201–220. [Google Scholar] [CrossRef] [PubMed]
  4. Bourguet, W.; Germain, P.; Gronemeyer, H. Nuclear receptor ligand-binding domains: Three-dimensional structures, molecular interactions and pharmacological implications. Trends Pharmacol. Sci. 2000, 21, 381–388. [Google Scholar] [CrossRef]
  5. Helsen, C.; Kerkhofs, S.; Clinckemalie, L.; Spans, L.; Laurent, M.; Boonen, S.; Vanderschueren, D.; Claessens, F. Structural basis for nuclear hormone receptor DNA binding. Mol. Cell. Endocrinol. 2012, 348, 411–417. [Google Scholar] [CrossRef]
  6. Claessens, F.; Gewirth, D.T. DNA recognition by nuclear receptors. Essays Biochem. 2004, 40, 59–72. [Google Scholar] [CrossRef] [Green Version]
  7. Khorasanizadeh, S.; Rastinejad, F. Nuclear-receptor interactions on DNA-response elements. Trends Biochem. Sci. 2001, 26, 384–390. [Google Scholar] [CrossRef]
  8. Perlmann, T.; Umesono, K.; Rangarajan, P.N.; Forman, B.M.; Evans, R.M. Two distinct dimerization interfaces differentially modulate target gene specificity of nuclear hormone receptors. Mol. Endocrinol. 1996, 10, 958–966. [Google Scholar] [CrossRef]
  9. Germain, P.; Bourguet, W. Dimerization of nuclear receptors. In Methods in Cell Biology; Academic Press Inc.: Cambridge, MA, USA, 2013; Volume 117, pp. 21–41. [Google Scholar] [CrossRef]
  10. Sainath, S.B.; André, A.; Castro, L.F.C.; Santos, M.M. The evolutionary road to invertebrate thyroid hormone signaling: Perspectives for endocrine disruption processes. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 2019, 223, 124–138. [Google Scholar] [CrossRef]
  11. Benoit, G.; Cooney, A.; Giguere, V.; Ingraham, H.; Lazar, M.; Muscat, G.; Perlmann, T.; Renaud, J.P.; Schwabe, J.; Sladek, F.; et al. International union of pharmacology. LXVI. Orphan nuclear receptors. Pharmacol. Rev. 2006, 58, 798–836. [Google Scholar] [CrossRef] [Green Version]
  12. Germain, P.; Staels, B.; Dacquet, C.; Spedding, M.; Laudet, V. Overview of nomenclature of nuclear receptors. Pharmacol. Rev. 2006, 58, 685–704. [Google Scholar] [CrossRef] [PubMed]
  13. Laudet, V.; Hanni, C.; Coll, J.; Catzeflis, F.; Stehelin, D. Evolution of the nuclear receptor gene superfamily. EMBO J. 1992, 11, 1003–1013. [Google Scholar] [CrossRef] [PubMed]
  14. Lecroisey, C.; Laudet, V.; Schubert, M. The cephalochordate amphioxus: A key to reveal the secrets of nuclear receptor evolution. Brief. Funct. Genom. 2012, 11, 156–166. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  15. Bridgham, J.T.; Eick, G.N.; Larroux, C.; Deshpande, K.; Harms, M.J.; Gauthier, M.E.A.; Ortlund, E.A.; Degnan, B.M.; Thornton, J.W. Protein evolution by molecular tinkering: Diversification of the nuclear receptor superfamily from a ligand-dependent ancestor. PLoS Biol. 2010, 8. [Google Scholar] [CrossRef] [Green Version]
  16. Markov, G.V.; Laudet, V. Origin and evolution of the ligand-binding ability of nuclear receptors. Mol. Cell. Endocrinol. 2011, 334, 21–30. [Google Scholar] [CrossRef]
  17. Fonseca, E.S.S. Nuclear receptors in metazoan lineages: The cross-talk between evolution and endocrine disruption. Ph.D. Thesis, Faculdade de Ciências da Universidade do Porto, Porto, Portugal, 2020. [Google Scholar]
  18. Santos, M.M.; Ruivo, R.; Capitão, A.; Fonseca, E.; Castro, L.F.C. Identifying the gaps: Resources and perspectives on the use of nuclear receptor based-assays to improve hazard assessment of emerging contaminants. J. Hazard. Mater. 2018, 358, 508–511. [Google Scholar] [CrossRef]
  19. Vogeler, S.; Galloway, T.S.; Lyons, B.P.; Bean, T.P. The nuclear receptor gene family in the Pacific oyster, Crassostrea gigas, contains a novel subfamily group. BMC Genom. 2014, 15. [Google Scholar] [CrossRef] [Green Version]
  20. Khalturin, K.; Billas, I.M.L.; Chebaro, Y.; Reitzel, A.M.; Tarrant, A.M.; Laudet, V.; Markov, G.V. NR3E receptors in cnidarians: A new family of steroid receptor relatives extends the possible mechanisms for ligand binding. J. Steroid Biochem. Mol. Biol. 2018, 184, 11–19. [Google Scholar] [CrossRef]
  21. Baker, M.E. Trichoplax, the simplest known animal, contains an estrogen-related receptor but no estrogen receptor: Implications for estrogen receptor evolution. Biochem. Biophys. Res. Commun. 2008, 375, 623–627. [Google Scholar] [CrossRef] [Green Version]
  22. Wu, W.; Niles, E.G.; Hirai, H.; LoVerde, P.T. Evolution of a novel subfamily of nuclear receptors with members that each contain two DNA binding domains. BMC Evol. Biol. 2007, 7. [Google Scholar] [CrossRef] [Green Version]
  23. Kaur, S.; Jobling, S.; Jones, C.S.; Noble, L.R.; Routledge, E.J.; Lockyer, A.E. The nuclear receptors of Biomphalaria glabrata and Lottia gigantea: Implications for developing new model organisms. PLoS ONE 2015, 10. [Google Scholar] [CrossRef] [Green Version]
  24. Huang, W.; Xu, F.; Li, J.; Li, L.; Que, H.; Zhang, G. Evolution of a novel nuclear receptor subfamily with emphasis on the member from the Pacific oyster Crassostrea gigas. Gene 2015, 567, 164–172. [Google Scholar] [CrossRef] [PubMed]
  25. Holzer, G.; Markov, G.V.; Laudet, V. Evolution of nuclear receptors and ligand signaling: Toward a soft key–lock model? In Current Topics in Developmental Biology; Academic Press Inc.: Cambridge, MA, USA, 2017; Volume 125, pp. 1–38. [Google Scholar] [CrossRef]
  26. Robinson-Rechavi, M.; Garcia, H.E.; Laudet, V. The nuclear receptor superfamily. J. Cell Sci. 2003, 116, 585–586. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  27. Bertrand, S.; Brunet, F.G.; Escriva, H.; Parmentier, G.; Laudet, V.; Robinson-Rechavi, M. Evolutionary genomics of nuclear receptors: From twenty-five ancestral genes to derived endocrine systems. Mol. Biol. Evol. 2004, 21, 1923–1937. [Google Scholar] [CrossRef] [PubMed]
