Asymmetric Expression of Specific Ephrins and Ephs Is Required at the Dorsal Ectoderm–Mesoderm Boundary

To address the issue of repulsion restriction, we conducted a comprehensive characterization of the ephrin–Eph system in the early gastrula. We first compared quantitatively the transcripts of all ephrins and Ephs in the dorsal ectoderm and in the mesoderm. Both tissues expressed multiple ephrins and Ephs. Although both A and B types are involved at the ectoderm–mesoderm boundary ([5] and unpublished data), the contribution of the A type appears less important, as they are not sufficient on their own to induce separation ([4] and unpublished data). We thus focused on the ephrin B subfamily and their receptors, which showed predominantly asymmetric patterns (Figure S1B). Thus, ephrinB2 and EphA4 were strongly enriched in the mesoderm, whereas the ectoderm selectively accumulated ephrinB3 and EphB2–4. We next wanted to test whether asymmetric expression is functionally relevant for tissue separation. Interestingly, ephrinB2, ephrinB3, and EphA4 start to be expressed just at the onset of gastrulation (Figure S1A), coinciding with the appearance of the ectoderm–mesoderm boundary and the onset of separation behavior [19]. We thus focused on ephrinB2, ephrinB3, EphA4, EphB2, and EphB4 as strongly asymmetrically distributed molecules and included in our analysis ephrinB1 as an example of an evenly expressed ligand (Figure 1C).

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Figure 1. Multiple ephrins and Eph receptors are specifically required for ectoderm–mesoderm separation.

(A) Ephrin–Eph signaling is required for ectoderm–mesoderm separation. Sagittal section of gastrula embryos, injected with antisense MOs. Inserts show detail of the dorsal side. The boundary separating ectoderm (E) from mesoderm (M) is marked by arrowheads. The concave arrowhead indicates the position of bottle cells, and the arrow the anterior edge of the anterior mesoderm. COMO, control morpholinos; EphrinB3 MO, The boundary is largely absent. Arrowheads point to a fuzzy remnant of ectoderm–mesoderm interface. Mesoderm involution is strongly impaired (arrow and concave arrowhead). (B) Diagram of the early Xenopus gastrula and of the in vitro tissue separation assay. In this assay, test explants obtained by dissection from the inner layer of the ectoderm or from the involuting mesoderm (blastopore lip) are placed on an ectoderm substrate. Ectoderm explants rapidly integrate into the substrate, whereas mesoderm aggregates remain well separated. The percentage of explants remaining separate is scored after 45–60 min. (B′) Ectoderm injected with ephrinB3 MO fails to maintain separation and incorporate mesoderm explants (arrows). Arrowheads, explants remaining separated. (C) Asymmetric expression of multiple ephrins and Ephs across the ectoderm–mesoderm boundary. Schematic representation of the relative expression of ephrins and Eph receptors analyzed in this study, based on RT-qPCR (Figure S1A). The same color code is used in all figures. (D) Each ephrin/Eph is specifically required. In vitro tissue separation assay. Depletion of ephrinB1 or ephrinB3 in the ectoderm (eB1MO, eB3MO) or EphA4 in the mesoderm (A4MO) led to inhibition of separation. Separation could be restored by treating directly the surface of the other tissue explant for 15 min with soluble Fc fragments of the corresponding ephrin/Eph molecule. Other ephrins/Ephs failed to rescue, with the exception of eB2, which could rescue eB3 depletion. cFc, control anti-human Fc antibody. Numbers on top indicate ratios of separated explants to total number of explants. p values = Student's t test. (E) The specific role of Eph receptors requires their extracellular domain, but the cytoplasmic tails are interchangeable. Chimeras were designed in which cytoplasmic domains of EphA4 and EphB4 were swapped. Each chimera was tested for the ability to rescue depletion of endogenous EphA4 in the mesoderm or EphB4 in the ectoderm. Depleted Ephs are represented by “ghost” shapes. Separation could be rescued by the constructs that contained the corresponding extracellular domain (AB for EphA4, BA for EphB4), but not by the constructs that contained the cytoplasmic domain. A* and B* were control constructs, which were wild type except for two amino acids within the end of the transmembrane domain, which had to be substituted in AB and BA to produce the chimeric constructs (see Materials and Methods).

https://doi.org/10.1371/journal.pbio.1001955.g001

Although ephrin/Eph depletion severely disrupts the endogenous ectoderm–mesoderm boundary ([4] and Figure 1A), the embryonic phenotype is difficult to interpret, due to multiple functions of ephrins and Ephs in various aspects of gastrulation [5],[20]–[22] (Winklbauer unpublished). We thus performed most of our study on a reconstituted boundary produced by apposition of ectoderm and mesoderm explants (Figure 1B,B′), an assay that allows an in-depth dissection of tissue separation [4],[19],[23]. By systematic depletions using antisense morpholino oligonucleotides (MOs) (Figure S2A,D), we established that each ephrin and each Eph is required, either in the ectoderm, in the mesoderm, or in both. Their depletion inhibited tissue separation to a degree that generally correlated with their relative tissue enrichment (Figures S1B and S2A). Interference with ephrins or with Ephs on one side of the boundary, by single or multiple depletions (Figure S2A) or dominant negative constructs [4], led to a maximal reduction of separation to 30–40%, whereas simultaneous interference on both sides of the boundary led to a significantly stronger inhibition (Figure S2A), consistent with a requirement for two antiparallel forward signals [4].

After having established that each ephrin and Eph subtype is required, we asked next whether a given subtype could be replaced by another member of the family. An ephrin or Eph was depleted, and rescue was attempted by mRNA injection (Figure S2B), or by direct activation at the boundary through incubation with soluble preclustered ephrin or Eph extracellular domains (Figure 1D). Results from both kinds of rescues were in perfect agreement. Subtypes typically failed to substitute for each other, as observed for ephrinB1 and B3, or for EphA4 and B4. The only exception was ephrinB2, which could substitute for ephrinB3 (Figure 1D), a result that will be explained in later experiments. The apparent lack of rescue was not due to a lower activity of a particular construct or Fc-fragment, as rescue of the same subtype was in all cases efficient. This was confirmed by comparing two different amounts of ephrinB1 and B3 mRNAs (Figure S2B) and verifying their expression levels (Figure S2C). The lower amount was sufficient to rescue depletion of the same subtype, but depletion of the other subtype could only be marginally rescued with the highest amount. These results demonstrated an unexpectedly tight, although not absolute, specificity of the requirements for each ephrin and Eph. Note that the ability of the EphA4-Fc fragment to rescue loss of EphA4 also uncovered a contribution from reverse signaling to tissue separation.

