International Doctoral School
Laura Cal Delgado
Doctoral dissertation
Fish Pigmentation: Functional And Evolutionary Characterization Of The Agouti Locus
Supervised by Josep Rotllant Moragas
2017
International Mention
The present thesis has been assessed by two scientific experts from non- Spanish research institutions for obtaining the international mention: Dr. Neil Ruane (Fish Helath Unit, Marine Institute, Oranmore, Ireland) and Prof. Oriol Sunyer (School of Veterinary Medicine, University of Pennsylvania, Philadelphia, USA) This work was carried out in the Institute of Marine Research of Vigo (IIM- CSIC) and funded by a grant from the Spanish Ministry of Economy and Competitiveness (AGL2011-23581). Additionally, it was partially funded by a grant from Xunta de Galicia (program AGI/CISIC (I+D+I) 2014-2016) Laura Cal Delgado was a beneficiary of a grant from Spanish Ministry of Economy and Competitiveness (FPI: BES-2012-055414) Laura Cal Delgado research stays in Eugene (Oregon, USA, 2014) and Tübingen (Germany, 2016) were funded by research stay grants of the Spanish Ministry of Economy and Competitiveness (EEBB-I-14-08631, EEBB-I-16-11802). Table of contents
TABLE OF CONTENTS
Summary…………………………………………………………………………………………. 1
Chapter 1. General Introduction and Objectives ……………………………………………. 5
General Introduction……………………………………………………………………………… 7
I. Pigment pattern……………………………………………………………………………. 9 a) Pigment cells ………………………………………………………………………………. 9 b) Pigment pattern mechanisms……………………………………………………………… 12 II. Melanocortin system…….. ……………………………………………………………… 16 a) Melanocortin receptors…………………………………………………………………….. 17 b) Melanocortin agonists…………. ………………………………………………………….. 20 c) Melanocortin antagonists…………………………………………………………………... 25
III. Pigmentation in aquaculture fish species………………………………………………… 29 a) Factors involved in pigmentary defects…………………………………………………… 30 b) Causes of pigment abnormalities: hypothesis……………………………………………. 32 c) Asip1 and pigment abnormalities………………………………………………………….. 33
Objectives…………………………………………………………………………………………. 34
Chapter 2. Genetic Analysis Of Dorsal-Ventral Pigment Pattern Formation In Zebrafish: Requirement Of Asip1………………………………………………………………………….. 37 Abstract………………………………………………………………………………………. 39
I. Introduction………………………………………………………………………………… 41 II. Material and Methods…………………………………………………………………….. 42 III. Results……………………………………………………………………………………. 45 IV. Discussion………………………………………………………………………………... 55
Chapter 3. Genetic Analysis Of Dorsal-Ventral Pigment Pattern Formation In Zebrafish: Requirement Of Mc1r ………………………………………………………………………….. 59 Abstract………………………………………………………………………………………. 61
I. Introduction………………………………………………………………………………… 63 II. Material and Methods……………………………………………………………………… 65 III. Results…………………………………………………………………………………….. 68 IV. Discussion………………………………………………………………………………… 80 Table of Contents
Chapter 4. Transcriptome Profiling Identifies Genes And Pathways Deregulated In Transgenic Fish Overexpressing asip1 ………………………………………………………………………. 85 Abstract……………………………………………………………………………………….. 87
I. Introduction…………………………………………………………………………………. 89 II. Material and Methods……………………………………………………………………… 90 III. Results…………………………………………………………………………………….. 93 IV. Discussion………………………………………………………………………………… 102
Chapter 5. Evolutionary Ancestry Of Dorsal-Ventral Pigment Asymmetry In Fish ………. 111 Abstract………………………………………………………………………………………. 113
I. Introduction………………………………………………………………………………… 115 II. Material and Methods……………………………………………………………………... 117 III. Results……………………………………………………………………………………. 122 IV. Discussion………………………………………………………………………………... 134
Chapter 6. General Discussion and Conclusions………………………………………………. 141
General Discussion……………………………………………………………………………….. 143
Conclusions……………………………………………………………………………………….. 149
References………………………………………………………………………………………… 151
Annex. Supplementary Information……………………………………………………………. 168
Summary 3
Summary
Pigment pattern formation is a classic problem in developmental biology and has been the subject of extensive experimental investigation and mathematical modelling. Vertebrate pigment patterns frequently show a clear distinction between a pale ventral area and a darker dorsal area. Thanks to the work of Greg Barsh’s lab, we have known for some time that in mammals this pattern results from spatially regulated expression of Agouti-signaling protein. The molecular basis of pigment pattern has also been widely studied in fish, but to date the emphasis has been exclusively on the mechanisms controlling striped and spotted patterns in zebrafish and its sister species. However, fish also show a pronounced dorsal-ventral pigment pattern.
In this PhD dissertation project, we show that the dorsal-ventral patterning in zebrafish is also Agouti-signaling protein-dependent, and that the striping mechanism is largely independently superimposed upon this evolutionarily-conserved dorsal-ventral patterning mechanism. Our observations are, we believe, the first to show a “loss-of- function” mutation of the signaling molecule (Agouti-signaling protein, Asip1) and its receptor (Mc1r) and to demonstrate that disruption of their expression abolishes the dorsal-ventral pigment pattern difference. Combined with the data from mammals, our data strongly suggests that an Agouti-signaling protein-dependent mechanism regulating dorsal-ventral pigment pattern is a conserved feature of vertebrates, so that other patterning mechanisms, whether generating stripes or spots or other patterns, are likely generated by superimposing secondary patterning mechanisms. Our conclusion is particularly interesting because the cellular basis for the patterns in mammals (and birds) and in fish appear very different, with the former depending on a shift in the biochemistry of melanin produced within melanocytes, whereas the latter seems to affect the ratios of black melanocytes and other pigment cell-types, the yellow xanthophores and the silver iridophore. Our data indicates that at the cellular level there is some conservation of the mechanisms, with Asip1 and Mc1r affecting melanin production (total levels, not melanin type), but also likely affecting the ratio of pigment cell types in different body regions. Additionally, our results also provide evidence that the mechanism leading to the dorsal-ventral pigment pattern was likely in place before 4 Summary the origin of teleosts. Finally, the results of this PhD project unquestionably confirmed the pleiotropic nature of the agouti gene. Thus revealing the ability of asip1 gene to simultaneously regulate pigmentation and other physiological processes in fish skin.
Lastly, we have to say that this work has been a cross-disciplinary and combined cutting-edge approaches from comparative genomics, cell and developmental biology, reverse genetics, and evolutionary biology.
Overall, we feel this PhD project not only provide basic knowledge on vertebrate pigmentation development, but also provide information relevant to the design of new strategies for intensive fish culture.
General Introduction and Objectives 7
GENERAL INTRODUCTION
Color patterns in animal species are involved in a wide range of functions. Although the significance of animal coloration is poorly understood, pigment patterns have been associated with camouflage (Dice and Blossom 1937; Belk and Smith 1996; Protas and Patel 2008; Rudh and Qvarnström 2013), forage success (Tso et al. 2002), thermoregulation (Ellis 1980; Rudh and Qvarnström 2013), photoprotection (Brenner and Hearing 2008; Rudh and Qvarnström 2013) or mate selection (Kodric-Brown and Nicoletto 2001; Protas and Patel 2008; Bajer et al. 2011; Maan and Sefc 2013). Coloration is composed of structural and pigmentary colors. Structural colors are produced by the interaction between the light and tissue nanostructures (Parker and Martini 2006; Roberts et al. 2012). Examples of structural colors are light reflection in iridophores (fish, amphibians and reptiles) (Kawaguti 1965; Olsson et al. 2013; Rudh and Qvarnström 2013), light scattering in bird barbules (Roulin and Ducrest 2013), or diffraction gratings in antenna hairs of some crustaceans (Parker and Martini 2006). Pigmentary colors are based on chemical pigments (Prum 2006; Olsson et al. 2013; Roulin and Ducrest 2013), deposited in specific organelles (chromatosomes) contained within pigment cells (chromatophores) (Fujii 2000). Pigmentary colors are frequently associated with structural colors since pigment cells are distributed in layers in the integument structures (Prum 2006; Olsson et al. 2013; Roulin and Ducrest 2013).
Fish, especially teleosts, show high pigment diversity. Teleost genomic duplication (TGD) and the functional divergence of pigmentation ohnologue genes have been suggested as the possible causes of this diversity (Braasch et al. 2008). The variability of pigment patterns is extraordinary with the dorsal-ventral countershading (dark dorsum and pale ventrum) pigment pattern being one of the most distinguishable color patterns in vertebrates.
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General Introduction and Objectives 9
I. Pigment pattern
Unlike in mammals, the color of adult fish is determined by the superposition of two different pigmentation patterns. For example, it has been shown that zebrafish have two distinct adult pigment patterning mechanisms - an ancient dorsal-ventral patterning mechanism based on dorsal-ventral differential asip1 gene expression, and a more recent striping mechanism based on cell-cell interactions (Ceinos et al. 2015). However, these genetically shaped pigment-patterning mechanisms can be changed to some extent in response to different environmental stimuli and represent one of the most captivating features in fish pigmentation. Additionally, fish can exhibit diverse patterns of pigmentation depending on their state of development. The larval color phenotype, for example, is commonly different from the adult phenotype (Kelsh et al. 2009; Parichy et al. 2011). Many pigmentary defects may result from alterations during metamorphosis from the larval phenotype to the adult phenotype because an apparently normal body color pattern in fish larvae occasionally results in abnormal adult pigmentation (Bolker and Hill 2000; Darias et al. 2013b; Ceinos et al. 2015).
a) Pigment cells
Pigment cells derive from neural crest cell population, a type of multipotent stem cells which produce intermediate precursors of specific cell types (Le Douarin and Dupin 2003). Those intermediate precursors migrate to their final position whereas they develop to pigment cells, glia cells and neurons of peripheral system (Raible and Eisen 1994; Le Douarin and Kalcheim 1999; Le Douarin and Dupin 2003).
Pigment pattern formation in vertebrates has been widely studied. Mammals and birds only have one pigment cell type in the skin, the melanocyte, which produces two types of melanin: dark/brown eumelanin or yellow/red pheomelanin (Barsh 1996; Lin and Fisher 2007; Hubbard et al. 2010). In contrast, amphibians, reptiles and fish present several pigment cell types (chromatophores) each one synthesizing chemically distinct pigment (Bagnara and Matsumoto 2006): melanophores (eumelanin), xanthophores and erythrophores (pteridines and/or carotenoids) and iridophores (guanine light-reflecting plateles). Interestingly, contrary to birds and 10 Chapter 1
mammals, pheomelanin is not produced by fish or reptiles (Olsson et al. 2013; Kottler et al. 2015). Additionally, some fish species also show leucophores (guanine light-reflecting plateles) (Fujii 1993) and cyanophores (unknown blue pigment) (Goda and Fujii 1995; Kelsh 2004; Bagnara and Matsumoto 2006) (Fig. 1.1).
Figure 1.1. Phylogenetic relationships, pigment cell repertoires and genome assemblies of important teleost fishes and other vertebrate groups. The timings of the fish-specific genome duplication (FSGD), the two rounds of genome duplication early in the vertebrate lineage (1R and 2R) as well as the salmonid-specific tetraploidization (T) are indicated. Black circle are melanophore- melanocytes, yellow/orange circles are xanhtophore- erythrophores, grey circles are iridophores, white circles are leucophores and blue circles are cyanophores. (Braasch et al. 2008).
The pigment pattern in mammals is mainly dependent on the type of melanin that is synthesized in melanocytes and also on the shape of melanocyte (Fig 1.2A) (Slominski 2004; Ito and Wakamatsu 2011; Roulin and Ducrest 2013). Fish pigment patterns result by the combination and patterned distribution of different pigment cell types (Fig. 1.2B) (Fujii 2000; Kelsh 2004; Kelsh et al. 2009; Olsson et al. 2013). General Introduction and Objectives 11
Figure 1.2. Comparisons of the cellular composition of adult color pattern in mammals and fish. (A) In mouse, melanocytes contain melanosomes that can produce eumelanin (dark pigment) or pheomelanin (light pigment), different ratios resulting in darker or lighter coloration. (B) In zebrafish, the adult pigment pattern is based on dark stripes and light interstripes. Different fish chromatophores produce chemically different pigments, so the fish pigment pattern is obtained by a specific distribution of different pigment cells types. Dark stripes are formed by four layers of chromatophores: type S iridophores (high number and uniform size of platelets), xanthophores , melanophores and type L iridophores (low number and large size of platelets), while light interstripes are composed of two layers: a xanthophore layer and a type S iridophore layer (Adapted from Kelsh (2004) and Irion et al. (2016)).
The dark blue coloration of stripes in zebrafish (Danio rerio), for example, is formed by four layers of pigment cells. The most external layer is composed of stellate, scattered and pale yellow xanthophores, which is followed by a thin layer of “loose” bluish type S iridophores (uniform size and high number of platelets). A melanophore layer appears beneath of iridophores type S, and the deepest layer is an iridophores Type L (large size and low number of platelets) layer (Hirata et al. 2003; Frohnhöfer et al. 2013; Mahalwar et al. 2014; Watanabe and Kondo 2015). Conversely, the iridiscent/golden coloration of interstripes in zebrafish is achieved by two layers: a layer of dense xanthophores covering the deepest layer of dense iridophores type S (Fig. 1.3) (Hirata et al. 2003; Frohnhöfer et al. 2013; Mahalwar et al. 2014; Watanabe and Kondo 2015). A similar organization of pigment cells has been found in Turbot (Scophthalmus maximus). Turbot melanophores and xanthophores were found in the most superficial pigment layer and the iridophores in the deepest layer in dorsal skin; only a single layer of iridophores exists in light 12 Chapter 1
ventral skin (Faílde et al. 2014). These different colorations of dorsal skin in turbot are reached by different quantities pigment cell types. Dark turbots show higher number of melanophores, brown-yellowish fish show higher number of xanthophores and brown-greyish turbots show similar number of melanophores and xanthophores (Faílde et al. 2014). The specific organization and amount of the different pigment cell types is fundamental for different pigment patterns and colorations.
Fig. 1.3. Scheme of layered organization of pigment cells in the striped pattern of zebrafish. Stripes are formed by the following layers from top to bottom: a thin layer of stellate xanthophores, a thin layer of “loose” S-iridophores, a layer of melanophores and a final layer of L-iridophores. Interstripes are organized in two layers: a more superficial dense xanthophore layer and a deeper dense S- iridophore layer.
b) Pigment pattern mechanisms
i) The cell-cell interaction mechanism (Turing mechanism)
The cellular and molecular mechanisms underlying the pigment pattern formation have been widely studied in the last decade. In fish, most studies on pigment patterning have focused on stripe formation of zebrafish, in which a core striping mechanism dependent upon interactions between different pigment cell types has been identified (Maderspacher and Nüsslein-Volhard 2003; Takahashi and Kondo 2008; Frohnhöfer et al. 2013; Singh et al. 2014). Each pigment cell type tends to General Introduction and Objectives 13
completely cover the skin; however, the presence of different pigment cell types forces them to interact with each other, producing the stripe pattern (Frohnhöfer et al. 2013; Irion et al. 2016; Mahalwar et al. 2016). Thus, zebrafish iridophores from the first interstripe are responsible for setting the pattern of adult stripe formation at the beginning of adulthood (Frohnhöfer et al. 2013), and the complete striped pattern results from heterotypic and homotypic interactions between every pigment cell (Frohnhöfer et al. 2013; Mahalwar et al. 2016; Walderich et al. 2016). Iridophores and melanophores present a short-range repulsion which causes the melanophores aggregation; moreover, iridophores and xanthophores exhibit mutual attraction, while xanthophores and melanophores repel each other (Frohnhöfer et al. 2013). Iridophores and xanthophores have a positive long-range effect on melanophores aggregation and survival (Frohnhöfer et al. 2013; Irion et al. 2016) (Fig. 1.4). Furthermore, feed-back mechanisms also interact between pigment cells. Thus, iridophores are required for xanthophores organization while xanthophores and melanophores are responsible for the location of iridophores (Frohnhöfer et al. 2013; Irion et al. 2014, 2016). Sole (Solea senegalensis) iridophores are restricted to melanophore-free areas, and xanthophores seems to repel melanophores (Darias et al. 2013). Therefore, although some of the cellular and molecular features of pattern formation still remain to be elucidated, present evidence strongly suggests that the underlying mechanism is mathematically equivalent to the mechanism postulated by Adan Turing more than half a century ago (Watanabe and Kondo 2015).
Figure 1.4. Scheme of interactions between chromatophores cell types. Red curved arrows are long-range intractions; black arrows, short-range interactions. (Frohnhöfer et al. 2013). Iridophores are represented as grey double stripes in the up-left position; xanthophores are represented as little yellow circles in the up-right position; melanophores are represented as larger black circles in the down position.
14 Chapter 1
ii) The countershading patter mechanism.
Most of vertebrate pigment pattern include a light ventrum and dark dorsum. Although the most thoroughly studied mechanism in fish is the striped pattern of zebrafish, it has only recently been demonstrated that zebrafish use two distinct adult pigment patterning mechanisms: a more recent striping mechanism based on cell-cell interactions, and an ancient dorsal-ventral patterning mechanism, present in all vertebrates (Ceinos et al. 2015). The agouti signaling protein has been identified as the paracrine factor that drives this common dorsal-ventral countershading pattern in mammals and also in birds (Lu et al. 1994; Siracusa 1994; Le Pape et al. 2008; Norris and Whan 2008; Fontanesi et al. 2012; Oribe et al. 2012; Chandramohan et al. 2013). A mammalian agouti orthologue has also been identified in fish (asip1) (Cerdá-Reverter et al. 2005; Kurokawa et al. 2006). In amphibians, the melanisation inhibiting factor (MIF) (Fukuzawa and Ide 1988; Fukuzawa et al. 1995) has been proposed as the potential mammalian agouti orthologue (Cerdá-Reverter et al. 2005). All this suggests that the melanocortin system is the key system responsible for the dorsal-ventral countershading pattern in vertebrates, which is widely described in the section II.
iii) Physiological and morphological color changes
Apart from the different genetically shaped pigment-patterning mechanisms that determine the final color of an organism, some organisms have evolved body coloration plasticity, so called color change, in order to cope with distinct backgrounds. Fish color changes are produced by two different mechanisms: physiological and morphological. Physiological color change is caused by short-term stimuli and is based on the aggregation or dispersion of pigment granules (chromatosomes) in skin chromatophores (Sköld et al. 2016) or changes in the distance and angle between light-reflectin platelets in motile iridophores (Fujii 2000). Physiological color change is controlled by both sympathetic and endocrine systems (Fujii 2000). In the former, noradrenalin has been shown to induce chromatosome aggregation (Fujii 2000; Aspengren et al. 2003; Biswas et al. 2014), while in the endocrine system, several hormones are involved in pigmentation. Two melanin concentrating hormones (Mch1 and Mch2) play an important role in pigment granule General Introduction and Objectives 15
aggregation (Fujii 2000; Berman et al. 2009; Kang and Kim 2013; Mizusawa et al. 2013, 2015). The α-melanocyte stimulating hormone (α-Msh) induces pigment granule dispersion (Fujii and Miyashita 1982; Fujii 2000; Yamanome et al. 2007). During physiological color change, noradrenalin and MCH are commonly released on light backgrounds (Fig. 1.5A). By contrast, α-Msh plasma levels are increased on dark backgrounds (Fig. 1.5B) (Kawauchi et al. 1983; Sugimoto 2002; Logan et al. 2006; Mizusawa et al. 2013).
Fig. 1.5. Schematic diagram of sympathetic and endocrine control of skin pigmentation on (A) light and (B) dark backgrounds. α-Msh, α-Melanocyte stimulating hormone; Mch, Melanin concentrating hormone; Na, Noradrenalin; black circles represent melanosomes.
Morphological color change is caused by long-term stimuli and it is mediated by apoptosis or proliferation of skin chromatophores as well as changes in their morphology and amounts of pigment (Sköld et al. 2016). Several studies have demonstrated that fish change their color by decreasing or increasing the number and size of melanophores under long-term adaptation to light or dark background, respectively; the opposite response is also observed in the number of iridophores (Sugimoto et al. 2000, 2005; van der Salm et al. 2005). Both physiological and morphological color changes appear to be controlled by similar mechanisms that include the melanocortin system.
16 Chapter 1
II. Melanocortin system
The teleost fish melanocortin system consists of five melanocortin receptor subtypes (Mc1r, Mc2r, Mc3r, Mc4r, Mc5r), the endogenous agonists alpha and beta melanocyte stimulating hormone (Msh) and adrenocorticotropic hormone (Acth), which are melanocortin peptides derived from the proopiomelanocortin (pomc) gene, and the endogenous antagonists of the agouti signaling protein family (Asip1, Asip2, Agrp1 and Agrp2) (see figure 1.6).
Figure 1.6. The melanocortin system and fish pigmentation. Schematic drawing with a hypothetical regulatory model in the control of pigmentation through different melanocortin system mechanisms in fish chromatophores; (A) melanophores, (B) xantophores and (C) iridophores. (↑), stimulation; (┬), inhibition; (dotted line), unknown.
General Introduction and Objectives 17
a) Melanocortin Receptors
The melanocortin system exerts its multiple functions via a number of G-protein couplet receptor (GPCRs), the melanocortin receptors (Mcrs). Within the GPCR family, MCRs are included in the rhodopsin class, family A-13. So far five Mcrs have been identified (Mc1r-Mc5r), each differing in its spatial distributin, ligand affinities and specificity, and therefore with a distinctive physiological role. Mcrs show the characteristic structure of GPCR family: seven transmembrane helix domains connected by three extra- and intracellular loops, an extracellular N- terminus and a intracellular C-terminus (Abdel-Malek 2001; Kobilka 2007). A high degree of identity and conservation in structural characteristics and pharmacology exists among Mcrs from fish and mammals (Schiöth et al. 2005). In fish, at least five melanocortin receptors also mediate the diverse actions of melanocortins (Mc1r, Mc2r, Mc3r, Mc4r, Mc5r). In mammals Mc1r is mainly expressed in skin and it is involved in coloration, Mc2r is expressed in adrenal gland and controls steroidogenesis, Mc3r and Mc4r are expressed in the brain and they are involved in the regulation of energy balance, and Mc5r is expressed in different tissues and it is involved in exocrine secretion (Cone 2006; Cooray and Clark 2011). In fish, the number, affinity, specificity, tissue distribution and the physiological roles are far from defined and they appear to be species-specific. In this context, for example, pufferfish (Tetraodon nigroviridis) has only four receptors with no Mc3r (Schiöth et al. 2005), while zebrafish has six receptors with two copies of the Mc5r (Kumar et al. 2011). Thus, the greater number of Mcr paralogs found in some telost species were probably originated in the extra round of whole genome duplication that occurred during early teleost evolution (3R or TGD; Teleost Genome Duplication) (Braasch et al. 2008). In fish, mc1r is expressed in skin of the sea bass (Dicentrarchus labrax) and of the Mexican tetra cavefish (Astyanax mexicanus) and it is also involved in pigmentation in cavefish (Gross et al. 2009; Sánchez et al. 2010); mc2r is expressed in zebrafish interrenal tissue and controls cortisol synthesis (To et al. 2007); mc3r is mainly expressed in the pufferfish (Takifugu rubripes) central nervous system (CNS) (Klovins et al. 2004b); mc4r is also expressed in the sea bass CNS (Sánchez et al. 2009b) and in the zebrafish peripheral tissues and it is involved in food intake (Cerdá-Reverter et al. 2003a; Forlano and Cone 2007); and finally mc5r is expressed 18 Chapter 1
in different sea bass tissues and it is involved in the regulation of hepatic lipid metabolism and, probably, pigmentation in sea bass (Sánchez et al. 2009a). Although mc5r is duplicated in zebrafish, only one form is present in most teleost genomas (Ringholm et al. 2002).
In mammals, the melanocortin-1 receptor (MC1R) is a key regulator of pigmentation. Both in mammals and fish, the melanocortin-1 receptor (MC1R/Mc1r) is activated by α-Msh, which induces cAMP production (García-Borrón et al. 2005; Sánchez et al. 2010). In turn, in mammals this cAMP increase activates, via protein kinase A (PKA), the melanin synthesis pathway that involves the melanogenic enzimes (tyrosinase (TYR), Tyrosinase-related protein 1 (TYRP1) and dopachrome tautomerase (DCT)), which result in de novo dark brown/black eumelanin pigment synthesis (Buscà and Ballotti 2000; García-Borrón et al. 2005). This pathway is assumed to be the same in teleosts, but the best-known effect of α-Msh-Mc1r binding in fish is the pigment dispersion within melanophores (Fujii 2000). In fish and mammals, melanin is produced and deposited within specific organelles called melanosmomes (Hearing 2000). Interestingly, melanosomes are dispersed by the effect of α-Msh also via cAMP pathway in fish melanophores (Sugimoto et al. 1997; Logan et al. 2006; Kobayashi et al. 2009), which results in the skin darkening usually involved in physiological coloration.
Sequence polymorphisms in the Mc1r gene are associated with differences in pigmentation in amniotes (Valverde et al. 1995; Dun et al. 2007) and various mutations can result in an overactive MC1R causing hyperpigmentation (melanism) in several mammalian and bird species (Robbins et al. 1993; Newton et al. 2000; Eizirik et al. 2003; Ling et al. 2003; Nadeau et al. 2006); in turn several polymorphisms related with “loss-of-function” mutations produce hypopigmentation (Valverde et al. 1995; Dun et al. 2007). Interestingly, a two base pair deletion mc1r mutant apparently seems to produce hypopigmentation in cavefish Astyanax mexicanus (Gross et al. 2009).
The physiological function of Mc1r can be antagonized by the paracrine signaling molecule protein Agouti (Asip, see below), which is responsible for the switch to the production of yellowish red pheomelanin pigment in amniotes (Cone et General Introduction and Objectives 19
al. 1996) and for the inhibition of melanosome dispersion in fish (Cerdá-Reverter et al. 2005). As mentioned above, it is widely accepted that fish only synthesize eumelanin. Thus, pheomelanin has not been identified in fish (Adachi et al. 2005; Braasch et al. 2009a; Kottler et al. 2015). As in mammals, Mc1r is encoded for a single exon gene in fish and it is present as a single gene with no extra paralog even in teleost fish (Selz et al. 2007). The mc1r gene encodes a protein with the common structure of GPCR family: three intracellular loops, three extracellular loops and seven transmembrane helix domains (García-Borrón et al. 2005). It has been suggested that its C-terminus has an important role in coupling, transport and correct anchoring in the plasma membrane (Selz et al. 2007). Moreover, the intracellular phosphorylation sites are important for controlling Mc1r activity and numerous conserved sites have been identified in several vertebrate species (Fig. 1.7) (García- Borrón et al. 2005; Selz et al. 2007). The expression of mc1r gene has been reported in the skin of most fish species studied so far: zebrafish, medaka (Oryzias latipes), platyfish (Xiphophorus maculatus) (Selz et al. 2007); cavefish (Gross et al. 2009); Japanese flounder (Paralichthys olivaceus) (Kobayashi et al. 2012a); goldfish (Carassius auratus) (Kobayashi et al. 2011b) and Japanese ornamental carp (Cyprinus carpio var. Koi) (Bar et al. 2013); and sea bass (Sánchez et al. 2010). mc1r expression has also been found in other tissues and its expression levels vary among fish species (Klovins et al. 2004a; Selz et al. 2007; Sánchez et al. 2010; Kobayashi et al. 2010; Takahashi et al. 2014).
In fish, α-Msh and Acth affinity to Mc1r differs among species and depends on their acetylation level (Klovins et al. 2004a; Sánchez et al. 2010; Kobayashi et al. 2010, 2012a). Interestingly, mc1r expression has been reported in zebrafish, barfin flounder (Verasper moseri) and Japanese flounder melanophores (Kobayashi et al. 2010, 2012a; Higdon et al. 2013), in goldfish xanthophores (Kobayashi et al. 2011a) and in zebrafish iridophores (Higdon et al. 2013). Moreover, mc1r is not the only melanocortin receptor gene expressed in fish chromatophores. The expression of mc5r was also detected in melanophores and xanthophores in barfin flounder and Japanese flounder (Kobayashi et al. 2010, 2012b), and shows strong expression in the skin of goldfish (Cerdá-Reverter et al. 2003a) and sea bass (Sánchez et al. 2009a). Additionally, taking into account that Acth (a melanocortin agonist) may 20 Chapter 1
have a role in regulating fish pigmentation (Fujii 2000) and that Acth acts on the Mc2r (Aluru and Vijayan 2008) and low levels of mc2r expression, were detected in the skin of sea bass (Agulleiro et al. 2013), the involvement of Mc2r in fish pigmentation might be a possibility and merits study in more detail.