  28. Laumer, C.E.; Fernández, R.; Lemer, S.; Combosch, D.; Kocot, K.M.; Riesgo, A.; Andrade, S.C.S.; Sterrer, W.; Sørensen, M.V.; Giribet, G. Revisiting metazoan phylogeny with genomic sampling of all phyla. Proc. R. Soc. B 2019, 286. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  29. Novotnỳ, J.P.; Chughtai, A.A.; Kostrouchová, M.; Kostrouchová, V.; Kostrouch, D.; Kaššák, F.; Kaňa, R.; Schierwater, B.; Kostrouchová, M.; Kostrouch, Z. Trichoplax adhaerens reveals a network of nuclear receptors sensitive to 9-cis-retinoic acid at the base of metazoan evolution. PeerJ 2017, 2017. [Google Scholar] [CrossRef] [Green Version]
  30. Reitzel, A.M.; Tarrant, A.M. Nuclear receptor complement of the cnidarian Nematostella vectensis: Phylogenetic relationships and developmental expression patterns. BMC Evol. Biol. 2009, 9. [Google Scholar] [CrossRef] [Green Version]
  31. Schubert, M.; Brunet, F.; Paris, M.; Bertrand, S.; Benoit, G.; Laudet, V. Nuclear hormone receptor signaling in amphioxus. Dev. Genes Evol. 2008, 218, 651–665. [Google Scholar] [CrossRef]
  32. Dehal, P.; Boore, J.L. Two rounds of whole genome duplication in the ancestral vertebrate. PLoS Biol. 2005, 3. [Google Scholar] [CrossRef] [Green Version]
  33. Baker, M.E. Steroid receptors and vertebrate evolution. Mol. Cell. Endocrinol. 2019, 496, 110526. [Google Scholar] [CrossRef] [Green Version]
  34. Robinson-Rechavi, M.; Maina, C.V.; Gissendanner, C.R.; Laudet, V.; Sluder, A. Explosive lineage-specific expansion of the orphan nuclear receptor HNF4 in nematodes. J. Mol. Evol. 2005, 60, 577–586. [Google Scholar] [CrossRef] [PubMed]
  35. Kostrouchova, M.; Kostrouch, Z. Nuclear receptors in nematode development: Natural experiments made by a phylum. Biochim. Biophys. Acta Gene Regul. Mech. 2015, 1849, 224–237. [Google Scholar] [CrossRef] [PubMed]
  36. Dehal, P.; Satou, Y.; Campbell, R.K.; Chapman, J.; Degnan, B.; De Tomaso, A.; Davidson, B.; Di Gregorio, A.; Gelpke, M.; Goodstein, D.M.; et al. The draft genome of Ciona intestinalis: Insights into chordate and vertebrate origins. Science 2002, 298, 2157–2167. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  37. Chung, A.C.; Cooney, A.J. The varied roles of nuclear receptors during vertebrate embryonic development. Nucl. Recept. Signal. 2003, 1, nrs.01007. [Google Scholar] [CrossRef] [PubMed]
  38. Mangelsdorf, D.J.; Evans, R.M. The RXR heterodimers and orphan receptors. Cell 1995, 83, 841–850. [Google Scholar] [CrossRef] [Green Version]
  39. Evans, R.M.; Mangelsdorf, D.J. Nuclear receptors, RXR, and the big bang. Cell 2014, 157, 255–266. [Google Scholar] [CrossRef] [Green Version]
  40. Gronemeyer, H.; Gustafsson, J.Å.; Laudet, V. Principles for modulation of the nuclear receptor superfamily. Nat. Rev. Drug Discov. 2004, 3, 950–964. [Google Scholar] [CrossRef]
  41. Zoeller, R.T.; Brown, T.R.; Doan, L.L.; Gore, A.C.; Skakkebaek, N.E.; Soto, A.M.; Woodruff, T.J.; Vom Saal, F.S. Endocrine-disrupting chemicals and public health protection: A statement of principles from the Endocrine Society. Endocrinology 2012, 153, 4097–4110. [Google Scholar] [CrossRef]
  42. Toporova, L.; Balaguer, P. Nuclear receptors are the major targets of endocrine disrupting chemicals. Mol. Cell. Endocrinol. 2020, 502, 110665. [Google Scholar] [CrossRef]
  43. Balaguer, P.; Delfosse, V.; Bourguet, W. Mechanisms of endocrine disruption through nuclear receptors and related pathways. Curr. Opin. Endocr. Metab. Res. 2019, 7, 1–8. [Google Scholar] [CrossRef]
  44. Katsiadaki, I. Are marine invertebrates really at risk from endocrine-disrupting chemicals? Curr. Opin. Environ. Sci. Health 2019, 11, 37–42. [Google Scholar] [CrossRef]
  45. Fernandez, M.A. Populations collapses in marine invertebrates due to endocrine disruption: A cause for concern? Front. Endocrinol. Lausanne 2019, 10, 1–14. [Google Scholar] [CrossRef]
  46. Ford, A.T.; Leblanc, G.A. Endocrine disruption in invertebrates: A survey of research progress. Environ. Sci. Technol. 2020. [Google Scholar] [CrossRef] [PubMed]
  47. Vogeler, S.; Bean, T.P.; Lyons, B.P.; Galloway, T.S. Dynamics of nuclear receptor gene expression during Pacific oyster development. BMC Dev. Biol. 2016, 16, 1–13. [Google Scholar] [CrossRef] [Green Version]
  48. Bodofsky, S.; Koitz, F.; Wightman, B. Conserved and exapted functions of nuclear receptors in animal development. Nucl. Recept. Res. 2017. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  49. Handberg-Thorsager, M.; Gutierrez-Mazariegos, J.; Arold, S.T.; Nadendla, E.K.; Bertucci, P.Y.; Germain, P.; Tomançak, P.; Pierzchalski, K.; Jones, J.W.; Albalat, R.; et al. The ancestral retinoic acid receptor was a low-affinity sensor triggering neuronal differentiation. Sci. Adv. 2018, 4. [Google Scholar] [CrossRef] [Green Version]
  50. André, A.; Ruivo, R.; Fonseca, E.; Froufe, E.; Castro, L.F.C.; Santos, M.M. The retinoic acid receptor (RAR) in molluscs: Function, evolution and endocrine disruption insights. Aquat. Toxicol. 2019, 208, 80–89. [Google Scholar] [CrossRef]
  51. Gutierrez-Mazariegos, J.; Nadendla, E.K.; Lima, D.; Pierzchalski, K.; Jones, J.W.; Kane, M.; Nishikawa, J.I.; Hiromori, Y.; Nakanishi, T.; Santos, M.M.; et al. A mollusk retinoic acid receptor (RAR) ortholog sheds light on the evolution of ligand binding. Endocrinology 2014, 155, 4275–4286. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  52. Holzer, G.; Roux, N.; Laudet, V. Evolution of ligands, receptors and metabolizing enzymes of thyroid signaling. Mol. Cell. Endocrinol. 2017, 459, 5–13. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Gomes, I.D.L.; Gazo, I.; Besnardeau, L.; Hebras, C.; McDougall, A.; Dumollard, R. Potential roles of nuclear receptors in mediating neurodevelopmental toxicity of known endocrine-disrupting chemicals in ascidian embryos. Mol. Reprod. Dev. 2019, 86, 1333–1347. [Google Scholar] [CrossRef] [PubMed]
  54. Holzer, G. Origin of thyroid hormone signalling in metazoans and implications in their metamorphosis. Ph.D. Thesis, Ecole Normale Supérieure de Lyon—ENS Lyon, Lyon, France, 2015. [Google Scholar]