The Specificity of Eph Receptors Resides in Their Extracellular Domain

The specific requirement for all ephrinB subtypes and their receptors could be due to specific differences in downstream signaling and hence to different roles during tissue separation. Alternatively, signaling could be uniform and additive, but depend on specific receptor–ligand interactions. To determine whether Eph receptor specificity resided in their cytoplasmic tail or in their extracellular domains, we constructed EphA4/B4 chimeras, where the respective cytoplasmic domains were swapped, which we tested for the ability to substitute for endogenous EphA4 or EphB4 proteins. We verified that these chimeric constructs were properly expressed at the cell surface (Figure S3A) and activated by ephrin-Fc fragments (Figure S3B). The functional assays gave clear-cut results (Figure 1E): The AB chimeric construct, which contained the extracellular domain of EphA4 and the intracellular domain of EphB4, could perfectly rescue the loss of EphA4, but was unable to substitute for EphB4. Reciprocally, the BA construct could rescue EphB4 but not EphA4 depletion. These results showed that the extracellular domains were responsible for the Eph specificity. The cytoplasmic domains appeared interchangeable, suggesting that the requirement for each of these Eph receptors was probably not due to differences in signaling but rather reflected the ability to bind selected ligands. We postulated that specific combinations of receptor–ligand pairs could underlie the restriction of repulsion to the boundary.

Complementarily Expressed Ephrins and Ephs Selectively Interact to Form Functional Pairs

If indeed specific ephrin–Eph pairs formed preferentially at the boundary, we predicted that ectopic addition of ephrins normally enriched in the mesoderm should induce artificial separation of two ectoderm explants. This prediction was fully verified. Indeed, ectoderm explants could be induced to repel each other by treatment with soluble Fc fragments for mesoderm-enriched ephrinB2 but not ephrinB1 (Figure 2A). Similarly, the separation of two mesoderm explants was efficiently induced by incubation with ephrinB3, which is normally expressed only in the ectoderm, but not by ephrinB2 (Figure 2A′). Ectoderm–ectoderm separation could also be induced by “mesodermal” EphA4-Fc, but not by EphB4-Fc (Figure 2A″ and A″′).

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Figure 2. Characterization of ephrin–Eph specificity.

(A) Identification of functional cognate receptors. The in vitro separation assay was performed using two explants from the same tissue (ectoderm–ectoderm or mesoderm–mesoderm). Under normal conditions these explants mix completely. Separation could be induced when these explants were exposed to soluble Fc fragments corresponding to ephrins or Ephs normally enriched in the other tissue (see Figure 1C), thus mimicking the endogenous asymmetric ephrin/Eph expression. The endogenous functional partners were then identified by depleting single candidate ephrins/Ephs in the receiving explant and determining which of them was required for ectopic separation. Note that in this set of experiments, Fc fragments were present during the assay, ensuring that both explants were continuously exposed. Separation can also be induced by treating only one explant, although the penetrance is lower [4]. (A) Separation of ectoderm explants was induced by soluble Fc fragments corresponding to “mesoderm-specific” ephrinB2, but not ephrinB1, already endogenously enriched in the ectoderm. EphrinB2-Fc–induced separation was strongly inhibited by EphB4MO and more weakly by EphB2MO. (A′) Mesoderm aggregates were tested on large mesoderm layers artificially produced in the animal pole of the embryo (see Materials and Methods). Separation was induced by Fc fragments of “ectoderm-specific” ephrinB3, but not ephrinB2. Ephrin3-induced separation was reversed by EphA4MO but not by EphB4MO (red arrow). (A″) Ectopic separation between ectoderm explants was induced specifically by “mesodermal” EphA4 but not by “ectodermal” EphB4. Separation was inhibited only by ephrinB3MO (red arrow). (A″′) Examples of mixing of ectoderm explants incubated with control Fc (arrows) and of separation of ectoderm explants treated with EphA4-Fc (arrowheads). (B) Biochemical analysis of differential activation of Eph receptors by ephrin ligands. Ectoderm explants were treated with 40 nM ephrinB1, B2, B3 Fc fragments or with control anti-Fc antibodies for 1 h, and then lysed. Endogenous EphB2 and EphB4, and EphA4 were immunoprecipitated and analyzed by immunoblotting for phospho-tyrosine and for total Eph levels. In the case of EphA4, which is only expressed at low levels in the ectoderm, EphA4-GFP was ectopically expressed. EphA4 was strongly phosphorylated in response to both ephrinB2 and ephrinB3, but not ephrinB1, which gave levels similar to negative controls. EphB2 responded strongly to ephrinB2, and only weakly to ephrinB1. EphB4 was highly phosphorylated in response to ephrinB2, whereas activation by ephrinB1 or ephrinB3 was negligible. IgG, control immunoprecipitation with nonimmune IgGs. (C) Summary of the main ectoderm and mesoderm-enriched ephrins/Ephs and of the preferred functional interactions.

https://doi.org/10.1371/journal.pbio.1001955.g002

The sensitivity of ectoderm and mesoderm to respond to a subset of ephrin/Eph fragments implied that each of these two tissues must specifically express respective partners. We used the induction of ectopic separation as a functional assay to identify these endogenous receptors. Explants depleted of single ephrins or Ephs were tested for their ability to separate upon incubation with an Fc fragment. Among potential candidate receptors for ephrinB2 in the ectoderm, EphB4 depletion strongly inhibited ephrinB2-Fc–induced separation, whereas EphB2 depletion had a weaker effect (Figure 2A). The ability of ephrinB3 to induce mesoderm separation was entirely dependent on the presence of EphA4 (Figure 2A′). Finally, of all the three ephrin ligands expressed in the ectoderm, ephrinB3 was clearly the one responsible for EphA4-Fc–induced separation (Figure 2A″). Although some of these results were consistent with the relative mRNA enrichments of the various ligands/receptors (Figure S1B), others clearly implied functional selectivity. For instance, in EphA4-induced ectoderm–ectoderm separation, an explanation based only on relative expression levels and under promiscuous binding could explain the minimal role of ephrinB2, which is scarce in the ectoderm, compared to abundantly expressed ephrinB3. However, this explanation would predict a much stronger effect of ephrinB1 depletion, as the latter is present at significant levels. Thus, the best explanation was that ephrinB3 and EphA4 tended to interact preferentially.