Figure 1.7. Predicted structure of platyfish melanocortin type 1 receptor (Accession number DQ866828). The positions of the transmembrane (TM) domains have been drawn according to human MC1R (www.GPCR.org; Accession number AF326275). Potential N- terminal glycosylation sites are shown. (Selz et al. 2007)
b) Melanocortin Agonists
Melanocortin receptors are stimulated by agonists, the melanocortins, a group of peptide hormones that include ACTH and the different forms of MSH (Dores and Lecaude 2005). Melanocortins derive from the post-translational cleavage of proopiomelanocortin (POMC) (Nakanishi et al. 1979) and are expressed mainly in corticotrope and melanotrope cells of the pituitary gland (Cerdá-Reverter and Canosa 2009). Comparisons of Pomc post-translational processing in several tissues showed that the precursor is cleaved in a tissue specific (pituitary, skin, central nervous tissue and placenta) manner in both mammals and fish (Eberle 2000; Cerdá-Reverter and Canosa 2009; Takahashi and Mizusawa 2013). The main enzymes cleaving POMC General Introduction and Objectives 21
precursors are prohormone convertase 1 (PC1) and prohormone convertase 2 (PC2) (Bertagna 1994). In corticotrope cells, PC1 cleaves POMC to produce ACTH and β- lipotrophin, but in the melanotrope cells the combined action of PC1 and PC2 cleave the precursor to produce α-MSH and β-endorphin (Barr 1991; Bertagna 1994; Metz et al. 2006). After cleavage, POMC derived peptides are under the control of different modifications including acetylation and amidation, which modulate peptide activity and receptor binding (Wilkinson 2006)
Acth is considered the main factor mediating the pituitary control of corticosteroid synthesis/secretion during fish stress response (Wendelaar-Bonga et al. 1994). Although it has been shown to mediate the pituitary control of fish stress response (Wendelaar-Bonga et al. 1994), Msh, is best known for the induction of melanosome dispersion in fish melanophores (Bagnara and Hadley 1973). The precursor Pomc includes different Mshs, i.e. α-melanocyte stimulating hormone (α- Msh), β-melanocyte stimulating hormone (β-Msh) and γ-melanocyte stimulating hormone (γ-Msh) (which is lost in teleosts) all of them characterized by a core HFRW sequence and localized in different domains of the precursor, i.e. γ-Msh in the N-terminal pro-γ-Msh domain, α-Msh in the Avth domain and β-Msh in the C- terminal β-lipotrophin domain (Fig. 1.8) (Barr 1991; Bertagna 1994; Metz et al. 2006).
Figure 1.8. Scheme of POMC postraslational modifications. POMC is the precursor of several peptides hormones. POMC is modified differently in corticotrope cells than in melanotrope cells. This scheme summarizes the tissue-specific POMC-derived products in fish. In corticotrope cells the ACTH is the major product produced, but it is possible to find other products depending on the specie. In melanotrope cells many different products are produced, some of them class-specific. An asterisk (*) indicates a region in the POMC sequence that is lost in teleosts. Two asterisks (**) indicate a specific region that appears only in condrichcyes.
22 Chapter 1
α-MSH induces the synthesis of the dark brown/black eumelanin pigment in vivo and in vitro disperses melanosomes by MC1R binding in vertebrates (Sugimoto et al. 1997; García-Borrón et al. 2005; Yamanome et al. 2007). In vitro experiments in mammals have shown that α-MSH-MC1R binding results in an increase of intracellular cAMP levels as a result of the stimulation of adenylyl cyclase (AC),
Figure 1.9. The melanocortin system in mammalian pigmentation. Schematic drawing with the regulatory pathway in the control of pigmentation of α-MSH through MC1R in mammalian melanocyte. Abbreviations: α-MSH, α-Melanocyte Stimulating Hormone; AC, Adenylyl Cyclase; ASIP, Agouti Signalling Protein; cAMP, Cyclic Adenosine Monophosphate; CRE, cAMP Responsive Element; CREB, cAMP Responsive Element Binding Protein; Cys, Cysteine; DCT, Dopachrome Tautomerase; MITF, Microphthalmia Transcription Factor; MC1R, Melanocortin 1 Receptor; PKA, Protein Kinase A; TYR, Tyrosinase; TYRP1, Tyrosinase Related Protein 1.
which is coupled to MC1R (Kreiner et al. 1973). Additionally, high levels of intracellular cAMP activate PKA (Roesler et al. 1998) and induce the activation of melanogenic enzimes mediated by microphtalmia transcription factor (MITF) (Bertolotto et al. 1998). In vitro tests showed that tyrosinase is the essential and rate- limiting enzyme in the melanin biosynthesis (Kobayashi et al. 1995). Tyrosinase- related protein 1 (TRP1) and dopachrome tautomerase (DCT) are another two well- characterized melanogenic enzimes essential for eumelanin production (Buscà and Ballotti 2000). Trp1 and Dct expressions are also mediated by MITF and they are General Introduction and Objectives 23
also induced by intracellular cAMP increase (Buscà and Ballotti 2000). This is the classical pathway by which α-MSH is assumed to mediate its melanogenic effects on melanocytes in mammals (Fig. 1.9).
In fish, α-Msh disperses melanosomes in both dermal and epidermal melanophores in vitro (Fujii and Miyashita 1982) and stimulates melanophore proliferation and melanin synthesis in vivo (Fujii 2000; Yamanome et al. 2007). A high degree of conservation of melanin pathway has been reported in vertebrates (Hoekstra 2006; Braasch et al. 2009b). It seems that the melanogenic effect of α-Msh is also mediated by binding to Mc1r. In vivo, α-Msh stimulates Mc1r signal transduction, which is coupled to the activation of adenylyl cyclase, resulting in increased cytosolic levels of cAMP, which activates PKA (Sugimoto et al. 1997; Sánchez et al. 2010). Moreover, teleostean microphtalmia transcription factor a (mitfa) gene also presents a conserved cAMP responsive element (CRE) region in the promoter (Li et al. 2013) and, like mammalian Mitf, induces melanization in vivo (Lister et al. 1999). Nevertheless in fish, in vitro experiments have related α-Msh with rapid melanosome dispersion within melanophores (Sugimoto et al. 1997; Logan et al. 2006; Kobayashi et al. 2009; Mizusawa et al. 2013). In vitro experiments showed that such melanosome dispersion is also regulated through of the cAMP pathway (Rozdzial and Haimo 1986), and is dependent on extracellular Ca2+ ions (Fujii 2000; Logan et al. 2006). This suggests differences in function between fish and mammalian Mc1r. However, long-term effects of α-Msh include melanophore proliferation and melanin synthesis in vivo (Yamanome et al. 2007), which is consistent with the α-Msh effect in mammals. Therefore, it should also be noted that there are significant differences in the physiology of pigmentation between fish and mammals: the presence of different pigment cells types, the production of only eumelanin in fish melanophores and the reversible dispersion and aggregation of melanosomes for rapid color change. Moreover, it has been shown that, in vitro, α- Msh stimulates not only melanophores but also other chromatophores – xanthophores and iridophores (Matsumoto et al. 1977; Iga and Matsuno 1986; Kobayashi et al. 2011a). In vitro, α-Msh promotes xanthosome dispersion (Kobayashi et al. 2011a, 2012a) and may promote platelet aggregation in specific type of iridophores (motile iridophores (Fujii 2000)). 24 Chapter 1
In fish, αMSH is involved in the process of physiological (short-term) (Mizusawa et al. 2013) and morphological (long-term) (Arends et al. 2000) background adaptation. The transfer of white-adapted animals to a black background results in dispersion of melanophore pigment within a few hours (short-term) and increases plasma α-Msh levels (Mizusawa et al. 2013). However, the role of α-Msh in fish long-term background adaptation has been studied for several species (van Eys and Peters 1981; Baker et al. 1984; Gilham and Baker 1984; Rotllant et al. 2003; van der Salm et al. 2005; Mizusawa et al. 2013), but no consensus regarding α-Msh behavior in each type of background has emerged, which suggests a possible multiple endocrine control of background adaptation and/or species-specific role of α-Msh in long-term background adaptation.
A crucial factor that may determine α-Msh melanogenic activity is the acetylation degree at its N-terminal domain (Wilkinson 2006; Takahashi and Mizusawa 2013). Lower levels of monoacetyl-α-Msh have been found in sea bream adapted to white compared with a black backgound (Arends et al. 2000). In tilapia (Oreochromis mossambicus), the greatest pigment dispersion was produced by monoacetyl-α-Msh (van der Salm et al. 2005). On the contrary, monoacetyl-α-Msh had no effect on melanosome movements in barfin flounder and Japanese flounder, but did affect pigment dispersion in flounder xanthophores (Kobayashi et al. 2010, 2012b). In goldfish, both diacetyl-α-and monoacetyl-α-Msh stimulate pigment dispersion in xanthophores (Kobayashi et al. 2012b, a). It therefore appears that there is no consistent response between α-Msh acetylation levels and pigment dispersion among different fish species (Arends et al. 2000; Rotllant et al. 2003; van der Salm et al. 2005; Mizusawa et al. 2013). Additionally, recent in vitro pharmacological receptor-binding studies in Japanese flounder have shown that dimerization of Mc1r and Mc5r creates a ligand-dependant signal modulation (Kobayashi et al. 2016). The authors specifically showed that heterodimerization of Mc1r and Mc5r controls the inhibition of cAMP production in Japanese flounder melanophores as a result of α-Msh but not of diacetyl-α-Msh activation.
General Introduction and Objectives 25
c) Melanocortin Antagonists
Another interesting feature of melanocortin receptors is the presence of endogenous antagonists that include agouti signaling protein (ASIP) and the agouti- related protein (AGRP). ASIP is a paracrine signaling protein involved in mammalian pigmentation, which competes with α-MSH by binding to MC1R and MC4R (Lu et al. 1994). AGRP is a neuropeptide involved in energy balance, body weight regulation and metabolism in mammals, acting as antagonist of MC3R and MC4R (Ollmann et al. 1997). It has been extensively reported that the ASIP regulates pigmentation in mammals (Siracusa 1994; Lu et al. 1994; Norris and Whan 2008; Fontanesi et al. 2012; Chandramohan et al. 2013). For its part, Asip is a gene composed of four exons that encode a 131 amino acid protein. This protein can be structurally divided into a N-terminal basic domain that is rich in lysine (K) residues, followed by a proline (P)-rich region localized just before the cysteine C-terminal domain (Bultman et al. 1992). The cysteine rich C-terminus was identified as the key region for the competitive antagonism with MC1R (Ollmann and Barsh 1999) and the C-terminal loop was shown to be the crucial domain for the inhibition of MC1R (Patel et al. 2010), while the amino-terminal residues were seen to be essential down- regulating MC1R (Ollmann and Barsh 1999).
Asip is expressed in the mammalian dermal papillae of hair follicles (Millar et al. 1995) and promotes the switch from eumelanin (dark/brown pigment) to pheomelanin (yellow/red pigment) synthesis (Miller et al. 1993) acting as a α-MSH competitive antagonist of the MC1 receptor (Ollmann et al. 1998). Additionally, ASIP inhibits also the basal melanogenesis, acting as α-MSH inverse agonist (Aberdam et al. 1998; Chai et al. 2003) (Fig. 1.10). 26 Chapter 1
Figure 1.10. The melanocortin system in mammalian pigmentation. Schematic drawing with the regulatory pathway in the control of pigmentation of ASIP through MC1R in mammalian melanocyte. Abbreviations: α-MSH, α-Melanocyte Stimulating Hormone; AC, Adenylyl Cyclase; ASIP, Agouti Signalling Protein; cAMP, Cyclic Adenosine Monophosphate; CRE, cAMP Responsive Element; CREB, cAMP Responsive Element Binding Protein; Cys, Cysteine; DCT, Dopachrome Tautomerase; MITF, Microphthalmia Transcription Factor; MC1R, Melanocortin 1 Receptor; PKA, Protein Kinase A; TYR, Tyrosinase; TYRP1, Tyrosinase Related Protein 1.
Additionally, in mouse (Mus musculus) some studies have also reported the effect of ASP (ASIP) on melanocyte differentiation and maturation (Aberdam et al. 1998; Manceau et al. 2011). Furthermore, dorsal-ventral variations in mouse hair coloration were shown to be due to different ASP expression levels (Manceau et al. 2011), and the expression of different ASP isoforms in dorsal and ventral skin respectively as result of the use of alternative promoters (Vrieling et al. 1994).
In contrast to the two Agouti family genes in tetrapods, the teleostean agouti gene family is more complex with four of these melanocortin antagonist genes that were, initially termed asip1, asip2 agrp1, and agrp2 (Kurokawa et al. 2006). The connectivity of teleost to mammalian gene function depends on understanding gene orthologies and it was suggested that asip2 and agrp2 are ohnologs of asip/asip1 and agrp1, respectively, from the teleost genome duplication (TGD) (Kurokawa et al. 2006). This interpretation of the evolutionary relationships of the four genes in teleosts, however, is controversial. Phylogenetic analyses alone are unlikely to General Introduction and Objectives 27
resolve their evolutionary history (Schioth et al. 2011; Braasch and Postlethwait 2011), and alternative data are needed.
A conserved synteny-based model (Braasch and Postlethwait 2011) proposes that the agrp/asip precursor duplicated in the R1 and R2 rounds of vertebrate genome duplication and that Agrp2 and Asip2 went missing from tetrapods but asip2 was retained in the lineage leading to teleosts. This asip2 gene was then further duplicated in the TGD, giving rise to the teleostean genes initially called asip2 and agrp2 (Kurokawa et al. 2006) and which should be called asip2a and asip2b (Braasch and Postlethwait 2011). According to this model, the second duplicates from the TGD of both asip1 and agrp1 were lost in teleosts, so that there is only one agrp type gene in teleosts but three of the asip type genes (asip1, asip2a, asip2b) (Braasch and Postlethwait 2011) (Table.1.1). This conserved synteny-based model, however, has been criticized by some (Schioth et al. 2011; Västermark et al. 2012) and will require further analysis in the future. Of relevance for our discussion here is that there are three teleostean genes with known functions for pigmentation patterning: asip1, asip2b (aka agrp2), and agrp1. To avoid confusion we have followed the conventional Kurokawa’s et al. (2006) gene nomenclature.
Table 1.1. Two different nomenclatures for the Agouti gene family have been proposed: the earlier Kurokawa nomenclature and the recently proposed Braasch & Postlethwait nomenclature. In order to avoid confusion, we have used the conventional Kurokawa nomenclature.
28 Chapter 1
Dorsal-ventral countershading in mammals is driven by the regulation of Asip expression and the resulting type of melanin synthesis in differentiated melanocytes. In fish, the dorsal-ventral countershading is, in contrast, achieved by differences in the distribution of different types of pigment cells (Bagnara and Matsumoto 2006). Despite the different countershading mechanisms in fish and in mammals, it has recently been shown that fish agouti (asip1) shows a dorsal-ventral gradient expression in the skin of different adult fish species (with lower levels in the dorsum and high levels in the belly (Cerdá-Reverter et al. 2005; Kurokawa et al. 2006; Guillot et al. 2012; Agulleiro et al. 2014; Ceinos et al. 2015). This expression pattern correlates with the dorsal-ventral pigment pattern showing darkly colored skin in the dorsal area and a light ventrum.
Additionally, it has also been shown that overexpression of asip1 or injection of capped asip1 mRNA induces lightening of dorsal skin in zebrafish (Fig. 1.11) (Ceinos et al. 2015) and flatfish (Guillot et al. 2012) respectively. Moreover, asip1
Figure 1.11. Disruption of the dorsal-ventral expression gradient in transgenic zebrafish overexpressing asip1 leads to the elimination of dorsal melanophores pigmentation Lateral view of adult pigment pattern of wild type (WT) and transgenic agouti zebrafish [asip1-Tg; line Tg(Xla.Eef1a1:Cau.Asip1)iim05;(Ceinos et al. 2015)] (A) Wildtype fish have a pattern with dark stripes and light interstripes. (B) Asip1-Tg fish show a dramatic dorsal-ventral reduction in melanophore number. Scale bar: 1cm.
mRNA ectopic overexpression and injection in zebrafish and turbot results in reduced expression of tyrosinase-related protein-1 (tyrp1) gene, a key enzyme of melanin synthesis (Guillot et al. 2012; Ceinos et al. 2015). It should also be General Introduction and Objectives 29
mentioned that Asip1 inhibits α-Msh-induced melanosome movement in the medaka scales via Mc1r (Cerdá-Reverter et al. 2005). All this evidence suggests that agouti (Asip1) has a conserved function in bony vertebrates, but that it may act through a different cellular mechanism in mammals and in ray-finned fish.
Apart from the obvious role of Asip1 in fish pigmentation, it has been reported that zebrafish Asip2b/Agrp2, which is mainly expressed in the pineal gland and brain antagonizes Mc1r. This melanocortin receptor is highly expressed in zebrafish hypothalamus and up-regulates proMch (pmch) and proMch-like (pmchl) genes, which regulates background adaptation (Zhang et al. 2010). The agrp2/asip2b gene is also expressed in the skin of different teleost species like zebrafish (Zhang et al. 2010), sea bass (Agulleiro et al. 2014), Atlantic salmon (Salmo salar) (Murashita et al. 2009) and fugu (Takifugu rubripes) (Kurokawa et al. 2006). Another endogenous antagonist probably involved in pigmentation is Agrp1/Agrp, which has been reported as inverse agonist of Mc1r in sea bass, where it is thought to affect pigmentation (Sánchez et al. 2010). Histological studies have revealed that agrp1 is expressed in the skin of fugu (Kurokawa et al. 2006), sea bass (Agulleiro et al. 2014), Atlantic salmon (Murashita et al. 2009) and goldfish (Cerdá-Reverter et al. 2003b). Despite of the above evidence, and the fact that induced stable overexpression of Agrp1/Agrp does not alter zebrafish pigment pattern phenotype (Song and Cone 2007), the role of Agrp1/Agrp on fish pigmentation has yet to be adequately explored.
III. Pigmentation in Aquaculture fish species
Malpigmentation is a common problem in fish production and a common cause of losses in aquaculture. Abnormal pigmentation has been reported in several species of fish, but particularly in flatfish (Purchase et al. 2002; Macieira et al. 2006; Akyol and Şen 2012). Normal pigmentation in flatfish is based on high number of melanophores on the ocular side and low number on the blind side, which results in a dark ocular side and light blind side (Venizelos and Benetti 1999; Bolker and Hill 2000). The most common pigment defects in flatfish are: 30 Chapter 1
Albinism, pseudo-albinism or hypomelanosis: from complete absence of pigmentation to white areas on the ocular side (Venizelos and Benetti 1999; Bolker and Hill 2000).
Hypermelanosis or ambicoloration: from complete pigmentation to pigmented areas on the blind side (Venizelos and Benetti 1999; Bolker and Hill 2000).
It has been shown that pigment defects are not restricted to cultured populations. Several cases of wild malpigmented adults have been reported in many species of flatfish like Solea solea (Akyol and Şen 2012; Cerím et al. 2016), Achirus declivis (Macieira et al. 2006), Achirus lineatus (Macieira et al. 2006), Trinectes maculatus (Moore and Posey 1974) Kareius bicoloratus (Fujita 1980), Paralichthys patagonicus (Astarloa et al. 2006), Solea vulgaris (Paris and Quignard 1968), among others. However, the best-known incidence of pigment abnormalities have been reported in flatfish hatcheries (Bolker and Hill 2000).
a) Factors involved in pigmentary defects
Numerous studies have suggested different putative factors that may be involved in pigment abnormalities. In flatfish, the developmental stage preceding metamorphosis was identified as the crucial period when environmental factors can strongly disturb the standard pigment patter formation (Næss and Lie 1998). This period is also known as “pigmentation window” (Izquierdo and Koven 2011).
One of the main factors implicated in flatfish malpigmentation is feeding, and the diet composition is often considered as essential aspect for improving pigment development in cultured flatfish species. The correct ratio and absolute amount of docosahexaenoic acid (DHA), eicosapentaenoic acid (EPA) and arachidonic acid (ARA) has been considered essential for achieving the standard skin pigmentation (Sargent et al. 1999; Izquierdo and Koven 2011). The specific moment for feeding with the optimum ratio during the “pigmentation window” is also a key aspect to take in account (Næss and Lie 1998; Izquierdo and Koven 2011; Boglino et al. 2014). Vitamin A, retinoic acid, has also been found highly relevant for pigment development in flatfish. Optimal levels of vitamin A prevents abnormal pigmentation General Introduction and Objectives 31
but an excess produces developmental problems, e.g. skeletal malformation (Miki et al. 1990; Takeuchi et al. 1995). Vitamin A is involved in multiple biological functions, so it is difficult to establish the optimal level for this nutrient (Bolker and Hill 2000).
Other aspects identified as important factors in fish pigment development are stress conditions and endocrine disorders. Ambicoloration in flatfish has been associated with high levels of cortisol, which indicates that stress during rearing conditions is a possible factor for pigment abnormalities (Yamada et al. 2011). This could also be related with the malpigmentation linked to substratum type. It was suggested that flatfish that have the possibility to bury themselves in a sandy substratum show lower pigment defects than others without such an opportuinty (Stickney and White 1975; Ottesen and Strand 1996). Endocrine disorders play a role in pigmentation through thyroid hormone, which produces pigment defects when it is over- or underexpressed (Yoo et al. 2000; McMenamin et al. 2014). Thyroid hormone was identified as a factor involved in adult pigment pattern development through its inhibition of melanophore number and stimulation of xanthophore differentiation in zebrafish (McMenamin et al. 2014). Additionally, in flatfish, an excess of thyroid hormone results in high incidence of albinism, which suggests that thyroid hormone also inhibitd melanophore proliferation in flatfish (Yoo et al. 2000). Other endocrine disorders could be the deregulation of α-Msh production, which has been suggested as a factor of malpigmentation in flatfish after methamorphosis (Bolker and Hill 2000; Itoh et al. 2012). Itoh et al. (2012) describe that some of the environmental factors which result in a high incidence of pigment defects are related with the light-brain-pituitary axis and seem to be able to disrupt normal α-Msh synthesis and secretion in flatfish. Moreover, Mch1 and Mch2, which are also endocrine hypothalamo-pituitary peptides, may be also involved in flatfish pigmentary defects. Low mch gene expression was suggested as the potential cause of malpigmentation in flounder (Kang and Kim 2013). It is clear that further studies into α-MSH and MCHs regulation and pigment abnormalities are needed.
32 Chapter 1
b) Causes of pigment abnormalities: hypothesis
The complexity of pigment pattern establishment mechanisms in fish makes it difficult to reduce the likelihood of pigment defects occurring in aquaculture fish species. Two hypotheses have been put forward to explain the pigment abnormalities in flatfish.
The first suggests that nutritional deficits result in visual defects, which interfere with the endocrine signaling pathway necessary for melanophore differentiation (Kanazawa 1993). Kanazawa (1993) suggested that deficient ratios of DHA, EPA and Vitamin A produce rhodopsin malformation in flatfish eye. Subsequent studies on the lipid composition of neural tissues in abnormally pigmented fish showed that they have different contents of DHA, EPA and ARA from normal pigmented fish (Estevez and Kanazawa 1996). Additionally, neural alterations and eye degeneration were reported in flatfish as a result of deficient intake of amino and fatty acids (Estevez et al. 1997). Taking together, these data support the first hypothesis: the signal pathway from the retina to the central nervous system could be disturbed by nutritional insufficiencies. This disturbance might be the cause of inappropriate production of α-Msh in the pituitary, which could produce pigment abnormalities in flatfish (Bolker and Hill 2000).
The second hypothesis proposes that pigment defects in flatfish are the consequence of the deregulation of the mechanisms responsible for the establishment of the ocular- and blind-side skin structures during methamorphosis (Seikai and Matsumoto 1994). Bolker et al. (2005) histologically analyzed normal pigmented and malpigmented skin and also propose that the pigment abnormalities may be caused by the normal regulatory pathway in the incorrect side. Recently, other studies showed that the establishment of both the asymmetrical body conformation and the adult pigment pattern is synchronized (Darias et al. 2013a), and that several deregulated genes are responsible for the pigmentary defects in adult pigment pattern in flatfish such as asip1 (Guillot et al. 2012; Darias et al. 2013b), somatolactin, paired box protein 3, tyrosinase, mitf, mc1r, mast/stem cell growth factor receptor Kit, sodium/potassium/calcium exchanger 5 and tyrosinase-related protein 1 (Darias et al. 2013b). Therefore, it has also hypothesized by Guillot et al General Introduction and Objectives 33
(2012) and Darias et al (2013b) that pigmentary defects could be the result of the establishment of pigment pattern mechanisms on the wrong side of flatfish.
These two hypotheses are not mutually exclusive (Bolker and Hill 2000). Indeed, both hypotheses suggest an effect on different mechanisms, among them the regulation of different melanogenic genes and different melanocortin system participants (α-Msh, Mc1r, Asip1), which are known to be involved in dorsal-ventral countershading pattern in mammals.
c) Asip1 and pigment abnormalities
As stated previously, several deregulated genes are responsible for the pigmentary defects in adult pigment pattern in flatfish (Guillot et al. 2012; Darias et al. 2013b). Among others, asip1, that is commonly more expressed in blind-side skin, is deregulated in malpigmented fish. In adult sole and turbot, ocular-side non- pigmented spots show abnormally high levels of asip1 mRNA (Guillot et al. 2012). Moreover, Guillot et al. (2012) noted that higher levels in ocular-side light spots are similar to blind-side asip1 levels. Interestingly, the malpigmentation caused by nutritional defects in sole is accompanied by asip1 mRNA up-regulation compared to normally pigmented fish ((Darias et al. 2013b). Therefore, asip1 deregulation has been proposed as a potential cause of pseudoalbinism (Guillot et al. 2012; Darias et al. 2013b). Whatever the case, asip1 regulation seems to have a crucial role in pigment abnormalities.
34 Chapter 1
OBJECTIVES
The fundamental scientific objective of this PhD dissertation is to do an integrated study that evaluates the molecular regulation of the dorsal-ventral pigment patterning in fish through the characterization of the agouti signaling protein (asip1) gene and its receptor, melanocortin receptor 1 (mc1r). The specific aims of this thesis are to characterise the Asip1-Mc1r system and determine how this system is regulated and functions in fish pigmentation.
To do that end, a model vertebrate specie, the zebrafish (Danio rerio) and spotted gar (Lepisosteus oculatus) were used due to the availability of genomic resources, embryos and larvae morphological characteristics and the availability of newly developed genetic manipulation tools (transgenic and knockout gene approaches).
The project focuses on three major objectives:
1. Genetic analysis of dorsal-ventral pigment pattern formation. The first objective of this PhD Project was to determine and understand the molecular genetic mechanisms that determines the dorsal-ventral pigment pattern in fish. To achive that, we created “loss-of-function” mutations in the asip1 (Chapter 2) and mc1r (Chapter 3) gene using CRISPR/Cas9 system. Results from Chapter 2 and Chapter 3 reveal the molecular genetic mechanism that establishes the dorsal-ventral pigmentation asymmetry in fish.
2. A comparative transcriptomic analysis of an altered dorsal- ventral pattern of color in zebrafish. Using this transgenic model that ectopically overexpresses asip1, we analyzed the transcriptomic profiling induced by decreased activity of the central melanocortin system, and characterized the different biological processe and/or gene networks downstream of the melanocortin system in zebrafish skin. Results form Chapter 4 uncover that dorsal-ventral pigment disruption-associated asip1 overexpression results in several other key General Introduction and Objectives 35
regulatory processes strongly affected, such as immune system, cell cycle and energetic metabolism.
