Correspondence to: Jean-Pierre Saint-Jeannet, New York University, College of Dentistry. Department of Basic Science & Craniofacial Biology, 345 East 24th Street, New York, NY 10010. E-mail: email@example.com
Located in the thoracic cavity of most vertebrates, the thymus is a primary lymphoid organ. It provides a unique microenvironment for T lymphocyte maturation, an essential step in the development of the adaptive immune system (Miller, 1961). Failure to organize or to maintain a proper thymic structure can have serious health consequences ranging from immunodeficiency to autoimmunity.
The thymus is one of four glands, along with the parathyroid, thyroid, and ultimobranchial bodies, that are derived from the foregut endoderm in the pharyngeal region. The pharyngeal apparatus is a transient structure composed of a series of bulges that form on either side of the developing head: the pharyngeal arches. Each pharyngeal arch is separated externally by ectoderm-lined pharyngeal clefts and internally by endoderm-lined pharyngeal pouches. Each pouch/cleft gives rise to a specific set of organs, although there is variation between species (reviewed in Grevellec and Tucker, 2010). In most organisms, thymus development is tightly linked to formation of the parathyroid glands (reviewed in Blackburn and Manley, 2004; Rodewald, 2008; Gordon and Manley, 2011). The two organs develop from a single primordium arising bilaterally from an outpocketing of the third pharyngeal pouch endoderm. The epithelial cells of the common primordium are initially indistinguishable, and patterning of the third pharyngeal pouch endoderm into distinct domains becomes apparent with the expression of two transcription factors, Foxn1 and Gcm2, that have restricted and complementary expression patterns in the prospective thymus and parathyroid domains, respectively (Gordon et al., 2001). Foxn1-expressing cells will form the thymus epithelial cells (TEC), the major component of the thymic stroma required for all stages of thymocyte development and differentiation. Lineage and functional analyses in mice have shown unambiguously that TEC from both the cortex and the medulla are exclusively derived from the pharyngeal endoderm (Gordon et al., 2004).
Each thymic primordium is surrounded by mesenchyme derived from the neural crest (NC) that migrates from the posterior hindbrain into the pharyngeal region. In chicken and mouse, these NC cells contribute to the connective tissue of the thymic capsule and become associated with the vasculature of the fetal and adult thymus (Le Lievre and Le Douarin, 1975; Le Douarin and Jotereau, 1975; Bockman and Kirby, 1984; Jiang et al., 2000, Foster et al., 2008; Muller et al., 2008). The development of a mature thymus depends on interactions between the NC-derived mesenchyme and the pharyngeal endoderm-derived TEC, NC cells stimulating proliferation and maturation of the epithelial compartment of the thymus (Itoi et al., 2007). Among the factors implicated are Fgf7 and Fgf10 produced by the NC mesenchyme, which promote both TEC proliferation and differentiation (Suniara et al., 2000; Revest et al., 2001; Dooley et al., 2007). The extent to which the NC participates in early thymus development is not as well described. In chicken embryos, NC ablation results in small thymi with delayed development and poor function (Bockman and Kirby, 1984). In the classic mouse mutants, Splotch, carrying a deletion of Pax3, the embryos have a dramatically reduced number of NC cells in the vicinity of the third pharyngeal pouch. However, the thymus and parathyroid primordia are normally specified (Griffith et al., 2009). A more recent study looking at compound mutant embryos homozygous for a NC-specific Foxd3 mutation and heterozygous for Pax3 reported severe thymus hypoplasia (Nelms et al., 2011). Moreover, DiGeorge syndrome patients have a range of congenital defects involving structures influenced by the cardiac NC, including cardiac deficiencies and craniofacial abnormalities but they also show thymus hypoplasia leading to T-cell deficiency and increased susceptibility to infections (reviewed in Wurdak et al., 2006).