We directly assessed functional selectivity at the level of Eph activation (Figure 2B). After treatment of ectoderm explants with equal concentrations of ephrinB1, B2, or B3 Fc fragments, each Eph receptor was immunoprecipitated, and the levels of phosphorylation were monitored by Western blot using an anti–phospho-tyrosine antibody [11]. Because of poor immunoprecipitation of the endogenous EphA4 protein with available antibodies, EphA4-YFP was ectopically expressed and immunoprecipitated with an anti-GFP antibody. In these experiments, cells were stimulated for 30 min, a time that should be sufficient to reach a bona fide steady state in terms of repulsive behavior, as separation can be induced already after a few minutes [4]. The results were clear cut: EphA4 was highly phosphorylated in response to ephrinB2 or ephrinB3 but not ephrinB1. EphB2 responded strongly to ephrinB2 and weakly to ephrinB1. EphB4 was activated by ephrinB2 but neither ephrinB1 nor ephrinB3. These results correlate well with ephrin–Eph affinities measured in vitro [13], with the notable exception of ephrinB1–EphB2, for which the affinity was reported to be similar to ephrinB2–EphB2 [24]. Combined with the asymmetric distribution of the various molecules, the differences in binding largely explained our functional data. For instance, the failure of EphB4 to induce ectoderm separation was due to the low levels of ephrinB2 in this tissue, which is its only strong interactor. On the other hand, the ectoderm could respond to EphA4 via ephrinB3, but not ephrinB1.

Taken together, our results show that each Eph receptor is selective for one or at most two ephrins. These results support the hypothesis that tissue separation is driven by asymmetric expression of specific receptor–ligand pairs. Most of the ephrin/Eph pairs identified functionally show indeed partially complementary expression (Figure 2C and Figure S1B). This is clearly the case for ectodermal ephrinB3 and mesodermaly enriched EphA4. Likewise, ectodermaly enriched EphB2 and EphB4 interact best with mesodermaly enriched ephrinB2. However, not all factors are expressed in simple complementary patterns. In particular, EphA4 interacts not only with ectodermal ephrinB3 but equally well with ephrinB2, which is abundant in the mesoderm. In contrast, ephrinB1, evenly expressed in both tissues, can only weakly activate EphB2, which is enriched in the ectoderm. Note that ephrinB1 depletion in either tissue had a rather strong phenotype (Figure 1D and Figure 2A) considering that it seemed to only interact weakly with EphB2. This may be indicative of the occurrence of an additional receptor for ephrinB1 yet to be identified (see also supplementary discussion in Text S1).

Complementary Expression of Specific Ephrin–Eph Pairs Is Sufficient to Account for Tissue Separation

Our results suggested that tissue separation may be simply explained by the complementary expression of selective ephrin/Eph pairs, which would generate an excess of repulsive signal at the boundary (Figure 3A). Nevertheless, it is conceivable that particular ephrins and Ephs would also be specifically needed on one or the other side of the boundary. We examined this possibility for the ephrinB3–EphA4 pair, which is closest to a fully complementary expression pattern. We depleted ephrinB3 in the ectoderm and EphA4 in the mesoderm and asked whether separation could be rescued by ectopic re-expression of these two molecules in the opposite tissues. The results were unambiguous: Swapping ephrinB3 and EphA4 efficiently restored separation (Figure 3A′). The same result was obtained when ephrinB3 was provided as a soluble ligand to the surface of the ectoderm (Figure 3B′). Thus, the presence of ephrinB3 and EphA4, respectively, in the ectoderm and the mesoderm is not required for separation; the complementary expression appears sufficient.

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Figure 3. Ectoderm–mesoderm separation relies on asymmetric expression of specific ephrin/Eph pairs across the boundary, irrespective of the direction of the asymmetry.

(A, A′, A″) Reciprocal replacement of mesodermal ephrinB3 and ectodermal EphA (mRNA injection). (A, A′) Diagram describing the experiment. (A) Endogenous ephrinB3 and EphA4 were depleted in the mesoderm and ectoderm, respectively (ghost labels). (A′) EphrinB3 (or B2) was then overexpressed in the ectoderm and EphA4 (or B4) in the mesoderm, thus effectively swapping the ligand and receptor (red double arrow). (A″) Quantification. Swapping ephrinB3 and EphA4 efficiently restored separation. EphrinB3 could be replaced by ephrinB2, consistent with the latter also being a ligand for EphA4. However, EphA4 could not be substituted by EphB4, in agreement with EphA4 being the only receptor of ephrinB3. The weak nonsignificant rescue was likely due to a slight boost in the ephrinB2–EphB4 signal. (B, B′) Similar experiment, but with the ephrin ligand substituted by direct incubation of the receiving explant with the corresponding soluble Fc fragment. EphrinB3 and EphA4 were depleted, EphA4 was overexpressed in the ectoderm, and ectoderm explants were incubated with the indicated Fc fragments. Overexpression of EphA4 led to a partial rescue of separation (control Fc), likely by activating ephrinB2–EphA4 signaling across the boundary (red dashed double arrow). Incubation with ephrinB3-Fc fully rescued separation. EphrinB2-Fc but not the ephrinB1-Fc fragment could also rescue, in agreement with the selectivity of EphA4.

https://doi.org/10.1371/journal.pbio.1001955.g003

We tested the specificity of the process by attempting the rescue with other ephrins or Ephs. EphrinB2, which is another good partner for EphA4, could substitute for ephrinB3 (Figure 3A′,B′). No rescue was observed with ephrinB1, which only activates EphB2, nor with EphB4, which cannot function as an ephrinB3 receptor (Figure 3A″,B′).

Separation Requires the Kinase Activity of Eph Receptors

Although EphA4 and B4 cytoplasmic tails appear interchangeable (Figure 1E), we also know that this tail is required for Eph function in separation [4]. We thus asked whether the kinase activity was involved by using kinase dead (KD) variants, where the ATP binding site was mutated [25],[26]. We found that both EphA4KD and EphB4KD failed to rescue depletion of the corresponding endogenous receptors (Figure S4A). On the contrary, they acted as dominant negatives, decreasing levels of receptor phosphorylation (Figure S4D) and inhibiting separation (Figure S4A) to similar levels as MO depletion or expression of cytoplasmic truncated forms (Figure S2 and [4]). This inhibition was fully rescued by co-expression of wild-type EphA4/B4 (Figure S4A). We further tested the effect of these KD mutants in two other situations—that is, (a) in ectopic separation of two ectoderm, or of two mesoderm explants by soluble ephrinB2, respectively, B3 Fc fragments (as in Figure 2A), or (b) in rescues of ephrinB2/B3 depletions by the corresponding Fc fragments (as in Figure 1D). In both types of experiments, expression of the EphA4KD blocked the action of ephrinB3 Fc, and EphB4KD that of ephrinB2 (Figure S4C). Thus, these KD forms behaved in all cases as strong dominant-negative mutants, demonstrating that their kinase activity is essential for ectoderm–mesoderm separation.

