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Barros 2011 Bastos FIN DEVELOPMENT Ana Maria FUNCTIONAL ANALYSIS OF ODD‐SKIPPED RELATED GENES: INSIGHTS FROM RENAL AND

Functional Comparative Analysis of Odd-skipped related genes : insights from renal and fin development 2011 2011 Porto Ana Neto Maria Barros Bastos

UNIVERSIDADE DO PORTO INSTITUTO DE CIENCIAS BIOMEDICAS DE ABEL SALAZAR

FUNCTIONAL ANALYSIS OF ODD-SKIPPED RELATED GENES: INSIGHTS FROM RENAL AND FIN

DEVELOPMENT

Ana Maria Bastos Barros Neto

Tese de doutoramento em Ciências Biomédicas 2011

Ana Maria Bastos Barros Neto

FUNCTIONAL COMPARATIVE ANALYSIS OF ODD- SKIPPED RELATED GENES: INSIGHTS FROM RENAL AND FIN DEVELOPMENT

Tese de Candidatura ao grau de Doutor em Ciências Biomédicas submetida ao Instituto de Ciências Biomédicas de Abel Salazar da Universidade do Porto.

Orientador – Doutor José Luís Gomez Skarmeta

Investigator Principal, Andalusian Center for Developmental Biology, CSIC-Universidad Pablo de Olavide, Seville, Spain

Co-orientador - Professor Doutor Cláudio Sunkel Professor Catedratico, Instituto de Ciências Biomédicas Abel Salazar, da Universidade do Porto, Portugal. Co-orientador - Fernando Casares Investigator Principal, Andalusian Center for Developmental Biology, CSIC-Universidad Pablo de Olavide, Seville, Spain

Este trabalho foi financiado pela Fundação para a Ciência e Tecnologia.

De acordo como disposto no 2o Artigo 8 o do Decreto Lei no 388/70, nesta dissertação foram utilizados resultados das publicações abaixo indicadas. No cumprimento do disposto no referido Decreto-Lei o autor desta dissertação declara que interveio na concepção e execução do trabalho experimental, na interpretação dos resultados e na redacção dos manuscritos publicados ou em preparação, sob o nome de Neto A. :

Tena, J. J., Neto, A., de la Calle-Mustienes, E., Bras-Pereira, C., Casares, F. and Gomez-Skarmeta, J. L. (2007). Odd-skipped genes encode repressors that control kidney development. Dev Biol 301, 518-31.

Neto, A., Casares F. and Gomez-Skarmeta, J. L. (2011). The osr1 and osr2 genes are essential to trigger limb development by responding to retinoic acid and controlling wnt2 expression in the pronephric anlage (submitted).

Outras publicações:

Roman A.C., Gonzalez-Rico F.J., Molto E., Hernando H., Neto A., Vicente C., Ballestar E., Gomez-Skarmeta J.L., Vavrova J., White R.J., Montoliu L., Fernandez-Salguero P.M. (2010) Dioxin receptor and Slug transcription factors regulate the insulator activity of B1 SINE retrotransposons via an RNA polymerase switch. Genome Res Mar;21(3):422-32.

Molina M.D., Neto A., Maeso I., Gómez-Skarmeta J.L., Saló E., Cebrià F. (2010) Noggin and noggin-like genes control dorsoventral axis regeneration in planarians. Current Biol. Feb 22;21(4):300-5.

ABSTRACT

Odd-skipped family of proteins (Odd in Drosophila and Osr in vertebrates) encode evolutionary conserved zinc-finger (ZF) transcription factors. In mammalians two odd- skipped related (osr) genes were found, Osr1 and Osr2. At the moment we start this study the function of the vertebrate proteins was completely unknown.

In the first part of this PhD project, we characterize the expression pattern of osr1 and osr2 genes in Xenopus and in zebrafish. We reported the role of osr genes in the development of renal structures (pronephros) in vertebrates. We found that osr1 and osr2 are both necessary and sufficient for the development of pronephros. Molecularly, osr proteins function as transcriptional repressors in renal development. We also showed that Drosophila odd induces the formation of kidney tissue in Xenopus. This ability is dependent of the Engrailed homology1 (eh1) domain. Odd genes may also be required for proper development of the Malpighian tubules, the Drosophila renal organs. Our results highlight the evolutionary conserved involvement of Odd-skipped transcription factors in the development of kidneys.

In the second part of this PhD project we set up to study the involvement of these genes in pectoral fin development, unravelling their place in the genetic cascade. Osr genes are expressed in the pectoral fin domain around 48hpf onwards. We found, by loss of function experiments, that osr genes are required for the formation of the pectoral fin bud. Strikingly, effect of loss of function is traced back to the 21 somites stage, affecting the expression of the earliest fin/limb specification marker, tbx5. We hypothesized that osr, expressed in the intermediate mesoderm, might act indirectly in the lateral plate mesoderm. We found that osr1 and both osr genes are able to regulate wnt2ba expression, a diffusible molecule, with a described role in fin development. Moreover, RA signalling is able to modulate osr genes expression, this controlling kidney and fin formation. Taken together, these results bring up a novel function of osr genes in the early steps of fin development.

In collaboration with Rankin and colleagues, we revealed the requirement of osr genes in the development of some endoderm structures such as lung and liver. The involvement of osr genes in the formation of endoderm derivatives as well as the molecular signalling, using wnt2b, seem to be conserved in Xenopus and zebrafish.

i RESUMO

A família de proteínas “Odd-skipped” é denominada como Odd na mosca da fruta (Drosophila) e em vertebrados por Osr, e são dedos de zinco altamente conservados durante a evolução. Em mamíferos dois genes “odd-skipped related” (osr) foram descobertos, Osr1 and Osr2. No princípio deste estudo a função das proteínas osr era totalmente desconhecida.

Na primeira parte deste projecto de doutoramento, o padrão de expressão dos genes osr1 e osr2 foram caracterizados em Xenopus e no peixe zebra. Neste estudo, a função destes genes foi descrita no desenvolvimento de estruturas renais (pronefros) em vertebrados. Podemos concluir que ambos genes, osr1 e osr2, são necessários e suficientes para o desenvolvimento de pronefros. As proteínas osr actuam a nível molecular como repressores da transcrição durante o desenvolvimento renal. Também demonstrámos que a proteina odd de Drosophila tem a propriedade de induzir a formação de tecido renal em Xenopus. Esta capacidade é dependente do domínio “Engrailed homology1“ (eh1). Os genes Odd podem tambem ser necessários para o correcto desenvolvimento dos tubulos de Malpighi, os orgãos renais da Drosophila. Os resultados apresentados neste trabalho ressaltam a conservação a nivel evolutivo dos factores de transcrição “Odd-skipped” no desenvolvimento renal.

Na segunda parte deste projecto de doutoramento proposemos o estudo da função destes genes no desenvolvimento das barbatanas peitorais, clarificando o lugar destes genes na via de sinalização. Os genes osr genes são expressos no domínio das barbatanas peitorais a partir de cerca de 48 horas depois da fertilização. Neste estudo demonstrámos por experiências de perda de função que os genes osr são necessários para a formação do primórdio das barbatanas peitorais. O efeito da perda de função é detectado no estadio de 21 somitos, afectando a expressão do marcador mais precoce de especificação da barbatana/membro, tbx5. Neste contexto os genes osr s ão expressos na mesoderme intermédia e poderiam actuar indirectamente na placa de mesoderme lateral. Neste trabalho descrevemos que osr1 e ambos genes osr regulam a expressão de wnt2ba, que é uma molécula difusível e com uma função já descrita no desenvolvimento das barbatanas. A sinalização do ácido retinóico é capaz de regular a expressão dos genes osr, e consequentemente a formação dos rins e das barbatanas. Em conjunto estes resultados demonstram a importância dos genes osr nas primeiras etapas de desenvolvimento das barbatanas.

ii Em colaboração com Scott Rankin e o grupo de Aaron Zorn, estudámos a importância dos genes osr no desenvolvimento de estruturas derivadas da endoderme como o pulmão e o fígado. A função dos genes osr na formação de derivados da endoderme assim como a sinalização molecular, via wnt2b, parece estar conservada em Xenopus e no peixe zebra.

iii

GENERAL INTRODUCTION

2

III. GENERAL INTRODUCTION

1. Principles of Developmental Biology

In 2002 Lewis Wolpert defined Developmental Biology as “the core of all biology, dealing with the process by which the genes in the fertilized egg control cell behaviour in the embryo and so determine its pattern, its form, and much of its behaviour” (Adapted from Wolpert L., 2002). Therefore, one of the key questions addressed by Developmental Biology is how different cells acquire unique morphologies and functional properties in diverse organs and tissues of the body, starting with an identical gene pool (Carroll, 2001)(Fig. 1). Remarkable advances in cell and molecular biology have allowed an exponential progress in Developmental Biology during the last years, resulting in the generation of an enormous volume of information (Wolpert L., 2002).

Currently, these complex developmental processes are being explained at the genomic level through the selective expression of distinct subsets of the many thousand of genes. However, the features of the complex regulatory program that controls gene expression in different cells over the course of animal development are only now starting to be unravelled. Most genetic networks involved in the development of certain tissues and organs remain largely unknown (Carroll, 2001).

Figure 1 - Differentiation of cell types of the vertebrate body. Development of a multicellular organism begins with the formation of a single cell, the zygote, when a sperm fertilizes an egg. The zygote has the potential to form an entire organism. In the first hours after fertilization, this cell divides into identical cells. These cells then begin to specialize, forming a hollow sphere of cells, called a blastocyst. Three germ layers can be identified in the blastocyst: ectoderm, light blue; mesoderm, light green; and endoderm, light yellow. The coloured panels illustrate some of the differentiated cell types formed from the zygote, including nerve cells, muscle cells, skin cells, blood cells, bone cells, and cartilage. Adapted from

3 http://www.ncbi.nlm.nih.gov/About/primer/genetics_cell.html

In this thesis we contribute to the understanding of these genetic networks by characterizing the function of odd-skipped related (osr) genes function in kidney and appendages development, using as model systems Xenopus and zebrafish.

2. Embryogenesis of Xenopus and zebrafish: useful models for developmental questions

Over the years several animal models have been used to address a great variety of questions in the Developmental Biology field. The choice of the model system used depends on the subject of study, availability of techniques and phylogenetic position, among others parameters (Drummond, 2000; Jones, 2005).

From the diversity of model systems, two vertebrates organisms that are used frequently and possess great advantages to address developmental questions are Xenopus and zebrafish (Danio rerio). The current knowledge of their genomes provides a great amount of information about the relation between genes, their genomic organization, and the degree of conservation during evolution, creating new areas of study, such as functional genomics that feed Developmental Biology with new data (Hellsten et al., 2010) (http://www.sanger.ac.uk/Projects/D_rerio/).

From a practical point of view, Xenopus and zebrafish are both very amenable for developmental studies since they combine a number of experimental advantages, such as high fecundity, easy manipulation of embryos with fast and external development (Fig. 2 and 3) and the use of conventional staining techniques. Moreover, the effect of drugs can be tested in vivo by adding them directly into the medium where the embryos grow (Drummond, 2000; Jones, 2005; Wingert and Davidson, 2008). A great diversity of functional techniques have been developed for the manipulation of genes, allowing the study of their specific role in development. In both models, the knockdown of a gene can be obtained by the microinjection of gene-specific morpholinos. Morpholinos are modified antisense oligonucleotides that lead to downregulation of a specific gene function. They act using two distinct strategies: blocking translation of the desired mRNA, when are designed to be complementary to the translation initiation sequence of the target gene, or preventing RNA processing, when designed to be complementary to a specific exon-intron splicing sites (Nasevivious and Ekker, 2000). Apart

4 from the use of morpholinos, in frogs and fishes, it is also possible to express ectopically a gene by direct microinjection of its mRNA, while the injection of a dominant-negative version of the mRNA of interest antagonizes its expression (Drummond, 2005; Jones, 2005). All these techniques are very useful and are based in the microinjection of different molecules in early embryos, which can cause general early embryogenesis effects, without a temporal or spatial resolution.

Figure 2 - Developmental events of the Xenopus life cycle. The major steps in development are represented: fertilization, cleavage, gastrulation, where the different germ layers are formed, neurulation during which organogenesis occurs and metamorphosis that marks the last step before the formation of the reproductive adult (Modified from Mereau et al., 2007).

To avoid this drawback, one strategy to control the timing of action is the use of mRNAs with hormone-inducible domains (Kolm and Sive, 1995). In this case, the fusion protein only will be activated when the hormone is added to the medium. In order to increase the resolution in space and time, other techniques were developed such as the generation of mosaics (zebrafish) or grafts (Xenopus) that allow the ectopic expression or gene knock- down in groups of cells (Carmany-Rampey and Moens, 2006; Grainger, 2000). Additionally, and with the same purpose, inducible heat-shock promoters or the GAL4/UAS system can also be used in combination with transient DNA injections and transgenic strains (Jones, 2005; Udvadia and Linney, 2003).

5 cell

yolk

(D) 25 hpf (A) 1 cell (B) bud

(C) 21-somite

(E) 35 hpf

(F) 72 hpf

Figure 3 - Diagram illustrating several stages of the zebrafish development. (A) Immediately after fertilization, two structures can be identified in 1-cell stage embryo: the yolk that represents 4/5 of the embryo and on top of the yolk, the cell. Developing at standard temperature of 28.5oC, at 10 hours post fertilization the tail bud is prominent (B). At the 21-somites stage (C), 19.5 hours after fertilization, structures such as the pronephros, the lens and the otic vesicle are already formed. The development of zebrafish is very fast and about 24 hours post-fertilization the majority of the organs are already formed (D). Around 35 hours post-fertilization, the pectoral fin is visible (arrow). At 72 hours after fertilization the zebrafish embryo hatches from the chorion (not represented here) that surrounds the embryo since 1-cell stage and becomes a free-swimming larvae. This organism reaches sexual maturity after 3 months of development (Modified from Kimmel et al., 1995)

In addition to their similarities as animal models, Xenopus and zebrafish have also specific advantages. Thus, zebrafish reaches its sexual maturity in three months, while X. tropicalis and X. laevis require six and twelve months, respectively (Fig. 2). During zebrafish meroblastic division, only a small portion of the egg is used to build the embryo, and all the cells remain connected until 8-cell stage allowing the microinjected molecule, morpholinos and/or mRNA, to diffuse freely through all the cells of the embryo. Moreover, zebrafish embryos are transparent allowing direct visualization of developmental processes and the in vivo dynamic expression of fluorescent reporter genes. One technical alternative to the use of

6 morpholinos in zebrafish are short interference RNAs (siRNA) that are also able to efficiently knock-down gene expression (Boonanuntanasarn et al., 2003; Dodd et al., 2004).

The fact that these small freshwater tropical fishes can originate a huge number of offspring, usually fertile after the exposure to random mutagenic agents, makes them suitable for mutagenesis screenings (Stemple and Driever, 1996). In 1996, Nusslein-Volhard’s and Fishman’s laboratories joint their efforts in a large scale mutagenesis screen in embryos. The resulting data were presented in a special issue of Development (Development. 1996 Dec; 123), in which the generation and characterization of several embryonic lethal mutations were described. This pioneer work fed the developmental biology field with a very useful collection of mutations that affect various systems, such as: fin (Amsterdam et al., 2004; Haffter et al., 1996; van Eeden et al., 1996), cardiovascular system (Chen et al., 1996; Stainier et al., 1996), gut, liver (Pack et al., 1996), blood cells (Ransom et al., 1996), and kidney (Drummond et al., 1998), among many others (Sprague et al., 2006). All the advantages described above make zebrafish a very useful vertebrate model for classical genetic approaches to study organogenesis.

Xenopus, on the other side, possesses several features as a model system, which are advantageous for some developmental studies. Xenopus cleavage is holoblastic where all the egg divides. The first cleavage originates two cells, one gives rise to the right side of the embryo and the other to the left side, without a significant mix of contents during the following divisions throughout development. This specific property makes possible the specific targeting of one side of the embryo by co-injecting mRNA and/or morpholinos with a lineage tracer (GFP, β-galactosidase, fluorescein) in one of the two cells. Therefore it is possible to compare the effect of the exogenous molecule with the uninjected contralateral control side (Fig.4A). Moreover, these early cleavage divisions divide the embryo into regions that are fated to become specific structures. Microinjection of molecules into individual blastomeres can direct its delivery with high probability to specific developing organs, for example, at 16- cell stage the blastomeres V2.2 contribute mainly to the formation of the pronephric tissue (Fig.5). The easy and directed microinjection of Xenopus embryos together with the accessible evaluation of the expression of specific gene markers, make these animals excellent model systems to carry out large-scale screenings (Grammer et al., 2000).

7 In situ analysis Uninjected (A) Expression screen contralateral control side mRNA Injected experimental side

Lineage tracer

!"#$ Epidermis (B) Animal Caps !#%&'()$

*"#$ *#%&'()$ Pronephros

Figure 4 - Techniques available using Xenopus embryos. (A) Co-injection of one cell from two–cell stage embryo with mRNA and/or morpholino and a lineage tracer (GFP in this case), allows the comparison between the injected/experimental side with the non-injected/control side of the same embryo. Injected embryos can be assayed for different probes, in this case lim1 probe was used, that labels the kidney domain. (B) Animal caps can be dissected with great accuracy from blastula embryos. The requirement of certain molecules for organogenic processes can be assayed. For example adding activin and retinoic acid at an appropriate concentration leads with high probability to the formation of the pronephric tissue (Modified from Jones, 2005).

!"#"$

Figure 5 - The Xenopus embryo suffers early cleavage divisions that originate blastomeres that are fated to become specific structures. Injection in specific blastomeres directs the molecule to the region of interest. For example, at 16-cell stage ventral vegetal cells, more precisely, blastomeres V2.2 are fated to contribute to pronephros formation (Adapted from Jones 2005, http://www.bio.davidson.edu).

8 Other manipulative techniques in Xenopus embryos are animal caps, grafts and explants studies that are very useful for the definition of the timing of requirement of different factors for the distinct processes of organogenesis (Brennan et al., 1999; Okabayashi and Asashima, 2003) (Fig.4B).

Although, the pseudo-tetraploid Xenopus laevis is usually the experimental amphibian of choice, its “cousin” Xenopus tropicalis with a diploid genome increases the possibilities of successful genetic screens (Grammer et al., 2000; Jones, 2005), and also has an increased probability of morpholino efficiency when compared with X. laevis or even with zebrafish, whose genome underwent duplication originating several paralog genes with possible redundant functions. Moreover, X. tropicalis allowed the identification of recessive mutations with a wide range of developmental defects (Grammer et al., 2005; Noramly et al., 2005).

In this thesis, we took advantage of these models to functionally characterize odd-skipped related (osr) genes in different developmental processes. To study the role of these genes in kidney development we used both model systems: Xenopus and zebrafish, which share similar developmental mechanisms in kidney formation. These models are very amenable for this study, due to the diversity of technical approaches available, but also due to their external development. Both models have simple excretory systems, the pronephros. These structures have striking similarities regarding the cell types and the patterning mechanisms with the mammalian metanephric kidney (Vize et al., 1997). Therefore, the knowledge acquired in these models can be then extrapolated to the understanding of more complex systems.

To study the role of osr genes in fin/limb development we used exclusively zebrafish. Xenopus is not a very practical model for this study. The Xenopus limb buds appear quite late in development (stage 44) where the action of morpholinos is difficult to relate to a specific limb phenotype (Tarin and Sturdee, 1971). Zebrafish was chosen as our model system due to the fast development of pectoral fin structures, which at 24 hours post fertilization are already recognizable, and also this simple vertebrate possesses a great variety of genetic tools advantageous for the study the genetic hierarchy of genes involved in fin formation (Kimmel et al., 1995). On the other hand, there are astonishing similarities at the molecular and anatomical level between the zebrafish-paired fins and the tetrapod limbs. The high degree of homology between these processes implies that, at least, part of the data obtained with zebrafish can be applied to other vertebrates, keeping in mind that there are particular features that might be important for the diversity of vertebrate appendages (reviewed in Mercader, 2007).

9 3. Early steps of kidney development

The kidney is the organ that performs the essential functions of blood filtration and osmoregulation (Howland, 1921; Tytler, 1988; Tytler, 1996; Vize et al., 1997). Early kidney development is an ideal system to explore several processes such as cell specification, morphogenesis, and organogenesis, among others. Advances in the understanding of kidney development can give some clues for stem cell renewal, segmentation and boundary formation, signalling pathways and disease mechanisms (Dressler, 2009).

In vertebrates, the intermediate mesoderm (IM), which lies between the somites and the lateral plate, will originate three types of kidneys of increasing complexity: pronephros, mesonephros and metanephros (Adapted from Brandli, 1999; Vize et al., 1997). These renal structures form sequentially through an inductive process that requires the formation of the previous structure. These kidneys have a different organization but they share the same structural unit: the nephron (Zhou et al.) (Fig. 6). The nephron is composed of three fundamental parts: a glomerulus used for filtration, a tubule where reabsortion and secretion of solutes occurs, and a collecting duct that transports the resulting solution to be eliminated (Mobjerg et al., 2000; Richards, 1929).

Figure 6 - Organization of a (non-integrated) nephron. Blood travels from the dorsal aorta to the glomus where it suffers selective filtration. Wastes filtered into the coelom are swept into kidney tubules by nephrostomes, which are thin ciliated funnels. Reabsorbed solutes are returned to the bloodstream via blood sinus (purple), which derives from the posterior cardinal vein that surrounds the tubules. The remaining excretory products pass along the pronephric duct, which is connected to the cloaca (Dressler, 2006).

3.1 Embryonic kidney structures in vertebrates

In fish and amphibians, the pronephros is the embryonic functional kidney, and is formed by a pair of bilateral non-integrated nephrons (Brandli, 1999; Burns, 1955; Drummond, 2003; Jones, 2005; Reimschuessel, 2001; Vize et al., 1997). Later, during larval life, the surrounding mesenchyme forms additional nephrons resulting in the formation of the

10 mesonephros, which are the adult kidney. In zebrafish mesonephros, the number of nephrons can reach to around 150 nephrons (Adapted from Vize et al., 1997), whereas the adult amphibian mesonephros is formed by around 2500 nephrons (Raciti et al., 2008; Reggiani et al., 2007; Tytler, 1988; Wingert and Davidson, 2008; Wingert et al., 2007; Zhou and Vize, 2004). In these organisms the metanephros is never formed.

In amniotes, the pronephros is just a transient structure and the mesonephros is the embryonic functional kidney. Later, the metanephros replaces the previous structures that degenerate or become part of the male reproductive system (Adapted fromWingert and Davidson, 2008). Metanephros development results from an interaction between the ureteric bud and the surrounding mesenchyme leading to branching morphogenesis, originating a complex structure with millions of nephrons (Fig. 7). This structure becomes the functional kidney during adulthood (Brennan et al., 1998).

(A) (B) (C)

Figure 7 - Scheme of different types of kidneys in different vertebrates. (A) In fishes and amphibians, the pronephros is the embryonic kidney. (B) In these organisms, the mesonephros replaces the pronephros in the osmorregulatory function. (C) However, in mammals the mesonephros are the embryonic kidneys that later are replaced by the adult kidneys, the metanephros (Carroll et al., 1999b; Dressler, 2006; Ryffel, 2003).

11 The nephrons from different kinds of kidneys present not only structural similarities but also physiological homologies. Indeed, the same types of solute transporters are present in the same proximo-distal (P/D) regions not only in Xenopus and zebrafish pronephros but also in mammalian metanephros (Carroll et al., 1999a; Heller and Brandli, 1999) (Fig.8).

(A) (B) (C)

Figure 8 - The segmentation pattern is shared by pronephric and metanephric nephrons. (A) Mammalian metanephric kidney. The enlargement shows a description of the segmental organization of only one nephron. (B) Lateral view of a zebrafish embryo where pronephros is indicated in blue. A dorsal view of the pronephros is shown in the enlargement, with indications of the segmental organization of each nephron. (C) Lateral view of a Xenopus embryo with the pronephros represented in blue. Enlargement details a lateral view of a single nephron. The color-coding of analogous segment identities used for all nephrons is based on comparative expression pattern of genes. Abbreviations: ATL, ascending thin limb; C, cloaca; CD, collecting duct; CNT, connecting tubule; CS, corpuscle of Stannius; DCT, distal convoluted tubule; DE, distal early; DL, distal late; DTL, descending thin limb; G, glomus; MD, macula densa; N, neck; Ne, nephrostome; P, podocytes of renal corpuscle; PCT, proximal convoluted tubule; PD, pronephric duct; PE, proximal early; PL, proximal late; PST, proximal straight tubule; T, tubule; TAL, thick ascending limb (Carroll and Vize, 1999; Heller and Brandli, 1999).

3.2. Genes involved in pronephros development

Pronephros development can be divided into three recognizable steps: specification, which occurs during gastrulation (Carroll and Vize, 1999; Chan et al., 2000); morphogenesis, triggered by the differentiation of different precursor regions of the nephron (tubules, duct and glomus); and segmentation, characterized by the specialization of different parts of the nephron along the proximo-distal (P/D) axis (Carroll et al., 1999a; Raciti et al., 2008). Functional studies in different vertebrates indicate that most of the genes essential for the formation of the simplest kidney, the pronephros, are also required for the formation of the mesonephros and the metanephros (Bouchard et al., 2002; Kobayashi et al., 2005; Pedersen et al., 2005; Porteous et al., 2000; Shawlot and Behringer, 1995; Torres et al., 1995).

In Xenopus, Xlim1 and Xpax8 were described as the first specification genes expressed in the prospective pronephric field (Maden, 1999; Ruiz i Altaba and Jessell, 1991). These

12 genes are transcribed in the intermediate mesoderm at early neurula stage, prior to any morphological signal of kidney formation (Cartry et al., 2006; Grandel and Brand; Wingert et al., 2007) Xlim1 and Xpax8 are both essential for pronephros formation (Duester, 2007; Keegan et al., 2005; Stafford and Prince, 2002). Indeed only co-expression of Xpax8 (or the partial redundant Xpax2) and Xlim1 is able to promote the formation of ectopic and enlarged patches of kidney tissue (Carroll and Vize, 1999).

The expression pattern of Xlim1 and Xpax8 in the intermediate mesoderm, fated to give rise to the pronephros, is conserved from zebrafish to mammals (Barnes et al., 1994; Bouchard et al., 2002; Fujii et al., 1994; Pfeffer et al., 1998; Plachov et al., 1990; Shawlot and Behringer, 1995; Toyama and Dawid, 1997; Tsang et al., 2000). In zebrafish, pax2.1, pax8 and lim1 are expressed since early stages of development in the prospective pronephros domain and at least the role of pax2.1 has been described to be fundamental for proper nephron patterning and the definition of the nephron tubule-glomerulus boundary (Majumdar et al., 2000; Pfeffer et al., 1998; Toyama and Dawid, 1997). Mice mutant for Lim1 or for both Pax2 and Pax8 show strong impairment in the formation of pronephros or any nephric structure (Chen et al., 2001; Hollemann et al., 1998). In double mutants for Pax genes, the intermediate mesoderm does not undergo a mesenchymal-epithelial transition required for the formation of the nephric duct and consequently fail to exhibit any nephric gene expression (Bouchard et al., 2002; Mansouri et al., 1999).

Retinoic acid (RA) is a molecule also described to be fundamental for the development of the vertebrate embryo. There are good evidences that RA is an essential signaling molecule for mesoderm patterning (Cartry et al., 2006; Grandel et al., 2002). In fact, the RA signaling pathway shapes different derivatives of mesoderm such as pronephros- and fin/limb- precursors (Brandli, 1999) but also derivatives from other origins (Brandli, 1999; Wessely and Tran, 2011). The gradient of RA results from the expression profile of its synthesizing (Raldh2) and degradating (Cyp26) enzymes. This gradient modulates gene expression in the mesoderm during gastrulation and early neurula, when pronephros specification occurs (Carroll and Vize, 1996; Majumdar et al., 2000). Treatment of Xenopus blastula animal caps with activin A and RA leads to the formation of kidney tissue that recapitulates the normal pronephric development at the molecular level, expressing the correct differentiation markers (Drummond et al., 1998). More recently, in vivo experiments demonstrated that RA signaling is required for pronephros specification and the regulation of the expression of Xlim1 and Xpax8 in the kidney anlage (Cartry et al., 2006). In mice mutant for Raldh2, the specification

13 of the pronephric lineage is severely compromised and the prospective mesonephros formation is impaired (Cartry et al., 2006; Fujii et al., 1997; Niederreither et al., 1997). In RA attenuated/ depleted zebrafish embryos the anterior tip of the pronephros, marked by pax2.1 expression, is reduced (Grandel et al., 2002). The similar kidney phenotype produced by the impairment of RA signalling in different models reflects the conservation of the role of RA in the early steps of kidney formation among vertebrates (Brandli, 1999; Saulnier et al., 2002).

After specification of the pronephros anlage, two major steps can be recognized in the formation of this structure such as morphogenesis and nephron patterning (Raciti et al., 2008; Zhou and Vize, 2004). These processes require the combined activity of Wnt, BMP, Hedgehog and Notch signalling cascades (Kriz and Bankir, 1988; Mobjerg et al., 2000). The pronephric region defined by Xlim1 and Xpax8 extends through the precursors of the tubules, duct and the glomus. During morphogenesis, this original pronephric domain is subdivided by additional patterning signals that define a medial region that will then form the glomus and establish distinct dorsolateral domain that will form the tubules and ventrolateral domain that will form the duct. For example, the expression of wt1a is localized in the medial pronephric mesoderm in Xenopus and zebrafish (Vize, 2003b). Its expression labels prospective glomus and promotes the restriction of mesoderm genes to the tubules and duct of the pronephros (Christensen et al., 2008; Eid et al., 2002; Raciti et al., 2008; Zhou and Vize, 2004; Zhou and Vize, 2005). Formation of the pronephric tubule is a process that requires changes in cell shape and extensive cell rearrangements. These processes are very similar to the ones occurring during the metanephric mesenchymal-epithelial conversion, and there are also homologies at the molecular level (Raciti et al., 2008; Reggiani et al., 2007).

3.3. Proximo/Distal (P/D) Patterning of the nephron

The patterning of the nephron along the P/D axis results in its organization into proximal tubule, intermediate tubule and distal tubule, which connect with the collecting duct (Raciti et al., 2008; Vize, 2003a; Zhou and Vize, 2004; Zhou and Vize, 2005). Moreover, distinct segments can be identified in each tubule based on histological characteristics (Adapted from Reggiani et al., 2007) and gene expression profiles (Van Campenhout et al., 2006).

The segments are formed by specialized epithelial cell types, which perform precise functions such as solute transport, pH regulation and water absorption. Three domains are

14 considered in the proximal tubules, PT1, PT2 and PT3, with functions in the reabsortion of ions, amino acids, glucose and water (Reggiani et al., 2007) (Fig.9). The intermediate tubules show two domains, IT1 and IT2, which share some markers with the distal tubule, and are functionally adapted to the reabsorption of salt and ions (Alarcon et al., 2008; Reggiani et al., 2007; Van Campenhout et al., 2006). The distal tubules composed by two domains, DT1 and DT2, perform several roles such as reabsorption of magnesium ions, urine acidification and ammonium transport (Deconinck et al., 2000; Tran et al., 2007; Weber et al., 1996) (Fig.9).

Figure 9 –Segments of the tubular portion of the Xenopus pronephric kidney are represented schematically. This model illustrates a pronephric kidney from stage 35/36 embryo. The four tubular compartments have a specific color. Subdivisions can be recognized in each tubule: proximal tubule (yellow; PT1, PT2, PT3), intermediate tubule (green; IT1, IT2, IT3), distal tubule (orange; DT1, DT2), and connecting tubule (gray; CT). The connection between the nephron and the coelomic cavity is made by nephrostomes (NS), which are ciliated peritoneal funnels (Grote et al., 2008; Grote et al., 2006).

Several molecular markers allow the identification of specialized segments in the nephron indicating a possible functional role. One example is Xevi1 a transcription factor that is expressed in the distal tubule and the pronephric duct (Cheret et al., 2002; Pontoglio et al., 1996). The Iroquois genes, Xirx1, Xirx2 and Xirx3, are expressed in other domains of the nephron such as proximal tubule segment (PT3) and intermediate tubule (IT1 and IT2) (Pontoglio et al., 2000; Tanaka et al., 2010). Functional studies associated Xirx1 and Xirx3 genes with a role in the specification of the segments where they are expressed (Kobayashi et al., 2005). Moreover, other transcription factors were identified as markers of pronephric duct and proximal tubules, Xgata3 and Xhnf1α, respectively (Nakai et al., 2003). Indeed the function of both genes was studied in mice. Gata3 was characterized as a regulator of

15 proliferation and guidance of the nephric duct (Colas et al., 2008; Grieshammer et al., 2005; Urban et al., 2006) and Hnf1α was described as a regulator of specific proximal tubular transporters (Bracken et al., 2008), controlling the renal glucose re-absorption (Cheng et al., 2007; Cheng et al., 2003; McLaughlin et al., 2000; Naylor and Jones, 2009; Taelman et al., 2006; Wang et al., 2003; White et al., 2010).

The transcription factor Lim1, already mentioned in the specification of the kidney anlage, is also necessary for the formation of the proximal segments, probably acting as upstream regulator of the Notch signalling (Coates, 1995). In mice, the transcription factor Brn1/ Pou3f3 seems to be required for the specification of the more distal nephron segments (Tanaka M., 2007).

Other players are necessary for the development of the pronephric tubule and duct such as Fgf8 (Tanaka M., 2007). This molecule is important in later tubule development, particularly regulating tubule epithelialization and is also involved in cell survival during nephrogenesis. BMP signalling is also involved in the differentiation of the pronephric tubules and duct but not in glomal development (Burke et al., 1995; Gaunt, 2000).

Several components of the Notch signalling are involved in the specification of a subpopulation of cells in the pronephros. This pathway specifies proximo-distal fates and one of its components, Notch2, is required for proximal differentiation (Cohn and Tickle, 1999).

4.Fin/Limb development

More than 400 million years ago, paired locomotion appendages appeared in the gnathostome lineage (jawed vertebrates). These structures are commonly known as fins in fishes and limbs in tetrapods. In all species known, the anterior set of appendages is located at the level of the heart and the posterior one is located at the level of the hindgut. During the “water to land transition” fins evolved into limbs in stem tetrapod groups presenting, as major morphological novelties, digits and wrists (Deschamps et al., 1999). Interestingly fins and limbs have been highly conserved since their origin in terms of position, morphology and development. These may reflect a tight conservation of the molecular networks involved in their formation (Capdevila and Izpisua Belmonte, 2001). In this section we will review the fin/limb developmental events with emphasis in the early steps of this process.

16

4.1 Fin/Limb development in different vertebrate models

The three vertebrate models most commonly used to study of fin/limb development are zebrafish, chicken and mouse. In all these models the early formation of the fin/limb bud is similar pointing to a common molecular mechanism of development established in their common ancestral (McPherron et al., 1999).

4.2 Fin/Limb positioning along main body axis

The positional identity along the embryonic anterior/posterior (A/P) axis is conferred by a specific and combinatory expression of Hox genes that have different levels of expression along the lateral plate mesoderm (LPM). The anterior expression of Hoxc6, Hoxc8, Hoxb5 and Hox9 genes coincides with forelimb/pectoral fin level (Nelson et al., 1996; Oliver et al., 1990; Rancourt et al., 1995). In fact, Hoxb5 mutants have a shift in the shoulder girdle and Hoxd9 mutants have forelimb malformations (Fromental-Ramain et al., 1996; Rancourt et al., 1995). Interestingly, differences in the position of the forelimb between chicken and mouse show a relation with changes of Hox genes expression domains (reviewed in Deschamps et al., 1999). Moreover, in snakes the expansion of Hox expression domains in both paraxial and LPM gives to the entire trunk a “flank identity” that correlates with the absence of limbs (Cohn et al., 1997).

Deschamps considered that “Hox expression is still labile while being established“ (Cohn et al., 1997). In the embryonic A/P axis, the control of Hox gene expression is exerted by a group of factors, namely retinoid acid receptors (RARs), Cdx/Caudal proteins, members of the PBX/Exd family of cofactors, Hox genes themselves, and members of FGF and TGF-β signalling (Capdevila and Izpisua Belmonte, 2001). Interaction of Hox genes with these different regulators results in the pre-patterning of the embryonic axis, contributing for the allocation of the limb fields in a variety of vertebrates. Mice with the TGFβ factor Gdf11 functionally inactivated show a posterior displacement of the hindlimbs. This change in the limb position is associated with changes in the axial Hox gene expression, suggesting that Gdf11 acts upstream of Hox genes to define positional identity along the A/P axis (Capdevila and Izpisua Belmonte, 2001; Johnson and Tabin, 1997; Tickle, 1999).

The positioning of the limbs along the main body axis is modulated by the expression of fibroblast growth factors (FGFs, (Agarwal et al., 2003; Begemann and Ingham, 2000;

17 Ruvinsky et al., 2000; Saito et al., 2006; Tamura et al., 1999)). Application of FGF beads in the interlimb region lead to the development of ectopic limbs in chicken (Naiche and Papaioannou, 2003; Takeuchi et al., 1999). This change of identity of flank cells towards limb cells displaces the limit of Hox9 expression in the LPM (Agarwal et al., 2003; Ahn et al., 2002; Garrity et al., 2002; Minguillon et al., 2005; Ng et al., 2002; Rallis et al., 2003; Takeuchi et al., 2003).

4.3 Fin/limb induction and initiation

Fin and limb initiation is characterized by a thickening of the somatopleura portion of the lateral plate mesoderm (LPM). This process implies specification of a particular group of cells on either side of the embryo at precise positions along the A/P axis (Mercader, 2007; Ng et al., 2002). Then coordinated signals for proliferation are maintained exclusively in these cells that will form the fin/limb primordia (Begemann et al., 2001; Gibert et al., 2006; Mic et al., 2004). These initial processes seem to be highly conserved in all models studied.

The earliest molecular marker of limb bud formation is the T-box transcription factor Tbx5, expressed in anterior appendage precursor cells in all vertebrate studied, from sharks to mice (Grandel and Brand). The location of the posterior appendages is marked by the expression of other member of the T-box family, Tbx4 (Grandel and Brand). Tbx5 is both necessary and sufficient for forelimb formation (Grandel and Brand). In zebrafish, tbx5 mutant heartstrings (hst) and the embryos injected with tbx5 antisense morpholino show loss of pectoral fin formation (Ahn et al., 2002; Garrity et al., 2002; Ng et al., 2002). The cells of the LPM expressing tbx5 are not able to migrate, falling to aggregate to form the compact circular structure of the wild type fin bud (Ahn et al., 2002; Garrity et al., 2002). Moreover, inactivation of Tbx5 or Tbx4 led to a limbless phenotype, in mice and chicken (Agarwal et al., 2003; Naiche and Papaioannou, 2003; Rallis et al., 2003; Takeuchi et al., 2003).

Studies in zebrafish indicate that tbx5 is an upstream regulator of fgf10 (Ng et al., 2002). This molecule is then crucial for the formation of the Apical Ectodermal Ridge (AER, see next section) and consequent fin/limb outgrowth. Interestingly fgf10 also contributes for the maintenance of tbx5 expression (Emoto et al., 2005). Surprisingly, it has been observed that transplantation of somitic cells, from the neighbourhood of the forelimb field, to the hindlimb level lead to Tbx5 expression instead of Tbx4. Therefore, somitic signals should have some functions along the A/P axis to specify the anterior and posterior limb fields (Saito et al.,

18 2006). Candidate(s) to be this/these somitic signals should be diffusible factors, such as retinoic acid (RA), expressed in the anterior region and able to modulate gene expression.

Indeed recently, in zebrafish, RA was shown to be an upstream regulator of tbx5 during limb initiation (Sirbu and Duester, 2006). There is a repeated requirement of RA during the fin determination process (Capdevila and Izpisua Belmonte, 2001). RA signalling is active during gastrulation participating in the determination program of different pools of LPM derivatives (Keegan et al., 2005; Waxman et al., 2008). During somitogenesis, this signalling pathway maintains fin precursors and then controls fin bud growth (Geduspan and Solursh, 1992; Stephens and McNulty, 1981). As mentioned above (section 3.2) RA levels result from synthesis/degradation process (Negishi et al.). The null mutant for the degradating enzyme Cyp26 (giraffe) shows a fin bud in a position shifted anteriorly (Agarwal et al., 2003; Galceran et al., 1999). The medaka, zebrafish and mice mutants for aldh1a2 are not capable of RA synthesis during early development, resulting in the absence of fins/limbs, and also do not express tbx5. These phenotypes presented by the aldh1a2 mutants can be rescued by the exogenous application of RA or by transplantation of wild-type cells (Begemann et al., 2001; Gibert et al., 2006; Grandel et al., 2002; Linville et al., 2004; Mercader et al., 2006; Mic et al., 2004; Negishi et al.; Niederreither et al., 1999; Zhao et al., 2009).

Interestingly the source of RA acting in the LPM during limb development seems to be in the somitic cells (Martin, 1998). Once produced, this molecule has to cross the intermediate mesoderm (IM) in order to reach the LPM and promote fin/limb initiation. Classical experiments proposed the IM, where kidney precursors are located, as a key tissue required for limb induction (Cohn et al., 1995; Kawakami et al., 2001; Ohuchi et al., 1997; Ohuchi et al., 1995; Yonei-Tamura et al., 1999). When the IM is removed or its contact with LPM is physically blocked, limb formation is impaired (Min et al., 1998; Sekine et al., 1999). However, the role of IM in limb formation is still controversial, given that other authors argue that the IM is indeed dispensable for limb outgrowth (Bouchard et al., 2002; Fernandez-Teran et al., 1997)

The Wnt signalling pathway is also required for normal fin/limb outgrowth (Kawakami et al., 2001; Ng et al., 2002). Wnt2b is detected in the IM, adjacent to the Tbx5-expressing area, and is required, in zebrafish fin and chicken, for fins and forelimbs development, respectively. In zebrafish, wnt2b seems to act downstream of RA and upstream of the tbx5 during the early pectoral fin formation (Mercader et al., 2006; Ng et al., 2002). Recently, a homolog of wnt2b was identified in zebrafish, wnt2bb (Ober et al., 2006) and the previous known gene renamed

19 wnt2ba. The role of wnt2bb was described in liver specification and is also regulated by RA signalling (Fischer et al., 2003; Ng et al., 2002). Studies in chicken indicate that the hindlimb level, the Wnt gene involved in the activation of the T-box transcription factor Tbx4 is Wnt8c (Ng et al., 2002), both Wnt members act through the canonical pathway.

There seems to be some species-specificity in the function of Wnt pathway during the limb formation. In mice, the expression of Wnt2ba is not detected in the LPM (Ng et al., 2002). Moreover, Tcf-/-/Lef-/- mice, nuclear transducers of the Wnt pathway, develop very rudimentary limb buds although they still show Tbx5 expression (Galceran et al., 1999; Kawakami et al., 2001; Norton et al., 2005). These data indicates that Wnt signalling plays a role in mice limb initiation through other(s) component(s) than Wnt2b (Mercader, 2007; Ng et al., 2002). In contrast zebrafish and chicken both use Wnt2ba signalling. In addition, only chicken has a second phase of Wnt2ba expression detected in the early wing bud (Kawakami et al., 2001), which is required for Tbx5 maintenance (Ng et al., 2002; Takeuchi et al., 2003).

Interestingly, a novel secreted protein called Fibin was identified in zebrafish, mice and human genomes (Tanaka M., 2007; Wakahara et al., 2007). In zebrafish, its expression is detected in the LPM since 14hpf, between the somites and the presumptive pectoral fin area. In zebrafish, fibin seems to be active downstream of RA and wnt2ba signalling and upstream of tbx5. In mice, Fibin expression was described in the forelimb buds but its function is still unknown (Wakahara et al., 2007).