  55. André, A.; Ruivo, R.; Gesto, M.; Castro, L.F.C.; Santos, M.M. Retinoid metabolism in invertebrates: When evolution meets endocrine disruption. Gen. Comp. Endocrinol. 2014, 208, 134–145. [Google Scholar] [CrossRef] [PubMed]
  56. Vogeler, S.; Galloway, T.S.; Isupov, M.; Bean, T.P. Cloning retinoid and peroxisome proliferatoractivated nuclear receptors of the Pacific oyster and in silico binding to environmental chemicals. PLoS ONE 2017, 12, 1–21. [Google Scholar] [CrossRef]
  57. Fonseca, E.; Ruivo, R.; Borges, D.; Franco, J.N.; Santos, M.M.; Castro, L.F.C. Of retinoids and organotins: The evolution of the retinoid x receptor in metazoa. Biomolecules 2020, 10, 594. [Google Scholar] [CrossRef] [Green Version]
  58. Pechenik, J.A. On the advantages and disadvantages of larval stages in benthic marine invertebrate life cycles. Mar. Ecol. Prog. Ser. 1999, 177, 269–297. [Google Scholar] [CrossRef] [Green Version]
  59. Hickman, C.S. Larvae in invertebrate development and evolution. In The Origin and Evolution of Larval Forms; Elsevier: Amsterdam, The Netherlands, 1999; pp. 21–59. [Google Scholar] [CrossRef]
  60. Hadfield, M.G. Why and how marine-invertebrate larvae metamorphose so fast. Semin. Cell Dev. Biol. 2000, 11, 437–443. [Google Scholar] [CrossRef] [PubMed]
  61. Leclère, L.; Horin, C.; Chevalier, S.; Lapébie, P.; Dru, P.; Peron, S.; Jager, M.; Condamine, T.; Pottin, K.; Romano, S.; et al. The genome of the jellyfish Clytia hemisphaerica and the evolution of the cnidarian life-cycle. Nat. Ecol. Evol. 2019, 3, 801–810. [Google Scholar] [CrossRef] [Green Version]
  62. Alié, A.; Hiebert, L.S.; Scelzo, M.; Tiozzo, S. The eventful history of nonembryonic development in tunicates. J. Exp. Zool. Part B Mol. Dev. Evol. 2020. [Google Scholar] [CrossRef]
  63. Scelzo, M.; Alié, A.; Pagnotta, S.; Lejeune, C.; Henry, P.; Gilletta, L.; Hiebert, L.S.; Mastrototaro, F.; Tiozzo, S. Novel budding mode in Polyandrocarpa zorritensis: A model for comparative studies on asexual development and whole body regeneration. Evodevo 2019, 10, 1–13. [Google Scholar] [CrossRef] [Green Version]
  64. Fuchs, B.; Wang, W.; Graspeuntner, S.; Li, Y.; Insua, S.; Herbst, E.M.; Dirksen, P.; Böhm, A.M.; Hemmrich, G.; Sommer, F.; et al. Regulation of polyp-to-jellyfish transition in Aurelia aurita. Curr. Biol. 2014, 24, 263–273. [Google Scholar] [CrossRef] [Green Version]
  65. Brekhman, V.; Malik, A.; Haas, B.; Sher, N.; Lotan, T. Transcriptome profiling of the dynamic life cycle of the scypohozoan jellyfish Aurelia aurita. BMC Genom. 2015, 16. [Google Scholar] [CrossRef] [Green Version]
  66. Wray, G.A.; Raff, R.A. The evolution of developmental strategy in marine invertebrates. Trends Ecol. Evol. 1991, 6, 45–50. [Google Scholar] [CrossRef]
  67. Taneja, R. Nuclear Receptors in Development; Elsevier: Amsterdam, The Netherlands, 2006; Volume 16, ISBN 9780444528735. [Google Scholar]
  68. Bunce, C.M.; Campbell, M.J. Nuclear receptors an introductory overview. In Nuclear Receptors; Springer: Berlin/Heidelberg, Germany, 2010; pp. 1–13. [Google Scholar] [CrossRef]
  69. Yaguchi, S.; Morino, Y.; Sasakura, Y. Development of marine invertebrates. In Japanese Marine Life; Springer: Singapore, 2020; pp. 109–124. [Google Scholar] [CrossRef]
  70. Huang, W.; Wu, Q.; Xu, F.; Li, L.; Li, J.; Que, H.; Zhang, G. Functional characterization of retinoid X receptor with an emphasis on the mediation of organotin poisoning in the Pacific oyster (Crassostrea gigas). Gene 2020, 753. [Google Scholar] [CrossRef] [PubMed]
  71. Pereira, F.A.; Tsai, M.J.; Tsai, S.Y. COUP-TF orphan nuclear receptors in development and differentiation. Cell. Mol. Life Sci. 2000, 57, 1388–1398. [Google Scholar] [CrossRef] [PubMed]
  72. Tang, K.; Tsai, S.Y.; Tsai, M.J. COUP-TFs and eye development. Biochim. Biophys. Acta Gene Regul. Mech. 2015, 1849, 201–209. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Gauchat, D.; Escriva, H.; Miljkovic-Licina, M.; Chera, S.; Langlois, M.C.; Begue, A.; Laudet, V.; Galliot, B. The orphan COUP-TF nuclear receptors are markers for neurogenesis from cnidarians to vertebrates. Dev. Biol. 2004, 275, 104–123. [Google Scholar] [CrossRef] [Green Version]
  74. Islam, M.M.; Zhang, C.L. TLX: A master regulator for neural stem cell maintenance and neurogenesis. Biochim. Biophys. Acta Gene Regul. Mech. 2015, 1849, 210–216. [Google Scholar] [CrossRef] [Green Version]
  75. Kaltenbach, S.L.; Yu, J.K.; Holland, N.D. The origin and migration of the earliest-developing sensory neurons in the peripheral nervous system of amphioxus. Evol. Dev. 2009, 11, 142–151. [Google Scholar] [CrossRef]
  76. Watt, A.J.; Garrison, W.D.; Duncan, S.A. HNF4: A central regulator of hepatocyte differentiation and function. Hepatology 2003, 37, 1249–1253. [Google Scholar] [CrossRef]
  77. Laws, K.M.; Drummond-Barbosa, D. Control of germline stem cell lineages by diet and physiology. In Results and Problems in Cell Differentiation; Springer: Berlin/Heidelberg, Germany, 2017; Volume 59, pp. 67–99. [Google Scholar] [CrossRef] [Green Version]
  78. Weber, H.; Holewa, B.; Jones, E.A.; Ryffel, G.U. Mesoderm and endoderm differentiation in animal cap explants: Identification of the HNF4-binding site as an activin A responsive element in the Xenopus HNFIα promoter. Development 1996, 122, 1975–1984. [Google Scholar]
  79. Chen, W.S.; Manova, K.; Weinstein, D.C.; Duncan, S.A.; Plump, A.S.; Prezioso, V.R.; Bachvarova, R.F.; Darnell, J.E. Disruption of the HNF-4 gene, expressed in visceral endoderm, leads to cell death in embryonic ectoderm and impaired gastrulation of mouse embryos. Genes Dev. 1994, 8, 2466–2477. [Google Scholar] [CrossRef] [Green Version]