Enhanced Eph Signaling and Myosin Activation at the Ectoderm–Mesoderm Boundary

Immunostaining of sections from wild-type gastrulae with an antibody recognizing a conserved phosphorylated site present in all EphBs demonstrated that Eph signaling was indeed activated in both ectoderm and mesoderm, but was significantly stronger at the boundary (Figure 4A,A′). A similarly increased signal was observed with an anti–phospho-EphA antibody (Figure 4A′). We confirmed biochemically the existence of basal signaling in the tissues and enhanced activity at ectoderm–mesoderm contacts. Such contacts were maximized by mixing dissociated ectoderm and mesoderm cells to produce heterogeneous aggregates (Figure 4B). The levels of phosphorylated EphAs and EphBs in extracts of mixed aggregates were compared with those of pure ectoderm and mesoderm aggregates. Mixed aggregates showed higher p-EphA and p-EphB signals than combined homogenous ectoderm and mesoderm aggregates. We further showed that EphA phosphorylation in these ectoderm–mesoderm aggregates required ephrinB3, but not ephrinB1, further confirming the specificity of EphA4 (Figure 2D′). Thus, high local Eph activation is consistent with cell repulsion being restricted to the boundary, due to the preferential interactions between complementary pairs of ephrins and Ephs enriched on opposite sides of the boundary.

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Figure 4. Increased Eph receptor and myosin activation at the ectoderm–mesoderm contacts.

(A) Detection of phospho-EphB by immunofluorescence of a sagittal section from wild-type early gastrula embryo. The upper panel shows a general view of the dorsal region, and the lower panel an enlarged view of the ectoderm–mesoderm boundary. (A′) Quantification of relative signal intensity (arbitrary units) measured for phospho-EphB and phospho-EphA at cell–cell contacts along the boundary and inside each tissue. Averages from five embryos for phospho-EphA and nine embryos for phospho-EphB. p values = Student t test. (B, B′) Biochemical comparison of Eph phosphorylation levels between homogenous tissue aggregates and mixed ectoderm mesoderm aggregates. (B) Schematic description of the experiment. Dissociated ectoderm and mesoderm cells were mixed and left to reaggregate for 30 min, which produced a maximal number of “heterotypic” ectoderm–mesoderm contacts mimicking contacts at the boundary. Homogenates from these mixed aggregates (E/M mix) were compared to the same amount of cells assembled into separate ectoderm and mesoderm aggregates, thus forming only “homotypic” contacts, and combined during homogeneization (E+M ctrl). (B′) Western blots were probed for total and phospho-EphA, phospho-EphB, total EphA, and EphB. Mixed aggregates showed a reproducibly higher level of p-Eph signals (arrows), which indicates stronger activation at boundary contacts. Arrowheads, nonspecific bands. GAPDH was used as loading control. (C, C′, C″) Selective accumulation of p-MLC along the ectoderm–mesoderm boundary and its dependence on ephrin/Eph signaling. Ectoderm and mesoderm explants were combined, incubated for 1 h, and fixed. Cryosections were immunostained for p-MLC. (C) Normal boundary (underlined by a dashed line) between wild-type tissues. p-MLC levels are higher in the ectoderm than the mesoderm, but highest along the boundary (arrows). Arrowheads point to p-MLC signal along membranes within each tissue. (C′) Loss of p-MLC staining at the tissue interface upon Eph depletion: EphB4-depleted ectoderm and EphA4-depleted mesoderm showed largely unchanged tissue staining, but p-MLC was prominently missing from the fused interface, delineated by the dashed line. (C″) Quantification of p-MLC signal intensity at cell–cell contacts at the boundary and inside the tissues in control and Eph morpholino conditions. Average from nine embryos. p values = Student t test.

https://doi.org/10.1371/journal.pbio.1001955.g004

The typical mechanical output of Ephrin/Eph signaling in repulsive behavior involves myosin-based contraction [27],[28]. We determined the distribution of phosphorylated myosin light chain (p-MLC) in the ectoderm and mesoderm tissues and at their interface (Figure 4C,C″). We observed p-MLC at cell–cell contacts within each tissue (arrowheads). The signal was significantly stronger in the ectoderm than in the mesoderm, but by far the most intense signal was consistently found in patches along the boundary (arrows), as expected from strong bursts of ephrin/Eph signaling at this interface. EphA4/EphB4 depletion strongly decreased p-MLC staining at the boundary and in the mesoderm, but not in the ectoderm (Figure 4C′,C″). These results confirm that the boundary is a site of significantly high contractile activity that largely depends on Eph signaling.

Separation Is Controlled by the Balance Between Ephrin–Eph-Dependent Repulsion and Cadherin-Mediated Adhesion

Significant Ephrin–Eph signaling also takes place within each of the two tissues (Figure 3A,B). To account for the fact that the tissues remained coherent and overt cell detachments occurred only at the interface, we considered the role of cell–cell adhesion. We proposed that cohesion or separation is determined by a balance between cadherin adhesion and Eph signaling-dependent contractility. Although cadherin levels are lower in the most anterior mesendoderm ([29]–[31] and unpublished data), they are quite similar between the ectoderm and the mesoderm analyzed in our experiments (Figure S5B), implying that the balance would mostly depend on the strength of Eph signaling. This signaling would take place at all cell contacts, but only at the boundary would the signal be sufficiently intense to overcome adhesion, thus causing cell detachment.

This model predicted that increasing cadherin levels or decreasing contractility should inhibit detachments between ectoderm and mesoderm cells, while on the contrary decreasing cell adhesion or increasing Eph signaling should lead to visible detachments between mesoderm cells. This hypothesis was consistent with the observation that ectoderm–mesoderm separation was impaired upon cadherin overexpression, which could be rescued by boosting Eph signaling across the tissue interface by incubation with soluble ephrin Fc fragments (Figure S5A). Note that Eph depletion did not significantly affect cadherin levels (Figure S5B), arguing against a direct regulation of cadherins by ephrin–Eph signaling.

To explore the behavior at individual homotypic and heterotypic cell–cell contacts, we juxtaposed single cells obtained by dissociation of early gastrula tissues (Figure 5A–I and Movies S1–S7). Our dissociation conditions fully preserved both the capacity of cells of the same tissue to rapidly re-establish stable adhesions (Figure 5D–G) and the ability for contacts between ectoderm and mesoderm cells to reproduce the alternating cycles of attachment/detachment characteristic of the separation behavior (Figure 5A–B and Movie S1). We first tested the effect of increasing cadherin levels on the normal repulsion between ectoderm and mesoderm cells. We also tested the effect of incubating the cells with the myosin inhibitor blebbistatin. Both conditions severely reduced the frequency of detachments (Figure 5B,C,H and Movies S2 and S3), consistent with the inhibition of tissue separation observed under the same manipulations (Figure S5A).

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Figure 5. Tissue separation is controlled by a balance between ephrin/Eph-mediated repulsion and cadherin adhesion.