The African clawed frog, Xenopus laevis, has been a model of choice for comparative and developmental studies of the immune system (reviewed in Du Pasquier et al., 1989; Robert and Ohta, 2009; Robert and Cohen, 2011). Its adaptive immune system in the adults is remarkably similar to that of mammals, and Xenopus has been an especially valuable system to analyze T-cell ontogeny and explore self-tolerance (reviewed in Du Pasquier et al., 1989; Robert and Ohta, 2009). While a great deal has been learned about the development and maturation of the immune system in Xenopus, less is known about early thymus development. A few reports published in the late 1960s briefly described thymus development in Xenopus (Nieuwkoop and Faber, 1967; Manning and Horton, 1969); however, these studies have provided only limited visual information on thymus organogenesis. Here we present a histological and molecular characterization of the emergence and development of the thymic rudiments in Xenopus. In particular, we examined the expression of three transcription factors that have been associated with pharyngeal gland development, gcm2, hoxa3, and foxn1, and highlight some of the more unique features of these genes as they relate to thymus development in frogs. Moreover, using transplantation experiments we analyzed the NC contribution to early thymus development.
Anatomy of the Developing Thymus
The thymus primordia were first morphologically recognizable at around stage 47 as two spherical masses on each side of the hindbrain. They were located dorsal-laterally between the developing inner ear and the eye (Fig. 1A–F). This is the stage at which the thymus rudiment is initially colonized by lymphoid precursors, which express the cell surface marker, RC47 (Du Pasquier and Flajnik, 1990). At stage 50, the thymi had increased in size and they retained their dorsolateral position (Fig. 1G–J), and expressed p63 (Fig. 1K,L), a factor essential for the proliferative capacity of the epithelial stem cells of the thymus (Senoo et al., 2007). By stage 55, the thymus rudiments were extensively colonized by NC-derived pigment cells that reside in the cortex (Fig. 1M,N), and the rudiments had shifted into a more superficial position, underneath the skin (Fig. 1O,P). In a transverse section, the cortex and medulla of the thymus were clearly visible (Fig. 1P,Q). While morphologically recognizable at this stage, the architecture of the cortex and medulla can be identified as early as stage 48/49 with the differential expression of MHC class II molecules by TEC (Du Pasquier and Flajnik, 1990) and CTX, a cell surface marker, by cortical thymocytes (Robert and Cohen, 1998).
Histological Analysis of Thymus Organogenesis
We next analyzed initial thymus development in Xenopus using serial sections prepared from tadpoles at stage 41 through stage 45 (Fig. 2). The transverse sections were made at a level posterior to the eye and anterior to the developing inner ear. Dorsal bilateral outpocketings of the second pharyngeal pouch endoderm were first visible at stage 41 (Fig. 2). As development proceeds, the thymus primordia became more rounded in shape but still connected to the pharyngeal pouch (stage 42 to stage 44; Fig. 2). The connection with the epithelium of the pharyngeal pouch was completely lost by stage 45 (Fig. 2). These findings are largely consistent with previous studies (Nieuwkoop and Faber, 1967; Manning and Horton, 1969).
Expression of gcm2, hoxa3, and foxn1 During Thymus Development
The three transcription factors Gcm2, Hoxa3, and Foxn1 have been functionally associated with pharyngeal gland development in mammals (reviewed in Manley and Condie, 2010). Here we analyzed the expression of these three genes as they relate to Xenopus thymus development.
gcm2. The vertebrate homologue of Drosophila gcm2 (glial cells missing 2) Gcm2, is exclusively expressed in the parathyroid glands and in the pharyngeal pouches from which they derive: the third and fourth pouches in chicken (Okabe and Graham, 2004; Neves et al., 2011; Grevellec et al., 2011), and the third pharyngeal pouch in mouse (Gordon et al., 2001). In fish that lack parathyroid glands, gcm2 is expressed in the developing gills (Hogan et al., 2004; Okabe and Graham, 2004).