3. Characterization of conserved regulatory elements required for the evolutionary conserved dorsal-ventral asip1 expression in the adult fish. The Third objective of this PhD Project was to determine the evolutionary ancestry of dorsal-ventral pigment asymmetry in fish. To achive that, we compared two ray-finned fishes: the teleost zebrafish (Danio rerio) and the non-teleost spotted gar (Lepisosteus oculatus) and to generate zebrafish transgenic lines carrying a bacterial artificial chromosome (BAC) from zebrafish or spotted gar. Results form Chapter 5 reveal that the mechanism leading to an important dorsal- ventral pigment pattern was likely in place before the origin of teleosts.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 39
ABSTRACT
Vertebrate pigment patterns frequently show a clear distinction between a pale ventral region and a darker dorsal region. We have known for some time that in mammals this pattern results from spatially regulated expression of Agouti-signaling protein (ASIP). In fish, little is known about the nature and interaction of the genes controlling this process. So far only asip1 have been identified that, when over- expressed, can disrupt the dorsal-ventral countershading pattern in adult zebrafish. However, “loss-of-function” analysis has not yet been reported. Using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tools, we introduced a mutation into the zebrafish genome, creating an asip1 zebrafish homozygous knockout mutant. Knockout mutant zebrafish that were homozygous for the asip1 allele displayed disorganized dorsal-ventral pigment pattern characterized by a dorsalization (i.e. a darkening due to an increased ratio of melanophores and xanthophores) of the ventral pigment pattern. These data suggest that a mutation in asip1 is associated with the disruption of dorsal-ventral pigment pattern in zebrafish, likely through a mechanism that affect melanophore, xanthophore and iridophore number and/or differentiation in the ventral region.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 41
I. Introduction
In teleost fish, the color pattern is obtained by the interaction and specific distribution of different pigment cell types. The widespread countershading pattern, based on a dark dorsum and light ventrum is reached by high numbers of melanophores in the dorsal skin and a high numbers of iridophores in the ventral regions (Hirata et al. 2005). Recently, the existence of two independent pigmentation mechanisms has been suggested – an ancient and conserved Asip1-dependent mechanism (Ceinos et al. 2015) and a more recent mechanism based on cell-cell interactions (Frohnhöfer et al. 2013; Irion et al. 2016). To date, most studies have focused on the stripped pigment pattern of zebrafish (Danio rerio), and how the black melanophores, yellow xanthophores and silver iridophores interact with each other to establish the pattern by their specific location in the fish skin (Patterson and Parichy 2013; Frohnhöfer et al. 2013; Singh et al. 2014; Irion et al. 2016). The homotypic and heterotypic cell-cell interactions between all these three pigment cell types is essential for the stripe pattern formation (see Chapter 1) (Maderspacher and Nüsslein-Volhard 2003; Nakamasu et al. 2009; Frohnhöfer et al. 2013; Watanabe and Kondo 2015). However, the dorsal-ventral pigment pattern establishment has been largely overlooked. In zebrafish, a high number of melanophores are accumulated in the dorsal skin, but they are almost completely absent in the ventral skin. Furthermore, accumulation of iridophores are present in the abdominal wall producing the light color of the zebrafish belly (Hirata et al. 2005). In mammals, it is well known that the Asip-Mc1r interaction is the key mechanism, which produces the dorsal-ventral pigment pattern. In fish, the Asip1 capability to bind to Melanocortin 1 receptor (Mc1r) and to inhibit melanin dispersion has already been shown in in vitro experiments (Cerdá-Reverter et al. 2005) and high ventral levels of asip1 expression were found in several fish species (Cerdá-Reverter et al. 2005; Kurokawa et al. 2006; Guillot et al. 2012; Agulleiro et al. 2014; Ceinos et al. 2015). Previously, the effect of asip1 ectopic overexpression in zebrafish was analyzed. Ceinos et al. (2015) showed that asip1 gene presented a dorsal-ventral gradient expression in WT zebrafish skin, with higher expression ventrally and decreased toward the dorsum. The disruption of this dorsal-ventral expression gradient by ectopic overexpression 42 Chapter 2 of asip1 in the whole body results in a loss of the dorsal-ventral countershading pattern. asip1 overexpression significantly reduces the melanophore number in dorsal skin by inhibition of melanophore proliferation/differentiation (Ceinos et al. 2015). Nevertheless, a “loss-of-function” approach is needed to entirely understand the mechanism that control regional differences in pigmentation. Thus, in this chapter we have investigated further the in vivo functional role of asip1 in zebrafish by generating asip1 zebrafish homozygous knockout mutants, using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tool (Bassett et al. 2013). We demonstrate that asip1 knockout mutant zebrafish displayed a disorganized dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (i.e. ventral skin becomes nearly as dark colored as dorsal skin due to an increased ratio of melanophores and xanthophores) of skin color. In contrast, we do not see any effects on the core-striped pattern in asip1 mutant fish, which all stripes still visible.
These results indicate that “loss-of-function” mutation of asip1 gene is associated with the disruption of dorsal-ventral pigment pattern in zebrafish, likely through a mechanism that affect melanophore, xanthophore and iridophores number and/or differentiation and therefore provide new insights into the mechanism that underlie region-specific difference in body morphology.
II. Materials and methods
a) Fish
Zebrafish were cultured as previously described (Westerfield 2007) and staged according to Kimmel et al., (1995). Fish of the following genotypes were used: TU strain (Tübingen, Nüsslein-Volhard Lab), Tg(TDL358:GFP) (Levesque et al. 2013) and Tg(kita:GalTA4:UAS:mCherry) (Anelli et al. 2009). Ethical approval (Ref. Number: AGL2011-23581) for all studies was obtained from the Institutional Animal Care and Use Committee of the IIM-CSIC Institute in accordance with the National Advisory Committee for Laboratory Animal Research Guidelines licensed by the Spanish Authority (RD53/2013) and conformed to European animal directive (2010/63/UE) for the protection of experimental animals.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 43
b) Generation and analysis of asip1 knockout mutants
The asip1 “loss-of-function” mutation was generated using the CRISPR-Cas9 system. The CRISPR protocol, originally adapted from Bassett et al. (2013), was kindly provided by Sam Peterson (U. Oregon). The potential target sequence was identified with the ChopChop web tool (Montague et al. 2014). Two long oligonucleotides (Scaffold oligo: 5`- GATCCGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCT TATTTTAACTTGCTATTTCTAGCTCTAAAAC-3`, and gene-specific oligo 5’- AATTAATACGACTCACTATAGCACACACACATGCCAATGGGTTTTAGAGC TAGAAATAGC-3’) were used to perform a DNA-free PCR to obtain a 125 bp DNA fragment that includes the previously identified target site sequence (5’- GCACACACACATGCCAATGG-3’). The PCR reaction was performed in 20 μL containing 10 μL of 2x Phusion High-Fidelity PCR Master Mix Buffer (New England Biolabs, UK), 1 μL of gene specific oligo (10 μM), 1 μL of gRNA scaffold oligo (10 μM) and H2O nuclease free to 20 μL. PCR conditions were 98ºC for 30 sec, 40 cycles of 98ºC for 10 sec, 60ºC for 10 sec, 72ºC for 15 sec, and a final step of 72ºC for 10 min. The PCR product was purified using DNA Clean&Concentration-5 Kit (Zymo Research, USA) according to the manufacturer’s instructions. Purified PCR product was used as template for in vitro transcription with MEGAscript T7 High yeld transcription Kit (Ambion, USA) according to the manufacturer’s instructions. The gRNA was purified with RNA Clean&Concentrator 5 (Zymo Research, USA) before to use it.
The gRNA was injected in a concentration of 25 ng/µL together with Cas9 mRNA (transcribed from the linearized pT3TS-nCas9n plasmid) in a concentration of 50 ng/µL and Phenol red solution (0,1%). Around 2 nL of this mix was microinjected into the cytoplasm of zebrafish eggs at the one- or two-cell stage. Dissection microscope (MZ8, Leica) equipped with a MPPI-2 pressure injector (ASI systems) was used for microinjection. Different mutations were found and three different potential nonfunctional mutations were raised as different asip1 knockout lines. The phenotype in each knockout stable line was exactly the same. For microscope imaging, zebrafish of 5dpf, 15 dpf, 30dpf and 180dpf were anesthetized
44 Chapter 2 with tricaine methasulfonate (MS-222-, Sigma-Aldrich) scales were isolated from the belly and immersed in PBS on a glass slide. Scales and fish were photographed with a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera.
Double reporter transgenic/asip1 mutant lines were obtained by setting up crosses between the asip1 mutant line and a reporter transgenic line Tg(TDL358:GFP); which label iridophores (Levesque et al. 2013), or a reporter transgenic line Tg(kita:GalTA4:UAS:mCherry), which label melanophores (Anelli et al. 2009). The offspring of these crosses were incrossed to obtain homozygous asip1 knockout mutants. Confocal imaging was carried out in a Leica TCS SP5 confocal microscope. 5dpf, 15dpf and 30 dpf transgenic zebrafish were anesthetized and photographed. Adult zebrafish (180dpf) were anesthetized with MS-222 and decapitated to sample a ventral skin section including the abdominal wall. This section was placed in PBS and photographed.
c) Melanophore and xanthophore counts
The melanophore pattern of asip1 knockout mutant fish was compared with control fish by quantification of melanized melanophores in both groups. Selected region for melanophore counting was different in each stage of development. In early stage (5dpf), we counted melanophores in a 1mm2 area in a dorsal view on the head and the dorsal area, on the horizontal myoseptum and in a ventral view of the head. In early metamorphic stage (15dpf), we counted melanophores in a 1 mm2 area in a dorsal view on the head, on the dorsal area, on the horizontal myoseptum and in a ventral view of the head and the belly. In late metamorphic stage (30dpf), we counted melanophores in a 1 mm2 area in a dorsal view on the head, dorsal area, horizontal myoseptum and in a ventral view on the head and belly. In adult fish (60 and 210 dpf) melanophores within 1 mm2 area were counted in several distinct positions: in a dorsal view on the head (head area) and on the dorsal area (from the edge of the head to edge of the dorsal fin); in a lateral view, on the 2D, 1D, 1V and 2V anterior areas (pectoral to pelvic fin); and finally, in a ventral view of the head and the belly (pectoral to pelvic fin) (see Fig. 2.4A). The dorsal-ventral xanthophore pattern of asip1 knockout mutant fish was compared with control fish by quantification of pigmented xanthophores in postmetamorphic fish (60 and 210 dpf). Selected regions for xanthophore counting were on the dorsal area (from the edge of
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 45 the head to edge of the dorsal fin), and in a ventral view of the belly (pectoral to pelvic fin) (see Fig. 2.4A). To analyze the number of melanophores and xanthophores, six fish per group were anesthetized with tricaine methasulfonate (MS-222-, Sigma-Aldrich) and immersed in 10 mg/ml epinephrine (Sigma) solution for 30 min to contract melanosomes. Fish photographed with a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera. Melanophores were counted using ADOBE PHOTOSHOP CS2 software (Adobe Systems Software Adobe Systems Ibérica SL, Barcelona, Spain) and the ImageJ software (National Institutes of Health, NIH, Maryland, USA).
III. RESULTS
a) Analysis and selection of induced “loss-of-function” mutations
“Loss-of-function” mutations in the asip1 gene were generated using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tools. We selected the target site in the first coding exon (60 bp after ATG) (Fig. 2.1A,B). We found ten different mutated alleles (Fig. 2.1B). Alleles M1, M3, M5 and M6 conserved the original open reading frame; therefore, they could generate a functional protein lacking only one or two amino acids and keeping almost the complete amino acid sequence. Alleles M2, M4, M7, M8, M9 and M10 show different reading frames downstream of the target site. The asip1 gene encodes a predicted protein of 125 amino acids. We selected three potential frameshift mutations, which usually the resulting protein is nonfunctional and raised the asip1K.O. lines to characterize the phenotype: M2 (CRISPR1-asip1.iim02 zebrafish line), M7 (CRISPR1-asip1.iim07 zebrafish line) and M8 (CRISPR1-asip1.iim08 zebrafish line) (Fig. 2.1B). The asip1iim02 allele lacks 11 bp (Del 76-86), the asip1iim07 allele has lost 4 bp (Del 77-81), and asip1iim08 lacks 16 bp (Del 62-76) and carries a 15 bp insertion at position 62 downstream of the predicted ATG (Fig. 2.1B). In those three alleles, the mutation lead to a frameshift and a premature stop codon. The asip1iim02 encodes a 71 amino acids mutant protein, the asip1iim07 encodes a predicted mutant protein with 38 amino acids, and asip1iim08 encodes a mutant protein with 39
46 Chapter 2 amino acids (Fig. 2.1C). All mutated proteins have lost the most of their basic central domain and, more interestingly, the C-terminal poly-cysteine domain, which is the crucial region for protein activity (Manne et al. 1995; McNulty et al. 2005; Patel et al. 2010). All asip1 knockout mutant zebrafish lines examined resulted in the same specific color phenotype.
Figure 2.1. CRISPR/Cas9-induced mutations at asip1 locus. (A) Scheme of the asip1 gene showing the target site mutation (black arrowhead). Coding exons are represented as white boxes and 5’ UTR and 3’UTR are shown as black boxes. (B) Sequence of induced deletions in asip1 locus. The first line shows the wild-type sequence. Black arrowhead labels the protospacer-adjacent motif (PAM). Next lines show different induced mutations. Italic lower case letters represent inserted new sequence. The number of deleted (-) and inserted (+) bases are marked in the right side of each sequence. Selected mutations are labeled by red arrowheads. (C) Predicted amino acid sequence encoded for asip1 loci. The first line shows the wild type protein (black), and following lines show the potential protein sequence of each selected mutation (blue). Green boxes show the wild type sequence conserved. Asterisk represents the stop codon. Cysteine residues from the putative C-terminal poly- cysteine domain of the Asip1 protein are labeled in red.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 47
b) asip1 function in dorsal-ventral pigment patterning.
To investigate the role of Asip1 in dorsal-ventral pigment pattern formation in zebrafish, we created “loss-of-function” mutations in the asip1 gene using CRISPR/Cas9 system. Fish homozygous for these knockout alleles (asip1K.O.) develop exactly the same phenotype as adults, with an irregular dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (i.e. ventral skin becomes nearly as dark colored as dorsal skin due to an increased ratio of melanophores and xanthophores) of skin color (Fig. 2.2). In the following analyses, we focus on line CRISPR1-asip1.iim08. Compared to WT sibling asip1K.O. fish had increased pigmentation over the entire ventral region (Fig. 2.2A-D), including the head (Fig.2.2. E,F). In asip1K.O. fish, melanophores and xanthophores seem to be able to proliferate in all ventral regions. This is contrary to WT fish, where the ventral regions are a restricted area for melanopohores, and xanthophores are mostly limited to the posterior area of the belly, around of the pelvic fins (Cal, personal observations). WT phenotype shows low number of melanophores in the ventral head region and high number of iridophores around branchiostegals and operculum (Fig. 2.2E). Nevertheless, asip1K.O. phenotype shows melanophores all around the jaws, branchiostegals and operculum (Fig. 2.2F). The ventral skin of WT fish shows almost a total absence of melanophores, which allows seeing the bright white of an iridophore sheet from the abdominal wall (Fig. 2.2G). Conversely, asip1K.O. fish have melanophores and xanthophores in the ventral skin; besides, the abdominal wall is affected: iridophore sheet is broken up into small fragments due to a decreasing of iridophores number. This confers darker color to ventral region of asip1K.O. fish (Fig. 2.2H). All those pigment defects caused by a “loss-of-function” mutation of asip1 gene were restricted to dorsal-ventral pigment pattern.
48 Chapter 2
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dominal wall seems to be also affected because it looks yellower than WT. Scale bar: (A,B) 5 mm, (C,D, E, F, G, H) 2 mm. 2 H) G, E, F, (C,D, mm, 5 (A,B) bar: Scale WT. than yellower it looks because affected also be to seems wall dominal
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Figure Figure 2 views views of 180 dpf WT and stripe is named a with are code: called stripes two primary 1D and and 1V, the are stripes 2V. 2D and two Thenamed secondary The is modification that 2V is than in stripe WT, the wider and 3V is dark The ventral darker belly stripe of developed. best operculum and jaw branchiostegal, however, and jaws branchiostegals; the around number reduced in appear melanophores WT, the asip1 abdominal wall. However, the ab
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 49
c) The development of the asip1K.O. phenotype.
To establish the time point when the phenotype of the asip1 mutants (asip1K.O.) becomes first apparent, melanophores were counted at larval (5dpf, SL 3 mm), metamorphic (15 dpf, SL 6.3 mm and 30 dpf, SL 7 mm) and two adult stages (60 dpf, SL 13 mm and 210 dpf, SL 25 mm) (Figures 2.3 and 2.4). It has been shown that pigment pattern changes during development can be distinguish by an increase in the melanophores number and changes in their distribution (Kelsh 2004; Parichy et al. 2009) We have quantified the distribution of melanophores in WT and asip1K.O. fish along the dorsal-ventral axis. No differences in melanophore numbers were found at larval stages (5dpf, SL 3 mm) (data not shown). At the beginning of the metamorphosis (15dpf, SL 6.3 mm), the dorsal-ventral pigment abnormalities begin to be visible. At 15dpf there are no differences in melanophore number in the belly between asip1K.O. and WT fish. However, the number of melanophores in the ventral area of the head was 68.7% higher in asip1K.O. fish (P<0.05) than in WT fish (Fig. 2.3A). At 30dpf pigment abnormalities finally appear also in the belly. The number of melanophores in the ventral view of the head was 63% higher (P<0.05) than in WT fish, and in the asip1K.O. belly was 41% higher than WT belly (P<0.05) (Fig. 2.3B).
Figure 2.3. Dorsal-ventral distribution of melanophores during metamorphosis. (A) Distribution and number of melanophores in WT and asip1K.O. 15dpf fish. At this stage, asip1K.O. shows significantly higher number of melanophores in the ventral view of the head (B) Distribution and number of melanophores in WT and asip1K.O. 30 dpf fish. At this stage, asip1K.O. shows significantly higher number of melanophores in the ventral view of the head, but also in the belly. Data are the mean ±SEM, n=7. Asterisks indicate significant differences between WT and asip1K.O. fish. Scale bar: (A) 200 μm, (B) 500 μm.
50 Chapter 2
The asip1K.O. fish at 60 and 210 dpf showed significant pigment pattern alterations, particularly in the ventral region (Fig. 2.4B). At 60 dpf, the number of skin melanophores in dark stripe 2V of asip1K.O. fish was 47% higher (P<0.001) than in WT fish, in the ventral region of the head was 86% higher (P<0.001) than in WT fish, and in the belly of asip1.KO. fish was 98% higher (P<0.001) than in WT belly. No differences were found in dorsal regions or in other dark stripes (Fig. 2.4C). Furthermore, we found that the number of xanthophores was also affected in ventral regions. At 60 dpf, the distribution of xanthophores in anterior area of the belly was 98% higher (P<0.05) than in WT. No differences were found in dorsal region (Fig. 2.4D). At 210 dpf, the same pattern of a higher number of melanophores in the ventral region was found. The number of melanpohores in the dark stripe 2V of asip1K.O. fish was 38% higher (P<0.001) than in WT fish, in the dark stripe 3V was 78.6% higher (P<0.001), in the ventral region of the head was 84% higher (P<0.001), and in the belly of asip1 KO fish was 99% higher than in WT belly (P<0.001). Similar to 60 dpf fish, the pigment defects were restricted to ventral regions (Fig. 2.4E). At 210 dpf, the distribution of xanthophores in the belly region was the same as at 60 dpf. No differences of number of xanthophores were found in dorsal regions. However, the number of xanthophores in the anterior region of the belly was 96% higher than in WT fish (P<0.001) (Fig. 2.4F).
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 51
Figure 2.4. Dorsal-ventral distribution of melanophores and xanthopores in WT and asip1K.O. fish. (A) Lateral view of zebrafish showing the body regions selected for melanophore and xanthophore count. (B) Ventral view of the WT and asip1K.O. 210 dpf zebrafish fish belly. (C) Distribution and number of melanophores in WT and asip1K.O. 60dpf fish. At this stage, asip1K.O. shows a significantly higher number of melanophores in the black stripe 2V, ventral head and belly. (D) Number of xanthophores in the dorsal and ventral skin of WT and asip1K.O. 60dpf fish. At this stage, asip1K.O. shows a significantly higher number of xanthophores in the belly region. (E) Distribution and number of melanophores in WT and asip1K.O. 210 dpf fish. At this stage, asip1K.O. shows significantly higher number of melanophores also in black stripe 2V, 3V, ventral head and belly. (F) Number of xanthophores in dorsal and ventral skin of WT and asip1K.O. 210 dpf fish. These fish showed highly significant higher number of xanthophores in belly region than WT. Data are the mean ±SEM, n=7. Asterisks indicate significant differences between WT and asip1K.O. fish. Scale bar (A,C,E) 1mm, (B) 100 μm.
52 Chapter 2
To allow a more detailed visualization of the pigment cells in asip1 mutants, we imaged fish carrying the Tg(Kita:Gal4,UAS:mCherry) transgene, which labels melanophores with membrane-bound mCherry (Anelli et al. 2009). In WT, melanophores are no detectable in the ventral skin region (Fig. 2.5A,B). However, in asip1 mutants, we observed an increase of melanophores in the ventral skin region (Fig. 2.5C,D). This is in agreement with the observed increase in the number of melanophores in asip1K.O. at later stages of development (Fig. 2.4). Analysing fish carrying Tg(TDL358:GFP) transgene, which label iridophores and glia with cytosolic GFP (Levesque et al. 2013), we observed that in asip1 mutants, the dense and uniform sheet of iridophores found in the abdominal wall of WT fish (Fig. 2.5F) is broken up into small fragments (Fig. 2.5G,H). Thus, asip1 mutants showed a strong reduction of the iridophores number interspersed with many melanophores (Fig. 2.5G, black arrow) and some xanthophores (Fig. 2.5G, orange arrow) along the abdominal wall.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 53
Figure 2.5. Detailed visualization of ventral pigment cells in WT and asip1 mutants . (A) Ventral view of 210 dpf WT belly. (B) Belly of 210 dpf WT fish carrying Tg(Kita:GAL4;UAS:mCherry) (labels melanophores) transgene shows no melanophores in ventral skin. (C) Ventral view of 210 dpf asip1K.O. belly. (D) Belly of 210 dpf asip1K.O. fish carrying Tg(Kita:GAL4;UAS:mCherry) transgene shows high number of melanophores in ventral skin. (E) Internal view of 210 dpf WT abdominal wall shows a white sheet of iridophores with few internal melanophores (black arrow). (F) Abdominal wall of 210 dpf WT fish carrying Tg(TDL358:GFP) (labels iridophores and glia) transgene displays a uniform and continuous sheet of iridophores. (G) Internal view of 210 dpf asip1K.O. abdominal wall shows a disrupted and discontinuous sheet of iridophores with high number of melanophores (black arrow) and some xanthophores (orange arrow). (H) Abdominal wall of 210 dpf asip1K.O. fish carrying Tg(TDL358:GFP) transgene exhibits a broken sheet of iridophores. Scale bars: 100 μm.
54 Chapter 2
To further characterize the asip1 mutant color phenotype, dorsal and ventral scales of WT and asip1 mutant adult fish were examined. Scales isolated from the belly of asip1 mutants displayed a dorsalized color pattern (i.e. ventral scales becomes nearly as dark colored as dorsal scales due to an increased number of pigment cells) (Fig 2.6A). Contrary to what was observed in WT ventral scales (Fig.6B), asip1 mutant presented various melanophores (Fig. 2.6A, black arrowheads), a significant number of xanthophores (Fig 2.6A, yellow arrowheads) and several silvery patches of iridophores on the ventral scale (Fig. 2.6A, white arrows).
Figure 2.6. Adult asip1K.O. ventral scales displayed a dorsalized color pattern. (A) 210 dpf asip1K.O. ventral scale exhibit a pattern of melanophores (black arrowheads), xanthophores (yellow arrowheads) and also iridophores (white arrowheads). (B) 210 dpf WT ventral scale does not exhibit any chromatophores. Scale bars: 100 μm.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 55
IV. Discussion
In this chapter, we have investigated further the in vivo functional role of asip1 in zebrafish by generating asip1 zebrafish homozygous knockout mutants, using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tool (Hwang et al. 2013). In mice (Mus musculus), a “loss-of- function” mutation of the agouti gene has been reported, which results from an 11 kb insertion and is named the nonagouti allele (a) (Siracusa 1994; Matsunaga et al. 2000). The a/a genotype is ASP negative (Matsunaga et al. 2000) and produces a black phenotype (Bultman et al. 1992) with the loss of dorsal-ventral pigment patterning. Our “loss-of-function” asip1 mutation results in a similar phenotypic effect found in the a/a genotype in mice. The asip1 mutants results from 16 bp deletion accompanied by a 15 bp insertion, which produces an early stop codon and resulting in a truncated protein of only 20 amino acids followed by 54 unrelated residues (Fig. 2.1).
The asip1 mutants exhibit a normal pigmentation pattern during larval development. The first signs of the color phenotype become visible at metamorphic stage (15 dpf, SL 6.3 mm). 15 dpf asip1 mutants develop higher number of melanophores in the ventral region of the head and belly than WT fish and at 30 dpf (SL 7 mm) pigmentation abnormalities are observables in the belly. At 60 dpf (SL 13 mm), when in WT the adult dorsal-ventral pigment pattern are already formed, in the asip1 “loss-of-function” mutants an irregular dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (i.e. ventral skin becomes nearly as dark colored as dorsal skin) is already clearly apparent but they show a normal organization of stripes and the patterning of the fins are not affected. However, we have to highlight a minor alteration defected in all asip1 mutants, the increase of the melanophore number in the ventral dark stripe 2V and the presence of a much more developed 3V dark stripe. The abnormal dark colored ventral skin is due to a marked increase of melanophore and xanthophore number, together with a strong reduction of iridophores in the abdominal wall. Melanophores, xanthophores and few iridophores cover the ventral skin region. This dorsalization of the ventral skin confers a darkened appearance to the belly. Additionally, asip1 mutants showed
56 Chapter 2 a strong reduction of the iridophores number interspersed with melanophores and some xanthophores with a clear disruption of the dense and uniform sheet of iridophores found along the abdominal wall in WT fish. The disruption of this dense and uniform sheet of iridophores in asip1 mutant results in the loss of the light color of the belly.
Based on gain-of-function experiments, the development of iridophores was reported to be largely independent of asip1 levels (Ceinos et al. 2015). It has been also reported that ectopic overexpression of asip1 results in a reduction of the number of melanophores in dorsal regions of flatfish skin, but it does not result in an obvious increase of iridophores (Guillot et al. 2012). Therefore, it was suggested that the apparent increase of iridophores in the asip1 transgenic fish could result from the underlying iridophores becoming more visible in the absence of melanophores (Ceinos et al. 2015). We repeated the experiment with the “loss-of-function” mutation of asip1 gene. In agreement with the previous study, we show that asip1 seems to affect melanophores and xanthophore numbers and their differentiation in adult fish, but we also found that asip1 has a role in iridophores. Therefore, the results of our reporter transgenic line experiments and scales analysis show that there is a contribution of iridophores to asip1 mutant phenotype. Our findings that asip1 mutation produces a disruption of the iridophores sheet in the abdominal wall could reflect a direct asip1 contribution to iridophores development. It is well documented that melanophores migrate and or differentiate within iridophores-free places (Patterson and Parichy 2013) and also melanophores seem to be able to repulse iridophores to some extent (Fadeev et al. 2015), therefore, it is also possible that disruption of the iridophore sheet was the result of the negative short-rang effect due to the increase of melanophores number found in the abdominal wall of asip1K.O.. However, further analyses are necessary to clarify the nature of asip1 mutated phenotype. On the other hand, our examination of asip1 mutant ventral scales revealed an increased number of all pigment cells, raising the possibility that asip1 could affect melanophore, xanthophore and iridophores number and differentiation. In the light of our data, it is required a severe examination of this issue using transgenic markers to specifically clarify the asip1 function in each pigment cell type.
Genetic analysis of dorsal-ventral pigment pattern formation: Asip1 57
In conclusion, we demonstrate that the graded expression of asip1 along the dorsal-ventral axis functions to establish the dorsal-ventral pigmentation pattern in fish, because the lost-of-function mutation of asip1 gene leads to phenotypic elimination of the dorsal-ventral color pattern, but has no effect on the embryonic pigment pattern and the stripe patterning. Furthermore, asip1 seems to affect melanophores, xanthophore and iridophores number and differentiation. Additionally, we must emphasize that, the viability of asip1 mutants presents exciting opportunities for investigating the factors involved in the generation of natural variations in pigment pattern formation in fish.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 57
ABSTRACT
The Melanocortin 1 receptor (MC1R) is the central melanocortin receptor involved in vertebrate pigmentation. Mutations in this gene are related to variations in coat coloration in amniotes. Additionally, in mammals the MC1R is the main receptor for agouti signaling protein (ASIP), which relates it as the critical receptor for dorsal-ventral countershading establishment. In fish, Mc1r is also involved in pigmentation but it has been essentially studied in relation to melanosome dispersion activity and as a putative mutated genetic factor responsible for dark/light phenotypes. However, its role as the crucial component for the Asip1-dependent dorsal-ventral pigmentation is still unclear. Using the clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tools (CRISPR/Cas9) we have generated “loss-of-function” mc1r mutant zebrafish. We demonstrate that Mc1r is the key factor for the establishment of the countershading pigment pattern in fish. Our observations of the mc1rK.O. phenotype include the drastic reduction of the melanophore number in the whole body in all stages of development and a disorganized dorsal-ventral pigment pattern characterized by a darkening of the belly due to an increasing ratio of melanophores and xanthophores. Furthermore, functional rescue of dorsal-ventral pigment disruption-associated asip1 ectopic overexpression, demonstrate that Asip1/Mc1R system modulates dorsal-ventral pigmentation in fish.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 59
I. Introduction
The melanocortin system is a complex neuroendocrine signaling mechanism involved in numerous physiological processes in vertebrates, including pigmentation, steroidogenesis and metabolic control. The melanocortin system exerts its multiple functions via a number of G- protein coupled receptors (GPCRs), the melanocortin receptors (Mcrs). To date, five Mcr types have been identified (Mc1r-Mc5r), each differing in its spatial distribution, ligand affinities and specificity, and therefore with a distinctive physiological role. A high degree of identity and conservation in structural characteristics and pharmacology exists between Mcrs from fish and mammals (Schiöth et al. 2005). In fish, at least five melanocortin receptors also mediate the diverse actions of melanocortins (Mc1r, Mc2r, Mc3r, Mc4r, Mc5r). In mammals, the melanocortin 1 receptor (MC1R) is a key regulator of pigmentation (García-Borrón et al. 2005). Mutations in MC1R are associated with hyper or hypopigmentation (Robbins et al. 1993; Newton et al. 2000; Eizirik et al. 2003; Ling et al. 2003; Nadeau et al. 2006). In mammals, skin darkening is produced through MC1R activation by the α-melanocyte stimulating hormone (α-MSH), which stimulates the eumelanin (dark melanin) production (Buscà and Ballotti 2000); and the light coloration is generated by agouti signaling protein (ASIP), which promotes pheomelanin (light melanin) synthesis by MC1R binding (see Chapter 1) (Miller et al. 1993; Lu et al. 1994; Ollmann et al. 1998). Therefore, the dorsal- ventral pigment pattern is established by ventrally higher Asip expression (Vrieling et al. 1994; Lightner 2009; Manceau et al. 2011). However, in fish, Mc1r shows a similar role but with some variations due to the different mechanism for fish pigment pattern establishment. Most of studies have focused on the melanosome dispersion in melanophores during black background adaptation, wherein α-Msh-Mc1r interaction displays a key role (Fujii and Miyashita 1982; Fujii 2000; Yamanome et al. 2007). Thus, Mc1r has been found to be an essential element for the melanosome dispersion in melanophores during fish color change (Richardson et al. 2008). However, few studies have focused on Mc1r role on countershading pattern in fish.