(A–C) Ectoderm–mesoderm repulsion requires myosin activity and is antagonized by cadherin adhesion. Selected frames from time lapse confocal movies (Materials and Methods) showing dynamics of cell–cell contacts between single embryonic cells. Cells dissociated from ectoderm and mesoderm tissues were plated on glass coated with low amounts of fibronectin (see Materials and Methods). Ectoderm cells expressed membrane-GFP, and mesoderm cells membrane-Cherry. (A) Wild-type ectoderm and mesoderm cells stably attached to cells from the same tissue, but contacts between ectoderm and mesoderm cells exhibited cycles of attachments (arrowheads) and detachments (arrows) recapitulating the separation behavior observed at the boundary between the two tissues. (B) After treatment with 100 µM blebbistatin, most ectoderm mesoderm contacts remained stable. (C) C-cadherin overexpression in both ectoderm and mesoderm strongly decreased detachments. Note that mesoderm cells tended then to surround ectoderm cells. (D–G) Evidence for subthreshold levels of Ephrin/Eph-mediated repulsive signals between mesoderm cells. (D) Mesoderm cells (here control MO-injected) established stable contacts (arrowheads) that were maintained (concave arrowheads) throughout the duration of the recording (1 h). (E) Cadherin-depleted mesoderm cells (cadherin MO) showed frequent figures of redetachments (arrows), indicating the existence of repulsive signals. (F) Stable contacts between cadherin-depleted mesoderm cells were rescued by simultaneous Eph depletion, demonstrating that the repulsion observed between mesoderm cells was due to ephrin–Eph signaling. (G) Detachment between mesoderm cells could be induced by increased ephrin–Eph-mediated repulsion through ectopic expression of ephrinB3 and EphB4, the “preferred” ectoderm partners for “mesodermal” EphA4 and ephrinB2. Frame numbers of the corresponding movies are indicated. (H) Quantification of the rate of attachment/detachment per cell per hour. Numbers indicate number of events per number of cells analyzed.

https://doi.org/10.1371/journal.pbio.1001955.g005

Because we expected mild repulsion between mesoderm cells, we predicted that detachments should become more prominent if adhesion would be experimentally decreased. We subjected mesoderm cells to a mild cadherin depletion (∼30%, not shown) and observed that contacts were now much less stable and cells displayed the typical repulsive behavior normally observed at contacts with ectoderm cells (Figure 5E,H and Movies S4 and S5). These retractions were entirely dependent on intact ephrin/Eph signaling (Figure 5F,H and Movie S6). We also predicted that mesoderm cells could become repellent even with normal cadherin levels if repulsive signals would be increased. We chose to express ephrinB3 and EphB4, because they are normally enriched in the ectoderm, and are the respective specific partners for mesoderm-enriched EphA4 and ephrinB2 (Figure 2C). EphrinB3/EphB4-expressing mesoderm cells readily showed strong repulsion (Figure 5G,H and Movie S7). This balance between repulsion forces and cell–cell adhesion was also observed at the tissue level in reaggregation assays (Figure S5C–E), in which diminished cohesion induced by cadherin depletion was rescued by co-depletion of EphB4, whereas ectopic expression of ephrinB3 and EphB4 decreased cohesion of cells with wild-type cadherin levels.

Simulating the Ephrin/Eph Network Reveals the Robustness and the Limits of the Model

Our results suggested that separation could be explained by the asymmetric expression of a subset of specific ephrin–Eph pairs, which resulted in widespread ephrin–Eph signaling, but with an altogether higher output across the boundary. Although the concept is intuitively coherent, the actual system is complex, with many ephrins and Ephs, including pairs that could interact extensively within one tissue (e.g., ephrinB2–EphA4 in the mesoderm). To better estimate the validity of this model, we simulated it by computing the contribution of all the various ephrin–Eph interactions established at homotypic contacts within the tissues and heterotypic contacts between tissues (Figure S7). We considered a minimal model taking into account two parameters: (a) the affinities between the extracellular domains of ephrins and Ephs and (b) their relative expression in the different tissues (Text S1 and Figure S7). Affinities for the various ephrin–Eph pairs have been measured in vitro [13]. These values remain imprecise, but were globally consistent with our data, with few exceptions (e.g., ephrinB1–EphB2; see Text S1). Endogenous levels for the different ephrin and Eph proteins could not be measured directly, due to lack of adequate antibodies, but were approximated based on relative mRNA levels determined by real-time PCR (Figure S1B) and on a global estimate obtained indirectly (see Text S1 and Figure S6). Note that these “apparent” affinities and concentrations are here purely operational terms. The actual system is hugely complex, constituted of multiple ligands and receptors, all of them membrane proteins, capable of spontaneous or ligand-induced clustering, and potentially also of cis-interactions [15],[32]–[37]. Most of these additional parameters and their impact on ephrin–Eph function are still ill-defined, and a formal description of such a system is a largely unresolved problem.

However, this simple model where receptor activation depends on its own concentration, the concentrations of its various potential ligands, and the corresponding affinities turned out to describe with surprising accuracy the situation at this boundary. The signal output turned out indeed to be clearly highest at the tissue boundary compared to ectoderm–ectoderm and mesoderm–mesoderm contacts (Figure S7D). The system appeared very robust, predicting a highest signal at the boundary over a broad range of ephrin–Eph apparent concentrations and of apparent KDs (Figure S7D). Quite strikingly, the result of the simulation closely resembled the relative levels of phosphorylated receptors in our immunofluorescence images (Figure S7D and E, reproduced from Figure 4A). The model was further tested for the ability to predict the outcome of loss- and gain-of-function experiments (Figure S7F–J). The simulation recapitulated very well the major characteristics of the system. In particular, the specific induction of “separation” between ectoderm cells by activation with mesoderm-enriched ephrinB2 or EphA4, the inhibition of normal separation upon ephrin or Eph depletions (Figure S7G,H), and most importantly the fact that rescue could not be obtained indifferently by any ephrin (or Eph) (Figure S7H) argued against widespread ligand–receptor promiscuity. It also accounted for our original observation that strong inhibition of separation required interfering with receptors on both sides of the boundary (Figure S7J) (see [4] and Figure S2A).

We conclude that a system of partially complementary and semiselective ephrin–Eph pairs is indeed sufficient to explain ectoderm–mesoderm separation. The robustness of the simulation provides strong support for the general principles of this model. Although multiple additional factors are likely to impact on the apparent concentrations and affinities, on the response curves of Eph activation, and more downstream on the contractile activity of boundary cells, such factors may not affect significantly the final global pattern. The reason for this robustness appears to be the very limited set of pairs that can effectively interact and thus influence the system, and the fact that three out of five of these pairs are asymmetrically expressed (ephrinB3–EphA4, ephrinB2–EphB2, and ephrinB2–B4). The simulation indicated that, for these five pairs, differences up to 10-fold in apparent affinities would change the relative strength of the outputs, but the signal remained strongest at the boundary under most conditions. The few aspects of our experimental data that were not well simulated are discussed in Text S1. Discrepancies were certainly expected considering the many additional mechanisms that can modulate ephrin–Eph signaling. Nevertheless, the predictions of the simulation were surprisingly good given the simplicity of the assumptions. We conclude that a combination of multiple semiselective pairs is sufficient to explain a large part of the system's behavior.