Purely aquatic amphibians lack parathyroid glands, and in Xenopus the parathyroid glands develop during metamorphosis (Shapiro, 1933; Srivastav et al., 1995). By whole mount in situ hybridization, Xenopus gcm2 was first detected at stage 31 in two vertical stripes in the pharyngeal region, with stronger expression in the most posterior stripe (Fig. 3A,B). At stage 32, gcm2 was also weakly expressed in a third stripe posterior to the first two (Fig. 3C,F). In situ hybridization on longitudinal sections of stage-32 and -34 embryos indicated that these stripes corresponded to gcm2 expression in the endoderm (pouch) and ectoderm (cleft) components of the second, third, and fourth pharyngeal arches (Fig. 3J–M). Later in development, gcm2 expression appeared confined to the developing gills (Fig. 3 G–I), as seen in teleost fish (Hogan et al., 2004; Okabe and Graham, 2004). We also assessed whether gcm2 was expressed in the parathyroid glands after metamorphosis. In 3-month-old post-metamorphic froglets, parathyroid glands were identified in the thoracic cavity based on their position relative to the carotid arteries, as seen on transverse sections (Fig. 3N,O; Shapiro, 1933; Coleman, 1969). We performed real time RT-PCR on dissected tissues from 3-month-old froglets and found that gcm2 was specifically expressed in the parathyroid glands and was virtually undetectable in other tissues such as brain, heart, and thymus (Fig. 3P). This result indicates that gcm2 expression in the parathyroid is conserved across many species.
In mice, the homeobox-containing gene, Hoxa3, is expressed in the third and fourth pharyngeal pouches endoderm and in the NC-derived mesenchyme of the third and fourth pharyngeal arches (Hunt et al., 1991; Manley and Capecchi, 1995, 1998). HOXA3 is essential for pharyngeal glands development as Hoxa3 mutant mouse embryos lack both the thymus and parathyroid (Manley and Capecchi, 1995).
In Xenopus embryos, hoxa3 was expressed at the early neurula stage in the developing spinal cord and migrating NC cells (Fig. 4A,B). hoxa3 showed an anterior boundary of expression corresponding to rhombomere 5 (McNulty et al., 2005). At stage 32, (Fig. 4E,F) and stage 35 (Fig. 4I,J) the anterior boundary of expression was maintained with hoxa3-positive NC cells populating the third pharyngeal arch. In situ hybridization on section at these stages showed that hoxa3 was expressed in the mesenchyme of the third and fourth pharyngeal arches, as well as in the endoderm-derived epithelium of the third and fourth pharyngeal pouches (Fig. 4G,H, K,L). By contrast, the population of NC cells that migrated into the second pharyngeal arch did not express hoxa3 (Fig. 4G,H, K,L). At stage 41, the thymus primordia were exclusively surrounded by hoxa3-negative NC cells (Fig. 4M–P), confirming that the thymus in Xenopus is exclusively derived from the second pharyngeal pouch.
foxn1. The winged helix/forkhead transcription factor Foxn1 (formerly called whn/Hfh11; Kaestner et al., 2000) is highly expressed in the thymus in mouse, birds, and fish (Nehls et al., 1996; Neves et al., 2011; Gordon et al., 2001 Schorpp et al., 2002). FOXN1 is not required for thymus specification (Nehls et al., 1996); however, it is essential later for TEC differentiation in mice (Blackburn et al., 1996).
A PCR product corresponding to Xenopus laevis foxn1 was amplified from stage-55 thymus cDNA. Sequence analysis (Fig. 5A; Table 1) indicated that at the amino acid level, the partial sequence of Xenopus laevis Foxn1 shared 53% identity with human FOXN1 (Schorpp et al., 1997), 53% identity with mouse FOXN1 (Nehls et al., 1996), 57% identity with chicken Foxn1 (Neves et al., 2011), and 42% with zebrafish Foxn1 (Schorpp et al., 2002). Xenopus laevis and Xenopus tropicalis Foxn1 sequences showed a very similar identity to that of other species (Table 1). The forkhead DNA-binding domain shows 100% conservation between human, mouse, chicken, and Xenopus (Fig. 5A). While in most species Foxn1 is expressed early in the pharyngeal pouch endoderm, in Xenopus foxn1 was first detected at stage 47, when the thymus primordia had already detached from the pharyngeal pouch. Whole-mount in situ hybridization shows that foxn1 expression was restricted to the thymus (Fig. 5B). This expression was confirmed by in situ hybridization on sections (Fig. 5C,D). At stage 48, the thymus primordia had increased in size and foxn1 expression appeared to be stronger (Fig. 5E,F). foxn1 can be detected up to stage 57 (the last stage examined in this study). However, at this stage the expression was more diffuse and appeared scattered throughout the entire organ (Fig. 5G), presumably due to the expansion of the lymphoid cells in the thymus (Du Pasquier and Weiss, 1973; Robert and Ohta, 2009).