  80. Zhong, W.; Sladek, F.M.; Darnell, J.E. The expression pattern of a Drosophila homolog to the mouse transcription factor HNF-4 suggests a determinative role in gut formation. EMBO J. 1993, 12, 537–544. [Google Scholar] [CrossRef] [PubMed]
  81. Carroll, S.B. Evo-devo and an expanding evolutionary synthesis: A genetic theory of morphological evolution. Cell 2008, 134, 25–36. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  82. Vella, K.R.; Hollenberg, A.N. The actions of thyroid hormone signaling in the nucleus. Mol. Cell. Endocrinol. 2017, 458, 127–135. [Google Scholar] [CrossRef] [PubMed]
  83. Taylor, E.; Heyland, A. Evolution of thyroid hormone signaling in animals: Non-genomic and genomic modes of action. Mol. Cell. Endocrinol. 2017, 459, 14–20. [Google Scholar] [CrossRef]
  84. Liu, Y.Y.; Milanesi, A.; Brent, G.A. Thyroid hormones. In Hormonal Signaling in Biology and Medicine: Comprehensive Modern Endocrinology; Elsevier: Amsterdam, The Netherlands, 2019; pp. 487–506. ISBN 9780128138151. [Google Scholar]
  85. Wu, W.; Niles, E.G.; LoVerde, P.T. Thyroid hormone receptor orthologues from invertebrate species with emphasis on Schistosoma mansoni. BMC Evol. Biol. 2007, 7. [Google Scholar] [CrossRef] [Green Version]
  86. Carosa, E.; Fanelli, A.; Ulisse, S.; Di Lauro, R.; Rall, J.E.; Jannini, E.A. Ciona intestinalis nuclear receptor 1: A member of steroid/thyroid hormone receptor family. Proc. Natl. Acad. Sci. USA 1998, 95, 11152–11157. [Google Scholar] [CrossRef] [Green Version]
  87. Patricolo, E.; Cammarata, M.; Dagati, P. Presence of thyroid hormones in ascidian larvae and their involvement in metamorphosis. J. Exp. Zool. 2001, 290, 426–430. [Google Scholar] [CrossRef] [PubMed]
  88. D’Agati, P.; Cammarata, M. Comparative analysis of thyroxine distribution in ascidian larvae. Cell Tissue Res. 2006, 323, 529–535. [Google Scholar] [CrossRef]
  89. Wang, S.; Zhang, S.; Zhao, B.; Lun, L. Up-regulation of C/EBP by thyroid hormones: A case demonstrating the vertebrate-like thyroid hormone signaling pathway in amphioxus. Mol. Cell. Endocrinol. 2009, 313, 57–63. [Google Scholar] [CrossRef]
  90. Paris, M.; Hillenweck, A.; Bertrand, S.; Delous, G.; Escriva, H.; Zalko, D.; Cravedi, J.P.; Laudet, V. Active metabolism of thyroid hormone during metamorphosis of amphioxus. Integr. Comp. Biol. 2010, 50, 63–74. [Google Scholar] [CrossRef] [Green Version]
  91. Taylor, E.; Heyland, A. Thyroid hormones accelerate initiation of skeletogenesis via MAPK (ERK1/2) in larval sea urchins (Strongylocentrotus purpuratus). Front. Endocrinol. Lausanne 2018, 9, 1–16. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  92. Holzer, G.; Morishita, Y.; Fini, J.B.; Lorin, T.; Gillet, B.; Hughes, S.; Tohmé, M.; Deléage, G.; Demeneix, B.; Arvan, P.; et al. Thyroglobulin represents a novel molecular architecture of vertebrates. J. Biol. Chem. 2016, 291, 16553–16566. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Klootwijk, W.; Friesema, E.C.H.; Visser, T.J. A nonselenoprotein from amphioxus deiodinates TRIAC but not T3: Is TRIAC the primordial bioactive thyroid hormone? Endocrinology 2011, 152, 3259–3267. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Huang, W.; Xu, F.; Qu, T.; Zhang, R.; Li, L.; Que, H.; Zhang, G. Identification of thyroid hormones and functional characterization of thyroid hormone receptor in the pacific oyster Crassostrea gigas provide insight into evolution of the thyroid hormone system. PLoS ONE 2015, 10, 1–20. [Google Scholar] [CrossRef]
  95. Pakharukova, M.Y.; Ershov, N.I.; Vorontsova, E.V.; Shilov, A.G.; Merkulova, T.I.; Mordvinov, V.A. Identification of thyroid hormone receptor homologs in the fluke Opisthorchis felineus (Platyhelminthes). Mol. Biochem. Parasitol. 2014, 194, 64–68. [Google Scholar] [CrossRef]
  96. Cornford, E.M. Schistosomatium douthitti: Effects of thyroxine. Exp. Parasitol. 1974, 36, 210–221. [Google Scholar] [CrossRef]
  97. Holzer, G.; Laudet, V. Thyroid hormones: A triple-edged sword for life history transitions. Curr. Biol. 2015, 25, R344–R347. [Google Scholar] [CrossRef] [Green Version]
  98. Osz, J.; McEwen, A.G.; Bourguet, M.; Przybilla, F.; Peluso-Iltis, C.; Poussin-Courmontagne, P.; Mély, Y.; Cianférani, S.; Jeffries, C.M.; Svergun, D.I.; et al. Structural basis for DNA recognition and allosteric control of the retinoic acid receptors RAR–RXR. Nucleic Acids Res. 2020, 48, 9969–9985. [Google Scholar] [CrossRef]
  99. Zieger, E.; Schubert, M. New insights into the roles of retinoic acid signaling in nervous system development and the establishment of neurotransmitter systems. In International Review of Cell and Molecular Biology; Academic Press: Cambridge, MA, USA, 2017; Volume 330, pp. 1–84. [Google Scholar] [CrossRef]
  100. Ghyselinck, N.B.; Duester, G. Retinoic acid signaling pathways. Development 2019, 146, 1–7. [Google Scholar] [CrossRef] [Green Version]
  101. Albalat, R.; Cañestro, C. Identification of Aldh1a, Cyp26 and RAR orthologs in protostomes pushes back the retinoic acid genetic machinery in evolutionary time to the bilaterian ancestor. Chem. Biol. Interact. 2009, 178, 188–196. [Google Scholar] [CrossRef]
  102. Urushitani, H.; Katsu, Y.; Ohta, Y.; Shiraishi, H.; Iguchi, T.; Horiguchi, T. Cloning and characterization of the retinoic acid receptor-like protein in the rock shell, Thais clavigera. Aquat. Toxicol. 2013, 142–143, 403–413. [Google Scholar] [CrossRef] [PubMed]
  103. Fonseca, E.S.S.; Hiromori, Y.; Kaite, Y.; Ruivo, R.; Franco, J.N.; Nakanishi, T.; Santos, M.M.; Castro, L.F.C. An orthologue of the retinoic acid receptor (RAR) is present in the ecdysozoa phylum Priapulida. Genes 2019, 10, 985. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  104. Nagatomo, K.I.; Ishibashi, T.; Satou, Y.; Satoh, N.; Fujiwara, S. Retinoic acid affects gene expression and morphogenesis without upregulating the retinoic acid receptor in the ascidian Ciona intestinalis. Mech. Dev. 2003, 120, 363–372. [Google Scholar] [CrossRef]