Cerdá-Reverter et al. (2005) demonstrated Asip1 antagonism of Mc1r signaling in fish. They showed that Asip1 is able to inhibit the NDP-Msh-induced melanosome dispersion through Mc1r binding and its consecutive reduction of intracellular cAMP levels. Additionally, several studies have also reported high expression levels of asip1 gene in the 60 Chapter 3 ventral skin region of goldfish (Carassius auratus) (Cerdá-Reverter et al. 2005), turbot (Scophthalmus maximus) (Guillot et al. 2012), zebrafish (Ceinos et al. 2015) and fugu (Takifugu rubripes) (Kurokawa et al. 2006) among others. They suggest that the Asip1-Mc1r interaction plays an important role on dorsal-ventral pigment pattern establishment in fish. However, Asip1 is able to bind and to antagonize other melanocortin receptors like Mc4r, Mc3r and Mc5r (Cerdá-Reverter et al. 2005; Guillot et al. 2016); and, additionally, other melanocortin receptors have been reported in fish skin like Mc4r in Schizothorax prenanti (Wei et al. 2013), Mc5r in barfin flounder (Verasper moseri) and Japanese flounder (Paralichthys olivaceus) (Kobayashi et al. 2012) or Mc2r in sea bass (Dicentrarchus labrax) (Agulleiro et al. 2013). Therefore, it is also important to consider the possibility that the Mc1r may not be the only receptor involved in dorsal-ventral pigmentation in fish.
In this chapter we have investigated the in vivo functional role of mc1r in zebrafish by generating mc1r zebrafish homozygous knockout mutants, using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tool (Bassett et al. 2013). We demonstrate that mc1r knockout mutant zebrafish displayed a disorganized dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (ie. ventral skin becomes nearly as dark colored as dorsal skin due to an increased ration of melanophores and xanthophores) of skin color. In contrast, we do not see any effects on the core-striped pattern in mc1r mutant fish, which all stripes still visible.
Additionally, functional rescue of dorsal-ventral pigment disruption-associated asip1 overexpression by crossing the asip1-Tg zebrafish with “loss-of-function” mc1r mutant fish demonstrate that Asip1/Mc1R system modulates fish dorsal-ventral pigmentation in fish.
II. Materials and methods
a) Fish
Zebrafish were cultured as previously described (Westerfield 2007) and staged according to Kimmel et al., (1995). Fish of the following genotypes were used: TU strain (Tübingen, Nüsslein-Volhard Lab), Tg(Xla.Eef1a1:Cau.Asip1)iim05 (Ceinos et al. 2015), Tg(TDL358:GFP) (Levesque et al. 2013) and Tg(kita:GalTA4:UAS:mCherry) (Anelli et al. 2009). Ethical approval (Ref. Number: AGL2011-23581) for all studies was obtained from
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 61 the Institutional Animal Care and Use Committee of the IIM-CSIC Institute in accordance with the National Advisory Committee for Laboratory Animal Research Guidelines licensed by the Spanish Authority (RD53/2013) and conformed to European animal directive (2010/63/UE) for the protection of experimental animals.
b) Generation and analysis of mc1r knockout mutants
The mc1r “loss-of-function” mutation was generated using the CRISPR-Cas9 system The CRISPR protocol, originally adapted from Bassett et al. (2013), was kindly provided by Sam Peterson (U Oregon). The target sequence was identified with the ChopChop web tool (Montague et al. 2014). Two long oligonucleotides (Scaffold oligo: 5`- GATCCGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTT TAACTTGCTATTTCTAGCTCTAAAAC-3`, and two different gene-specific oligo GS1: 5’- AATTAATACGACTCACTATAGCTAGTGAGCGTCAGTAATGGTTTTAGAGCTAGAA ATAGC-3’ and GS2: 5’- AATTAATACGACTCACTATAGGGCCAAGATGAACATGTGCAGGGTTTTAGAGCT AGAAATAGC-3’ ) were used to perform a DNA-free PCR to obtain a 125 bp DNA fragment that includes the two previously identified target site sequences (TS1: 5’- GCTAGTGAGCGTCAGTAATG-3’ and TS2: 5’-GGGCCAAGATGAACATGTGCAGG- 3’). The PCR reaction was performed in 20 μL containing 10 μL of 2x Phusion High-Fidelity PCR Master Mix Buffer (New England Biolabs, UK), 1 μL of gene specific oligo (10 μM), 1
μL of gRNA scaffold oligo (10 μM) and H2O nuclease free to 20 μL. PCR conditions were 98ºC for 30 sec, 40 cycles of 98ºC for 10 sec, 60ºC for 10 sec, 72ºC for 15 sec, and a final step of 72ºC for 10 min. The PCR product was purified using DNA Clean&Concentration-5 Kit (Zymo Research, USA) according to the manufacturer’s instructions. Purified PCR product was used as template for in vitro transcription with MEGAscript T7 High yeld transcription Kit (Ambion, USA) according to the manufacturer’s instructions. The gRNA was purified with RNA Clean&Concentrator 5 (Zymo Research, USA) before to use it.
The gRNA was injected in a concentration of 25 ng/µL together with Cas9 mRNA (from the pT3TS-nCas9n plasmid) in a concentration of 50 ng/µL and Phenol red solution (0,1%). Around 2 nL of this mix was microinjected into the cytoplasm of zebrafish eggs at the one- or two-cell stage. Dissection microscope (MZ8, Leica) equipped with a MPPI-2 pressure injector (ASI systems) was used for microinjection. Different mutations were found and three different potential nonfunctional mutations were raised as different knockout lines. The
62 Chapter 3 phenotype in each knockout stable line was exactly the same. For microscope imaging, zebrafish of 5dpf, 15 dpf, 30 dpf and 210 dpf were anesthetized with tricaine methasulfonate (MS-222-, Sigma-Aldrich) scales were isolated from the belly and immersed in PBS on a glass slide. Scales and fish were photographed with a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera.
Transgenic/knockout lines were obtained by setting up crosses between the mc1r knockout mutant lines and the reporter transgenic line Tg(TDL358:GFP); which label iridophores (Levesque et al. 2013), or Tg(kita:GalTA4:UAS:mCherry), which label melanophores (Anelli et al. 2009). The offspring of these crosses were incrossed to obtain homozygous mc1r knockout mutans. Confocal imaging was carried out in a Leica TCS SP5 confocal microscope. 5dpf, 15dpf and 30 dpf transgenic zebrafish were anesthetized and photographed. Adult zebrafish (210dpf) were anesthetized with MS-222 and decapitated to sample a ventral section including the abdominal wall. This section was placed in PBS and photographed.
c) Melanophore and xanthophore counts
The melanophore pattern of mc1r knockout mutant fish was compared with control fish by quantification of melanized melanophores in both groups. Selected region for melanophore counting was different in each stage of development. In early stage (5dpf), we counted melanophores in a 1mm2 area in a dorsal view on the head and the dorsal area, on the horizontal myoseptum and in a ventral view of the head. In early metamorphic stage (15dpf), we counted melanophores in a 1 mm2 area in a dorsal view on the head, on the dorsal area, on the horizontal myoseptum and in a ventral view of the head and the belly. In late metamorphic stage (30dpf), we counted melanophores in a 1 mm2 area in a dorsal view on the head, dorsal area, horizontal myoseptum and in a ventral view on the head and belly. In adult fish (60 and 210 dpf) melanophores within 1 mm2 area were counted in several distinct positions: in a dorsal view on the head (head area) and on the dorsal area (from the edge of the head to edge of the dorsal fin); in a lateral view, on the 2D, 1D, 1V and 2V anterior areas (pectoral to pelvic fin); and finally, in a ventral view of the head and the belly (pectoral to pelvic fin) (see Fig. 2.4A). The dorsal-ventral xanthophore pattern of asip1 knockout mutant fish was compared with control fish by quantification of pigmented xanthophores in postmetamorphic fish (60 and 210 dpf). Selected regions for xanthophore counting were on the dorsal area (from the edge of the head to edge of the dorsal fin), and in a ventral view of
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 63 the belly (pectoral to pelvic fin) (see Fig. 2.4A). To analyze the number of melanophores and xanthophores, six fish per group were anesthetized with tricaine methasulfonate (MS-222-, Sigma-Aldrich) and immersed in 10 mg/ml epinephrine (Sigma) solution for 30 min to contract melanosomes. Fish photographed with a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera. Melanophores were counted using ADOBE PHOTOSHOP CS2 software (Adobe Systems Software Adobe Systems Ibérica SL, Barcelona, Spain) and the ImageJ software (National Institutes of Health, NIH, Maryland, USA).
d) Functional rescue experiment
Transgenic/knockout line were obtained by setting up crosses between the mc1r knockout mutant line and a reporter transgenic line Tg(Xla.Eef1a1:Cau.Asip1)iim05 line; which ectopically overexpresses asip1 and produces a dorsal-ventral disruption of pigment pattern phenotype with dorsal skin as light colored as ventral skin (Ceinos et al. 2015). The offspring was then incrossed to obtain the F2 generation and mc1r locus was sequenced to find homozygous knockout mutation (mc1r K.O.) that carries the dominant asip1 transgene, localized by PCR using specific primers for the Tol2 vector (forward: 5’- GCCCCTCTGCTAACCATGTTC-3’, reverse: 5’- TCATCAATGTATCTTATCATGTCTGG-3’). Adult double transgenic/mutant zebrafish (100dpf) were anesthetized with MS-222 and photographed. Microscope imaging was carried out in a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera.
III. RESULTS
a) Analysis and selection of induced “loss-of-function” mutations.
“Loss-of-function” mutations in the mc1r gene were generated using CRISPR/Cas9 genome engineering tool. We selected two different target sites in the coding exon (286 bp after ATG and 636 bp after ATG) (Fig. 3.1A,B). We found six different mutated alleles (Fig. 3.1B). Alleles M1 and M6 conserved the original open reading frame; therefore, they could generate a functional protein lacking only one or two amino acids and keeping most of the complete amino acid sequence. Alleles M2, M3, M4 and M5 show different reading frame downstream of the target site. The mc1r gene encodes a predicted protein of 323 amino acids. We selected
64 Chapter 3 three mutations with predicted potentially non-functional protein and raised the mutant zebrafish stable line of each one to verify the phenotype: M2 (CRISPR1-mc1r.iim02 zebrafish line), M3 (CRISPR1-mc1r.iim03 zebrafish line) and M4 (CRISPR1-mc1r.iim04 zebrafish line) (Fig. 3.1B). The mc1riim02 allele lacks 7 bp (Del 264-271) and carries 11 bp insertion at position 264 downstream of the predicted ATG. The mc1riim03 allele has lost 2 bp (Del 642-644) and mc1riim04 lacks 2 bp (Del 642-644) and carries 16 bp insertion at position 642 downstream of the predicted ATG (Fig. 3.1B). In those three alleles, the deletions/insertions lead to a frameshift and a new stop codon. The mc1riim02 encodes a 123 amino acids mutant protein, the mc1riim03 encodes a predicted mutant protein with 231 amino acids, and mc1riim04 encodes a mutant protein with 243 amino acids (Fig. 3.1C). These three mutant zebrafish lines resulted in the same specific phenotype.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 65
Figure 3.1. CRISPR/Cas9-induced deletions at mc1r locus. (A) Scheme of the mc1r gene showing the target sites (black arrowheads). Coding exons are represented as white boxes and 5’ UTR and 3’UTR are shown as black boxes. (B) Sequence of induced deletions in mc1r locus. The first and fourth lines show the wild-type target sequences. Black arrowheads label the protospacer-adjacent motif (PAM). The following lines show different induced mutations. Italic lower case letters represent inserted new sequence. The number of deleted (-) and inserted (+) bases are marked in the right side of each sequence. Selected mutations are labeled by white arrowheads. (C) Predicted amino acid sequence encoded for mc1r locus. The first line shows the wild type protein (black), and following lines show the potential protein sequence of each selected mutation. Green boxes show the conserved wild type sequence and blue letters the new mutated sequence. Asterisk represents the stop codon.
66 Chapter 3
b) mc1r function in dorsal-ventral pigment patterning.
In order to determine the Mc1r role in in fish pigmentation, we created a “loss-of- function” mutantion in the mc1r gene using CRISPR/Cas9 system. Fish homozygous for these mutated alleles (mc1rK.O.) exhibited the same color phenotype as adults, with an irregular dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (ie. Ventral skin becomes nearly as dark colored as dorsal skin) (Fig.3.2). Therefore, in the subsequent analyses, we focus just on line CRISPR1-mc1r.iim02.
Mutated mc1r. fish displayed an augmented pigmentation over the entire ventral region (Fig.3.2 A-H), including the head area (Fig.3.2. E,F), and a significant reduction over the dorsal and lateral regions compared to WT siblings (Fig. 3.2 C,D,I,J). Therefore, In mc1rK.O. fish, melanophores and xanthophores increased in all ventral regions (Fig 3.2. G,H) together with a concomitant reduction in all dorsal regions (Fig.3.2 I,J), resulting in an apparent homogenization of the color of the animal with its corresponding loss of countershading.
Mutation of mc1r gene has no significant effects on the typical zebrafish stripe patterning (Fig.3.2 A-D). It has only to be emphasized the significant decrease of melanophores in the dorsal 2D and 1D dark stripes of mc1rK.O fish (Fig.3.2 C,D).
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 67
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68 Chapter 3
c) The development of the mc1rK.O. phenotype.
To determine the time point when the phenotype of the mc1r mutants (mc1rK..O.) becomes first apparent, melanophores were counted at larval (5dpf, SL 3 mm), metamorphic (15 dpf, SL 6.3 mm and 30 dpf, SL 7 mm) and two adult stages (60 dpf, SL 13 mm and 210 dpf, SL 25 mm) (Figures 3.3 and 3.4). It is well known that pigment pattern changes during development can be differentiate by an increase in the melanophores number and variations in their distribution (Kelsh 2004; Parichy et al. 2009) We have quantified the melanophore distribution in mc1rK..O and WT sibling. along the dorsal-ventral axis. A significant general reduction of melanophore number were found in mc1rK..O. at all developmental stages evaluated. At pre-metamporhic (5dpf, data not shown) and metamorphic stages, the melanophore numbers were significantly lower in all areas examined. During early metamorphosis (15dpf), the number of melanophores were 45% lower (P<0.01) in dorsal region, 27% lower (P<0.001) in lateral region and 32% lower (P<0.05) in the belly of mc1rK..O. fish than their WT siblings (Fig. 3.3A). During late metamorphosis (30dpf), the number of melanophores were 31% lower (P<0.05) in dorsal region of the head, 50% lower (P<0.001) in dorsal region, 55% lower (P<0.001) in the lateral region and 36% lower (P<0.01) in the belly of mc1rK..O. fish than their WT siblings (Fig. 3.3B).
Figure 3.3. Dorsal-ventral distribution of melanophores during metamorphosis. (A) Distribution and number of melanophores in WT and mc1rK.O. 15dpf fish. At this stage, mc1rK.O. shows significantly lower number of melanophores in the dorsal region, lateral region and in the belly (B) Distribution and number of melanophores in WT and mc1rK.O. 30 dpf fish. At this stage, mc1rK.O. shows significantly lower number of melanophores in the dorsal view of the head, in the dorsal region, in the lateral region and in the belly. Data are the mean ±SEM, n=6. Asterisks indicate significant differences between WT and mc1rK.O. fish. Scale bar: (A) 200 μm, (B) 500 μm.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 69
At 60 and 210 dpf mc1rK.O. exhibited an obvious dorsal-ventral color defect characterized by an evident proliferation on melanophore and xantophores on the ventral areas together by a significant decrease on melanophore and xantophores number on the dorsal and lateral body sections (Fig. 3.4B). At 60 dpf, the number of melanophores in mc1rK.O. in relation to WT fish was 47% lower (P<0.001) in dorsal region of the head, 53% lower (P<0.001) in dorsal region, 65% lower (P<0.001) in the dark stripe 2D, 35% lower (P<0.001) in the dark stripe 1D and 24% lower (P<0.001) in the dark stripe 1V. However, in the ventral region of the head, the number of melanophores was 64% higher (P<0.001) in mc1rK.O. fish than in WT fish. No differences were found in the dark stripe 2V and the belly (Fig. 3.4C). Additionally, the number of xanthophores were also affected. mc1rK.O. fish shows a 82% higher (P<0.01) number of xanthophores in the belly than WT fish (Fig. 3.4D). A similar phenotype was found in 210 dpf mc1rK.O. fish . Therefore, melanophore number in mc1rK.O. fish compared to WT were 62% lower (P<0.001) in the dorsal region of the head, 30% lower (P<0.01) in the dorsal region, 52% lower (P<0.001) in the dark stripe 2D and 20% lower (P<0.001) in the dark stripe 1D. In contrast, the number of melanophores in the ventral region of mc1rK.O. fish were 97% higher (P<0.001) than their WT sibling, 51% higher (P<0.01) in the dark stripe 3V and 98% higher (P<0.001) in the belly (Fig. 3.4E). Regarding the xantophore counts. The number of xanthophores in the dorsal region of mc1rK.O. fish were 52% lower (P<0.01) than their WT siblings. However, the number of xantophores in the ventral regions of mc1rK.O. fish were 95% higher (P<0.01) than in their WT siblings (Fig. 3.4F).
70 Chapter 3
Figure 3.4. Dorsal-ventral distribution of melanophores and xanthopores in WT and mc1rK.O. fish. (A) Lateral view of zebrafish showing the body regions selected for mealnophore and xanthophore count. (B) Ventral view of the WT and mc1rK.O. 210 dpf zebrafish fish bellies. (C) Distribution and number of melanophores in WT and mc1rK.O. 60dpf fish. At this stage, mc1rK.O. shows significantly lower number of melanophores in dorsal head, dorsal region and black stripes 2D, 1D and 1V. However, mc1rK.O. shows significantly higher number of melanophores in the ventral region of the head. (D) Number of xanthophores in dorsal and ventral skin of WT and mc1rK.O. 60dpf fish. At this stage, mc1rK.O. shows significantly higher number of xanthophores in the belly region. (E) Distribution and number of melanophores in WT and mc1rK.O. 210 dpf fish. At this stage, mc1rK.O. shows significantly lower number of melanophores in dorsal head, dorsal region and black stripes 2D and 1D; conversely, mc1rK.O. shows significantly higher number of melanophroes in black stripe 3V, ventral head and belly. (F) Number of xanthophores in dorsal and ventral skin of WT and mc1rK.O. 210 dpf fish. The mc1rK.O. fish showed highly significant lower number of xanthophores in dorsal regions and significantly higher number of xanthophores in belly region than WT. Data are the mean ±SEM, n=7. Asterisks indicate significant differences between WT and mc1rK.O. fish. Scale bar (A,C,E) 1mm, (B) 100 μm.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 71
In order to obtain a better assessment of the pigment cells in mc1rK.O. fish, we imaged fish carrying the Tg(Kita:Gal4,UAS:mCherry) transgene, which labels melanophores (Anelli et al. 2009). WT fish showed no melanophores in ventral skin (Fig. 3.5A,B), however, mc1rK.O. fish showed an increase in melanophores in the ventral skin region (Fig. 3.5C,D). This is in agreement with the detected increase in the number of melanophores in ventral regions together with the identified decrease in the number of melanophores in dorsal and lateral regions of mc1rK.O. fish at later stages of development (Fig. 3.4). Analysing fish carrying Tg(TDL358:GFP) transgene, which label iridophores and glia with cytosolic GFP (Levesque et al. 2013), we observed that in mc1r mutants, the dense and uniform sheet of iridophores found in the abdominal wall of WT fish (Fig.3.5 E,F) is broken up (Fig. 3.5 E,H). Consequently, mc1r mutants displayed an important reduction of the iridophores number interspersed with a significant increased number of melanophores (Fig. 3.5 G) along the abdominal wall.
72 Chapter 3
Figure 3.5. Ventral pattern disruption. (A) Ventral view of 210 dpf WT belly. (B) Belly of 210 dpf WT fish carrying Tg(Kita:GAL4;UAS:mCherry) transgene shows no melanophores in ventral skin. (C) Ventral view of 210 dpf mc1rK.O. belly. (D) Belly of 210 dpf mc1rK.O. fish carrying Tg(Kita:GAL4;UAS:mCherry) transgene shows higher number of melanophores in ventral skin. (E) Internal view of 210 dpf WT abdominal wall shows a white sheet of iridophores with few internal melanophores (black arrow). (F) Abdominal wall of 210 dpf WT fish carrying Tg(TDL358:GFP) transgene displays a uniform and continuous sheet of iridophores. (G) Internal view of 210 dpf mc1rK.O. abdominal wall shows a disrupted and discontinuous sheet of iridophores with high number of melanophores. (H) Abdominal wall of 210 dpf mc1rK.O. fish carrying Tg(TDL358:GFP) transgene exhibits a broken sheet of iridophores. Scale bars: 100 μm.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 73
To further characterize the mc1r mutant pigment phenotype, dorsal and ventral scales of WT and mc1rK.O. from adult fish were examined. Scales isolated from mc1rK.O. belly show an anomalous pigment pattern. The scale area is occupied by some melanophores (Fig. 3.6A, black arrowheads) and a high number of xanthophores (Fig 3.6A, yellow arrowheads); contrary to ventral WT scale (Fig. 3.6B). No iridophores were found in mc1r mutant ventral scales (Fig.3.6 A)
Figure 3.6. Adul mc1rK.O. fish displays an anomalous pigment pattern in ventral scales. (A) 210 dpf mc1rK.O. ventral scale exhibits a pattern of melanophores (black arrowheads) and xanthophores (yellow arrowheads) (B) 210 dpf WT ventral scale does not exhibit any chromatophores. Scale bars: 100 μm.
e) Asip1 is acting as an inverse agonist of Mc1r
To determine if the Mc1r mediates the Asip1 signaling, we analyzed the combination of the mc1rK.O. with the asip1-Tg zebrafish line. Both mc1rK.O. and asip1-Tg zebrafish lines differ in their phenotype to the WT fish (Fig. 3.7). WT phenotype (Fig. 3.7 A) was described above; it shows a specific striped pattern (Fig. 3.7 B), light belly (Fig. 3.7 C) and darker dorsum (Fig. 3.7 D). The mc1rK.O. phenotype (Fig. 3.7 E) was also reported previously; it shows a barely affected stripped pattern (Fig. 3.7 F), darker belly than WT fish (Fig. 3.7 G) and dark dorsum but with less melanophores than WT fish (Fig. 3.7 H). The asip1-Tg zebrafish line phenotype was previously described by Ceinos et al. (2015) (Fig. 3.7 I); it presents a slightly affected striped pattern (Fig 3.7 J), light belly like the WT fish (Fig. 3.7 K)
74 Chapter 3 and a strongly affected dorsal pattern: asip1-Tg fish present a drastic reduction of dorsal melanophores (Fig. 3.7 L) due to the ectopic overexpression of asip1. Finally, the cross of the mc1rK.O. with the asip1-Tg zebrafish reverts the asip1-Tg phenotype to the mc1rK.O. phenotype (Fig. 3.7 M). The mc1rK.O. +asip1-Tg zebrafish line presents the same barely affected striped pattern than mc1rK.O. (Fig. 3.7 N), the pigmented belly (Fig. 3.7 O) and the dark dorsum with lesser melanophores than WT fish, but undoubtedly pigmented, contrary to asip1-Tg (Fig. 3.7 P). Fish overexpressing Asip1 have paler pigmentation than fish that lack Mc1r. This suggests that Asip1 does more than simply antagonize Mc1r or block basal signalling, as the overexpression phenotype is stronger than absence of receptor. Fish that overexpress Asip1 but also lack Mc1r have a dorsalized phenotype, similar to fish only lacking Mc1r, indicating that Asip1 is acting as an inverse agonist of Mc1r to elicit the opposite response from that produced by α-MSH.
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 75
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76 Chapter 3
Figure 3.7. (cont’d) (N) best developend and the dark stripe 2D worse developed than WT fish, (O) hyperpigmented belly and (P) a lesser pigmented dorsum. Scale bar: (A, E, I, M) 2 mm, (B, C, D, F, G, H, J, K, L, N, O, P) 1 mm.
IV. Discussion
In this chapter, we have investigated the in vivo functional role of Mc1r in zebrafish pigmentation by generating mc1r zebrafish homozygous knockout mutants, using clustered regularly interspaced short palindromic repeats (CRISPR)/clustered regularly interspaced short palindromic repeats-associated protein-9 nuclease (Cas9) genome engineering tool (Bassett et al. 2013). In amniotes, several “loss-of-function” alleles of the extension locus, which encodes MC1R, have been reported in different vertebrate species (Klungland et al. 1995; Takeuchi et al. 1996; Kijas et al. 1998; Schiöth et al. 1999; Newton et al. 2000). They have been characterized by the synthesis of red/yellow pheomelanin instead of dark eumelanin. Initially, the “loss-of-function” allele of the extension locus was found in mice (Mus musculus) and named the nonextension allele (e) (Robbins et al. 1993). The 1 bp deletion resulted in a frameshift and in a premature stop codon after the fourth transmembrane domain of the protein (Robbins et al. 1993). The mutated protein was not able to react to α-MSH binding, which results in yellow coloration (Robbins et al. 1993). Subsequently, similar “loss-of-function” alleles associated with yellow or red coloration have been reported in several other species such as humans (Schiöth et al. 1999), dogs (Newton et al. 2000), pigs (Kijas et al. 1998), cattle (Klungland et al. 1995) and chicken (Takeuchi et al. 1996). Our study reports an interesting and different effect of the mc1r mutation in fish. The mc1rK.O. zebrafish line results from a 7 bp deletion and a 11 bp insertion, which produces a frameshift and an early stop codon after the second transmembrane domain of the protein. The truncated protein have the first 95 amino acids followed by 28 unrelated residues (Fig. 3.1).
The mutant mc1r. pre-metamorphic larvae (15dpf, SL 6.3 mm) exhibit an apparent normal pigment pattern but with a significant reduction in the number of melanophores. Similar to the pre-metamorphic larvae, post-metamorphic larvae (30 dpf, SL 7mm) shows also a significant decrease of melanophores number; this is especially evident in the dorsal region. Juvenile (60 dpf, SL 13 mm) and adult (210 dpf, SL 25 mm) mutant mc1r fish develop an
Genetic analysis of dorsal-ventral pigment pattern formation: Mc1r 77 irregular dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (ie. Ventral skin becomes nearly as dark colored as dorsal skin). The ventral regions of the adult mc1rK.O. fish presents a higher number of melanophores and xanthophores than their WT siblings. However, they show a significant reduction on the number of melanophores and xanthophores on dorsal and lateral skin regions. The abnormal ventral skin darkening is due to a marked increase of melanophores and xanthophore number, together with a strong reduction of iridophores in the abdominal wall. Melanophores and xanthophores cover ventral skin regions . and the strong reduction of the iridophore number in the abdominal wall is interspersed with melanophores. Therefore, the abdominal wall presents a clear disruption of the dense and uniform sheet of iridophores found along the abdominal wall in WT fish. The disruption of this dense and uniform sheet of iridophores in mc1rK.O. fish results in the loss of the light color of the belly and in an apparent dorsal-to- ventral transformation of the ventral skin. Therefore, mc1rK.O. fish phenotype results in an apparent pale homogenization of the color of the animal with its corresponding loss of countershading. Mutation of mc1r gene has no significant effects on the typical zebrafish stripe patterning.
In fish, it has already been shown that Mc1r signaling is involved in the control of pigmentation as nonsense mutations in Mc1r have been reported to result in hypopigmentation in cavefish (Astyanax mexicanus) (Gross et al. 2009)
We show that mc1r “loss-of-function” mutation results in a reduction of melanophore and xanthophore number in the dorsal and lateral fish body regions concomitant with a significant increase in all ventral regions, resulting in an apparent pale homogenization of the color of the animal with its corresponding loss of countershading. We show that mc1r seems to affect melanophore and xanthophore number and their differentiation in larvae, juveniles and adult fish, but our data could also reflect a direct mc1r contribution to iridophores development. As mentioned previously (see Chapter 1), iridophores and melanophores show a negative short- rang effect (Patterson and Parichy 2013; Frohnhöfer et al. 2013; Fadeev et al. 2015); therefore, it is possible that disruption of the iridophore sheet was the result of the negative short-rang effect due to the increase of melanophores number found in the abdominal wall of mc1rK.O. or could reflect a direct mc1r contribution to iridophores development, since mc1r expression has been reported in zebrafish iridophores (Higdon et al. 2013). However, further analyses are needed to elucidate the nature of mc1r mutated phenotype. Therefore, our observations on the significant reduction of dorsal and lateral melanophore and xanthophore
78 Chapter 3 numbers indicate that the color pattern changes could derive from both an inhibition of melanization, directly comparable to mechanism of countershading documented in mammals, but importantly also from a novel mechanism regulating the number of melanophores, and thus the ration of melanophores and other pigment cells.