Table 1. Sequence Homology Comparison of Foxn1 in Different Species
The temporal expression of hoxa3, gcm2, and foxn1 genes was confirmed by RT-PCR (Fig 6). These results are consistent with our in situ hybridization data and further demonstrate the lack of foxn1 expression in the thymus primordium prior to its separation from the pharynx.
NC Contribution to Thymus Development
The cardiac NC is a subdivision of the cranial NC that makes an important contribution to the pharyngeal glands (Le Lievre and Le Douarin, 1975; Bockman and Kirby, 1984). To evaluate whether NC cells contribute to thymus development in Xenopus, we performed transplantation experiments. NC grafts isolated from stage-17 embryos injected with mRNA encoding Red Fluorescent Protein (RFP) were transplanted onto the same region of stage-matched unlabeled embryos (Fig. 7A; Lee and Saint-Jeannet, 2011). At stage 26, the RFP-labeled NC grafts displayed a normal pattern of cranial NC migration in the host embryo, and cells derived from the RFP-labeled NC graft contributed to the hyoid (pharyngeal arch 2), anterior (pharyngeal arch 3), and posterior (pharyngeal arch 4–5) branchial streams of the NC, but not to the mandibular (pharyngeal arch 1) stream of NC (Fig. 7B–E), as previously described (Lee and Saint-Jeannet, 2011).
The progeny of the RFP-labeled NC cells was analyzed in transplanted embryos at stage 41 and 47 (n=16; three independent experiments). On longitudinal sections through the pharyngeal region of stage-41 embryos, RFP-positive cells formed most of the mesenchyme surrounding the thymus primordium, which remained free of NC cells at this stage (Fig. 7F–O). In situ hybridization for hoxa3, which has an anterior boundary of expression in the third pharyngeal arch mesenchyme, indicated that the RFP-labeled NC cells surrounding the thymus primordia were hoxa3-negative, consistent with its derivation from the second pharyngeal pouch. In all cases examined (n=10), transverse sections of stage-47 tadpoles showed that RFP-labeled NC cells populated the thymus primordial visualized by foxn1 expression (Fig. 7P–Y).
The NC Is Not Required for Initial Thymus Development
We next performed NC ablation experiments to evaluate the role of the NC-derived mesenchyme, surrounding the pharyngeal pouch, during initial thymus development. A segment of the NC, corresponding to prospective cardiac NC, was ablated at stage 17 (Fig. 8A, B, E; Lee and Saint-Jeannet, 2011). The successful ablation of the NC was assessed by expression of the NC marker sox10 at stage 25, when the NC cells populated the pharyngeal arches in control embryos (Fig. 8C,D, F,G). In all NC-ablated tadpoles examined at stage 47 (n=13, in three independent experiments), the bilateral thymus primordia formed in their proper location and expressed foxn1 gene (Fig. 8H–M). These results indicate that NC cell–derived signals are not required for the specification and the initial organogenesis of the thymus primordium in Xenopus. To further evaluate the expression levels of foxn1 in control and manipulated embryos, we performed real time RT-PCR on thymi isolated from stage-48 tadpoles. We normalized foxn1 expression levels to the expression of the TEC-specific marker, keratin 8 (Klug et al., 2002), to account for any changes in TEC number between control and NC-ablated samples. We found that foxn1 expression in the NC-ablated thymi was not statistically different from that of control thymi, indicating that NC cells are not required for foxn1 expression in developing TEC (Fig. 8N). Next we determined whether NC cells were needed for the function of TEC to support T-cell development. To address this issue, we analyzed the expression of the recombination-activating gene 1 (rag1) in dissected thymi from stage-48 tadpoles following NC ablation (Greenhalgh et al., 1993). Rag1 is specifically expressed in lymphocytes as they initiate the assembly of the T-cell receptor genes, and is therefore a good marker to assess intrathymic T-cell development (reviewed in Schatz and Swanson, 2011). By real time RT-PCR, we found that rag1 expression in the thymi of NC-ablated tadpoles (n=9 tadpoles) was similar to that of control thymi (Fig. 8O), suggesting that intrathymic T-cell development was not impaired by NC ablation.