  105. Kaneko, N.; Katsuyama, Y.; Kawamura, K.; Fujiwara, S. Regeneration of the gut requires retinoic acid in the budding ascidian Polyandrocarpa misakiensis. Dev. Growth Differ. 2010, 52, 457–468. [Google Scholar] [CrossRef]
  106. Schubert, M.; Gibert, Y. Retinoids in embryonic development. Biomolecules 2020, 10, 1278. [Google Scholar] [CrossRef]
  107. Schubert, M.; Holland, N.D.; Laudet, V.; Holland, L.Z. A retinoic acid-Hox hierarchy controls both anterior/posterior patterning and neuronal specification in the developing central nervous system of the cephalochordate amphioxus. Dev. Biol. 2006, 296, 190–202. [Google Scholar] [CrossRef]
  108. Halme, A.; Cheng, M.; Hariharan, I.K. Retinoids regulate a developmental checkpoint for tissue regeneration in Drosophila. Curr. Biol. 2010, 20, 458–463. [Google Scholar] [CrossRef] [Green Version]
  109. Bui-Göbbels, K.; Quintela, R.M.; Bräunig, P.; Mey, J. Is retinoic acid a signal for nerve regeneration in insects? Neural Regen. Res. 2015, 10, 901–903. [Google Scholar] [CrossRef]
  110. Estephane, D.; Anctil, M. Retinoic acid and nitric oxide promote cell proliferation and differentially induce neuronal differentiation in vitro in the cnidarian Renilla koellikeri. Dev. Neurobiol. 2010, 70, 842–852. [Google Scholar] [CrossRef]
  111. Lefebvre, P.; Benomar, Y.; Staels, B. Retinoid X receptors: Common heterodimerization partners with distinct functions. Trends Endocrinol. Metab. 2010, 21, 676–683. [Google Scholar] [CrossRef] [Green Version]
  112. Krężel, W.; Rühl, R.; de Lera, A.R. Alternative retinoid X receptor (RXR) ligands. Mol. Cell. Endocrinol. 2019, 491, 110436. [Google Scholar] [CrossRef] [PubMed]
  113. Heyman, R.A.; Mangelsdorf, D.J.; Dyck, J.A.; Stein, R.B.; Eichele, G.; Evans, R.M.; Thaller, C. 9-cis retinoic acid is a high affinity ligand for the retinoid X receptor. Cell 1992, 68, 397–406. [Google Scholar] [CrossRef]
  114. Reitzel, A.M.; Macrander, J.; Mane-Padros, D.; Fang, B.; Sladek, F.M.; Tarrant, A.M. Conservation of DNA and ligand binding properties of retinoid X receptor from the placozoan Trichoplax adhaerens to human. J. Steroid Biochem. Mol. Biol. 2018, 184, 3–10. [Google Scholar] [CrossRef] [PubMed]
  115. Maeng, S.; Kim, G.J.; Choi, E.J.; Yang, H.O.; Lee, D.S.; Sohn, Y.C. 9-cis-retinoic acid induces growth inhibition in retinoid-sensitive breast cancer and sea urchin embryonic cells via retinoid X receptor α and replication factor C3. Mol. Endocrinol. 2012, 26, 1821–1835. [Google Scholar] [CrossRef] [Green Version]
  116. Capitão, A.; Lopes-Marques, M.; Páscoa, I.; Ruivo, R.; Mendiratta, N.; Fonseca, E.; Castro, L.F.C.; Santos, M.M. The Echinodermata PPAR: Functional characterization and exploitation by the model lipid homeostasis regulator tributyltin. Environ. Pollut. 2020, 263. [Google Scholar] [CrossRef]
  117. Iwema, T.; Billas, I.M.L.; Beck, Y.; Bonneton, F.; Nierengarten, H.; Chaumot, A.; Richards, G.; Laudet, V.; Moras, D. Structural and functional characterization of a novel type of ligand-independent RXR-USP receptor. EMBO J. 2007, 26, 3770–3782. [Google Scholar] [CrossRef]
  118. Wang, Y.H.; Wang, G.; LeBlanc, G.A. Cloning and characterization of the retinoid X receptor from a primitive crustacean Daphnia magna. Gen. Comp. Endocrinol. 2007, 150, 309–318. [Google Scholar] [CrossRef]
  119. Germain, P.; Chambon, P.; Eichele, G.; Evans, R.M.; Lazar, M.A.; Leid, M.; De Lera, A.R.; Lotan, R.; Mangelsdorf, D.J.; Gronemeyer, H. International union of pharmacology. LXIII. Retinoid X receptors. Pharmacol. Rev. 2006, 58, 760–772. [Google Scholar] [CrossRef] [Green Version]
  120. Heldring, N.; Pike, A.; Andersson, S.; Matthews, J.; Cheng, G.; Hartman, J.; Tujague, M.; Ström, A.; Treuter, E.; Warner, M.; et al. Estrogen receptors: How do they signal and what are their targets. Physiol. Rev. 2007, 87, 905–931. [Google Scholar] [CrossRef] [Green Version]
  121. Fuentes, N.; Silveyra, P. Estrogen receptor signaling mechanisms. In Advances in Protein Chemistry and Structural Biology; Academic Press: Cambridge, MA, USA, 2019; Volume 116, pp. 135–170. [Google Scholar] [CrossRef]
  122. Amenyogbe, E.; Chen, G.; Wang, Z.; Lu, X.; Lin, M.; Lin, A.Y. A review on sex steroid hormone estrogen receptors in mammals and fish. Int. J. Endocrinol. 2020, 2020. [Google Scholar] [CrossRef] [Green Version]
  123. Bardet, P.L.; Laudet, V.; Vanacker, J.M. Studying non-mammalian models? Not a fool’s ERRand! Trends Endocrinol. Metab. 2006, 17, 166–171. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Crevet, L.; Vanacker, J.M. Regulation of the expression of the estrogen related receptors (ERRs). Cell. Mol. Life Sci. 2020, 77, 4573–4579. [Google Scholar] [CrossRef] [PubMed]
  125. Huss, J.M.; Garbacz, W.G.; Xie, W. Constitutive activities of estrogen-related receptors: Transcriptional regulation of metabolism by the ERR pathways in health and disease. Biochim. Biophys. Acta Mol. Basis Dis. 2015, 1852, 1912–1927. [Google Scholar] [CrossRef] [Green Version]
  126. Fritsch, M.; Leary, C.M.; Furlow, J.D.; Gorski, J.; Ahrens, H.; Mueller, G.C.; Schuh, T.J. A ligand-induced conformational change in the estrogen receptor is localized in the steroid binding domain. Biochemistry 1992, 31, 5303–5311. [Google Scholar] [CrossRef]
  127. Katsu, Y.; Cziko, P.A.; Chandsawangbhuwana, C.; Thornton, J.W.; Sato, R.; Oka, K.; Takei, Y.; Baker, M.E.; Iguchi, T. A second estrogen receptor from Japanese lamprey (Lethenteron japonicum) does not have activities for estrogen binding and transcription. Gen. Comp. Endocrinol. 2016, 236, 105–114. [Google Scholar] [CrossRef] [Green Version]