Selective Ephrin–Eph Pairs at the Ventral Ectoderm–Mesoderm and at the Notochord Boundaries

We wondered whether the principles found for the early ectoderm–mesoderm boundary would similarly apply to other boundaries. Ephrin and Eph expression is indeed widespread in the gastrula embryo, and two other important boundaries form during this phase. A ventral boundary appears at midgastrula to separate the ectoderm for the ventral mesoderm (Figure 6B), which is known to have properties different from those of dorsal axial mesoderm. At the end of gastrulation, the dorsal mesoderm is partitioned into the axial notochord and lateral paraxial mesoderm (prospective somitic mesoderm) (Figure 6C). In the latter case, we recently showed that ephrins and Ephs were indeed involved in separation [38]. We decided to systematically analyze the patterns of ephrin/Eph expression in these two regions (Figure S1B). We found that the same set of ephrins and Ephs was expressed in those regions. The expression patterns revealed common themes as well as some significant differences (summarized in the diagrams of Figure 6). In the case of the ventral boundary, the ephrinB3–EphA4 complementarity and the mesoderm enrichment of ephrinB2 were preserved. However, EphB4, which was enriched in the ectoderm on the dorsal side, was now homogenously expressed, whereas on the contrary ephrinB1, equal on both sides of the dorsal boundary, was here strongly asymmetric, now fully complementary to its receptor EphB2. In the late gastrula, the two new structures emerging from the dorsal mesoderm showed very interesting expression patterns: The notochord had preserved strong EphA4 and ephrinB2 expression, which were the major characteristics of the early mesoderm. The paraxial mesoderm, however, had dramatically modified its ephrin/Eph expression, acquiring typical features of ectoderm—that is, low EphA4 and high EphB4. Note that the complementarity was strong for Eph receptors, but rather mild for the ephrins. However, a similar trend was observed, where the notochord remained more “mesoderm-like” (higher ephrinB2, lower ephinB3), and the paraxial mesoderm was now more “ectoderm-like” (lower ephrinB2, higher ephinB3).

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Figure 6. Schematic representation of the three boundaries forming during Xenopus gastrulation and simplified diagrams of the preferential ephrin–Eph pairs.

(A) Dorsal ectoderm–mesoderm boundary. (B) Ventral ectoderm–mesoderm boundary. (C) Notochord–paraxial mesoderm boundary. The top drawings represent sagittal sections of the corresponding stages and highlight the two tissues forming the corresponding boundary. At stage 14, the boundaries between the notochord (no) and the paraxial mesoderm (pm) form perpendicular to the plane of the section. Cross-section is shown in insert. ar, archenteron roof; ne, neuroderm. In the lower diagrams, the position of the boxes representing each molecule symbolizes its general distribution: A box placed on one side of the boundary corresponds to strong asymmetric distribution. Weakly asymmetric or homogenous distributed molecules are drawn overlapping the boundary. The double arrows link the functional pairs. The patterns at the dorsal and the ventral boundaries are similar, with two prominent differences: EphrinB1 shifts from equal in the two dorsal tissues to more mesodermal in the ventral side. EphB4, on the contrary, shifts from mainly ectodermal to equally distributed. At the end of gastrulation, the dorsal mesoderm experiences significant changes: Compared to the earlier dorsal mesoderm, the notochord (no) keeps EphA4 and ephrinB2, but the paraxial mesoderm (pm) loses EphA4 and acquires ephrinB3 and EphB4, which were until then typically ectodermal. Note that the major asymmetries concern EphA4 and B4; the other components are only slightly enriched in one or the other sides of the boundary.

https://doi.org/10.1371/journal.pbio.1001955.g006

These observations suggested that tissues that became separated by boundaries expressed some sort of “modules” characterized in particular in all cases by complementary ephrinB3–EphA4 expression, but had some flexibility for other components. We then tested the role of key players, at both boundaries. In the case of the ventral boundary, we used the classical separation assay, attempting both loss- and gain-of-function experiments (Figure S8). All the results were in perfect agreement with the expression patterns and with our model. Similar to the dorsal boundary, depletion of either the ectoderm-enriched ephrinB3 or of its mesoderm partner EphA4 inhibited separation. In addition, ephrinB1 depletion in the mesoderm, which had only a weak effect on the dorsal side, affected here separation much more drastically (Figure S8B). Depletion in the ectoderm of its receptor EphB2 similarly inhibited separation.

In gain-of-function experiments, both mesoderm-enriched ephrinB1 and B2 could induce robust separation between two ectoderm explants when added as soluble Fc fragments (Figure S8C), an effect that could not be obtained with ephrinB1 on the dorsal side (Figure 2A). We used this phenotype to identify the functional partner of ephrinB1 on the ectoderm surface. We found that EphB2, but not EphB4, depletion significantly inhibited separation, functionally validating our biochemical results (Figure 2B).

The role of ephrins and Ephs on the notochord boundary was examined by targeted depletion in a restricted region of the embryo [38]. The effect on the integrity of the boundary was examined on sections of whole embryos (Figure 7). We tested depletion of the two strongly asymmetric Eph receptors, A4 and B4, and of their ligands ephrinB2 and B3. We observed in each case strong disruption of the boundary. However, each ephrin/Eph MO disrupted the boundary only when targeted to the expressing tissue: ephrinB3 MO and EphB4 MO in the paraxial mesoderm, EphA4 MO in the notochord. EphrinB2 MO was the only one that had an effect on both sides. These phenotypes were perfectly consistent with the distribution and partial specificity of these molecules. For the receptors, which were strongly asymmetrically expressed, MO injection obviously had an effect only when targeted to the tissue where they were enriched. For the ligands, the results were explained by their specificity. Indeed, the only receptor for ephrinB3 is EphA4, enriched in the notochord; thus, despite the poor complementarity of ephrinB3 distribution, its depletion only impaired separation when targeted to the opposite tissue (paraxial mesoderm) (Figure 7D,E,L). EphrinB2, however, can stimulate both EphA4 and EphB4 and was thus predicted to significantly contribute to generate repulsion at both sides of the boundary, explaining the widespread effect of its depletion (Figure 7L). We further verified that this functional selectivity was purely due to extracellular interactions: We used the EphA4 and B4 chimeras to rescue disruption of the notochord boundary upon EphA4 or EphB4 depletion. In both cases, rescue was only achieved by expression of the chimeric construct that harbored the proper extracellular domain (Figure 7F–K,M).