In this study, we analyzed the development of the thymus in Xenopus laevis using histological and molecular tools. In agreement with earlier studies (Nieuwkoop and Faber, 1967; Manning and Horton, 1969), we found that the thymus rudiment arises from the second pharyngeal pouch endoderm at around stage 41, and that by stage 45 the rudiment separates from the pharynx to become a fully individualized paired organ. This was documented by the expression of hoxa3, a gene with an anterior boundary of expression in the third pharyngeal arch. While NC cells migrating into the third pharyngeal arch express Hoxa3, the population of NC cells that travels rostral to the otic vesicle into the second pharyngeal arch remains hoxa3-negative. In Xenopus, the thymus primordium is exclusively surrounded by hoxa3-negative NC cells consistent with its derivation from the second pharyngeal pouch. This contrasts with mouse, chicken, and zebrafish embryos in which the thymus primordium originates from the third and/or fourth pharyngeal pouch, and is surrounded by Hoxa3-positive NC cells (Hunt et al., 1991; Manley and Capecchi, 1995, 1998; Saldivar et al., 1996; Hogan et al., 2004; Chen et al., 2010; see Table 2). While in all species examined so far the thymus originates from pharyngeal pouches, its precise embryological origin and the number of thymus glands in the adult vary greatly. For example, sharks develop a thymic primordium in every pouch except the first one, resulting in five bilateral thymi. In reptiles, the thymus arises from the second and third pharyngeal pouches, and in chicken the thymus primordia are located in the third and fourth pharyngeal pouches and give rise to seven thymic lobes (reviewed in Rodewald, 2008; Ge and Zhao, 2012). These differences in the origin of the thymus presumably reflect the specialized characteristics of individual pharyngeal arches acquired during evolution (Grevellec and Tucker, 2010).
Table 2. Summary of the Developmental Expression of Gcm2, Hoxa3, and Foxn1 in Different Speciesa
pam, pharyngeal arch mesenchyme; pce, pharyngeal cleft ectoderm; ppe, pharyngeal pouch endoderm. -, not detected in the pharyngeal apparatus; nd, not determined.
apce expression has been reported in one study (Hogan et al., 2004).
In most species, Hoxa3 is expressed in the NC mesenchyme of the pharyngeal arches as well as in the pharyngeal pouches' endoderm (see Table 2). Hoxa3 mutant mouse embryos lack both the thymus and parathyroid. However, the amount of NC cells migrating in the pharyngeal arches is essentially normal in the mutants (Manley and Capecchi, 1995), suggesting that it is Hoxa3 expression in the pharyngeal pouch endoderm that is critical for thymus formation. This was recently confirmed by a NC-specific deletion of Hoxa3 in mice, which did not affect specification of the thymus and parathyroid, rather both organs failed to properly sever their connection with the pharynx and therefore were ectopically positioned (Chen et al., 2010). Because thymus organogenesis in Hoxa3 null mutants fails after formation of the third pharyngeal pouch and prior to organ specification, it has been proposed that HOXA3 functions to establish the identity of the third pharyngeal pouch to form the thymus and parathyroid territories (Blackburn and Manley, 2004; Manley and Condie, 2010). The fact that the thymus primodium arises from the second pharyngeal pouch endoderm in Xenopus, a tissue that is hoxa3-negative, clearly indicates that thymus organogenesis can occur independently of Hoxa3, raising the question of the precise function of HOXA3 during thymus development in mammals. If HOXA3 is indeed providing positional identity to the third pouch compatible with thymus formation, it is possible that this function may have been co-opted by another Hox gene product in frogs, presumably Hoxa2, which has an anterior boundary of expression in the second pharyngeal arch (Pasqualetti et al., 2000).