However, in the light of our data, it is required a severe examination of this issue using transgenic markers to specifically clarify the mc1r function in each pigment cell type.
We have also demonstrated that ectopic asip1 transgenic zebrafish, show rescue of mc1r deficiency to give a superficially mc1r mutant-type phenotype. We show that fish overexpressing ectopically asip1 have paler pigmentation than fish that lack Mc1r. This suggests that asip1 does more than simply antagonize Mc1r and blocks basal signaling, as the ectopic overexpression phenotype is stronger than absence of receptor. In fish, it has previously been shown that asip1 blocks α-Msh-induced melanin dispersion in medaka (Oryzias latipes) scale melanophore by acting as a competitive antagonist of Mc1r (Cerdá- Reverter et al. 2005). Additionally, the presence of other receptors in zebrafish melanophores that alleviate the Mc1r loss and promote melanophore development is a plausible possibility. The presence of other melanocortin receptors in fish skin has already been demonstrated. Thus, mc5r expressions have been reported in barfin and Japanese flounder melanophores (Kobayashi et al. 2012), the mc2r expression (Agulleiro et al. 2013) and mc4r expression (Wei et al. 2013) were detected in sea bass and in Schizothorax prenanti skin respectively. Another explanation for this phenomenon could be that our mc1r mutation results in a reduced protein function and not in an abolished protein.
We also show that fish that ectopically overexpress asip1 but that also lack mc1r have an apparent pale homogenization of the color of the animal with its corresponding loss of countershading phenotype, similar to fish only lacking mc1r, indicating that Asip1 is acting as an inverse agonist of Mc1r to elicit the opposite response from that produced by α-Msh.
It is, therefore, possible that the constitutive activation of a functional Mc1r leads to dorsal melanogenesis, but the ventral expression of asip1 blocks the constitutive activity, thus inhibiting ventral melanogenesis.
Gene expression analysis of asip1 effect in zebrafish skin 83
ABSTRACT
The melanocortin system is a neuroendocrine system that regulates a wide range of endocrine and homeostatic processes. Using a transgenic model that ectopically overexpresses asip1, we examined the extent to which the melanocortin system simultaneously regulates pigmentation and other possible physiological processes in fish skin. We applied skin tissue transcriptomics to identify and profile the genes and gene networks associated with the decreased activity of the central melanocortin system, and to study the different signaling pathways downstream of the melanocortin system. We found a total of 3506 genes were differentially expressed (DEG) by asip1 ectopic overexpression. Females exhibited 2478 DEGs, and males exhibited 1765 DEGs. Only 737 genes were significantly regulated independently of the biological sex. Our data demonstrate that asip1 gene simultaneously regulates pigmentation and other physiological processes in fish skin.
Gene expression analysis of asip1 effect in zebrafish skin 84
Gene expression analysis of asip1 effect in zebrafish skin 85
I. Introduction
The teleost fish melanocortin system consists of five melanocortin receptor subtypes (Mc1r, Mc2r, Mc3r, Mc4r, Mc5r), the endogenous agonists alpha and beta melanocyte stimulating hormone (Msh), adrenocorticotropic hormone (Acth) which are melanocortin peptides derived from the proopiomelanocortin gene and the endogenous antagonists agouti signaling protein (Asip) and the agouti-related protein (Agrp). In fish like mammals, this system is also involved a wide range of biological processes that includes food intake, stress response and pigmentation (Cerdá- Reverter et al. 2011).
One of the interesting features of the melanocortin system is the presence of endogenous receptor antagonists (Lu et al. 1994; Ollmann et al. 1997) that in fish includes Asip1, Asip2, Agrp1 and Agrp2 (suggested alternative nomenclature from Braasch & Postlethwait (2011) is reviewed in the Chapter 1. These antagonists compete with melanocortin agonists by binding to melanocortin receptors. Asip has been reported as a melanocortin antagonist of Mc1r but it is able to antagonize different melanocortin receptors (Mc1r, Mc3r, Mc4r, Mc5ra, Mc5rb) (Cerdá- Reverter et al. 2005; Guillot et al. 2016). In mammals, the lethal yellow mice (Ay) that ubiquitously overexpresses Asp display a yellow coat color phenotype, but also hyperinsulinemia, hyperglycemia and obesity (Klebig et al. 1995). These mice show an ectopic overexpression of Asp as result of a 5’ deletion, which allows the Raly promoter control the Asp coding region (Michaud et al. 1993; Miller et al. 1993). These pleiotropic effects are linked to MC4R binding (Ste Marie et al. 2000). Pleiotropic effects have also been found in asip1-Transgenic zebrafish that ubiquitously overexpresses asip1(Guillot et al. 2016). These transgenic zebrafish ectopically overexpress asip1 under the control of the constitutive promoter (Eef1α) (Ceinos et al. 2015). The asip1-Tg fish display disorganized dorsal-ventral pigment pattern characterized by a ventralization (i.e. lightening due to an descreased ratio of melanophores and xanthophores) of the dorsal pigment pattern. (Ceinos et al. 2015) and also show enhanced growth (Guillot et al. 2016). Additionally, studies on the asip1-Tg zebrafish brain showed that asip1 ectopic overexpression regulates several genes involved in the control of food intake and stress response(Guillot et al. 2016). This model offers an excellent scenario to analyze the phenotype induced by
86 Chapter 4 decreased activity of the central melanocortin system, and to study the different signaling pathways downstream of the melanocortin system.
Using this transgenic model that ectopically overexpresses asip1, we analyzed the transcritomic profiling induced by decreased activity of the central melanocortin system, and characterized the different biological processes and /or signaling pathways downstream of the melanocortin system in zebrafish skin.
II. Materials and methods
a) Fish
Zebrafish were cultured as previously described (Westerfield 2007). Fish of the following genotypes were used: TU strain (Tübingen, Nüsslein-Volhard Lab) and Tg(Xla.Eef1a1:Cau.Asip1)iim04 (asip1-Tg) (Ceinos et al. 2015). Prior to any manipulation, animals were netted and anaesthetized for 1 min in 2-phenoxy-ethanol (0.05%) in the sampling tank. When required, animals were sacrificed by rapid decapitation after anesthesia. Ten asip1-Tg (5 males+5 females) and ten WT (5 males+5 females) zebrafish were placed in 6-liter tanks and acclimated for 30 days to minimize physiological differences. On the sampling day, animals were sacrificed and the skin from dorsal region to ventral region was dissected under a stereo microscope. Samples were placed in Trizol and stored at -80ºc until RNA extraction. Ethical approval (Ref. Number: AGL2013-46448-C3-3-R) for all studies was obtained from the Institutional Animal Care and Use Committee of the IIM-CSIC Institute in accordance with the National Advisory Committee for Laboratory Animal Research Guidelines licensed by the Spanish Authority (RD53/2013) and conformed to European animal directive (2010/63/UE) for the protection of experimental animals
b) Microarray
RNA extraction was carried out with the RNA purification kit (Sigma, St Louis, MO, USA) (following the manufacturer’s protocol) and samples were treated with DNAse (Promega). Total RNA was quantified by spectrometry (NanoDrop ND1000) and quality was confirmed by an RNA 6000 Nano Bioanalyzer (Agilent
Gene expression analysis of asip1 effect in zebrafish skin 87
Technologies) assay. Cyanine 3-CTP-labeled cRNA was produced from 150 ng of total RNA using the Low Input Quick Amp Labelling Kit, One-Color (Agilent p/n 5190-2305) according to the manufacturer’s instructions. The cRNA (1650 ng) was hybridized with the Zeno Custom Gene Expression Microarray (Agilent p/n G2514F- 044138) following One-Color Microarray-Based Gene Expression Analysis protocol Version 6.6 (Agilent p/n G4140-90040). Arrays were scanned in an Agilent Microarray Scanner (Agilent G2565C) according to the manufacturer’s protocol and data extracted using Agilent Feature Extraction Software 11.5.1.1 following the Agilent protocol GE1_1050_Oct12. Six biological replicates (3 females and 3 males) of each group (asip1-Tg and Control) were analyzed in a total of 3 slides containing 4 microarrays in the 4x44 k format. Microarrays included 60-mer probes to the unique transcript from Ensembl and Unigene, which were annotated by functional pathways of Kyoto encyclopedia of genes and genomes (KEGG) with bioinformatics package STARS, which was also used for processing the data. Spot intensities were filtered by the criterion (I/B)>8, where I is the median signal and B the background intensity. The arrays were equalized according to the mean intensities. For each gene, the individual values were divided by the mean value of all samples. The log2-ER (Expression Ratios) were normalized by locally weighted non-linear regression (Lowless). The differentially expressed genes (DEG) were selected by comparison with WT group according to the criteria p < 0.05 after t-test and log2-ER≥ |0.8| (1.7- fold), and analyzed using the newly developed Gene-Hub software (Afanasyev and Krasnov, unpublished data). Hierarchical clustering analysis of the samples followed Ward’s method. Enrichment analysis was performed by comparison of counts per pathway in the lists of DEGs and all microarray features that passed the quality of control. Difference was assessed with Yate’s corrected chi square test (p < 0.05).
b) qRT-PCR validation and expression levels
Up to six differentially expressed genes were validated by real-time qPCR on the same individual RNA samples used for cDNA labeling for microarray analysis. The six selected DEG genes are Asip1-modulated independently of the sex. One microgram of total RNA was used for cDNA synthesis with superscript III reverse transcriptase (Invitrogen) primed with random hexaamers and oligo (dT)12-18 (Invitrogen). Subsequently, one microliter of pure or diluted cDNA was added to
88 Chapter 4
12.5 μL of 2 x SYBR Green/ROX qPCR master mix (2x) (ThermoFisher), 0.5 μL of each primer (10 μM) and H2O nuclease free to 20 μL. Reactions were carried out in triplicate and the thermal protocol was: 95ºC for 10 min and 40 cycles of 95ºC for 15 sec and 60ºC for 1 min. Specific primers were designed with the Primer3 program (Rozen and Skaletsky 2000).: nr1d2a (Fw:5’- GCTGCTCGCCTTCAAAATAC-3’, Rv:5’- TTTCGCTTGTGCGTCATTAG-3’), rell1 (Fw:5’-TGAAGCCAACTCAGATGCAC-3’, Rv:5’- GCGGTGGCAGATGTTTTTAT-3’), lmf2b (Fw:5’-GCCAAATGCAATGACAAATG-3’, Rv:5’- AGCACGAGTACCGCTCATCT-3’), idi1 (Fw:5’-AATACCGCTGGAGAAGCTCA-3’, Rv:5’- CTGAACGCTCGATGCAATAA-3’), agr1 (Fw:5’-GCAGTTGCTGTCAGCTTGAG-3’, Rv:5’- TTTTGGCTGCTTTAGCCAGT-3’) and pgm2 (Fw:5’-GAGTCCTGACGCTGTCCTTC-3’, Rv:5’-AGCATTGGAAAATCCACCAG-3’). Primer and cDNA concentrations were tested for each gene and reaction efficiencies belong 90% were not accepted. The housekeeping gene elongation factor-1α (EF-1α) was used as internal reference to normalize the cDNA template between samples (McCurley and Callard 2008). The melting curves of the products were verified to confirm the specificity of PCR products. RNA extraction, DNAse treatment, cDNA synthesis and calculations were carried out as above. For qPCRs, normalized relative quantities of mRNA expression were calculated with the mathematical method of ΔΔCt (cycle threshold). Additionally the Spearman’s Rho test analysis was carried out to validate the microarray results. Differences were considered statistically significant at p<0.05.
Gene expression analysis of asip1 effect in zebrafish skin 89
III. RESULTS
Figure 4.1. Transcriptome change between wild type fish (C) and asip1-Tg fish (A). Hierarchical clustering of 5549 genes altered by asip1 ectopic overexpression using Ward’s method. Abbreviations: TF, asip1-Tg female; TM, asip1-Tg male; CF, control (WT) female; CM, control (WT) male.
a) Overview of the transcriptomic profile and pathway analysis.
We applied skin tissue transcriptomics to identify and profile the genes and gene networks associated with the decreased activity of the central melanocortin system, and to study the different signaling pathways downstream of the melanocortin system. We analyzed the functional significance of DEGs in zebrafish skin when asip1 is ectopically overexpressed. A total of 3506 genes are significantly regulated by Asip1. Hierarchical clustering analysis grouped WT and asip1 transgenic fish in separated sets. Interestingly, WT and asip1-Tg fish were correctly grouped by sex, but asip1-Tg males and females were grouped closer than WT males and females (Fig. 4.1). It seems that asip1 ectopic overexpression reduces the genetic differences between males and females. As result, the transcriptome of males and females was also compared. A total of 1741 genes are significantly regulated by Asip1 only in females, 1028 genes only in males and 737 genes are significantly regulated by Asip1 independently of the sex (Fig. 4.2).
90 Chapter 4
Figure 4.2. Asip1-derregulated DEG genes in both males and females.
The distribution of fold-changes in expression is similar for males and females in respect of induced and repressed genes (Fig. 4.3). Around 50% of DEGs are induced or repressed in both males and females by asip1 ectopic overexpression, and also the distribution of the degree of induction/repression (1 to 2 Log2-ER, 2 to 5 Log2- ER…etc.) is similar. Despite of females show higher number of DEGs, the proportions of induced and repressed genes are mostly similar.
Gene expression analysis of asip1 effect in zebrafish skin 91
Figure 4.3. The number of differentially expressed genes and the Log2-ER in asip1-Tg males and females. In males, 929 genes were induced and 873 genes were repressed by Asip1. In females, 1190 genes were induced and 1288 genes were repressed by asip1 ectopic overexpression. Numbers indicate the number of genes in each interval of Log2-ER. Percentages indicate the percentage of genes included in each interval of Log2-ER degree in relation to the total of DEGs of that gender.
The enrichment analysis of KEGG pathways was carried out for each gender separately. This analysis results in a significant enrichment in 11 pathways in both males and females, 23 pathways enriched only in males and 17 pathways enriched only in females (Fig. 4.4).
Figure 4.4. Comparison of significantly enriched pathways KEGG in asip1-Tg males and females. Complete lists of KEGG enriched pathways can be obtained in Supplementary table 1.
92 Chapter 4
b) Validation of microarray results.
To validate microarray results, 6 genes were selected among the 43776 genes represented on the microarray and their significant association was analyzed by Spearman’s Rho test (analyses not shown here). Therefore, we selected six Asip1- modulated DEGs independently of the sex; therefore, we do not distinguish between males and females in this analysis. Expression levels obtained by microarrays were calculated as the log ratio sample vs universal reference RNA. On the other hand, expression levels of these genes were assessed by qPCR starting with the same RNA that was used for microarrays. Our results show that qRT-PCR data obtained for all RNA are consistent with microarray results (Fig. 4.6).
Figure 4.6. Differentially expressed genes from microarray analysis analyzed by qPCR. The qPCR results are in good agreement with the microarray findings, all genes show significant changes in the proper direction. Data are the mean ±SEM from six samples after triplicate PCR analysis.
c) Deregulated genes in asip1 transgenic zebrafish skin.
Among the ten most significantly up-regulated genes by asip1 ectopic overexpression in both sexes in zebrafish skin we find genes involved in immune response, apoptosis, cell proliferation, cell migration, carcinogenesis or circadian pathway (Table 4.1). The greatest ten induced gene are: Zgc:162941, an orthologue of mammalian g-type lysozyme; relt-like1 (rell1) a gene involved in immune protection in mammals (Sica et al. 2001); death effector domain-containing 1
Gene expression analysis of asip1 effect in zebrafish skin 93
(dedd1) gene, which is involved in apoptosis in mammals (Valmiki and Ramos 2009); Si:ch211-127b11.1 gene, which is an orthologue of mammalian Plexin b2 (Plxb2) and it is involved in cell migration (Halloran and Wolman 2006); the nuclear receptor subfamily 1 group d member 2a (nr1d2a) a gene involved in circadian rhythm (Amaral and Johnston 2012). Other high induced genes are widely known for its role in carcinogenesis: the eukaryotic translation initiation factor 1A X-linked b (eif1axb), v-myb avian myeloblastosis viral oncogene homolog-like 2b (mybl2b) and leucine rich repeat containing 58b (lrrc58b) (Ewens et al. 2014; Dmitriev et al. 2015; Kunstman et al. 2015; Tao et al. 2015). Furthermore, a membrane anchor protein encoded by phosphatidylinositol glycan anchor biosynthesis class P (pigp) gene is also highly up-regulated in asip1-Tg fish skin and it is involved in embryonic development (Shao et al. 2009).
Log2-ER p-Value p-Value Log2-ER Accession ID Gene Name in in in in Males Females Females Males ENSDARG00000071701 Zgc:162941 6,54 4,37 3,84E-05 7,43E-03 ENSDARG00000018312 relt-like 1 3,65 4,35 3,23E-02 3,04E-02 leucine rich repeat ENSDARG00000063509 4,26 4,07 2,22E-05 1,77E-04 containing 58b v-myb avian ENSDARG00000032264 myeloblastosis viral 2,56 3,74 2,50E-03 9,50E-04 oncogene homolog-like 2b nuclear receptor ENSDARG00000003820 subfamily 1, group d, 3,61 3,37 2,25E-02 6,91E-03 member 2a ENSDARG00000079345 si:ch211-217k17.10 3,91 3,25 9,16E-04 7,68E-03 eukaryotic translation ENSDARG00000057912 initiation factor 1a, x- 3,51 3,13 6,97E-04 1,47E-04 linked, b phosphatidylinositol ENSDARG00000038805 glycan anchor 2,80 3,01 4,30E-04 5,14E-04 biosynthesis, class p ENSDARG00000077090 Si:ch211-127b11.1 2,83 2,92 4,25E-05 7,41E-05 death effector domain- ENSDARG00000002758 3,07 2,85 4,91E-03 1,86E-03 containing 1 Table 4.1. The top 10 most significantly up-regulated genes by asip1 ectopic overexpression in zebrafish skin.
94 Chapter 4
Regarding the most significantly down-regulated genes by asip1 ectopic overexpression in zebrafish skin, the top 10 are involved in neural degeneration, cell differentiation, regeneration and metabolism. The top one is the anterior gradient 1 (agr1) gene, which is involved in tissue regeneration (Ivanova et al. 2015). The other one’s are lipase maturation factor 2b (lmf2b), isopentenyl-diphosphate delta isomerase 1(idi1), integral membrane protein 2Bb (itm2bb) and acetyl-Coa acetyltransferase 2 (acat2) involved in different metabolic pathways (Berthelot et al. 2012; Péterfy 2012; Schröder and Saftig 2016). The (ENSDARG00000026821) gene associated with the mammalian transmembrane protein 106B (Tmem106B) and
Log2-ER p-Value Log2-ER p-Value Accession ID Gene Name in in in Males in Males Females Females ENSDARG00000060682 anterior gradient 1 -4,23 -4,02 3,11E-04 4,42E-04 transmembrane ENSDARG00000074390 -3,69 -3,91 1,24E-02 1,94E-03 protein 176l.4 lipase maturation ENSDARG00000025859 -3,09 -3,41 2,85E-02 2,70E-04 factor 2b ring finger protein ENSDARG00000068851 -3,00 -3,06 9,60E-04 3,60E-04 183 ENSDARG000000268 ENSDARG00000026821 -2,64 -2,89 4,38E-06 7,29E-04 21.1 ENSDARG00000035088 si:ch211-254c8.3 -2,60 -2,76 9,11E-04 2,22E-04 ENSDARG00000076386 ependymin-like 1 -3,09 -2,75 1,75E-06 3,16E-04 integral membrane ENSDARG00000041505 -2,64 -2,67 8,02E-06 4,94E-04 protein 2Bb isopentenyl- ENSDARG00000019976 diphosphate delta -2,61 -2,64 1,21E-02 1,63E-02 isomerase 1 acetyl-coa ENSDARG00000007127 -2,98 -2,42 2,41E-02 4,72E-02 acetyltransferase 2 Table 4.2. The top 10 most significantly down-regulated genes by asip1 ectopic overexpression in zebrafish skin.
involved in transport and neuronal dendricity (Schröder and Saftig 2016). The ring finger protein 183 (rnf183) gene associated to inflammation in mammals (Yu et al. 2016), the ependymin-like 1 (epdl1) associated to neural regeneration (Shashoua
Gene expression analysis of asip1 effect in zebrafish skin 95
1991) and finally, the transmembrane protein 176l.4 (tmem176l.4) and the si:ch211-254c8.3 gene without any known associated function.
d) asip1 target genes relevant to pigmentation
Log2-ER Log2- p-Value in p-Value Accession ID Gene Name in ER in Females in Males Females Males ENSDARG00000040314 psph -0,96 -1,73 9,65E-03 2,75E-02 ENSDARG00000018178 pgm2 -0,96 -2,10 1,91E-02 7,75E-03 ENSDARG00000005423 pgam1a -0,87 -0,95 1,11E-03 2,83E-02 ENSDARG00000005897 dera -1,05 -0,56 9,76E-03 1,44E-01 ENSDARG00000033539 paics -0,93 -0,69 1,67E-02 8,52E-02 ENSDARG00000012399 adssl -0,83 -0,64 5,09E-03 5,87E-03 ENSDARG00000004517 ppat -0,81 -0,43 9,19E-05 4,06E-02 ENSDARG000000 ENSDARG00000017738 -1,06 -2,48 1,83E-01 2,77E-02 17738.1 ENSDARG00000001950 ak1 -0,24 1,68 4,33E-01 1,19E-03 ENSDARG00000039007 eno3 -0,68 0,99 5,62E-02 1,92E-02 ENSDARG00000016706 atic -0,15 -0,98 5,87E-01 2,48E-02 ENSDARG00000028000 pfkpa -0,56 -0,90 1,45E-02 5,30E-03 ENSDARG00000040492 mthfd1b -0,11 -0,83 7,92E-01 3,61E-03 Table 4.3. Iridophore-related DEGs from Higdon et al. (2013) list. The values that fit criteria are underlined.
Analysis of the KEGG pathways showed significantly affected guanine synthesis related pathways in both males and females. Unexpectedly, the classical pathways of melanogenesis were not significantly affected. Several genes involved in purine metabolism and pentose phosphate pathway were also affected by Asip1 gene ectopic overexpression, some of the them were: pgm2, dck, pgd, tktb and fbp2,; and genes involved in glycolysis such as adma, eno3, pgam1a, aldob and aaldocb (see Supplementary table 2). The purine metabolism is differentially affected in both males and females. However, pentose phosphate pathway and glycolysis are modulated only in males (Supplementary table 1).
Previously, a list of enriched genes in iridophores has been reported (Higdon et al. 2013). We have compared that list with our microarray results and found thirteen members of that list affected by asip1 ectopic overexpression (Table 4.3). These
96 Chapter 4 thirteen DEG belong to purine metabolism (pgm2, paics, adssl, ppat, ak1, atic ), pentose phosphate pathway (pgm2, dera, pfkpa), glycolysis pathway (pgm2, pgam1a, eno3, pfkpa), citrate cycle (ENSDARG00000017738.1) and Glycine/serine metabolism (psph, pgam1a). Therefore, they are also involved in guanine synthesis cycle.
Four DEGs from females (hbp1, cyhr1, il10rb and mbd2) were previously reported as genes highly expressed in zebrafish melanophores, iridophres and retinal pigmented ephitelial cells (Higdon et al. 2013), with hbp1, cyhr1, il10rb and mbd2 up-regulated and mbd2 down-regulated (Supplementary table 3). Additionally, genes related to neural crest processes are deregulated in asip1-Tg with sox9a up-regulated in both males and females, coro1ca and sox9b downregulated in males, nrg2a, lbh and lef1 up-regulated in females and finally ovol1 downregulated in females (Supplementary table 4).
e) Pleiotropic effects of Asip1
Genes related to immune system seems to be severely affected in asip1-Tg fish. A number of intermediates in the NOD-like receptor signaling pathway, which is responsible for pathogen detection in innate immune response (Broz and Monack 2013), are deregulated in asip1-Tg fish independently of biological sex (nfkbiab, mapk3, ccl38.1, hsp90b1, lpp). This pathway is the only immune-related pathway affected in both males and females. Males do not show other affected immune- related pathway. However, females show some other affected immune-related pathways such as Fc gamma R-mediated phagocytosis, which is an important mechanism involved in the immune system (Stuart and Ezekowitz 2005) (rps6kb1b, wasb, ncf1, dnm1l). In relation with Fc gamma R-mediated phagocytosis, genes involved in phagosome formation are also significantly deregulated in females (acta1a, tubb5, calr, tuba81). Other two adaptive immune-related pathways are deregulated in females: B cell receptor signaling pathway, which is essential for the survival and proliferation of B-cells and for the mediation of the antigen processing (Niiro and Clark 2002) (chp1, nfkbiab, chp2, vav2, mapk3); and T cell receptor signaling pathway, which is necessary for T cell proliferation and cytokine production by T cells (Huang and Wange 2004) (chp1, nfkbiab, chp2, lcp2a, mapk14b). Females also show several genes of the chemokine signaling pathway
Gene expression analysis of asip1 effect in zebrafish skin 97 significantly deregulated. This pathway is essential to drive leukocytes to the site of inflammation (Wong and Fish 2003) (nfkbiab, ccl38.1, rasgrp4, cxcr4b, ccr9a). And finally, several gene related to leukocyte transendothelial migration pathway (cxcr4b, myl9a, actn2b, acta1a, actn1), which is essential to immune response (van Buul and Hordijk 2004), are also deregulated in females (Supplementary table 1 and 5).
Several specific genes that encode proteins implicated in endocrine and metabolic pathways are also deregulated in Asip1-Tg fish.
Insulin signaling pathways are significantly affected in females (socs3a, bada, rps6kb1b, eif4e1c, socs1a, mknk2a, irs2a, mknk2b, irs1). In male only specific genes implicated in type I diabetes mellitus pathway (mfaa, mapk3) are significantly affected. Several genes implicated in PPAR signaling pathway (fads2, scd, fabp7a, fabp7b, acaa1, slc27a2, cyp27a1.2, cyp27a1.4, me3), an important group of nuclear hormone receptors activated by fatty acids (Feige et al. 2006), are also significantly affected in asip1-Tg males,(Supplementary table 1 and 6).
The decreased activity of the central melanocortin system also affects several genes implicated in the cell cycle signaling pathways (mcm5, orc6, mcm4, cdkn2c) in males, RNA polymerase signaling pathway (polr2h, ube2r2, klf13) in both, DNA replication and mismatch repair signaling (acta1b, eif2s1a, actc1) and focal adhesion signaling pathways (acta1b, actn3a, myl9a, mylpfb) in males. In females, several gene implicated in apoptosis (bada, casp611, chp1, nfkbiab, cycsb), p53 signaling (cycsb, ccng2, sesn1), and cancer signaling (zbtb16a, rps6kb1b, pim3, bada, foxo3b, tp53, stk3) pathways are significantly affected (Supplementary table 1 and 7).
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IV. Discussion
In this chapter, we examined, using a transgenic model that ectopically overexpresses asip1, the extent to which the melanocortin system simultaneously regulates pigmentation and other physiological processes in fish skin. Specifically, we applied skin tissue transcriptomics to identify and profile the genes and gene networks associated with the decreased activity of the central melanocortin system, and to study the different signaling pathways downstream of the melanocortin system. A formal proof of the pleiotropic nature of Asip1 function in vivo is whether ubiquitous ectopic overexpression of Asip1 protein could results in a simultaneous deregulation of different physiological processes. In fact, several studies have made transgenic constructs that place the agouti gene under the transcriptional control of ubiquitous promoters (Robbins et al. 1993; Newton et al. 2000; Bonilla et al. 2005; Guillot et al. 2016). It has previously been shown that asip1 overexpression resulted in both increased linear growth and body weight in zebrafish (Guillot et al. 2016). Furthermore, brain transcriptomic studies revealed expression differences in several central circuits involved in the control of food intake as well as modifications in the central responsiveness to neurotransmitter, neuropeptides and peripheral hormones known to be involved in control of the energy balance (Guillot et al. 2016).
Our skin transcriptomic studies reveal a total of 3506 genes differentially expressed (DEG) by asip1 ectopic overexpression. Females exhibits 2478 DEGs, and males exhibits 1765 DEGs. Only 737 genes are significantly regulated independently of the biological sex.
Our data in transgenic models demonstrate that asip1 overexpression results in a significant sex dependent differential regulation. Only the 21% of the Asip1- modulated DEGs are sex-independent regulated. The 29% of DEGs are Asip1- affected only in males and the 50% of DEGs are Asip1-modulated only in females. The total deregulated genes are significantly higher in asip1-Tg female fish. However, our results also show that asip1-Tg males and females are transcriptionally more similar than their respective WT. These results are in agreement with a previous asip1-Tg brain transcriptomic study in which reduced differences among transgenic males and females transcriptomes were found compared to their
Gene expression analysis of asip1 effect in zebrafish skin 99 respective WT (Guillot et al. 2016). Interestingly, results shown in brain by Guillot et al. (2016) (males modulate more genes than females) was contradictory to what we have actually found in skin (females modulate more genes than males). This data suggests that asip1 overexpression effect is tissue-specific. Interestingly, the percentage of genes up-regulated and down-regulated is similar in females and males. Finally, the KEGG enriched analysis shows that only the 22% of the significantly affected pathways are modified independently of the biological sex. However, the pathways altered by Asip1 depending of the sex affect similar gene networks (metabolism, cell cycle, immune system, etc.).
Our skin transcriptomic study shows that the guanine synthesis pathway is modulated by asip1 overexpression. The purine metabolism, pentose phosphate and glycolysis metabolism pathways are significantly modulated by asip1 ectopic overexpression in zebrafish skin. Among the most affected genes of these pathways, there are several enzymes comprising translation initiation factor activity (eif2s1a), catalytic activity (pgam1a, adma, aldob, aldocb, aldoaa, aldoab, pgd, tktb, dhcr24), transferase activity (aprt, pgm2), polymerase activity (polr2h), mediated signal transduction activity (arf5), phosphatase activity (fbp2), and kinase activity (dck) in addition to others. Interestingly, Higdon et al. (2013) suggested that ADP Ribosylation Factors likely contribute to the formation of the membranous platelets containing the reflective guanine crystals. Arf proteins display a role on membrane traffic and organelle structure (Donaldson and Honda 2005). Our data show that one of the most affected genes from the guanine synthesis pathway in both sexes is the arf5 gene, a member of the ADP ribosylation Factor family. Higdon et al. (2013) reported a list of genes that are highly expressed in zebrafish pigment cells. We have compared that list with our DEGs data and found four members of that list significantly deregulated by asip1 ectopic overexpression in female only. The hbp1, which is a transcription factor involved in cell cycle progression in mammals (Li et al. 2010; Watanabe et al. 2015); the cyhr1, with an inhibitor role on apoptosis in mammals (Desaki et al. 2015); the il10rb, a receptor involved in immune response signaling (Grayfer et al. 2011); and mbd2, a regulator of the transcription by methyl- CpG binding (Hendrich and Tweedie 2003). These genes were found by Higdon et al. (2013) as candidate genes to identify pigment cell identity because they were especially induced in pigment cells.
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Several genes related with neural crest cells are also found to be deregulated by asip1 ectopic overexpression. The most significantly up-regulated gene is a type of plexines involved in cell migration (Halloran and Wolman 2006) including neural crest migration (Fiore and Püschel 2003). Sox9 is a transcription factor that plays a pivotal role in neural crest development and male sexual development (Weider and Wegner 2016); and both sox9a and sox9b are deregulated by asip1 ectopic overexpression in zebrafish. Other genes related to neural crest migration and deregulated in asip1-tg fish are coro1ca, nrg2a, lbh, ovol1 and lef1 (Honjo et al. 2008; Piloto et al. 2010; Degenhardt et al. 2011; Powder et al. 2014; Williamson et al. 2014). Interestingly, one of the top ten up-regulated genes, the pigp, is essential for embryonic development but also regulates the Wnt signaling (Shao et al. 2009). Wnt signaling has been related to pigment pattern establishment (Rawls et al. 2001), in addition to other functions. These findings could indicate that the pigment cell development is altered to some extent at least.
We expected that the melanogenesis pathway was modulated by asip1 ectopic overexpression because it was previously reported that several melanogenic genes were down-regulated by Asip1 in asip1-Tg fish (Guillot et al. 2012; Ceinos et al. 2015). Unexpectedly, we found that melanogenesis was not affected in males nor females in our skin transcriptomic study. An explanation for this could be that the skin was sampled from the dorsal region to the ventral region, including stripes. Therefore, it is possible that the asip1 repressing effect found in the dorsal skin area in asip1-Tg fish (Ceinos et al. 2015), had been masked for the presence of adjacent regions in the skin sample analysed. The properties and performance characteristics of our microarray system could be another explanation for these unexpected results. Further transcritopic analyses of asip1-Tg fish are needed to fully explain the structure and complexity of the gene network regulating expression of pigment cell genes.
Our skin transcriptomic study shows a deregulated expression of numerous immune related genes. Among these genes are: transcription factors (nfkbiab), protein kinases (mapk3, rps6kb1b, mapk14b), guanine-nucleotide exchange activity proteins (vav2, rasgrp4), chemokines and chemokine receptors (ccl39.1, cxcr4b, ccr9a), GTPases (dnm1l, tubb5, tuba81) and chaperones (hsp90b1, calr) in addition to others. Interestingly, the top up-regulated gene in both biological sexes is a type of
Gene expression analysis of asip1 effect in zebrafish skin 101 g-lysozyme. Lysozymes have shown antibacterial properties in fish (Saurabh and Sahoo 2008) and usually they are induced by several bacterial infections (Yin et al. 2003; Sha et al. 2012). Other significantly up-regulated gene is the rell1 and the Si:ch211-127b11.1 gene. Rell1 gene function is related to T-cells proliferation in mammals (Sica et al. 2001), and Si:ch211-127b11.1 is an orthologue of mammalian Plexin b2 (Plxb2) involved in cell migration (Halloran and Wolman 2006) and in several other functions of the immune system (T-cells, dendritic cells and macrophages) (Potiron et al. 2007; Roney et al. 2013).
Functional enrichment analysis reveal that biological process terms related to innate and adaptive immune response are enriched. Regarding the innate response, the NOD-like receptor signaling pathway is significantly deregulated, which is involved in the recognition of pathogens and in the initiation of the inflammatory response (Broz and Monack 2013). Concerning the adaptive response, several genes related B cell receptor and T cell receptor signaling pathways are significantly deregulated. Additionally, genes involved in phagocytosis, chemokine signaling and transendothelial migration of leukocytes signaling are also deregulated by asip1 ectopic overexpression.
Several studies have shown important roles of the melanocortin system in inflammation. The anti-inflammatory and immunomodulatory properties revealed the anti-inflammatory potential of α-MSH and its cognate receptors (Cooper et al. 2005; Maaser et al. 2006; Ahmed et al. 2013). Our skin transcriptomic study also supports the role of melanocortin system in inflammation in fish. Both inflammatory response and lymphocyte-related genes are affected. Interestingly, the anti-inflammatory effect of α-MSH is commonly produced by the action of NFκB cascade (Ahmed et al. 2013), and we have found in our skin transcriptomic study some NFκB-related genes significantly deregulated such as traf4a, traf4b, or also the nfkbiab (Supplementary Table 8). Another gene involved in inflammation, the nr1d2a, it is also one of the top 10 up-regulated genes by asip1 ectopic overexpression. nr1d2a is a clock gene involved in the circadian cycle (Amaral and Johnston 2012) and inflammatory response (Morioka et al. 2016). rnf183 gene, identified in mammals as an important factor on inflammation regulation (Yu et al. 2016) was significantly deregulated in asip1-Tg fish.
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Our skin transcriptomic study showed that a common term/pathway, which is noticeably affected by asip1 overexpression especially in female but also in male skin, is the inflammatory response. It is well-known that the immunological defenses present in skin mucosa are a very important part of the fish immune system, serving as an anatomical and physiological barrier against external hazards. Our skin transcriptomic study also reveals an important effect of Asip1 on the energy metabolism, as previously showed in zebrafish brain (Guillot et al. 2016). The most deregulated genes related to energy metabolism pathways are several protein kinases and protein kinase inhibitors genes (socs3a, socs1a, rps6kb1b, mknk2a, mknk2b, mapk3), oxidoreductases genes (fads2, scd, cyp27a1.2, cyp27a1.4, me3), fatty acid transpoters and fatty acid binding proteins (fabp7a, fabp7b, slc27a2), insulin receptor substrate proteins (irs2a, irs1), translation initiation and transcription factors (eif4e1c, mafia) in addition to others.
Several insulin signaling pathways-related genes are also affected by asip1 ectopic overexpression. This effect is not surprising because it was already reported in mice. The phenotype of the lethal yellow mice shows yellow coloration and obesity, apart from hyperinsulemia, hyperglycemia and type II diabetes mellitus (Klebig et al. 1995). Our study also shows that one of the top down-regulated genes independently of the gender is itm2bb, which is a gene related to the promoteion of the Insulin Degrading Enzyme (IDE) secretion (Schröder and Saftig 2016). Additionally, several energy metabolism related pathways (glycolysis/gluconeogenesis, glyoxylate and dicarboxylate metabolism, pyruvate metabolism and pentose phosphate metabolism) are significantly deregulated in asip1 transgenic male fish, and amino sugar and nucleotide sugar metabolism, and starch and sucrose metabolism are modified in both males and females (Supplementary Table 1). The key role of the central melanocortin system in the control of food intake and energy metabolism is well known. In fish, the involvement of melanocortin systems in fish feeding behavior and growth has been shown (Guillot et al. 2016). They demonstrated that asip1 overexpression resulted in increased growth but not obesity. They hypothesized that the increased growth observed in asip1-Tg zebrafish could be mediated by Asip1 binding to central Mc4r. (Guillot et al. 2016). Interestingly, asip1 overexpression induced downregulation of lipase maturation factors (lmf2b) which have a role on maturation and transport of lipoprotein lipases and its deficiency produces
Gene expression analysis of asip1 effect in zebrafish skin 103 hypertriglyceridemia (Péterfy 2012). The PPAR are a type of receptors activated by lipids (Feige et al. 2006). Accordingly, KEGG pathway analysis revealed an alteration of the peroxisome proliferator-activated receptor (PPAR) signaling pathway in males. The role of ASP in lipid metabolism has previously been reported in lethal yellow mice (Klebig et al. 1995) and in transgenic mice carrying the aP2- promoter driving agouti expression on adipose tissue (Mynatt and Stephens 2003). These transgenic mice show a substantial effect over PPAR signaling pathway (Mynatt and Stephens 2003). Interestingly, the transgenic mice carrying the aP2- pomoter driving Asip show an increasing of fat mass (Mynatt and Stephens 2003), contrary to our asip1-Tg fish which only show an increase of the growth rate, but not of the fat mass (Guillot et al. 2016). Our skin transcriptomic study also reveals some pathways of the lipid metabolism affected such as biosynthesis of unsaturated fatty acids and steroid biosynthesis (Supplementary Table 1). Moreover, it was shown in mammals that ASIP induces lipogenesis via a Ca2+-dependent mechanism (Xue et al. 1998); and, interestingly, our data also show the calcium signaling pathway affected in males (Supplementary Table 1).
Other genes and gene networks associated with the decreased activity of the central melanocortin are related to cell cycle, cell proliferation and cell survival. The top up-regulated genes in these pathways are protein kinases and kinase inhibitors (rps6kb1b, pim3, stk3, ccng2, cdkn2c), transcription and translation factors (foxo3b, klf13, nfkbiab, zbtb16a, eif2s1a), apoptotic proteins and peptidases (bada, casp611), members of the minichromosome maintenance protein complex (mcm5, mcm4) and actin binding proteins (myl9a, milpfb). Several of the top up-regulated genes independently of the biological sex are related to cell cycle such as dedd1, which in mammals induces apoptosis by activation of caspases (Valmiki and Ramos 2009); rell1, which is also involved in cell death (Cusick et al. 2012); or the mybl2 gene; which is an oncogene associated to breast cancer (Tao et al. 2015), but also to cell migration, cell cycle and cell differentiation (Maruyama et al. 2014; Tao et al. 2014, 2015). However, the top down-regulated gene is the agr1. The agr1 gene is involved in regeneration and cell differentiation in fish (Ivanova et al. 2015).
Cell adhesion is essential for cell survival. Failed adhesion results in cell death (Stupack and Cheresh 2002). Our transcriptomic results show that Asip1 alters the focal adhesion. This is in agreement to other investigations that show the role of α-
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MSH-MC1R system in cell adhesion in mammals (Scott et al. 1997). Furthermore, DNA replication, RNA polymerase and the cell cycle are also affected in our study. This could be related to cell growth or cell proliferation. It has been shown that α- MSH, ACTH and MC3R are able to stimulate cellular proliferation in mammals (Zohar and Salomon 1992; Chai et al. 2007). Interestingly, it has also been shown that ASIP can also to inhibit the differentiation of mammalian immortal melanoma cells; the opposite role was reported by α-MSH (Sviderskaya et al. 2001).
The melanocortin system has been related to apoptosis in mammals. α-MSH, MC1R and MC4R inhibit or decrease apoptosis in different cell types such as neurons, astrocytes or melanocytes (Kadekaro et al. 2005; Chai et al. 2006; Caruso et al. 2007). Additionally, the p53 pathway, which mediates apoptosis, cell cycle arrest and cell senescence in response to different stresses (Levine et al. 2006), is also under the control of asip1 ectopic expression in zebrafish skin. In mammals, the p53 is a target gene of the downstream MC1R signaling pathway (Kadekaro et al. 2012). The α-MSH binds to MC1R and activates p53 pathway, which reduces oxidative stress in melanocytes (Kadekaro et al. 2012). Our transcriptomic results shows that the mismatch repair of DNA pathway is also regulated by asip1 ectopic overexpression.
Our skin transcriptomic studies revealed Asip1-induced deregulated expression of some neuropeptides involved in the control of fish reproduction, including the gonadotropin releasing hormone system (Senthilkumaran et al. 1999) (Supplementary Table 1). The interaction between melanocortin system and the GnRH pathway has been already reported in mammals (Chai et al. 2006; Zandi et al. 2014) and also in fish (Jiang et al. 2017). Angiogenesis, the development of new vasculature from preexisting blood vessels, is also affected by asip1 ectopic overexpression in zebrafish skin. The genes related to VEGF signaling pathway were significantly deregulated in asip1-tg female fish. In mammals, in vitro experiments showed that α-MSH strongly affects different angiogenic processes (Weng et al. 2014). In addition, several genes involved in cardiac muscle contraction and vascular smooth muscle contraction are significantly deregulated in males. In mammals, in vitro experiments have shown that the hypothalamus peptide nesfatin-1 produces arterial contractility and it might be possible that this effect is mediated by MC3R (Yamawaki et al. 2012). Additionally,
Gene expression analysis of asip1 effect in zebrafish skin 105 several genes typically involved in dilated cardiomyopathy and hypertrophic cardiomyopathy (HCM) are also significantly affected in asip1-Tg male zebrafish (Supplementary Table 1). Other affected pathways are related to intracellular and environmental processing information, for example protein processing in endoplasmic reticulum, protein export or phosphatidylinositol signaling system. The intracellular communication has been also affected via Gap junction genes in both males and females (Supplementary Table 1).
Taking together, our skin transcriptomic study shown that most of the common term/pathway that are noticeably affected by asip1 overexpression in fish are also affected in similar overexpression approaches in mammals (Klebig et al. 1995). Our results also confirm the pleiotropic nature of the agouti gene. This reveals the ability of asip1 gene to simultaneously regulate pigmentation and other physiological processes in fish skin.
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 113
ABSTRACT
Dorsal-ventral pigment patterning, characterized by a light ventrum and a dark dorsum, is one of the most widespread chromatic adaptations in vertebrate body coloration. In mammals, this countershading depends on differential expression of agouti-signaling protein (ASIP), which drives a switch of synthesis of one type of melanin to another within melanocytes. Teleost fish share countershading, but the pattern results from a differential distribution of multiple types of chromatophores, with black-brown melanophores most abundant in the dorsal body and reflective iridophores most abundant in the ventral body. We previously showed that Asip1 (a fish ortholog of mammalian ASIP) plays a role in patterning melanophores. This observation leads to the surprising hypothesis that agouti may control an evolutionarily conserved pigment pattern by regulating different mechanisms in mammals and fish. To test this hypothesis, we compared two ray-finned fishes: the teleost zebrafish and the non-teleost spotted gar (Lepisosteus oculatus). By examining the endogenous pattern of asip1 expression in gar, we demonstrate a dorsal-ventral graded distribution of asip1 expression that is highest ventrally, similar to teleosts. Additionally, is the first reported experiments to generate zebrafish transgenic lines carrying a bacterial artificial chromosome (BAC) from spotted gar, we show that both transgenic zebrafish lines replicate the endogenous asip1 expression pattern in adult zebrafish, showing that BAC transgenes from both species contain all of the regulatory elements required for regular asip1 expression within adult ray-finned fishes. These experiments provide evidence that the mechanism leading to an important pigment pattern was likely in place before the origin of teleosts.
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 115
I. Introduction
The origin of vertebrate pigment patterns – the differential pigmentation of skin, scales, hair, or feathers - is a classic problem in developmental biology. The extension and agouti genes are both key loci involved in the regulation of body pigmentation. The agouti locus encodes the Agouti-signaling protein (ASIP), a ligand protein that competes with α-Melanocyte stimulating hormone (α-MSH) in binding to the Melanocortin 1 receptor (MC1R), which, in turn, is encoded by the extension locus (see Chapter 1). This competitive antagonism results, in amniotes, in melanocytes switching between the production of eumelanin (black/brown pigment) and pheomelanin (red/yellow pigment) (Lu et al. 1994). In amniotes, high Asip expression in the ventral region inhibits Mc1r signaling, which results in a pale belly coloration (phaeomelanin>>eumelanin) in contrast to the darker dorsal region (eumelanin>>phaeomelanin); the dorsal-ventral pigment pattern gradient is adaptive and widely conserved in the animal kingdom, including fish (Vrieling et al. 1994; Manceau et al. 2011). Curiously, this conserved pattern of dorsal-ventral pigment polarity occurs by a different mechanism in fish and mammals: in fish, the pattern appears by the distribution of different pigment cell types that produce chemically distinct pigments rather than by the switching of melanin types within the same kind of pigment cell as in mammals (Kelsh 2004). In zebrafish (Danio rerio), a teleost fish, the pigment pattern depends on the patterned distribution of three different pigment cell types: dark eumelanin-producing melanophores (homologous to the mammalian melanocytes, but without the ability to synthesize the lighter pheomelanin (Kottler et al. 2015), yellow to orange xanthophores, which produce pteridines, and silver shiny iridophores, which produce reflective plates of guanine crystals. These different pigment cell types derive from embryonic neural crest (Le Douarin and Kalcheim 1999; Dupin et al. 2000).
In adult zebrafish, melanophores are distributed in four horizontal stripes alternating with three white interstripes that are rich in iridophores and xanthophores; the ventrum is pale due to an abundance of iridophores and the absence of melanophores (Kelsh et al. 2009). Recent studies have characterized asip1 genes in several teleost fish species, showing that its high ventral expression is a conserved
116 Chapter 5 feature (Cerdá-Reverter et al. 2005; Klovins and Schiöth 2005; Kurokawa et al. 2006; Guillot et al. 2012; Ceinos et al. 2015). Our previous experiments demonstrated a gradient of asip1 expression with lowest expression levels in the dorsal region (Ceinos et al. 2015). Disruption of this expression gradient by gain-of- function approaches using ubiquitous promoters (Tol2-Eef1α1/asip1) interrupted pigment patterns by inhibiting dorsal melanogenesis and melanophore differentiation. Asip1 overexpression, however, had a minor impact on the pattern of stripes revealing the existence of two different pigment patterns in zebrafish: a dorsal-ventral pattern and stripe pattern, which are regulated by different mechanisms (Ceinos et al. 2015). Consequently, these results suggest that Asip- dependent countershading and its molecular machinery are conserved between mammals and teleost fish, but, rather surprisingly, the cellular basis for pigment pattern formation is different in the two groups.
We ask here the question: is the patterning mechanism found in teleosts specific to that taxon, perhaps arising as a feature of the teleost genome duplication (TGD) (Amores et al. 1998; Postlethwait et al. 1999; Taylor et al. 2003; Jaillon et al. 2004), or alternatively, is it a more ancient feature shared by other groups of ray-finned fishes? To address this question, we investigated the regulation of asip1 in spotted gar, which represents a ray-finned lineage that diverged from teleosts before the TGD (Amores et al. 2011).
To gain insight into the evolutionary conservation of these processes in ray- finned fish, we compare here the promoter regions of zebrafish and spotted gar (Lepisosteus oculatus). Gars diverged from the teleost lineage around 350 million years ago (Hoegg et al. 2004; Vandepoele et al. 2004; Hurley et al. 2007) just before the teleost specific genome duplication in a teleost ancestor (TGD), and gar is therefore considered a useful outgroup to the teleost lineage (Amores et al. 2011; Braasch et al. 2016). The use of a lineage closely related to teleosts, but diverging before the TSGD, facilitates connectivity, the sharing of biological understanding among ray-finned fish and the translation of knowledge to tetrapods including mammalian species (Braasch et al. 2016). Spotted gar can be reared in the laboratory and is thereby amenable for the study of spatial-temporal patterns of gene expression during development and the functional testing of hypotheses about the origin of gene activities in divergent fish lineages (Amores et al. 2011; Braasch et al. 2015, 2016).
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 117
Exploiting these advantages, we studied the functional evolution of the asip1 locus in ray-finned fish. We describe here the generation of bacterial artificial chromosome (BAC) transgenic (Tg) zebrafish lines, which harbor zebrafish or spotted gar BACs carrying the asip1 gene modified to drive enhanced green fluorescent protein (eGFP) expression. The resulting transgenic animals carrying asip1 BAC transgenes from either species showed eGFP expression in a pattern that mirrors that of endogenous asip1 gene expression in adult zebrafish and, also of spotted gar, and illustrate that the transgenic zebrafish is advantageous for identifying the conserved regulatory elements required for normal asip1 expression within adult fish. Thereby, our study is the first to generate zebrafish transgenic lines carrying a BAC clone from spotted gar. Additionally, we show a dorsal-ventral gradient of asip1 expression in the skin of adult spotted gar with lower levels in the dorsum and higher the belly, similar to the expression gradient in zebrafish, and other teleosts.
II. Materials and methods
a) Fish
Zebrafish of TU strain (Tübingen, Nüsslein-Volhard Lab) were cultured as previously described (Westerfield 2007) and staged according to Kimmel et al. (1995). Ethical approval (Ref. Number: AGL2011-23581) for all studies was obtained from the Institutional Animal Care and Use Committee of the IIM-CSIC Institute in accordance with the National Advisory Committee for Laboratory Animal Research Guidelines licensed by the Spanish Authority (RD53/2013) and conformed to European animal directive (2010/63/UE) for the protection of experimental animals. Spotted gar (Lepisosteus oculatus) were reared and collected at the Institute of Neuroscience (University of Oregon). This work was approved by the University of Oregon institutional Animal Care and Use Committee (Animal Welfare Assurance Number A-3009-01, IACUC protocol 12-02RA).
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b) Comparative genomic analysis and sequence conservation
To analyze the conserved sequences between zebrafish and spotted gar and identify any conserved non-coding elements (CNEs), repeat-masked sequences were aligned using MLAGAN alignment software in m-VISTA (Mayor et al. 2000; Brudno et al. 2003). Conserved sequences were predicted under 65% identity and 50bp window. Sequences analyzed were zebrafish masked sequence (http://www.ensembl.org/Danio_rerio/Info/Index) as reference genome and spotted gar masked sequence (http://www.ensembl.org/Lepisosteus_oculatus/Info/Index). The protein alignment was performed using the Expresso alignment with the T- coffee software. The predicted protein sequence was obtained from the Ensembl zebrafish genome database (ENSDART00000113083) and the GenBank database for spotted gar (XM_015365014.1). We used the web-based Genomicus synteny browser [http://www.dyogen.ens.fr/lcgi-bin/search.pl] and Ensembl genome browsers to assess the synteny between regions from zebrafish BAC clone (16N21) from DanioKey Library and spotted gar BAC clone (VMRC-56-161-M17) from BAC library VMRC-56 (Postlethwait Lab, Institute of Neuroscience, University of Oregon). Only genes annotated in these assembly versions were considered for our synteny comparison.
c) RNA isolation, RT-PCR and qRT-PCR
Animals were anesthetized with MS-222 and decapitated. Total RNA was extracted from adult fish (210 dpf zebrafish and 270 dpf spotted gar) from eye, gonads, liver, brain, gill, intestine, muscle, heart and dorsal and ventral skin. Furthermore, dorsal, stripes (1D, 1V, 2D and 2V in zebrafish; 2D, Medium and 2V in spotted gar) and belly skin samples of eight fish of both species were excised with a scalpel for qRT-PCR (see Fig.5.4 A, C). Tissue samples were frozen at -80ºC in RNAlater (Ambion, USA) until analysis. Total RNA was extracted with Direct-zol RNA miniprep kit (Zymo research) according to the manufacturer’s instructions. The Maxima First Strand cDNA Synthesis Kit (Fermentas) was used to synthetize cDNA from 100 ng of total RNA. Tissue expression patterns were analyzed by RT-PCR. A pair of specific primers was designed for spotted gar asip1 and spotted gar β-actin as internal control from the mRNA sequence published in Ensembl database (ENSLOCT00000008713 and ENSLOCT00000005654 respectively). Published
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 119 zebrafish asip1 and zebrafish eEf1α primers were used for RT-PCR (McCurley and Callard 2008; Ceinos et al. 2015). PCR reactions were performed in 25 µL volumes containing 1 µL of cDNA, 0’5 µL of each primer (10 µM), 12’5 µL of Dream Taq green PCR Master Mix (2x) and H2O nuclease free to 25 µL. PCR conditions were 94ºC for 5 min, 35 cycles of 94ºC for 30 sec, 60ºC for 30 sec, 72ºC for 2 min, and a final step of 72ºC for 7 min. PCR products were run on 1% agarose gels. Quantitative asip1 mRNA expression in the skin was determined by qRT-PCR. Each set of skin samples from zebrafish and spotted gar was set up in triplicate. The qRT- PCR reaction was carried out in 20 µL volumes, containing 1 µL of cDNA (diluted 1:10), 0’5 µL of each primer (10 µM), 12’5 µL of SYBR Green/ROX qPCR Master Mix (2x) (ThermoFisher) and H2O nuclease free to 20 µL. The thermal protocol was: 95ºC for 10 min and 40 cycles of 95ºC for 15 sec and 60ºC for 1 min. Primers are listed in Table 5.1.
d) RNA in situ hybridization and confocal imaging
RNA in situ hybridization on zebrafish (210 dpf; 22mm) and spotted gar (270 dpf; 130 mm) paraffin sections were carried out following methods as described (Cerdá-Reverter et al. 2000; Rotllant et al. 2008). Fish (n=4; 2 of each fish species) were fixed in 4% paraformaldehyde and 0.1 M phosphate buffer, for 2 days at 4ºC, dehydrated and embedded in Paraplast (Sherwood, St Louis, MO, USA). Serial 12μm transverse anatomical sections (rostrocaudal direction) were cut using a rotary microtome. Sections were mounted on 3-aminopropyltriethoxylane (TESPA)-treated slides, air-dried at room temperature (RT) overnight and stored at 4ºC under dry conditions and used for hybridization within one month. RNA in situ hybridization was performed using digoxigenin-labeled antisense riboprobes as previously described (Rotllant et al. 2008). Antisense and sense riboprobes were made from linearized full-length zebrafish asip1 and partial-length spotted gar asip1. For cryostat sectioning, 150 dpf Tg(Drer.asip1-iTol2-eGFP-BAC) and Tg(Loc.asip1- iTol2-eGFP-BAC) fish were fixed in 4% paraformaldehyde overnight at 4ºC, washed with PBS, transferred to 15% sucrose, followed by 30% sucrose, and then embedded and frozen in OCT media. Serial 25μm rostro-caudal transverse sections were collected on Poly-Lysine slides (Thermo scientific) and allowed to dry. Slides were
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washed in PBS, dyed with 4,6-diamidino-2-phenylindole (DAPI, 3μM) to stain cell nuclei, mounted in mowiol and imaged on a Leica TCS SP5 confocal microscope.
e) Isolation of BAC clones containing the asip1 locus and BAC transgene construction
The zebrafish BAC clone (16N21, 214 Kb) from Danio Key Library, which includes the asip1 locus, was ordered from Source BioScience (Nottingham, UK). Spotted gar BAC clone (VMRC-56-161-M17, 130 Kb) from BAC library VMRC-56 (Braasch et al. 2016) was identified by PCR screening with a pair of asip1-specific primers that were designed using the asip1 gene sequence in the spotted gar Ensembl database (accession no. ENSLOCG00000007193). BAC transgene construction was performed as previously described (Suster et al. 2011). We used homologous recombination in E.coli to introduce the iTol2-amp cassette from the pCR8GW-iTol2- amp plasmid (Suster et al. 2011) into the BAC plasmid backbone, which contains the inverted minimal cis-sequences required for Tol2 transposition. The insertion of the reporter gene eGFP into the Drer.asip1 and Loc.asip1 locus of Drer.asip1-iTol2- BAC clone and Loc.asip1-iTol2-BAC clone, respectively, was carried out by homologous recombination. Briefly, the eGFP-Kan reporter gen cassette was amplified from eGFP-pA-FRT-Kan-FRT plasmid by PCR, together with a sequence of 50bp homologies to the Drer.asip1 or Loc.asip1 translation start site. After transformation of Drer.asip1-iTol2 or Loc.asip1-iTol2 BAC clone-containing cells with the PCR product, homologous recombination took place between the PCR product and the Drer.asip1-iTol2 or Loc.asip1-iTol2 BAC clone resulting in integration of the vector insert into the BAC clone, placing the eGFP reporter gene under control of the Drer.asip1 or Loc.asip1 regulatory regions within the clone (Drer.asip1-iTol2-eGFP-BAC or Loc.asip-iTol2-eGFP-BAC). Primers used are listed in Table 5.1.
f) Generation of BAC-transgenic zebrafish and analysis of transgene expression
BACs functionalized with iTol2kan were prepared in a concentration of 200 ng/µL together with Tol2 transposase mRNA in a concentration of 200 ng/µL and Phenol red solution (0,1%). Around 2nL of this mixture was microinjected into the
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 121 cytoplasm of zebrafish embryos at the one- or two-cell stage. A dissection microscope (MZ8, Leica) equipped with a MPPI-2 pressure injector (ASI systems) was used for microinjection. To observe pigment cells, double transgenic zebrafish lines (Tg(Drer.asip1-iTol2-eGFP-BAC;sox10:mRFP) and Tg(Loc.asip1-iTol2-eGFP- BAC;sox10:mRFP)) were obtained by setting up crosses between the asip1 BAC reporter gene transgenic lines and Tg(sox10:mRFP) (M. Levesque; obtained from the Nüsslein-Volhard Lab). For fluorescent microscope imaging, transgenic zebrafish of 3-days post-fertilization (dpf) (n=50), 30 dpf (n=20) and 100 dpf (n=15) were anesthetized with tricaine methasulfonate (MS-222-, Sigma-Aldrich) and photographed with a Leica M165FC stereomicroscope equipped with a Leica DFC310FX camera. Confocal imaging was carried out in a Leica TCS SP5 confocal microscope.
Table 5.1. Gene-specific primer sequences used for RT-PCR and qRT- PCR analysis. Dre (Danio rerio), Loc (Lepisosteus oculatus).
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III. RESULTS
a) Genomic and peptide comparison
We determined the organization of the asip1 locus in gar and compared it to zebrafish. In both gar and zebrafish, the asip1 gene has three coding exons distributed over regions of about 3.6 and 7.4 kb, respectively. Peptide alignment shows an identity level of 47% and a similarity of 62% between zebrafish and spotted gar Asip1 (Fig. 5.1A). Comparative mapping of asip1 regions in zebrafish and spotted gar shows high conserved genomic organization (Fig. 5.1B). Sequence conservation analysis of the genomic region upstream and downstream of asip1 was performed to learn if any non-coding elements are conserved between zebrafish and spotted gar. MLAGAN alignments of genomic sequence spanning 6:49873114 to 6:49991996 for Zebrafish (Zv9), and 18:14032011 to 18:14456756 for gar (LepOcu1) show few conserved non-coding regions (red arrows) and coding regions (blue arrows) (Fig.5.1C,D).
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Figure 5.1. Identification of zebrafish and spotted gar BACs likely to recapitulate asip1 expression. (A) T-coffee Asip1 peptide sequence alignment of Drer.asip1 (ENSDART00000113083.2) and Loc.asip1. (ENSLOCT00000008713.1). Grey boxes show conserved amino acid residues. (B) Genomic organization and gene synteny comparison for genes annotated from zebrafish BAC clone (16N21) from DanioKey Library and spotted gar BAC clone (VMRC-56-161-M17) from BAC library VMRC-56 (Postlethwait Lab; Institute of Neuroscience, University of Oregon). Picture style was adapted from Genomicus [http://www.dyogen.ens.fr/genomicus-57.01/cgi-bin/search.pl]. Genes are depicted by colored polygons and transcriptional orientation is indicated at the angled end of each gene. Gene names are indicated above. Orthologs across the two species are depicted in the same colors. (C, D) Distribution of the spotted gar conserved coding and non-coding regions. Vista browser representation of the genomic region surrounding the spotted gar asip1 gene (LepOcu1, chr18:14032011-14456756), using Zv9 zebrafish genome (chr6:49873114-49991996) as reference. Conservation is represented as vertical peaks. Conserved non-coding elements (CNEs) and coding elements with possible enhancer activity are marked as red and blue bars, respectively.
b) Expression of asip1 in spotted gar compared to zebrafish
To investigate the role of Asip1 in establishing the spotted gar dorsal-ventral pigment pattern, we first compared the tissue expression distribution of asip1 expression as detected by RT-PCR in zebrafish (Fig. 5.2A) and spotted gar (Fig. 5.2B).
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Figure 5.2. Spatial expression of asip1 gene in zebrafish (A) and spotted gar (B). RT-PCR analysis of tissue specific expression pattern of asip1 in adults. Scale bar: (A)2 mm, (B) 1 cm.
In 270 dpf adult gar, high levels of asip1 mRNA were found only in the eye and ventral skin and were present at lower levels in the heart (Fig. 5.2B). In 11 dpf spotted gar larvae, asip1 expression was not found in the skin (Fig. 5.3). Detecting asip1 expression particularly in the ventral skin of zebrafish and adult gar (Fig.5.2A and 5.2B), was consistent with a possible role in adult pigment pattern formation, but did not have the quantitative resolution to decide whether that expression might show a spatial difference.
Figure 5.3. Spatial expression of asip1 gene spotted gar larva. RT-PCR analysis of tissue specific expression pattern of asip1 in 11 dpf larva.
To test this question, we investigated whether there were dorsal-ventral differences in expression levels by qRT-PCR. We measured asip1 transcript levels in skin from different dorsal-ventral positions, specifically, in the belly, each body stripe (2V, 1V, D and 2D) and dorsal skin regions for zebrafish (Fig. 5.4A) and belly, each body stripe (2V, Medium and 2D) and dorsal skin regions for spotted gar (Fig. 5.4C). Figure 5.4B shows the expression levels of asip1 in skin samples obtained
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 125 along the dorsal-ventral axis in wild-type adult zebrafish. The highest levels of expression of asip1 were found on the belly; however, asip1 transcripts showed a gradual decrease from the belly to the dorsal midline of the fish. Figure 5.4D shows the expression levels of asip1 in skin samples obtained along the dorsal-ventral axis in adult gar. Equivalent to what is found in zebrafish, the highest levels of expression of asip1 in gar were also found in the belly; and asip1 transcripts similarly showed a gradual decrease from the belly to the dorsal midline of the fish. Additionally, we use mRNA in situ hybridization to examine whether asip1 mRNA was dorsal-ventrally expressed. In adult fish, we only saw detectable asip1 expression in the ventral skin area in both species (Fig. 5.4E,F,G,H,I). This expression analysis suggests that asip1 may have a conserved function among ray-finned fish and that function might be conserved with mammals, which also suggests that the regulation and the asip1- dependent mechanism might be conserved among ray-finned fish and mammals.
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Figure 5.4. Dorsal-ventral gradient expression of asip1 in adult zebrafish (A) and spotted gar skin (C). Normalized gene expression levels of (B) zebrafish and (D) spotted gar asip1 in skin samples from belly to dorsum (locations of skin sample points are indicated in the picture). Shown are log 10- transformed ΔCt values of asip1 relative to eEf1α and β-actin, respectively. Data are the mean ±SEM from eight samples after triplicate PCR analysis. (E) Schematic representation of the rostro-caudal transverse sections. (F-I) Rostro-caudal transverse sections (12 μm thick) of paraffin embedded, in situ hybridization of asip1 expression in 210 dpf zebrafish (F,H) and 270dpf spotted gar (G,I). The sections in (H,I) shows the expression of asip1 transcripts in the dermis region of zebrafish (H) and spotted gar (I) ventral skin (black arrows). No aisp1 expression was found in the dermis region of dorsal skin of both species (C,D). Superscripts a, b, c and d indicate statistical differences (P<0.05) in gene expression levels among skin region (statistics data are similar if share at least one letter not sure what you mean). Scale bar: (A) 2 mm, (C) 1 cm, (F,G,H,I) 50 μm. d, dermis; m, muscle.
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c) Generation and molecular analysis of BAC transgenic zebrafish
Having shown that asip1 was expressed with dorsal-ventral asymmetry in the skin of both zebrafish (Fig. 5.4B) and gar (Fig. 5.4D), we generated transgenic zebrafish lines to characterize conserved regulatory elements required for the evolutionarily conserved dorsal-ventral asip1 expression pattern found in zebrafish and spotted gar. We first tried conventional reporter transgene approaches generating several constructs of different sizes up to 13kb of the promoter sequence upstream of the zebrafish asip1 TSS (Transcription Starting Site) (data not shown). Injection of these reporter constructs did not show eGFP expression, suggesting that the regulatory elements of asip1 are not located in the regions used in the reporter assay. Therefore, we created four independent BAC transgenic zebrafish lines, containing either a BAC from a zebrafish or a spotted gar library carrying the asip1 gene modified to drive enhanced green fluorescent protein (eGFP) (two lines per BAC/species). In stable transgenic lines, eGFP expression was first detectable in 3 dpf embryos, when the developing opercle was the initial site of expression for Drer.asip1-iTol2-eGFP-BAC (Fig. 5.5A–B, White arrows), which mimicked the already well described expression pattern of the endogenous asip1 mRNAs in zebrafish (Ceinos et al. 2015) and in the developing pectoral fins for the spotted gar BAC (Fig. 5.5C-D, white arrows). At 30 dpf, eGFP expression was still present in the opercle, dermal branchiostegal rays, quadrate dermal bone and pectoral fin folds for Drer.asip1-iTol2-eGFP-BAC (Fig. 5.5E-F). In the Loc.asip1-iTol2-eGFP-BAC transgenic line, eGFP was mainly expressed in the pectoral fin folds, interopercular bone and pectoral girdle (Fig. 5.5G-H). Furthermore, eGFP was also detectable in ventral skin area (Fig. 5.5F-H, white arrows) in both BAC transgenic lines. At 100 dpf, strong eGFP expression was seen in the ventral skin area, in the base of pectoral and pelvic fins, and eGFP expression was still detectable in the pharyngeal region in both BAC transgenic lines (Fig. 5.5I-P).
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See legend on the next page next the on legend See
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5 Figure 5. Figure
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Figure 5.5. BAC transgenes exhibit endogenous asip1-like expression in most tissues. Shown are representative eGFP fluorescence images in Drer.asip1-iTol2-eGFP-BAC (A,B,E,F,I,J,M,N)) and Loc.asip1-iTol2-eGFP-BAC (C,D,G,H,K,L,O,P) stable transgenic zebrafish lines. (A,C,I,K,M,O) are lateral views anterior to the left and (B,D,E,F,G,H,J,L,N,P) ventral views anterior to the left. eGFP fluorescence is first detected at 3dpf (A,B,C,D). (A, B) lateral and ventral views of eGFP expression in larval heads of Drer.asip1-iTol2-eGFP-BAC transgenic zebrafish showing specific expression restricted to the developing opercle. (C,D) lateral and ventral views of eGFP expression in larval heads of Loc.asip1-iTol2-eGFP-BAC transgenic zebrafish showing expression limited to the pectoral fins. (E-H) ventral views of eGFP expression in 30dph larvae, showing increased expression in different pharyngeal bone structures (opercle, dermal branchiostegal rays, quadrate dermal bone and pectoral fin folds) and in ventral skin area (white arrows) in Drer.asip1-iTol2-eGFP-BAC (E,F) and in the pectoral fin folds, interopercular bone, pectoral girdle and ventral skin area (white arrows) in Loc.asip1-iTol2-eGFP-BAC transgenic zebrafish line (G-H). (I-P) eGFP expression in 100dph fish, showing increased expression in the ventral skin region and the base of pectoral and pelvic fins of both BAC transgenic lines (M-P), eGFP was also detectable in different bone structures of pharyngeal region in both BAC Transgenic lines (I-L). Scale bars: (A-H) 200 μm; (I-P) 1mm. br, brachiostegals; e, eye; iop, interopercular bone; m, Meckel’s cartilage; op, operculum; pf, pectoral fin; pop, preopercle; pvf, pelvic fin; pg, pectoral girdle q, quadrate; 1D, 1V, 2V stripes.
eGFP expression in adult zebrafish (150 dpf) monitored by direct observation under confocal fluorescence was strongly detected in the ventral skin region and peritoneal membrane, while low expression levels were found in the dorsal skin region in both BAC transgenic lines (Fig. 5.6A-D). Therefore, at 100-150 dpf, both transgenic zebrafish lines containing either a BAC from a zebrafish or a spotted gar library showed highly similar eGFP expression patterns recapitulating the spatial pattern of endogenous asip1 transcript in adult zebrafish (Fig. 5.4B,F,H).
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Figure 5.6. Comparison of eGFP expression in rostro-caudal transverse sections (25 μm) from 150dpf Drer.asip1-iTol2-eGFP-BAC (A,C) and the Loc.asip1-iTol2-eGFP-BAC (B,D) transgenic zebrafish. Drer.asip1-iTol2-eGFP-BAC and Loc.asip1-iTol2-eGFP-BAC zebrafish show strong eGFP expression in the ventral skin region (dermis) and peritoneal membrane (C,D). Variegated eGFP fluorescence is seen in Loc.asip1-iTol2-eGFP-BAC dorsal dermis area (B). Cells that do not express eGFP are revealed with DAPI staining (blue color). Scale bars: 50 μm. d, dermis; ms, muscle; pc, peritoneal cavity; pm, peritoneal membrane.
To further characterize cells expressing asip1 in the developing opercle, dermal branchiostegal rays, quadrate dermal bone, pectoral fins and ventral skin, we crossed BAC transgenic lines to a sox10:mRFP line, which marks neural crest derivatives with membrane-bound monomeric red fluorescent protein (mRFP) (Fadeev et al. 2015) (Fig. 5.7). Interestingly, although both asip1 and sox10 transgenes were expressed in the pharyngeal bones and pectoral fins, they were expressed in different cell types (Fig. 5.7A-F, revealing that the asip1 gene (eGFP positive cells) was not expressed in the pre- and post-migratory neural crest cells (mRFP positive cells) in both zebrafish or spotted gar BAC transgenic lines. Figure 5.7G-H shows the sox10:mRFP positive cell localization in the ventral skin region (E,G) and dorsal-
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 131 ventral sagittal section (H) in Drer.asip1-iTol2-eGFP-BAC 30dph zebrafish transgenic line. mRFP-labeled neural crest derivatives were predominantly present in the peritoneal membrane region where a reflecting iridophore layer is located. In contrast, eGFP-labeled asip1 cells in the peritoneal membrane region were present, but remained mostly in the ventral dermis area (Fig. 5.7H).
Figure 5.7. See legend on next page.
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Figure 5.7. Confocal image projections of live Tg(Drer.asip1-iTol2-eGFP-BAC/sox10:mRFP) (A,C,E,G,H) and Tg(Loc.asip1-iTol2-eGFP-BAC/sox10:mRFP) (B,D,F) 5dpf larvae (A,B) and 30dpf juvenile zebrafish. (C,D,E,F). Ventral view of a live 5 dpf larvae, anterior is to the left. sox10:mRFP specifically labels the pre- and post-migratory neural crest cells. (A,B) mRFP strongly labels the cartilages of maxillary, mandibular arches and pectoral fins. (A) Drer.asip1-iTol2-eGFP-BAC labels the developing opercle and dermal branchiostegal rays. (B) Loc.asip1-iTol2-eGFP-BAC labels the pectoral gridle. (A,B) The pectoral fin is labeled by both fluorescent proteins at 5 dpf. (C,D) Ventral view of a live 30dpf larvae heads, anterior is to the left. sox10:mRFP labels the cartilages of maxillary and mandibular arches. Drer.asip1-iTol2-eGFP-BAC labels different pharyngeal bone structures (interopercle, dermal branchiostegal rays, quadrate dermal bone) (C), and Loc.asip1-iTol2-eGFP-BAC also labels some pharyngeal bone structures (interopercle and dermal branchiostegal rays) (D). (E,F) Ventral dermis region was strongly labeled by Drer.asip1-iTol2-eGFP-BAC and Loc.asip1-iTol2- eGFP-BAC. (G) Magnified view of ventral skin region from (E), showing dermis, peritoneal membrane and peritoneal cavity region. (H) Fluorescent dorsal-ventral sagittal projection image of ventral skin area of 30 dpf juvenile showing clear sox10:mRFP expression in peritoneal membrane region where a reflecting iridophore layer is located. The peritoneal membrane region is also positive for Drer.asip1-iTol2-eGFP-BAC. However, the dermis region is exclusively labeled by Drer.asip1- iTol2-eGFP-BAC. (I) Schematic representation of pictures G and H imaging. (J) Scheme of different layers in I. Scale bar: (A,B) 100 μm, (C,D,E, F) 200 μm, (H,J) 50 μm. br, brachiostegals; ch, ceratohyal; cs, coraco scapular cartilage; d, dermis; e, eye; iop, interopercular bone; m, Meckel’s cartilage; ms, muscle; o, opercle; pc, peritoneal cavity; pf, pectoral fin; pg, pectoral gridle; pm, peritoneal membrane; pq, palatoquadrate; q, quadrate; sk, skin.
This revealed that asip1 gene was not detected in pre- and post-migratory neural crest cells and their derivatives, which strongly suggests that asip1 is not expressed by the pigment cells itself but by surrounding dermal cells. Therefore, we show that the asip1 gene is specifically expressed in fish dermal fibroblasts (Fig. 5.8).
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Figure 5.8. Image projections of rostro-caudal transverse sections of ventral skin region (40 μm) from Drer.asip1-iTol2-eGFP-BAC. Expression of eGFP (green) together with DAPI (blue) was analyzed in 1 year old fish. Stereomicroscope image projection of the ventral region of skin (A, B) shows the different layers of the skin (epidermis and dermis) around ventral scales with DAPI staining (A) and eGFP fluorescence (B). eGFP labels a thin layer of fibroblast within the dermis layer of skin next to the scales. Confocal image projection of the ventral skin (C) shows the different layers of the skin (epidermis and dermis) in a region without scales. eGFP labels dermal fibroblasts within the dermis layer of skin. Scale bars: (A, B) 50 μm, (C) 25 μm.
In the peritoneal cavity, the thin layer of eGFP expression detected in the peritoneal membrane is not coincident with the mRFP-labeled neural crest derivatives, but it is also expressed in adjacent peritoneal fibroblasts (Fig. 5.9).
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Figure 5.8. Confocal Image projection of rostro-caudal transverse sections of intraperitoneal membrane region (40 μm) from Drer.asip1-iTol2-eGFP-BAC/sox10:mRFP. Expression of eGFP (green) and sox10 (red) together with DAPI (blue) was analyzed in 1 year old fish. eGFP labels dermal fibroblasts within the extracellular matrix. mRFP labels neural crest derivatives, including iridophores. Scale bars: 25 μm.
IV. Discussion
It is well known that the dorsal-ventral countershading pigment pattern, which is one of the most common pigmentary adaptations in vertebrates, is driven by differential expression of agouti-signaling protein Asip, with high ventral Asip expression driving pale belly color (Manceau et al. 2011; Ceinos et al. 2015). In mammals, pigment pattern formation depends on the regulation of the production of different melanin types and levels in melanocytes (Lin and Fisher 2007), whereas in teleostean fish, it results from a patterned distribution of multiple types of chromatophores (Hirata et al. 2005; Kelsh et al. 2009; Parichy and Spiewak 2015; Irion et al. 2016). It would be reasonable to hypothesize that the conserved expression of Asip reflects a conserved role in pigment pattern formation in vertebrates, and specifically that the pale ventrum of vertebrates depends upon elevated levels of Asip expression.
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Whereas in mammals, ASIP influences the type of melanin produced, in teleost fish it is likely to affect the ratios between different cell-types, i.e. promoting development of iridophores and suppressing development of melanophores (Guillot et al. 2012; Ceinos et al. 2015). Thus, rather unexpectedly, Asip appears to control an evolutionarily conserved pattern by regulating different cellular mechanisms in mammals and in teleost fish. To test of this model, we analyzed and compared the agouti locus in zebrafish and spotted gar using a BAC transgenic approach. As mentioned above, within ray-finned fish, spotted gar is a member of holostean linage that diverged from the teleost linage before the TGD; consequently, the spotted gar genome provides an outgroup to the teleost lineage with a genome duplication status at the same level as mammals and thus spotted gar provides information regarding vertebrate genome evolution (Amores et al. 2011; Braasch et al. 2016). We report here the cloning, genomic organization and spatial and temporal expression of the asip1 gene in spotted gar.
To characterize conserved regulatory elements required for the evolutionary conserved asip1 expression in the adult fish, we report the generation and expression analysis of two independent BAC transgenic zebrafish lines, containing a zebrafish and/or spotted gar BAC carrying the asip1 gene modified to drive enhanced green fluorescent protein (eGFP). Our gar asip1 transgenic zebrafish line is the first one ever to introduce a holostean BAC clone into the zebrafish genome. Because asip1 transcriptional control is complex, we decided to use BACs, which is likely to reduce potential problems associated with “standard” reporter transgene approaches, including positional effects (Giraldo and Montoliu 2001; Suster et al. 2011) and the isolation of isolated and individualized elements that may not recapitulate the complete endogenous cooperative interactions between regulatory elements of a larger genomic region.
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Figure 5.10. Alignment of agouti-signaling protein (ASIP) and agouti-related protein (AGRP) amino acid sequences. Gar sequences are in bold letters. Dashes were introduced to improve alignment. Black boxes show amino acid residues conserved in all sequences. Green boxes show residues only conserved in Asip1 sequences. Red boxes indicate residues only conserved in Agrp1 sequences. Blue boxes show residues of the short tail present in all AGRP sequences. Lines joining cysteine residues indicate putative disulfide bonds forming the cysteine domain. Arrow shows conserved motif for AGRP post-transcriptional processing.
The spotted gar asip1 gene shows a similar genomic organization present in other vertebrate species (Bultman et al. 1992; Kwon et al. 1994; Cerdá-Reverter et al. 2005; Guillot et al. 2012; Agulleiro et al. 2014). It consists of three coding exons that encode a putative peptide of 127 amino acids. Multiple sequence alignments indicated that the deduced amino acid sequence of spotted gar Asip1 was also highly conserved compared to orthologs from other vertebrate lineages (Fig. 5.10).
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 137
Spatial expression analysis revealed that the asip1 gene in adult spotted gar is expressed in diverse tissues as shown in other vertebrate species (Girardot et al. 2005; Cerdá-Reverter et al. 2005; Guillot et al. 2012; Agulleiro et al. 2014; Ceinos et al. 2015), therefore suggesting other possible roles of the asip1 gene in addition to its role in pigmentation, such as energy balance (Miller et al. 1993; Guillot et al. 2016). We also show conservation of a dorsal-ventral gradient of asip1 expression in the skin of adult spotted gar with lower levels in the dorsum and higher levels in the belly. This expression pattern correlates with the countershading coloration in which the dorsal side (upper side) of the animal is darker than its ventral (lower) side. In contrast to the high expression level of asip1 found in adult spotted gar skin, our data give no evidence of asip1 expression in 11 dpf pre-metamorphic spotted gar larvae skin. Therefore, our results reveal that asip1 is important for adult spotted gar pigment pattern formation with a reduced or non-existent effect in embryonic and larval pigment pattern as it has been described in other fish species like zebrafish (Ceinos et al. 2015).
Asip transcriptional control is complex because the gene uses at least two alternatives promoters in different mammalian species (Bultman et al. 1992; Vrieling et al. 1994; Girardot et al. 2005; Fontanesi et al. 2010; Oribe et al. 2012). Our previous attempts to characterize the conserved regulatory elements of asip1 in zebrafish by conventional reporter transgene assays did not show eGFP expression, which suggests that essential regulatory elements of asip1 must be located further from the locus. Alternatively, BAC transgenesis showed that 214 Kb and 130 Kb genomic sequences including the asip1 locus of zebrafish or spotted gar, respectively were capable of recapitulating the endogenous dorsal-ventral expression pattern of asip1 in adult zebrafish and similar to the dorsal-ventral asip1 expression pattern in adult spotted gar. Therefore, our BAC transgenesis approach identified large genomic regions carrying divergent sequences but a conserved function in adult ray- finned fish. These results confirm the complex regulation of the asip gene and show the conserved evolutionary roots of this complex regulation among ray-finned fish. Surprisingly, our observations also indicate that this conserved evolutionary asip1 expression pattern in adult zebrafish and spotted gar is not found in larvae and/or pre-metamorphic fish, raising the possibility that different regulatory elements could be involved in the larval control of asip1 gene expression in both species. Further
138 Chapter 5 analyses are necessary to fully characterize the specific regulatory elements (enhancers) that control the spatial and temporal gene expression of asip1 and its sequence and function conservation.
In most mammalian species ASIP is a protein secreted by dermal papilla cells (Bultman et al. 1992; Miller et al. 1993) that acts as a paracrine factor for melanocyte pigment production (Millar et al. 1995). In adult teleost fish, Asip1 is produced mainly by dermal fibroblasts in the ventral skin and by peritoneal fibroblasts in the intraperitoneal membrane, and possibly acts nearby (in a paracrine fashion) to reduce melanophore number and melanophore differentiation in the ventrum, both in the skin and in the intraperitoneal membrane, but has no effect on larvae and/or pre- metamorphic pigment pattern (Ceinos et al. 2015), likely because asip1 in zebrafish larvae is not expressed in the skin.
Using transgenic reporter lines expressing eGFP and mRFP under the asip1 regulatory machinery and sox10 promoters, respectively, we demonstrated here that, although both transgenes were expressed in the pharyngeal bones, pectoral fins, and peritoneal membrane, they were expressed in distinctive cell types. asip1/green was expressed in dermal fibroblast, however, sox10/red was expressed in neural crest derivatives, including craniofacial chondrocytes and pigment cells, suggesting that asip1 transgenes (eGFP positive cells) and thus endogenous asip1 was not expressed in the pre- and post-migratory neural crest cells (sox10:mRFP positive cells).
In conclusion, we demonstrate for the first time the existence of a dorsal-ventral gradient of asip1 expression in the skin of adult spotted gar with a high level in the ventrum and lower levels in the dorsum, conserved with the dorsal-ventral gradient in zebrafish and other teleosts. We describe the generation of bacterial artificial chromosome (BAC) transgenic (Tg) zebrafish lines that harbor zebrafish or spotted gar BACs carrying the asip1 gene modified to drive enhanced green fluorescent protein (eGFP). The resulting transgenic lines showed eGFP expression in a pattern that mirrors that of endogenous asip1 gene expression in adult zebrafish and spotted gar and illustrated that these transgenic zebrafish lines are a promising tool for future identification of the regulatory elements required for regular dorsal-ventrally gradients of asip1 expression in adult fish. The development and the pigment pattern of both transgenic lines were totally normal. Therefore, our study provides new
Evolutionary ancestry of dorsal-ventral pigment asymmetry in fish 139 molecular genetic tools for functional investigations of the regulatory elements that control spatial and temporal pattern of asip1 expression and its role in pigment pattern formation. The work emphasizes the need to characterize asip1 transcriptional regulation to further elucidate the conserved functions of ASIP protein among bony vertebrates despite the different cellular mechanisms of pigment pattern formation in mammals and fish.
General discussion and conclusions 143
GENERAL DISCUSSION
Pigment abnormalities are one of the most important factors affecting the economic viability of flatfish aquaculture. The reason for the relatively high incidence of pigment abnormalities in aquaculture flatfish species is unknown Thus, although pigment alterations can be hereditable (Hoekstra 2006) they are often linked as a results caused by inappropriate rearing conditions (light intensity, nutritional insufficiencies, stress, etc.) or special post-harvesting manipulation (use of cold and/or of ice) (Boglione and Costa 2011). Currently, there is scanty information about the molecular regulation of pigmentation in aquaculture fish species, which makes the control of this problem difficult if not impossible. Although more than 100 loci have been identified as being involved in vertebrate pigmentation, the melanocortin system is consistently a key determinant of the pigment phenotype. In fact, recent studies from our laboratory have demonstrated that the melanocortin system is also a key player in the establishment of the adult pigment pattern in fish and it has been shown that components of the melanocortin system controls light deviations of pigment from wild standard in turbot (Scophthalmus maximus), sole (Solea senegalensis) (Guillot et al. 2012) and zebrafish (Danio rerio) (Ceinos et al. 2015). However, we do not know its molecular regulation. Better understanding the molecular regulation of pigment pattern development and pigment abnormalities in fish will not only provide information relevant to the design of new strategies for intensive fish culture, but will also provide basic knowledge on vertebrate pigmentation development.
Therefore, the fundamental scientific objective of this PhD Thesis was to do an integrated study to evaluate the molecular regulation of pigmentation in fish through the characterization of a key factor of the melanocortin system, the endogenous antagonist agouti signaling protein (asip1) gene, and its receptor, the melanocortin 1 receptor (mc1r) gene. Due to the conservation of cell biological and developmental processes across all vertebrates, the studies presented in this thesis were carried out in zebrafish, a prominent model animal for understanding fish pigmentation (Irion et al. 2016). 144 Chapter 6
To that end, we investigated the functional role of Asip1 in fish pigment pattern establishment. Second, we determine the in vivo functional role of mc1r in fish pigmentation and define the receptor functionality and the contribution of the Asip1- Mc1r signaling to fish pigmentation. Third, we analyzed the transcriptomic profiling induced by decreased activity of the central melanocortin system, and characterized the different biological processes and/or signaling pathways downstream of the melanocortin system in zebrafish skin. Finally, we determine the conserved regulatory elements and the evolutionary ancestry of the dorsal-ventral pigmentation mechanism in fish.
Our results demonstrate that Asip1-Mc1r signaling play a critical role in dorsal- ventral pigment pattern establishment in adult zebrafish. From an evolutionary point of view, our results show both the novel and conserved aspects of the molecular mechanisms of asip1 function. We show in fish that Asip1 regulates the dorsal-ventral pigment pattern likely through a mechanism that affects melanophore, xanthophore and iridophore number and/or differentiation. In contrast, mammalian countershading, an equivalent dorsal-ventral pigment patterning process, comes about by regulating the differential pigment production and differentiation of their only chromatophore type, the melanocyte. Consequently, our results suggest that Asip1-dependent countershading and its molecular machinery are conserved between mammals and teleost fish, but, rather surprisingly, the cellular basis for pigment pattern formation is different in the two groups. Additionally, our results also provide evidence that the mechanism leading to the dorsal-ventral pigment pattern was likely in place before the origin of teleosts.
Finally, our results confirm the pleiotropic nature of the agouti gene. Thus, this reveals the ability of asip1 gene to simultaneously regulate pigmentation and other physiological processes in fish skin.
As reviewed in Chapter 1, Pigment pattern formation – e.g. the differential pigmentation of skin, hair or feathers – is a classic problem in pattern formation. In mammals pigment pattern formation depends upon the regulation of the production of melanin types and levels in melanocytes. To date, the most important regulator of skin and hair pigmentation in mammals is the ASIP-MC1R system: the agouti locus encodes the Agouti signaling protein (ASIP), which antagonizes the effects of α-melanocyte stimulating hormone (α-MSH) binding to the MC1R on the follicular melanocyte, resulting in a decrease in the production of eumelanins (dark pigments) and an increase
General Discussion and conclusions 145 in pheomelanins (red/yellow pigments) (Lu et al. 1994). A key aspect of mammalian pigment pattern and one that is widely conserved is the pale ventrum and darker dorsum; the underlying mechanism is regulation of ASIP expression, with high ventral Agouti driving pale belly color (Vrieling et al. 1994; Manceau et al. 2011).
Fish often show a pale ventrum, but here the pigmentation mechanism is very different with independent cell-types, each expressing chemically distinct pigments (not just melanins), being differentially distributed. Thus, recent studies showing that not only the agouti gene, but even the elevated ventral expression levels, is conserved in different adult fish species (Cerdá-Reverter et al., 2005; Tadahide Kurokawa et al., 2006; Klovins and Schiöth, 2005). It is, therefore, of exceptional interest to establish whether Asip1 might also function in regulating fish pigment pattern formation, and if so how?
In adult zebrafish, the pigment pattern depends on the number and distribution of three kinds of cells, dark melanophores (homologous to mammalian melanocytes, orange xanthophores, and silver, shiny iridophores). Like mammalian melanocytes, these cells are derived from embryonic neural crest cells (Le Douarin and Kalcheim 1999; Le Douarin and Dupin 2003). In zebrafish, melanophores are distributed in four horizontal stripes alternating with three white interstripes rich in iridophores; the ventrum is also white due to an abundance of iridophores (Hirata et al. 2003, 2005; Kelsh et al. 2009; Guillot et al. 2016b).
We propose that the conserved expression of agouti may reflect a conserved role in pigment pattern formation in vertebrates, and specifically that the pale ventrum of all vertebrates depends upon elevated levels of agouti expression. We suggest that, whilst in mammals ASIP influences the type of melanin produced, in fish it is likely to affect the ratios of different cell-types (melanophores, xanthophores and iridophores).
Consistent with this suggestion, our previous results indicate that adult zebrafish show graded agouti expression, with highest levels in ventral regions (Ceinos et al., 2015). Surprisingly, our earlier observations also indicated that in more dorsal regions, both silver interstripes and black stripes show low levels of agouti, indicating that another mechanism, likely independent of agouti expression, may act in parallel to impose the striped pattern on the dorsal body.
146 Chapter 6
As a direct test of our model we have previously used an overexpression approach, having generated transgenic zebrafish (asip1-Tg) in which agouti is overexpressed using a ubiquitous promoter (Tol2-Eef1α/asip1). Our initial observations showed these fish to have decreased melanophores in dorsal, but not ventral, regions, suggesting that elevated Asip1 levels may inhibit melanophore development.
In order to dissect the role of Asip1 in zebrafish pigment pattern formation we have perform a series of experiments focused on understanding the molecular basis for the pigment pattern changes we observed.
Therefore, in Chapter 2 and 3 we study the genetic analysis of dorsal-ventral pigment pattern formation and the involvement of Asip1-Mc1r signaling in the countershading pattern by analyzing “loss-of-function” pigment phenotypes in zebrafish. We demonstrate that “loss-of-function” mutation of asip1 and mc1r genes result in the disruption of dorsal-ventral pigment pattern characterized by an apparent dorsal-to-ventral transformation (i.e. ventral skin becomes nearly as dark colored as dorsal skin). The ventral regions of the adult asip1 and mc1r mutant fish show a higher number of melanophores and xantophores than their WT siblings. The abnormal ventral skin darkening is due to a marked increase of melanophore and xanthophore number, together with a strong reduction of iridophores in the abdominal wall. The disruption of this dense and uniform sheet of iridophores in both mutants results in the loss of the light color of the belly. This decrease in the iridophore number could reflect a direct asip1 and mc1r contribution to iridophores development or it is possible that disruption of the iridophores sheet was the result of the negative short-rang effect due to the increase of melanophore number found in the abdominal wall of both mutants. Those two possibilities are not exclusive.
Therefore, our observations indicate that the color pattern changes could derive from both an inhibition of melanization, directly comparable to the mechanism of countershading documented in mammals, but importantly also from a novel mechanism regulating the number of melanophores and other pigment cells.
It was previously reported that high levels of Asip1 inhibit melanogenesis and melanophore differentiation (Guillot et al. 2012; Ceinos et al. 2015). We show that fish overexpressing asip1 has paler pigmentation than fish that lack mc1r. This suggests that asip1 does more than simply antagonize mc1r or block basal signaling, as the
General Discussion and conclusions 147 overexpression phenotype is stronger than absence of receptor. We also shown that fish that overexpress asip1 but also lack mc1r have an apparent pale homogenization of the color of the animal with its corresponding loss of countershading phenotype, similar to fish only lacking mc1r, indicating that asip1 is acting as an inverse agonist of mc1r to elicit the opposite response from that produced by α-Msh. It is, therefore, possible that the constitutive activation of Mc1r leads to dorsal melanogenesis, but the ventral expression of asip1 blocks the constitutive activity, thus inhibiting ventral melanogenesis. Overall, these results reveal that the mc1r is a crucial factor for asip1 signaling in fish pigmentation. Indicating that the molecular machinery of asip1-mc1r dependent dorsal-ventral pigment patterning is conserved between mammals and teleost fish, but, rather surprisingly, the cellular basis for pigment pattern formation is different in the two groups.
Additionally, our results reveal that both mutants exhibit a normal organization of stripes and the patterning of the fins are not affected. Thus, suggesting that zebrafish adult pigment pattern formation involved two largely independent processes: a dorsal- ventral pigment pattern mechanisms that utilizes Asip1, and a striping mechanism based on cell-cell interactions. These observations are consistent with our previous study, which showed that the stripe pattern and pigmentation of fin stripes of asip1-Tg fish was indistinguishable from that observed in WT fish (Ceinos et al. 2015).
To determine the time point when the phenotype of the asip1 and mc1r mutants becomes first apparent, we evaluate the pigment phenotype at different stage of development. asip1 mutants exhibit a normal pigmentation pattern during larval development, therefore suggesting that only adults pigment pattern is affected. In contrast, mutation of the mc1r gene produces a significant pigmentation changes with general reduction of melanophore number at all developmental stages.
From an evolutionary point of view, our results showed that the dorsal-ventral graded distribution of asip1 gene is also conserved in two distantly related adult ray- finned fish (Zebrafish and Spotted gar (Lepisosteus oculatus)), therefore showing that the dorsal-ventral patterning mechanism found in teleost is an ancient feature shared by other groups of ray-finned fish and not a specific feature arising as result of the teleost genoma duplication (TGD). Therefore, our results demonstrate that the mechanism leading to the dorsal-ventral pigment pattern was likely in place before the origin of teleosts.
148 Chapter 6
Finally, we show the extent to which the melanocortin system simultaneously regulates pigmentation and other possible physiological processes in fish skin. It has already been shown that the melanocortin system is a neuroendocrine system that regulates a wide range of endocrine and homeostatic processes (Cerdá-Reverter et al. 2011). In fish, melanocortin system pleiotropic effects have been previously described, showing that decreased activity of the central melanocortin system regulates several genes involved in the control of food intake and stress response (Guillot et al. 2016a). Our results show that asip1 simultaneously regulates pigmentation and other physiological processes including inflammation, energy metabolism, cell cycle and reproduction in fish skin. It has to be emphasized that most of the common term/pathway that are noticeably affected by asip1 overexpression in fish are also affected in similar overexpression approaches in mammals (Klebig et al. 1995). Therefore, we confirm the pleiotropic nature of the agouti gene. This reveals the ability of asip1 gene to simultaneously regulate pigmentation and other physiological processes in fish skin.
General Discussion and conclusions 149
CONCLUSSIONS
The graded expression of asip1 along the dorsal-ventral axis functions to establish the dorsal-ventral asymmetric pigmentation in fish.
asip1 gene is expressed in dermal and peritoneal fibroblasts in zebrafish
“Loss-of-function” mutation of asip1 gene has no effect on the embryonic pigment pattern and the stripe patterning.
Knockout mutant zebrafish that were homozygous for the asip1 allele displayed disorganized dorsal-ventral pigment pattern characterized by a dorsalization (i.e. a darkening due to an increased ratio of melanophores and xanthophores) of the ventral pigment pattern.
“Loss-of-function” mutation of mc1r gene has significant effect on the embryonic pigment pattern but do not affect the stripe patterning.
Knockout mutant zebrafish that were homozygous for the mc1r allele displayed disorganized dorsal-ventral pigment pattern characterized by a dorsalization (i.e. a darkening due to an increased ratio of melanophores and xanthophores) of the ventral pigment pattern together with an apparent pale homogenization of the color of the animal.
Asip1 is acting as an inverse agonist of Mc1r to elicit the opposite response from that produced by α-MSH.
Asip1-Mc1r signaling is the key molecular signaling pathway controlling dorsal- ventral pigment pattern establishment in adult fish.
150 Chapter 6
asip1 has the ability to simultaneously regulate pigmentation and other physiological processes in fish skin.
Most of the common biological processes that are noticeably affected by asip1 overexpression in fish are also affected in similar overexpression approaches in mammals
Spotted gar, one of Darwin's defining examples of 'living fossils', shows a dorsal-ventral gradient of asip1 expression with lower levels in the dorsum and higher in the belly, similar to the expression gradient in zebrafish, and other teleosts.
BAC transgenesis approach identified large asip1 genomic regulatory regions in zebrafish and spotted gar, carrying divergent sequences but a conserved function in adult ray-finned fish.
The conserved evolutionary asip1 expression pattern in adult zebrafish and spotted gar is not found in larvae and/or pre-metamorphic fish
Conserved expression of asip1 reflects a conserved role in pigment pattern formation in vertebrates, and specifically that the pale ventrum of vertebrates depends upon elevated levels of agouti gene expression.
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Annex: Supplementary Information 169
ANNEX: SUPPLEMENTARY INFORMATION
Supplementary table 1. KEGG pathway analysis. The p-value was obtained by Yate’s corrected chi square test.
Pathway p-Value Gender
Protein processing in endoplasmic reticulum 2,50E-06 F Protein processing in endoplasmic reticulum 3,67E-04 M Purine metabolism 3,54E-06 F Purine metabolism 1,00E-03 M Pyrimidine metabolism 2,69E-05 F Pyrimidine metabolism 3,14E-02 M Arginine and proline metabolism 6,96E-05 F Arginine and proline metabolism 6,48E-05 M Amino sugar and nucleotide sugar metabolism 1,50E-04 F Amino sugar and nucleotide sugar metabolism 6,86E-03 M NOD-like receptor signaling pathway 1,72E-03 F NOD-like receptor signaling pathway 3,14E-02 M RNA polymerase 1,97E-03 F RNA polymerase 3,78E-02 M Gap junction 1,40E-02 F Gap junction 2,96E-02 M Steroid biosynthesis 2,35E-02 F Steroid biosynthesis 9,51E-05 M GnRH signaling pathway 2,68E-02 F GnRH signaling pathway 1,82E-02 M Starch and sucrose metabolism 3,28E-02 F Starch and sucrose metabolism 3,77E-02 M Riboflavin metabolism 1,07E-03 F Acute myeloid leukemia 2,02E-03 F Apoptosis 2,63E-03 F Insulin signaling pathway 2,86E-03 F Phagosome 4,92E-03 F T cell receptor signaling pathway 9,36E-03 F Non-small cell lung cancer 1,02E-02 F Fc gamma R-mediated phagocytosis 1,25E-02 F N-Glycan biosynthesis 1,39E-02 F Alanine_ aspartate and glutamate metabolism 1,39E-02 F VEGF signaling pathway 1,45E-02 F B cell receptor signaling pathway 1,67E-02 F 170 Annex: Supplementary Information
Protein export 2,59E-02 F Leukocyte transendothelial migration 2,87E-02 F Chemokine signaling pathway 3,94E-02 F Vibrio cholerae infection 4,25E-02 F p53 signaling pathway 4,81E-02 F Dilated cardiomyopathy 1,61E-06 M Hypertrophic cardiomyopathy (HCM) 8,80E-06 M Drug metabolism - cytochrome P450 1,18E-04 M Biosynthesis of unsaturated fatty acids 1,33E-04 M Focal adhesion 2,78E-04 M One carbon pool by folate 3,86E-04 M Terpenoid backbone biosynthesis 5,18E-04 M Pentose phosphate pathway 5,33E-04 M Glutathione metabolism 6,52E-04 M Metabolism of xenobiotics by cytochrome P450 8,11E-04 M DNA replication 1,03E-03 M Type II diabetes mellitus 3,21E-03 M Vascular smooth muscle contraction 4,76E-03 M Glycolysis / Gluconeogenesis 5,60E-03 M PPAR signaling pathway 7,63E-03 M Mismatch repair 1,09E-02 M Cell cycle 1,12E-02 M Cardiac muscle contraction 1,29E-02 M Pyruvate metabolism 1,80E-02 M Phosphatidylinositol signaling system 2,11E-02 M Glyoxylate and dicarboxylate metabolism 2,16E-02 M Calcium signaling pathway 2,19E-02 M Proteasome 4,57E-02 M Protein processing in endoplasmic reticulum 2,50E-06 F
Annex: Supplementary Information 171
Supplementary table 2. Guanine synthesis pathway. The 10 most affected DEGs in each pathway. The values that fit criteria are underlined. Abbreviations: G, Glycolysis; PM, Purine metabolism; PPP, Pentose phosphate pathway.
p-Value Gene Log2-ER Log2-ER p-Value Accession ID in Pathway Name in Females in Males in Males Females
ENSDARG00000005827 eif2s1a +1,01 +3,01 2,59E-03 2,80E-02 PM ENSDARG00000038505 polr2h -1,53 -2,01 3,52E-04 2,90E-03 PM ENSDARG00000003519 aprt +1,36 +1,32 7,62E-03 2,21E-02 PM ENSDARG00000036371 acta1a -2,15 -0,96 3,96E-02 3,90E-02 PM ENSDARG00000070472 arf5 -1,07 -0,88 1,70E-03 3,45E-03 PM,G ENSDARG ENSDARG00000017820 000000178 -1,35 -0,88 9,34E-05 1,13E-02 PM 20 PM,PPP, ENSDARG00000018178 pgm2 -0,96 -2,10 1,91E-02 7,75E-03 G ENSDARG00000044807 dck -0,85 -1,13 2,51E-02 2,32E-03 PM,PPP ENSDARG00000055618 acta1b +2,47 +5,15 1,24E-01 2,56E-03 PM ENSDARG00000013236 dhcr24 -1,22 -2,55 1,17E-01 1,55E-02 PM ENSDARG00000015343 pgd -1,44 -2,13 3,27E-03 6,70E-03 PM, PPP ENSDARG00000006019 tktb -0,06 -1,91 8,28E-01 3,43E-03 PM, PPP PM, PPP, ENSDARG00000012366 fbp2 +0,11 +1,78 8,68E-01 1,83E-03 G ENSDARG ENSDARG00000012256 000000122 -1,00 -1,29 8,77E-05 2,33E-03 PM, PPP 56 PM, PPP, ENSDARG00000034470 aldoab -0,09 +1,15 8,88E-01 2,94E-02 G PM, PPP, ENSDARG00000011665 aldoaa -0,84 +1,08 7,46E-02 1,09E-02 G PM, PPP, ENSDARG00000019702 aldocb +0,19 -1,05 4,92E-01 4,54E-02 G PM, PPP, ENSDARG00000053684 aldob 0,00 +0,95 9,92E-01 1,39E-02 G ENSDARG00000015263 adma +0,54 1,36 5,52E-02 2,16E-02 G ENSDARG00000039007 eno3 -0,68 +0,99 5,62E-02 1,92E-02 G ENSDARG00000005423 pgam1a -0,87 -0,95 1,11E-03 2,83E-02 G
172 Annex: Supplementary Information
Supplementary table 3. DEGs in pigment cells (Higdon et al. 2013) affected by asip1. The values that fit criteria are underlined.
Log2-ER in Log2-ER p-Value in p-Value Accession ID Gene Name Females in Males Females in Males
ENSDARG00000028517 hbp1 +2,26 +0,40 4,30E-04 5,40E-01 ENSDARG00000061057 cyhr1 +0,94 -0,07 6,31E-03 7,71E-01 ENSDARG00000078042 il10rb +0,92 +0,33 3,20E-03 1,30E-01 ENSDARG00000075952 mdb2 -0,88 +0,41 6,31E-03 7,71E-01
Supplementary table 4. DEGs in neural crest cells affected by asip1. The values that fit criteria are underlined.
Log2-ER in Log2-ER p-Value in p-Value Accession ID Gene Name Females in Males Females in Males
ENSDARG00000003293 sox9a +1,85 +1,44 1,22E-02 2,01E-02 ENSDARG00000077818 nrg2a +1,57 +0,38 5,33E-03 3,30E-01 ENSDARG00000076472 ovol1 -1,07 -0,53 1,27E-02 2,82E-01 ENSDARG00000087377 lbh +1,02 -0,57 4,32E-03 1,99E-01 ENSDARG00000031894 lef1 +0,93 +0,27 2,19E-02 5,98E-01 ENSDARG00000043923 sox9b -0,61 -0,93 1,83E-01 2,66E-02 ENSDARG00000035598 coro1ca -0,55 -0,87 1,75E-02 1,74E-03
Annex: Supplementary Information 173
Supplementary table 5. Immune system. The 5 most affected DEGs in each pathway. The values that fit criteria are underlined. Abbreviations: BCRSP, B cell receptor signaling pathway; CSP, Chemokine signaling pathway; FGRMP, Fc gamma R-mediated phagocytosis; LTM, Leukocyte transendothelial migration, NlRS, NOD- like receptor signaling pathway; P, Phagosome; TCRSP, T cell receptor signaling pathway.
p-Value Gene Log2-ER Log2-ER p-Value Accession ID in Pathway Name in Females in Males in Males Females
BCRSP, ENSDARG00000052859 chp1 +1,47 -0,05 4,04E-02 7,99E-01 TCRSP BCRSP, CSP, ENSDARG00000007693 nfkbiab +1,47 +1,23 1,60E-02 4,45E-02 NlRS, TCRSP BCRSP, ENSDARG00000025788 chp2 +1,30 +0,43 1,82E-03 8,29E-02 TCRSP ENSDARG00000044718 vav2 +0,87 +0,80 8,00E-03 3,10E-02 BCRSP BCRSP, ENSDARG00000070573 mapk3 -0,84 -0,88 5,16E-03 4,45E-03 NlRS CSP, ENSDARG00000041919 ccl38.1 +2,14 +0,80 4,60E-03 4,14E-03 NlRS ENSDARG00000061796 rasgrp4 +1,96 +0,41 9,28E-03 1,90E-01 CSP CSP, ENSDARG00000041959 cxcr4b +1,53 +0,55 1,17E-02 2,98E-01 LTM ENSDARG00000055186 ccr9a +1,50 +1,03 2,85E-02 1,98E-01 CSP ENSDARG00000058230 rps6kb1b -1,91 -0,91 3,65E-04 3,71E-02 FGRMP ENSDARG00000026350 wasb 1,28 0,33 2,55E-03 2,36E-01 FGRMP ENSDARG00000033735 ncf1 1,18 1,02 2,83E-03 6,81E-03 FGRMP ENSDARG00000015006 dnm1l -1,10 -0,29 7,97E-04 8,67E-02 FGRMP ENSDARG00000090369 zgc:86896 -1,01 -0,08 7,36E-03 8,68E-01 FGRMP ENSDARG00000038123 myl9a +3,12 +2,53 7,70E-04 3,14E-03 LTM ENSDARG00000071090 actn2b +2,30 +0,54 1,43E-02 2,28E-01 LTM ENSDARG00000036371 acta1a -2,15 -0,96 3,96E-02 3,90E-02 LTM, P ENSDARG00000007219 actn1 +1,75 +1,10 7,65E-04 9,76E-02 LTM ENSDARG00000003570 hsp90b1 -1,11 -1,30 3,27E-02 1,53E-03 NlRS ENSDARG00000023578 lpp +0,88 +1,01 3,26E-02 1,11E-02 NlRS ENSDARG00000037997 tubb5 -2,26 -1,21 6,67E-04 4,04E-02 P ENSDARG00000076290 calr -1,99 -4,00 4,13E-03 7,10E-03 P 174 Annex: Supplementary Information
si:ch73- ENSDARG00000094466 -1,95 -1,48 1,84E-03 1,54E-02 P 199e17.1 ENSDARG00000042708 tuba8l -1,66 -1,36 6,74E-03 8,89E-03 P ENSDARG00000055955 lcp2a +1,09 +0,19 2,78E-03 4,60E-01 TCRSP ENSDARG00000028721 mapk14b +1,04 +0,67 3,08E-03 1,24E-02 TCRSP
Supplementary table 6. Endocrine system. The 10 most affected DEGs in each pathway. The values that fit criteria are underlined. Abbreviations: IP, Insuline pathway; PPAR, PPAR Signaling pathway; T2DM, Type II diabetes mellitus.
p-Value Gene Log2-ER Log2-ER p-Value Accession ID in Pathway Name in Females in Males in Males Females
ENSDARG00000025428 socs3a +2,85 +2,64 1,90E-02 6,54E-02 IP ENSDARG00000087993 bada +1,95 +0,47 1,13E-04 3,40E-01 IP ENSDARG00000058230 rps6kb1b -1,91 -0,91 3,65E-04 3,71E-02 IP ENSDARG00000012274 eif4e1c -1,79 -1,28 6,76E-03 2,82E-02 IP ENSDARG00000038095 socs1a +1,70 +0,81 7,56E-03 8,64E-02 IP ENSDARG00000011373 mknk2a +1,64 +0,67 3,66E-04 1,34E-02 IP ENSDARG00000037099 irs2a +1,47 -0,63 2,20E-03 2,87E-02 IP ENSDARG00000015164 mknk2b +1,34 +1,72 5,86E-03 5,14E-02 IP ENSDARG00000054087 irs1 +1,28 +0,49 2,98E-02 5,46E-02 IP ENSDARG ENSDARG00000076338 000000763 +1,21 +0,49 7,87E-04 2,31E-02 IP 38 ENSDARG00000019532 fads2 -1,80 -2,40 9,22E-02 7,60E-03 PPAR ENSDARG00000033662 scd -2,37 -2,14 1,39E-03 3,56E-02 PPAR ENSDARG00000007697 fabp7a -1,70 -1,85 3,51E-03 1,17E-02 PPAR ENSDARG00000034650 fabp7b -0,39 -1,69 2,06E-01 5,33E-03 PPAR ENSDARG00000004687 acaa1 -0,67 -1,59 5,38E-02 5,20E-03 PPAR ENSDARG00000067520 slc27a2 +0,92 +1,57 3,81E-02 1,80E-02 PPAR ENSDARG00000069186 cyp27a1.2 -1,28 -1,29 9,23E-05 2,51E-02 PPAR ENSDARG00000055159 cyp27a1.4 -1,39 -1,29 2,37E-04 2,12E-02 PPAR ENSDARG00000002305 me3 +0,58 +0,96 2,33E-02 2,22E-02 PPAR ENSDARG00000086961 ENSDARG -0,21 +0,87 6,04E-01 3,38E-02 PPAR Annex: Supplementary Information 175
000000869 61 ENSDARG00000044155 mafaa -0,29 +2,39 5,59E-01 1,12E-02 T2DM ENSDARG00000070573 mapk3 -0,84 -0,88 5,16E-03 4,45E-03 T2DM
Supplementary table 7. Cell cycle. The 5 most affected DEGs in each pathway. The values that fit criteria are underlined. Abbreviations: AML, Acute myeloid leukemia; A, Apoptosis; CC, Cell cycle; DR, DNA replication; FA, Focal adhesion; MR, Mismatch repair; NSCLC, Non-small cell lung cancer; RP, RNA polymerase; PSP, p53 signaling pathway.
p-Value Gene Log2-ER Log2-ER p-Value Accession ID in Pathway Name in Females in Males in Males Females
ENSDARG00000007184 zbtb16a +2,86 +1,82 2,73E-02 1,29E-03 AML ENSDARG ENSDARG00000059120 000000591 +2,65 +2,21 2,69E-02 3,63E-02 AML 20 AML, A, ENSDARG00000087993 bada +1,95 +0,47 1,13E-04 3,40E-01 NSCLC ENSDARG00000058230 rps6kb1b -1,91 -0,91 3,65E-04 3,71E-02 AML ENSDARG00000055129 pim3 +1,80 +1,06 4,85E-03 2,54E-02 AML ENSDARG00000025608 casp6l1 -2,12 +0,01 3,39E-02 9,82E-01 A ENSDARG00000044562 cycsb -1,81 -0,18 4,75E-03 7,11E-01 A, PSP ENSDARG00000052859 chp1 +1,47 -0,05 4,04E-02 7,99E-01 A ENSDARG00000007693 nfkbiab +1,47 +1,23 1,60E-02 4,45E-02 A ENSDARG00000019507 mcm5 -1,48 -1,47 2,61E-03 1,37E-02 CC ENSDARG00000075682 orc6 -0,99 -1,34 2,67E-03 1,14E-02 CC ENSDARG00000040041 mcm4 -0,62 -1,32 1,48E-01 1,67E-02 CC ENSDARG00000057610 cdkn2c -1,82 -1,31 3,32E-02 1,60E-03 CC DR, FA, ENSDARG00000055618 acta1b +2,47 +5,15 1,24E-01 2,56E-03 MR ENSDARG00000005827 eif2s1a +1,01 +3,01 2,59E-03 2,80E-02 DR, MR si:ch73- DR, FA, ENSDARG00000076126 -0,48 +2,31 2,60E-01 1,94E-03 187m15.4 MR ENSDARG00000079111 actc1 -0,31 +1,72 4,89E-01 7,21E-03 DR, MR ENSDARG ENSDARG00000015911 -1,43 -1,71 5,24E-03 7,45E-03 DR, CC 000000159 176 Annex: Supplementary Information
11 ENSDARG00000013755 actn3a -0,27 +2,77 5,40E-01 9,14E-03 FA ENSDARG00000038123 myl9a +3,12 +2,53 7,70E-04 3,14E-03 FA ENSDARG00000002589 mylpfb -0,01 +2,27 9,89E-01 1,13E-03 FA ENSDARG ENSDARG00000037840 000000378 -0,06 +1,20 8,53E-01 1,72E-04 MR 40 ENSDARG NSCLC, ENSDARG00000035750 000000357 -1,42 -0,16 9,64E-04 5,54E-01 PSP 50 ENSDARG00000042904 foxo3b +1,17 +0,46 9,56E-04 5,51E-02 NSCLC ENSDARG00000035559 tp53 -1,16 -0,78 3,37E-05 1,25E-03 NSCLC ENSDARG00000011312 stk3 +0,96 +0,34 6,87E-03 1,37E-01 NSCLC ENSDARG00000038505 polr2h -1,53 -2,01 3,52E-04 2,90E-03 RP ENSDARG ENSDARG00000017820 000000178 -1,35 -0,88 9,34E-05 1,13E-02 RP 20 ENSDARG ENSDARG00000006926 000000069 +1,32 +1,45 1,85E-02 1,44E-02 RP 26 ENSDARG00000058740 ube2r2 +1,18 +1,08 3,52E-03 8,77E-03 RP ENSDARG00000061368 klf13 +1,31 +1,68 1,75E-01 1,05E-02 RP ENSDARG00000017602 ccng2 +4,68 +2,41 6,22E-04 5,78E-02 PSP ENSDARG ENSDARG00000078084 000000780 +2,59 +0,89 2,89E-04 2,32E-01 PSP 84 ENSDARG00000020693 sesn1 +1,27 +0,46 2,16E-02 2,86E-01 PSP
Annex: Supplementary Information 177
Supplementary table 8. DEGs involved in inflammatory response affected by asip1. The values that fit criteria are underlined.
Log2-ER in Log2-ER p-Value in p-Value Accession ID Gene Name Females in Males Females in Males
ENSDARG00000007693 nfkbiab +2,77 +2,35 1,60E-02 4,45E-02 ENSDARG00000003963 traf4a +1,76 -1,12 5,79E-03 4,52E-01 ENSDARG00000038964 traf4b -1,03 -2,15 9,44E-01 1,99E-02