  128. Bridgham, J.T.; Keay, J.; Ortlund, E.A.; Thornton, J.W. Vestigialization of an allosteric switch: Genetic and structural mechanisms for the evolution of constitutive activity in a steroid hormone receptor. PLoS Genet. 2014, 10. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  129. Gomes, I.D.L.; Gazo, I.; Nabi, D.; Besnardeau, L.; Hebras, C.; McDougall, A.; Dumollard, R. Bisphenols disrupt differentiation of the pigmented cells during larval brain formation in the ascidian Phallusia mammillata. Aquat. Toxicol. 2019, 216, 105314. [Google Scholar] [CrossRef] [PubMed]
  130. Bardet, P.L.; Schubert, M.; Horard, B.; Holland, L.Z.; Laudet, V.; Holland, N.D.; Vanacker, J.M. Expression of estrogen-receptor related receptors in amphioxus and zebrafish: Implications for the evolution of posterior brain segmentation at the invertebrate-to-vertebrate transition. Evol. Dev. 2005, 7, 223–233. [Google Scholar] [CrossRef] [PubMed]
  131. Palanker, L.; Necakov, A.S.; Sampson, H.M.; Ni, R.; Hu, C.; Thummel, C.S.; Krause, H.M. Dynamic regulation of Drosophila nuclear receptor activity in vivo. Development 2006, 133, 3549–3562. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  132. Tennessen, J.M.; Baker, K.D.; Lam, G.; Evans, J.; Thummel, C.S. The Drosophila estrogen-related receptor directs a metabolic switch that supports developmental growth. Cell Metab. 2011, 13, 139–148. [Google Scholar] [CrossRef] [Green Version]
  133. Paris, M.; Pettersson, K.; Schubert, M.; Bertrand, S.; Pongratz, I.; Escriva, H.; Laudet, V. An amphioxus orthologue of the estrogen receptor that does not bind estradiol: Insights into estrogen receptor evolution. BMC Evol. Biol. 2008, 8, 1–20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  134. Markov, G.V.; Gutierrez-Mazariegos, J.; Pitrat, D.; Billas, I.M.L.; Bonneton, F.; Moras, D.; Hasserodt, J.; Lecointre, G.; Laudet, V. Origin of an ancient hormone/receptor couple revealed by resurrection of an ancestral estrogen. Sci. Adv. 2017, 3, 1–14. [Google Scholar] [CrossRef] [Green Version]
  135. Jones, B.L.; Walker, C.; Azizi, B.; Tolbert, L.; Williams, L.D.; Snell, T.W. Conservation of estrogen receptor function in invertebrate reproduction. BMC Evol. Biol. 2017, 17, 1–10. [Google Scholar] [CrossRef] [Green Version]
  136. Keay, J.; Thornton, J.W. Hormone-activated estrogen receptors in annelid invertebrates: Implications for evolution and endocrine disruption. Endocrinology 2009, 150, 1731–1738. [Google Scholar] [CrossRef] [Green Version]
  137. García-Alonso, J.; Hoeger, U.; Rebscher, N. Regulation of vitellogenesis in Nereis virens (Annelida: Polychaeta): Effect of estradiol-17β on eleocytes. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2006, 143, 55–61. [Google Scholar] [CrossRef]
  138. Lidke, A.K.; Bannister, S.; Löwer, A.M.; Apel, D.M.; Podleschny, M.; Kollmann, M.; Ackermann, C.F.; García-Alonso, J.; Raible, F.; Rebscher, N. 17β-Estradiol induces supernumerary primordial germ cells in embryos of the polychaete Platynereis dumerilii. Gen. Comp. Endocrinol. 2014, 196, 52–61. [Google Scholar] [CrossRef]
  139. Balbi, T.; Ciacci, C.; Canesi, L. Estrogenic compounds as exogenous modulators of physiological functions in molluscs: Signaling pathways and biological responses. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 2019, 222, 135–144. [Google Scholar] [CrossRef]
  140. Tran, T.K.A.; MacFarlane, G.R.; Kong, R.Y.C.; O’Connor, W.A.; Yu, R.M.K. Potential mechanisms underlying estrogen-induced expression of the molluscan estrogen receptor (ER) gene. Aquat. Toxicol. 2016, 179, 82–94. [Google Scholar] [CrossRef]
  141. Balbi, T.; Franzellitti, S.; Fabbri, R.; Montagna, M.; Fabbri, E.; Canesi, L. Impact of bisphenol A (BPA) on early embryo development in the marine mussel Mytilus galloprovincialis: Effects on gene transcription. Environ. Pollut. 2016, 218, 996–1004. [Google Scholar] [CrossRef]
  142. Yan, X.H.; Liu, H.L.; Huang, H.; Li, X.B.; Guo, Y.W. Steroids with aromatic A-rings from the Hainan soft coral Dendronephthya studeri Ridley. J. Nat. Prod. 2011, 74, 175–180. [Google Scholar] [CrossRef]
  143. Satya, S.; Wade, M.; Hass, U.; Holbech, H.; Løfstedt, M.; Vinggaard, A.M.; Tyle, K.H.; Nielsen, P.J.; Holmer, M.L.; Christiansen, S. Guidance Document on Standardised Test Guidelines for Evaluating Chemicals for Endocrine Disruption; Organisation for Economic Cooporation and Development: Paris, France, 2014. [Google Scholar] [CrossRef]
  144. Horiguchi, T. Contamination by organotins and its population-level effects involved by imposex in prosobranch gastropods. In Biological Effects by Organotins; Springer: Tokyo, Japan, 2017; pp. 73–99. ISBN 978-4-431-56451-5. [Google Scholar] [CrossRef]
  145. Cuvillier-Hot, V.; Lenoir, A. Invertebrates facing environmental contamination by endocrine disruptors: Novel evidences and recent insights. Mol. Cell. Endocrinol. 2020, 504, 110712. [Google Scholar] [CrossRef] [PubMed]
  146. Zou, E. Invisible endocrine disruption and its mechanisms: A current review. Gen. Comp. Endocrinol. 2020, 293, 113470. [Google Scholar] [CrossRef] [PubMed]
  147. Hartenstein, V. The neuroendocrine system of invertebrates: A developmental and evolutionary perspective. J. Endocrinol. 2006, 190, 555–570. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  148. Wingfield, J.C. Environmental endocrinology: Insights into the diversity of regulatory mechanisms in life cycles. Integr. Comp. Biol. 2018, 58, 790–799. [Google Scholar] [CrossRef] [PubMed]
  149. Norris, D.O. Comparative endocrinology: Past, present, and future. Integr. Comp. Biol. 2018, 58, 1033–1042. [Google Scholar] [CrossRef] [PubMed]
Genes 12 00083 g001 550
Figure 1. General structure of nuclear receptor (NR) proteins. Different domains of the NR protein are shown in different colors. A schematic representation of the ligand-binding pocket (LBP) is included in the ligand binding domain.
Figure 1. General structure of nuclear receptor (NR) proteins. Different domains of the NR protein are shown in different colors. A schematic representation of the ligand-binding pocket (LBP) is included in the ligand binding domain.