In non-aquatic vertebrates, the thymus and parathyroid glands are derived from a common primordium and their segregation into two distinct domains can be first visualized by the expression of two transcription factors, Foxn1 and Gcm2, which are largely thymus- and parathyroid-specific, respectively. Gcm2 is critically required for parathyroid gland formation. It is expressed in the pharyngeal pouches and the forming parathyroid glands (see Table 2), and Gcm2 mutant mice fail to form parathyroid glands (Günther et al., 2000; Liu et al., 2007). While fish and purely aquatic amphibians lack parathyroid glands, Xenopus develops parathyroid glands during metamorphosis (Shapiro, 1933; Srivastav et al., 1995). In Xenopus embryos, gcm2 is initially detected in the second, third, and fourth pharyngeal pouches and clefts and then its expression becomes restricted to the gills. This is consistent with previous reports in teleost fish showing that gcm2 was expressed within the pharyngeal apparatus and internal gill buds (Hogan et al., 2004; Okabe and Graham, 2004; see Table 2). Interestingly, parathyroid hormone-encoding genes were also detected in the gills, suggesting that the tetrapod parathyroid glands and the gills of fish are evolutionarily related and may play a similar role in controlling calcium levels (Okabe and Graham, 2004). Because in post-metamorphic froglets gcm2 is specifically expressed in the parathyroid glands, it is likely that Gcm2 in Xenopus may carry a similar function to that of higher vertebrates (Liu et al., 2007).
The transcription factor Foxn1 is the gene mutated in the nude mice (Nehls et al., 1994; Blackburn et al., 1996). While Foxn1 is not required for initial thymus specification and organogenesis (Nehls et al., 1996), it is one of the earliest genes expressed in the thymus primordium. Foxn1 is first detected in the presumptive thymic epithelium of the third pharyngeal pouch at 72 hpf in zebrafish, and at E11.25 in the mice (Gordon et al., 2001; Schorpp et al., 2002). In birds, it is detected in the dorsal-anterior region of the third and fourth pharyngeal pouch endoderm as early as at E5 (Neves et al., 2011). In these species Foxn1 is detected in the pharyngeal pouch endoderm well before the thymus primordium detaches from the pharynx and starts to differentiate (Gordon et al., 2001; Schorpp et al., 2002; Neves et al., 2011; see Table 2). Interestingly, in Xenopus, foxn1 was not detected until the thymus primordium separated from the pharynx at around stage 47. This is approximately the stage at which the TEC starts to differentiate and express MHC class II molecules, and when RC47-positive leukocytes colonize the thymus (Du Pasquier and Flajnick, 1990). This suggests that initial TEC development in Xenopus is Foxn1-independent.
The NC contributions to thymus development have been somewhat controversial. Initial NC ablation experiments in chicken embryos suggest that thymus organogenesis depends on signals derived from the NC cells surrounding the third and fourth pharyngeal pouches (Bockman and Kirby, 1984). In the mouse mutants Splotch, which have a dramatically reduced number of NC cells in the third pharyngeal pouch, the thymus and parathyroid primordia are normally specified as determined by the expression of Gcm2 and Foxn1 (Griffith et al., 2009). In these mutants, the boundary between the two glands' primordia is shifted such that the presumptive thymic region is expanded at the expense of the parathyroid, resulting in a larger fetal thymus (Griffith et al., 2009). A more recent study suggests that thymus formation in mouse embryos is NC-dependent (Nelms et al., 2011). Compound mutant embryos homozygous for a NC-specific Foxd3 mutation and heterozygous for Pax3, which lack NC cells caudal to the first pharyngeal arch, show severe thymus hypoplasia (Nelms et al., 2011). It is unclear why these two mouse models have somewhat different outcomes with respect to thymus development. Our findings are consistent with the former observations since NC ablation did not affect specification and initial organogenesis of the thymus. Moreover, the thymus primordium expressed foxn1 in the absence of NC cells, suggesting that NC-mesenchyme signals are not required for these processes. We also found that ablation of NC cells did not alter the expression of rag1, an essential factor for thymic T-cell development. Altogether these results indicate that the NC cells are largely dispensable for early thymus development and differentiation in Xenopus.
Although the mechanisms regulating thymus development have been extensively studied in the last decade (reviewed in Blackburn and Manley, 2004; Hollander et al., 2006; Rodewald, 2008; Gordon and Manley, 2011), less is known about the factors that initiate the specification of the thymus primordium. Because Xenopus is an excellent model to analyze cell fate choices during embryogenesis, the work presented here provides the basis for future investigations of thymus development in vertebrates.
Embryos, Injections, Tissues Labeling, and Transplantation
Xenopus laevis embryos were staged according to Nieuwkoop and Faber (2011) and raised in 0.1× NAM (Normal Amphibian Medium; Slack and Forman, 1980). Monomeric Red Fluorescent Protein (RFP) mRNAs were synthesized in vitro using the Message Machine kit (Ambion, Austin, TX). Labeling and transplantation of the NC explants were performed as previously described (Lee and Saint-Jeannet, 2011). Briefly, embryos were injected into the animal pole at the 2-cell stage with 1 ng of RFP mRNA. NC-labeled explants, corresponding to prospective cardiac NC, were dissected at stage 17 in 1× NAM supplemented with 50 μg/ml of gentamycin using a 25-gauge syringe needle. Grafts were then transplanted onto the same region of unlabeled stage-matched sibling embryos. Grafted embryos were allowed to heal for 30 min in the dissection medium, before being transferred for long-term culture into 0.1× NAM. Embryos were observed under an epifluorescence microscope (Nikon, Melville, NY; Eclipse E800).
Isolation of Xenopus laevis foxn1 and gcm2
Xenopus laevis foxn1 was amplified by PCR from stage-55 thymus cDNA generated by the First-Strand cDNA Synthesis Kit (GE Healthcare, Piscataway, NJ). The primers (F: 5′-TTTCATCCGTATAAGAGG-3′; and R: 5′-ATCTGTAAGAGAAGGATT-3′) were designed based on the sequence of Xenopus tropicalis foxn1 (accession no. XM_002937144). These primers were located at position 478 and 1,764 of Xenopus tropicalis sequence. The PCR conditions were as follows: denaturation 94°C (1 min), annealing at 60°C (1 min), and extension at 72°C (1 min) for 35 cycles. A 1,287-bp PCR product was purified, ligated into pGEMTeasy (Promega, Madison, WI), and sequenced. The partial sequence of Xenopus laevis foxn1, which covers 129 amino acids, has been deposited into GeneBank (accession no. JQ182306). Xenopus laevis gcm2 was amplified by PCR from stage-35 cDNA using primers (F: 5′-TCCAAAGATGCCTCAGGACT-3′; and R: 5′-TATATCCGTG CATTGGAGCA-3′) designed based on the published sequence (accession no. AB175676; Okabe and Graham, 2004). The PCR conditions were as follows: denaturation 94°C (1 min), annealing at 55°C (1 min), and extension at 72°C (1 min) for 35 cycles. A 1,100-bp PCR product was purified, ligated into pGEMTeasy (Promega), and sequenced. Xenopus laevis hoxa3 cDNA (NP_001080293) was purchased from Open Biosystems (Thermo Scientific, Swedesboro, NJ).
Total RNAs were extracted from embryos or dissected tissues using an RNeasy micro RNA isolation kit (Qiagen, Valencia, CA). The RNA samples were treated with RNase-free DNase I before RT-PCR. The reactions were performed using the One Step RT-PCR kit (Qiagen) according to the manufacturer's instructions using the following primer sets: hoxa3 (F: 5′-TCCCTACCAAGGAGCAAATG-3′; R: 5′-GCAGGCAGGTCGATGATATT-3′), gcm2 (F: 5′-GATGCCTCAGGACTTC AAGC-3′; R: 5′-CATGGCCCAACCACTTAGAT-3′), foxn1 (F: 5′-ATCCGTGAGA AAAAGCATGG-3′; R: 5′-TAGTGGGGGAGTTCACAAGG-3′), rag1 (F: 5′-GA GGTGGCATTGCCTAATGT-3′; R: 5′-TTCCCTCTGTTTCCTCCTCA-3′); keratin 8 (F: 5′-AGGCTGCAGTCCGAAATAGA-3′; R: 5′-CTCCTTCCAGCAGTTTCCTG-3′), and odc (F: 5′-ACATGGCATTCTCCCTGAAG-3′; R: 5′-TGGTCCCAAGGCTAAAGTTG-3′). The PCR conditions were as follows: denaturation 94°C (30 sec), annealing at 55°C (30 sec), and extension at 72°C (30 sec) for 30 cycles. For real-time RT-PCR, the reaction was performed using the QuantiTect SYBR Green RT-PCR kit (Qiagen) on a LightCycler (Roche Diagnostics, Indianapolis, IN). The reaction mixture consisted of 10 μl of QuantiTect SYBR Green RT-PCR Master Mix, 500 nM forward and reverse primers, 0.2 μl of RT, and 60 ng of template RNA in a total volume of 20 μl. The cycling conditions were as follows: denaturation at 95°C (15 sec), annealing at 55°C (20 sec), and extension at 72°C (15 sec). By optimizing primers and reaction conditions, a single specific product was amplified as confirmed by melting curve analysis. Each reaction included a control without template and a standard curve of serial dilutions (in 10-fold increments) of test RNAs. In each case, ornithine decarboxylase (odc) was used as an internal reference (data not shown). Each bar on the histograms has been normalized to the level of odc.
In Situ Hybridization
Digoxygenin (DIG)-labeled antisense RNA probes (Genius Kit, Roche) were transcribed from linearized plasmids using the indicated RNA polymerase: pGEMT-foxn1 (SacII, SP6 polymerase), pGEMT-gcm2 (SalI, T7 polymerase) and pSport6-hoxa3 (SalI, T7 polymerase). For each gene, sense probes were used as negative control (not shown). Whole-mount in situ hybridization was performed as previously described (Harland, 1991). For in situ hybridization on sections, embryos were fixed in 4% paraformaldehyde in phosphate buffer saline (PBS; Gibco, Gaithersburg, MD) for 1 hr, embedded in Paraplast+ and 12-µm sections hybridized with the appropriate DIG-labeled probes as described (Henry et al. 1996). Because RFP can no longer be visualized following this procedure, sections of RFP-labeled embryos were first individually photographed and then processed for in situ hybridization. Sections were then briefly counterstained with eosin.
Histology and Immunohistochemistry
Embryos were fixed in 4% paraformaldehyde in PBS for 1 hr. After dehydration through a graded series of ethyl-alcohol, embryos were embedded in Paraplast+. Transverse serial sections (12 μm) were performed on an Olympus (Center Valley, PA) rotary microtome and stained with Hematoxylin and Eosin. For immunohistochemistry, after rehydration in PBS, 10-μm sections were boiled for 20 min in sodium citrate buffer (10 mM sodium citrate, 0.05% Tween 20, pH 6.0) to retrieve antigens. The sections were then blocked in PBST (0.1% Triton X-100 in PBS, containing 0.2% bovine serum albumin and 5% inactivated lamb serum) and incubated successively with mouse anti-p63 (Santa Cruz Biotechnology, Santa Cruz, CA; clone 4A4; Senoo et al., 2007) antibody overnight at 4°C (1:50 dilution), and an anti-mouse IgG-FITC conjugated secondary antibody (Jackson ImmunoResearch, West Grove, PA; 1:100 dilution) for 1 hr at room temperature. All antibodies were diluted in PBST.
We are grateful to Dr. Patricia Labosky for comments on the manuscript. We thank Drs. Dominique Alfandari and Helene Cousin for reagents. This work was supported by a grant from the National Institutes of Health to J.-P. S.-J. (R01-DE014212).