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Figure 7. Partially complementarily expressed ephrins and Ephs control notochord–paraxial mesoderm separation.

(A) Summary of ephrin and Eph expression and of the functional pairs (double arrows) in the dorsal mesoderm at stage 14 (see Figure S1B). (B–M) Manipulations were targeted to a restricted region of the dorsal mesoderm. Embryos were fixed at stage 14, and the dorsal structure was analyzed on sections. Injected cells were detected by the tracer Myc-GFP (red). Only the strongest signal is visible on these images but is sufficient to indicate the position of the injected area. Membranes were labeled with an anticadherin antibody (green). In (D) and (E), FoxA4-positive nuclei appear in green–cyan. FoxA4 is used as the notochord marker. (B) Control. Normal boundaries (highlighted by dashed lines) were characterized by a smooth alignment of the notochord and paraxial mesoderm cells. (C) Strong boundary disruption in an ephrinB2 MO-injected embryo. The position of the left boundary can still be located (arrowheads), but the alignment is jagged (arrowhead). On the right side, the two tissues are continuous, without a detectable boundary. Arrows point to the approximate limit of the notochord. (D and E) The boundary is effectively disrupted by ephrinB3 MO targeted to the paraxial mesoderm (arrows in D), but not to the notochord (E). (F–H) EphA4 MO injection in the notochord, the tissue fused with the paraxial mesoderm (F, arrows). A boundary can be rescued by the coinjection of mRNA coding for the AB Eph chimera (G, dashed lines) but not by the BA chimera (H, boundaries lack on both sides, arrows). Thus, the extracellular domain of EphA4 is necessary and sufficient for its function in the notochord. (I) Inhibition of separation by EphB4 MO targeted to the paraxial mesoderm. Left boundary is missing (arrows). (J) Rescue by the BA chimera. The injection was targeted to the right boundary. (K) The AB chimera fails to rescue. The injection was targeted to the left side (arrows). The right boundary is intact (dashed line). (L) Quantification of boundary disruption by ephrin/Eph depletion. Individual boundaries were scored as follows: 1, intact boundary; 0.5, partially disrupted boundary (rare cases); 0, fully disrupted boundary. The first column (t) compiles data of all embryos, (no) and (pm) the number of boundaries where injection was mainly targeted to the notochord or to the paraxial mesoderm. Numbers on top are numbers of embryos (2–6 independent experiments). EphrinB2, which can interact with both EphA in the notochord and EphB4 in the paraxial mesoderm, was required in both tissues. EphrinB3 and EphB4 depletion strongly disturbed the boundary when targeted to the paraxial mesoderm, but had no effect in the notochord. The opposite was observed for EphA4, consistent with the expression patterns and the selective interactions. (M) Quantification of rescues by wild type and chimera forms of EphA4 and EphB4. In all cases, complete rescue was obtained with the corresponding wild-type proteins and with the chimera containing the correct extracellular domain. The nature of the intracellular domain was indifferent. Numbers on top indicate total number of boundaries in each category.

https://doi.org/10.1371/journal.pbio.1001955.g007

Ethics Statement

All animal studies were approved by the McGill University Animal Care Committee (permit 4869 “Cellular mechanisms of embryonic boundary formation”) and the University of Toronto Animal Care Committee (permit 20010074 “Analysis of gastrulation movements in Xenopus”).

Antibodies

Affinity-purified antibodies specific for EphB4 and EphB2 were generous gifts from Dr. Elena Pasquale (Burnham Medical Institute, [48]). Anti–phospho-EphB4, anti–phospho-EphA4, and anti-EphA4 antibodies were gifts from Dr. Greenberg [49]. Mouse anti-EphA4 was purchased from BD, pan anti-mouse ephrinB antibody from Zymed, mouse anti-alpha tubulin from Cell Signalling, mouse anti-GAPDH from ABi, mouse and rabbit anti-GFP from Invitrogen, affinity-purified polyclonal goat EphB4 from R&D, and mouse anti–phospho-tyrosine PY-20 HRP-conjugated from Santa Cruz.

Recombinant Proteins

Recombinant mouse ephrinB2-Fc, ephrinB1-Fc, human ephrinB3, and mouse EphB4-Fc and EphA4-Fc (R&D Systems) comprising the extracellular domain of ephrin/Eph fused to C-terminal 6× histidin-tagged Fc region of human IgG were preclustered by 1 h incubation with anti-human Fc antibody (Jackson ImmunoResearch Laboratories) at a 1∶2 ratio in MBSH [4] before application. Anti-human Fc IgGs alone were used as control.

Plasmids and Constructs

Membrane-targeted GFP and Cherry and Xenopus ephrinB1 and ephrinB2 were described previously [4]. Xenopus EphB4 and ephrinB3 in pCS2+ were gifts from Dr. A. Brändli [50]. Xenopus EphA4 (Pagliaccio) in pBluescript KS by DR Bob Winning [51] was a gift from Dr. T. Sargent. Chicken EphA4-GFP in peGFPN2 was a gift from Dr. A. Kania [52]. EphA4 was subcloned into the pCS2+and C-terminally fused to eYFP. Mouse EphA4 Y928F KD mutant in pCS2+ was a gift from Dr. Ira Daar [25]. Xenopus EphB4 KD mutant was constructed by substituting arginine to lysine at position 645 (ATP-binding) using the QuickChange site-directed mutagenesis kit. Alk4*, a constitutively active Activin receptor, was a gift from Dr. J. Smith [53]. β-catenin in pSP36T was a gift from Drs. P. McCrea and B.M. Gumbiner [54].

To construct Eph chimeras with swapped cytoplasmic domains, a restriction site was introduced at the end of the transmembrane domain in each original receptor (Nhe1 for EphB4 and Spe1 for EphA4) by site-directed mutagenesis (Quick Change XII, Stratagene). The two resulting mutants, called EphA4* and EphB4*, had, respectively, a change from bp1926 tgtcat to actagt and from bp 1763 ggtggt to gctagc corresponding to a V to L and V to A substitution corresponding to the last hydrophobic amino acid of the transmembrane domain amino acid 558–559 of Xenopus EphB4 and EphA4. EphA4* was subcloned into pCS2+ for consistency with EphB4. EphA4* and EphB4* in pCS2+ rescued loss of the corresponding endogenous Ephs with the same efficiency as the original A4 and B4 constructs (Figure 1E). The newly introduced Spe1 and Nhe1 sites were used to cut and exchange the fragments corresponding to the sequences of the cytoplasmic tails, yielding EphA4B4 (extracellular and transmembrane domains of A4 and cytoplasmic tail of B4) and EphB4A4 (extracellular and transmembrane domains of B4 and cytoplasmic tail of A4).

Injections

mRNAs were synthesized according to the manufacturer's instructions (mMessage mMachine kit, Ambion). MOs and mRNAs were injected animally in the two blastomeres of two-cell stage embryos to target the ectoderm, and equatorially in the two dorsal blastomeres of four-cell stage embryos to target the mesoderm. MO sequences. The amounts of MO and of mRNA injected are listed in the Text S1.

In Vitro Separation Assay

mRNA was injected animally at the two-cell stage for ectoderm expression and dorsally at the four-cell stage for dorsal mesoderm expression. Dissections and assays were performed in Modified Barth Solution (MBSH) containing: 88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 0.82 mM MgSO4, 0.33 mM Ca(NO3)2, 0.41 mM CaCl2, 10 mM Hepes, adjusted to pH 7.4 with NaOH and supplemented with 10 µg/ml streptomycin sulfate and penicillin. For the standard assay, explants were dissected at stage 10+ (early gastrula). Ectoderm or mesoderm aggregates were laid on ectoderm caps, and the degree of separation was scored as the percentage of aggregates that did not incorporate into the cap after 45–60 min incubation [31]. In some cases, animal caps were induced into mesoderm by injection of 120 pg β-catenin and 1 ng constitutively active Activin receptor mRNAs.

Separation of ventral tissue was similarly performed using stage 11 ectoderm and mesoderm from the ventral lip. For in vitro activation using soluble ephrins/Ephs, explants were either preincubated with preclustered ephrinB/Eph-Fc fragments (40 nM unless specified otherwise) in MBSH for 15 min at room temperature (e.g., Figure 1D) or the entire assay was performed in the presence of Fc fragments (e.g., Figure 2A). For the statistical analysis, results were compared using the two-tailed paired-sample Student's t test.

Reaggregation Assay

Cells from dissected mesoderm and inner layer ectoderm were dissociated in alkaline calcium-free buffer (88 mM NaCl, 1 mM KCl, 10 mM NaHCO3, pH 9.3). For reaggregation, cells were transferred to agarose-coated Petri dishes containing MBSH and incubated for 1 h under mild rotation (10 rpm) on an orbital shaker. Images were taken under a dissecting microscope at a12× magnification using a Micropublished RTV3.3 camera (Qimaging) and were analyzed for object size using ImageJ software. Two parameters were measured: average object area and area/perimeter ratio. Results of six independent experiments were normalized using wild-type ectoderm or mesoderm as reference.

Immunofluorescence

Wild-type embryos were fixed at stage 10.5, and sagittal cryosections were prepared as described [55],[56]. Sections were stained with anti–phospho-EphB antibody and Alexa488-coupled anti-rabbit IgG (Invitrogen). Images were collected with a DMIRE2 epifluorescence microscope (Leica) equipped with a 20×/0.70IMM Corr CS oil immersion objective and an ORCA-ER camera (Hamamatsu Photonics), controlled with Metamorph software.

Live Imaging

Tissues expressing membrane-targeted Cherry or GFP as well as various mRNAs/MOs were dissociated in dissociation buffer, cells were transferred to glass-bottom petri dishes (Fluoro dish, World Precision Instruments) coated with 0.01 mg/ml Fibronectin, and cell behavior was filmed for 2 h. Cells were imaged with a WaveFX spinning disc confocal (Quorum Technologies) mounted on an automated DMI6000B Leica microscope, controlled with Volocity 3DM software (Improvision), using a 40× HCX PL APO CS, NA = 1.25 oil objective. Images were acquired every 2 to 5 min using an EM CCD 512×512 BT camera. Image processing was performed with Metamorph (Universal Imaging Corporation) and Adobe Photoshop7 software. Processing consisted of merging two to three planes from z stacks, assigning pseudo-colors, and adjusting image contrast.

Analysis of Eph and p-Eph Levels by Western Blot

Cells from 14 ectoderm- and mesoderm-dissected explants were dissociated and an equal amount of cells were reaggregated either as mixed ectoderm/mesoderm aggregates, or as separate ectoderm and mesoderm aggregates for 1 h on agarose-coated plates in 1× MBSH. Mixed aggregates formed both homotypic ectoderm–ectoderm and mesoderm–mesoderm contacts as well as heterotypic ectoderm–mesoderm contacts, which mimicked the contacts at the boundary. The reaggregation time was set to maximize heterotypic contacts. Separate ectoderm and mesoderm aggregates, which formed only homotypic contacts, were combined for extraction, thus yielding the same amount of material as the mixed aggregates, and served as control. Extraction was performed in 1% NP40 containing buffer as previously described [46]. Extracts were probed by Western blot for EphA, EphB, p-EphA, and p-EphB.

Immunoprecipitation

Ectoderm tissues were dissected from wild-type 40 embryos (for EphB2 and EphB4 immunoprecipiation) or from 40 embryos injected with EphA4-YFP mRNA (250 pg per blastomere) for EphA4 immunoprecipitation. Dissected tissues were treated for 40 min with an equal concentration (40 nM) of EphrinB1/B2/B3-Fc fragments, or control goat anti-human Fc. Tissues were extracted in 10 mM Hepes, 150 mM Nacl, 2 mM EDTA, 1% NP40, supplemented with protease and phosphatase inhibitors [57]. Cleared lysates were incubated for 4–5 h with rabbit anti-EphB2, anti-EphB4, or anti-GFP (for EphA4-YFP), followed by 1 h incubation with protein A-Sepharose beads (Thermo Pierce) at 4°C. The beads were washed four times with 10 mM Hepes, 150 mM NaCl, 2 mM EDTA, 1% NP40, phosphatase inhibitors+0.5% sodium deoxycholate, 0.1% SDS. Immunoprecipitates were analyzed by Western blot for phospho-Tyrosine as well as for total EphA4, EphB2, and EphB4.

RT-PCR and qPCR

RT-PCR was performed using mRNA extracted from whole-stage 9–12 embryos. Loading was equalized by comparing levels of FGFR. Two independent experiments showed identical temporal patterns of expression. RT-qPCR was performed using mRNA extracted from ectoderm and dorsal mesoderm at stage 10.5. qPCR was carried out using CFX 96 Thermo cycler (Biorad). The PCR reactions were set up using 5 µl of RT (20 to 50 times diluted) with 5 µl of SYBR green (Biorad) ½ dilution, 5 µl of 3× PCR-MgCl₂ buffer (Invitrogen), and 5 µl of a 6µM solution of primers. Cycling conditions were as follows: 94°C for 15 s, and 58°C for 30 s, 72.0°C for 1 min. Quantification was based on a dilution series (five fold steps) of the whole embryo RT. Relative expression levels were normalized as ratio to ODC, a ubiquitous gene with homogenous distribution in Xenopus embryos. The sequences of the primers are listed in Text S1.