Genes 12 00083 g001
Genes 12 00083 g002 550
Figure 2. Evolution of the nuclear receptor (NR) superfamily. Subfamily origin and taxonomic distribution in metazoans. Animal phylogeny is based on Laumer et al. (2019) [28]. Subfamily diversification is based on data reported in the text [17,18,19,20,21,22,23,24,27]. Classification of NR subfamilies is as defined in Table 1. Red dots indicate events of subfamily diversification. The dotted red line, on the right, marks the NR expansion event at the base of bilaterians. Black thin dotted lines, on the right, indicate the separation between different animal clades (non-Bilateria/Bilateria and Protostomia/Deuterostomia). Grey bars, on the right, indicate the origin and conservation of NR subfamilies in metazoans. Light grey bars indicate NR subfamilies that were already present before the origin of bilaterians. Dark grey bars highlight bilaterian-specific NR subfamilies.
Figure 2. Evolution of the nuclear receptor (NR) superfamily. Subfamily origin and taxonomic distribution in metazoans. Animal phylogeny is based on Laumer et al. (2019) [28]. Subfamily diversification is based on data reported in the text [17,18,19,20,21,22,23,24,27]. Classification of NR subfamilies is as defined in Table 1. Red dots indicate events of subfamily diversification. The dotted red line, on the right, marks the NR expansion event at the base of bilaterians. Black thin dotted lines, on the right, indicate the separation between different animal clades (non-Bilateria/Bilateria and Protostomia/Deuterostomia). Grey bars, on the right, indicate the origin and conservation of NR subfamilies in metazoans. Light grey bars indicate NR subfamilies that were already present before the origin of bilaterians. Dark grey bars highlight bilaterian-specific NR subfamilies.
Genes 12 00083 g002
Genes 12 00083 g003 550
Figure 3. Examples for nuclear receptor (NR) duplications and losses during metazoan evolution. (A) Non-bilaterians. (B) Protostomes. (C) Deuterostomes. Gene expansions are shown in green and gene losses are shown in red. WGD: Whole genome duplications. Animal phylogenies are based on Laumer et al. (2019) [28]. NR nomenclature is as defined in Table 1.
Figure 3. Examples for nuclear receptor (NR) duplications and losses during metazoan evolution. (A) Non-bilaterians. (B) Protostomes. (C) Deuterostomes. Gene expansions are shown in green and gene losses are shown in red. WGD: Whole genome duplications. Animal phylogenies are based on Laumer et al. (2019) [28]. NR nomenclature is as defined in Table 1.
Genes 12 00083 g003
Genes 12 00083 g004 550
Figure 4. Nuclear receptor (NR) expression during development of marine invertebrates. (A) Cnidarians. (B) Bivalve mollusks. (C) Ascidian tunicates. NRs dynamically expressed during embryonic development (ED) (green), embryo to larva transition (EtL) (red), larval development (LD) (blue), metamorphosis (Met) (violet) and during cnidarian strobilation (orange). NR nomenclature is as defined in Table 1. Data for the cnidarians Nematostella vectensis and Aurelia aurita from Reitzel et al. (2009), Fuchs et al. (2014) and Brekhman et al. (2015) [30,64,65], for the bivalve mollusk Crassostrea gigas from Vogeler et al. (2016) and Huang et al. (2015; 2020) [19,24,70]. Raw data for the ascidian tunicates Ciona intestinalis and Phallusia mammillata are from the Aniseed database (https://www.aniseed.cnrs.fr/) and from Gomes et al. (2019) [53].
Figure 4. Nuclear receptor (NR) expression during development of marine invertebrates. (A) Cnidarians. (B) Bivalve mollusks. (C) Ascidian tunicates. NRs dynamically expressed during embryonic development (ED) (green), embryo to larva transition (EtL) (red), larval development (LD) (blue), metamorphosis (Met) (violet) and during cnidarian strobilation (orange). NR nomenclature is as defined in Table 1. Data for the cnidarians Nematostella vectensis and Aurelia aurita from Reitzel et al. (2009), Fuchs et al. (2014) and Brekhman et al. (2015) [30,64,65], for the bivalve mollusk Crassostrea gigas from Vogeler et al. (2016) and Huang et al. (2015; 2020) [19,24,70]. Raw data for the ascidian tunicates Ciona intestinalis and Phallusia mammillata are from the Aniseed database (https://www.aniseed.cnrs.fr/) and from Gomes et al. (2019) [53].
Genes 12 00083 g004
Genes 12 00083 g005 550
Figure 5. Ancestral sets of nuclear receptors (NRs) define the development of marine invertebrates. Members of the NR2 subfamily (COUP-TF, TLX/PNR, HNF4, RXR) are dynamically expressed during development of cnidarians, mollusks and tunicates. Albeit their presence in the genomes of at least medusozoan cnidarians, NR3 receptors only contribute to the developmental NR complements of the two bilaterian taxa, mollusks and tunicates. Members of the NR1 subfamily (THR, PPAR, RAR) are dynamically expressed during development of both protostomes and deuterostomes, further expanding the bilaterian set of NRs with developmental functions.
Figure 5. Ancestral sets of nuclear receptors (NRs) define the development of marine invertebrates. Members of the NR2 subfamily (COUP-TF, TLX/PNR, HNF4, RXR) are dynamically expressed during development of cnidarians, mollusks and tunicates. Albeit their presence in the genomes of at least medusozoan cnidarians, NR3 receptors only contribute to the developmental NR complements of the two bilaterian taxa, mollusks and tunicates. Members of the NR1 subfamily (THR, PPAR, RAR) are dynamically expressed during development of both protostomes and deuterostomes, further expanding the bilaterian set of NRs with developmental functions.
Genes 12 00083 g005
Table
Table 1. Nuclear receptor (NR) complements characterized in metazoan genomes.
Table 1. Nuclear receptor (NR) complements characterized in metazoan genomes.