4.4 Formation of the Apical Ectodermal Ridge (AER)

After an initial growth of the fin/limb bud the Tbx5-expressing cells start to express Fgf10. This marks a new step on the development of the bud mainly characterized by the formation of the AER. The AER is basically a thickening of the most distal ectoderm that starts acting as a signalling centre for proper outgrowth and patterning. Several pathways are involved in the AER establishment, that implicate a feedback loop between mesenchyme and ectoderm: Fgf signaling (Bell et al., 2003; Kawakami et al., 2004; Treichel et al., 2003), Wnt pathway (Galceran et al., 1999; Norton et al., 2005) and BMP family of proteins (Ahn et al., 2001; Pizette et al., 2001).

4.5 Fin/Limb outgrowth

The outgrowth of limbs is known to be controlled by members of the Fgf pathway, in particular Fgf8 and Fgf10, as mentioned before (reviewed in (Adapted from Tanaka M.,

20 2007). Several works described increased proliferation and consequent formation of ectopic limbs with the application of Fgf beads in the flank of chicken and mouse (Camarata et al., 2006; Camarata et al.). Moreover, Fgf10 null mice show initiation of the limb bud formation, but the following step of outgrowth did not occur (Min et al., 1998; Sekine et al., 1999).

The expression of Fgf10 in the prospective limb mesenchyme promotes maintenance of LPM proliferation to form a bud and after triggers the expression of Fgf8 in the most distal ectoderm, the AER (Harvey and Logan, 2006). The AER then secrets Fgf8 to the adjacent cells maintaining them in a proliferate state and promoting limb outgrowth. Interestingly Fgf8 has the ability to maintain the expression of Fgf10 under the AER. Therefore these molecules act by a feedback loop regulatory mechanism (Ohuchi et al., 1997b).

In chicken Fgf10 expression is regulated by members of the Wnt pathway. Wnt2b and Wnt8c are able to restrict Fgf10 expression at, respectively, forelimb and hindlimb levels (Kawakami et al., 2001; Takeuchi et al., 2003). Tbx5 and Tbx4 are upstream of the second round of Wnt expression in the limb and Fgf10 in the genetic cascade that regulates limb induction (Ahn et al., 2002; Minguillon et al., 2005; Ng et al., 2002; Rallis et al., 2003; Takeuchi et al., 2003). Distinct mechanisms are described in zebrafish whereas wnt2ba is required for tbx5 expression that triggers fgf24 expression and then activates fgf10 (Lee and Roy, 2006; Mercader et al., 2006; Wilm and Solnica-Krezel, 2005) (Fig.10).

Lateral plate mesoderm Limb ectoderm

Wnt2b Wnt3a/3 Anterior Tbx5 paired appendages Fgf10 Fgf8

zebrafish Wnt2b(a) Tbx5 Fgf10

Posterior Wnt8c Wnt3a/3 paired Tbx4 appendages Fgf10 Fgf8

Figure 10 - Scheme of some of the key genes involved in fin/limb development. Tbx5 and Tbx4 activate Wnt2b/Fgf10 and Wnt8c/Fgf10 signals, respectively in anterior and posterior paired

21 appendages. The Wnt/Fgf signalling is able to feedback Tbx genes to maintain their expression. Fgf10 activates Wnt3a/3/Fgf8 signals in the limb ectoderm and induces AER formation. It should be noted that most of this data are from chick. Wnt2b and Wnt8c have not been described in mouse limb development. In zebrafish, wnt2b(a) is expressed in the intermediate mesoderm and acts upstream of tbx5, represented in the light blue boxes (Saunders J.W., 1968).

Interestingly, other distinct wnt3 molecules play a similar role in different organisms. These wnt3 molecules mediate fgf8 activation in the ectoderm and establish the positive FGF feedback loop between mesenchyme and ectoderm (reviewed by Tabin and Wolpert, 2007). Other molecules, which will not be described here, have been described to be required for proper outgrowth of the fin/limb. Some examples of that are pdlim7 (Riddle et al., 1993) , Trap 230/Med12 (Rau et al., 2006), sall4, sall1a (Summerbell et al., 1973), prdm1 (Mercader et al., 2000; Tabin and Wolpert, 2007) and histone deacetylase-1 (Pillai et al., 2004).

4.6 Patterning of Fin/Limb Bud

The vertebrate limb undergoes development and results in a highly organized structure whereas three different axes can be distinguished: The back of the hand (dorsal) is different from the palm (ventral)(D/V), the thumb (anterior) is distinct from the little finger (posterior) (A/P), and the upper arm (proximal) is different from the lower arm and the hand (distal) (P/D) (Tabin and Wolpert, 2007).

The zone of polarizing activity (ZPA) is a signalling centre localized in the posterior limb mesenchyme that acts organizing the polarity A/P of the limb bud (Cooper et al.; Rosello-Diez et al.). Several signalling molecules that are important for early processes of development are again required for the A/P patterning of the limb such as Sonic hedgehog (Shh) (Helms et al., 1996; Lopez-Martinez et al., 1995; Lu et al., 1997; Riddle et al., 1993; Stratford et al., 1996; Tickle, 2004; Yang et al., 1997), RA (Eichele and Thaller, 1987; Helms et al., 1996; Lu et al., 1997; Mic et al., 2004; Stratford et al., 1996; Yang et al., 1997) and Hox genes (Charite et al., 1994; Laufer et al., 1994; Niswander et al., 1994; Zakany et al., 2004). Shh mediates the polarizing activity of the ZPA (ten Berge et al., 2008). Grafting of an additional ZPA or of cells that express Shh results in similar mirror-image duplication (Helms et al., 1996; Lopez- Martinez et al., 1995; Lu et al., 1997; Riddle et al., 1993; Stratford et al., 1996; Tickle, 2004; Yang et al., 1997).

Two general models have been proposed to explain the specific patterning of the P/D axis of the limb, the “Progress Zone model” (Cooper et al.; Mackem and Lewandoski; Rosello-

22 Diez et al.) and the “Early Specification model” (Chen and Johnson, 1999; Niswander, 2003). The first postulates that progressive acquisition of distal identity occurs through an autonomous mechanism associated with the undifferentiated mesenchymal cells. These mesenchymal cells are immediately under the AER, where the cells are kept in a proliferative state through the action of this signaling centre. The acquisition of proximal or distal identity is regulated by an internal clock mechanism that tracks the time spent in this zone under the influence of distal FGF signals. On the other hand, the second model claims that the specification of the proximo-distal segments results from opposite gradients of proximal and distal-inducing factors. Late limb buds of all vertebrates express identity markers, such as Meis1, Hoxa11 and Hoxa13, which are correlated with the stylopod (proximal), zeugopod and autopod (distal), respectively. Although none of the segmental markers are required for P/D specification (Chen et al., 1998; Riddle et al., 1995; Vogel et al., 1995)(Fig.11).

Figure 11 – The limb bud progenitors are kept in an undifferentiated and proliferative state through the balance of Wnt and Fgf signaling. During outgrowth the differential exposure to proximal RA and distal Fgfs will result in the formation of the distinct limb segments. The progenitors of each segment will express specifically molecular markers, as observed in the image with Meis, Hoxa11 and Hoxa13 (Wang and Coulter, 1996).

23 Recent reports support the second model and show that initially the early limb mesenchyme is exposed to proximal and distal signals, such as Wnt3a, Fgf8 and RA, that maintains the cells in a competent state for the formation all the segments (Cygan et al., 1997; Logan et al., 1997; Loomis et al., 1996; Loomis et al., 1998). During limb bud growth, exposure of continued signals from the flank establish the proximal limb segment while the medial and distal segments are formed outside the range of the proximal signaling. In this context the proximal cells, without suffering the action of distal signals, are kept in an undifferentiated state (Ahn et al., 2001; Pizette et al., 2001). The action of proximal signals leads to differentiation of the cells adjacent to the flank resulting in the formation of proximal structures. As the limb grows, the distal mesenchymal cells are beyond the reach of the RA action produced at the proximal region, restricting the distal cells to the zeugopod and the autopod fates. The emerging model of P/D patterning proposes that a temporal dynamic balance between the proximal RA signal and the distal FGF levels regulates the proximo- distal identity of the limb bud during outgrowth and the progenitor pool is maintained by distal Wnt and Fgf signals beyond the RA proximal influence (reviewed by Niswander, 2003).

Experimental manipulations of the chick embryonic limb and genetic manipulation in mice unravelled the molecular network that regulates the limb D/V patterning (Adapted from Mackem and Lewandoski). Lmx1b is one of the players involved in this process, is expressed in the dorsal mesenchyme and is necessary for the acquisition of a dorsal fate. During limb bud formation Lmx1b expression is induced by Wnt7a, which is expressed in the dorsal ectoderm (Nusslein-Volhard and Wieschaus, 1980). The restricted expression of Wnt7a in the dorsal ectoderm results from the repression in the ventral ectoderm by Engrailed1 (En1)(Nusslein-Volhard and Wieschaus, 1980). The expression of En1 is itself induced by Bmpr1a (Bone Morphogenetic Protein Type I Receptor)(Coulter et al., 1990). The description of the zebrafish orthologs wnt7a and eng1a suggest a conserved role in the patterning of the dorsal and ventral ectoderm, respectively (Hatta et al., 1991; Norton et al., 2005). The same occurs with lmx1b, which expression of in the periferic muscle of the pectoral fin resembles the dorsal limb muscle of amniotes (Uemura et al., 2005). Thus, the players of the molecular network that regulate the patterning of D/V axis of the fin/limb bud are well conserved indicating a possible functional homology (Green et al., 2002).

24 5. Odd-skipped family genes

Odd-skipped family of proteins (Odd in Drosophila and Osr in vertebrates) are evolutionary conserved zinc-finger (ZF) transcription factors, with a variable number of ZFs among the members of this family of proteins (Goldstein et al., 2005). The first gene identified from this family was odd-skipped (odd) in a large mutagenesis screen for developmental control genes in Drosophila (Kawai et al., 2005). The gene name and its designation as a pair-rule gene was due to the cuticular defects present in the trunk of Odd mutant embryos and that are limited to alternate, odd-numbered segments (Gao et al., 2009). Following the above-mentioned discovery, three other members of this gene family were identified: sister of odd and bowl (sob), drumstick (drm) and bowel (bowl) (Green et al., 2002; Hart et al., 1996; Iwaki et al., 2001; Liu et al., 1999). These genes comprise four transcription factors that share high homology in their ZF domains, with a variable number of motifs: Odd has four, Sob and Bowl have five whereas Drm has only two motifs (Fig.12). odd, sob and drm are clustered together on the second chromosome and are similarly expressed in a specific segment of the gut, where midgut and hindgut connect. Their expression has also been observed in the ureters of Malpighian tubules (Green et al., 2002; Ward and Coulter, 2000), while bowl expression is detected along the hindgut (Green et al., 2002; Hart et al., 1996; Iwaki et al., 2001; Johansen et al., 2003; Ward and Coulter, 2000).

Figure 12- Members of the Odd family of proteins. Drm, Odd, Sob and Bowl proteins are similar in their ZF domains. In the remaining protein sequence they share a little degree of sequence similarity. The protein lengths are indicated on the right (Adapted from Green et al., 2002).

Although odd, sob and drm are expressed in almost identical patterns (Hart et al., 1996; Johansen et al., 2003), the analysis of the available mutants show different phenotypes (Coulter et al., 1990; Green et al., 2002; Johansen et al., 2003; Nusslein-Volhard and Wieschaus, 1980; Wang and Coulter, 1996). Odd mutant embryos show pattern defects in the anterior regions of odd-numbered segments (Lan et al., 2001; Lan et al., 2004; So and Danielian, 1999; Stricker et al., 2006). Drm is required for the establishment of the small

25 intestine (Lan et al., 2001). Bowl is expressed in the hindgut and the mutants for this protein die in late embryogenesis due to defects in terminal derivatives such as hindgut and proventriculus (Gao et al., 2009; Lan et al., 2001; Lan et al., 2004; Wang et al., 2005). Altogether, these data indicate that the members of the odd family have distinct functions during Drosophila development. Odd and Bowl proteins share an Engrailed homology 1 (Eh1) like motif. This common motif allows the recruitment of the co-repressor, Groucho, which downregulates gene expression during embryonic segmentation (Goldstein et al., 2005).

Two Odd-skipped related genes were found in the mammalian genome: Osr1 and Osr2 (Lan et al., 2001; So and Danielian, 1999). The protein coded by the Osr1 gene has three ZF (Osr1). The Osr2 gene encodes two proteins: Osr2B, with three zinc fingers; and Osr2A with five zinc fingers, resulting from alternative splicing of the transcripts (Katoh, 2002; Kawai et al., 2005; Lan et al., 2001; So and Danielian, 1999). It has been reported that when fused to the Gal4 DNA binding domain, the two Osr2 isoforms display opposite transcriptional activity in a cell culture based reporter assay (Lan et al., 2001). However, a recent work claims that Osr2A and Osr2B have the same functional potential when expressed in the same tissue (Gao et al., 2009). The structural homology of the mammalian Osr proteins with the Drosophila protein is limited to the ZF domains and the Eh1-like domain. Interestingly, Osr1 and Osr2B present a 65% amino acid sequence identity overall, and considering the amino acid sequence in the ZF domains present an identity of 98% (Fig.13).

Figure 13- Alignment of the zinc fingers from Drm, Odd, Sob, Bowl, mOsr1, mOsr2 and mOsr2 alt (the alternative splicing form). The latter three are mouse proteins. Identical and similar residues are shaded black and grey, respectively. Dashes indicate gaps in the alignment, and dots indicate amino acid residues (not shown) outside the zinc finger domains. Conserved residues in the canonical C2H2 zinc finger are shown below (Adapted from Green et al., 2002).

26 During development and organogenesis, the mouse Osr1 and Osr2 show both common and specific expression patterns (Gao et al., 2009). In mice, the onset of Osr1 expression occurs very early (E7.5) in the intermediate mesoderm, which is the region that renal structures originate from (Mugford et al., 2008; So and Danielian, 1999). This expression is maintained until kidney organogenesis occurs. Osr2 expression starts at stage E9.25 in the mesonephros, and at stage E14.5 it is expressed in the mesenchyme that surrounds the ducts of the mesonephros and metanephros (Gao et al.)(Fig.14). The expression of these genes is also detected in other tissues, such as heart, limbs and craniofacial structures. Targeted null mutations of each gene in mice give rise to distinct phenotypes probably related to differences in their expression pattern. The Osr1-/- mice show heart, kidney and urogenital defects whereas the Osr2-/- mice show cleft palate and open eyelids at birth (James et al., 2006; Lan et al., 2004; Wang et al., 2005). In fact, individual knockouts present phenotypes correlated with their specific expression pattern (James et al., 2006; Lan et al., 2004; Wang et al., 2005). Moreover, mutants for the individual genes do not show any phenotype in tissues where Osr1 and Osr2 are co-expressed suggesting that their roles are partially redundant during mouse embryonic development (Stricker et al., 2006).

A B

Fig.14- Expression patterns of Osr genes are very dynamic in early mouse embryos. (A) At E9.25 Osr1 expression is detected in the intermediate mesoderm (arrow) and in the mesoderm at the base of the heart primordium (arrowhead). Despite Osr1 being expressed in the intermediate mesoderm since E7.5, Osr2 is first mOsr1 E9.25 mOsr2 E9.25 expressed at E9.25 in the mesonephros (arrow). (C, D) At E10.5 Osr1 and Osr2 C D show distinct expression patterns in the maxillary (mx) and mandibular (mb) processes, the second branchial arch (b2), forelimb (fl) and hindlimb (hl) buds, and somites (s). Note that all images are lateral views of whole mount embryos (Adapted from Lan et al., 2001; Lan et al., 2004; So and Danielian, 1999; Stricker et al., 2006; Wang et al., 2005).

mOsr1 E10.5 mOsr2 E10.5

27 Indeed, mice with the endogenous Osr2 coding region replaced by either Osr1 cDNA or Osr2A cDNA are able to express these genes in most of the tissues where the endogenous Osr2 was expressed. The knock-in alleles Osr1 and Osr2A completely rescue cleft palate and cranial skeleton defects present in Osr2-/- mice but are not able to rescue the phenotype of open eyelids at birth. This can be explained by the differences in expression of the endogenous gene and the knock-in alleles during eyelid development (Buckley et al., 2004). Moreover, the phenotype rescue of the Osr2 -/- mutant mice by Osr1 indicates that both mammalian Osr genes evolved into different functional identities probably through changes in cis-regulatory regions that modulate their spatio-temporal expression patterns rather than coding sequences (Stricker et al., 2006). Recently, conditional inactivation of Osr1 in the developing limb mesenchyme in Osr2 -/- mutant mice resulted in the fusion of multiple joints. This phenotype strengths the hypothesis that Osr genes act redundantly in developmental processes where they are co-expressed such as synovial joints, kidney, craniofacial structures and limb (Gao et al.; Kawai et al., 2007; Lan et al., 2004; Mudumana et al., 2008; Wang et al., 2005; Zhang et al., 2009).

The expression patterns of Osr genes in chicken include the developing kidney, heart, gut, eye, branchial arches, the trunk dermis and the limbs (Fig.15) (James et al., 2006; Wang et al., 2005). These expression patterns are highly conserved with Osr expression patterns observed in mice (Fig.14). In C. elegans, two odd-skipped genes, which play essential and distinct roles in gut development were identified {Gao, 2009 #130;Lan, 2001 #166;Lan, 2004 #158;Wang, 2005 #152}.

Taken together, the expression patterns of vertebrate Osr genes in different tissues as well as functional (knock-out and knock-down) data from several studies indicate that Osr genes are required for correct formation and/or patterning of the heart, kidney, the endoderm, the teeth, the palate, the bones and the synovial joints in limbs (Gao et al., 2011; Kawai et al., 2007; Lan et al., 2004; Mudumana et al., 2008; Wang et al., 2005; Zhang et al., 2009)

28

Fig.15- Chicken Osr genes are expressed in A B the kidney, heart and limbs development. (A, B) At HH19, both Osr genes are expressed in the mesonephros (black arrowheads). In the ventral view inset Osr2 expression can be observed in nephric tubules. Osr1 expression is detected in the sinus venosus (white arrow). Osr2 is also expressed in the branchial arches (white arrowheads) and in a domain posterior to the eye (red arrowhead). (C, D) Later in development, at HH28, Osr1 and Osr2 expression is detected in the fore and C D hindlimbs, their expression in the flank is distinct, Osr2 is expressed dorsally (arrows), and Osr1 is expressed in a lateral domain (arrow) (Adapted from Stricker et al., 2006).

Taken together, the expression patterns of vertebrate Osr genes in different tissues as well as functional (knock-out and knock-down) data from several studies indicate that Osr genes are required for correct formation and/or patterning of the heart, kidney, the endoderm, the teeth, the palate, the bones and the synovial joints in limbs (Gao et al., 2011; Kawai et al., 2007; Lan et al., 2004; Mudumana et al., 2008; Wang et al., 2005; Zhang et al., 2009).

In this thesis, we addressed the role of osr genes in early kidney development taking advantage of the simple and valuable kidney system of zebrafish and Xenopus. Simultaneously with our studies, the mice phenotype of the Osr1 targeted mutation was published, unraveling that this gene was fundamental for kidney organogenesis as we also proposed in our study (James et al., 2006; Wang et al., 2005). In addition, we also determined the function of osr genes in fin development, unraveling an essential requirement of these genes during early stages of limb formation.

29 References:

The Danio rerio Sequencing Project Agarwal, P., J.N. Wylie, J. Galceran, O. Arkhitko, C. Li, C. Deng, R. Grosschedl, and B.G. Bruneau. 2003. Tbx5 is essential for forelimb bud initiation following patterning of the limb field in the mouse embryo. Development. 130:623-33. Ahn, D.G., M.J. Kourakis, L.A. Rohde, L.M. Silver, and R.K. Ho. 2002. T-box gene tbx5 is essential for formation of the pectoral limb bud. Nature. 417:754-8. Ahn, K., Y. Mishina, M.C. Hanks, R.R. Behringer, and E.B. Crenshaw, 3rd. 2001. BMPR-IA signaling is required for the formation of the apical ectodermal ridge and dorsal-ventral patterning of the limb. Development. 128:4449-61. Alarcon, P., E. Rodriguez-Seguel, A. Fernandez-Gonzalez, R. Rubio, and J.L. Gomez- Skarmeta. 2008. A dual requirement for Iroquois genes during Xenopus kidney development. Development. 135:3197-207. Amsterdam, A., R.M. Nissen, Z. Sun, E.C. Swindell, S. Farrington, and N. Hopkins. 2004. Identification of 315 genes essential for early zebrafish development. Proc Natl Acad Sci U S A. 101:12792-7. Begemann, G., and P.W. Ingham. 2000. Developmental regulation of Tbx5 in zebrafish embryogenesis. Mech Dev. 90:299-304. Begemann, G., T.F. Schilling, G.J. Rauch, R. Geisler, and P.W. Ingham. 2001. The zebrafish neckless mutation reveals a requirement for raldh2 in mesodermal signals that pattern the hindbrain. Development. 128:3081-94. Boonanuntanasarn, S., G. Yoshizaki, and T. Takeuchi. 2003. Specific gene silencing using small interfering RNAs in fish embryos. Biochem Biophys Res Commun. 310:1089-95. Bouchard, M., A. Souabni, M. Mandler, A. Neubuser, and M. Busslinger. 2002. Nephric lineage specification by Pax2 and Pax8. Genes Dev. 16:2958-70. Bracken, C.M., K. Mizeracka, and K.A. McLaughlin. 2008. Patterning the embryonic kidney: BMP signaling mediates the differentiation of the pronephric tubules and duct in Xenopus laevis. Dev Dyn. 237:132-44. Brandli, A.W. 1999. Towards a molecular anatomy of the Xenopus pronephric kidney. Int J Dev Biol. 43:381-95. Brennan, H.C., S. Nijjar, and E.A. Jones. 1998. The specification of the pronephric tubules and duct in Xenopus laevis. Mech Dev. 75:127-37. Buckley, M.S., J. Chau, P.E. Hoppe, and D.E. Coulter. 2004. odd-skipped homologs function during gut development in C. elegans. Dev Genes Evol. 214:10-8. Burke, A.C., C.E. Nelson, B.A. Morgan, and C. Tabin. 1995. Hox genes and the evolution of vertebrate axial morphology. Development. 121:333-46. Camarata, T., B. Bimber, A. Kulisz, T.L. Chew, J. Yeung, and H.G. Simon. 2006. LMP4 regulates Tbx5 protein subcellular localization and activity. J Cell Biol. 174:339-48. Camarata, T., D. Snyder, T. Schwend, J. Klosowiak, B. Holtrup, and H.G. Simon. Pdlim7 is required for maintenance of the mesenchymal/epidermal Fgf signaling feedback loop during zebrafish pectoral fin development. BMC Dev Biol. 10:104. Capdevila, J., and J.C. Izpisua Belmonte. 2001. Patterning mechanisms controlling vertebrate limb development. Annu Rev Cell Dev Biol. 17:87-132. Carmany-Rampey, A., and C.B. Moens. 2006. Modern mosaic analysis in the zebrafish. Methods. 39:228-38. Carroll, S.B., Grenier J.K., Weatherbee S.D. 2001. From DNA to diversity: molecular genetics and the evolution of animal design. Blackwel Science. Carroll, T., J. Wallingford, D. Seufert, and P.D. Vize. 1999a. Molecular regulation of pronephric development. Curr Top Dev Biol. 44:67-100.

30 Carroll, T.J., and P.D. Vize. 1996. Wilms' tumor suppressor gene is involved in the development of disparate kidney forms: evidence from expression in the Xenopus pronephros. Dev Dyn. 206:131-8. Carroll, T.J., and P.D. Vize. 1999. Synergism between Pax-8 and lim-1 in embryonic kidney development. Dev Biol. 214:46-59. Carroll, T.J., J.B. Wallingford, and P.D. Vize. 1999b. Dynamic patterns of gene expression in the developing pronephros of Xenopus laevis. Dev Genet. 24:199-207. Cartry, J., M. Nichane, V. Ribes, A. Colas, J.F. Riou, T. Pieler, P. Dolle, E.J. Bellefroid, and M. Umbhauer. 2006. Retinoic acid signalling is required for specification of pronephric cell fate. Dev Biol. 299:35-51. Chan, T.C., S. Takahashi, and M. Asashima. 2000. A role for Xlim-1 in pronephros development in Xenopus laevis. Dev Biol. 228:256-69. Chen, H., and R.L. Johnson. 1999. Dorsoventral patterning of the vertebrate limb: a process governed by multiple events. Cell Tissue Res. 296:67-73. Chen, H., Y. Lun, D. Ovchinnikov, H. Kokubo, K.C. Oberg, C.V. Pepicelli, L. Gan, B. Lee, and R.L. Johnson. 1998. Limb and kidney defects in Lmx1b mutant mice suggest an involvement of LMX1B in human nail patella syndrome. Nat Genet. 19:51-5. Chen, Y., N. Pollet, C. Niehrs, and T. Pieler. 2001. Increased XRALDH2 activity has a posteriorizing effect on the central nervous system of Xenopus embryos. Mech Dev. 101:91-103. Cheng, H.T., M. Kim, M.T. Valerius, K. Surendran, K. Schuster-Gossler, A. Gossler, A.P. McMahon, and R. Kopan. 2007. Notch2, but not Notch1, is required for proximal fate acquisition in the mammalian nephron. Development. 134:801-11. Cheng, H.T., J.H. Miner, M. Lin, M.G. Tansey, K. Roth, and R. Kopan. 2003. Gamma- secretase activity is dispensable for mesenchyme-to-epithelium transition but required for podocyte and proximal tubule formation in developing mouse kidney. Development. 130:5031-42. Cheret, C., A. Doyen, M. Yaniv, and M. Pontoglio. 2002. Hepatocyte nuclear factor 1 alpha controls renal expression of the Npt1-Npt4 anionic transporter locus. J Mol Biol. 322:929-41. Christensen, E.I., D. Raciti, L. Reggiani, P.J. Verroust, and A.W. Brandli. 2008. Gene expression analysis defines the proximal tubule as the compartment for endocytic receptor-mediated uptake in the Xenopus pronephric kidney. Pflugers Arch. 456:1163- 76. Coates, M.I. 1995. Limb evolution. Fish fins or tetrapod limbs--a simple twist of fate? Curr Biol. 5:844-8. Cohn, M.J., J.C. Izpisua-Belmonte, H. Abud, J.K. Heath, and C. Tickle. 1995. Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell. 80:739-46. Cohn, M.J., K. Patel, R. Krumlauf, D.G. Wilkinson, J.D. Clarke, and C. Tickle. 1997. Hox9 genes and vertebrate limb specification. Nature. 387:97-101. Cohn, M.J., and C. Tickle. 1999. Developmental basis of limblessness and axial patterning in snakes. Nature. 399:474-9. Colas, A., J. Cartry, I. Buisson, M. Umbhauer, J.C. Smith, and J.F. Riou. 2008. Mix.1/2- dependent control of FGF availability during gastrulation is essential for pronephros development in Xenopus. Dev Biol. 320:351-65. Cooper, K.L., J.K. Hu, D. ten Berge, M. Fernandez-Teran, M.A. Ros, and C.J. Tabin. 2011. Initiation of proximal-distal patterning in the vertebrate limb by signals and growth. Science. 332:1083-6.

31 Coulter, D.E., E.A. Swaykus, M.A. Beran-Koehn, D. Goldberg, E. Wieschaus, and P. Schedl. 1990. Molecular analysis of odd-skipped, a zinc finger encoding segmentation gene with a novel pair-rule expression pattern. EMBO J. 9:3795-804. Cygan, J.A., R.L. Johnson, and A.P. McMahon. 1997. Novel regulatory interactions revealed by studies of murine limb pattern in Wnt-7a and En-1 mutants. Development. 124:5021-32. Deconinck, A.E., P.E. Mead, S.G. Tevosian, J.D. Crispino, S.G. Katz, L.I. Zon, and S.H. Orkin. 2000. FOG acts as a repressor of red blood cell development in Xenopus. Development. 127:2031-40. Deschamps, J., E. van den Akker, S. Forlani, W. De Graaff, T. Oosterveen, B. Roelen, and J. Roelfsema. 1999. Initiation, establishment and maintenance of Hox gene expression patterns in the mouse. Int J Dev Biol. 43:635-50. Dodd, A., S.P. Chambers, and D.R. Love. 2004. Short interfering RNA-mediated gene targeting in the zebrafish. FEBS Lett. 561:89-93. Dressler, G.R. 2006. The cellular basis of kidney development. Annu Rev Cell Dev Biol. 22:509-29. Dressler, G.R. 2009. Advances in early kidney specification, development and patterning. Development. 136:3863-74. Drummond, I.A. 2000. The zebrafish pronephros: a genetic system for studies of kidney development. Pediatr Nephrol. 14:428-35. Drummond, I.A. 2005. Kidney development and disease in the zebrafish. J Am Soc Nephrol. 16:299-304. Drummond, I.A., A. Majumdar, H. Hentschel, M. Elger, L. Solnica-Krezel, A.F. Schier, S.C. Neuhauss, D.L. Stemple, F. Zwartkruis, Z. Rangini, W. Driever, and M.C. Fishman. 1998. Early development of the zebrafish pronephros and analysis of mutations affecting pronephric function. Development. 125:4655-67. Duester, G. 2007. Retinoic acid regulation of the somitogenesis clock. Birth Defects Res C Embryo Today. 81:84-92. Eid, S.R., A. Terrettaz, K. Nagata, and A.W. Brandli. 2002. Embryonic expression of Xenopus SGLT-1L, a novel member of the solute carrier family 5 (SLC5), is confined to tubules of the pronephric kidney. Int J Dev Biol. 46:177-84. Emoto, Y., H. Wada, H. Okamoto, A. Kudo, and Y. Imai. 2005. Retinoic acid-metabolizing enzyme Cyp26a1 is essential for determining territories of hindbrain and spinal cord in zebrafish. Dev Biol. 278:415-27. Fischer, S., B.W. Draper, and C.J. Neumann. 2003. The zebrafish fgf24 mutant identifies an additional level of Fgf signaling involved in vertebrate forelimb initiation. Development. 130:3515-24. Galceran, J., I. Farinas, M.J. Depew, H. Clevers, and R. Grosschedl. 1999. Wnt3a-/--like phenotype and limb deficiency in Lef1(-/-)Tcf1(-/-) mice. Genes Dev. 13:709-17. Gao, Y., Y. Lan, H. Liu, and R. Jiang. 2011. The zinc finger transcription factors Osr1 and Osr2 control synovial joint formation. Dev Biol. 352:83-91. Gao, Y., Y. Lan, C.E. Ovitt, and R. Jiang. 2009. Functional equivalence of the zinc finger transcription factors Osr1 and Osr2 in mouse development. Dev Biol. 328:200-9. Garrity, D.M., S. Childs, and M.C. Fishman. 2002. The heartstrings mutation in zebrafish causes heart/fin Tbx5 deficiency syndrome. Development. 129:4635-45. Gaunt, S.J. 2000. Evolutionary shifts of vertebrate structures and Hox expression up and down the axial series of segments: a consideration of possible mechanisms. Int J Dev Biol. 44:109-17. Geduspan, J.S., and M. Solursh. 1992. A growth-promoting influence from the mesonephros during limb outgrowth. Dev Biol. 151:242-50.

32 Gibert, Y., A. Gajewski, A. Meyer, and G. Begemann. 2006. Induction and prepatterning of the zebrafish pectoral fin bud requires axial retinoic acid signaling. Development. 133:2649-59. Grainger, R.S., H.L. Harland, R.M. 2000. Early Development of Xenopus laevis: A Laboratory Manual. Cold Spring Harbor Laboratory Press. 338 pp. Grammer, T.C., M.K. Khokha, M.A. Lane, K. Lam, and R.M. Harland. 2005. Identification of mutants in inbred Xenopus tropicalis. Mech Dev. 122:263-72. Grammer, T.C., K.J. Liu, F.V. Mariani, and R.M. Harland. 2000. Use of large-scale expression cloning screens in the Xenopus laevis tadpole to identify gene function. Dev Biol. 228:197-210. Grandel, H., and M. Brand. 2011. Zebrafish limb development is triggered by a retinoic acid signal during gastrulation. Dev Dyn. Grandel, H., K. Lun, G.J. Rauch, M. Rhinn, T. Piotrowski, C. Houart, P. Sordino, A.M. Kuchler, S. Schulte-Merker, R. Geisler, N. Holder, S.W. Wilson, and M. Brand. 2002. Retinoic acid signalling in the zebrafish embryo is necessary during pre-segmentation stages to pattern the anterior-posterior axis of the CNS and to induce a pectoral fin bud. Development. 129:2851-65. Green, R.B., V. Hatini, K.A. Johansen, X.J. Liu, and J.A. Lengyel. 2002. Drumstick is a zinc finger protein that antagonizes Lines to control patterning and morphogenesis of the Drosophila hindgut. Development. 129:3645-56. Grieshammer, U., C. Cebrian, R. Ilagan, E. Meyers, D. Herzlinger, and G.R. Martin. 2005. FGF8 is required for cell survival at distinct stages of nephrogenesis and for regulation of gene expression in nascent nephrons. Development. 132:3847-57. Grote, D., S.K. Boualia, A. Souabni, C. Merkel, X. Chi, F. Costantini, T. Carroll, and M. Bouchard. 2008. Gata3 acts downstream of beta-catenin signaling to prevent ectopic metanephric kidney induction. PLoS Genet. 4:e1000316. Grote, D., A. Souabni, M. Busslinger, and M. Bouchard. 2006. Pax 2/8-regulated Gata 3 expression is necessary for morphogenesis and guidance of the nephric duct in the developing kidney. Development. 133:53-61. Haffter, P., M. Granato, M. Brand, M.C. Mullins, M. Hammerschmidt, D.A. Kane, J. Odenthal, F.J. van Eeden, Y.J. Jiang, C.P. Heisenberg, R.N. Kelsh, M. Furutani-Seiki, E. Vogelsang, D. Beuchle, U. Schach, C. Fabian, and C. Nusslein-Volhard. 1996. The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development. 123:1-36. Harvey, S.A., and M.P. Logan. 2006. sall4 acts downstream of tbx5 and is required for pectoral fin outgrowth. Development. 133:1165-73. Heller, N., and A.W. Brandli. 1999. Xenopus Pax-2/5/8 orthologues: novel insights into Pax gene evolution and identification of Pax-8 as the earliest marker for otic and pronephric cell lineages. Dev Genet. 24:208-19. Hellsten, U., R.M. Harland, M.J. Gilchrist, D. Hendrix, J. Jurka, V. Kapitonov, I. Ovcharenko, N.H. Putnam, S. Shu, L. Taher, I.L. Blitz, B. Blumberg, D.S. Dichmann, I. Dubchak, E. Amaya, J.C. Detter, R. Fletcher, D.S. Gerhard, D. Goodstein, T. Graves, I.V. Grigoriev, J. Grimwood, T. Kawashima, E. Lindquist, S.M. Lucas, P.E. Mead, T. Mitros, H. Ogino, Y. Ohta, A.V. Poliakov, N. Pollet, J. Robert, A. Salamov, A.K. Sater, J. Schmutz, A. Terry, P.D. Vize, W.C. Warren, D. Wells, A. Wills, R.K. Wilson, L.B. Zimmerman, A.M. Zorn, R. Grainger, T. Grammer, M.K. Khokha, P.M. Richardson, and D.S. Rokhsar. 2010. The genome of the Western clawed frog Xenopus tropicalis. Science. 328:633-6. Hollemann, T., Y. Chen, H. Grunz, and T. Pieler. 1998. Regionalized metabolic activity establishes boundaries of retinoic acid signalling. EMBO J. 17:7361-72.

33 James, R.G., C.N. Kamei, Q. Wang, R. Jiang, and T.M. Schultheiss. 2006. Odd-skipped related 1 is required for development of the metanephric kidney and regulates formation and differentiation of kidney precursor cells. Development. 133:2995-3004. Johnson, R.L., and C.J. Tabin. 1997. Molecular models for vertebrate limb development. Cell. 90:979-90. Jones, E.A. 2005. Xenopus: a prince among models for pronephric kidney development. J Am Soc Nephrol. 16:313-21. Kawai, S., T. Kato, H. Inaba, N. Okahashi, and A. Amano. 2005. Odd-skipped related 2 splicing variants show opposite transcriptional activity. Biochem Biophys Res Commun. 328:306-11. Kawai, S., M. Yamauchi, S. Wakisaka, T. Ooshima, and A. Amano. 2007. Zinc-finger transcription factor odd-skipped related 2 is one of the regulators in osteoblast proliferation and bone formation. J Bone Miner Res. 22:1362-72. Kawakami, Y., J. Capdevila, D. Buscher, T. Itoh, C. Rodriguez Esteban, and J.C. Izpisua Belmonte. 2001. WNT signals control FGF-dependent limb initiation and AER induction in the chick embryo. Cell. 104:891-900. Keegan, B.R., J.L. Feldman, G. Begemann, P.W. Ingham, and D. Yelon. 2005. Retinoic acid signaling restricts the cardiac progenitor pool. Science. 307:247-9. Kimmel, C.B., W.W. Ballard, S.R. Kimmel, B. Ullmann, and T.F. Schilling. 1995. Stages of embryonic development of the zebrafish. Dev Dyn. 203:253-310. Kobayashi, A., K.M. Kwan, T.J. Carroll, A.P. McMahon, C.L. Mendelsohn, and R.R. Behringer. 2005. Distinct and sequential tissue-specific activities of the LIM-class homeobox gene Lim1 for tubular morphogenesis during kidney development. Development. 132:2809-23. Kolm, P.J., and H.L. Sive. 1995. Efficient hormone-inducible protein function in Xenopus laevis. Dev Biol. 171:267-72. Kriz, W., and L. Bankir. 1988. A standard nomenclature for structures of the kidney. The Renal Commission of the International Union of Physiological Sciences (IUPS). Kidney Int. 33:1-7. Lan, Y., P.D. Kingsley, E.S. Cho, and R. Jiang. 2001. Osr2, a new mouse gene related to Drosophila odd-skipped, exhibits dynamic expression patterns during craniofacial, limb, and kidney development. Mech Dev. 107:175-9. Lan, Y., C.E. Ovitt, E.S. Cho, K.M. Maltby, Q. Wang, and R. Jiang. 2004. Odd-skipped related 2 (Osr2) encodes a key intrinsic regulator of secondary palate growth and morphogenesis. Development. 131:3207-16. Lee, B.C., and S. Roy. 2006. Blimp-1 is an essential component of the genetic program controlling development of the pectoral limb bud. Dev Biol. 300:623-34. Logan, C., A. Hornbruch, I. Campbell, and A. Lumsden. 1997. The role of Engrailed in establishing the dorsoventral axis of the chick limb. Development. 124:2317-24. Loomis, C.A., E. Harris, J. Michaud, W. Wurst, M. Hanks, and A.L. Joyner. 1996. The mouse Engrailed-1 gene and ventral limb patterning. Nature. 382:360-3. Loomis, C.A., R.A. Kimmel, C.X. Tong, J. Michaud, and A.L. Joyner. 1998. Analysis of the genetic pathway leading to formation of ectopic apical ectodermal ridges in mouse Engrailed-1 mutant limbs. Development. 125:1137-48. Mackem, S., and M. Lewandoski. 2011. Development. Limb cells don't tell time. Science. 332:1038-9. Maden, M. 1999. Heads or tails? Retinoic acid will decide. Bioessays. 21:809-12. Majumdar, A., K. Lun, M. Brand, and I.A. Drummond. 2000. Zebrafish no isthmus reveals a role for pax2.1 in tubule differentiation and patterning events in the pronephric primordia. Development. 127:2089-98.

34 Martin, G.R. 1998. The roles of FGFs in the early development of vertebrate limbs. Genes Dev. 12:1571-86. McLaughlin, K.A., M.S. Rones, and M. Mercola. 2000. Notch regulates cell fate in the developing pronephros. Dev Biol. 227:567-80. McPherron, A.C., A.M. Lawler, and S.J. Lee. 1999. Regulation of anterior/posterior patterning of the axial skeleton by growth/differentiation factor 11. Nat Genet. 22:260-4. Mercader, N. 2007. Early steps of paired fin development in zebrafish compared with tetrapod limb development. Dev Growth Differ. 49:421-37. Mercader, N., S. Fischer, and C.J. Neumann. 2006. Prdm1 acts downstream of a sequential RA, Wnt and Fgf signaling cascade during zebrafish forelimb induction. Development. 133:2805-15. Mercader, N., E. Leonardo, M.E. Piedra, A.C. Martinez, M.A. Ros, and M. Torres. 2000. Opposing RA and FGF signals control proximodistal vertebrate limb development through regulation of Meis genes. Development. 127:3961-70. Mereau, A., C. Le Sommer, H. Lerivray, M. Lesimple, and S. Hardy. 2007. Xenopus as a model to study alternative splicing in vivo. Biol Cell. 99:55-65. Mic, F.A., I.O. Sirbu, and G. Duester. 2004. Retinoic acid synthesis controlled by Raldh2 is required early for limb bud initiation and then later as a proximodistal signal during apical ectodermal ridge formation. J Biol Chem. 279:26698-706. Min, H., D.M. Danilenko, S.A. Scully, B. Bolon, B.D. Ring, J.E. Tarpley, M. DeRose, and W.S. Simonet. 1998. Fgf-10 is required for both limb and lung development and exhibits striking functional similarity to Drosophila branchless. Genes Dev. 12:3156-61. Minguillon, C., J. Del Buono, and M.P. Logan. 2005. Tbx5 and Tbx4 are not sufficient to determine limb-specific morphologies but have common roles in initiating limb outgrowth. Dev Cell. 8:75-84. Mobjerg, N., E.H. Larsen, and A. Jespersen. 2000. Morphology of the kidney in larvae of Bufo viridis (Amphibia, Anura, Bufonidae). J Morphol. 245:177-95. Mudumana, S.P., D. Hentschel, Y. Liu, A. Vasilyev, and I.A. Drummond. 2008. odd skipped related1 reveals a novel role for endoderm in regulating kidney versus vascular cell fate. Development. 135:3355-67. Naiche, L.A., and V.E. Papaioannou. 2003. Loss of Tbx4 blocks hindlimb development and affects vascularization and fusion of the allantois. Development. 130:2681-93. Nakai, S., Y. Sugitani, H. Sato, S. Ito, Y. Miura, M. Ogawa, M. Nishi, K. Jishage, O. Minowa, and T. Noda. 2003. Crucial roles of Brn1 in distal tubule formation and function in mouse kidney. Development. 130:4751-9. Naylor, R.W., and E.A. Jones. 2009. Notch activates Wnt-4 signalling to control medio-lateral patterning of the pronephros. Development. 136:3585-95. Negishi, T., Y. Nagai, Y. Asaoka, M. Ohno, M. Namae, H. Mitani, T. Sasaki, N. Shimizu, S. Terai, I. Sakaida, H. Kondoh, T. Katada, M. Furutani-Seiki, and H. Nishina. 2010. Retinoic acid signaling positively regulates liver specification by inducing wnt2bb gene expression in medaka. Hepatology. 51:1037-45. Ng, J.K., Y. Kawakami, D. Buscher, A. Raya, T. Itoh, C.M. Koth, C. Rodriguez Esteban, J. Rodriguez-Leon, D.M. Garrity, M.C. Fishman, and J.C. Izpisua Belmonte. 2002. The limb identity gene Tbx5 promotes limb initiation by interacting with Wnt2b and Fgf10. Development. 129:5161-70. Niswander, L. 2003. Pattern formation: old models out on a limb. Nat Rev Genet. 4:133-43. Noramly, S., L. Zimmerman, A. Cox, R. Aloise, M. Fisher, and R.M. Grainger. 2005. A gynogenetic screen to isolate naturally occurring recessive mutations in Xenopus tropicalis. Mech Dev. 122:273-87.

35 Norton, W.H., J. Ledin, H. Grandel, and C.J. Neumann. 2005. HSPG synthesis by zebrafish Ext2 and Extl3 is required for Fgf10 signalling during limb development. Development. 132:4963-73. Nusslein-Volhard, C., and E. Wieschaus. 1980. Mutations affecting segment number and polarity in Drosophila. Nature. 287:795-801. Ohuchi, H., T. Nakagawa, A. Yamamoto, A. Araga, T. Ohata, Y. Ishimaru, H. Yoshioka, T. Kuwana, T. Nohno, M. Yamasaki, N. Itoh, and S. Noji. 1997. The mesenchymal factor, FGF10, initiates and maintains the outgrowth of the chick limb bud through interaction with FGF8, an apical ectodermal factor. Development. 124:2235-44. Ohuchi, H., T. Nakagawa, M. Yamauchi, T. Ohata, H. Yoshioka, T. Kuwana, T. Mima, T. Mikawa, T. Nohno, and S. Noji. 1995. An additional limb can be induced from the flank of the chick embryo by FGF4. Biochem Biophys Res Commun. 209:809-16. Pack, M., L. Solnica-Krezel, J. Malicki, S.C. Neuhauss, A.F. Schier, D.L. Stemple, W. Driever, and M.C. Fishman. 1996. Mutations affecting development of zebrafish digestive organs. Development. 123:321-8. Pedersen, A., C. Skjong, and W. Shawlot. 2005. Lim 1 is required for nephric duct extension and ureteric bud morphogenesis. Dev Biol. 288:571-81. Pizette, S., C. Abate-Shen, and L. Niswander. 2001. BMP controls proximodistal outgrowth, via induction of the apical ectodermal ridge, and dorsoventral patterning in the vertebrate limb. Development. 128:4463-74. Pontoglio, M., J. Barra, M. Hadchouel, A. Doyen, C. Kress, J.P. Bach, C. Babinet, and M. Yaniv. 1996. Hepatocyte nuclear factor 1 inactivation results in hepatic dysfunction, phenylketonuria, and renal Fanconi syndrome. Cell. 84:575-85. Pontoglio, M., D. Prie, C. Cheret, A. Doyen, C. Leroy, P. Froguel, G. Velho, M. Yaniv, and G. Friedlander. 2000. HNF1alpha controls renal glucose reabsorption in mouse and man. EMBO Rep. 1:359-65. Porteous, S., E. Torban, N.P. Cho, H. Cunliffe, L. Chua, L. McNoe, T. Ward, C. Souza, P. Gus, R. Giugliani, T. Sato, K. Yun, J. Favor, M. Sicotte, P. Goodyer, and M. Eccles. 2000. Primary renal hypoplasia in humans and mice with PAX2 mutations: evidence of increased apoptosis in fetal kidneys of Pax2(1Neu) +/- mutant mice. Hum Mol Genet. 9:1-11. Raciti, D., L. Reggiani, L. Geffers, Q. Jiang, F. Bacchion, A.E. Subrizi, D. Clements, C. Tindal, D.R. Davidson, B. Kaissling, and A.W. Brandli. 2008. Organization of the pronephric kidney revealed by large-scale gene expression mapping. Genome Biol. 9:R84. Rallis, C., B.G. Bruneau, J. Del Buono, C.E. Seidman, J.G. Seidman, S. Nissim, C.J. Tabin, and M.P. Logan. 2003. Tbx5 is required for forelimb bud formation and continued outgrowth. Development. 130:2741-51. Ransom, D.G., P. Haffter, J. Odenthal, A. Brownlie, E. Vogelsang, R.N. Kelsh, M. Brand, F.J. van Eeden, M. Furutani-Seiki, M. Granato, M. Hammerschmidt, C.P. Heisenberg, Y.J. Jiang, D.A. Kane, M.C. Mullins, and C. Nusslein-Volhard. 1996. Characterization of zebrafish mutants with defects in embryonic hematopoiesis. Development. 123:311-9. Reggiani, L., D. Raciti, R. Airik, A. Kispert, and A.W. Brandli. 2007. The prepattern transcription factor Irx3 directs nephron segment identity. Genes Dev. 21:2358-70. Richards, A.N. 1929. Methods and results of direct investigations of the fucntion of the kidney. Williams and Wilkins Company, Baltimore. Riddle, R.D., M. Ensini, C. Nelson, T. Tsuchida, T.M. Jessell, and C. Tabin. 1995. Induction of the LIM homeobox gene Lmx1 by WNT7a establishes dorsoventral pattern in the vertebrate limb. Cell. 83:631-40. Riddle, R.D., R.L. Johnson, E. Laufer, and C. Tabin. 1993. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell. 75:1401-16.

36 Rosello-Diez, A., M.A. Ros, and M. Torres. 2011. Diffusible signals, not autonomous mechanisms, determine the main proximodistal limb subdivision. Science. 332:1086- 8. Ruiz i Altaba, A., and T. Jessell. 1991. Retinoic acid modifies mesodermal patterning in early Xenopus embryos. Genes Dev. 5:175-87. Ruvinsky, I., A.C. Oates, L.M. Silver, and R.K. Ho. 2000. The evolution of paired appendages in vertebrates: T-box genes in the zebrafish. Dev Genes Evol. 210:82-91. Ryffel, G.U. 2003. What can a frog tell us about human kidney development. Nephron Exp Nephrol. 94:e35-43. Saito, D., S. Yonei-Tamura, Y. Takahashi, and K. Tamura. 2006. Level-specific role of paraxial mesoderm in regulation of Tbx5/Tbx4 expression and limb initiation. Dev Biol. 292:79-89. Saulnier, D.M., H. Ghanbari, and A.W. Brandli. 2002. Essential function of Wnt-4 for tubulogenesis in the Xenopus pronephric kidney. Dev Biol. 248:13-28. Saunders J.W., G.M.T. 1968. Epithelial mesenchymal interactions, Baltimore. Sekine, K., H. Ohuchi, M. Fujiwara, M. Yamasaki, T. Yoshizawa, T. Sato, N. Yagishita, D. Matsui, Y. Koga, N. Itoh, and S. Kato. 1999. Fgf10 is essential for limb and lung formation. Nat Genet. 21:138-41. Shawlot, W., and R.R. Behringer. 1995. Requirement for Lim1 in head-organizer function. Nature. 374:425-30. Sirbu, I.O., and G. Duester. 2006. Retinoic-acid signalling in node ectoderm and posterior neural plate directs left-right patterning of somitic mesoderm. Nat Cell Biol. 8:271-7. So, P.L., and P.S. Danielian. 1999. Cloning and expression analysis of a mouse gene related to Drosophila odd-skipped. Mech Dev. 84:157-60. Stafford, D., and V.E. Prince. 2002. Retinoic acid signaling is required for a critical early step in zebrafish pancreatic development. Curr Biol. 12:1215-20. Stemple, D.L., and W. Driever. 1996. Zebrafish: tools for investigating cellular differentiation. Curr Opin Cell Biol. 8:858-64. Stephens, T.D., and T.R. McNulty. 1981. Evidence for a metameric pattern in the development of the chick humerus. J Embryol Exp Morphol. 61:191-205. Stricker, S., N. Brieske, J. Haupt, and S. Mundlos. 2006. Comparative expression pattern of Odd-skipped related genes Osr1 and Osr2 in chick embryonic development. Gene Expr Patterns. 6:826-34. Summerbell, D., J.H. Lewis, and L. Wolpert. 1973. Positional information in chick limb morphogenesis. Nature. 244:492-6. Tabin, C., and L. Wolpert. 2007. Rethinking the proximodistal axis of the vertebrate limb in the molecular era. Genes Dev. 21:1433-42. Taelman, V., C. Van Campenhout, M. Solter, T. Pieler, and E.J. Bellefroid. 2006. The Notch- effector HRT1 gene plays a role in glomerular development and patterning of the Xenopus pronephros anlagen. Development. 133:2961-71. Takeuchi, J.K., K. Koshiba-Takeuchi, K. Matsumoto, A. Vogel-Hopker, M. Naitoh-Matsuo, K. Ogura, N. Takahashi, K. Yasuda, and T. Ogura. 1999. Tbx5 and Tbx4 genes determine the wing/leg identity of limb buds. Nature. 398:810-4. Takeuchi, J.K., K. Koshiba-Takeuchi, T. Suzuki, M. Kamimura, K. Ogura, and T. Ogura. 2003. Tbx5 and Tbx4 trigger limb initiation through activation of the Wnt/Fgf signaling cascade. Development. 130:2729-39. Tamura, K., S. Yonei-Tamura, and J.C. Izpisua Belmonte. 1999. Differential expression of Tbx4 and Tbx5 in Zebrafish fin buds. Mech Dev. 87:181-4. Tanaka, K., S. Terryn, L. Geffers, S. Garbay, M. Pontoglio, and O. Devuyst. 2010. The transcription factor HNF1alpha regulates expression of chloride-proton exchanger ClC-5 in the renal proximal tubule. Am J Physiol Renal Physiol. 299:F1339-47.

37 Tanaka M., T.C. 2007. The development of Fins and Limbs. In Fins into Limbs: Evolution, Development, and Transformation. B.K. Hall, editor. The University of Chicago Press, Chicago and London. 65-78. Tarin, D., and A.P. Sturdee. 1971. Early limb development of Xenopus laevis. J Embryol Exp Morphol. 26:169-79. ten Berge, D., S.A. Brugmann, J.A. Helms, and R. Nusse. 2008. Wnt and FGF signals interact to coordinate growth with cell fate specification during limb development. Development. 135:3247-57. Tickle, C. 1999. Morphogen gradients in vertebrate limb development. Semin Cell Dev Biol. 10:345-51. Torres, M., E. Gomez-Pardo, G.R. Dressler, and P. Gruss. 1995. Pax-2 controls multiple steps of urogenital development. Development. 121:4057-65. Tran, U., L.M. Pickney, B.D. Ozpolat, and O. Wessely. 2007. Xenopus Bicaudal-C is required for the differentiation of the amphibian pronephros. Dev Biol. 307:152-64. Tytler, P. 1988. Morphology of the pronephros of the juvenile brown trout, Salmo trutta. J Morphol. 195:189-204. Urban, A.E., X. Zhou, J.M. Ungos, D.W. Raible, C.R. Altmann, and P.D. Vize. 2006. FGF is essential for both condensation and mesenchymal-epithelial transition stages of pronephric kidney tubule development. Dev Biol. 297:103-17. Van Campenhout, C., M. Nichane, A. Antoniou, H. Pendeville, O.J. Bronchain, J.C. Marine, A. Mazabraud, M.L. Voz, and E.J. Bellefroid. 2006. Evi1 is specifically expressed in the distal tubule and duct of the Xenopus pronephros and plays a role in its formation. Dev Biol. 294:203-19. van Eeden, F.J., M. Granato, U. Schach, M. Brand, M. Furutani-Seiki, P. Haffter, M. Hammerschmidt, C.P. Heisenberg, Y.J. Jiang, D.A. Kane, R.N. Kelsh, M.C. Mullins, J. Odenthal, R.M. Warga, and C. Nusslein-Volhard. 1996. Genetic analysis of fin formation in the zebrafish, Danio rerio. Development. 123:255-62. Vize, P.D. 2003a. The chloride conductance channel ClC-K is a specific marker for the Xenopus pronephric distal tubule and duct. Gene Expr Patterns. 3:347-50. Vize, P.D., D.W. Seufert, T.J. Carroll, and J.B. Wallingford. 1997. Model systems for the study of kidney development: use of the pronephros in the analysis of organ induction and patterning. Dev Biol. 188:189-204. Vize, P.D., Woolf A. S., Bard J.B.L 2003b. The Kidney: from Normal Development to Congenital Disease. Academic Press, San Diego, CA. Vogel, A., C. Rodriguez, W. Warnken, and J.C. Izpisua Belmonte. 1995. Dorsal cell fate specified by chick Lmx1 during vertebrate limb development. Nature. 378:716-20. Wang, L., and D.E. Coulter. 1996. bowel, an odd-skipped homolog, functions in the terminal pathway during Drosophila embryogenesis. EMBO J. 15:3182-96. Wang, P., F.A. Pereira, D. Beasley, and H. Zheng. 2003. Presenilins are required for the formation of comma- and S-shaped bodies during nephrogenesis. Development. 130:5019-29. Wang, Q., Y. Lan, E.S. Cho, K.M. Maltby, and R. Jiang. 2005. Odd-skipped related 1 (Odd 1) is an essential regulator of heart and urogenital development. Dev Biol. 288:582-94. Weber, H., B. Holewa, E.A. Jones, and G.U. Ryffel. 1996. Mesoderm and endoderm differentiation in animal cap explants: identification of the HNF4-binding site as an activin A responsive element in the Xenopus HNF1alpha promoter. Development. 122:1975-84. Wessely, O., and U. Tran. 2011 . Xenopus pronephros development-past, present, and future. Pediatr Nephrol.

38 White, J.T., B. Zhang, D.M. Cerqueira, U. Tran, and O. Wessely. 2010. Notch signaling, wt1 and foxc2 are key regulators of the podocyte gene regulatory network in Xenopus. Development. 137:1863-73. Wilm, T.P., and L. Solnica-Krezel. 2005. Essential roles of a zebrafish prdm1/blimp1 homolog in embryo patterning and organogenesis. Development. 132:393-404. Wingert, R.A., and A.J. Davidson. 2008. The zebrafish pronephros: a model to study nephron segmentation. Kidney Int. 73:1120-7. Wingert, R.A., R. Selleck, J. Yu, H.D. Song, Z. Chen, A. Song, Y. Zhou, B. Thisse, C. Thisse, A.P. McMahon, and A.J. Davidson. 2007. The cdx genes and retinoic acid control the positioning and segmentation of the zebrafish pronephros. PLoS Genet. 3:1922-38. Wolpert L., B.R., Jessell T., Lawrence P., Elliot Meyerowitz, Jim Smith. 2002. Principles of Development. Oxford University Press. Yonei-Tamura, S., T. Endo, H. Yajima, H. Ohuchi, H. Ide, and K. Tamura. 1999. FGF7 and FGF10 directly induce the apical ectodermal ridge in chick embryos. Dev Biol. 211:133-43. Zhang, Z., Y. Lan, Y. Chai, and R. Jiang. 2009. Antagonistic actions of Msx1 and Osr2 pattern mammalian teeth into a single row. Science. 323:1232-4. Zhou, W., R.C. Boucher, F. Bollig, C. Englert, and F. Hildebrandt. Characterization of mesonephric development and regeneration using transgenic zebrafish. Am J Physiol Renal Physiol. 299:F1040-7. Zhou, X., and P.D. Vize. 2004. Proximo-distal specialization of epithelial transport processes within the Xenopus pronephric kidney tubules. Dev Biol. 271:322-38. Zhou, X., and P.D. Vize. 2005. Amino acid cotransporter SLC3A2 is selectively expressed in the early proximal segment of Xenopus pronephric kidney nephrons. Gene Expr Patterns. 5:774-7.

39 40

AIMS

The general aim of this thesis is to characterize the role of osr genes in renal and fin development.

Specific aims

• Characterize the expression pattern of osr genes during kidney development using Xenopus and zebrafish as model systems. • Study the function of osr genes during kidney development. • Identify the molecular behaviour of osr genes during renal development.

• Characterize the expression of osr genes throughout zebrafish pectoral fin formation. • Study the role of osr genes in the pectoral fin formation. • Unravel the place of osr1 and osr2 in the fin/limb genetic cascade.

41

42

CHAPTER I

“Odd-skipped genes encode repressors that control kidney development”

In this chapter, we characterized the expression pattern of osr genes in Xenopus and zebrafish. Here, we show that osr1 and osr2 are both necessary and sufficient for kidney development. These genes behave as molecular repressors in the context of renal development. Drosophila odd is also able to induce kidney development.

I contributed to this work performing the experiments using Xenopus and zebrafish.

43

44 Developmental Biology 301 (2007) 518–531 www.elsevier.com/locate/ydbio

Odd-skipped genes encode repressors that control kidney development

Juan J. Tena a, Ana Neto a, Elisa de la Calle-Mustienes a, Catarina Bras-Pereira a,c, ⁎ ⁎ Fernando Casares a,b, , José Luis Gómez-Skarmeta a,

a Centro Andaluz de Biología del Desarrollo, Consejo Superior de Investigaciones Científicas/Universidad Pablo de Olavide, Carretera de Utrera Km1, 41013 Sevilla, Spain b Instituto de Biologia Molecular e Celular, Universidade do Porto, Rua do Campo Alegre 823, 4150-180, Porto, Portugal c PDBEB, University of Coimbra, Portugal

Received for publication 23 May 2006; revised 18 August 2006; accepted 25 August 2006 Available online 1 September 2006

Abstract

Odd-skipped family of proteins (Odd in Drosophila and Osr in vertebrates) are evolutionarily conserved zinc finger transcription factors. Two Osr genes are present in mammalian genomes, and it was recently reported that Osr1, but not Osr2, is required for murine kidney development. Here, we show that in Xenopus and zebrafish both Osr1 and Osr2 are necessary and sufficient for the development of the pronephros. Osr genes are expressed in early prospective pronephric territories, and morphants for either of the two genes show severely impaired kidney development. Conversely, overexpression of Osr genes promotes formation of ectopic kidney tissue. Molecularly, Osr proteins function as transcriptional repressors during kidney formation. We also show that Drosophila Odd induces kidney tissue in Xenopus. This might be accomplished through recruitment of Groucho-like co-repressors. Odd genes may also be required for proper development of the Malpighian tubules, the Drosophila renal organs. Our results highlight the evolutionary conserved involvement of Odd-skipped transcription factors in the development of kidneys. © 2006 Elsevier Inc. All rights reserved.

Keywords: Drosophila; Odd-skipped; Kidney; Repressor; Xenopus; Zebrafish

Introduction by the mesonephros. In these organisms, a metanephros does not develop. The three kidney types differ in their organization, During vertebrate development, three renal structures of but share the same structural unit, the nephron. The number of increasing complexity form successively from the intermediate nephrons varies from 1 to 50 in simple kidneys to a million in mesoderm: pronephros, mesonephros and metanephros (Saxén, the mammalian ones. The nephron is divided in three basic 1987). Each of these develops by an inductive process mediated segments: the corpuscle, the tubules and the duct. The corpuscle by the previous structure. In mammals, the pronephros is not or glomerulus filters the blood, the tubular epithelium is the site functional but is required for mesonephros formation, which of selective re-absorption and secretion and the duct collects and will execute renal functions in the embryo. Later in develop- excretes the urine (Brandli, 1999; Burns, 1955; Saxén, 1987; ment, the mesonephros will be replaced by the metanephros, the Vize et al., 1997). adult functional kidney. In fish and amphibians, the pronephros In Xenopus and in zebrafish (D. rerio), the pronephros is is the functional embryonic kidney, being replaced in the adults formed by a pair of unique non-integrated nephrons, symmet- rically localized in the embryo (Brandli, 1999; Burns, 1955; Saxén, 1987; Vize et al., 1997). Most of the genes necessary for ⁎ Corresponding authors. J.L. Gómez-Skarmeta and F. Casares are to be the formation of the Xenopus and zebrafish pronephros are also contacted at Centro Andaluz de Biología del Desarrollo, Consejo Superior de crucial for the formation of the more complex mammalian Investigaciones Científicas/Universidad Pablo de Olavide, Carretera de Utrera Km1, 41013 Sevilla, Spain. kidneys (reviewed in Carroll et al., 1999; Ryffel, 2003). These E-mail addresses: [email protected] (F. Casares), [email protected] similarities at the molecular level correlate with physiological (J.L. Gómez-Skarmeta). homologies. Thus, the tubules of all nephrons have similar

0012-1606/$ - see front matter © 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ydbio.2006.08.063 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 519 subdivisions along the anterior–posterior axis with an analo- In Drosophila, the odd-skipped (Odd) family of genes gous distribution of transporters of small molecules and ions comprises four transcription factors with high homology in their along this axis (Zhou and Vize, 2004). zinc finger domains: Odd, Sob, Drm and Bowl (Hart et al., In Xenopus, the transcription factors XPax8 and Xlim1 are 1996; Iwaki et al., 2001). odd, sob and drm are similarly the earliest known genes to be expressed in the pronephric expressed in the segment of the gut where midgut–hindgut join, primordium. Their expression in the intermediate mesoderm at and in the ureters of the mature tubules (Ward and Coulter, early neurulation stage precedes any morphological indication 2000), while bowl is expressed along the hindgut (Hart et al., of pronephros formation (Carroll and Vize, 1999; Heller and 1996; Iwaki et al., 2001; Ward and Coulter, 2000). No renal Brandli, 1999). Both genes are essential for tubule and duct tubules phenotype has been described for odd-family mutants. formation (Carroll and Vize, 1999; Chan et al., 2000). Two mammalian odd-skipped related genes, Osr1 and Osr2, Moreover, only the combined overexpression of XPax8 (or have been described (Lan et al., 2001; So and Danielian, 1999). the partially redundant XPax2) and Xlim1 efficiently forms In the mouse, Osr1 expression starts early (E8.0) in the ectopic renal tissue (Carroll and Vize, 1999). Early expression intermediate mesoderm, from where renal structures derive (So of Lim1 and Pax2/8 in the pronephric territory and functional and Danielian, 1999), and is maintained until kidney organo- requirement for at least Pax2 have been reported in zebrafish genesis occurs. Osr2 is activated at stage E9.25 in the (Majumdar et al., 2000; Pfeffer et al., 1998; Toyama and Dawid, mesonephros, and later (stage E14.5) in the mesenchyme that 1997). Consistently with these results, mice lacking Lim1 or surrounds the ducts of the mesonephros and metanephros (Lan Pax2/8 have severe kidney malformations (Bouchard et al., et al., 2001). Osr1 knock-outs lack renal structures (Wang et al., 2002; Porteous et al., 2000; Shawlot and Behringer, 1995; 2005; James et al., 2006), while Osr2 mutants have normal Torres et al., 1995). kidney development (Lan et al., 2004). In Drosophila, the renal (or Malpighian) tubules are the Here we report that both Osr1 and Osr2 function as major excretory and osmoregulatory organs. They originate transcriptional repressors required for pronephros development from the embryonic proctodeum, a posterior region of the in Xenopus and zebrafish. When overexpressed, both lead to ectoderm that gives rise to the hindgut. After specification, they formation of ectopic renal tissue. Moreover, Drosophila Odd proliferate and evaginate from the gut epithelium as four buds, genes may be also necessary for renal tubule formation and can which later extend by cell rearrangement to form the four slim generate renal tissue when overexpressed in Xenopus. There- renal tubules (Jung et al., 2005). In addition, cells from the fore, Odd/Osr genes are utilized to generate filtration organs in caudal visceral mesoderm migrate into the tubules to give rise to both insects and vertebrates. the stellate cells (Denholm et al., 2003). Stellate cells transport water and chloride anions, while the rest of the tubule's cells (so-called ‘principal cells’) transport organic solutes and Materials and methods cations. The transcription factors Kruppel (Kr) and Cut mark the renal tubules primordium within the proctodeum, and both Plasmid constructions are required for normal proliferation and eversion of the renal The following cDNA clones were obtained from the I.M.A.G.E. Lawrence tubules. Still, Kr and cut mutant embryos form uric acid Livermore National Laboratory Consortium: XOsr2 (IMAGE 4405046), zOsr1 excreting cells – therefore with renal tubules characteristics – (IMAGE 7226990) and zOsr2 (IMAGE 7406070). The XOsr1 cDNA clone on the hindgut wall (reviewed in Ainsworth et al., 2000). This (Mochii XL211m14) was a kind gift from N. Ueno and the NIBB/NIG Xenopus suggests the existence of other genes involved in renal tubule laevis EST project. The pCS2-XOsr1 construct was generated by inserting the specification in Drosophila. full-length cDNA into EcoRI site of pCS2+ (Turner and Weintraub, 1994). The pCS2-XOsr2 construct was generated by inserting the full-length cDNA into Despite major differences in embryonic origin, general EcoRI and XhoI sites of pCS2+. To generate the pCS2-zOsr1 and pCS2- organization and physiology, vertebrate kidneys and Droso- zOsr2 constructs, we cloned the corresponding cDNAs into EcoRI and XbaI phila renal tubules share certain developmental and genetic sites of pCS2+. To generate the MT-Osr and Osr-MT constructs, we PCR- aspects. For instance, in Drosophila, renal tubules arise from amplified the corresponding Osr coding regions with the following pairs of ′ ′ ′ the hindgut primordium, which expresses brachyenteron primers: 5 -GAATTCGATGGGGAGCAAGACGCTTCC-3 and 5 - CTCGAGGCATTTGATTTTGGAAGGCTTGAGTTC-3′ for XOsr1;5′- (Singer et al., 1996). Its vertebrate homologue, Brachyury, GAATTCGATGGGCAGCAAAGCTCTTCCAG-3′ and 5′-CTCGA- is required to specify mesoderm and is thus necessary for GAATCGCAATTTCTCCGGAAAACTTTTC-3′ for XOsr2;5′-GAATTCG- kidney development (Technau, 2001). The Kr and cut GAATTAGTCATGGGTAGTAAGACG-3′ and 5′-CTCGAGCTTTATCTTGG homologues Glis2 and Cux-1, respectively, also play a role CTGGCTTGAG-3′ for zOsr1;5′-GAATTCTGCACCGGGAATGG-3′ and 5′- ′ in kidney formation in mammals (Sharma et al., 2004; CTCGAGGACTGTGGCGCCGC-3 for zOsr2. The corresponding EcoRI and XhoI sites are shown in bold. The different PCR fragments were subcloned in Vanden Heuvel et al., 1996; Zhang et al., 2002). The Wnt pGEMT-Easy (Promega) and sequenced. For generating the MT-Osr or the Osr- pathway is required for the specification and proliferation of MT constructs, we cloned the PCR fragments between the EcoRI and XhoI sites the renal tubules (Wan et al., 2000), while Wnt-4 knock-out of pCS2 MT or the pCS2p+MTC2, respectively. These vectors were kindly mice develop small and dysgenic kidneys (Stark et al., 1994). provided by D. Turner. To generate the MT-XOsr-EnR and MT-XOsr-E1A Moreover, hibris, a fly homologue of vertebrate nephrin constructs, we removed a XhoI and SacII fragment containing SV40 polyA region from the MT-XOsr construct and replaced it with a XhoI and SacII (Kestila et al., 1998), is expressed in prospective stellate cells fragment containing the EnR or E1A and the SV40 polyA region. These and is required for their colonizing of the tubules (Denholm fragments were obtained from the pCS2-MT-NLS-EnR and pCS2-MT-NLS- et al., 2003). E1A plasmids kindly donated by N. Papalopulu. The complete open reading 520 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 frame (ORF) from the Drosophila drm gene was excised with XhoI and XbaI odd and uncovers approximately 30 predicted genes, including drm, sob and from the drm cDNA and cloned into the XhoI and XbaI sites from pCS2+ vector odd. Mutant chromosomes were balanced over the 2nd marked balancer to generate the drm construct. To generate the odd construct, we amplified the chromosomes CyO, Kr-GFP; homozygous mutant embryos were detected as ORF from the Drosophila odd gene with the following primers: 5′- GFP-negative. To trace the lineage of odd-expressing cells in the RTs, we GAATTCAATGTCTTCCACATCGGCCTC-3′ and 5′-TCTAGA- crossed odd-Gal4 (a Gal4 insertion in odd that faithfully recapitulates its TATCTGCTCATGATCTCATCGATG-3′. The PCR fragment was subcloned expression, gift from M. Calleja and G. Morata, CBM, Madrid) into UAS-flip; into pGEMT-Easy (Promega), sequenced and then transferred to pCS2 MT act>Draf>LacZ (Campbell and Tomlinson, 1998). In the resulting odd-Gal4/ between the EcoRI and XbaI sites. The oddΔeh1 construct was generated by UAS-flip; act>Draf>LacZ cells derived from odd-expressing cells are subcloning an EcoRI–XhoI fragment from the Drosophila odd gene into the constitutively marked by the expression of lacZ. The expression of the odd EcoRI and XhoI sites from the pCS2+ vector. This fragment encodes a lineage (odd>lineage) was compared to the actual expression of UAS-lacZ; odd- truncated Odd protein that lacks the last 19 amino acids (SSEKPKRMLGFTI- Gal4 larvae. DEIMSR), which include the eh1 domain (underlined). To overexpress Odd-family genes in the RT primordial and hindgut, we used a brachyenteron (byn)-Gal4 driver (Iwaki et al., 2001). byn-Gal4/TM3, ftz-Z DNA sequencing females were crossed to homozygous UAS-bowl (de Celis Ibeas and Bray, 2003), UAS-sob or UAS-odd/TM6B (Hao et al., 2003)orUAS-drm (Green et al., β DNA sequencing was performed with ABI chemistry in an automatic DNA 2002) males. Embryos carrying byn-Gal4 were detected as LacZ ( - sequencer using T3 and T7 oligonucleotides. Custom synthesized oligonucleo- galactosidase)-negative. Those expressing odd were detected using an anti- tides were obtained from Sigma. Odd specific antibody. UAS-src-GFP is described in Kaltschmidt et al. (2000).

Xenopus, zebrafish and Drosophila in situ hybridization, X-Gal and Results antibody staining Osr genes are expressed in the renal primordium of Xenopus Antisense RNA probes were prepared from cDNAs using digoxigenin or and zebrafish fluorescein (Boehringer Mannheim) as labels. Xenopus, zebrafish and Droso- phila specimens were prepared, hybridized and stained as described (Hao et al., 2003; Harland, 1991; Jowett and Lettice, 1994). Xenopus and Drosophila X- A search in databanks identified two X. laevis and two Gal staining was performed according to Coffman et al. (1993). Xenopus, zebrafish EST clones that correspond to genes encoding zebrafish and Drosophila antibody staining was performed as described orthologues of human and mouse Osr1 and Osr2 (Supple- (Gómez-Skarmeta et al., 2001; Hernandez et al., 2005; Sanchez-Herrero, mentary Fig. 1). We named these genes XOsr1, zOsr1, β 1991). Antibodies used in this study were rabbit anti- -galactosidase (Cappel), XOsr2 and zOsr2. No additional Osr genes were detected by rabbit anti-GFP (Molecular Probes) and guinea pig anti-Odd (Kosman et al., 1998). The monoclonal antibody 3G8 were kindly provided by E. Jones. The Blast searches in these species, suggesting that, as in monoclonal antibodies 12/101 and 2B10 (developed by J. P. Brockes) and mammals, they have two Osr genes. Both XOsr genes are Mouse anti-Cut (developed by I. Rebay, G. Dailey, K. Lopardo and G. Rubin) initially detected during early gastrulation in the involuting were obtained from the Developmental Studies Hybridoma Bank developed mesoderm and endoderm (Figs. 1A, E). At the end of under the auspices of the NICHD and maintained by The University of Iowa, gastrulation, XOsr2 is detected in the intermediate mesoderm Department of Biological Science, Iowa City, IA 52242. (inset in Fig. 1F) preceding the activation of the early In vitro RNA synthesis and microinjection of mRNA and morpholinos pronephric markers XPax8 and Xlim1 (not shown and inset in Fig. 1I). During neurulation, this expression resolves in a All DNAs were linearized and transcribed as described (Harland and broad domain largely overlapping that of Xlim1 and XPax8 Weintraub, 1985) with a GTP cap analog (New England Biolabs), using SP6, (Figs. 1F, G, I–K, M–O) (Carroll and Vize, 1999). XOsr1 is T3 or T7 RNA polymerases. After DNAse treatment, RNA was extracted with similarly expressed although at lower levels (Figs. 1B, C). phenol-chloroform, column purified and precipitated with ethanol. mRNAs for During tailbud (stage 35), XOsr1 is expressed in the rectal injection were resuspended in water. X. laevis and Xenopus tropicalis embryos were injected in the marginal region at the 2-cell stage using a volume of 10 or diverticulum and in the ducts (Fig. 1D). At this stage, XOsr2 2–5 nl, respectively. V2.2 blastomeres of X. tropicalis 8–16 cell stage mRNA is also expressed in the duct but in a broader domain. embryos were injected with 1–2 nl of morpholino solution. In these In addition, XOsr2 is also expressed in the tubules (Fig. 1H). experiments, embryos were co-injected with Dextran-Fluorescein (10,000 See for comparison the expression of Xlim1 and XPax8 in the MW, Molecular probes). Embryos showing fluorescence in the prospective tubules and duct at this stage (Figs. 1L, P). kidney domain but not in the somites were selected under a fluorescent dissecting scope and further processed for in situ hybridization. The In zebrafish, zOsr1 also precede the expression of the early localization of Fluorescein was later determined with anti-Fluorescein antibody pronephros marker zlim1 and zPax2, while the expression of coupled to alkaline phosphatase (Roche). The following morpholinos were zOsr2 appears at the 8-somite stage, once zlim1 and zPax2 used in this study: MOXOsr1: 5′-TGCTGGAAGGGTCTTGCTCCCCATC-3′, are transcribed but prior to any sign of pronephros ′ ′ ′ MOXOsr2: 5 -GGCTGGAAGAGCTTTGCTGCCCATT-3 ,MOzOsr1:5- histogenesis (see Fig. 2 for a full description of zOsr1 and GCGTCTTACTACCCATGACTAATTC-3′ and MOzOsr2: 5′-AGAGTCT- TACTGCCCATTCCCGGT-3′. The Xenopus morpholinos were designed to zOsr2 expression patterns). The staggered expression of target Osr1 or Osr2 genes from both X. laevis and X. tropicalis. X. tropicalis Osr1 and Osr2 we observe in zebrafish is similar to that embryos were injected with 10–20 ng of morpholinos at the two cell stage and recently described in mice and chicken (So and Danielian, with 2 ng at the 8–16 cell stage. Zebrafish embryos were injected in the yolk at 1999; Lan et al., 2001; Wang et al., 2005; James et al., 2006; – – 1 2 cell stage with 10 20 ng of morpholinos. Stricker et al., 2006). This situation is reversed in Xenopus, where Osr2 expression in the prospective renal territory seems Drosophila strains and genetic manipulations to precede that of Osr1, even if both genes are co-expressed The following mutant alleles are described in Flybase (http://flybase.org/): by the time the early renal markers XPax8 and Xlim1 begin to odd5, bowl1, drm6. Deficiency drmP2 (Green et al., 2002) deletes from tim to be expressed. J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 521

Fig. 1. Expression pattern of XOsr genes. Panels A, E are vegetal and panels B–D, F–P are lateral views. Insets in panels A, C, D, E, G, H, L, M and N are transverse vibratome sections through the dashed lines in the main panels. (A) Early gastrula stage (stg). XOsr1 is expressed in the involuting mesoderm and endoderm (arrowhead and arrow in inset, respectively). (B, C) During neurulation, XOsr1 mRNA is detected in the pronephric territory. (D) At tailbud, XOsr1 is expressed in the ducts (arrowhead in inset) and in the rectal diverticulum (arrow). (E–G) Expression of XOsr2 is similar, but stronger. In the prospective kidney territory, XOsr2 is detected earlier than XOsr1 (stage 11.5–12, inset in panel F; arrowhead marks the prospective kidney domain), and earlier than other pronephric markers (see inset in panel I for the expression of Xlim1 at this stage; arrowhead marks the prospective kidney domain). (H) At tailbud, XOsr2 is expressed in the tubules (arrow) and in a broad domain adjacent to the ducts (arrowhead in inset). (I, J, M, N) During neurula, expressions of XOsr2 and Xlim1 (I, M) largely overlap in the pronephric region (double in situ hybridization, J, N). (K, O) During neurula, XPax8 is also detected in the pronephric territory. (L, P) At tailbud, Xlim1 and XPax8 are expressed in the tubules and ducts. Inset in panel L show Xlim1 expression in the duct (arrowhead).

Morpholino knockdown of Osr1 and Osr2 impairs renal respectively; Figs. 3J–L, N–P and not shown). To avoid development in Xenopus and zebrafish possible kidney defects caused by altered muscle development, we co-injected the XOsr MOs with Dextran-Fluorescein in a In mouse, Osr1, but not Osr2, is essential for kidney single blastomere (V2.2) of 8–16 cell stage embryos, and development (Lan et al., 2004; Wang et al., 2005). We have analyzed tailbud-stage embryos showing Fluorescein signal examined whether Osr genes are required for pronephric restricted to the kidney territory, but not in the somites. In these development in X. tropicalis and zebrafish by blocking the embryos, injection of XOsr1 or XOsr2 MOs promoted a clear translation of Osr1 and Osr2 mRNAs with specific morpholinos downregulation of XSGLT1K and XNKCC2 (Figs. 3Q–T), two (MOs) (Supplementary Fig. 2A). genes encoding pronephric epithelial transporters that are X. tropicalis embryos injected with 10–20 ng of XOsr1 or specifically expressed in the proximal and distal tubule, XOsr2 MOs show similar downregulation of the early pro- respectively (Zhou and Vize, 2004), without any visible effect nephric territory markers Xlim1 and XPax8 (84% and 71%, on somites formation. n=68 and 66, respectively; Figs. 3A–I, M). This down- In zebrafish, MOs targeting zOsr1 or zOsr2 genes caused regulation was not associated with an expansion of muscle downregulation of the early pronephric markers zlim1 (76% and tissue as determined by the muscle specific antibody 12/101 46%, n=85 and 93, respectively; Figs. 4A, E, I) or zPax2.1 (Figs. 3A–I, M). Indeed, in some cases, muscle size was reduced (73% and 38%, n=81 and 77, respectively; Figs. 4B, F, J) and (see Fig. 3M). Moreover, a strong defect in, or the disappearance induced defects in the differentiated renal structures (Figs. 4C, of, the differentiated embryonic kidneys was observed, as G, K). The observed downregulation of zlim1 at 4-somite stage, determined by the pronephros monoclonal antibody 3G8 (Vize though, was more pronounced in MOzOsr1 morphant embryos et al., 1995) (92 and 78% with reduced kidneys, n=175 and 166, (compare Figs. 4E and I). In addition, at 72 hpf these morphant 522 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531

Fig. 2. Expression pattern of zOsr genes. Dorsal views are shown, except (A), a vegetal view and (K, O) and insets and (I, M, P), transverse sections through the pectoral fin buds or the posterior spinal cord, respectively. (A) At shield stage, zOsr1 mRNA is detected in the shield and in a ventro-lateral ring. (B, E, F) At tailbud, zOsr1 (B) is expressed in the pronephric territory (arrowhead), preceding the expression of zlim1 (E) and zPax2.1 (F) (arrowheads mark the prospective pronephric territory at this stage). (C, G, H) During early somitogenesis, zOsr1 (C), zlim1 (G) and zPax2.1 (H) show similar expression domains within the pronephros territories, although zOsr1 seems to extend more rostrally. (D, L) At the 8-somite stage, both zOsr1 (D) and zOsr2 (L) mRNAs are expressed in the pronephros, zOsr2 being a transcription domain weaker and shorter. In addition, zOsr2 also shows a weak generalized expression. (I, M) At 24 hpf, the expression of zOsr1 in the pronephros starts to be downregulated (I, red arrowhead). At this stage, zOsr2 is detected in the tubules and in the anterior duct (M, red arrowhead). In addition, zOsr1 is expressed in two rows that run parallel to the pronephros (I, black arrowheads and inset) while zOsr2 is found in the gut (M, black arrowheads and inset). (J, K, N, O) Expression of zOsr1 (J, K) and zOsr2 (N, O) at 60 hpf. zOsr1 is detected in the glomerulus, in some patches in the eye and brain, and weakly in the pectoral fin buds (J). The expression in the glomerulus is clearly visible in a transverse section (arrowhead in panel K). zOsr2 is expressed in the tubules and the pectoral fin buds (N). The expression in the tubules is more evident in a transverse section (arrowhead in panel O). (P) Expression of zPax2 at 24 hpf. The expression in the pronephros is pointed at by an arrowhead and can be visualized in a transverse section in the inset. embryos showed pericardial edemas and kidney cysts (Figs. 4D, tagged versions of either Xenopus or zebrafish Osr1 and Osr2 H, L), defects characteristic of renal failure (Drummond et al., mRNAs yielded similar results. Many of the Xenopus Osr- 1998; Hostetter et al., 2003). injected embryos showed gastrulation defects that were the In zebrafish and Xenopus, we have compared the effect of more severe the higher the doses of mRNA. However, in targeting both Osr genes at the same time (with half the dose of embryos injected with 100 pg of mRNA, about 30% showed no each MO) with reducing individual Osr gene function. No gastrulation defects. In most of these (75%, n>200; Figs. 5A– synergistic effect was observed by reducing Osr1 and Osr2 F), Xlim1 and XPax8 were expressed in ectopic patches of cells. function simultaneously (not shown). Thus, in contrast to mice Other pronephric markers such as XNHF1β or Gata3 were (Lan et al., 2004; Wang et al., 2005; James et al., 2006), in similarly ectopically expressed, but not the glomerulus marker Xenopus and zebrafish, both Osr genes seem to be required XWt1 (not shown). We also examined the effect of Osr for development of kidney structures. overexpression on genes encoding pronephric epithelial transporters. Late neurula injected embryos showed ectopic Osr1 and Osr2 gain of function promotes ectopic renal tissue patches of XSGLT1 and XNKCC2 expression at similar frequencies (Figs. 5G, H). These patches differentiate as We next examined the effects of overexpressing Osr genes pronephric structures later, as determined by the tubule-specific on Xenopus kidney development. Either wild-type or Myc- monoclonal antibody 3G8 (Figs. 5I, J). Morpholino-insensitive J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 523

Fig. 3. Xenopus Osr morphant embryos have severely impaired kidneys. (A–H) Lateral views of stage 25 Xenopus tropicalis embryos injected with 20 ng of MOXOsr1 (A–D) or 20 ng of MOXOsr2 (E–H) and 300 pg of LacZ mRNA to determine the injected side. Purple staining shows the expression of Xlim1 (A, B, E, F) or XPax8 (C, D, G, H), and brown staining the somitic muscles, labeled with the monoclonal antibody 12/101. The MO injected embryos show a reduced expression of the kidney markers on the injected sides (arrows in panels B, D, F and H; compare with the control sides shown in panels A, C, E and G). (I, M) Transverse section of stage 25 MOXOsr1 (I) or MOXOsr2 (M) injected embryos triple labeled for Xlim1 (pronephros, purple), muscles (brown) and Sox2 (neural tissue, cyan). Note the strong reduction of the pronephric tissue in the injected sides (arrows). (J–L and N–P) Stage 37 Xenopus tropicalis embryos injected with MOXOsr1 (J–L) or MOXOsr2 (N–P) and stained with the monoclonal antibody 3G8. Note the strong reduction of the kidney tissue in the injected sides (arrows in panels K, L, O, P). Insets are closer views. This reduction is clearly visible in transverse sections (arrow in panels L and P). (Q–T) Lateral views of stage 35 Xenopus tropicalis embryos co-injected with MOXOsr1 (Q, R) or MOXOsr2 (S–T) and Dextran-Fluorescein in the V2.2 blastomere at the 8–16 cell stage. The expression of XSGLT1K (Q, S; purple) and XNKCC2 (R, T; purple) is impaired in the injected side (Fluorescein distribution is visible in cyan). Brown staining in panels Q and T shows the somitic muscles labeled with the monoclonal antibody 12/101. Insets show the control un-injected side. (U) Target sequences for Xenopus Osr Morpholinos (MOs). In all sequences, the first methionine of the corresponding gene is underlined. Identical bases are in blue and mismatches in red. Note that the MOs for each Xenopus Osr gene have one mismatch with the corresponding Xenopus laevis and Xenopus tropicalis target sequences. In contrast, the MO against one of the paralogues has five or more mismatches with the sequence of the other gene. MOs with only one mismatch can efficiently block translation while five or more mismatches make an MO inactive. 524 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531

Fig. 4. Zebrafish Osr morphant embryos have severely impaired kidneys. (A–D) Wild-type zebrafish embryos. Embryos injected with 20 ng of MOzOsr1 (E–H) or 20 ng of MOzOsr2 (I–L). These injected embryos show a reduction of the pronephric markers zlim1 at the 4-somite stage (A, E, I; arrowheads), zPax8 at 24 hpf (B, F, J; arrowheads) and reduced kidney tissue at 48 hpf, as determined by the staining with the monoclonal antibody 3G8, which labels tubules and anterior ducts (C, G, K; arrowheads). At 70 hpf, pericardial edemas (arrowheads) and kidney cysts (arrow) are visible (D, H). These are characteristic of renal failure. (M) Target sequences for zebrafish Osr Morpholinos (MOs). As in Fig. 3 in all sequences, the first methionine of the corresponding gene is underlined. Identical bases are in blue and mismatches in red.

MT-XOsr1 and MT-XOsr2 could also rescue the impaired to downregulate target genes during embryonic segmentation development of renal tissue of embryos injected with XOsr1 (Goldstein et al., 2005). Therefore, in this context, Odd and XOsr2 MOs, respectively (50% or 37% with rescued proteins work as repressors. In contrast, the molecular function kidneys, Figs. 5K, L). We also examined the effect of of mammalian Osr proteins is unclear. Osr2 mRNA generates overexpressing Osr genes in transverse sections of stage 22– two protein splicing variants, one containing three zinc fingers 25 embryos triply stained for pronephros, somitic muscle and and the other five, that function as activator and repressor, neural ectoderm. The ectopic renal tissue was always found respectively, in cell culture assays (Kawai et al., 2005). To close to the neural tube, which in some cases was strikingly further investigate this question, we injected X. laevis embryos enlarged in the direction of the ectopic pronephros. The somitic with mRNAs (100 pg) encoding Osr proteins fused to either muscles were normal or slightly reduced (Figs. 5M, N). At stage the Engrailed repressor domain (EnR) or the E1A activation 38, we also observed a clear enlargement of the endogenous domain. Similarly to wild-type Osr mRNAs, XOsr1-EnR or renal tissue and ectopic pronephric structures in the proximity of XOsr2-EnR mRNAs induced patches of ectopic expression the spinal cord (Figs. 5O, P). of Xlim1 and XPax8 (Figs. 6A, B and not shown) that In zebrafish embryos, both Osr mRNAs promoted enlarge- differentiated into renal tissue (Fig. 6C). In contrast, XOsr1- ment of the pronephric domains of zlim1 and zPax2.1 markers E1A or XOsr2-E1A mRNAs (500 pg) downregulated Xlim1 (Figs. 5Q–S and not shown). At later stages, the differentiated and XPax8 and strongly reduced differentiated kidney kidney tissue was also expanded (Fig. 5T). In addition, some structures (Figs. 6D–F, and not shown). Thus, vertebrate Osr embryos displayed ectopic renal tissue (Fig. 5T). proteins appear to act as transcriptional repressors during kidney development. Osr proteins function as transcriptional repressors during The zinc fingers of Drosophila and vertebrate Odd/Osr renal development proteins are largely identical in sequence, although the number of zinc fingers varies among them. Vertebrate Osr proteins Two Drosophila Odd proteins, Odd and Bowl, harbor an contain three (except the mammalian Osr2A splice variant that eh1-like motif that helps recruiting the Groucho co-repressor contains five), while Drosophila Drm contains two, Odd four J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 525

Fig. 5. Overexpression of Osr genes promotes ectopic kidney development. (A–L) Lateral views of stage 25 (A–F), stage 30 (G, H) or stage 37 (I–L) Xenopus embryos, or 48 hpf zebrafish embryo (T). (M–P) Transverse sections of stage 25 (M, N) or stage 37 (O, P) Xenopus embryos. (Q–T) Dorsal views of four somites (Q) or 24 hpf (R, S) zebrafish embryos. Embryos were injected with 50–100 pg of Xenopus or zebrafish Osr mRNAs. Xenopus embryos were co-injected with 300 pg of LacZ mRNA as a lineage tracer. (A–D) Embryos injected with XOsr1 mRNA showed ectopic patches of Xlim1 (A, B) or XPax8 (C, D) expression in the injected sides (arrowheads in panels B, D). In addition, many embryos have enlarged pronephros (arrows in panels B and D; compare with control sides in panels A and C). (E, F) Stage 25 Xenopus embryos injected with XOsr2 (E) or zOsr1 (F) mRNAs and doubly hybridized for Xlim1 and XPax8. The first chromogenic reaction, to detect Xlim1 expression, is shown in the main panels (cyan), and the second chromogenic reaction, to detect XPax8, in the insets (purple). Note that the same cells express ectopically both markers (arrowheads). (G, H) Embryos injected with 100 pg of Xenopus Osr1 mRNA showed ectopic patches of XSGLT1K (G, arrowheads) and XNKCC2 (H, arrowhead). Note that these embryos have gastrulated properly. (I, J) Enlarged (arrow) and ectopic (arrowhead) kidney tissue, as determined by 3G8 staining, in stage 37 Xenopus embryos injected with XOsr1 (I) or XOsr2 (J) mRNAs. Insets show magnification of ectopic renal tissue in other injected embryos. (K, L) Stage 37 Xenopus embryos co-injected with MOXOsr1 and MTXOsr1 mRNA (K) or MOXOsr2 and MTXOsr2 mRNA (L) and stained for 3G8 monoclonal antibody. Note that these MO insensitive mRNAs rescue the MO-induced kidney marker reduction (arrow) (see panels K and O in Fig. 3 for comparison) and promote ectopic renal tissue (arrowhead). (M–P) Transverse sections on stage 25 (M, N) or stage 37 (O, P) Xenopus embryos injected with XOsr1 (M, O) or XOsr2 (N, P) mRNAs. The embryos in panels M and N show a triple staining for Xlim1 (pronephros, purple), monoclonal antibody 12/101(somitic muscles, brown) and Sox2 (neural tube, magenta). The embryos in panels O, P show differentiated kidneys labeled with the monoclonal antibody 3G8. Note that the ectopic renal tissue is always found close to the neural tube (arrowhead). In addition, these embryos show a clear enlargement of the neural tube and the endogenous pronephros (arrows). (Q–T) Zebrafish embryos injected with zOsr1 (Q, R) or zOsr2 (S, T) mRNAs showing zlim1 expression at 4-somite stage (Q), zPax2.1 expression at 24 hpf (R, S) and differentiated renal structures, as determined by 3G8 monoclonal antibody staining (T). Note the enlarged pronephros (arrowheads) and the ectopic renal tissue (T, arrow). Insets in panels Q, R and T show control non-injected embryos. 526 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531

Fig. 6. Osr proteins function as repressors during kidney development. All panels show lateral views of late neurula (left and middle panels) or tailbud (right panels) Xenopus embryos. The left, middle and right panels show Xlim1, XPax8 and 3G8 staining, respectively. Cartoons on the left indicate the proteins encoded by the injected mRNAs. (A–C) Injection of 100 pg of XOsr2-EnR mRNA promotes ectopic pronephros (arrowheads). In contrast, injection of 500 pg of XOsr2-E1A mRNA downregulates pronephric markers (D–F). Inset in panel F shows the control non-injected side. (G–I) Overexpression of Drosophila odd mRNA (500 pg) promotes ectopic renal tissue (arrowheads). This activity depends on its eh1 domain (orange) as the deletion of this motif (oddΔeh1) impairs its ability to activate renal markers (J–L). (M–O) Drosophila drm mRNA (1 ng) is unable to promote kidneys when overexpressed in Xenopus. and Bowl and Sob five. In addition, Drosophila Odd and Bowl Drosophila Odd genes are expressed during RT formation and function in some contexts as repressors by recruiting Groucho, may be required for their formation but Sob and Drm do not bind this co-repressor (Goldstein et al., 2005). We examined whether Drosophila Odd proteins promote The ability of odd to promote renal tissue in Xenopus ectopic kidney differentiation in Xenopus. odd, but not drm prompted us to determine whether this family of genes is mRNA, promoted ectopic renal tissue (Figs. 6G–I, M–O). This required for renal tubules formation in Drosophila. We re- ability depended on the eh1 domain as its removal abolished it examined the expression of the different Odd genes in (Figs. 6J–L). These data strongly suggest that, to function in embryogenesis. The similar expression of odd, drm and sob renal development, vertebrate Osr proteins may also require a in the gut suggested that the three genes might also be Groucho-like co-repressor. expressed in the renal tubules ureters. This was the case, as J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 527 detected by coexpression with Cut along the proximal ureteric When overexpressed, none of the four odd genes induced tubes (Figs. 7A–C, I). This expression was detected at least or expanded the renal tissue. Only the overexpression of odd from embryonic stage 12 as a stripe of cells at the base of the (Figs. 7I–K), and to a lesser extent that of sob (not shown), budding RT primordia (not shown). We did not detect bowl resulted in a widening and shortening of the tubules and larger transcription at significant levels in the Cut-expressing cells at ureters, consistent with an alteration of tubule extension. Our any stage. results suggest that drm, odd and sob may be required for The expression pattern of drm, sob and odd argues against proper renal tubules development. This requirement is likely a role in early stages of renal tubules specification, but suggests a redundant function later in renal tubules development. To test this hypothesis, we examined the renal tubules in odd, drm and bowl single mutants, and in embryos homozygous for a large deficiency (DfdrmP2), that uncover at least 30 predicted genes, including drm, sob and odd (Green et al., 2002). (No sob single mutation is available, which prevented analysis of its individual mutant phenotype.) None of the three individual mutants affected renal tubules specification, growth or extension. Nevertheless, in DfdrmP2 embryos, renal tubules were singled out as Cut- expressing buds, but failed to grow or extend further (Fig. 7D). The secretory activity of the remaining rudiments in these mutant embryos, monitored by the production of uric acid, was greatly reduced when compared with normal tubules (Figs. 7E, F). The general defective development of renal tubules was not anticipated by the localized expression of odd genes in just the ureters primordia. If odd, sob and drm genes were indeed responsible for the phenotype observed in DfdrmP2 embryos, this might be explained if odd genes were controlling the production of non-autonomous growth signals. In addition, the odd-expressing cells could contribute to the tubules them- selves. We tested the second possibility by following the lineage of odd-expressing cells by using a lineage tracing system (see Materials and methods). While in odd>lacZ larvae, X-gal positive cells were confined to the ureters, in odd>lineage-lacZ embryos, positive cells were found along the distal tubules, indicating that drm/sob/odd-positive ureteric cells give rise to tubule cells that lose expression of odd genes (Figs. 7G, H).

Fig. 7. Odd genes expression and requirement for renal tubule development in Drosophila. (A) Schematic representation of the late embryonic RTs. The domains of expression of cut (orange) and of odd, sob and drm (blue) are shown. cut and drm/sob/odd overlap in the ureters, shown in gray. Mg: midgut; hg: hindgut; rt: renal tubules; u: ureter. (B, C, I) Expressions of sob (B), drm (C) and odd (I) are similar and co-localize with Cut in the ureters (arrows). (B, C) sob and drm expression is detected by in situ hybridization (purple) and that of cut by immunohistochemistry (orange). Overlap is seen as dark gray. (I) Odd (green) and Cut (red) expression is detected by immunofluorescence. Overlap is seen in yellow. Ureters are marked by arrows. (D) DfdrmP2 mutant embryo (labeled as drm− sob− odd−), showing rudimentary Cut-expressing tubules. (E, F) Uric acid excretion in wild-type (E) and DfdrmP2 (F) late embryos, observed under phase contrast optics. (G, H) Histochemical X-Gal staining of RTs of odd>LacZ (G) and odd>lineage (H; see Materials and methods) L3 larvae. Nuclei of X-Gal positive cells (blue) are seen along the distal tubules in odd>lineage (arrows; H) but not in odd>lacZ tubules. (J) byn>odd late embryo, co-stained for Odd (green) and Cut (red). Tubules (red) and ureters (yellow) are wider, and tubules are shorter. (K, L, M) Early stage 13 byn>srcGFP (green) embryo, co-stained for Cut (red). The Cut- positive RT buds are included within the byn domain. (*) marks the Cut- expressing posterior spiracles in all panels. 528 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 to occur after the Malpighian tubule primordia have been ectopic renal tissue. Osr proteins activate Pax2/8, which can specified. downregulate Wt1 (Majumdar et al., 2000). Therefore, strategies devised at inducing functional renal tissue by Discussion making use of Osr expression should overcome this problem. A transient Osr expression might solve it as it would allow Osr1 and 2 genes function at the top of the genetic hierarchy early specification of the whole pronephric primordium, and controlling pronephric development not interfere with the later formation of the glomerulus. Our results showing that Osr genes can drive the Here we show that the two paralogues Osr1 and Osr2 are development of ectopic pronephros, together with the expres- expressed at early stages in the intermediate mesoderm. Osr1 in sion and functional data, suggest that they lay atop the kidney zebrafish and Osr2 in Xenopus are first detected before the genetic program. Nevertheless, only the dorsal region of the earliest markers of kidney development. This is similar to what embryo was competent to develop ectopic renal tissue upon Osr was described for mouse Osr1 (So and Danielian, 1999; Wang mRNA injection. Similar results were found with Xlim1 and et al., 2005; James et al., 2006). However, in contrast to the Pax8 co-injection experiments (Carroll and Vize, 1999). In situation found in mammals, where Osr2 seems dispensable for chick embryos, a gradient of BMP activity patterns mesodermal kidney development (Lan et al., 2004), our morpholino fates with highest signaling levels at the lateral mesoderm experiments indicate that both Osr1 and Osr2 are required for inhibiting intermediate fates, including Osr1 expression and proper pronephros development in Xenopus and zebrafish. In renal development (James and Schultheiss, 2005). Intermediate Xenopus, both genes are coexpressed just at the time the levels would allow acquisition of intermediate mesoderm fates pronephros territory is being defined, as determined by the indirectly, through the relief of a transcriptional repressor expression of Xlim1 and Pax8. This is consistent with both activity on intermediate mesoderm genes, such as Osr1 (James genes being required for the early specification of the kidney and Schultheiss, 2005). Therefore, intermediate and medial anlage. The fact that we do not detect synergistic defects when (dorsal) regions would be competent to develop renal tissue. impairing simultaneously both genes indicates that Osr1 and This coincides with the regions in Xenopus where Osr mRNA Osr2 are required additively for this specification. In contrast, injection widens the endogenous pronephros or induces ectopic in zebrafish, Osr1 precedes the activation of early kidney pronephric tissue. There was no correlation between ectopic markers while the onset of Osr2 expression is delayed until the pronephros and muscle loss (a derivative of dorsal mesoderm), 8-somite stage, when the early kidney markers are already arguing against a muscle-to-kidney transformation. In zebrafish, activated but still there is no histological sign of kidney tissue injected zOsr mRNAs enlarged the pronephros, but only (Drummond et al., 1998). In mice, the onset of Osr2 is further occasionally induced ectopic tissue. This suggests strong delayed, only appearing at the 18-somite stage, when restrictions in the competence of the dorsal mesoderm in mesonephros are already differentiating (Lan et al., 2001). zebrafish. The degree of delay in the activation of Osr2 expression The ectopic renal tissue was patchy, while the distribution of correlates with the functional requirement of these genes: while ectopic Osr protein was broader and continuous (not shown). in zebrafish knockdown of Osr2 mildly affects the expression Possibly, a lateral inhibition process prevents subsets of Osr of early pronephric markers, but severely impairs differentiation expressing cells from differentiating as kidney tissue. Evidently, of the kidney, the knock-out of Osr2 in mice seems not to have some sort of signaling, of unknown nature, occurs between the any effect (Lan et al., 2004). Recent experiments show that the ectopic developing pronephros and the neighboring cells, as overexpression of Osr1 is able to induce ectopic kidney shown by the neural tube overgrowths associated with the markers in chicken (James et al., 2006). It will be interesting ectopic renal tissue. to assay if overexpression of Osr2 can also promote kidney formation in chick to determine whether the Osr2 gene of Osr proteins act as transcriptional repressors during kidney higher vertebrates retains the functional capabilities of its development paralogue Osr1. In both Xenopus and zebrafish, the expression of both Native XOsr proteins and the constitutive repressors XOsr- genes diverges during pronephros formation, one paralogue EnR similarly induced ectopic kidney tissue in Xenopus, which being expressed in more proximal segments than the other. indicates that vertebrate Osr proteins function as transcriptional Thus, Osr genes may provide distinct late functions during repressor in vivo. Drosophila Odd also acts as a transcriptional pronephric organogenesis. This functional diversification repressor during embryonic segmentation by directly binding to seems to have proceeded further in the lineage leading to the Groucho co-repressor (Goldstein et al., 2005). This inter- mammals as Osr1 has an additional role in heart development action occurs through, and requires the C-terminal “engrailed (Lan et al., 2004). homology 1” (eh1) motif. In Xenopus, we show that odd mRNA The knockdown of Osr1 and Osr2 results in the loss of all also induces ectopic nephrogenesis, an ability that depends on pronephric structures, including the glomerulus. However, the eh1 domain. This suggests that some member(s) of the their ectopic expression activates several early and late vertebrate Transducin-like Enhancer of Split (TLS) family of markers, but not the glomerulus-specific marker Wt1 (not Groucho homologues (Chen and Courey, 2000) is recruited by shown). Hence, this structure seems to be missing in the Odd. Moreover, the repressor activity of vertebrate Osr products J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 529 may similarly require interaction with TLS co-repressors. molecular cassette engaged in forming and/or patterning tubular Indeed, we identified a putative eh1 motif in vertebrate Osr1 organs, as they do during foregut, hindgut (reviewed in Lengyel and Osr2 (Supplementary Fig. 1) that is located N-terminal to the and Iwaki, 2002) and renal tubules (this work) development in zinc fingers, instead of at the C-terminal end as in Odd. Drosophila, or nephron formation in vertebrate kidneys. That the activation of the kidney genes Pax8 and lim1 requires the repressor activity of Osr proteins implies the Acknowledgments existence of at least one additional intermediate repressor in the cascade. Foxc1 and/or Foxc2 are possible candidates. These We are most grateful to M. Allende, T. Becker, M. Brand, S. transcription factors are required for somites development Bray, M. Chung, I. Dawid, E. Jones, N. Papalopulu, C. (Topczewska et al., 2001) and are necessary and sufficient to Rauskolb, D. Turner, P. Vize, F. Serluca for reagents. We repress intermediate mesoderm markers, such as Osr1 and lim1 specially thank T. Becker, J. Castelli-Gair, J. Modolell, P. Vize (Wilm et al., 2004). Still, we do not favor this hypothesis as and anonymous referees for helpful criticisms. We also thank N. Osr1 and Osr2 morphants did not show expanded somites Ueno and the NIBB/NIG X. laevis EST project for the Mochii associated to the loss of pronephros. clone XL211m14. This work was supported by grants from: Spanish Ministry of Education and Science (BFU2004-00310 Odd genes are expressed in the renal organs of Drosophila and and GEN2001-4846-C05-01) to J.L.G-S. and BMC2003-06248 may be required for their development to F.C., and a Junta de Andalucía Proyecto de Excelencia (CVI260) to J.L.G-S. Ana Neto is funded by Fundaçao para a We find that Odd genes may also be required for the Ciencia e Tecnologia from Portugal (SFRH/BD/15242/2004) development of the renal organs of Drosophila. drm, odd and and belongs to the Graduate Program in Areas of Basic and sob are expressed in the ureters of the Malpighian tubules, Applied Biology from Oporto University. and embryos homozygous for a deficiency that removes at least 30 predicted genes, including drm, odd and sob,form rudimentary renal structures with impaired excretory activity, Appendix A. Supplementary data a phenotype reminiscent of that seen in Kr and cut mutants (Harbecke and Janning, 1989). While this may reflect a Supplementary data associated with this article can be found, requirement for these genes in Drosophila renal development, in the online version, at doi:10.1016/j.ydbio.2006.08.063. other genes within this deficiency may also contribute to the phenotype. We have found that cells born in the drm/sob/odd expression domain are incorporated into the tubules. Neverthe- References less, a failure in this cell contribution does not seem to explain the dramatic reduction of Cut cells in the DfdrmP2 mutants and Ainsworth, C., Wan, S., Skaer, H., 2000. Coordinating cell fate and therefore it is likely that, in these embryos, an additional cell morphogenesis in Drosophila renal tubules. Philos. Trans. R. Soc. Lond., – non-autonomous growth signal is defective. B. Biol. Sci. 355, 931 937. Bouchard, M., Souabni, A., Mandler, M., Neubuser, A., Busslinger, M., 2002. Renal organs, in charge of nitric waste excretion and Nephric lineage specification by Pax2 and Pax8. Genes Dev. 16, 2958–2970. osmoregulation, are pervasive among metazoans. Although it Brandli, A.W., 1999. Towards a molecular anatomy of the Xenopus pronephric is conceivable that a kidney precursor existed in the common kidney. Int. J. Dev. Biol. 43, 381–395. ancestor of both insects and vertebrates, embryological studies Burns, R.K., 1955. Urogenital system. In: Willier, P.A.W. a. V.H.B.H. (Ed.), – indicated otherwise. Vertebrate kidneys have a mesodermal Analysis of Development. Saunders, Philadelphia, pp. 462 491. Campbell, G., Tomlinson, A., 1998. The roles of the homeobox genes aristaless origin, while insect renal tubules are formed mostly as an and Distal-less in patterning the legs and wings of Drosophila. Development ectodermal derivative. Nevertheless, recent work raises again 125, 4483–4493. the subject of homology. Cells of mesodermal origin undergo a Carroll, T.J., Vize, P.D., 1999. Synergism between Pax-8 and lim-1 in embryonic mesenchymal to epithelial transition and then give rise to the kidney development. Dev. Biol. 214, 46–59. stellate cells (Denholm et al., 2003). Mesenchymal to epithelial Carroll, T., Wallingford, J., Seufert, D., Vize, P.D., 1999. Molecular regulation of pronephric development. Curr. Top. Dev. Biol. 44, 67–100. transition is also characteristic of mesoderm mesenchymal cells Chan, T.C., Takahashi, S., Asashima, M., 2000. A role for Xlim-1 in pronephros while forming the vertebrate kidney. In addition, several fly development in Xenopus laevis. Dev. Biol. 228, 256–269. renal tubules genes such as Kr, cut and hibris have vertebrate Chen, G., Courey, A.J., 2000. Groucho/TLE family proteins and transcriptional homologues (Glis2, Cux-1 and nephrin, respectively) either repression. Gene 249, 1–16. expressed or having a role in kidney development (Sharma et Coffman, C.R., Skoglund, P., Harris, W.A., Kintner, C.R., 1993. Expression of an extracellular deletion of Xotch diverts cell fate in Xenopus embryos. Cell al., 2004; Vanden Heuvel et al., 1996; Zhang et al., 2002). 73, 659–671. However, these genes are expressed too late to play a role in the de Celis Ibeas, J.M., Bray, S.J., 2003. Bowl is required downstream of Notch for specification of vertebrate renal organs. Therefore, Odd/Osr elaboration of distal limb patterning. Development 130, 5943–5952. genes are the first ones that seem to participate during early Denholm, B., Sudarsan, V., Pasalodos-Sanchez, S., Artero, R., Lawrence, P., stages of renal development in both vertebrates and inverte- Maddrell, S., Baylies, M., Skaer, H., 2003. Dual origin of the renal tubules in Drosophila: mesodermal cells integrate and polarize to establish secretory brates. This molecular conservation might underlie a deep function. Curr. Biol. 13, 1052–1057. evolutionary homology between different kidney types. Alter- Drummond, I.A., Majumdar, A., Hentschel, H., Elger, M., Solnica-Krezel, L., natively, Odd/Osr genes might be used in a conserved Schier, A.F., Neuhauss, S.C., Stemple, D.L., Zwartkruis, F., Rangini, Z., 530 J.J. Tena et al. / Developmental Biology 301 (2007) 518–531

Driever, W., Fishman, M.C., 1998. Early development of the zebrafish skipped related 2 (Osr2) encodes a key intrinsic regulator of secondary pronephros and analysis of mutations affecting pronephric function. palate growth and morphogenesis. Development 131, 3207–3216. Development 125, 4655–4667. Lengyel, J.A., Iwaki, D.D., 2002. It takes guts: the Drosophila hindgut as a Goldstein, R.E., Cook, O., Dinur, T., Pisante, A., Karandikar, U.C., Bidwai, A., model system for organogenesis. Dev. Biol. 243, 1–19. Paroush, Z., 2005. An eh1-like motif in odd-skipped mediates recruitment of Majumdar, A., Lun, K., Brand, M., Drummond, I.A., 2000. Zebrafish no isthmus Groucho and repression in vivo. Mol. Cell. Biol. 25, 10711–10720. reveals a role for pax2.1 in tubule differentiation and patterning events in the Gómez-Skarmeta, J.L., de la Calle-Mustienes, E., Modolell, J., 2001. The Wnt- pronephric primordia. Development 127, 2089–2098. activated Xiro1 gene encodes a repressor that is essential for neural Pfeffer,P.L.,Gerster,T.,Lun,K.,Brand,M.,Busslinger,M.,1998. development and downregulates BMP4. Development 128, 551–560. Characterization of three novel members of the zebrafish Pax2/5/8 family: Green, R.B., Hatini, V., Johansen, K.A., Liu, X.J., Lengyel, J.A., 2002. dependency of Pax5 and Pax8 expression on the Pax2.1 (noi) function. Drumstick is a zinc finger protein that antagonizes Lines to control Development 125, 3063–3074. patterning and morphogenesis of the Drosophila hindgut. Development 129, Porteous, S., Torban, E., Cho, N.P., Cunliffe, H., Chua, L., McNoe, L., Ward, T., 3645–3656. Souza, C., Gus, P., Giugliani, R., Sato, T., Yun, K., Favor, J., Sicotte, M., Hao, I., Green, R.B., Dunaevsky, O., Lengyel, J.A., Rauskolb, C., 2003. The Goodyer, P., Eccles, M., 2000. Primary renal hypoplasia in humans and mice odd-skipped family of zinc finger genes promotes Drosophila leg with PAX2 mutations: evidence of increased apoptosis in fetal kidneys of segmentation. Dev. Biol. 263, 282–295. Pax2(1Neu)+/− mutant mice. Hum. Mol. Genet. 9, 1–11. Harbecke, R., Janning, W., 1989. The segmentation gene Kruppel of Drosophila Ryffel, G.U., 2003. What can a frog tell us about human kidney development. melanogaster has homeotic properties. Genes Dev. 3, 114–122. Nephron, Exp. Nephrol. 94, e35–e43. Harland, R., 1991. In situ hybridization: an improved whole mount method for Sanchez-Herrero, E., 1991. Control of the expression of the bithorax complex Xenopus embryos. Methods Cell Biol. 36, 685–695. genes abdominal-A and abdominal-B by cis-regulatory regions in Droso- Harland, R., Weintraub, H., 1985. Translation of mRNA injected into Xenopus phila embryos. Development 111, 437–449. oocytes is specifically inhibited by antisense RNA. J. Cell Biol. 101, Saxén, L., 1987. Organogenesis of the Kidney. Cambridge Univ. Press, Cambridge. 1094–1099. Sharma,M.,Fopma,A.,Brantley,J.G.,VandenHeuvel,G.B.,2004. Hart, M.C., Wang, L., Coulter, D.E., 1996. Comparison of the structure and Coexpression of Cux-1 and Notch signaling pathway components during expression of odd-skipped and two related genes that encode a new family of kidney development. Dev. Dyn. 231, 828–838. zinc finger proteins in Drosophila. Genetics 144, 171–182. Shawlot, W., Behringer, R.R., 1995. Requirement for Lim1 in head-organizer Heller, N., Brandli, A.W., 1999. Xenopus Pax-2/5/8 orthologues: novel insights function. Nature 374, 425–430. into Pax gene evolution and identification of Pax-8 as the earliest marker for Singer, J.B., Harbecke, R., Kusch, T., Reuter, R., Lengyel, J.A., 1996. Droso- otic and pronephric cell lineages. Dev. Genet. 24, 208–219. phila brachyenteron regulates gene activity and morphogenesis in the gut. Hernandez, P.P., Moreno, V., Olivari, F.A., Allende, M.L., 2005. Sub-lethal Development 122, 3707–3718. concentrations of waterborne copper are toxic to lateral line neuromasts in So, P.L., Danielian, P.S., 1999. Cloning and expression analysis of a mouse gene zebrafish (Danio rerio). Hear. Res. 213, 1–10. related to Drosophila odd-skipped. Mech. Dev. 84, 157–160. Hostetter, C.L., Sullivan-Brown, J.L., Burdine, R.D., 2003. Zebrafish pronephros: Stark, K., Vainio, S., Vassileva, G., McMahon, A.P., 1994. Epithelial a model for understanding cystic kidney disease. Dev. Dyn. 228, 514–522. transformation of metanephric mesenchyme in the developing kidney Iwaki, D.D., Johansen, K.A., Singer, J.B., Lengyel, J.A., 2001. drumstick, bowl, regulated by Wnt-4. Nature 372, 679–683. and lines are required for patterning and cell rearrangement in the Droso- Stricker, S., Brieske, N., Haupt, J., Mundlos, S., 2006. Comparative expression phila embryonic hindgut. Dev. Biol. 240, 611–626. pattern of Odd-skipped related genes Osr1 and Osr2 in chick embryonic James, R.G., Schultheiss, T.M., 2005. Bmp signaling promotes intermediate development. Gene Expr. Patterns 6, 826–834. mesoderm gene expression in a dose-dependent, cell-autonomous and Technau, U., 2001. Brachyury, the blastopore and the evolution of the mesoderm. translation-dependent manner. Dev. Biol. 288, 113–125. BioEssays 23, 788–794. James, R.G., Kamei, C.N., Wang, Q., Jiang, R., Schultheiss, T.M., 2006. Odd- Topczewska, J.M., Topczewski, J., Shostak, A., Kume, T., Solnica-Krezel, L., skipped related 1 is required for development of the metanephric kidney and Hogan, B.L., 2001. The winged helix transcription factor Foxc1a is essential regulates formation and differentiation of kidney precursor cells. Develop- for somitogenesis in zebrafish. Genes Dev. 15, 2483–2493. ment 133, 2995–3004. Torres, M., Gomez-Pardo, E., Dressler, G.R., Gruss, P., 1995. Pax-2 controls Jowett, T., Lettice, L., 1994. Whole-mount in situ hybridizations on zebrafish multiple steps of urogenital development. Development 121, 4057–4065. embryos using a mixture of digoxigenin- and fluorescein-labelled probes. Toyama, R., Dawid, I.B., 1997. lim6, a novel LIM homeobox gene in the Trends Genet. 10, 73–74. zebrafish: comparison of its expression pattern with lim1. Dev. Dyn. 209, Jung, A.C., Denholm, B., Skaer, H., Affolter, M., 2005. Renal tubule 406–417. development in Drosophila: a closer look at the cellular level. J. Am. Soc. Turner, D.L., Weintraub, H., 1994. Expression of achaete–scute homolog 3 in Nephrol. 16, 322–328. Xenopus embryos converts ectodermal cells to a neural fate. Genes Dev. Kaltschmidt, J.A., Davidson, C.M., Brown, N.H., Brand, A.H., 2000. Rotation 12, 1434–1447. and asymmetry of the mitotic spindle direct asymmetric cell division in the Vanden Heuvel, G.B., Bodmer, R., McConnell, K.R., Nagami, G.T., Igarashi, P., developing central nervous system. Nat. Cell Biol. 2, 7–12. 1996. Expression of a cut-related homeobox gene in developing and Kawai, S., Kato, T., Inaba, H., Okahashi, N., Amano, A., 2005. Odd-skipped polycystic mouse kidney. Kidney Int. 50, 453–461. related 2 splicing variants show opposite transcriptional activity. Biochem. Vize, P.D., Jones, E.A., Pfister, R., 1995. Development of the Xenopus Biophys Res. Commun. 328, 306–311. pronephric system. Dev. Biol. 171, 531–540. Kestila, M., Lenkkeri, U., Mannikko, M., Lamerdin, J., McCready, P., Putaala, Vize, P.D., Seufert, D.W., Carroll, T.J., Wallingford, J.B., 1997. Model systems H., Ruotsalainen, V., Morita, T., Nissinen, M., Herva, R., Kashtan, C.E., for the study of kidney development: use of the pronephros in the analysis of Peltonen, L., Holmberg, C., Olsen, A., Tryggvason, K., 1998. Positionally organ induction and patterning. Dev. Biol. 188, 189–204. cloned gene for a novel glomerular protein-nephrin-is mutated in congenital Wan, S., Cato, A.M., Skaer, H., 2000. Multiple signalling pathways establish nephrotic syndrome. Mol. Cell 1, 575–582. cell fate and cell number in Drosophila malpighian tubules. Dev. Biol. 217, Kosman, D., Small, S., Reinitz, J., 1998. Rapid preparation of a panel of 153–165. polyclonal antibodies to Drosophila segmentation proteins. Dev. Genes Wang, Q., Lan, Y., Cho, E.S., Maltby, K.M., Jiang, R., 2005. Odd-skipped related Evol. 208, 290–294. 1 (Odd1) is an essential regulator of heart and urogenital development. Dev. Lan, Y., Kingsley, P.D., Cho, E.S., Jiang, R., 2001. Osr2, a new mouse gene related Biol. 288, 582–594. to Drosophila odd-skipped, exhibits dynamic expression patterns during Ward, E.J., Coulter, D.E., 2000. odd-skipped is expressed in multiple tissues craniofacial, limb, and kidney development. Mech. Dev. 107, 175–179. during Drosophila embryogenesis. Mech. Dev. 96, 233–236. Lan, Y., Ovitt, C.E., Cho, E.S., Maltby, K.M., Wang, Q., Jiang, R., 2004. Odd- Wilm, B., James, R.G., Schultheiss, T.M., Hogan, B.L., 2004. The forkhead J.J. Tena et al. / Developmental Biology 301 (2007) 518–531 531

genes, Foxc1 and Foxc2, regulate paraxial versus intermediate mesoderm functions. Roles in kidney development and neurogenesis. J. Biol. Chem. cell fate. Dev. Biol. 271, 176–189. 277, 10139–10149. Zhang, F., Nakanishi, G., Kurebayashi, S., Yoshino, K., Perantoni, A., Kim, Y.S., Zhou, X., Vize, P.D., 2004. Proximo-distal specialization of epithelial transport Jetten, A.M., 2002. Characterization of Glis2, a novel gene encoding a Gli- processes within the Xenopus pronephric kidney tubules. Dev. Biol. 271, related, Kruppel-like transcription factor with transactivation and repressor 322–338. Supplementary data

Supplementary Fig. 1. Alignment of vertebrate Osr proteins. (A) Identical and similar amino acid residues are shaded in black and gray, respectively. Dashes indicate gaps in the alignment. The three canonical C2H2 zinc finger (C-X2-C-X12-H-X3-H) are underlined in blue. Asterisks mark a candidate eh1 motif. The accession numbers are: mOsr1 (AAD37115), XOsr1 (BN000922), zOsr1 (AAH83241), mOsr2B (AAL07364), XOsr2 (AAI08580), zOsr2

59 (AAH93148). (B) Phylogenetic tree showing the relative evolutionary distance between the different Osr proteins.

Supplementary Fig. 2. Specificity of Osr morpholinos. To test the specificity of the Xenopus and zebrafish Osr1 and Osr2 morpholinos (MOs), we generated Xenopus laevis or zebrafish Osr constructs bearing an Myc epitope either at the amino-(MT-XOsr1, MT-zOsr1, MT-XOsr2 and MT-zOsr2) or at the carboxy-terminus (XOsr1-MT, zOsr1-MT, XOsr2-MT and zOsr2-MT). MOs are antisense oligonucleotides that block translation efficiently only if their target sequence is within 25 bases from the translation start site. This is the case for Osr-MT mRNA but not for the MT-Osr transcripts. As controls, we examined if, for each species, the MO against one of the paralogues affects the translation of the other. Assays were performed in Xenopus laevis embryos. (A, D G and J) Myc staining of stage 12 embryos injected with 0.5 ng of MT-Osr mRNAs and 10 ng of the corresponding MO. The MOs are unable to block MT-Osr translation. (B, E, H and K) Myc staining in stage 12 embryos injected with 0.5 ng of Osr-MT mRNAs and 10 ng of the corresponding MO. Each MO effectively blocks the translation of its corresponding Osr-MT mRNA. Note that Xenopus MOs, which have one mismatch with the Xenopus laevis targeted sequence, are perfectly efficient in blocking

60 translation. However, the MO against one Osr-MT mRNA, which differs in five or more bases with the target sequence, cannot impair the translation of the other gene (C, F, I and L).

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62 CHAPTER II

“The osr1 and osr2 transcription factors are essential to trigger limb development by responding to retinoic acid and controlling wnt2ba expression in the pronephric anlage”

In this chapter, we characterized the expression pattern of osr genes in the pectoral fin domain in zebrafish. We show that osr1 and osr2 are necessary for early pectoral fin development. Both osr genes are able to modulate the earliest marker of fin formation, tbx5. This regulation is indirect and mediated, at least by osr1, through wnt2ba. Osr genes are also targets of the RA signaling.

I contributed to this work performing and assisting in the experimental design of the included studies.

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64

The osr1 and osr2 genes are essential for pectoral fin development by responding to retinoic acid and controlling wnt2ba expression in the pronephric anlage

Ana Neto, and José Luis Gómez-Skarmeta*

Centro Andaluz de Biología del Desarrollo, Consejo Superior de Investigaciones

Científicas/Universidad Pablo de Olavide, Carretera de Utrera Km1, 41013 Sevilla,

Spain.

Running title: pronephric osr expression is essential for limb development

* Author for correspondence

E-mails: [email protected]

1 Neto and Gómez-Skarmeta.

Abstract

Vertebrate Odd-skipped related genes (Osr) have an essential function during the formation of the intermediate mesoderm and the kidney structures derived form it. Here we show that these genes are also essential for triggering limb bud formation. Reduction of zebrafish osr function with specific morpholinos causes impairment of fin development due to the early downregulation of tbx5a in the lateral plate mesoderm

(LPM) at the level of the forming pectoral fin bud. The effects on tbx5a are indirect and mediated through Wnt2ba. Osr morphant embryos show a reduction of wnt2ba and increasing Wnt signaling in these morphant embryos partially rescue tbx5a expression.

Finally, we demonstrate that osr genes are downstream targets of retinoic acid signaling

(RA). Therefore, osr genes act as a relay within the genetic cascade of fin bud initiation formation: by controlling the expression of the signaling molecule Wnt2ba in the intermediate mesoderm they play an essential function transmitting the RA signaling originated in the somites to the LPM.

2 Neto and Gómez-Skarmeta.

Introduction

The odd-skipped (Odd/Osr) family of genes comprises evolutionary conserved zinc- finger transcription factors that lie at the top of the genetic hierarchy required for renal development in vertebrates and likely in Drosophila as well (James et al., 2006; Tena et al., 2007; Wang et al., 2005). Mammalian genomes contain two paralogues, Osr1 and

Osr2 (Lan et al., 2001; So and Danielian, 1999). In the mouse, Osr1 expression starts early (E7.5) in the intermediate mesoderm (IM), from where renal structures derive

(Mugford et al., 2008; So and Danielian, 1999), and is maintained until kidney organogenesis occurs. Osr2, in contrast, is activated at stage E9.25 in the mesonephros, and later (stage E14.5) in the mesenchyme that surrounds the ducts of the mesonephros and metanephros (Lan et al., 2001). Osr1 knock-outs lack renal structures (James et al.,

2006; Wang et al., 2005), while Osr2 mutants have apparently normal kidney development (Lan et al., 2004). Xenopus and zebrafish genomes also contain two Osr genes, and, in contrast to the mouse genes, both of them seem to contribute to some extend to the formation of the kidney (Tena et al., 2007). Indeed, knock-down of both genes generate stronger kidney defects than single depletions indicating partial redundancy between both genes. A partial redundancy of these two genes is further observed in gain of function assays in Xenopus and zebrafish (Tena et al., 2007), and can also be observed in knock-in experiments in mice (Gao et al., 2009).

Beside the kidney, vertebrate Osr genes are expressed in many other tissues. Analysis of mutant lines has indicated that these genes are required for proper formation and/or patterning of the endoderm, the heart, the teeth, the palate, the bones and the synovial joints in the limbs (Gao et al., 2011; Kawai et al., 2007; Lan et al., 2004; Mudumana et al., 2008; Wang et al., 2005; Zhang et al., 2009). Expression studies in mice and chicken indicate that in the limb Osr genes are expressed from very early stages in a

3 Neto and Gómez-Skarmeta.

highly dynamic pattern (Lan et al., 2001; So and Danielian, 1999; Stricker et al., 2006).

These genes start to be expressed at E11.5 in mouse or HH22 stage in chick limb buds in largely overlapping domains in the bud mesenchyme, the tissue that will form, among other cell types, the bones. Slightly later, the expression of Osr1 and Osr2 becomes largely complementary, whereby Osr2 is expressed more proximally and Osr1 more distally. Finally, at later stages during limb development, both genes are again co- expressed in the developing joints. Despite this complex and dynamic expression patterns covering most of the developing limb, tissue-specific ablation using a LPM specific Cre line (Prx::Cre) has recently indicated that Osr1 and Osr2 genes are only required for joint development (Gao et al., 2011). To get insight about other possible function of these genes during limb formation, and the degree of conservation of Osr function during the development of the vertebrate appendages, we have examined the requirement of both osr1 and osr2 genes during development of the zebrafish pectoral fins, the structures that are equivalent to forelimbs in tetrapods. Surprisingly, we find an essential function of these genes at the earliest stages of fin bud formation.

During early stages of development, the pectoral fin/forelimb field is induced with the specification of a group of LPM cells on either side of the embryo’s trunk at precise positions along the Anterior/Posterior (A/P) axis (Capdevila and Izpisua Belmonte,

2001; Johnson and Tabin, 1997; Mercader, 2007; Tickle, 1999). Interactions between the mesenchyme and the overlying ectoderm triggers the outgrowth of the fin/limb bud.

The earliest molecular marker described as an initiator of limb bud formation, is the T- box transcription factor Tbx5a that is expressed in the mesenchymal precursors of all tetrapods as well as all fish species analyzed so far (Agarwal et al., 2003; Begemann and Ingham, 2000; Gibson-Brown et al., 1996; Isaac et al., 1998; Logan et al., 1998;

Ruvinsky et al., 2000; Saito et al., 2006; Tamura et al., 1999). Indeed, Tbx5a is both

4 Neto and Gómez-Skarmeta.

necessary and sufficient for forelimb formation (Agarwal et al., 2003; Garrity et al.,

2002; Minguillon et al., 2005; Ng et al., 2002; Rallis et al., 2003; Takeuchi et al., 2003).

In zebrafish, tbx5a mutant heartstrings (hst) and tbx5a morphants show loss of pectoral fin formation (Garrity et al., 2002; Ng et al., 2002). The cells of the LPM expressing tbx5a fail to aggregate to form the compact circular structure of the wild type fin bud.

An upstream regulator of Tbx5a in the establishment of the limb field is the retinoic acid (RA) pathway (Begemann et al., 2001; Gibert et al., 2006; Grandel and Brand,

2010; Grandel et al., 2002; Mercader et al., 2006; Mic et al., 2004; Zhao et al., 2009).

The limiting step in the synthesis of RA is catalyzed by Aldehyde dehydrogenase 1a2, and the action of RA is limited mostly by its degradation through Cyp26a1. The medaka, zebrafish and mice mutants for aldh1a2 are not capable of RA synthesis during early development, resulting in the absence of fins/limbs, and also do not express Tbx5a

(Begemann et al., 2001; Grandel et al., 2002; Mic et al., 2004; Niederreither et al.,

1999). These phenotypes can be rescued by the exogenous application of RA or by transplantation of wild-type cells (Gibert et al., 2006; Linville et al., 2004; Mercader et al., 2006; Mic et al., 2004; Negishi et al., 2010; Niederreither et al., 1999; Zhao et al.,

2009).

The Wnt signaling pathway is also required for normal fin/limb outgrowth. In the chick, Wnt2b is detected in the IM as well as in the lateral plate mesoderm of the wing bud field (Kawakami et al., 2001). In the zebrafish, two wnt2b orthologs have been described, wnt2ba and wnt2bb. wnt2ba expression is restricted to the IM, while wnt2bb expression has been described in the LPM at the time of fin formation (Ng et al., 2002;

Ober et al., 2006). Wnt2b gain-of-function in the chick leads to ectopic limb formation

(Kawakami et al., 2001), while morpholino mediated wnt2ba gene silencing in the zebrafish impedes proper fin bud outgrowth (Ng et al., 2002). wnt2ba expression is

5 Neto and Gómez-Skarmeta.

regulated by RA signaling (Mercader et al., 2006; Negishi et al., 2010), suggesting that

RA might control limb initiation by controlling wnt2b expression.

The role of the IM, a kidney precursor tissue, in controlling limb outgrowth is still controversial. Classical experiments involving surgical ablation of the mesonephros in the chick led to impaired limb outgrowth (Geduspan and Solursh, 1992; Stephens and

McNulty, 1981). On the contrary, physical block of IM and LMP did not interfere with limb development (Fernandez-Teran et al., 1997). Also, genetic ablation of the kidney anlage in mouse appears to be compatible with the development of a normal limb

(Bouchard et al., 2002).

Here we report that in zebrafish osr1 and, to a lesser extend osr2, are required at early stages during limb development. We show that these genes are necessary within the IM for proper activation of wnt2ba. We also show that osr1 expression is controlled by RA signaling. Therefore, our studies allow connecting the RA and the Wnt pathways during early limb formation through Osr function.

Material and Methods zebrafish in situ hybridization

Antisense RNA probes were prepared from cDNAs using digoxigenin or fluorescein

(Boehringer Mannheim) as labels. Zebrafish specimens were prepared, hybridized and stained as described (Harland, 1991; Jowett and Lettice, 1994). For zwnt2ba in situ, the blocking solution used was 2% blocking powder (Roche) in Maleic Acid solution

(0.1M, pH7.5) and the antibody was diluted in the same blocking solution. For in situ hybridization of sections, embryos were sectioned in a Leica VT100S vibratome to a thickness of 30 µm and examined under the microscope.

6 Neto and Gómez-Skarmeta.

In vitro RNA synthesis and microinjection of mRNA and morpholinos

All DNAs were linearized and transcribed as described (Harland and Weintraub, 1985) with a GTP cap analog (New England Biolabs), using SP6, T3 or T7 RNA polymerases.

After DNAse treatment, RNA was extracted with phenol-chloroform, column purified and precipitated with ethanol. mRNAs for overexpression studies were resuspended in water and injected at the desired concentration in the yolk at 1–2 cell stage. For knock- down experiments, zebrafish embryos were injected in the yolk at 1–2 cell stage with

10–20 ng of morpholinos. The MOosr1 and MOosr2 morpholinos and the osr overexpression constructs have been described previously (Tena et al., 2007).

Pharmacological treatments

Embryos were incubated in the dark at 28 °C in 10–8 M all-trans retinoic acid (Sigma), diluted in embryo E3 medium, from a 10–5 M stock solution in DMSO. DEAB (4- diethylaminobenzaldehyde) (Sigma Aldrich), a competitive reversible inhibitor of retinaldehyde dehydrogenases (Begemann et al., 2004), was applied at a concentration of 10–4 M diluted from a 0.1 M stock in DMSO in E3 media. As controls, wild-type embryos were treated with similar dilutions of DMSO without drugs. Embryos injected with inducible domain of beta-catenin, were incubated with dexamethasone (4 µg/ml) in

E3 embryo medium (Kolm and Sive, 1995).

Results osr genes are required for fin development in zebrafish

We have previously reported two zebrafish morpholinos (MOosr1and MOosr2) that impair osr1 and osr2 gene function in zebrafish (Tena et al., 2007). Embryos injected with 20 ng of any of the individual MOs show reduction of fin buds at 5 dpf (MOosr1:

7 Neto and Gómez-Skarmeta.

60.5%, n=89; MOosr2: 46.5%; n=33), although the limb phenotype was stronger for osr1 morphant embryos (Fig. 1B and C). Moreover, this phenotype was stronger and more penetrant in embryos co-injected with 10 ng of each MO (71%; n=201, Fig. 1D).

These results indicate that both genes participate at some step along the genetic cascade that operates during fin development. We then examined the expression patterns of these genes during stages of early fin bud formation and compared them with that of tbx5a, an essential early gene for limb formation (Ahn et al., 2002; Garrity et al., 2002;

Rallis et al., 2003). At the level of the fin primordium, at 24 hours post fertilization

(hpf), both osr1 and osr2 genes were found at the developing pronephric anlage, and osr1 also show expression in endodermal cells (Mudumana et al., 2008; Tena et al.,

2007); Fig.2A, B). These domains are adjacent to the tbx5a-expressing territory that will form the limbs (Fig. 2C). At 48 hpf, while osr1 is very weakly expressed in the growing limb bud (Fig. 2D), osr2 is observed in two patches, one in the anterior and another in the posterior edges of this territory (Fig. 2E). A stronger and broader expression of osr2 was observed in the fins at 72 hpf, while osr1 is only moderately expressed in the fin at this stage (Fig. 2F-H).

osr genes are necessary at the pronephric anlage for early events during limb formation

The rather late expression of osr genes in the fin fields let us examined at what stage of pectoral fin development was affected in the osr morphant embryos. Surprisingly, at 24 hpf the expression of tbx5a was already affected in embryos injected with MOosr1

(60%; n=90) or MOosr2 (56%; n=88) (Fig. 3B and C). Correlating with the pectoral fin phenotype, tbx5a downregulation was stronger in osr1 morphant embryos than in embryos injected with MOosr2. Also as before, this effect that was even stronger in

8 Neto and Gómez-Skarmeta.

double morphant embryos (67%, n=162; Fig. 3D). In these experiments we used as an internal control tbx5a expression in the eye (Fig. 3E-H), which was not affected in injected embryos. This indicates that osr genes are required for limb formation before they are expressed in the fins. This was further demonstrated by examining tbx5a expression at the onset of pectoral fin induction. Indeed, in osr morphant embryos, tbx5a expression was already reduced at the 21 somites stage (MOosr1: 57%, n=113;

MOosr2: 47%; n=203, MOosr1+ MOosr21: 68%, n=147, Fig. 3I-L). The fact that the combination of both osr MOs produced stronger defects that individual MOs suggests that both Osr factors are required, in a partially redundant manner, for pectoral fin development. To assess this, we evaluate the ability of individual osr genes to rescue tbx5a expression in double osr-morphant embryos. As expected, both osr genes were similarly capable of rescuing the expression of tbx5a in the pectoral fin territory, as well as the expression of the pronephric marker pax2, which was used as a control in this experiment (Fig. 4A-D). Thus, while 93,5% of the embryos morphant for both osr genes showed reduced tbx5a and pax2 expression, this proportion was reduced to 83%

(n=138) or 76% (n=250) in embryos co-injected with both MOs and MTosr1 or MTosr2 mRNAs, respectively.

We conclude that both osr genes are required, in a partially redundant manner, for pectoral fin formation at the time they are not expressed in this territory but are co- expressed in the kidney anlage (Tena et al., 2007).

osr genes are necessary for wnt2ba expression at the intermediate mesoderm

Both in zebrafish and chick embryos, it has been reported that wnt2ba, which is expressed in the IM, is essential to trigger limb formation by activating tbx5a expression (Kawakami et al., 2001; Ng et al., 2002). Since this is precisely the tissue

9 Neto and Gómez-Skarmeta.

expressing osr genes, we examined wnt2ba expression in embryos with impaired osr function. In morphant embryos for osr1 function wnt2ba expression was downregulated

(60%, n=47; Fig. 5A-C). Embryos injected with MOzosr2 did not show reduced wnt2ba, although in double morphant embryos wnt2ba downregulation was stronger than in those injected with MOzosr1 alone (59%, n=138; Fig. 5D). To confirm that downregulation of tbx5a observed in the osr morphant embryos is dependent on Wnt signaling, we overexpressed this pathway in embryos co-injected with both osr MOs. In this experiment, we used an inducible form of beta-catenin fused to the

Dexamethasone-regulated Glucocorticoid Receptor domain (Afouda et al., 2008). This allowed us, upon adding Dexamethasone (Dex) to the injected embryos at tailbud stage, to activate the pathway only after gastrulation preventing early defects associated with increased Wnt signaling during this critical period of development. The expression of tbx5a was not affected in embryos injected with beta-catenin mRNA in the absence of

Dex but was slightly expanded upon Dex addition (38%, n=65; Fig. 5E, F). In embryos co-injected with both osr MOs and beta-catenin mRNA, tbx5a was downregulated in the absence of Dex (56%; n=66). However, tbx5a downregulation was observed in significantly fewer embryos upon adding this hormone (30%, n=71; Fig. 5G,H). These results strongly suggest that, at least in part, the Wnt signaling mediates the effect of osr genes on tbx5a and limb development. Accordingly, sections through 22 hpf embryos show that osr genes and wnt2ba are both expressed in the pronephric area (Fig. 6).

Retinoic acid triggers limb formation through osr genes

Retinoic acid (RA) is essential for proper kidney development and also for limb formation through the regulation of tbx5a (Begemann et al., 2001; Cartry et al., 2006;

Gibert et al., 2006; Grandel and Brand, 2010; Mic et al., 2004). Since osr genes are key

10 Neto and Gómez-Skarmeta.

regulators of both processes, we examined the mutual relationship between these genes and the RA pathway. We first determined if the expression of raldh2a, an enzyme required for RA production was affected in osr morphant embryos. As expected for downstream factors of the RA pathway, the expression of raldh2a was not affected in embryos injected with any of the osr MOs or the combination of both of them (MOosr1:

3%, n=53; MOosr2: 2%; n=41, MOosr1+ MOosr2: 1%, n=40, Fig. 7 A-D). This suggests that osr genes may be downstream of the RA signaling cascade. Further indicating this, while increasing RA signaling promotes a slight enlargement of the tbx5a expression territory (48%, n=128; Fig. 8E, F), this treatment was unable to rescue tbx5a expression in osr morphant embryos (95% of the embryos with reduced tbx5a expression, n=47; Fig. 8G, H). We then examined the effect of increasing or reducing the RA signaling pathway on the expression of the osr genes. To that end, we incubated the embryos from tailbud to the 21-22 somites stages either with all-trans RA or with

DEAB (4-diethylaminobenzaldehyde), a competitive reversible inhibitor of retinaldehyde dehydrogenases. In embryos incubated with RA the osr1 expression domain became broader while the osr2 expression pattern was slightly extended anteriorly (45%, n=31; Fig.7J, M). In contrast, in embryos incubated with DEAB osr1 was strongly downregulated (96%, n=25; Fig.7K) and the osr2 expression domain was slightly compacted (100%, n=34; Fig. 7N). These results strongly indicate that ors1, and to a lesser extent osr2, are downstream of the RA signaling pathway. If so, it would be expected that overexpression of osr genes could be able to overcome the loss of tbx5a expression under conditions of reduced RA signaling. Indeed, this was the case since tbx5a expression was recovered in 30% of the embryos exposed to DEAB when injected with the osr mRNAs (Fig.7 O-R).

11 Neto and Gómez-Skarmeta.

Discussion

Novel function of osr genes during pectoral fin induction

Our work demonstrates an essential function of zebrafish Osr1 and Osr2 transcription factors during pectoral fin formation. osr genes are required at initial stages of fin outgrowth, for tbx5 expression, which is the earliest marker involved in limb bud formation (Agarwal et al., 2003; Ahn et al., 2002; Garrity et al., 2002; Rallis et al.,

2003). Interestingly, this requirement occurs at a developmental stage in which the osr genes are not expressed in the fin bud primordia, but in the adjacent IM. A role of the

IM during limb development has been previously been reported by others (Geduspan and Solursh, 1992; Stephens and McNulty, 1981). Our data further support an important function of kidney anlage for limb bud formation.

As it has been previously found during early kidney formation in zebrafish and Xenopus

(Tena et al., 2007), both osr genes seem to be partially redundantly required for to pectoral fin formation. A similar redundancy has also been shown during mouse joint formation (Gao et al., 2011) and in other knock-in studies (Gao et al., 2009).

Nevertheless, during pectoral fin formation, we systematically observed that loss of osr1 function seems to produce stronger defects that the reduction of osr2 activity. This may be due to broader and earlier expression of osr1 in the IM territory in most vertebrates (Lan et al., 2001; So and Danielian, 1999; Stricker et al., 2006; Tena et al.,

2007). Therefore, as proposed recently (Gao et al., 2009), the distinct developmental requirement exhibited by both genes in this and other processes is likely the result of divergence of the cis-regulatory regions that control the spatiotemporal expression of these genes, although the functional potential of both gene products is likely very similar.

Our results demonstrate that osr genes are required in the IM territory to indirectly

12 Neto and Gómez-Skarmeta.

promote the induction of the pectoral fin bud. This indirect action could be explained by the diffusible molecule wnt2ba, a factor essential for the activation of tbx5a (Ng et al.,

2002). We show that wnt2ba and osr genes show overlapping expression domains in the

IM, and that wnt2ba expression depends on the function of Osr transcription factors.

Although the functional reduction of both osr genes causes a stronger wnt2ba downregulation than the single osr1 knock-down, the loss of osr2 does not affect wnt2ba expression. This suggests a minor requirement of osr2 function for wnt2ba expression. Nevertheless, loss of osr2 does cause tbx5a downregulation and pectoral fin defects. Therefore, it is likely that osr genes are also necessary for the expression of signaling molecules at the IM, other than Wnt2ba, that also participate in pectoral fin formation.

Positioning osr genes in the signaling cascade involved in fin induction

The earliest known player of the signaling cascade that leads to pectoral fin induction is the RA signaling pathway. RA signaling during gastrulation has been shown to be important for the establishment of a tbx5a-positive fin field (Grandel and Brand, 2010;

Grandel et al., 2002). During somitogenesis, RA derived from the somites is involved in maintaining and expanding the tbx5-positive limb precursors (Begemann et al., 2001;

Gibert et al., 2006; Linville et al., 2004; Mercader et al., 2006). RA is also required for kidney formation (Cartry et al., 2006) and has been shown to be able to activate the expression of Osr genes when Xenopus animal caps or mouse ES cells differentiated to

IM or kidney identities (Drews et al., 2011; Mae et al., 2010). Here we show that in zebrafish embryos RA signaling is strongly required for osr1 expression, and to a lesser extend for osr2 activation. We therefore propose a model in which RA signaling, produced at the anterior somites, is required to activate the expression of osr genes in

13 Neto and Gómez-Skarmeta.

the IM. These transcription factors are then essential at the IM for renal organ formation but also for indirectly promoting pectoral fin development by controlling wnt2ba expression at the kidney anlage. Wnt2ba, produced in the osr-expressing domain, then diffuses to the LPM to maintain tbx5a expression and promote pectoral fin formation

(Fig. 8 Scheme). In this genetic cascade, in which the RA signaling is relayed through three different tissues, the osr genes in the IM play an essential linker function responding to a signal from the somites and transmitting it to the LPM. Moreover, by activating the osr genes, RA is capable to control in a coordinate way two different developmental processes, kidney formation and pectoral fin development.

How much of this genetic cascade is conserved during tetrapode limb development

Neither Osr1 nor Osr2 knock-out mouse embryos have been reported to present major limb defects (James et al., 2006; Lan et al., 2004; Wang et al., 2005). This could be due to a partial redundancy of both genes in the activation of Wnt2b, or another similar Wnt ligand, at the kidney anlage. A very recent report has examined the consequences of removing both genes at the early limb bud (Gao et al., 2011). In this study the impairment of both genes produced only late developmental defects in joint formation but it did not impair early limb bud development (Gao et al., 2011). However, in these experiments Osr1 was still present in the IM. To precisely examine Osr requirement in the activation of limb bud formation, it would be necessary to eliminate both Osr genes from the early kidney anlage. Nevertheless, there are several reasons to believe that Osr genes are likely necessary for limb bud formation in other vertebrates as well. Firstly, in all vertebrates examined, proper kidney and limb formation require RA signaling

(Begemann et al., 2001; Cartry et al., 2006; Gibert et al., 2006; Grandel and Brand,

2010; Grandel et al., 2002; Linville et al., 2004; Mercader et al., 2006; Mic et al., 2004;

14 Neto and Gómez-Skarmeta.

Negishi et al., 2010; Niederreither et al., 1999; Zhao et al., 2009). Secondly, as we report here in zebrafish, this signaling pathway also controls Osr expression in other vertebrates (Drews et al., 2011; Mae et al., 2010). Thirdly, it has also been reported that the kidney anlage is required for limb formation in other vertebrates (Geduspan and

Solursh, 1992; Stephens and McNulty, 1981), and kidney formation depends on Osr function in all vertebrates (James et al., 2006; Mudumana et al., 2008; Tena et al., 2007;

Wang et al., 2005). Finally, Osr expression at the IM precedes and overlaps that of

Wnt2b in all vertebrates examined (Kawakami et al., 2001; Mercader et al., 2006;

Mudumana et al., 2008; Ng et al., 2002; Stricker et al., 2006; Tena et al., 2007).

Therefore, in all vertebrates, Osr expression is in the right place (the kidney anlage) and at the right time to relay RA signaling from the somites to the lateral plate through

Wnt2b. Functional experiments would be required to demonstrate if Osr genes indeed participate in the genetic cascade that triggers limb bud formation in other vertebrates as well.

Acknowledgements

We would like to thank S. Hoppler, J. C. Izpisua-Belmonte, K. Takeshima and M.

Allende for reagents. We are especially grateful to F. Casares and N. Mercader for critical reading of the manuscript. We thank the Spanish and Andalusian Governments for grants BFU2010-14839, CSD2007-00008 and Proyecto de Excelencia CVI-3488.

Ana Neto is funded by Fundaçao para a Ciencia e Tecnologia from Portugal

(SFRH/BD/15242/2004) and belongs to the Graduate Program in Areas of Basic and

Applied Biology from Oporto University.

15 Neto and Gómez-Skarmeta.

References

Afouda, B. A., Martin, J., Liu, F., Ciau-Uitz, A., Patient, R. and Hoppler, S. (2008).

GATA transcription factors integrate Wnt signalling during heart development.

Development 135, 3185-3190.

Agarwal, P., Wylie, J. N., Galceran, J., Arkhitko, O., Li, C., Deng, C., Grosschedl,

R. and Bruneau, B. G. (2003). Tbx5 is essential for forelimb bud initiation following patterning of the limb field in the mouse embryo. Development 130, 623-633.

Ahn, D. G., Kourakis, M. J., Rohde, L. A., Silver, L. M. and Ho, R. K. (2002). T- box gene tbx5 is essential for formation of the pectoral limb bud. Nature 417, 754-758.

Begemann, G. and Ingham, P. W. (2000). Developmental regulation of Tbx5 in zebrafish embryogenesis. Mech Dev 90, 299-304.

Begemann, G., Marx, M., Mebus, K., Meyer, A. and Bastmeyer, M. (2004). Beyond the neckless phenotype: influence of reduced retinoic acid signaling on motor neuron development in the zebrafish hindbrain. Dev Biol 271, 119-129.

Begemann, G., Schilling, T. F., Rauch, G. J., Geisler, R. and Ingham, P. W. (2001).

The zebrafish neckless mutation reveals a requirement for raldh2 in mesodermal signals that pattern the hindbrain. Development 128, 3081-3094.

Bouchard, M., Souabni, A., Mandler, M., Neubuser, A. and Busslinger, M. (2002).

Nephric lineage specification by Pax2 and Pax8. Genes Dev 16, 2958-2970.

Capdevila, J. and Izpisua Belmonte, J. C. (2001). Patterning mechanisms controlling vertebrate limb development. Annu Rev Cell Dev Biol 17, 87-132.

Cartry, J., Nichane, M., Ribes, V., Colas, A., Riou, J. F., Pieler, T., Dolle, P.,

Bellefroid, E. J. and Umbhauer, M. (2006). Retinoic acid signalling is required for specification of pronephric cell fate. Dev Biol 299, 35-51.

16 Neto and Gómez-Skarmeta.

Drews, C., Senkel, S. and Ryffel, G. U. (2011). The nephrogenic potential of the transcription factors osr1, osr2, hnf1b, lhx1 and pax8 assessed in Xenopus animal caps.

BMC Dev Biol 11, 5.

Fernandez-Teran, M., Piedra, M. E., Simandl, B. K., Fallon, J. F. and Ros, M. A.

(1997). Limb initiation and development is normal in the absence of the mesonephros.

Dev Biol 189, 246-255.

Gao, Y., Lan, Y., Liu, H. and Jiang, R. (2011). The zinc finger transcription factors

Osr1 and Osr2 control synovial joint formation. Dev Biol. 352, 83-91

Gao, Y., Lan, Y., Ovitt, C. E. and Jiang, R. (2009). Functional equivalence of the zinc finger transcription factors Osr1 and Osr2 in mouse development. Dev Biol 328,

200-209.

Garrity, D. M., Childs, S. and Fishman, M. C. (2002). The heartstrings mutation in zebrafish causes heart/fin Tbx5 deficiency syndrome. Development 129, 4635-4645.

Geduspan, J. S. and Solursh, M. (1992). A growth-promoting influence from the mesonephros during limb outgrowth. Dev Biol 151, 242-250.

Gibert, Y., Gajewski, A., Meyer, A. and Begemann, G. (2006). Induction and prepatterning of the zebrafish pectoral fin bud requires axial retinoic acid signaling.

Development 133, 2649-2659.

Gibson-Brown, J. J., Agulnik, S. I., Chapman, D. L., Alexiou, M., Garvey, N.,

Silver, L. M. and Papaioannou, V. E. (1996). Evidence of a role for T-box genes in the evolution of limb morphogenesis and the specification of forelimb/hindlimb identity. Mech Dev 56, 93-101.

Grandel, H. and Brand, M. (2010). Zebrafish limb development is triggered by a retinoic acid signal during gastrulation. Dev Dyn. DOI: 10.1002/dvdy.22461

17 Neto and Gómez-Skarmeta.

Grandel, H., Lun, K., Rauch, G. J., Rhinn, M., Piotrowski, T., Houart, C., Sordino,

P., Kuchler, A. M., Schulte-Merker, S., Geisler, R. et al. (2002). Retinoic acid signalling in the zebrafish embryo is necessary during pre-segmentation stages to pattern the anterior-posterior axis of the CNS and to induce a pectoral fin bud.

Development 129, 2851-2865.

Harland, R. (1991). In situ hybridization:an improved whole mount method for

Xenopus embryos. Methods Cell Biol. 36, 685-695.

Harland, R. and Weintraub, H. (1985). Translation of mRNA injected into Xenopus oocytes is specifically inhibited by antisense RNA. J. Cell Biol. 101, 1094-1099.

Isaac, A., Rodriguez-Esteban, C., Ryan, A., Altabef, M., Tsukui, T., Patel, K.,

Tickle, C. and Izpisua-Belmonte, J. C. (1998). Tbx genes and limb identity in chick embryo development. Development 125, 1867-1875.

James, R. G., Kamei, C. N., Wang, Q., Jiang, R. and Schultheiss, T. M. (2006).

Odd-skipped related 1 is required for development of the metanephric kidney and regulates formation and differentiation of kidney precursor cells. Development 133,

2995-3004.

Johnson, R. L. and Tabin, C. J. (1997). Molecular models for vertebrate limb development. Cell 90, 979-990.

Jowett, T. and Lettice, L. (1994). Whole-mount in situ hybridizations on zebrafish embryos using a mixture of digoxigenin- and fluorescein-labelled probes. Trends Genet

10, 73-74.

Kawai, S., Yamauchi, M., Wakisaka, S., Ooshima, T. and Amano, A. (2007). Zinc- finger transcription factor odd-skipped related 2 is one of the regulators in osteoblast proliferation and bone formation. J Bone Miner Res 22, 1362-1372.

18 Neto and Gómez-Skarmeta.

Kawakami, Y., Capdevila, J., Buscher, D., Itoh, T., Rodriguez Esteban, C. and

Izpisua Belmonte, J. C. (2001). WNT signals control FGF-dependent limb initiation and AER induction in the chick embryo. Cell 104, 891-900.

Kolm, P. J. and Sive, H. L. (1995). Efficient hormone-inducible protein function in

Xenopus laevis. Dev. Biol. 171, 267-272.

Lan, Y., Kingsley, P. D., Cho, E. S. and Jiang, R. (2001). Osr2, a new mouse gene related to Drosophila odd-skipped, exhibits dynamic expression patterns during craniofacial, limb, and kidney development. Mech Dev 107, 175-179.

Lan, Y., Ovitt, C. E., Cho, E. S., Maltby, K. M., Wang, Q. and Jiang, R. (2004).

Odd-skipped related 2 (Osr2) encodes a key intrinsic regulator of secondary palate growth and morphogenesis. Development 131, 3207-3216.

Linville, A., Gumusaneli, E., Chandraratna, R. A. and Schilling, T. F. (2004).

Independent roles for retinoic acid in segmentation and neuronal differentiation in the zebrafish hindbrain. Dev Biol 270, 186-199.

Logan, M., Simon, H. G. and Tabin, C. (1998). Differential regulation of T-box and homeobox transcription factors suggests roles in controlling chick limb-type identity.

Development 125, 2825-2835.

Mae, S., Shirasawa, S., Yoshie, S., Sato, F., Kanoh, Y., Ichikawa, H., Yokoyama,

T., Yue, F., Tomotsune, D. and Sasaki, K. (2010). Combination of small molecules enhances differentiation of mouse embryonic stem cells into intermediate mesoderm through BMP7-positive cells. Biochem Biophys Res Commun 393, 877-882.

Mercader, N. (2007). Early steps of paired fin development in zebrafish compared with tetrapod limb development. Dev Growth Differ 49, 421-437.

19 Neto and Gómez-Skarmeta.

Mercader, N., Fischer, S. and Neumann, C. J. (2006). Prdm1 acts downstream of a sequential RA, Wnt and Fgf signaling cascade during zebrafish forelimb induction.

Development 133, 2805-2815.

Mic, F. A., Sirbu, I. O. and Duester, G. (2004). Retinoic acid synthesis controlled by

Raldh2 is required early for limb bud initiation and then later as a proximodistal signal during apical ectodermal ridge formation. J Biol Chem 279, 26698-26706.

Minguillon, C., Del Buono, J. and Logan, M. P. (2005). Tbx5 and Tbx4 are not sufficient to determine limb-specific morphologies but have common roles in initiating limb outgrowth. Dev Cell 8, 75-84.

Mudumana, S. P., Hentschel, D., Liu, Y., Vasilyev, A. and Drummond, I. A. (2008). odd skipped related1 reveals a novel role for endoderm in regulating kidney versus vascular cell fate. Development 135, 3355-3367.

Mugford, J. W., Sipila, P., McMahon, J. A. and McMahon, A. P. (2008). Osr1 expression demarcates a multi-potent population of intermediate mesoderm that undergoes progressive restriction to an Osr1-dependent nephron progenitor compartment within the mammalian kidney. Dev Biol 324, 88-98.

Negishi, T., Nagai, Y., Asaoka, Y., Ohno, M., Namae, M., Mitani, H., Sasaki, T.,

Shimizu, N., Terai, S., Sakaida, I. et al. (2010). Retinoic acid signaling positively regulates liver specification by inducing wnt2bb gene expression in medaka.

Hepatology 51, 1037-1045.

Ng, J. K., Kawakami, Y., Buscher, D., Raya, A., Itoh, T., Koth, C. M., Rodriguez

Esteban, C., Rodriguez-Leon, J., Garrity, D. M., Fishman, M. C. et al. (2002). The limb identity gene Tbx5 promotes limb initiation by interacting with Wnt2b and Fgf10.

Development 129, 5161-5170.

20 Neto and Gómez-Skarmeta.

Niederreither, K., Subbarayan, V., Dolle, P. and Chambon, P. (1999). Embryonic retinoic acid synthesis is essential for early mouse post-implantation development. Nat

Genet 21, 444-448.

Ober, E. A., Verkade, H., Field, H. A. and Stainier, D. Y. (2006). Mesodermal

Wnt2b signalling positively regulates liver specification. Nature 442, 688-691.

Rallis, C., Bruneau, B. G., Del Buono, J., Seidman, C. E., Seidman, J. G., Nissim,

S., Tabin, C. J. and Logan, M. P. (2003). Tbx5 is required for forelimb bud formation and continued outgrowth. Development 130, 2741-2751.

Ruvinsky, I., Oates, A. C., Silver, L. M. and Ho, R. K. (2000). The evolution of paired appendages in vertebrates: T-box genes in the zebrafish. Dev Genes Evol 210,

82-91.

Saito, D., Yonei-Tamura, S., Takahashi, Y. and Tamura, K. (2006). Level-specific role of paraxial mesoderm in regulation of Tbx5/Tbx4 expression and limb initiation.

Dev Biol 292, 79-89.

So, P. L. and Danielian, P. S. (1999). Cloning and expression analysis of a mouse gene related to Drosophila odd-skipped. Mech Dev 84, 157-160.

Stephens, T. D. and McNulty, T. R. (1981). Evidence for a metameric pattern in the development of the chick humerus. J Embryol Exp Morphol 61, 191-205.

Stricker, S., Brieske, N., Haupt, J. and Mundlos, S. (2006). Comparative expression pattern of Odd-skipped related genes Osr1 and Osr2 in chick embryonic development.

Gene Expr. Patterns. 6, 826-834.

Takeuchi, J. K., Ohgi, M., Koshiba-Takeuchi, K., Shiratori, H., Sakaki, I., Ogura,

K., Saijoh, Y. and Ogura, T. (2003). Tbx5 specifies the left/right ventricles and ventricular septum position during cardiogenesis. Development 130, 5953-5964.

21 Neto and Gómez-Skarmeta.

Tamura, K., Yonei-Tamura, S. and Izpisua Belmonte, J. C. (1999). Differential expression of Tbx4 and Tbx5 in Zebrafish fin buds. Mech Dev 87, 181-184.

Tena, J. J., Neto, A., de la Calle-Mustienes, E., Bras-Pereira, C., Casares, F. and

Gomez-Skarmeta, J. L. (2007). Odd-skipped genes encode repressors that control kidney development. Dev Biol 301, 518-531.

Tickle, C. (1999). Morphogen gradients in vertebrate limb development. Semin Cell

Dev Biol 10, 345-351.

Wang, Q., Lan, Y., Cho, E. S., Maltby, K. M. and Jiang, R. (2005). Odd-skipped related 1 (Odd1) is an essential regulator of heart and urogenital development. Dev Biol

288, 582-594.

Zhang, Z., Lan, Y., Chai, Y. and Jiang, R. (2009). Antagonistic actions of Msx1 and

Osr2 pattern mammalian teeth into a single row. Science 323, 1232-1234.

Zhao, X., Sirbu, I. O., Mic, F. A., Molotkova, N., Molotkov, A., Kumar, S. and

Duester, G. (2009). Retinoic acid promotes limb induction through effects on body axis extension but is unnecessary for limb patterning. Curr Biol 19, 1050-1057.

22 Neto and Gómez-Skarmeta.

Figure legends

Figure 1. Zebrafish osr genes are necessary for pectoral fin formation.

All panels show the dorsal view of 5 dpf zebrafish embryos with anterior to the top. (A)

Control uninjected embryos. (B, C) Embryos injected with 20 ng of MOosr1 (B) or

MOosr2 (C). (D, E) Embryos injected with 10 ng of each morpholino. Note that the reduction of the pectoral fin is stronger in double injected embryos (D, E) than in embryos morphant for each individual gene (B, C). Arrowheads point at reduced fins and the asterisk to the complete elimination of the fins in a double morphant embryo.

Figure 2. Comparative expression pattern of osr genes and tbx5a during pectoral fin development.

All panels show the dorsal view of zebrafish embryos with the anterior to the left. (A-C)

At 24hpf, osr1 (A) and osr2 are expressed at the kidney anlage (white arrowheads) adjacent to the tbx5a (C) domain in the forming limb bud. (D-F) At 48 hpf, osr1 expression is detected very weakly in the fin bud (D), while osr2 is found in two patched domains within the tbx5a (F) expressing territory. (G-I) At 72hpf, osr2 is expressed in broader domain (H) overlapping most of the tbx5a expressing territory (J), while osr1 expression occurs only in small patches in the developing limbs (G).

Figure 3. osr genes are required for early tbx5a expression at the fin bud formation.

Dorsal (A-D and I-L) and lateral (E-H) views of zebrafish embryos with the anterior to the left. (A-H) Effect of the injection of individual osr MOs (A-C and E-G) or the combination of both of them (D, H) on tbx5a expression in pectoral fin (A-D) or in the

23 Neto and Gómez-Skarmeta.

eye (E-H) at 24 hpf. The expression of tbx5a is reduced in the pectoral fins (encircle in red) but not in the eye of the morphant embryos. (I-L) Depletion of individual osr genes

(I-K), or both of them (L), already reduces tbx5a expression in the presumptive pectoral fin at the 21 somites stage (arrowheads).

Figure 4. Morpholinos insensitive osr mRNAs rescue tbx5a and pax2 expression in double morphants embryos.

All panels show the expression of tbx5a and pax2 from a dorso-lateral view at 24 hpf.

In each embryo the limb bud is encircle in red and the pronephos is pointed at with and arrowhead. (A) Control un-injected embryo. (B) Embryos co-injected with the two osr morpholinos. (C, D) Embryos co-injected with the two osr morpholinos and the MT- osr1 (C) or the MT-osr2 (D) mRNA. Note that both the pronephros and the limb buds are reduced in double morphant embryos, defects rescued by any of the two osr mRNAs. Note also that in (C, D) the limb buds seem slightly larger that in (A).

Figure 5. wnt2ba expression in the intermediate mesoderm is dowsntream of osr function.

All panels show dorsal views with anterior to the left. (A-D) wnt2ba expression at 22 hpf (arrowheads) in the IM is downregulated in single (B, C) and double (D) osr- morphant embryos. (E-H) Increasing Wnt signaling in double osr-morphant embryos rescue tbx5a expression in the limb buds at 30 hpf. (E, F) Embryos injected with

βcatGR mRNA show normal (E) or slightly larger (F) tbx5a expression domains

(encircle in red) in the absence or presence of Dexamethasone, respectively. (G, H)

Embryos co-injected with βcatGR mRNA and the two osr MOs show reduced (G) or

24 Neto and Gómez-Skarmeta.

rescued (H) tbx5a expression domains in the absence or presence of Dexamethasone, respectively.

Figure 6. osr, wnt2ba and pax2 genes are co-expressed in the kidney anlage

Transverse sections of 21-somites stage embryos with dorsal at the top. (A-C) Embryos showing the expression of osr1 (A), ors2 (B) or wnt2ba (C) genes in the kidney anlage.

(D-E). Double stained embryos for osr1 (D) or ors1 (E) and pax2 (osr genes in purple and pax2 in red) and wnt2ba (purple) and osr2 (red) (F). Note that the expression of all three genes overlaps at the pronephros (encircle).

Figure 7. Retinoic acid signaling is required for limb formation through the activation of the osr genes

All panels are dorsal view of embryos at the 21-somite stage (A-D, I-K and O-R) or at

24 hpf (E-H and L-N) with anterior to the left. (A-D) raldh2 expression is not affected in single or double osr-morphant embryos (arrowheads). (E-H) Increasing RA signaling is unable to rescue loss of tbx5a expression (encircle) in double osr-morphant embryos.

(I-L) RA signaling is required for the expression of the osr1 (I-K) and osr2 (L-N) genes.

Thus, increasing (J, M) or reducing (K, N) RA signaling, expanded or reduced, respectively, the expression domains of the osr genes (arrowheads). (O-R) osr genes can rescue the loss of tbx5a expression when RA signaling is reduced. (O, P) Reducing

RA signaling strongly impaired tbx5a expression (arrowheads). (Q-R) The expression of tbx5a is rescued when osr genes are overexpressed under these conditions

(arrowheads).

25 Neto and Gómez-Skarmeta.

Figure 8. Schematic representation of the relay mechanism that triggers pectoral fin development in zebrafish embryos.

Retinoic acid, generated at the somites (red), activates the osr genes (orange: osr1, yellow: osr2) at the pronephric territory. Osr transcription factors are then required for the expression of the signaling molecule wnt2ba expression (green) that induced tbx5a expression (blue) in the adjacent lateral plate mesoderm in which limb bud formation is initiated.

26 Neto and Gómez-Skarmeta.

GENERAL DISCUSSION

99

100 GENERAL DISCUSSION

Osr genes are essential genes for early kidney development

In this study we have demonstrated that osr genes are at the top of the genetic cascade required for kidney development in Xenopus and zebrafish.

We have first characterized the expression pattern of osr1 and osr2 in Xenopus and zebrafish and compare it with expression of early kidney markers. In Xenopus, the expression of both Xosr genes in the kidney anlage starts before all known early kidney markers such as Xpax8 and Xlim1 are activated. In zebrafish osr1 expression is detected before the activation of the first kidney markers, such as pax2.1 and lim1, while osr2 expression begins only at the 8 somites stage, when these early pronephric markers are already activated and before any histological signals of kidney tissue are detected (Drummond et al., 1998). Here, performing loss of function studies, we show that in zebrafish and Xenopus both genes are necessary for the kidney formation. These phenotypes are correlated with the spatiotemporal expression of osr genes in the early kidney domain. We also observed that the simultaneous knock down of osr1 and osr2 does not reveal any synergism indicating that they are required additively for early kidney formation.

A more recent independent study with zebrafish osr1 has largely confirmed our results (Mudumana et al., 2008), although Mudumana and colleagues have also shown that osr1 is not required for distal pronephros development. Moreover, instead of being essential for early glomerulus formation, it seems that osr1 plays more important role in the correct maintenance of the glomerulus tissue.

In contrast with zebrafish and Xenopus, in mice only Osr1 mutants present kidney defects (James et al., 2006; Wang et al., 2005). Indeed, in mice, the onset of Osr1 expression in the intermediate mesoderm (IM) precedes the detection of kidney markers (James et al., 2006; So and Danielian, 1999; Stricker et al., 2006; Wang et al., 2005). However, the onset of Osr2 expression in mice occurs even in a later step of kidney development than in zebrafish, at 18- somite stage, when the embryonic kidney is already differentiating (Lan et al., 2001; Stricker et al., 2006). This observation may be the explanation why in mice only Osr1 is essential for kidney development.

101 Osr genes are able to promote ectopic kidney expression

An important conclusion from our studies is that osr genes are not only necessary, but also sufficient for kidney development. Thus, we have shown that any of the osr genes from Xenopus, zebrafish or even the Drosophila odd can ectopically activate kidney markers.

Interestingly, the ectopic expression of osr genes is able to trigger early and late kidney markers but not a specific glomerulus marker, wt1a, suggesting that the ectopic renal tissue fails to form this structure. This could be a consequence of the activation of pax2/8, promoted by osr genes, that leads to wt1a downregulation (Majumdar et al., 2000). Moreover, osr1 function has been recently related to glomerular maturation rather than with progenitors specification (Mudumana et al., 2008). Similar to our results, it was demonstrated that overexpression of Osr1 in chicken is also able to induce early markers of kidney precursor cells, although, in contrast to what we found in Xenopus, it inhibits the production of differentiated kidney tubules (James et al., 2006).

Interestingly, the ectopic renal tissue originated by overexpression of Xosr or Xlim and Xpax8 is limited to the dorsal region of the embryo (our results and (Carroll and Vize, 1999)). This spatial restriction in the formation/expansion of kidney tissue could be related with the dorsal/ventral gradient of BMP signaling. Indeed, highest levels of BMP gradient seems to inhibit intermediate mesoderm fates, such as Osr1 expression and renal development (James and Schultheiss, 2005). Consistently with this, the domains competent for renal development correspond with intermediate and lower levels of the BMP gradient, at the medial and dorsal regions of the Xenopus embryo respectively.

Despite the intense effort devoted to understand the formation of the ectopic kidney, there are still some aspects of its formation that remain unclear today. First, this process cannot be explained by a change of fate of dorsal mesoderm derivatives, since the ectopic kidney is not correlated with the loss of muscle. Second, the formation of the ectopic kidney tissue occurs in patches although the osr protein distribution is broader. The differentiation of osr- expressing cells into kidney tissue can be blocked by some still unknown lateral inhibition processes, as suggested by the overgrowth of the neural tube in association with the ectopic renal tissue.

Role for osr genes during pectoral fin induction

We have been able to demonstrate in this study that both osr genes are crucial for

102 zebrafish early pectoral fin formation. Although the onset of expression of osr genes in the fin occurs only around 48 hpf, the effect of loss of function is traced back to the 21 somites stage. At this stage of development, the (individual and double) morphant embryos already show a reduction in the expression of tbx5, the earliest limb specification marker. Interestingly, this occurs when osr genes are expressed in the intermediate mesoderm at the pronephros level. How can these genes act in the lateral plate mesoderm (LPM) where they seem not to be expressed? The hypothesis raised to explain this observation was that osr genes could act indirectly in the induction of the pectoral fin bud. When these genes are expressed in the IM, the precursors of kidney structures, they could signal for the adjacent LPM. The effect of IM in limb development has been already suggested but interestingly osr genes were never considered before as candidates to mediate this role until our study (Geduspan and Solursh, 1992; Stephens and McNulty, 1981).

In this work we have demonstrated that wnt2ba expression was affected in the knock down of both genes and in the osr1 knock down. This molecule that has been previously reported to play an important role in the fin/limb genetic cascade was considered a candidate to mediate osr action because it is a diffusible molecule, able to transfer the signal information from IM to LPM, and it was also described to be essential for tbx5 activation (Ng et al., 2002). Indeed, supporting this hypothesis, we observed that wnt2ba has an overlapping expression pattern with both osr genes in the IM (Fig.6 Chapter 2). In agreement with our results, other studies shown that Osr genes start to be activated in the IM before Wnt2b onset of expression in other vertebrates and also that they have overlapping expression domains (Kawakami et al., 2001; Mercader et al., 2006; Mudumana et al., 2008; Ng et al., 2002; Stricker et al., 2006; Tena et al., 2007). The knockdown of both osr genes caused a stronger reduction (qualitatively) in wnt2ba expression than the loss of function of individual osr1. On the other hand, the loss of function of osr2 by itself did not alter wnt2ba expression suggesting that osr2 is not required for wnt2ba activation. Despite that, morphants of osr2 showed reduction of tbx5 expression and defects in the pectoral fin. This data suggests that osr can modulate the expression of other IM components that mediate the signalling that forms pectoral fins. Moreover, supporting that osr genes act in the fin formation through the Wnt pathway, the loss of function phenotype of osr genes is partially rescued when the canonical Wnt pathway is activated (Fig.5 Chapter 2).

We also demonstrate that expression of osr genes is regulated by the RA signaling. The loss of function of both osr genes that leads to fin impairment is not compensated by the

103 activation of the RA signaling. In addition, osr genes are not able to regulate the expression of the limitating enzyme of RA synthesis. These data strengthen the idea that RA signaling is the first player in the genetic fin/limb cascade modulating the expression of the downstream targets of the pathway such as osr genes. RA is able to modulate osr expression not only in zebrafish (Fig.7 Chapter 2) but also in other vertebrate systems such as Xenopus animal caps and mouse embryonic stem cells (Drews et al., 2011; Mae et al., 2010). The functional requirement of RA signaling has been already described for kidney and limb development and its function is conserved throughout studied vertebrates such as zebrafish, medaka, Xenopus and mice (Begemann et al., 2001; Cartry et al., 2006; Fujii et al., 1997; Gibert et al., 2006; Grandel and Brand, 2011; Grandel et al., 2002; Mic et al., 2004; Negishi et al.; Niederreither et al., 1997; Zhao et al., 2009).

Therefore we propose a model where the RA signaling, originated in the somites, is necessary for the expression of genes at the IM level such as osr genes (Fig.7 Chapter 2) and wnt2ba (Mercader et al., 2006). Osr are essential at the IM for kidney formation and are also involved indirectly in the pectoral fin development through the activation of wnt2ba. This diffusible molecule co-expressed with osr genes at the kidney anlage propagates to the LPM to keep tbx5 expression and induce pectoral fin formation (Fig. A). Moreover, osr genes are able to modulate the levels of fibin, which is positioned in this genetic cascade upstream of tbx5 (Wakahara et al., 2007).

tbx5

RA

wnt2ba osr1 fibin osr2

Figure A - Schematic representation of the genetic cascade that controls fin initiation in zebrafish embryos. The somites (red) are the source of retinoic acid (RA). This signalling activates the expression of osr genes (osr1 in orange and osr2 in yellow) in intermediate mesoderm. The expression of diffusible molecule wnt2ba (green) depends of osr genes. The wnt molecule is able to induce tbx5 (blue) expression, probably through fibin (light blue). Both fibin and tbx5 are expressed in the adjacent lateral plate mesoderm.

104 Contrary to our observations, Gibert and colleagues considered the role of IM for the pectoral fin initiation dispensable, since RA signaling in the somites is sufficient for triggering this process (Gibert et al., 2006). Nevertheless the recovery of somitic RA signaling in aldh1a2 mutants could be sufficient for the triggering of IM genes involved in fin initiation.

Partial redundant function of osr genes

Our results suggest that the function of the osr proteins can be largely interchangeable, resembling the previous observations made in mice, in which individual knockouts of Osr1 or Osr2 present phenotypes correlated with their specific expression pattern (James et al., 2006; Lan et al., 2004; Wang et al., 2005). The Osr1 null mice show heart, kidney and urogenital defects while the Osr2 null mice show cleft palate and open eyelids at birth (James et al., 2006; Lan et al., 2004; Wang et al., 2005). These individual mutants lack defects in the structures where both genes are co-expressed during embryonic development (Gao et al., 2009; Lan et al., 2001; Lan et al., 2004; Wang et al., 2005), suggesting the partial functional redundancy of Osr genes. Moreover, the knock-in of Osr1 and Osr2A in the Osr2 coding region are able to completely rescue cleft palate and cranial skeleton defects shown in Osr2- /- mice. Nevertheless, the knock-in alleles are not able to rescue the phenotype of open eyelids at birth, probably due to the differences in expression of the endogenous gene and the knock-in alleles during eyelid development (Gao et al., 2009). These results strongly suggest that the distinct functions of both mammalian Osr genes are a consequence of divergence of spatiotemporal expression rather than change of coding regions (Gao et al., 2009). In our work we observed that osr genes are required additively for the pronephros formation in zebrafish and Xenopus. Recently, a specific limb mesenchyme inactivation of Osr1 in Osr2 -/- mutant mice revealed the requirement of both Osr genes for joint development (Gao et al., 2011). Also the formation of irregular connective tissue (ICT) fibroblast differentiation in chicken is affected only when both Osr genes are affected (Stricker et al., 2011). These data supports the idea that Osr genes act redundantly in developmental processes where they are co-expressed such as ICT, synovial joints, kidney, craniofacial structures and limb (Gao et al., 2011; Stricker et al., 2011). However, in the pectoral fin formation the loss of function of osr1 produces continuously stronger phenotype than the osr2 loss of function. This difference can be explained by the broader and earlier expression of osr1 in the majority of vertebrates (Lan et al., 2001; So and Danielian, 1999; Stricker et al., 2006; Tena et al., 2007). Moreover, the idea of functional divergency of osr genes based on

105 differential expression pattern proposed by Gao and colleagues is also supported by the evidence that both proteins are able to partial rescue the loss of tbx5 expression in the embryos injected with both morpholinos (Fig.4 Chapter 2).

Involvement of osr genes in the formation of endoderm derivatives

In this work and in collaboration with Rankin and colleagues, we revealed the requirement of osr genes in the development of some endoderm structures such as lung and liver. Moreover, osr genes seem to use the same signaling molecule, wnt2b, to modulate the downstream targets that trigger lung development and fin/limb development, such as nkx2.1 and tbx5, respectively. The involvement of osr genes in the formation of endoderm derivatives as well as the molecular signaling seem to be conserved in Xenopus and zebrafish.

The expression pattern of osr genes comprehends also some endodermal territories adjacent to the pronephros, the liver and also the lung epithelium and surrounding mesenchyme (Mudumana et al., 2008; Tena et al., 2007) (Rankin, Neto et al, in preparation).

In collaboration with Rankin and colleagues, we performed loss of function studies and determine that both Xosr genes are required for the formation of the lung (Rankin, Neto et al, in preparation), but not for the expression of most of the endoderm markers assessed. In agreement with our zebrafish data, loss of function of both Xosr leads to the reduction of expression of Xwnt2b. Indeed, Wnt2/Wnt2b are required for the specification of lung endoderm progenitors within the anterior foregut and are also able to promote this fate (Goss et al., 2009). Also the double osr morphant embryos show downregulation of nkx2.1, the earliest marker of lung endoderm described.

The function of osr1 in the zebrafish renal development was linked with the vascular system, an endoderm derivative. Indeed osr1 seems to be required for a proper balance between kidney (mesoderm) and the vascular system (endoderm). The specific reduction of the anterior portion of the kidney showed by osr1 morphants is associated with a compensatory increase of the vascular system (Mudumana et al., 2008).

We also showed that osr genes are required for the LPM expression of the wnt2bb (Fig.X Chapter 2), as it occurs with the other homolog, wnt2ba, which is also regulated at least by osr1 expression. Nevertheless wnt2bb is involved in liver specification (Negishi et al.; Ober et

106 al., 2006) distinct from wnt2ba that triggers the formation of a LPM derivative, the fin.

The zinc-finger (ZF) transcription factors encoded by osr genes act as repressors in the context of kidney development

We have observed that XOsr transcription factors behave molecularly as repressors during kidney development. Overexpression of XOsr fusion proteins with the repressor domain or the activation domain allow us to identify that the fusion with the repressor domain behaves as the native protein, promoting the formation of the ectopic kidney tissue in Xenopus. Similar studies in chicken mesenchymal cells also concluded that Osr genes act predominantly as transcriptional repressors in the process of cartilage condensation (Stricker et al., 2011). The vertebrate ZF proteins behave as repressors similarly with the Drosophila odd that also acts as transcriptional repressor during embryonic segmentation. This process is mediated by engrailed-like domain 1 (eh1) that is able to recruit the co-repressor Groucho (Goldstein et al., 2005). In this work, we observed in Xenopus that nephrogenic capability of the Drosophila odd protein is also dependent of the eh1 domain. This data suggests that vertebrate proteins are able to recruit vertebrate homologous of Groucho, members of the family of Transducin-like Enhancer of Split (TLS) (Chen and Courey, 2000). In fact, a putative eh1 domain is present in both Xosr proteins and is located in the N-terminal to the zinc fingers while in odd is located at the C-terminal end.

There is some controversy about the functional role of the isoforms of Osr2. It has been reported that the two Osr2 isoforms when fused to the Gal4 DNA binding domain display opposite transcriptional activity in a cell culture based reporter assay (Kawai et al., 2005). However, a recent work claims that Osr2A and Osr2B have the same functional potential when expressed in the same tissue (Gao et al., 2009), suggesting that the distinct results obtained in cell culture assays represent a non-physiological activity.

Osr genes are able to activate the expression of early pronephric markers Xlim1 and Xpax8, although they have a repressor activity, which implicates the presence of an intermediate repressor in the genetic cascade. The transcription factors Foxc1 and Foxc2 present some characteristics that fulfill this role. They are necessary for somites formation (Topczewska et al., 2001), and are necessary and sufficient to repress the expression of genes of the intermediate mesoderm such as Osr1 and Lim1 (Wilm et al., 2004). However the loss of function of Osr1 and Osr2 promotes the reduction of the kidney tissue but is not correlated with the expansion of the somites.

107 Osr genes are involved in several developmental processes originating endoderm-, mesoderm- and lateral plate mesoderm-derivatives. We show that the IM acquires a relevant role in the link of the RA signaling pathway derived from the somites with the formation of two distinct structures, fin and kidney. Our data support a relay mechanism where RA activates osr genes that will induce the expression of the diffusible molecule wnt2ba, which is able to trigger the expression of tbx5, an essential gene for fin/limb formation. Moreover, we show that osr genes ability to regulate Wnt signaling is conserved in Xenopus and zebrafish and that this mechanism is also conserved in distinct developmental processes.

108 References:

Begemann, G., T.F. Schilling, G.J. Rauch, R. Geisler, and P.W. Ingham. 2001. The zebrafish neckless mutation reveals a requirement for raldh2 in mesodermal signals that pattern the hindbrain. Development. 128:3081-94. Carroll, T.J., and P.D. Vize. 1999. Synergism between Pax-8 and lim-1 in embryonic kidney development. Dev Biol. 214:46-59. Cartry, J., M. Nichane, V. Ribes, A. Colas, J.F. Riou, T. Pieler, P. Dolle, E.J. Bellefroid, and M. Umbhauer. 2006. Retinoic acid signalling is required for specification of pronephric cell fate. Dev Biol. 299:35-51. Chen, G., and A.J. Courey. 2000. Groucho/TLE family proteins and transcriptional repression. Gene. 249:1-16. Drews, C., S. Senkel, and G.U. Ryffel. 2011. The nephrogenic potential of the transcription factors osr1, osr2, hnf1b, lhx1 and pax8 assessed in Xenopus animal caps. BMC Dev Biol. 11:5. Drummond, I.A., A. Majumdar, H. Hentschel, M. Elger, L. Solnica-Krezel, A.F. Schier, S.C. Neuhauss, D.L. Stemple, F. Zwartkruis, Z. Rangini, W. Driever, and M.C. Fishman. 1998. Early development of the zebrafish pronephros and analysis of mutations affecting pronephric function. Development. 125:4655-67. Fujii, H., T. Sato, S. Kaneko, O. Gotoh, Y. Fujii-Kuriyama, K. Osawa, S. Kato, and H. Hamada. 1997. Metabolic inactivation of retinoic acid by a novel P450 differentially expressed in developing mouse embryos. EMBO J. 16:4163-73. Gao, Y., Y. Lan, H. Liu, and R. Jiang. 2011. The zinc finger transcription factors Osr1 and Osr2 control synovial joint formation. Dev Biol. 352:83-91. Gao, Y., Y. Lan, C.E. Ovitt, and R. Jiang. 2009. Functional equivalence of the zinc finger transcription factors Osr1 and Osr2 in mouse development. Dev Biol. 328:200-9. Geduspan, J.S., and M. Solursh. 1992. A growth-promoting influence from the mesonephros during limb outgrowth. Dev Biol. 151:242-50. Gibert, Y., A. Gajewski, A. Meyer, and G. Begemann. 2006. Induction and prepatterning of the zebrafish pectoral fin bud requires axial retinoic acid signaling. Development. 133:2649-59. Goldstein, R.E., O. Cook, T. Dinur, A. Pisante, U.C. Karandikar, A. Bidwai, and Z. Paroush. 2005. An eh1-like motif in odd-skipped mediates recruitment of Groucho and repression in vivo. Mol Cell Biol. 25:10711-20. Goss, A.M., Y. Tian, T. Tsukiyama, E.D. Cohen, D. Zhou, M.M. Lu, T.P. Yamaguchi, and E.E. Morrisey. 2009. Wnt2/2b and beta-catenin signaling are necessary and sufficient to specify lung progenitors in the foregut. Dev Cell. 17:290-8. Grandel, H., and M. Brand. 2011. Zebrafish limb development is triggered by a retinoic acid signal during gastrulation. Dev Dyn. Grandel, H., K. Lun, G.J. Rauch, M. Rhinn, T. Piotrowski, C. Houart, P. Sordino, A.M. Kuchler, S. Schulte-Merker, R. Geisler, N. Holder, S.W. Wilson, and M. Brand. 2002. Retinoic acid signalling in the zebrafish embryo is necessary during pre-segmentation stages to pattern the anterior-posterior axis of the CNS and to induce a pectoral fin bud. Development. 129:2851-65. James, R.G., C.N. Kamei, Q. Wang, R. Jiang, and T.M. Schultheiss. 2006. Odd-skipped related 1 is required for development of the metanephric kidney and regulates formation and differentiation of kidney precursor cells. Development. 133:2995-3004. James, R.G., and T.M. Schultheiss. 2005. Bmp signaling promotes intermediate mesoderm gene expression in a dose-dependent, cell-autonomous and translation-dependent manner. Dev Biol. 288:113-25.

109 Kawai, S., T. Kato, H. Inaba, N. Okahashi, and A. Amano. 2005. Odd-skipped related 2 splicing variants show opposite transcriptional activity. Biochem Biophys Res Commun. 328:306-11. Kawakami, Y., J. Capdevila, D. Buscher, T. Itoh, C. Rodriguez Esteban, and J.C. Izpisua Belmonte. 2001. WNT signals control FGF-dependent limb initiation and AER induction in the chick embryo. Cell. 104:891-900. Lan, Y., P.D. Kingsley, E.S. Cho, and R. Jiang. 2001. Osr2, a new mouse gene related to Drosophila odd-skipped, exhibits dynamic expression patterns during craniofacial, limb, and kidney development. Mech Dev. 107:175-9. Lan, Y., C.E. Ovitt, E.S. Cho, K.M. Maltby, Q. Wang, and R. Jiang. 2004. Odd-skipped related 2 (Osr2) encodes a key intrinsic regulator of secondary palate growth and morphogenesis. Development. 131:3207-16. Mae, S., S. Shirasawa, S. Yoshie, F. Sato, Y. Kanoh, H. Ichikawa, T. Yokoyama, F. Yue, D. Tomotsune, and K. Sasaki. 2010. Combination of small molecules enhances differentiation of mouse embryonic stem cells into intermediate mesoderm through BMP7-positive cells. Biochem Biophys Res Commun. 393:877-82. Majumdar, A., K. Lun, M. Brand, and I.A. Drummond. 2000. Zebrafish no isthmus reveals a role for pax2.1 in tubule differentiation and patterning events in the pronephric primordia. Development. 127:2089-98. Mercader, N., S. Fischer, and C.J. Neumann. 2006. Prdm1 acts downstream of a sequential RA, Wnt and Fgf signaling cascade during zebrafish forelimb induction. Development. 133:2805-15. Mic, F.A., I.O. Sirbu, and G. Duester. 2004. Retinoic acid synthesis controlled by Raldh2 is required early for limb bud initiation and then later as a proximodistal signal during apical ectodermal ridge formation. J Biol Chem. 279:26698-706. Mudumana, S.P., D. Hentschel, Y. Liu, A. Vasilyev, and I.A. Drummond. 2008. odd skipped related1 reveals a novel role for endoderm in regulating kidney versus vascular cell fate. Development. 135:3355-67. Negishi, T., Y. Nagai, Y. Asaoka, M. Ohno, M. Namae, H. Mitani, T. Sasaki, N. Shimizu, S. Terai, I. Sakaida, H. Kondoh, T. Katada, M. Furutani-Seiki, and H. Nishina. Retinoic acid signaling positively regulates liver specification by inducing wnt2bb gene expression in medaka. Hepatology. 51:1037-45. Ng, J.K., Y. Kawakami, D. Buscher, A. Raya, T. Itoh, C.M. Koth, C. Rodriguez Esteban, J. Rodriguez-Leon, D.M. Garrity, M.C. Fishman, and J.C. Izpisua Belmonte. 2002. The limb identity gene Tbx5 promotes limb initiation by interacting with Wnt2b and Fgf10. Development. 129:5161-70. Niederreither, K., P. McCaffery, U.C. Drager, P. Chambon, and P. Dolle. 1997. Restricted expression and retinoic acid-induced downregulation of the retinaldehyde dehydrogenase type 2 (RALDH-2) gene during mouse development. Mech Dev. 62:67-78. Ober, E.A., H. Verkade, H.A. Field, and D.Y. Stainier. 2006. Mesodermal Wnt2b signalling positively regulates liver specification. Nature. 442:688-91. So, P.L., and P.S. Danielian. 1999. Cloning and expression analysis of a mouse gene related to Drosophila odd-skipped. Mech Dev. 84:157-60. Stephens, T.D., and T.R. McNulty. 1981. Evidence for a metameric pattern in the development of the chick humerus. J Embryol Exp Morphol. 61:191-205. Stricker, S., N. Brieske, J. Haupt, and S. Mundlos. 2006. Comparative expression pattern of Odd-skipped related genes Osr1 and Osr2 in chick embryonic development. Gene Expr Patterns. 6:826-34.

110 Stricker, S., S. Mathia, J. Haupt, P. Seemann, J. Meier, and S. Mundlos. 2011. Odd-skipped related genes regulate differentiation of embryonic limb mesenchyme and bone marrow mesenchymal stromal cells. Stem Cells Dev. Tena, J.J., A. Neto, E. de la Calle-Mustienes, C. Bras-Pereira, F. Casares, and J.L. Gomez- Skarmeta. 2007. Odd-skipped genes encode repressors that control kidney development. Dev Biol. 301:518-31. Topczewska, J.M., J. Topczewski, A. Shostak, T. Kume, L. Solnica-Krezel, and B.L. Hogan. 2001. The winged helix transcription factor Foxc1a is essential for somitogenesis in zebrafish. Genes Dev. 15:2483-93. Wakahara, T., N. Kusu, H. Yamauchi, I. Kimura, M. Konishi, A. Miyake, and N. Itoh. 2007. Fibin, a novel secreted lateral plate mesoderm signal, is essential for pectoral fin bud initiation in zebrafish. Dev Biol. 303:527-35. Wang, Q., Y. Lan, E.S. Cho, K.M. Maltby, and R. Jiang. 2005. Odd-skipped related 1 (Odd 1) is an essential regulator of heart and urogenital development. Dev Biol. 288:582-94. Wilm, B., R.G. James, T.M. Schultheiss, and B.L. Hogan. 2004. The forkhead genes, Foxc1 and Foxc2, regulate paraxial versus intermediate mesoderm cell fate. Dev Biol. 271:176-89. Zhao, X., I.O. Sirbu, F.A. Mic, N. Molotkova, A. Molotkov, S. Kumar, and G. Duester. 2009. Retinoic acid promotes limb induction through effects on body axis extension but is unnecessary for limb patterning. Curr Biol. 19:1050-7.

111

112

CONCLUSIONS

1. Osr genes are expressed early in the intermediate mesoderm that will originate the embryonic kidney.

2. Osr1 and osr2 are both necessary and sufficient for the kidney development in Xenopus and zebrafish.

3. The Osr transcription factors behave molecularly as repressors during kidney development.

4. Osr genes are expressed in the pectoral fin at around 48 hpf onwards.

5. Osr1 and osr2 are required for early pectoral fin formation.

6. The morphant phenotypes are observed when osr genes are expressed in the intermediate mesoderm, a kidney precursor.

7. The effect of osr genes in fin development is indirect and is probably mediated by wnt2ba.

8. The RA signaling is able to regulate osr expression.

113

114 APPENDIX I

“Dioxin receptor and Slug transcription factors regulate the insulator activity of B1 SINE retrotransposons via an RNA polymerase

switch”

In this work, it was reported that a genome-wide B1 SINE (Short Interspersed Nuclear Element) retrotransposon (B1-X35S) functions as potent insulator when tested in cultured cells and live animals. This insulator activity is mediated by binding of the transcription factors dioxin receptor (AhR) and Slug (Snai2) to consensus elements present in the SINE.

I tested the insulator activity of B1-X35S and the mutant version for AhR and Slug binding sites (B1-Xmut35Smut) in zebrafish in vivo. Results of these experiments are shown on Figure 2 and Supplementar Figure 2 of the published report.

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Research Dioxin receptor and SLUG transcription factors regulate the insulator activity of B1 SINE retrotransposons via an RNA polymerase switch

Angel Carlos Roma´n,1 Francisco J. Gonza´lez-Rico,1 Eduardo Molto´,2,3 Henar Hernando,4 Ana Neto,5 Cristina Vicente-Garcia,2,3 Esteban Ballestar,4 Jose´ L. Go´mez-Skarmeta,5 Jana Vavrova-Anderson,6 Robert J. White,6,7 Lluı´s Montoliu,2,3 and Pedro M. Ferna´ndez-Salguero1,8 1Departamento de Bioquı´mica y Biologı´a Molecular, Facultad de Ciencias, Universidad de Extremadura, 06071 Badajoz, Spain; 2Centro Nacional de Biotecnologı´a (CNB), Consejo Superior de Investigaciones Cientı´ficas (CSIC), Department of Molecular and Cellular Biology, Campus de Cantoblanco, C/Darwin 3, 28049 Madrid, Spain; 3Centro de Investigacio´n Biome´dica en Red de Enfermedades Raras (CIBERER), ISCIII, Madrid, Spain; 4Chromatin and Disease Group, Cancer Epigenetics and Biology Programme, Bellvitge Biomedical Research Institute (IDIBELL), Barcelona 08907, Spain; 5Centro Andaluz de Biologı´a del Desarrollo, CSIC-Universidad Pablo de Olavide, 41013 Sevilla, Spain; 6College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow G12 8QQ, United Kingdom; 7Beatson Institute for Cancer Research, Glasgow, G61 1BD, United Kingdom

Complex genomes utilize insulators and boundary elements to help define spatial and temporal gene expression patterns. We report that a genome-wide B1 SINE (Short Interspersed Nuclear Element) retrotransposon (B1-X35S) has potent in- trinsic insulator activity in cultured cells and live animals. This insulation is mediated by binding of the transcription factors dioxin receptor (AHR) and SLUG (SNAI2) to consensus elements present in the SINE. Transcription of B1-X35S is required for insulation. While basal insulator activity is maintained by RNA polymerase (Pol) III transcription, AHR- induced insulation involves release of Pol III and engagement of Pol II transcription on the same strand. B1-X35S insulation is also associated with enrichment of heterochromatin marks H3K9me3 and H3K27me3 downstream of B1-X35S, an effect that varies with cell type. B1-X35S binds parylated CTCF and, consistent with a chromatin barrier activity, its positioning between two adjacent genes correlates with their differential expression in mouse tissues. Hence, B1 SINE retrotransposons represent genome-wide insulators activated by transcription factors that respond to developmental, oncogenic, or tox- icological stimuli. [Supplemental material is available for this article.]

Mammalian genes are often clustered at chromosomal locations whether they require the epigenetic machinery. In plants, epige- sharing common cis-regulatory elements (Gerasimova and Corces netics appear to play an important role in transposon-mediated 2001; Bushey et al. 2008; Molto´ et al. 2009). Such organization in control of gene expression (Lippman et al. 2004). However, in mouse expression domains poses the problem of misregulation due to and humans no direct link has yet been established between trans- inappropriate action of enhancers (Bell et al. 2001). To cope with poson epigenetics and control of gene expression (Aravin et al. this, insulators have evolved to delineate functionally independent 2007). chromosomal regions (Bell et al. 2001; Gerasimova and Corces A SINE B1-X35S retrotransposon was found in over 14,000 2001; Ohlsson et al. 2001; Mukhopadhyay et al. 2004) that might instances in the mouse genome that contains a dioxin receptor be exploited to improve the success of gene-therapy protocols and (AHR) binding site (XRE) (common to canonical B1 elements) and transgenesis experiments by modulating gene expression (Giraldo a SLUG (SNAI2) site for recruitment of the zinc-finger transcrip- et al. 2003; Recillas-Targa et al. 2004; Molto´ et al. 2009). tion factor SLUG. Sequence conservation analysis revealed that B1- It is known that transposable elements such as ERV1, ERVK, X35S is a new SINE-B1subfamily differing from canonical elements MIR1, LTR10, and MER61 contain binding sites for transcription by the presence of the SLUG site (Roman et al. 2008). We previously factors (Wang et al. 2007; Bourque et al. 2008; Kuwabara et al. observed that B1-X35S is enriched in promoter regions (1398 2009; Kunarso et al. 2010). Moreover, a B2 SINE retrotransposon genes) and that AHR and SLUG binding represses transcription of has been shown to insulate expression domains during murine downstream genes. However, the mechanism by which B1-X35S- organogenesis (Lunyak et al. 2007). Therefore, it is of great interest containing genes are repressed by AHR and SLUG remained un- to determine whether retrotransposons interact with specific tran- solved. Here, we demonstrate that B1-X35S SINE can also function scription factors to define genome-wide expression profiles and as an insulator. This insulation mechanism is complex, involving transcription by Pol III and Pol II, accumulation of heterochromatic marks at downstream regions, and binding of CTCF. AHR is a well- 8 Corresponding author. conserved and ubiquitously expressed basic-helix-loop-helix (bHLH) E-mail [email protected]; fax 34-924-289419. Article published online before print. Article, supplemental material, and pub- transcription factor with a prominent role in cell physiology and lication date are at http://www.genome.org/cgi/doi/10.1101/gr.111203.110. homeostasis and in the response to environmental toxins (Furness

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AHR and SLUG trigger insulation by B1 SINE repeats et al. 2007; Gomez-Duran et al. 2009). SLUG has a critical role in Mutation of the XRE site (B1-Xmut35S) reduces insulation, while epithelial-to-mesenchymal transitions that occur during devel- mutation of the E-box does not have a significant effect (Fig. 1A). opment and in certain pathological conditions, such as tumor me- Unlike AHR, basal levels of SLUG and SNAIL are very low in HEK tastasis (Thiery et al. 2009; Martinez-Estrada et al. 2010). B1-X35S 293 cells (Supplemental Fig. 1C), possibly accounting for the lack SINEs repress gene expression in vivo as the result of AHR and SLUG of effect of single E-box mutation under basal cell conditions. binding to their cognate consensus elements (Roman et al. 2008). However, double XRE+E-box mutation (B1-Xmut35Smut) repro- The abundance of B1-X35S in the mouse genome suggests a wide- ducibly reduced the insulation effect of single XRE mutation (Fig. spread impact on gene expression. This may be important during 1A), suggesting functional interactions between factors bound at normal development as well as in pathological conditions. these sites. SLUG and SNAIL might compete to bind B1-X35S because they share the same response element. To test this, mouse Hepa-1 Results cells were transiently transfected with SLUG or SNAIL and chro- matin immunoprecipitation (ChIP) was used to assay their binding B1-X35S is a SINE retrotransposon with basal and transcription to the promoter of five B1-X35S-containing genes (Dad1, Lpp, factor-inducible insulator activity Cabin1, Tbc1d1, and Rtl1; see Supplemental Fig. 1D for location). B1-X35S has consensus-binding sites for AHR (XRE) and SLUG and SLUG expression increased its own binding to B1-X35S and re- SNAIL (E-box) (Supplemental Fig. 1A). Genomic analysis showed duced that of SNAIL, while the converse was true for SNAIL ex- that B1-X35S has a prevalence of 14% in proximal promoters (<10 pression (Fig. 1B). Consistent with the EBA results, reducing basal kb), 18% in distal promoters (10–50 kb), 48% in nonpromoter gene AHR levels with a specific shRNA severely impaired binding of sequences, and 20% in any other genomic locations, additively both SLUG and SNAIL to B1-X35S in vivo (Fig. 1C), indicating that giving a 32% occurrence in upstream gene regulatory regions AHR facilitates SLUG and SNAIL recruitment to B1-X35S-con- (Supplemental Fig. 1B). To investigate the potential insulator ac- taining promoters (individual results for each gene and experi- tivity of B1-X35S, we performed enhancer-blocking assays (EBA) in mental condition shown in Fig. 1B,C are detailed in Supplemental human HEK 293 cells. EBA revealed that B1-X35S has a potent Table 1). Consistent with this, sequential ChIP (re-ChIP) demon- intrinsic insulator activity, similar to that of the chicken 1.2-kb strated that AHR co-occupies with SLUG the B1-X35S element of 59HS4 beta-globin insulator (Fig. 1A; Recillas-Targa et al. 1999). Dad1 and Tbc1d1 (Fig. 1D).

Figure 1. B1-X35S is a SINE with insulator activity. (A) Wild-type B1-X35S or its mutant forms Xmut35S (AHR site mutated), X35Smut (E-box mutated), and Xmut35Smut (both sites mutated) were transiently transfected and their insulator activity analyzed by EBA. The constructs are illustrated at the top of the figure. Data are presented as fold-enhancer blocking activity normalized to the reference pELuc vector. (B) Occupancy of SLUG and SNAIL on B1-X35S was analyzed by qChIP after transient transfection of either protein. (C ) Effects of AHR expression on SLUG and SNAIL recruitment to B1-X35S were determined by qChIP after transient transfection of AHR-specific shRNA or scrambled shRNA as negative control. (D) Simultaneous presence of AHR and SLUG on B1-X35S was analyzed by sequential qChIP. A first ChIP used anti-SLUG antibody and the immunoprecipitated DNA was re-ChIPed with anti-AHR antibody. Re-ChIP for GAPDH provides a negative control. Data were quantified with respect to input DNA from the first ChIP. Results for Dad1 and Tbc1d1 are shown. (E) The effect of AHR, SLUG, and SNAIL expression on the insulator activity of wild-type B1-X35S was analyzed by EBA and quantified as above. (F) The experiment in E was done using the double-mutant B1-Xmut35Smut. Human HEK 293 cells were used in A, E,andF, while mouse Hepa-1 cells were used in B, C,andD.(B,C) The average data from the five B1-X35S containing genes Dad1, Lpp, Cabin1, Tbc1d1,andRtl1. Individual results for each gene and experimental condition (B,C) are detailed in Supplemental Table 1. Data are shown as mean 6SD. (*) P = 0.033 and (**) P = 0.009 with respect to B1-X35S.

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As expected, individual increases in AHR or SLUG expression raise the in- sulator activity of B1-X35S, while coex- pression of both has an additive effect, further strengthening insulation (Fig. 1E). Surprisingly, SNAIL expression had little effect on the EBA (Fig. 1E), suggesting that even though it can bind B1-X35S, it does not promote insulation. We therefore fo- cused the remainder of the study on AHR and SLUG. The effects of these proteins require their recognition motifs, as trans- fection of AHR, SLUG, or SNAIL did not significantly alter the lower basal EBA of the double-mutant B1-Xmut35Smut trans- poson (Fig. 1F).

B1-X35S has insulator activity in zebrafish in vivo Zebrafish has been established as a pow- erful model to analyze cis-regulatory genomic elements in vivo (Bessa et al. 2009). Zebrafish is a good model because it expresses three functional forms of AHR (AhR1a, the ortholog of mammalian AHR, AhR1b, and AhR2) (Mathew et al. 2009) and an ortholog of SLUG (Katoh 2005). B1-X35S and its mutant version B1-Xmut35Smut were cloned in the enhancer+promoter-driven GFP vector, as indicated in Figure 2A. These con- structs were microinjected into one-cell zebrafish embryos and GFP expression quantified in CNS (enhancer-dependent and susceptible to insulator regulation) and somites (muscle promoter-driven and Figure 2. B1-X35S has insulator activity in zebrafish. (A) B1-X35S and mutant B1Xmut35Smut in the insulation-independent). B1-X35S has enhancer+promoter-driven GFP vector, as illustrated, were microinjected into one-cell zebrafish em- a potent insulator activity in zebrafish, bryos. Each animal was photographed and GFP signals quantified in CNS and somites. Measurements of down-regulating CNS enhancer-driven individual zebrafish are shown in Supplemental Figure 2. (B) Insulator activity was calculated as the GFP expression (Fig. 2A). Mutation of AHR median GFP signal in muscle/CNS for each experimental condition. Data are shown as mean 6SD. and SLUG-binding sites significantly re- duced insulation (Fig. 2A). GFP signals in CNS and somites were not have a significant effect. Consistent with the EBA response quantified in each individual animal as the GFP ratio in muscle/CNS (Fig. 1), XRE+E-box double mutation additively promotes Pol III (Supplemental Fig. 2). The inter-individual variation is due to the and TFIIIC binding to B1-X35S (Fig. 3A). This can be explained if integration of the transgene at different genomic locations in dif- binding of AHR and SLUG interferes with binding of TFIIIC and, ferent somatic cells—they are not stable lines resulting from a hence, Pol III. To further test this, Hepa-1 cells were transfected and unique integration in the germ line. The median fluorescence occupancy was analyzed by qChIP of five B1-X35S loci. Increased ratios revealed that wild-type B1-X35S, but not the double mu- expression of AHR, with or without SLUG, strongly suppresses tant, triggers a strong insulation response (Fig. 2B). Thus, B1-X35S binding to B1-X35S by Pol III, TFIIIC, and TFIIIB (Fig. 3B; Supple- is a B1 SINE with insulator activity in vivo. mental Table 1). These effects are specific, since histone H3 binding to B1-X35S is unaltered (Fig. 3C), as is Pol III binding to its un- related target tRNALeu gene (Fig. 3D). AHR or SLUG expression AHR and SLUG modulate Pol III binding to B1-X35S does not alter Pol III protein levels in Hepa-1 cells (Supple- Pol III and Pol II transcription are implicated in insulation mental Fig. 3). (Valenzuela and Kamakaka 2006; Lunyak et al. 2007; Lunyak 2008). Like canonical B1 repeats, B1-X35S has consensus A and B B1-X35S is transcribed by Pol III, and AHR enhances boxes for binding transcription factor TFIIIC, which then recruits transcription by promoting exchange of Pol III by Pol II transcription factor TFIIIB and Pol III (Supplemental Fig. 1A). DNA- binding affinity assays confirmed that Pol III and its assembly Although SLUG inhibits B1-X35S transcription in vitro, AHR factor TFIIIC can bind B1-X35S in vitro (Fig. 3A). Mutation of the produces an unexpected increase (Fig. 4A). In both cases, the re- XRE element increases binding of both, while E-box mutation does sponses depend on the respective binding motifs. SLUG ablates the

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least in part, on an increase in transcrip- tion due to a switch to Pol II from Pol III.

Pol III and Pol II transcribe B1-X35S in the same direction Some B2 SINEs carry a Pol II promoter that is orientated in the opposite direc- tion to the Pol III promoter (Ferrigno et al. 2001). Similarly, the insulator ac- tivity of a B2 SINE involves its transcrip- tion from opposite strands by Pol III and Pol II (Lunyak et al. 2007). To clarify the direction of transcription of B1-X35S, we used strand-specific RT–qPCR with sense and antisense primers, as schematized in Figure 5A. The level of RT-generated products was clearly detected in the an- tisense, but not the sense direction, sug- gesting that most expression is from one Figure 3. Pol III binds B1-X35S: role of AHR and SLUG. (A) In vitro binding of Pol III and TFIIIC to B1- strand. Pol II inhibition with a-AMA X35S, B1-Xmut35S, B1-X35Smut, or B1-Xmut35Smut was analyzed by DNA-binding affinity assays. (B) qChIP to address whether AHR and SLUG expression block Pol III recruitment to B1-X35S in vivo after decreased the amount of antisense RT transient transfection of Hepa-1 cells with AHR alone or with SLUG. (C,D) AHR was expressed with or product by 57%, but had no effect on the Leu without SLUG and their effects on histone H3 binding to B1-X35S (C ) or Pol III binding to tRNA genes low levels of sense RT product (Fig. 5A). (D) determined by qChIP. (B,C) Average data from five B1-X35S-containing genes (Dad1, Lpp, Cabin1, Our data therefore suggest that Pol III Tbc1d1,andRtl1). Data are shown as mean 6SD. and Pol II transcribe B1-X35S in the same direction (Fig. 5A, solid arrows). effect of AHR on B1-X35S transcription, perhaps because AHR fa- Occupancy of B1-X35S by Pol III and Pol II was further ana- vors binding of inhibitory SLUG (see Figs. 1C, 3A,B). The positive lyzed by sequential ChIP assays (Re-ChIP). Immunoprecipita- effect of AHR on B1-X35S transcription was also found in vivo in tion of Pol II on B1-X35S DNA from Pol III ChIPs revealed that transfected Hepa-1 (Fig. 4B). Therefore, AHR expression increases Pol III prevails under basal conditions over Pol II (Fig. 5B). Muta- B1-X35S transcription despite inhibiting Pol III binding. tion of the A and B boxes impairs Pol III binding to B1-X35S in Some B2 SINEs carry active Pol II promoters (Ferrigno et al. DNA-binding affinity assays (Fig. 5C). EBA experiments showed 2001). Furthermore, Pol II-dependent transcription may influence that A+B mutation severely impairs the intrinsic insulator activity B2 insulator function (Lunyak et al. 2007). We therefore tested of B1-X35S (Fig. 5D). This supports an important role for Pol III in whether AHR activates B1-X35S transcription through Pol II. basal SINE-mediated insulation. Moreover, mutation of the A and qChIP revealed that AHR expression increases Pol II binding to B boxes abolished the increase in insulator activity induced by B1-X35S (Fig. 4C; Supplemental Table 1) without raising Pol II AHR overexpression, suggesting that the enhanced insulation as- protein levels (Supplemental Fig. 4). sociated with Pol II requires the Pol III promoter. Overall, these In Hepa-1 cells, the Pol III inhibitor tagetitoxin (TGT) re- data strongly suggest that high AHR levels promote the release of duces B1-X35S transcription by 70%, whereas a-amanitin (a-AMA) Pol III and the engagement of Pol II on B1-X35S, with a concomi- at a concentration specifically inhibiting Pol II (Rollins et al. tant increase in transcription and in insulator activity. B1-X35S does 2007) decreases it by 50% (Fig. 4D). An increase in AHR expression not have a canonical TATA box, but it contains a similar sequence attenuates the inhibitory effect of TGT to 45%, but increases the (59-TTAAT-39). Whether this is the binding site for Pol II remains to inhibition caused by a-AMA to 80% (Fig. 4D). These data indicate be explored. that both Pol III and Pol II participate in B1-X35S transcription, with Pol III contributing more than Pol II. However, an increase B1-X35S insulation correlates with epigenetic marks in AHR expression changes the ratio in favor of Pol II (Fig. 4D). of heterochromatin and with CTCF binding, and both Indeed, a-AMA efficiently inhibits in vivo the increase in B1-X35S properties are modulated by AHR transcription caused by AHR expression (Fig. 4B), further sup- porting that such effect is Pol II dependent. Comparison of the To analyze the relationship between B1-X35S and epigenetic marks inhibition caused by a-AMA and the general transcriptional blocker that might sustain its insulator function, we analyzed by qChIP the actinomycin D also indicates that Pol II has a significant contri- profile of heterochromatin marks in histone H3 within a À1000 bution to AHR-dependent B1-X35S transcription (Fig. 4B). The im- to +1500 region upstream and downstream from B1-X35S in Dad1, portance of AHR in this Pol III–Pol II exchange is supported Lpp, and Cabin1 (Fig. 6A; Supplemental Table 2). Histone H3 tri- by two additional sets of experiments. First, XRE mutation (B1- methyl lysine 9 (H3K9me3) (Fig. 6B) and trimethyl lysine 27 Xmut35S) abolishes the inhibitory effect of a-AMA, but not that (H3K27me3) (Fig. 6C) marks are lower upstream of B1-X35S and of TGT (Fig. 4E). Second, B1-X35S transcription in extracts of cells increase downstream from the transposon. AHR+SLUG expression lacking AHR expression (T-FGM AhRÀ/À) is insensitive to a-AMA enhances significantly H3K9me3 and H3K27me3 levels down- but remains susceptible to TGT (Fig. 4F). It seems that the basal stream from the repeat (Fig. 6B,C). insulator activity of B1-X35S is maintained by Pol III transcription CTCF is a transcription factor with a role in the control of primarily, while the enhanced insulation caused by AHR relies, at genomic boundaries and insulators (Bushey et al. 2008). Binding of

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Figure 4. B1-X35S is transcribed by Pol III and AHR increases its transcription with exchange of Pol III by Pol II. (A) IVT using extracts of Hepa-1 cells transiently transfected with empty vector, AHR, or SLUG. Wild-type B1-X35S and mutants B1-Xmut35S and B1-X35Smut were used as templates. (B) B1- X35S expression analyzed by qRT–PCR using total RNA from Hepa-1 cells after transfection with empty vector or AHR. Concentrations of 20 mg/mL a-amanitin (a-AMA) or 5 mg/mL actinomycin-D were added where indicated. (C ) The effect of AHR on Pol II binding to wild-type B1-X35S in Hepa-1 cells was analyzed by qChIP. (D) IVT of B1-X35S with extracts of Hepa-1 cells transfected with empty vector or AHR. A total of 10 mM tagetitoxin (TGT) or 3 mg/ mL a-AMA were included where indicated. Percentages indicate the inhibition caused by each chemical relative to transcription in corresponding vehicle- treated reactions. (E) IVT as indicated in D using the mutant B1-Xmut35S. (F) IVT using wild-type B1-X35S and extracts of immortalized fibroblasts from AHR-null mice (T-FGM AhRÀ/À). (C ) Average data from the five B1-X35S containing genes Dad1, Lpp, Cabin1, Tbc1d1, and Rtl1. Data are shown as mean 6SD.

CTCF to insulator sequences correlates with an increase in het- ance in H3K9me3 content was calculated as the Manhattan dis- erochromatin marks such as H3K27me3 (Han et al. 2008; Li et al. tance in a 500-bp region downstream of B1-X35S using 700 2008). We therefore asked whether CTCF binds B1-X35S in vivo. common instances for ES and MEF (Fig. 7B). It was found that the qChIP showed that CTCF binds B1-X35S and that such binding is ES/MEF distance for H3K9me3 markedly increased downstream of significantly enhanced by AHR expression, but only marginally by B1-X35S (B1 to +500). These results agree with our experimental SLUG expression (Fig. 6D). Consistently, with the accumulation of data (see Figs. 6B,C, 7A) and suggest a role for B1-X35S in estab- H3K27me3, coexpression of AHR+SLUG additively increases CTCF lishing heterochromatic regions that vary with cell type, perhaps binding to B1-X35S. CTCF is activated by poly-ADP-ribose poly- depending on the differentiation status. In contrast, heterochro- merase (PARP1)-dependent parylation (Witcher and Emerson matic H3K9me3 regions were not differentially established by 2009) and, accordingly, PARP1 binding to B1-X35S follows a pat- murine ID SINE elements (Fig. 7B) in ES vs. MEF, indicating that tern close to that of CTCF (Fig. 6D). The PARP1 inhibitor 3-amino SINE elements differ in their ability to modulate chromatin com- benzamide (3-ABA) (Witcher and Emerson 2009) significantly re- paction and probably to insulate gene expression. Indeed, EBA duces CTCF binding to B1-X35S in vivo, suggesting that parylation assays reveal that insulation for an ID element is significantly facilitates the interaction (Fig. 6D). On the contrary, histone H3, lower than that to B1-X35S (Supplemental Fig. 5). a nucleosomal protein not expected to interact with CTCF nor We sought to determine whether the presence of B1-X35S PARP1, binds B1-X35S in an AHR- and SLUG-independent manner between adjacent genes induces divergent patterns of gene ex- (Fig. 3C). Therefore, the insulator activity of B1-X35S may in- pression. We configured three gene groups (700 gene pairs for each volve an increase in heterochromatin content downstream of condition) in which: (1) two adjacent genes have a B1-X35S in the transposon, perhaps through recruitment of parylated CTCF. between; (2) two genes are adjacent without a B1-X35S; or (3) the It has been suggested that insulators and SINEs have a role in two genes are randomly located. The median difference in ex- chromatin compartmentalization. Based on the accumulation of pression for the two genes in each of the 700 pairs for each con- heterochromatic marks downstream of B1-X35S, we performed dition (e.g., gene–B1-gene, or gene–gene, or random gene pair) was a comparative epigenomic analysis (from NCBI GEO repository obtained for 150 tissues of the mouse gene expression atlas (Fig. GSE12241 (Mikkelsen et al. 2007) for H3K9me3 in mouse embry- 7C). Manhattan distance analysis revealed that the presence of onic stem (ES) cells vs. embryonic fibroblasts (MEF). We focused on B1-X35S between contiguous genes significantly increases their a À500 to +500 region flanking B1-X35S in 75 instances. H3K9me3 median difference in expression (black), as compared with adjacent is markedly enriched downstream of B1-X35S in ES cells, but not in genes lacking any SINE (blue) or to randomly located gene pairs MEFs (Fig. 7A), suggesting a cell type-dependent function in set- (red). The use of the Mahalanobis algorithm gave a similar result, ting heterochromatin domains. To address this further, the vari- indicating that B1-X35S accentuates the difference in expression

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start site), further upstream (between À10 kb and À50 kb), or in- ternally within the locus. These three gene sets (available upon request) were subjected to gene expression analysis with the Ex- plain program (BioBase; http://biobase-international.com/pages/ index.php?id=286) to extract transcriptional networks that share a common subset of genes. Signaling networks were identified for the regulators of survival, proliferation, and apoptosis AKT and p62DOK1 (Supplemental Fig. 6) for the modulators of the stress response, apoptosis, and angiogenesis SGK-1 and JNK-1 (Supple- mental Fig. 7) and for the essential regulator of mitochondrial apoptosis caspase 6 (Supplemental Fig. 8). The functional relevance of B1-X35S in the control of gene expression was also estimated by gene-ontology analysis. We found that the occurrence of B1-X35S at different genomic loca- tions varies according to functional categories and is markedly under-represented in the promoters of genes involved in neuro- logical disorders, sensory perception, and G protein-coupled re- ceptors. Conversely, it is over-represented in the promoters of genes associated with ubiquitin ligase activity and ATP binding (Supplemental Table 3).

Discussion

A major finding of this work is that the recently identified SINE B1- X35S retrotransposon has potent intrinsic insulator activity in vitro and in vivo, and that this function can be enhanced by its interaction with the transcription factors AHR and SLUG. The in- volvement of these proteins in controlling insulation was un- expected and could have considerable biological significance. AHR is a key regulator of xenobiotic-induced carcinogenesis (Shimizu et al. 2000; Marlowe and Puga 2005) and has a role in cell proliferation (Barouki et al. 2007), immune T cell differentiation (Quintana et al. 2008), migration (Diry et al. 2006; Carvajal-Gonzalez et al. 2009), and angiogenesis (Roman et al. 2009). SLUG and SNAIL are im- portant in promoting migration during development and disease by Figure 5. The same strand of B1-X35S is transcribed by Pol III and Pol inducing EMT, the epithelial-to-mesenchymal transition (Thiery II. (A) IVT of B1-X35S analyzed using direction-specific sense and antisense et al. 2009). Although SLUG and SNAIL are closely related and with extracts of Hepa-1 cells treated with or without 3 mg/mL a-AMA. recognize the same consensus sequence, SNAIL did not stimulate A negative qPCR control was performed in the absence of RT template (ÀRT). (B) Binding of Pol III and Pol II on B1-X35S was determined by re- insulation despite its efficient binding to B1-X35S, suggesting dif- ChIP. First ChIP used anti-Pol III antibody, and the resulting DNA was ferences in the regulatory interactions that modulate insulation. immunoprecipitated again with anti-Pol II or anti-TFIIIC (positive control) AHR and SLUG have an additive effect on B1-X35S insulation, antibodies. Re-ChIP for GAPDH was used as negative control. Data are which might be relevant to pathological conditions such as met- presented as percentage of input from the first ChIP. (C ) Pol III binding in vitro to B1-X35S A+B mutant was determined by DNA affinity assays with astatic cancer, given the migration-promoting activity of AHR and anti-Pol III RPC32 antibody. (D) EBA performed as for Figure 1A using wild- SLUG in mesenchymal cells. Consistent with the insulator activity, type B1-X35S and B1-X35S A+B mutant with or without AHR transfection. AHR and SLUG cooperate to repress transcription of B1-X35S- (B) Average data from five B1-X35S containing genes (Dad1, Lpp, Cabin1, containing genes (Roman et al. 2008), possibly because AHR pro- Tbc1d1,andRtl1). Data are shown as mean 6SD. motes SLUG binding to B1-X35S. As for the B2 SINE at the growth hormone locus (Lunyak et al. 2007), B1-X35S-dependent insulation is associated with tran- between contiguous genes (Fig. 7D). While the presence between scription across the transposon. However, two main differences adjacent genes of the binding site for CTCF also increases their were found with respect to the B2 case: First, Pol III and Pol II difference in expression, the insertion of ID SINEs does not have transcribe B1-X35S from the same strand in the downstream di- a significant effect (Fig. 7D). Thus, these data suggest that two rection and, second, AHR increases B1-X35S transcription (which contiguous genes separated by a B1-X35S element acquire different is otherwise mainly dependent on Pol III) by an exchange mech- levels of expression, probably sustained by heterochromatin marks anism that recruits Pol II and releases Pol III from the SINE. The such as H3K9me3 at one side of the SINE. AHR-dependent increase in B1-X35S transcription by Pol II is as- We postulated that B1-X35S SINEs could have been selected sociated with enhanced insulator activity, indicating that SINE and fixed at certain genomic locations due to their intrinsic ca- transcription correlates with the degree of insulation. Recruitment pacities to attract AHR and SLUG and to recruit epigenetic modi- of Pol II to B1-X35S requires the A and B box Pol III promoter fications. To address this, we split the 14,000 known locations of elements and mutation of these abolishes the inducing effect of B1-X35S into three categories, depending on whether this SINE is AHR on B1-X35S insulation. Pol III and its associated factors located at promoter regions (up to À10 kb from the transcription bound to the A and B boxes may establish epigenetic marks that

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Figure 6. B1-X35S establishes a heterochromatic epigenetic mark that responds to AHR. The region from 1000 bp upstream (À1000) to 1500 bp downstream (+1500) from B1-X35S was analyzed for chromatin marks by qChIP in Hepa-1 cells with or without transfection of AHR+SLUG. (A) Location of B1-X35S in the promoter region of Dad1, Lpp,andCabin1 and of the PCR fragments analyzed is indicated. (B) H3K9me3 status. (C ) H3K27me3 status. Data were normalized to the amount of H3 immunoprecipitated. (D) In vivo binding of CTCF and PARP1 to B1-X35S was analyzed by qChIP in Hepa-1 cells with or without transfection of AHR+SLUG. PARP1 inhibitor 3-amino benzamide (3-ABA, 5 mM) was added for 24 h where indicated. Average results for genes Dad1, Lpp, Cabin1, Tbc1d1,andRtl1 are shown. Experiments were performed in Hepa-1 cells and data are shown as mean 6SD.

allow subsequent recruitment of Pol II (Kenneth et al. 2007; Barski was found downstream of B1-X35S, and this pattern was increased et al. 2010; Moqtaderi et al. 2010; Raha et al. 2010). It is striking by AHR+SLUG binding, suggesting that these transcription factors that AHR induces the replacement of Pol III by Pol II at B1-X35S. determine the recruitment of histone methyltransferases to spe- We are not aware of a precedent for this behavior. cific regions of the genome adjacent to the transposon. This is It seems paradoxical that replacement of Pol III by Pol II consistent with previous studies showing that nucleosomes at increases transcription of B1-X35S. Pol III transcription is generally transposons are often enriched in heterochromatin marks like very efficient due to a high rate of re-initiation, in which the H3K9me3 (Martens et al. 2005), and that a reporter gene intro- polymerase is recycled to the start site without dissociating from duced in the genome of ES cells by LINE-1 retrotransposons is the template DNA (Dieci and Sentenac 2003). For example, tRNA rapidly repressed by chromatin modifications at the insertion site, and 7SL RNA genes utilize this mechanism to maintain rapid while ES cell differentiation attenuates the silencing induced by transcription (Ferrari et al. 2004). Indeed, the four 7SL gene copies LINE-1 (Garcia-Perez et al. 2010). The ability of B1-X35S to provide in humans are sufficient to maintain transcript levels of ;106 a potential heterochromatin barrier could have substantial func- copies per cell (Liu et al. 1994). However, we found that Pol III tional impact across the genome. Because it displays markedly occupancy is much lower on B1 SINEs than on Pol III templates different H3K9me3 profiles in ES cells and fibroblasts, the insulator that are actively transcribed, such as 7SL or tRNA genes (Supple- activity of B1-X35S may regulate gene expression during devel- mental Fig. 9). This reflects a defect subsequent to TFIIIC binding, opment. Location of a B1-X35S SINE between two adjacent genes as the ratio of Pol III/TFIIIC ChIP signals is significantly lower on correlates with their differential expression in most adult mouse B1 when compared with 7SL (P = 0.002 by Student t-test). There- tissues, supporting the involvement of B1-X35S in establishing fore,ourdatasuggestthatPolIIIrecruitmentisrelativelyin- distinct expression domains for flanking genes, possibly by mod- efficient at B1 SINEs, which might explain why replacement of ulating chromatin. DNA methylation of B1-X35S could also play Pol III with a Pol II-dependent system can result in increased ex- a role in insulation, since changes in methylation of CpG di- pression. Replacement may occur by AHR interfering with TFIIIC nucleotides flanking human Alu repeats are implicated in epen- binding to the A and B boxes. dymomas (Xie et al. 2010). In addition, we have shown that the Transposable elements and insulators are closely linked to the insulator activity of B1-X35S involves binding of parylated CTCF, epigenetic regulation of the genome (Slotkin and Martienssen an established insulator-binding protein involved in loop forma- 2007; Bushey et al. 2008; Tomilin 2008). B1-X35S shares with other tion and the recruitment of repressive complexes to chromatin insulators, such as the Su(Hw), Mod(mdg) (Gerasimova and Corces (Han et al. 2008; Li et al. 2008). AHR-mediated binding of CTCF/ 1998), and the growth hormone B2 SINE (Lunyak et al. 2007), the PARP1 to B1-X35S-containing genes could contribute to their characteristic of heterochromatin marks capable of regulating cis- transcriptional repression (Roman et al. 2008). A similar mecha- acting genes. An elevation of heterochromatin-associated marks nism has been proposed for the transcriptional regulation of

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Figure 7. Distinct patterns of heterochromatin and gene expression upstream and downstream from B1-X35S. (A) H3K9me3 analyzed in mouse ES cells and MEFs at 75 B1-X35S loci between À500 bp (upstream) and +500 bp (downstream) from the SINE (box). ChIP-seq data were taken from NCBI GEO repository (GSE12241) (Mikkelsen et al. 2007). Red indicates higher and green lower H3K9me3 levels. (B) H3K9me3 levels downstream of B1-X35S and SINE ID elements (element to +500) in 700 instances common to ES and MEF cells were analyzed. Data are presented as the ES/MEF Manhattan distance ratio. A larger distance indicates higher H3K9me3 content in ES cells. Random sequences were also analyzed as negative controls. (C ) The effect of B1- X35S on the differential expression of flanking genes was analyzed in the following groups (700 gene pairs for each): gene pairs having a B1-X35S between them (gene-B1-gene); gene pairs lacking B1-X35S (gene-gene); randomly located gene pairs (random). The difference in expression for each pair in every group was analyzed in 150 tissues of the mouse gene expression atlas and the values obtained represented as the median Manhattan distance in expression. Higher values represent larger differences between adjacent genes. The mean distance for each condition is indicated by a horizontal dashed line. (D) The gene–B1X35S-gene, gene–ID-gene, and gene–gene groups were analyzed using the Mahalanobis algorithm. The impact of the binding sequence for CTCF on the expression of contiguous genes (gene–CTCF-gene) was also determined using ChIP-seq data from the NCBI GEO repository (GSE11431) (Chen et al. 2008).

p16ink4a expression in human tumor cells (Witcher and Emerson homeostasis. B1-X35S seems to be restricted to the mouse genus 2009). (Roman et al. 2008) and caution should be taken in extrapolating The clustering of genes containing B1-X35S elements the insulator activity of AHR and SLUG to other species, since the revealed the existence of transcriptional networks controlling the profile of occupancy of binding sites for transcription factors (e.g., cell cycle, proliferation, apoptosis, and cancer. Both AHR and OCT-8 and NANOG but not CTCF) in transposable elements vary SLUG have been implicated in cell growth and proliferation, ap- widely between mouse and human ES cells (Kunarso et al. 2010). optosis, and migration, thus supporting their functional inter- Interestingly, B1-X35S has insulator activity in both fish and hu- action with the AKT-1, p62DOK1, SGK-1, JNK-1, and caspase 6 man cells. While maintaining this property in different experi- pathways. Indeed, AHR seems to regulate dioxin-induced migra- mental systems helps establish the role of B1-X35S as a genome- tion through the JNK pathway in human breast tumor cells (Diry wide insulator, our conclusions are not necessarily applicable to et al. 2006), while AHR deficiency results in lower AKTactivation in repetitive elements and expression patterns of AHR and SLUG in mouse fibroblasts (Mulero-Navarro et al. 2005). Given the inter- other species. Certainly, a different repetitive element such as the action between these transcriptional networks and AHR and SLUG, murine SINE ID has significantly lower insulator activity than we envisage that most B1-X35S-linked genes may normally be re- B1-X35S. pressed by these factors, perhaps because increased SINE transcrip- In summary, we propose a model for the insulator activity of tion by Pol II enhances the insulator and chromatin barrier activi- B1-X35S SINE retrotransposons involving AHR and SLUG, Pol III, ties. In the absence of AHR and SLUG, B1-X35S transcription by and Pol II, and the epigenetic regulation of chromatin (Fig. 8). Pol II would be reduced and its insulator activity diminished, Under basal conditions, Pol III is predominant at B1-X35S and eventually resulting in gene derepression and in altered cellular provides its intrinsic insulator activity. Elevated AHR expression

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manufacturer’s recommendations. Protein expression and the anal- ysis of protein–DNA interactions in vitro were by Western blotting as indicated (Roman et al. 2008).

Quantitative Chromatin Immunoprecipitation (qChIP) qChIP was performed as described (Roman et al. 2008; Carvajal- Gonzalez et al. 2009). Positive controls were performed using input DNAs, whereas negative controls included binding reactions in the absence of specific antibodies. To quantify changes in protein binding to DNA, all amplifications were performed by real-time PCR (qPCR) using IQ-SYBR Green in an iCycler equipment (Roman Figure 8. Proposed model for the regulation of the B1-X35S insulator. et al. 2009). Data are presented as percentage of DNA input in the Arrows indicate entry and exit of proteins. Small gray circles indicate antibody-containing immunoprecipitates minus the percentage of H3K27me3 and small gray triangles H3K9me3. Binding sites for AHR, DNA input in the corresponding negative controls. Bars represent SLUG, SNAIL, and TFIIIC are indicated. Potential involvement of parylated the average amplification of B1-X35S elements located in the up- CTCF is also indicated. stream promoter region of the genes Dad1, Lpp, Cabin1, Tbc1d1, and Rtl1. Individual data for each gene and experimental condi- triggers recruitment of Pol II and the displacement of Pol III from tion are provided in Supplemental Table 1. In experiments map- the SINE, raising B1-X35S transcription and enhancing insulation. ping histone H3 modifications upstream and downstream of B1- AHR increases heterochromatin marks downstream from B1-X35S, X35S only Dad1, Lpp, and Cabin1 were analyzed. Individual data which establish a chromatin barrier that contributes to the in- for each gene and experimental condition are provided in Sup- hibition of neighboring genes. The insulator activity of B1-X35S plemental Table 2. also seems to involve parylated CTCF. Although further studies will be required to validate this model at specific genomic loci, it has Sequential ChIP (re-ChIP) potential relevance genome wide. It is now apparent that AHR and SLUG have unexpected functions beyond their classical role in Sequential ChIP was used to analyze simultaneous binding of two transcription. These molecular events may have an impact on proteins to a common DNA region. Re-ChIP was by q-PCR fol- development and cell physiology in health and disease. lowing a published protocol (Metivier et al. 2008) using primers for B1-X35S elements located in promoters of the genes listed in Supplemental Table 4. GAPDH was immunoprecipitated as nega- Methods tive control for the second ChIP. Results are presented as percent- age of DNA in the primary immunoprecipitates. Reagents, antibodies, plasmids, and cell lines 3-amino benzamide, a-amanitin, and anti-actin antibody were Enhancer blocking assay (EBA) purchased from Sigma. Tagetitoxin was from Epicentre Biotech- nologies. Protein A/G Plus Agarose, anti-SLUG (sc-10437), anti- Enhancer blocking assay to address the insulator activity of wild- Pol II (N-20), and anti-Pol III (RPC32) antibodies were from Santa type and mutant B1-X35S used the pELuc plasmid previously de- Cruz Biotechnology. Antibodies to histone H3 (H3), H3K9me3, scribed (Lunyak et al. 2007). B1-X35S elements were cloned be- H3K27me3, and H3K4me3 were purchased from Diagenode. Other tween the CMV enhancer and the promoter (XhoI) or upstream of commercial antibodies used include anti-AHR MA1-514 (Affinity the CMV enhancer (PstI). EBA was performed by transfecting the Bioreagents), anti-GAPDH (Cell Signaling), and anti-PARP1 (Roche). constructs into human embryonic kidney HEK 293 cells as re- Polyclonal antisera against Pol III (RPC155), TFIIIB (Brf1), and ported (Lunyak et al. 2007). Data are presented as fold-enhancer TFIIIC (102kD and 110kD) have been described (Kenneth et al. blocking activity normalized to the value achieved by the basal 2008). Anti-SNAIL and anti-CTCF antibodies were kindly provided pELuc vector. The 1.2-kb chicken 59HS4 beta-globin insulator ele- by Dr. Antonio Garcia de Herreros and Dr. Felix Recillas-Targa, ment was used as positive control. Similarly, the internal II/III respectively. boxes from the chicken 59HS4 beta-globin insulator element, both B1-X35S-containing plasmids were made by sequential PCR. wild type and mutated, were used as positive and negative con- A first amplification reaction was performed using long over- trols, respectively (Recillas-Targa et al. 1999). See schemes in lapping primers (B1-X35 and B1-35S and their Xmut, Smut, and Figure 1. ABmutants indicated in Supplemental Table 5 and Supplemental Fig. 1A) covering the full-length B1-X35S retrotransposon. The In vivo insulator activity in zebrafish products obtained were then used as templates in a second PCR reaction performed using a pair of common flanking primers The insulator activity of wild-type and mutant B1-X35S was also (Flank-F and Flank-R in Supplemental Table 5). The resulting analyzed in vivo by microinjection of the constructs drawn in products were A/T cloned into pGEM-T for further use (Promega). Figure 2 into one-cell zebrafish embryos as described (Bessa et al. Mouse Hepa-1 cells (Interlab Cell Line Collection ATL98016) 2009). Briefly, B1-X35S elements were cloned in a GFP reporter that were cultured in a-MEM (Invitrogen) as indicated (Roman et al. allows quantification of enhancer-blocking activity by measuring 2008). Immortalized fibroblasts from AhR+/+ and AhRÀ/À mice were the ratio of fluorescence in somites (driven by a cardiac actin pro- used as previously (Carvajal-Gonzalez et al. 2009). moter) vs. fluorescence in the central nervous system (regulated by the Z48 midbrain neuronal enhancer). Individual specimens were analyzed 36 h after microinjection. At least 20 zebrafish were Transfection and Western blotting photographed and quantified for each condition in two indepen- Hepa-1 cells were transiently transfected in a MicroPorator (Digi- dent experiments. LaserPix (Bio-Rad) image analysis software was talBio) using a single pulse of 1400 V for 30 msec, following the used for quantification.

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AHR and SLUG trigger insulation by B1 SINE repeats

Epigenomic bioinformatic analysis (BIO2009-1297); and by Grants to J.L.G.-S. from the Spanish and Andalusian Governments (BFU2007-60042/BMC, Petri Genomic data for H3K9me3 in embryonic stem cells (ES) and PET2007_0158, Proyecto de Excelencia CVI-3488 and CSD2007- mouse embryonic fibroblasts (MEF) were retrieved from NCBI GEO 00008). A.C.R. and F.J.G.-R. were supported by the MICINN, and repository (GSE12241) (Mikkelsen et al. 2007). Genomic location E.M. was supported by CIBERER (ISCIII). All Spanish funding is co- of all B1-X35S elements was obtained using a modified version sponsored by the European Union FEDER program. J.V.-A. and of our previously described script (Roman et al. 2008). We selected R.J.W. were funded by the Wellcome Trust and Cancer Research B1-X35S elements having significant differences in H3K9me3 in a UK. The generous gifts of Dr. Antonio Garcia de Herreros (Institut À500 bp to +500 bp window from the transposon position (paired Municipal d’Investigacio´ Me`dica, Barcelona, Spain) (anti-SNAIL t-test <0.001 between ES and MEF). Hierarchical clustering (com- antibody and pcDNA-SNAIL), Dr. Felix Recillas-Targa (Instituto de plete linkage) of ES cells and heat maps of ES and MEF cells were Fisiologı´a Celular, UNAM, Mexico) (anti-CTCF antibody), and Dr. done using R statistical package. The effect of B1-X35S on the ex- Amparo Cano (Instituto de Investigaciones Biome´dicas, Madrid, pression of adjacent genes was analyzed in 150 mouse tissues from Spain) (pcDNA-SLUG) are greatly appreciated. The helpful com- the mouse gene expression atlas (Su et al. 2002). The Manhattan ments and suggestions of Drs. R. Kamakaka, R. Ohlsson, and J.M. distance algorithm was used to quantify differences in H3K9me3 Merino are acknowledged. content upstream and downstream of B1-X35S in ES and MEF cells and to determine changes in the expression of pairs of genes lo- cated at both sides of the transposon. In some cases, the Mahala- References nobis distance was also calculated to address changes in gene ex- pression due to the presence of either B1-X35S or CTCF. Genomic Aravin AA, Sachidanandam R, Girard A, Fejes-Toth K, Hannon GJ. 2007. data for the presence of CTCF binding sites was retrieved from the Developmentally regulated piRNA clusters implicate MILI in transposon NCBI GEO repository (GSE11431) (Chen et al. 2008). control. Science 316: 744–747. Barouki R, Coumoul X, Fernandez-Salguero PM. 2007. The aryl hydrocarbon receptor, more than a xenobiotic-interacting protein. FEBS Lett 581: 3608–3615. DNA-binding affinity assays Barski A, Chepelev I, Liko D, Cuddapah S, Fleming AB, Birch J, Cui K, White Protein binding to B1-X35S in vitro was analyzed by DNA-binding RJ, Zhao K. 2010. Pol II and its associated epigenetic marks are present at Pol III-transcribed noncoding RNA genes. Nat Struct Mol Biol 17: 629– affinity as described (Roman et al. 2008). The primers indicated in 634. Supplemental Table 5 were biotin labeled and used to amplify by Bell AC, West AG, Felsenfeld G. 2001. Insulators and boundaries: Versatile PCR the wild-type and mutant forms of B1-X35S as above. Binding regulatory elements in the eukaryotic. Science 291: 447–450. reactions used streptavidin mMACS Factor Finder kit (Miltenyi Bessa J, Tena JJ, de la Calle-Mustienes E, Fernandez-Minan A, Naranjo S, Fernandez A, Montoliu L, Akalin A, Lenhard B, Casares F, et al. 2009. Biotec) following the manufacturer’s instructions. Proteins were Zebrafish enhancer detection (ZED) vector: A new tool to facilitate detected by Western blotting. transgenesis and the functional analysis of cis-regulatory regions in zebrafish. Dev Dyn 238: 2409–2417. Bourque G, Leong B, Vega VB, Chen X, Lee YL, Srinivasan KG, Chew JL, In vitro transcription (IVT) Ruan Y, Wei CL, Ng HH, et al. 2008. Evolution of the mammalian transcription factor binding repertoire via transposable elements. IVT was performed as previously described (Francis et al. 2005) Genome Res 18: 1752–1762. with some modifications. Wild-type or mutant pGEM-T-B1-X35S Bushey AM, Dorman ER, Corces VG. 2008. Chromatin insulators: constructs were used as templates. Hepa-1 protein extracts were Regulatory mechanisms and epigenetic inheritance. Mol Cell 32: 1–9. Carvajal-Gonzalez JM, Mulero-Navarro S, Roman AC, Sauzeau V,Merino JM, pre-mixed with vehicle or with specific inhibitors of Pol II (3 mg/ Bustelo XR, Fernandez-Salguero PM. 2009. The dioxin receptor regulates mL a-amanitin, a-AMA) (Rollins et al. 2007) or Pol III (10 mM the constitutive expression of the vav3 proto-oncogene and modulates tagetitoxin, TGT) (Perl et al. 2000). Treated extracts were then cell shape and adhesion. Mol Biol Cell 20: 1715–1727. mixed with templates and IVT started by addition of 0.4 mM Chen X, Xu H, Yuan P,Fang F, Huss M, Vega VB, Wong E, Orlov YL, Zhang W, Jiang J, et al. 2008. Integration of external signaling pathways with the rNTPs. Reactions proceeded for 60 min at 30°C. After treatment core transcriptional network in embryonic stem cells. Cell 133: 1106– with DNase I for 30 min, reactions were stopped and precipitated 1117. with ethanol. Purified RNA was subjected to RT–qPCR as described Dieci G, Sentenac A. 2003. Detours and shortcuts to transcription (Roman et al. 2009) using the primers in Supplemental Table 5. For reinitiation. Trends Biochem Sci 28: 202–209. Diry M, Tomkiewicz C, Koehle C, Coumoul X, Bock KW, Barouki R, Transy strand-specific IVT, reverse transcription used sense (B1-Flank-F) or C. 2006. Activation of the dioxin/aryl hydrocarbon receptor (AhR) antisense (B1-X35S-R)-specific primers. Negative controls included modulates cell plasticity through a JNK-dependent mechanism. reactions in the absence of protein, DNA, or RT product. Oncogene 25: 5570–5574. Ferrari R, Rivetti C, Acker J, Dieci G. 2004. Distinct roles of transcription factors TFIIIB and TFIIIC in RNA polymerase III transcription Statistical analyses reinitiation. Proc Natl Acad Sci 101: 13442–13447. Ferrigno O, Virolle T, Djabari Z, Ortonne JP, White RJ, Aberdam D. 2001. Experiments were performed three times with at least two repli- Transposable B2 SINE elements can provide mobile RNA polymerase II cates. Data are shown as mean 6SD. Statistical tests (unpaired and promoters. Nat Genet 28: 77–81. paired t-tests) were applied using the R statistical package. Wil- Francis J, Chakrabarti SK, Garmey JC, Mirmira RG. 2005. Pdx-1 links histone H3-Lys-4 methylation to RNA polymerase II elongation during coxon test was applied for median-based analyses. activation of insulin transcription. J Biol Chem 280: 36244–36253. Furness SG, Lees MJ, Whitelaw ML. 2007. The dioxin (aryl hydrocarbon) receptor as a model for adaptive responses of bHLH/PAS transcription Acknowledgments factors. FEBS Lett 581: 3616–3625. This work was supported by grants to P.M.F-S. from the Spanish Garcia-Perez JL, Morell M, Scheys JO, Kulpa DA, Morell S, Carter CC, Hammer GD, Collins KL, O’Shea KS, Menendez P, et al. 2010. Epigenetic Ministry of Science and Innovation (MICINN) (SAF2008-00462), silencing of engineered L1 retrotransposition events in human the Junta de Extremadura (GRU09001), and the Red Tema´tica de embryonic carcinoma cells. Nature 466: 769–773. Investigacio´n Cooperativa en Ca´ncer (RTICC) [RD06/0020/1016, Gerasimova TI, Corces VG. 1998. Polycomb and trithorax group proteins Cell Fondo de Investigaciones Sanitarias (FIS), Carlos III Institute, mediate the function of a chromatin insulator. 92: 511–521. Gerasimova TI, Corces VG. 2001. Chromatin insulators and boundaries: Spanish Ministry of Health]; by Grants to L.M. from the Spanish Effects on transcription and nuclear organization. Annu Rev Genet 35: Ministry of Education and Science (BFU2006-12185) and MICINN 193–208.

Genome Research 431 www.genome.org Downloaded from genome.cshlp.org on August 8, 2011 - Published by Cold Spring Harbor Laboratory Press

Roma´ netal.

Giraldo P, Rival-Gervier S, Houdebine LM, Montoliu L. 2003. The potential chromatin insulator protein CTCF map to DNA methylation-free benefits of insulators on heterologous constructs in transgenic animals. domains genome-wide. Genome Res 14: 1594–1602. Transgenic Res 12: 751–755. Mulero-Navarro S, Pozo-Guisado E, Perez-Mancera PA, Alvarez-Barrientos A, Gomez-Duran A, Carvajal-Gonzalez JM, Mulero-Navarro S, Santiago-Josefat Catalina-Fernandez I, Hernandez-Nieto E, Saenz-Santamaria J, Martinez B, Puga A, Fernandez-Salguero PM. 2009. Fitting a xenobiotic receptor N, Rojas JM, Sanchez-Garcia I, et al. 2005. Immortalized mouse into cell homeostasis: How the dioxin receptor interacts with TGFbeta mammary fibroblasts lacking dioxin receptor have impaired signaling. Biochem Pharmacol 77: 700–712. tumorigenicity in a subcutaneous mouse xenograft model. J Biol Chem Han L, Lee DH, Szabo PE. 2008. CTCF is the master organizer of domain- 280: 28731–28741. wide allele-specific chromatin at the H19/Igf2 imprinted region. Mol Cell Ohlsson R, Renkawitz R, Lobanenkov V. 2001. CTCF is a uniquely versatile Biol 28: 1124–1135. transcription regulator linked to epigenetics and disease. Trends Genet Katoh M. 2005. Comparative genomics on SNAI1, SNAI2, and SNAI3 17: 520–527. orthologs. Oncol Rep 14: 1083–1086. Perl A, Colombo E, Samoilova E, Butler MC, Banki K. 2000. Human Kenneth NS, Ramsbottom BA, Gomez-Roman N, Marshall L, Cole PA, White transaldolase-associated repetitive elements are transcribed by RNA RJ. 2007. TRRAP and GCN5 are used by c-Myc to activate RNA polymerase III. J Biol Chem 275: 7261–7272. polymerase III transcription. Proc Natl Acad Sci 104: 14917–14922. Quintana FJ, Basso AS, Iglesias AH, Korn T, Farez MF, Bettelli E, Caccamo M, Kenneth NS, Marshall L, White RJ. 2008. Recruitment of RNA polymerase III Oukka M, Weiner HL. 2008. Control of T(reg) and T(H)17 cell in vivo. Nucleic Acids Res 36: 3757–3764. differentiation by the aryl hydrocarbon receptor. Nature 453: 65–71. Kunarso G, Chia NY, Jeyakani J, Hwang C, Lu X, Chan YS, Ng HH, Bourque Raha D, Wang Z, Moqtaderi Z, Wu L, Zhong G, Gerstein M, Struhl K, Snyder G. 2010. Transposable elements have rewired the core regulatory M. 2010. Close association of RNA polymerase II and many network of human embryonic stem cells. Nat Genet 42: 631–634. transcription factors with Pol III genes. Proc Natl Acad Sci 107: Kuwabara T, Hsieh J, Muotri A, Yeo G, Warashina M, Lie DC, Moore L, 3639–3644. Nakashima K, Asashima M, Gage FH. 2009. Wnt-mediated activation Recillas-Targa F, Bell AC, Felsenfeld G. 1999. Positional enhancer-blocking of NeuroD1 and retro-elements during adult neurogenesis. Nat Neurosci activity of the chicken beta-globin insulator in transiently transfected 12: 1097–1105. cells. Proc Natl Acad Sci 96: 14354–14359. Li T, Hu JF, Qiu X, Ling J, Chen H, Wang S, Hou A, Vu TH, Hoffman AR. 2008. Recillas-Targa F, Valadez-Graham V, Farrell CM. 2004. Prospects and CTCF regulates allelic expression of Igf2 by orchestrating a promoter- implications of using chromatin insulators in gene therapy and polycomb repressive complex 2 intrachromosomal loop. Mol Cell Biol transgenesis. Bioessays 26: 796–807. 28: 6473–6482. Rollins J, Veras I, Cabarcas S, Willis I, Schramm L. 2007. Human Maf1 Lippman Z, Gendrel AV, Black M, Vaughn MW, Dedhia N, McCombie WR, negatively regulates RNA polymerase III transcription via the TFIIB Lavine K, Mittal V, May B, Kasschau KD, et al. 2004. Role of transposable family members Brf1 and Brf2. Int J Biol Sci 3: 292–302. elements in heterochromatin and epigenetic control. Nature 430: 471–476. Roman AC, Benitez DA, Carvajal-Gonzalez JM, Fernandez-Salguero PM. Liu WM, Maraia RJ, Rubin CM, Schmid CW. 1994. Alu transcripts: 2008. Genome-wide B1 retrotransposon binds the transcription factors Cytoplasmic localisation and regulation by DNA methylation. Nucleic dioxin receptor and Slug and regulates gene expression in vivo. Proc Natl Acids Res 22: 1087–1095. Acad Sci 105: 1632–1637. Lunyak VV. 2008. Boundaries. Boundaries...Boundaries??? Curr Opin Cell Roman AC, Carvajal-Gonzalez JM, Rico-Leo EM, Fernandez-Salguero PM. Biol 20: 281–287. 2009. Dioxin receptor deficiency impairs angiogenesis by a mechanism Lunyak VV,Prefontaine GG, Nunez E, Cramer T, Ju BG, Ohgi KA, Hutt K, Roy involving VEGF-A depletion in the endothelium and transforming R, Garcia-Diaz A, Zhu X, et al. 2007. Developmentally regulated growth factor-beta overexpression in the stroma. J Biol Chem 284: activation of a SINE B2 repeat as a domain boundary in organogenesis. 25135–25148. Science 317: 248–251. Shimizu Y, Nakatsuru Y, Ichinose M, Takahashi Y, Kume H, Mimura J, Marlowe JL, Puga A. 2005. Aryl hydrocarbon receptor, cell cycle regulation, Fujii-Kuriyama Y, Ishikawa T. 2000. Benzo[a]pyrene carcinogenicity is toxicity, and tumorigenesis. J Cell Biochem 96: 1174–1184. lost in mice lacking the aryl hydrocarbon receptor. Proc Natl Acad Sci Martens JH, O’Sullivan RJ, Braunschweig U, Opravil S, Radolf M, Steinlein P, 97: 779–782. Jenuwein T. 2005. The profile of repeat-associated histone lysine Slotkin RK, Martienssen R. 2007. Transposable elements and the epigenetic methylation states in the mouse epigenome. EMBO J 24: 800–812. regulation of the genome. Nat Rev Genet 8: 272–285. Martinez-Estrada OM, Lettice LA, Essafi A, Guadix JA, Slight J, Velecela V, Su AI, Cooke MP, Ching KA, Hakak Y, Walker JR, Wiltshire T, Orth AP, Vega Hall E, Reichmann J, Devenney PS, Hohenstein P, et al. 2010. Wt1 is RG, Sapinoso LM, Moqrich A, et al. 2002. Large-scale analysis of the required for cardiovascular progenitor cell formation through human and mouse transcriptomes. Proc Natl Acad Sci 99: 4465–4470. transcriptional control of Snail and E-cadherin. Nat Genet 42: 89–93. Thiery JP, Acloque H, Huang RY, Nieto MA. 2009. Epithelial-mesenchymal Mathew LK, Simonich MT, Tanguay RL. 2009. AHR-dependent transitions in development and disease. Cell 139: 871–890. misregulation of Wnt signaling disrupts tissue regeneration. Biochem Tomilin NV. 2008. Regulation of mammalian gene expression by Pharmacol 77: 498–507. retroelements and non-coding tandem repeats. Bioessays 30: 338–348. Metivier R, Gallais R, Tiffoche C, Le Peron C, Jurkowska RZ, Carmouche RP, Valenzuela L, Kamakaka RT. 2006. Chromatin insulators. Annu Rev Genet Ibberson D, Barath P, Demay F, Reid G, et al. 2008. Cyclical DNA 40: 107–138. methylation of a transcriptionally active promoter. Nature 452: 45–50. Wang T, Zeng J, Lowe CB, Sellers RG, Salama SR, Yang M, Burgess SM, Mikkelsen TS, Ku M, Jaffe DB, Issac B, Lieberman E, Giannoukos G, Alvarez P, Brachmann RK, Haussler D. 2007. Species-specific endogenous Brockman W, Kim TK, Koche RP, et al. 2007. Genome-wide maps of retroviruses shape the transcriptional network of the human tumor chromatin state in pluripotent and lineage-committed cells. Nature 448: suppressor protein p53. Proc Natl Acad Sci 104: 18613–18618. 553–560. Witcher M, Emerson BM. 2009. Epigenetic silencing of the p16(INK4a) Molto´ E, Fernandez A, Montoliu L. 2009. Boundaries in vertebrate genomes: tumor suppressor is associated with loss of CTCF binding and Different solutions to adequately insulate gene expression domains. a chromatin boundary. Mol Cell 34: 271–284. Brief Funct Genomics Proteomics 8: 283–296. Xie H, Wang M, Bonaldo Mde F, Rajaram V, Stellpflug W, Smith C, Arndt K, Moqtaderi Z, Wang J, Raha D, White RJ, Snyder M, Weng Z, Struhl K. 2010. Goldman S, Tomita T, Soares MB. 2010. Epigenomic analysis of Alu Genomic binding profiles of functionally distinct RNA polymerase repeats in human ependymomas. Proc Natl Acad Sci 107: 6952–6957. III transcription complexes in human cells. Nat Struct Mol Biol 17: 635–640. Mukhopadhyay R, Yu W, Whitehead J, Xu J, Lezcano M, Pack S, Kanduri C, Kanduri M, Ginjala V, Vostrov A, et al. 2004. The binding sites for the Received June 2, 2010; accepted in revised form December 16, 2010.

432 Genome Research www.genome.org A POL III complex AHR C AHR - + + + A‐BOX XRE 5´ SLUG - - + - GCCGGGCGTGGTGGCGCACGCCTTTAATCCCAGCACTTGGGAGGCAG SNAIL - - - +

5´ hAHR AHR SLUG/SNAIL POL III complex 3´ mAHR E‐box B‐BOX AGGCAGGTGGATTTCTGAGTTCGAGGCCAGCCTGGTCTACAGAGTGAG SLUG

3´ SNAIL A‐RICH 3´ TTCCAGGACAGCCAGGGCATACAGAGAAACCCTGTCTCG -ACTIN 5´ B D 20% Cabin1 Gene B1-X35S -1226 bp Proximal promoter Tbc1d1 48% B1-X35S (<10 kb) -4841 bp 18% Distal Promoter B1-X35S (<50 kb) Dad1 -3178 bp Other B1-X35S Rtl1 14% -120 Kbp Lpp -2460 bp

Supplementary Figure 1. Consensus sequences for DNA binding proteins in the B1-X35S retrotransposon and its prevalence in several genomic locations. (A) Elements for the dioxin binding AhR (XRE), the epithelial-to-mesenquimal regulators SLUG and SNAIL (E-box) and for RNA polymerase III complex (A and B boxes) are indicated. The long blue and red arrows indicate the sequence of the forward and reverse primers used to amplifly by PCR the full length B1-X35S retrotransposon (see Table SII). (B) B1-X35S localization was determined in the proximal promoter (red), distal promoter (green), non-promoter gene regions (blue) and any other location excluding all of the above (purple). (C) Western blot detection of AHR, SLUG and SNAIL in human HEK 293 cells under basal conditions and after transient transfection with the correspondingexpression plasmids. Note that the transfected high affinity AHRb1 mouse protein has a lower molecular weight (̴ 90 kDa) than its human endogenous homolog (̴ 106 kDa). (D) Location of the B1-X35S element in the Cabin1, Dad1, Lpp, Tbc1d1 and Rtl1 gene promoters. Nucleotide positions are refered to the TSS. 5000

4500

4000 /CNS) /CNS)

3500

muscle in in 3000

2500 expression

2000 (GFP (GFP

1500 activity

1000

500 Insulator 0 Empty vector B1-X35S B1-Xmut35Smut

Supplementary Figure 2. Analysis of insulator activity in individual specimens of zebrafish transfected with the indicated constructs. Insulator activity was measured as the ratio of GFP signal driven by a muscle promoter with respect to that modulated by a CNS enhancer. At least 19 animals were quantified for B1-X35S and 34 for B1-Xmut35Smut. Eighteen additional zebra fish specimens were analyzed as negative controls (empty vector without retrotransposon). Higher values indicate lower expression in CNS and thus stronger insulation from the Z48 enhancer. The experiment was repeated twice. SLUG - - + AHR - + + POL III (RPC32)

-ACTIN

Supplementary Figure 3. Western blot detection of RNA POL III complex subunit RPC32 in Hepa I cells under basal conditions and after transient transfection with expression plasmids for AHR and SLUG. Protein levels of -ACTIN were used as loading control. AHR - + POL II

-ACTIN

Supplementary Figure 4. Western blot detection of RNA POL II in Hepa I cells under basal conditions and after transient transfection with expression plasmids for AHR and SLUG. Protein levels of -ACTIN were used as loading control. PstI XhoI Ins Enh Pro EnhIns Pro 50 LUC LUC

40 XhoI PstI

30

20 (relative to pELuc)

Fold Enhancer Blocking 10

0

Supplementary Figure 5. Enhancer blocking assay to analyze the insulator activity of a SINE ID element. The SINE ID was cloned flanking the enhancer and promoter (PstI construct) or between both regulatory sequences (XhoI construct) and its ability to regulate luciferase activity measured in transfected MEK 293 cells. The SINE ID has a much lower insulator activity than B1- X35S. The experiment was repeated twice. Supplementary Figure 6. There were 2555 genes having a B1-X35S element in their promoters (-10 kbp to +1) and Explain identified two significant networks: AKT1 (false discovery rate FDR=0.01) and p62DOK1 (FDR=0.026). AKT1 is known as a thymoma viral proto-oncogene with serine-threonine kinase activity that acts in G protein-coupled receptor (GPCR) pathways regulating cell proliferation, apoptosis and angiogenesis. In humans, AKT1 is aberrantly expressed in ovarian neoplasms, gliomas, and various other cancers. p62DOK1 (docking protein 1) binds ABL and RASGAP and it is involved in cell adhesion to the extracellular matrix, migration and growth. Human p62DOK1 is associated with chronic lymphocytic leukemia. Explain program (BioBase; http://biobase-international.com/pages/index.php?id=286). Supplementary Figure 7. In 4997 genes the B1-X35S element was located within the transcriptional unit, mainly in introns but also in 5’UTR and 3’UTRs. Explain found two main transcriptional networks: SGK-1 (FDR=0.008) and JNK-1 (FDR=0.046) (Supplementary Fig. 6). SGK-1 (serum-glucocorticoid regulated kinase 1) is a protein serine-threonine kinase with anti- apoptotic activity that acts in neurite development. Human SGK1 is associated with hyper- insulinism, type 2 diabetes mellitus, pulmonary fibrosis, and several neoplasms. JNK-1 (c-Jun NH2 terminal protein kinase 1) is a mitogen activated protein kinase that regulates protein phosphorylation, JAK-STAT cascade, apoptosis, caspase activity and angiogenesis. Explain program (BioBase; http://biobase-international.com/pages/index.php?id=286). Supplementary Figure 8. A total of 5325 genes were identified between -10 kbp and -50 kbp encompassing transcriptional networks for CASPASE-6 (FDR=0) and, again, AKT-1 (FDR=0.012), although using other intermediate signaling molecules. Explain program (BioBase; http://biobase-international.com/pages/index.php?id=286). inputs

B1 SINE

tRNA

H19

Supplementary Figure 9. Comparison by ChIP of crosslinking of endogenous TFIIIB, TFIIIC and POL III at B1 SINEs and tRNALeu genes. The Pol II-dependent H19 locus is included as a control. Positive control antibodies recognize acetylated histones H3 (AcH3) and H4 (AcH4), whilst antibody against TFIIA and beads without antibody provide negative controls. Supplementary Table 1. Single-gene q-ChIP data for the results presented in the indicated Figures Data are shown as mean ± SD

Conditions ChIP Data in Figure Gene name Dad1 Lpp Tbc1d1 Cabin1 Dlk Mean SD Mean SD Mean SD Mean SD Mean SD Basal SLUG Figure 1B 1,42 0,3 2,82 0,12 0,81 0,04 1,12 0,09 2,11 0,25 Basal SNAIL Figure 1B 4,32 0,5 3,24 0,03 4,14 0,21 3,21 0,56 6,12 0,21 SLUG SLUG Figure 1B 6,41 1,1 10,2 0,84 8,12 1,12 7,65 0,69 8,54 1,25 SLUG SNAIL Figure 1B 0,15 0,07 1 0,23 0,07 0,05 0,3 0,1 0,17 0,06 SNAIL SLUG Figure 1B 0,23 0,1 0,41 0,03 0,32 0,04 0,09 0,02 0,12 0,02 SNAIL SNAIL Figure 1B 12,41 0,87 10,21 1,1 9,35 0,65 12,26 1,25 11,25 1,54 Scramble AHR Figure 1C 2,82 0,4 3,48 0,96 3,24 0,23 3,03 0,21 1,51 0,08 Scramble SLUG Figure 1C 1,42 0,3 2,82 0,12 0,81 0,04 1,12 0,09 2,11 0,25 Scramble SNAIL Figure 1C 4,32 0,5 3,24 0,03 4,14 0,21 3,21 0,56 6,12 0,21 shAHR AHR Figure 1C 1,11 0,23 0,21 0,05 0,53 0,1 1,03 0,08 0,11 0,03 shAHR SLUG Figure 1C 0,31 0,02 0,54 0,12 0,06 0,01 1,54 0,04 1,07 0,12 shAHR SNAIL Figure 1C 0,87 0,12 1,24 0,06 0,61 0,2 1,53 0,98 1,74 0,32 Basal TFIIIB Figure 3B 0,42 0,05 0 0 0,06 0,01 0,5 0,01 0,4 0,05 AHR TFIIIB Figure 3B 0 0 0,24 0,08 0 0 0,01 0 0,05 0,01 SLUG TFIIIB Figure 3B 0,05 0,02 0,6 0,01 0,01 0,01 0,04 0,01 0,06 0,01 A/SHRLUG TFIIIB Figure 3B 0,03 0,03 0,1 0 0 0 0,02 0,01 0,04 0,02 Basal TFIIIC Figure 3B 0,72 0,2 0,28 0,03 0,29 0,3 0,75 0,2 1 0,2 AHR TFIIIC Figure 3B 0 0 0,17 0,05 0 0 0,12 0,04 0,05 0,01 SLUG TFIIIC Figure 3B 0,01 0,02 0,02 0 0,03 0,02 0,03 0,01 0 0 AH/SRLUG TFIIIC Figure 3B 0,008 0 0,01 0,02 0,001 0 0,007 0,01 0 0 Basal POL III Figure 3B 4 0,5 2,6 0,4 10 2 2,3 0,5 4,4 0,8 AHR POL III Figure 3B 0,87 0,2 0,4 0,1 2,39 0,5 0,8 0,2 0,5 0,2 SLUG POL III Figure 3B 0,4 0,2 0,2 0,05 0,1 0,05 0,2 0,05 0,1 0,08 AH/SRLUG POL III Figure 3B 0,28 0,1 0,11 0,01 0,09 0 0,09 0,02 0,3 0,08 Basal H3 Figure 3C 0,35 0,03 0,5 0,1 0,3 0,01 0,4 0,01 0,6 0,03 AH/SLUGR H3 Figure 3C 0,4 0,02 0,3 0,1 0,4 0,05 0,3 0,04 0,4 0,01 Basal POL II Figure 4B 11 2,3 0 0 0 0 14 1,8 0,7 0,03 AHR POL II Figure 4B 12 2,1 12 3 15 2,5 70 12,4 10 0,8 Basal POL III / TFIIIC Figure 5B 4,2 0,5 1,95 0,4 2,24 0,3 2,24 0,4 2,76 0,6 Basal POL III / POL II Figure 5B 1,82 0,5 1,48 0,3 1,04 0,25 0,6 0,01 0,6 0,02 Basal POL III / GAPDH Figure 5B 1,95 0,4 1,12 0,3 1,58 0,2 0,79 0,02 0,37 0,01 Basal CTCF Figure 6D 0,43 0,1 0,43 0,08 0,08 0,02 4 0,5 0,17 0,03 AHR/SLUG CTCF Figure 6D 0,57 0,06 0,53 0,09 0,25 0,4 19 1,1 0,15 0,04 AH/SLUGR +ABA CTCF Figure 6D 0,49 0,2 0,48 0,09 0,14 0,2 8 0,8 0,16 0,05 Basal PARP1 Figure 6D 0,46 0,1 0,25 0,05 0,06 0,01 0,4 0,01 0,18 0,03 AH/SLUGR PARP1 Figure 6D 0,81 0,3 0,53 0,05 0,21 0,3 0,37 0,01 0,46 0,1 AH/SLUGR +ABA PARP1 Figure 6D 0,5 0,1 0,6 0,1 0,2 0,04 0,3 0,01 0,3 0,06 Supplementary Table 2. Single-gene q-ChIP data for the results presented in Figure 6A and 6B Data are shown as mean ± SD

ChIP B1-X35S Conditions Primer positions -1000 -500 500 1000 1500 Mean SD Mean SD Mean SD Mean SD Mean SD Figure 6A Basal 0 0 0,2 0,01 0,09 0,01 0,03 0,01 0,01 0,02 Dad1 Figure 6A A/SHRLUG 0,33 0,1 0,35 0,1 0,39 0,1 0,49 0,05 0,72 0,3 Figure 6A Basal 0,12 0,02 0,24 0,02 1,1 0,2 0,86 0,01 1,1 0,2 H3K9me3 Lpp Figure 6A A/HSLUGR 0,29 0,04 0,13 0,03 1,9 0,2 3,6 0,4 3,5 0,3 Figure 6A Basal 0,18 0,02 0,14 0,02 0,8 0,11 0,3 0,02 0,4 0,1 Cabin1 Figure 6A A/HSLUGR 0,29 0,03 1 0,1 2,8 0,12 0,27 0,1 0,37 0,2 Figure 6B Basal 0,39 0,1 0,91 0,1 0,5 0,03 0,1 0,05 0,19 0,05 Dad1 Figure 6B A/HSLUGR 1 0,2 0,82 0,09 0,96 0,04 0,4 0,03 1,1 0,2 Figure 6B Basal 1,28 0,21 1,68 0,12 2,78 0,8 1,84 0,4 1,5 0,1 H3K27me3 Lpp Figure 6B A/HSLUGR 1,46 0,18 1,8 0,11 3 0,4 3,6 0,5 4 0,06 Figure 6B Basal 0,34 0,08 0,41 0,09 2,74 0,6 0,92 0,2 1 0,02 Cabin1 Figure 6B A/HSLUGR 0,55 0,1 1,52 0,19 6,42 1 1,2 0,4 2,4 0,9

Supplementary Table 3

Gene Ontology comparison of genes with B1-X35S and the rest of the genome

B1-X35S prom2 B1-X35S intron3 B1-X35S prom4 GO Name1 GO ID vs vs vs rest of genome rest of genome B1-X35S intron

Neurological Process GO:0050877 31.2% vs 68.8% 25.1% vs 74.9% Primary Metabolic GO:0044238 53.1% vs 46.9% 53.0% vs 47.0% Process Sensory Perception GO:0007600 26.3% vs 73.7% 14.9% vs 85.2% Signal Transduction GO:0007165 44.2% vs 55.8% 46.4% vs 53.6% ATP binding GO:0005524 58.0% vs 42.1% 60.3% vs 39.7% G-protein coupled GO:0004930 30.5% vs 69.5% 21.1% vs 78.9% receptor activity Cell communication GO:0007154 44.3% vs 55.7% 47.2% vs 52.8% Cell Cycle GO:0007049 58.2% vs 41.8% Ubiquitin Ligase GO:0000151 73.5% vs 26.5% complex Nucleotide binding GO:0000166 57.4% vs 42.6% Establishment of GO:0051234 57.2% vs 42.9% localization Defense response GO:0006952 36.3% vs 63.7% Transport GO:0006810 57.3% vs 42.7% Secretion GO:0046903 60.1% vs 39.9% Cell motility GO:0006928 58.3% vs 41.7% Cell Adhesion GO:0007155 59.5% vs 40.5% 29.3% vs 70.7% Response to biotic GO:0009607 32.4% vs 67.6% 74.4% vs 25.7% stimulus Structural constituent GO:0003735 21.2% vs 78.8% 83.4% vs 16.6% of ribosome Protein binding GO:0005515 55.5% vs 44.5% 46.1% vs 53.9%

1 Gene ontology (GO) classification into different functional categories. 2 Percentage of genes having B1-X35S in upstream promoter regions (<10 kb) with respect to the percentage of genes not having B1-X35S. 3 Percentage of genes having B1-X35S in introns with respect to the percentage of genes not having B1-X35S. 4 Percentage of genes having B1-X35S in upstream promoter regions (<10 kb) with respect to the percentage of genes having B1-X35S in introns. All comparisons are statistically significant at p<0.001.

Supplementary Table 4

Primer sequences used in ChIP assays

Gene name* Primer sequence (5´-3´)

Dad1 Forward: CTGGCCTCCAACTCAGGAAT Reverse: GGGAGAGTAGAGGGGGCATA

Lpp Forward: GGTCCATCCTCTGGACAAAT Reverse: TACCAACACAGGTTGGAGGA

Cabin1 Forward: CAAACTCTTACAATATGGGGGA Reverse: CACCATGTTGGGAATTGAACT

Tbc1d1 Forward: AACACCCTGAGTCATTTCCC Reverse: CGCATGAGTTTCCTTTGAGA

Dlk1 Forward: ACATTTGCCCAGTGAGCTTT Reverse: GTTGGTTGGTTGGTTTGGTT

tRNA-Leu Forward: CTGTAAGTCAGGATGGCCG Reverse: TGGACAGACAAGCAGAAACAA

Dad1(-1000) Forward: TTGGAGAGCTGGCAACTATG Reverse: AGGAGGGGCAAATGAGTATG

Dad1(-500) Forward: AGGACAACAAAAGCTCCCATA Reverse: CGCTTATTTGGAACATGTGG

Dad1(+500) Forward: CCAATGTGTCTTTGCCAGTC Reverse: TCCAGAGCCCATGATAAATG

Dad1(+1000) Forward: TAATTTCCAGGCGAGGCTAC Reverse: GCTCATGTGCAGAGGGATAG

Lpp(-1000) Forward: TCCACACTGGCCATCATATT Reverse: GCCAAGTGCTAGGAGGTCTT

Lpp(-500) Forward: AGGGAAAATATGCCCACAAA Reverse: ATCATTTTGTCCCACTGCAA

Supplementary Table 4 (continued)

Lpp(+500) Forward: CATCAATGGGAGGAGAGGAC Reverse: TCAAATGCTATCCCCTTTCC

Lpp(+1000) Forward: GCAGCATTAGTTGGCACAGT Reverse: AATGAAGCTGCTTGTGATGC

Cabin1(-1000) Forward: TGTTCTGTTTGAGGCTCTGG Reverse: GTCAGGGCCACCTAGAAAGA

Cabin1(-500) Forward: CCGCATTTTGTTGTTGTTGT Reverse: ATTTAAAGCTGGGCATGGTG

Cabin1(+500) Forward: CCAGAAGAAGCCAGAAGAGG Reverse: TGGCAAAGTATCCCACAAGA

Cabin1(+1000) Forward: ACGTTTCTTAAGCGGGTCTG Reverse: CCTGGTCTGGAACTCCGTAT

*Each “gene name” indicates a gene having a B1-X35S element within 10 Kbp of genomic sequence upstream of the transcription start site.

Supplementary Table 5

Additional primer sequences used

Oligo name Primer sequence (5´-3´)

Primers for DNA binding affinity, enhancer blocking assays and in vitro transcription

B1-X35 GCCGGGCGTGGTGGCGCACGCCTTTAATCCCAGCACTTGGG AGGCAGAGGCAG

B1-35S CGAGACAGGGTTTCTCTGTATGCCCTGGCTGTCCTGGAACTC ACTCTGTAGACCAGGCTGGCCTCGAACTCAGAAATCCACCT GCCTCTGCCTCCCAAGT

B1-Xmut35 GCCGGGCGTGGTGGCGCTATCCTTTAATCCCAGCACTTGGG AGGCAGAGGCAG

B1-35Smut CGAGACAGGGTTTCTCTGTATGCCCTGGCTGTCCTGGAACTC ACTCTGTAGACCAGGCTGGCCTCGAACTCAGAAATCTGCCT GCCTCTGCCTCCCAAGT

B1-X35-ABmut GCCGGGGAATTCGGCGCACGCCTTTAATCCCAGCACTTGGG AGGCAGAGGCAG

B1-35S-ABmut CGAGACAGGGTTTCTCTGTATGCCCTGGCTGTCCTGGAACTC ACTCTGTAGACCAGGCTGGCCGAATTCTCAGAAATCCACCT GCCTCTGCCTCCCAAGT

Primers for DNA binding affinity, enhancer blocking assays and strand-specific in vitro transcription

B1-Flank-F CTCGAGCTGCAGGCCGGGCGTGGTGGC

B1-Flank-R CTCGAGCTGCAGCGAGACAGGGTTTCT

Primer for specific B1-X35S PCR amplification

B1-X35S-R TGGCCTCGAACTCAGAAATCCA

APPENDIX II

“Noggin and Noggin-Like genes control dorsoventral axis regeneration in planarians”

In this study, it was demonstrated that a BMP/ADMP organizer governs DV axis reestablishment during planarian regeneration. Also, planarian noggin genes were shown to function as canonical BMP inhibitors while the silencing of planarian nlg8 induces ectopic neurogenesis and enhances ventralizing bmp (RNAi) phenotypes. In addition, we revealed that noggin-like genes are conserved from cnidarian to vertebrates and that both planarian nlg8 and Xenopus nlg ventralize Xenopus embryos when overexpressed.

I tested the D/V function of planarian noggin (nog) and noggin-like (nlg) in gain-of- function studies using Xenopus laevis as model system. Using the same type of assays I checked the importance of the insertion within the noggin domain in the opposite function of nog and nlg. I also tested if the ventralizing activity of Xlnlg depended on BMP and ADMP signaling. Results of these experiments are shown on Figure 4 (exception 4.C), Supplementar Figure 4 (exception 4.C and 4D).

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Supplemental Information

Noggin and Noggin-Like Genes

Control Dorsoventral Axis

Regeneration in Planarians M. Dolores Molina, Ana Neto, Ignacio Maeso, José Luis Gómez-Skarmeta, Emili Saló, and Francesc Cebrià

Figure S1, Related to Figure 1. (A) Smed-admp-1 expression pattern in intact and regenerating planarians. Smed-admp-1 is expressed at the ventral midline and along the body margins in intact planarians. During regeneration admp-1 is detected within the anterior blastema from day 5 and in posterior blastemas from day 3. Ventral views. Scale bar, 250μm. (B) Graphical representation of the relative quantitative expression of Smed-nog1 and Smed- nog2 after nog1(RNAi);nog2(RNAi) silencing at 6 days of regeneration after the first round of injection and amputation. Expression levels relative to gfp(RNAi) organisms at the same point of regeneration (doted line). Data are means ± s.d. for at least three experiments. *, P<0.05; ***, P<0.001, Student‟s t-test. (C) Smed-ifb labelling the DV margin of control animals and nog1(RNAi);nog2(RNAi) organisms. Anteroventral expansion (red arrowheads) and broader labelling of the posterior marginal region is observed after nog1(RNAi);nog2(RNAi) silencing (compare yellow arrowheads). 19 days of regeneration after the third round of injection and amputation. Anterior to the left. Scale bar, 400μm.

1

2

Figure S2, Related to Figure 2. (A) Graphical representation of the relative quantitative expression of Smed-nlg8 after nlg8 silencing at 6 days of regeneration after the first round of injection and amputation. Expression levels relative to gfp(RNAi) organisms at the same point of regeneration (doted line). Data are means ± s.d. for at least three experiments. *** P < 0.001, Student‟s t-test. (B) Molecular characterization of nlg8(RNAi) anterior blastemas. nlg8-silenced planarians regenerated asymmetric blastemas with deficient differentiation of the DV boundary (labelled by Smed-ifb), mechanosensory cells (labelled by cintillo) and brain (labelled by Smed-GluR and anti-synapsin antibody). Some treated animals did not regenerate at all (right column). ifb and GluR expression at 18 days of regeneration. cintillo expression at 11 days of regeneration. Anti- synapsin immunostaining at 10 days of regeneration. Anterior to the top. Scale bar, 200μm. (C) nlg8(RNAi) resembles some bmp loss-of-function phenotypes [3]. Anterodorsal cintillo- positive mechanosensory cells (bracket, 6 days of regeneration) and brain lateral branches labelled for GluR (red arrowhead, 18 days of regeneration) spread dorsally after nlg8 silencing. Abnormal visual projections labelled with anti-VC-1 after nlg8(RNAi) treatment (purple arrowheads), compared to the normal stereotypical pattern of visual axonal projections found in control planarians (21 days of regeneration). All images show anterior blastemas. Anterior to the left. Scale bar, 100μm. (D) nlg8(RNAi) does not duplicate the planarian DV boundary. Even though there is a dorsal spreading of Smed-mag-1 labelled subepidermal marginal gland cells after both nlg8(RNAi) and bmp(RNAi) (red arrowheads), a clear DV border duplication is only observed through ifb expression after bmp(RNAi) (yellow arrowheads). 18-24 days of regeneration. Scale bar, 600μm.

3

Figure S3, Related to Figure 3. (A) Triple nog1/nog2/nlg8 silencing enhances nog1(RNAi);nog2(RNAi) phenotypes. After the second round of nog1/nog2/nlg8 dsRNA injection more than 80% of organisms developed head- like structures on their ventral side and about 65% differentiated ectopic ventral eyes within it. The silencing of nog1/nog2 at the concentration needed as control (1/3 of each noggin diluted with 1/3 of gfp dsRNA) did not produce ventral outgrowths and ectopic eyes after either the second or the third round of treatment. Therefore, to see whether bmp(RNAi) or admp-1(RNAi) rescued the dorsalization of the ventral side seen after nog1 and nog2 silencing, we co-silenced nog1/nog2/nlg8/bmp or nog1/nog2/nlg8/admp-1 instead of nog1/nog2/bmp or nog1/nog2/admp- 1. (B) Graphical representation of the relative quantitative expression of Smed-nlg8 and Smed-bmp at 6 days of regeneration after the second round of injection and amputation. Expression levels

4

relative to gfp(RNAi) organisms at the same point of regeneration (doted line). Data are means ± s.d. for at least three experiments. ***, P<0.001, Student‟s t-test. (C) Expression of bmp and admp-1 after nog1/nog2/nlg8 silencing. Smed-bmp is expressed at the dorsal midline of both control and triple RNAi organisms. In addition, ectopic bmp expression (arrowheads) is detected at the ventral midline after nog1/nog2/nlg8 silencing. On the other hand, the expression of admp-1 along the ventral midline disappears after triple nog1/nog2/nlg8 silencing. 5 days of regeneration after the third round of dsRNA injection and amputation. Scale bar, 150μm. (D) Graphical representation of the relative quantitative expression of Smed-nlg8, Smed-nog1 and Smed-nog2 at 6 days of regeneration after the first round of injection and amputation of different combinatorial RNAi experiments. Note that similar levels of inhibition are found in all the combinations. Expression levels relative to gfp(RNAi) organisms at the same point of regeneration (doted line). Data are means ± s.d. for at least three experiments. ***, P<0.001, Student‟s t-test.

5

6

7

E

Figure S4, Related to Figure 4. (A) Summary of the phenotypes obtained by injection of planarian nlgs into Xenopus embryos. The overexpression of planarian nlg in Xenopus resulted in normal and ventralized organisms. However, some dorsalized embryos were also obtained after nlg5 injection. (B) All panels show lateral views of Xenopus laevis embryos at stage 25 of development after overexpression of indicated mRNAs. Overexpression of planarian nog1 and Xenopus nog1 dorsalize the embryos, whereas overexpression of planarian nlg8 and Xenopus nlg ventralize them. Overexpression of Xlnog1_CnlgC (Xlnog1 mRNA with the W215-Q227 peptide domain of Xlnlg inserted between C197-C205) produces mild dorsalization. Overexpression of Xlnlg_CnogC (Xlnlg mRNA with the Q198-K204 peptide domain of Xlnog1 inserted between C214-C228) produces dorsalization. Anterior to the left. Scale bar, 1mm. (C) noggin and noggin-like gene complement across metazoans. A schematic tree on the left shows the phylogenetic relationships of the species analyzed in the present study. Non- eumetazoans are colored in gray, Cnidarians are shown in yellow, and the three main bilaterian clades, Deuterostomes, Ecdysozoans and Lophotrochozoans, are depicted in blue, green and purple, respectively. Figures on the right indicate the presence and number of nog and nlg genes. Asterisks mark the species for which there is no whole genome project available and therefore a complete nog and nlg complement could not be confidently determined. Ψ refers to a human nlg pseudogene. (D) Eight million-generation Bayesian inference phylogenetic reconstruction of the noggin and noggin-like genes found across metazoans estimated under the WAG + I + Γ model as recommended by Prottest. The noggin homologues found from cnidarians to vertebrates cluster into two different groups, the noggin (88% of posterior probability) and the noggin-like gene (99% of posterior probability) group. Placozoans (Tad) and Sponges (Aqu, Emu and Sdo) possess an ancestral nog-nlg homologue. Species are abbreviated as follows: Aca, Anolis carolinensis; Apa, Acropora palmata; Apy, Acyrtosiphon pisum; Aqu, Amphimedon queenslandica; Bfl, Branchiostoma floridae; Cte, Capitella teleta; Dja, Dugesia japonica; Dpu,

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Daphnia pulex; Dre, Danio rerio; Emu, Ephydatia muelleri; Gga, Gallus gallus; Hsa, Homo sapiens; Isc, Ixodes scapularis; Lgi, Lottia gigantea; Lsa, Lepeophtheirus salmonis; Mli, Macrostomum lignano; Nve, Nematostella vectensis; Pma, Petromyzon marinus; Sdo, Suberites domuncula; Sko, Saccoglossus kowalevskii; Sma, Schistosoma mansoni; Sme, Schmidtea mediterranea; Spu, Strongylocentrotus purpuratus; Tad, Trichoplax adhaerens; Tru, Takifugu rubripes; Rpr, Rhodnius prolixus; Xla, Xenopus laevis. (E) Table and graphical representation summarizing the statistical analysis of the ratio of activated SMAD (P-SMAD1/5/8) versus total SMAD1/5/8 levels after several combinatorial mRNA injections in Xenopus embryos. Values are relative to uninjected organisms at the same point of development. s.d., standard deviation; n, number of experiments; s.e.m., standard error of the means. Xlnog1, Xlbmp4, Xlbmp4+Xlnog, Xlbmp4+Xlnlg and Xladmp mRNA injections significantly modify the relative levels of P-SMAD1/5/8 when compared to uninjected organisms (asterisks). Note that not significant variations are detected after Xlnlg mRNA overexpression either alone (compared to uninjected organisms) or coinjected with Xlbmp4 (compared to Xlbmp4 mRNA injections). In contrast, Xlnog1 mRNA injections rescue the upregulation in the levels of P-SMAD1/5/8 caused by Xlbmp4 overexpression (indicated as and compared to Xlbmp4 mRNA overexpression). Neither Xlnog1 not Xlnlg mRNA injection modify P-SMAD1/5/8 levels when coinjected with Xladmp. The same results were obtained after injection of two different mRNA concentrations of each condition (see Supplemental Experimental Procedures). Data are means ± s.e.m. for at least two experiments. *, significant variations relative to controls. , significant variations relative to Xlbmp4 mRNA injection. n.s., non-significant *, P<0.05; **, P<0.001; , P<0.001. Student‟s t-test.

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Supplemental Experimental Procedures

Animals

The Schmidtea mediterranea clonal line BCN-10 was used for all experiments. Animals were maintained at 20˚C in a 1:1 (v/v) mixture of distilled water and tap water treated with AquaSafe (TetraAqua). Planarians were fed with beef liver and starved for at least one week prior to experiments.

Identification and Isolation of Smed-ifb and Smed-admp-1 Genes

Dugesia japonica Intermediate Filament b, DjIFb [30], Schmidtea mediterranea BMP, Smed- BMP [3] and ADMP sequences from several organisms were used in tblastn searches against the S.mediterranea genome assembly draft 3.1. (available at http://genome.wustl.edu). A single genomic contig encoding SMED-IFB (Contig1239.3) and two genomic contigs encoding SMED- ADMP-1 (Contigs 5853.2 and 7118.1) were found. Comparisons between the admp contigs revealed low variability within the exonic regions. Since very few changes were also found within the intronic regions (for instance, in intron 2, only five changes among 140 nucleotide positions) we considered that each contig represented different haplotypes of a single ADMP-1 locus. Sets of specific primers were designed to amplify candidate genes from cDNA made from total RNA using Superscript III (Invitrogen). Sequencing of the amplified admp-1 cDNAs revealed the existence of two additional haplotypes confirming the elevated degree of polymorphism of the ADMP-1 locus. Genes were named according to the S. mediterranea gene nomenclature guidelines [31]. GenBank accession numbers are: Smed-admp-1a HM135945, Smed-admp-1b HM135946, Smed-admp-1c HM135947, Smed-admp-1d HM135948, and Smed- ifb HM135949.

RNAi Analysis

Double-stranded RNAs [20] for planarian noggins Smed-nog1 and Smed-nog2 [7], noggin-like genes Smed-nlg1 to Smed-nlg8 [7], Smed-admp-1 and Smed-bmp [3] were synthesized by in vitro transcription and injected into the planarian digestive system using a Drummond Scientific Nanoject injector (Broomall, PA, USA). To avoid off-target effects of RNAi and confirm the specificity of the phenotypes observed, two different, non-overlapping regions of each gene were used to synthesize dsRNA. For simultaneous silencing of two or more genes, dsRNAs were proportionally diluted and animals with the same dose of each single dsRNA were injected in parallel as controls; for instance, the injection of 1/3 nlg8 + 2/3 gfp dsRNA and 1/3 nog1 + 1/3 nog2 + 1/3 gfp dsRNA were performed as controls of 1/3 nogl + 1/3 nog2 + 1/3 nlg8 dsRNA treatment (referred to as nog1/nog2/nlg8 within the text). Combinatorial RNAi experiments against nlg-1 to 7 and noggin genes are not shown as they did not produce an enhancement of the nog1(RNAi);nog2(RNAi) phenotype. Treated planarians were amputated pre- and post- pharyngeally three days after the first injection, and the trunk pieces were allowed to regenerate new heads and tails. In some experiments, animals were injected and amputated for two, three or four consecutive weeks, in each case with a three day gap between injection and amputation. In all images “X”d refers to days after amputation and “X”R to rounds of injection and amputation. For example 19d 3R (Fig. 1E) means 19 days of regeneration after the third round of injection

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and amputation. Control animals were injected either with water or, in most cases, with dsRNA against gfp, a gene not found in the planarian genome. Live animals were observed through a Zeiss Stemi SV6 stereomicroscope and images from representative organisms of each experimental condition were captured with a DeltaPix InfinityX camera.

The dsRNAs used in each experiment are summarized as follows: Fig.1A, E gfp dsRNA injection. Fig. 1B admp-1/gfp dsRNA injection. Fig. 1C bmp/gfp dsRNA injection. Fig. 1D bmp/admp-1 dsRNA injection. Fig. 1F nog1/nog2 dsRNA injection. Fig. 2A gfp dsRNA injection. Fig. 2B nlg8 dsRNA injection. Fig. 3A nog1/nog2/nlg8 and nlg8/2xgfp dsRNA injection. Fig. 3B nog1/nog2/nlg8 and nog1/nog2/gfp dsRNA injection. Fig. 3C gfp and nog1/nog2/nlg8 dsRNA injection. Fig. 3D nog1/nog2/nlg8 dsRNA injection. Fig. 3E bmp/2xgfp dsRNA injection. Fig. 3F nlg8/bmp/gfp dsRNA injection. Fig. 3G gfp, nog1/nog2/nlg8/bmp, nog1/nog2/nlg8/admp-1, nog1/nog2/nlg8/gfp dsRNA injection. Fig. 3H nog1/nog2/nlg8/gfp and nog1/nog2/nlg8/admp-1 dsRNA injection. Fig. S1B,C gfp and nog1/nog2 dsRNA injection. Fig. S2A-C gfp and nlg8 dsRNA injection. Fig. S2D gfp, nlg8 and bmp dsRNA injection. Fig. S3A gfp, nog1/nog2/gfp and nog1/nog2/nlg8 dsRNA injection. Fig. S3B gfp, bmp/2xgfp, nlg8/bmp/gfp dsRNA injection. Fig. S3C gfp and nog1/nog2/nlg8 dsRNA injection. Fig. S3D gfp, nog1/nog2/nlg8/bmp, nog1/nog2/nlg8/admp-1, nog1/nog2/nlg8/gfp dsRNA injection.

Quantitative Real-Time PCR

Total RNA was extracted from a pool of three head or trunk fragments of RNAi treated planarians using TRIzol® reagent according to manufacturer‟s instructions. Isolated RNA was treated for 20 min at 37oC with DNase I (Roche) to remove contaminating genomic DNA. First- Strand cDNA was reverse transcribed from l μg of total RNA using a First-Strand Synthesis System kit from Invitrogen. Real time PCR was performed using SYBR Green (Applied Biosystems) in an ABI Prism 7900HT Sequence Detection System (Applied Biosystems). Three samples for each condition were run in parallel. Samples without reverse transcriptase served as the negative control template. Data were normalized to the expression of the internal control UDP [32]. The lack of primer dimerization or nonspecific PCR product bands was tested. Statistical analyses were performed with SPSS software.

The following sets of specific primers were used: Smed-nlg8F 5‟ TATTACAGTGAATTCTCCTTCG 3‟

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Smed-nlg8R 5‟ ATTTAGAAACGTTATCAAAGTGG 3‟ Smed-nog1F 5‟ GTTCGATCTGAATGAAAAGCC 3‟ Smed-nog1R 5‟ CTCTCGGTTGGAAGATTCG 3‟ Smed-nog2F 5‟ GAAATCGTTTCTTCGGAAGGC 3‟ Smed-nog2R 5‟ CCAGAAACGTTCTTAATTTTCG 3‟ Smed-admp-1F 5‟ CCAATTATTTCAATTTACACTCG 3‟ Smed-admp-1R 5‟ TAGGATCCATATTTTCAAATGCC 3‟ Smed-bmpF 5‟ GTGAATATCGAAGGAAAACCTG 3‟ Smed.bmpR 5‟ TTGTGGATACCAACGGATGAC 3‟ Smed-udpF 5‟ CATTCACGTTGTCGATCTAGC 3‟ Smed-udpR 5‟ CCGAATATCCTCTGCCAGTG 3‟

Whole-Mount In Situ Hybridization

Gene expression analysis was carried out in an Intavis InsituPro hybridization robot as described elsewhere [3]. Digoxigenin-labeled riboprobes for Smed-bmp [3], Smed-admp-1, Smed-ifb (DV boundary), Smed-septin (dorsal mesenchymal cells) [33], Smed-eye53 [33], cintillo (mechanosensory cells around the cephalic ganglia) [34], Smed-GluR (lateral branches of the cephalic ganglia) and Smed-mag-1 (subepidermal marginal gland cells) [35] were synthesized using an in vitro transcription kit (Roche). All samples were observed through a Leica MZ16F stereomicroscope and images from representative organisms in each experiment were captured with a ProgRes®C3 camera from Jenoptik.

Immunostaining

Animals were killed by immersion in 2% HCl for five minutes on ice and then fixed in Carnoy‟s solution for two hours at 4oC. After fixation, samples were processed as described elsewhere [20]. The following primary antibodies were used: VC-1 [36], a mouse monoclonal antibody specific for planarian photosensitive cells (kindly provided by Hidefumi Orii, diluted 1:15000); anti-SYNORF1, a mouse monoclonal antibody specific for synapsin (Developmental Studies Hybridoma Bank, diluted 1:50) and used as a pan neural marker; and AA4.3, a mouse monoclonal antibody specific for α-tubulin (Developmental Studies Hybridoma Bank, diluted 1:20) and used to label the cilia found in both dorsal and ventral epidermis. Goat anti-mouse secondary antibody conjugated to Alexa 488 (Molecular Probes) was used at a 1:400 dilution. Samples were mounted in SlowFade® Gold antifade reagent (Invitrogen). Confocal laser scanning microscopy was performed with a Leica TCS-SPE (Leica Lasertechnik, Heidelberg) and images were processed using ImageJ 1.43m and Photoshop CS 8.0.1 software. Confocal stacks from representative organisms are shown for each experimental condition.

In Silico Identification and Comparison of noggin and noggin-like genes across Metazoans

Using previously described noggin and noggin-like genes from various metazoans (vertebrates, amphioxus, cnidarians and planarians) as queries, we performed tBLASTN and BLASTP searches against the genomes of Trichoplax adhaerens Grell-BS-1999 v1.0, Nematostella vectensis JGI v1.0, Branchiostoma floridae JGI v1.0, Takifugu rubripes JGI v4.0, Daphnia pulex JGI v1.0, Lottia gigantea JGI v1.0 and Capitella teleta JGI v1.0 through the JGI webpage

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(http://genome.jgi-psf.org/euk_home.html); of Strongylocentrotus purpuratus Build 2.1, Acyrthosiphon pisum Build 1.1, Apis mellifera Build 4.0, Nasonia vitripennis Build 1.1, Tribolium castaneum Build 2.1, Drosophila melanogaster Build Fb5.3, Homo sapiens Build GRCh37, Mus musculus Build 37.1, and Danio rerio Build Zv8 through the NCBI webpage (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi); of Rhodnius prolixus and Trichinella spiralis through the NCBI genomic BLAST webpage for unfinished eukaryotic genomes (http://www.ncbi.nlm.nih.gov/sutils/genom_table.cgi?organism=eukaryotes); of Petromyzon marinus through the Ensembl Pre! webpage (http://pre.ensembl.org/Multi/blastview); of Saccoglossus kowalevskii through the Baylor College of Medicine webpage (http://www.hgsc.bcm.tmc.edu/project-species-x-organisms.hgsc); of Ixodes scapularis through the VectorBase webpage (http://www.vectorbase.org/Tools/BLAST/); of Macrostomum lignano through the M. lignano genome initiative webpage (http://www.macgenome.org/blast/index.html); and of Schistosoma mansoni through the Sanger Institute webpage (http://www.sanger.ac.uk/cgi-bin/blast/submitblast/s_mansoni). For the sponge Amphimedon queenslandica we downloaded the genomic traces from the NCBI webpage (ftp://ftp.ncbi.nih.gov/pub/TraceDB/) and performed tBLASTN searches locally. We then downloaded each corresponding genomic region and built different gene models using GenScan [37] and GeneWise2 [38] software when necessary. We compared these predictions with ESTs and existing gene models when available. In general, both noggin and noggin-like genes lacked introns (45 out of 54 genes with available genomic data), and therefore the predicted coding sequences were reliable. To recover additional sequences from phylogenetically informative organisms without genome sequencing projects, we also performed tBLASTN searches against the nucleotide collection and the non-human, non-mouse ESTs databases through the NCBI webpage (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi), which allowed us to recover sequences from Acropora palmata, Dugesia japonica, Ephydatia muelleri, Lepeophtheirus salmonis and Suberites domuncula. Amino acid sequences for the whole sequence of the genes (available upon request) were aligned using MUSCLE [21] as implemented in Jalview 2.4 [39], and the resulting alignment was manually curated. The signal sequences preceding the „clip‟ and the cystine knot of the noggin domains could not be confidently aligned and were discarded for subsequent analyses. To avoid a possible artifactual grouping of the noggin-like genes due to the presence of their shared amino acid insertion between the fifth and sixth cysteines, this region of the alignment was also removed. A phylogenetic tree was then generated by the Bayesian method with MrBayes 3.1.2 [22, 23], with the model WAG + I + G, as recommended by ProtTest 1.4 [40-42] under the Akaike information and the Bayesian information criteria. Two independent runs were performed, each with four chains. By convention, convergence was reached when the value for the standard deviation of split frequencies stayed below 0.01. Burn-in was determined by plotting parameters across all runs: all trees prior to stationarity and convergence were discarded, and consensus trees were calculated for the remaining trees (from at least 1,000,000 generations).

Xenopus Microinjection of mRNA and In Situ Hybridization

The entire coding regions of Smed-noggins, Smed-noggin-like genes, Xlbmp4 [24, 25], Xlnog1 [16] and Xlnog4 [17], renamed Xlnlg after this work, were amplified by PCR and inserted into the multicloning site of pCS2+ [26]. The pCS2+ vectors containing the ORF of Xlnog1 and Xlnlg were used as a PCR template to swap the region located between the fifth and sixth cysteine

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residues within their noggin domains. Xlnog1_CnlgC: two consecutive PCR reactions (primer 5‟- CCACACAGCAATGTGCGTGGATAACCATTCAG-3‟ and primer 5‟- CCTTCATCCAACATCTCCACCTTAAGATGGTC-3‟ for the first PCR; primer 5‟- TTGGGCTGGGCCACACAGCAATGTGCGTGGA-3‟ and primer 5‟- ACCAGTGTCCTTCATCCAACATCTCCACCTT-3‟ for the second PCR) were used to insert the W215-Q227 peptide domain of Xlnlg and remove the Q198-K204 peptide domain of Xlnog1. Xlnlg_CnogC: a single PCR reaction (primer 5‟- TCAGCAGAAGTGTACCTGGAGGCAGGTACCCT-3‟ and primer 5‟- ACCCTGCGTTGGCAATGCCAGGCCAGCAGTTTA-3‟) was used to insert the Q198-K204 peptide domain of Xlnog1 and remove the W215-Q227 peptide domain of Xlnlg. The linearized PCR products were phosphorylated and religated. Fidelity of PCR reactions was verified by sequencing. For mRNA preparation, the plasmids were linearized with ApaI and transcribed with SP6 RNA polymerases and addition of GTP cap analog (New England Biolabs) as previously described [27]. After DNAse treatment, RNA was column purified, extracted with phenol- chloroform and precipitated with ethanol. mRNAs for injection were resuspended in water. Xenopus embryos were injected at the two-cell stage in a single blastomere. Images from representative organisms on stage 25-27 of each experiment were captured with an Olympus DP71 digital camera. X-Gal staining was performed as described elsewhere [28]. Antisense RNA probes were prepared from Sox2 and Szl cDNAs using digoxigenin (Roche). Xenopus specimens were hybridized as described [29]. The Xenopus (Xlnlg or Xlnog4) clone was obtained from the NIBB/NIG Xenopus laevis EST project (clone XL003k11).

Western Blotting

A pool of 10-20 Xenopus embryos from each condition were homogenized (1 µl per embryo) in lysis buffer (20 mM Tris pH 8, 50 mM NaCl, 50 mM NaF, 10 mM β-glycerophosphate, 2 mM EDTA, 1% NP40, 20 µg/ml aprotinin, 40µg/ml leupeptin, 4 µg/ml pepstatin, 0.75 mM PMSF, 1 mM Na3Vo4). Lysates were centrifuged for 10 minutes at 4°C and suspended in twice their volume of 2 x Laemmli buffer. 5µL of total protein extracts were denatured by boiling at 95 ºC for 5 minutes, separated by 10% SDS poly-acrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene fluoride (PVDF) membrane (Amersham HybondTM-P). Membranes were blocked for 1 hour with 5% non-fat milk dissolved in TBS-T buffer (20mM Tris; 150mM NaCl, 0.1% Tween 20, pH 7.5), rinsed with TPBS and incubated with primary antibody against phospho-SMAD1/5/8 ON at 4°C (Cell Signalling Technology, diluted 1:1000 in 5% BSA). Blots were washed in TBS-T buffer and incubated for 1h at RT with a goat anti-rabbit IgG peroxidase- conjugated antibody (Pierce Biotechnology, diluted 1:5000 in 5% non-fat milk). Immunoreactive protein was visualized by a chemiluminescence-based detection kit (EZ-ECL Kit from Biological Industries) with a LAS-4000 mini Imaging System from FujiFilm and semiquantitative values were measured with Quantity One software (Bio-Rad). After developing, the membrane was incubated at 60oC for 30min in stripping solution (62.5 mM Tris-HCl pH 6.8, 2% SDS, 10 mM β-Mercaptoethanol), at 60oC for 30min in 62.5 mM Tris-HCl pH 6.8, 2% SDS buffer, washed with TBST at RT and incubated ON at 4°C with the primary antibody against total SMAD1/5/8 (Smad1/5/8 (N-18)-R antibody from Santa Cruz Biotechnology, diluted 1:3000 in 5% non-fat milk). Blots were treated and developed as above. Statistical analyses were performed with SPSS software. Similar results were obtained after injection of two different mRNA concentrations for each condition:

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Xlnog1: 20pg and 80pg. Xlnlg: 500pg and 2ng. Xlbmp4: 250pg and 500ng. Xladmp: 100pg and 200pg. Xlbmp4+ Xlnog1: 250pg+20pg and 500pg+80pg. Xlbmp4+Xlnlg: 250pg+500pg and 500pg+2ng. Xladmp+Xlnog1: 100pg+20pg and 200pg+80pg. Xladmp+Xlnlg: 100pg+500pg and 200pg+2ng.

Supplemental References 30. Tazaki, A., Kato, K., Orii, H., Agata, K., and Watanabe, K. (2002). The body margin of the planarian Dugesia japonica: characterization by the expression of an intermediate filament gene. Dev Genes Evol 212, 365-373. 31. Reddien, P.W., Newmark, P.A., and Sanchez Alvarado, A. (2008). Gene nomenclature guidelines for the planarian Schmidtea mediterranea. Dev Dyn 237, 3099-3101. 32. Scimone, M.L., Meisel, J., and Reddien, P.W. The Mi-2-like Smed-CHD4 gene is required for stem cell differentiation in the planarian Schmidtea mediterranea. Development 137, 1231-1241. 33. Zayas, R.M., Hernandez, A., Habermann, B., Wang, Y., Stary, J.M., and Newmark, P.A. (2005). The planarian Schmidtea mediterranea as a model for epigenetic germ cell specification: analysis of ESTs from the hermaphroditic strain. Proc Natl Acad Sci U S A 102, 18491-18496. 34. Oviedo, N.J., Newmark, P.A., and Sanchez Alvarado, A. (2003). Allometric scaling and proportion regulation in the freshwater planarian Schmidtea mediterranea. Dev Dyn 226, 326-333. 35. Zayas, R.M., Cebria, F., Guo, T., Feng, J., and Newmark, P.A. (2010). The use of lectins as markers for differentiated secretory cells in planarians. Dev Dyn 239, 2888-2897. 36. Sakai, F., Agata, K., Orii, H., and Watanabe, K. (2000). Organization and regeneration ability of spontaneous supernumerary eyes in planarians -eye regeneration field and pathway selection by optic nerves. Zoolog Sci 17, 375-381. 37. Burge, C., and Karlin, S. (1997). Prediction of complete gene structures in human genomic DNA. J Mol Biol 268, 78-94. 38. Birney, E., and Durbin, R. (2000). Using GeneWise in the Drosophila annotation experiment. Genome Res 10, 547-548. 39. Waterhouse, A.M., Procter, J.B., Martin, D.M., Clamp, M., and Barton, G.J. (2009). Jalview Version 2--a multiple sequence alignment editor and analysis workbench. Bioinformatics 25, 1189-1191. 40. Drummond, A., and Strimmer, K. (2001). PAL: an object-oriented programming library for molecular evolution and phylogenetics. Bioinformatics 17, 662-663. 41. Guindon, S., and Gascuel, O. (2003). A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52, 696-704. 42. Abascal, F., Zardoya, R., and Posada, D. (2005). ProtTest: selection of best-fit models of protein evolution. Bioinformatics 21, 2104-2105.

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ABBREVIATIONS

A/P- Antero-Posterior

AER-Apical ectodermal ridge

Bowl- Bowel

BMP - Bone morphogenic protein

D/V- DORSO-VENTRAL

Cdx- Caudal

DNA- Deoxyribonucleic acid

Drm- Drumstick

DT- Distal tubules

D/V- dorso-ventral

Eh1- Engrailed homology 1

Exd- Extradenticle

FGF- Fibroblast growth factor

GAL4/UAS

GFP

Hpf- hours post fertilization

Hst- heartstrings ()

IM- Intermediate mesoderm

IT- Intermediate tubules

LPM- Lateral Plate Mesoderm

167 mRNA- messenger RNA

MO- Morpholino

MT- Myc tagged

Odd- Odd-skipped

Osr- Odd-skipped related

PBX/ – Pre-B cell Leukemia Transcription Factor/ P/D – Proximo-distal

Prdm1

PT- Proximal tubules

PZ - Progress zone

RA- Retinoic acid

RAR- Retinoid acid receptors

RNA- Ribonucleic acid

RT- Renal tubule

Sall1a

SalI4

SM- Somitic mesoderm

Shh - Sonic hedgehog

Sob- Sister of bowl

Tbx - T-box transcription factor

TGF-

TLS- Transducin-like Enhancer of Split

168 Wnt - Vertebrate homologue of the Wingless gene from Drosophila

ZF - ZINC FINGER

169

170

ACKNOWLEDGEMENTS

171