SubfamilyGroupNameNRNC SymbolAbbreviationPhysiological Ligand
# *ANRs with two DBDsNR#A12DBD-NRα
BNR#B12DBD-NRβ
CNR#C12DBD-NRγ
DNR#D12DBD-NRδ
0A *Zygotic gap protein NR0A1KNIx
Zygotic gap protein- relatedNR0A2KNRLx
EgonNR0A3EGx
ODR-7NR0A4ODR-7x
TrithoraxNR0A5TRXx
BDosage-sensitive sex reversal-adrenal hypoplasia congenital critical region on the X chromosome, gene 1NR0B1DAX1x
Small heterodimer partnerNR0B2SHPx
1AThyroid hormone receptorNR1A1,2THRα,βT3
BRetinoic acid receptorNR1B1-3RARα-γAll-trans-RA
CPeroxisome proliferator-activated receptorNR1C1-3PPARα-γFatty acids, Prostaglandins
DRev-ErbANR1D1,2Rev-ErbAα,βx
E *Ecdysone-regulated E78 geneNR1EE78
FRAR-related orphan receptorNR1F1-3RORα-γx
HR3 *NR1F4HR3
G *CNR14-likeNR1G1Sex-1x
HLiver X receptor-like NR1H1 *EcREcdysteroids
NR1H2,3LXRα,βOxysterols
NR1H4,5FXRα,βx
IVitamin D receptor-likeNR1I1VDRVitamin D
NR1I2PXRXenobiotics
NR1I3CARAndrostane
J *NHR96NR1J1DHR96
K *VDR/PXRα,βNR1K1,2VDRα,β-like
L *HNR-like 97NR1LHR97
M *HNR-like 19NR1M1HR10
N *HNR-like 11NR1N1HR11
O * NR1O
P * NR1P1-11
2AHepatocyte nuclear factor 4NR2A1-3HNF4α,γFatty acids
NR2A4 *HNF4
BRetinoid X receptorNR2B1-3RXRα-γx
NR2B4USPx
CTesticular receptorNR2C1TR2x
NR2C2TR4x
D *DHR78NR2D1HR78
ETailless / Photoreceptor cell-specific nuclear receptorNR2E1TLXx
NR2E2 *TLLx
NR2E3PNR/HR51 *x
Dissatisfaction nuclear receptor *NR2E4DSF
Nuclear hormone receptor FAX-1 *NR2E5FAX1
FChicken ovalbumin upstream promoter transcription factorNR2F1,2COUP-TFI,IIx
Seven-up *NR2F3SVP
Chicken ovalbumin upstream promoter transcription factor III *NR2F4COUP-TFIII
Seven-up related protein 46 *NR2F5SVP-46
V-erbA-related protein 2NR2F6EAR-2x
3AEstrogen receptorNR3A1,2ERα,βEstradiol
BEstrogen-related receptorNR3B1-3 ERRα-γx
NR3B4 *ERRx
CSteroid receptor / Ketosteroid receptorsNR3C1Glucocorticoid receptor, GRCortisol
NR3C2Mineralocorticoid receptor, MRAldosterone
NR3C3Progesterone receptor, PRProgesterone
NR3C4Androgen receptor, ARTestosterone
D *Estrogen receptor-like in ProtostomiaNR3DER-like
E *Estrogen receptor-like in CnidariaNR3EER-like
F *Estrogen receptor-like in PlacozoaNR3FER-like
4ANerve growth factor IBNR4A1NGFIBx
Nuclear receptor related 1NR4A2NURR1x
Neuron-derived orphan receptor 1NR4A3NOR1x
DHR38 *NR4A4HR38
5ASteroidogenic factor 1NR5A1SF1Phosphatidylinositols
Liver receptor homolog-1NR5A2LRH1Phosphatidylinositols
NHR FTZ1-α *NR5A3FTZ1-α
B *NHR39/FTZ1-βNR5B1HR39
6AGerm cell nuclear factorNR6A1GCNFx
HR4 *NR6A2HR4
7/8 *A NR7/8A1
Non-vertebrate NR subfamilies and receptors are indicated by (*). Physiological ligands refer to in vivo conditions in vertebrates and the (x) highlights orphan NRs [12,13,14,15,16,17,18,19,20,21,22,23,24,25,26]. NRs with two DBDs are listed as (#), because they do not yet have an unequivocal subfamily status. Like NR0s, NRs with two DBDs do not have the conventional NR structure as shown in Figure 1. The NR7/8 subfamily has been called either NR7 or NR8 [14,22,24]. When information is absent or disputed, cells are left empty.
Table
Table 2. Summary of thyroid hormone receptor (THR) functions in marine invertebrates.
Table 2. Summary of thyroid hormone receptor (THR) functions in marine invertebrates.
TaxonCladePhylumReceptor ActivityDevelopmental Function
DeuterostomiaChordataTunicataUnknownSuspected role in metamorphosis
CephalochordataActivated by TRIACPivotal regulator of metamorphosis
AmbulacrariaEchinodermataPresumably ligand-activated and/or controlled by alternative signaling pathwaysSuspected role in growth, metamorphosis, skeletogenesis
ProtostomiaLophotrochozoaAnnelidaLigand-activated by T3 or TRIACRegulator of developmental transition from trochophore to crawling larva
MolluscaPresumably ligand-activated and/or controlled by alternative signaling pathwaysSuspected role in growth and developmental transitions
PlatyhelminthesPresumably ligand-activated and/or controlled by alternative signaling pathwaysSuspected role in growth
Non-BilateriaRadiataCnidariaAbsent from the genomeTHs with a role in metamorphosis, strobilation, skeletogenesis
Table
Table 3. Summary of retinoic acid receptor (RAR) functions in marine invertebrates.
Table 3. Summary of retinoic acid receptor (RAR) functions in marine invertebrates.
TaxonCladePhylumReceptor ActivityDevelopmental Function
DeuterostomiaChordataTunicataLigand-activated by retinoic acidNeurogenesis, budding
CephalochordataLigand-activated by retinoic acidNeurogenesis, axial patterning
AmbulacrariaEchinodermataLigand-activated by high concentrations of retinoic acidPresumably involved in developmental growth
ProtostomiaEcdysozoaPriapulidaLigand-activated by high concentrations of retinoic acidUnknown
HexapodaAbsent from the genomeRA with role in nervous system regeneration, tissue repair
LophotrochozoaAnnelidaLigand-activated by high concentrations of retinoic acidNeurogenesis
MolluscaLigand-binding pocket occluded and potential activation by liganded RXRNeurogenesis
Non-BilateriaRadiataCnidariaAbsent from the genomeRA with role in neurogenesis, metamorphosis, strobilation
Table
Table 4. Summary of estrogen receptor (ER) and estrogen-related receptor (ERR) functions in marine invertebrates.
Table 4. Summary of estrogen receptor (ER) and estrogen-related receptor (ERR) functions in marine invertebrates.
TaxonCladePhylumReceptor ActivityDevelopmental Function
DeuterostomiaChordata
TunicataERR: Orphan receptor
ER: Lost		ERR: Suggested role in sensory cell differentiation in the larval brain
CephalochordataERR: Orphan receptor
ER: Unknown		ERR: Suspected role in establishment of neuromuscular contacts
ER: Unknown
ProtostomiaEcdysozoa
ArthropodaERR: Orphan receptor
ER: Lost 		ERR: Control of metabolism underlying larval growth and cell proliferation
Lophotrochozoa
AnnelidaERR: Unknown
ER: Ligand-activated receptor binding estrogens 		ER: Regulation of formation and proliferation of primordial germ cells
MolluscaERR: Orphan receptor
ER: Occluded ligand binding pocket, but constitutive transcriptional activity		ERR: Unknown
ER: Unknown
RotiferaERR: Unknown
ER: Ligand-activated receptor binding estrogens		ERR: Unknown
ER: Unknown
Non-BilateriaRadiata
CnidariaER-like: Ligand-activated receptor binding paraestrol A, an ancestral estrogenER-like: Unknown
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Miglioli, A.; Canesi, L.; Gomes, I.D.L.; Schubert, M.; Dumollard, R. Nuclear Receptors and Development of Marine Invertebrates. Genes 2021, 12, 83. https://0-doi-org.brum.beds.ac.uk/10.3390/genes12010083

AMA Style

Miglioli A, Canesi L, Gomes IDL, Schubert M, Dumollard R. Nuclear Receptors and Development of Marine Invertebrates. Genes. 2021; 12(1):83. https://0-doi-org.brum.beds.ac.uk/10.3390/genes12010083

Chicago/Turabian Style

Miglioli, Angelica, Laura Canesi, Isa D. L. Gomes, Michael Schubert, and Rémi Dumollard. 2021. "Nuclear Receptors and Development of Marine Invertebrates" Genes 12, no. 1: 83. https://0-doi-org.brum.beds.ac.uk/10.3390/genes12010083

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop