Mesodermal patterning in the amphibian embryo has been extensively studied in its dorsal aspects, whereas little is known regarding its ventrolateral regionalization due to a lack of specific molecular markers for derivatives of this type of mesoderm. Since smooth muscles (SM) are thought to arise from lateral plate mesoderm, we have analyzed the expression of an - actin isoform specific for SM with regard to mesoderm patterning. Using an antibody directed against -SM actin that recognized specifically this actin isoform in Xenopus, we have found that the expression of -SM actin is restricted to visceral and vascular SM with a transient expression in the heart. The overall expression of the -SM actin appears restricted to the ventral aspects of the differentiating embryo. -SM actin expression appears to be activated following mesoderm induction in animal cap derivatives. Moreover, at the gastrula stage, SM precursor cells are regionalized since they will only differentiate from ventrolateral marginal zone explants. Using the animal cap assay, we have found that -SM actin expression is specifically induced in treated animal cap with bFGF or a low concentration of XTC-MIF, which induce ventral structures, but not with a high concentration of XTC-MIF, which induces dorsal structures. Altogether, these results establish that -SM actin is a reliable marker for ven-trolateral mesoderm. We discuss the importance of this novel marker in studying mesoderm regionalization.

Cell-cell interactions are of major importance in the generation of cell-type diversity in the vertebrate embryo. One of the best documented examples of such interactive events is the control of muscle lineage determination in the amphibian embryo (reviewed in Gurdon, 1987, 1992; Woodland, 1989). Presumptive muscle cells arise from the equatorial region of the blastula under the control of inductive signals which emanate from the prospective endoderm (Nieuwkoop, 1969; reviewed in Smith, 1989). It is believed that mesoderm formation in Xenopus requires three sig-nalling pathways (Smith and Slack, 1983). Two signals derived from the dorsal and the ventral areas of the vegetal hemisphere specify, respectively, the dorsal mesoderm (organizer) and the “general” ventral mesoderm. Part of the ventral mesoderm adjacent to the organizer region then receives a dorsalizing signal which converts it to somitic and intermediate mesoderms, whereas the most remote tissue remains ventral (Smith and Slack, 1983; reviewed in Woodland, 1989).

Recent interest in mesoderm induction stemmed from the discovery that several peptide growth factors (PGFs), either of the transforming growth factor β (TGF-β) family or of the fibroblast growth factor (FGF) family, are able to mimic the action of prospective endoderm in the induction of mesodermal derivatives in blastula animal caps (reviewed in Withman and Melton, 1989; Dawid et al., 1990; Green and Smith, 1991; Jessell and Melton, 1992). Members of the TGF-β family, such as activin, can induce dorsal and anterior mesoderm (Rosa et al., 1988; Smith et al., 1990; van den Eijnden-Van Raaij, 1990; Sokol et al., 1990; Thomsen et al., 1990), whereas members of the FGF family, such as basic FGF (bFGF), can induce ventrolateral and posterior mesoderm (Slack et al., 1987; Kimelman and Kirschner, 1987; Amaya et al., 1991). Such factors can determine the balance of tissue types in a dose-dependent manner, suggesting that they might act as morphogens (Green and Smith, 1990, 1991).

The dorsal mesoderm gives rise to axial structures of the embryo such as the notochord and skeletal muscle. Specific markers are available to characterize the various structures derived from dorsal mesoderm, such as a keratan sulfate which is specific for the notochord (Zanetti et al., 1985), α-cardiac actin (Gurdon et al., 1985), Myo D (Hopwood et al., 1989), Xhox3 (Ruiz i Altaba and Melton, 1989), Brachyury T (Smith et al., 1991) and a 102×103Mr contractile apparatus-associated protein (Kintner and Brockes, 1984) for skeletal muscle differentiation. Since dorsal mesoderm possesses neural-inducing capacity, N-CAM can be used as indirect evidence of dorsal mesoderm formation (Kintner and Melton, 1987). Ventrolateral mesoderm forms the lateral plate mesoderm, which is thought to participate with other tissues in the formation of the heart and smooth muscles. The heart results from the fusion of the lateral plates in the ventral midline and smooth muscles derive from the splanchnic layer, which forms an outer coat around the primary epithelial lining of hollow internal organs (Carl-son, 1981). In contrast to the dorsal mesoderm, there are no molecular markers available for derivatives of ventro-lateral mesoderm, which are currently identified on the basis of morphological criteria such as the lack of axial structures or the presence of differentiated mesothelium and red blood cells (Boterenbrood and Nieuwkoop, 1973; Dale et al., 1985; Green et al., 1990). The availability of molecular markers would allow unambiguous definition of the derivatives of this type of mesoderm, which would be essential tools for a better understanding of the molecular mechanisms underlying mesodermal patterning.

Since smooth muscles (SM) are thought to arise from the lateral plate mesoderm, we made use of an anti-α-SM actin antibody (Skalli et al., 1986) to visualize the expression of this molecule during Xenopus laevis development with regard to early mesoderm regionalization. The pattern of expression of this molecule indicate that α-SM actin can be used as a reliable marker of ventrolateral mesoderm.

Animals

Adult Xenopus laevis were obtained from the “Service d’élevage de Xénope” of the CNRS (Montpellier). Mature oocytes were stripped from females injected 12 hours earlier with 1000 i.u. of chorionic gonadotropin (Sigma) and immediately fertilized with minced testis. Embryos were reared in Normal Amphibian Medium (NAM) 0.1× (Slack, 1985) and staged according to Nieuwkoop and Faber (1967).

Dissection of embryos

Embryos were dejellied by a 2- to 3-minute treatment in 2% cysteine pH 7.8. The vitelline membrane was removed using fine forceps in NAM 1×. Embryos at stage 8 and stage 11 were dissected with platinum threads. Explants were then individually cultured in NAM 0.5× containing penicillin and streptomycin on 1% agar-coated wells until control embryos reached appropriate stages, and were then fixed for immunohistochemistry. Isolated animal and ventral parts from stage-8 embryos were cultured in isolation or in conjugate as previously described (Gurdon et al., 1985). Animal caps isolated at stage 8 were treated for 1 hour either with XTC-GTX-11 (van den Eijden-Van Raaij et al., 1990) -conditioned medium (a gift of Dr van den Eijden-Van Raaij, Utrecht, The Netherlands) used undiluted and at 1:3 dilution, or with recombinant bFGF (a gift of Dr Prats, Toulouse, France) at 50 ng ml−1 and 200 ng ml−1 dilution. Controls were untreated animal caps. Dorsal, ventral and lateral mesoderm, endoderm and animal caps from stage-11 embryos were excised as previously reported (Dale and Slack, 1987).

Antibody

The characterization of the monoclonal antibody against a synthetic decapeptide corresponding to the NH2-terminal peptide of mammalian α-SM actin has been described elsewhere (Skalli et al., 1986). This antibody was purchased from Sigma (France) and used at a 1:1000 dilution for western blots and 1:400 dilution for immunofluorescence.

Western blots

Adult tissues were homogenized in PBS containing 0.5% NP40 and a mixture of protease inhibitors (phenylmethylsulfonyl fluo-ride, aprotinin, iodoacetamide). Homogenates were centrifuged at 10,000 revs/minute for 5 minutes. An aliquot of the supernatant was mixed with Laemmli sample buffer, boiled for 3 minutes, resolved by 10% SDS-PAGE (Laemmli, 1970) and electrotrans-ferred on nitrocellulose (Towbin et al., 1979). Nitrocellulose sheets were blocked with TBS containing 3% BSA for 3 hours and then incubated overnight with the anti-α-SM actin antibody. After extensive washings immunoreactivity was revealed using 125I-anti-mouse antibody (Amersham) followed by autoradiography.

Immunohistochemistry

Paraffin sections were prepared using a previously published procedure (Levi et al., 1987). Briefly, embryos and explants were fixed in isopentane, cooled in liquid nitrogen and transferred successively in methanol equilibrated at −80°C, −20°C, 4°C and room temperature. Embryos were then embedded in Paraplast and 10 μm sections collected on a glass slide. Sections were deparaf-finized in xylene, rehydrated and incubated overnight with primary antibody in PBS containing 5% fetal calf serum (FCS). Primary antibody was revealed by successive incubation with biotinylated anti-mouse IgG (Amersham) and streptavidin-FITC (Amersham). Slides were then mounted in MOWIOL 4-88 (Hoechst) and observed with an epifluorescence Leitz microscope.

Characterization of the anti-α -SM actin antibody

In this study, we have used a monoclonal antibody directed against the 10-amino acid peptide of the NH2 terminal region of mammalian α-SM actin (Skalli et al., 1986). Although this sequence appears to be very specific for this actin isoform (Vandekerckhove and Weber, 1979, 1981), it was important to demonstrate that this antibody did not recognize other members of the actin family in Xenopus. This was carried out on western blots of adult Xenopus laevis tissues. It appears on western blots (Fig. 1) that the anti-α-SM actin antibody reacts with a major band of apparent relative molecular mass 45×103 in extracts of aorta (lane 4), intestine (lane 8), skin (lane 9), lung (lane 3) and bladder (lane 10), but does not react with extracts of brain (lane 5), heart (lane 6) and skeletal muscles (lanes 1, 2). The liver (lane 7) gives a faint signal as a result of its vascularization. The intestine and the bladder give additional bands with lower relative molecular masses, which might possibly represent degradation products (lanes 8, 10).

Fig. 1.

Western blot analysis of extracts from Xenopus adult tissues. Skeletal muscles of the legs (lane 1) and of the back (lane 2), brain (lane 5), and heart (lane 6) do not present any reactivity to the anti-α-SM actin antibody. Lung (lane 3), aorta (lane 4), skin (lane 9), and liver to a lesser extent (lane 7), display a single band with an apparent relative molecular mass of 45 × 103. Intestine (lane 8) and bladder (lane 10) present also a major band at 45×103 and additional bands with lower relative molecular masses, which are probably degradation products. Relative molecular masses of markers are respectively, 106, 80, 49.5 and 32.5 ×103.

Fig. 1.

Western blot analysis of extracts from Xenopus adult tissues. Skeletal muscles of the legs (lane 1) and of the back (lane 2), brain (lane 5), and heart (lane 6) do not present any reactivity to the anti-α-SM actin antibody. Lung (lane 3), aorta (lane 4), skin (lane 9), and liver to a lesser extent (lane 7), display a single band with an apparent relative molecular mass of 45 × 103. Intestine (lane 8) and bladder (lane 10) present also a major band at 45×103 and additional bands with lower relative molecular masses, which are probably degradation products. Relative molecular masses of markers are respectively, 106, 80, 49.5 and 32.5 ×103.

These observations, together with immunohistochemistry on adult tissue sections (data not shown) clearly demonstrate the specificity of this antibody for adult Xenopus SM-containing tissues; α-SM actin expression is in perfect correlation with the patterns previously reported in other species (Vandekerckhove and Weber, 1979, 1981).

Developmental expression of -SM actin

Paraffin sections of Xenopus embryos at different stages of development were stained with the anti-α-SM actin antibody. The developmental pattern of expression of α-SM actin is summarized in Fig. 2A. The first expression of α-SM actin was detected at stage 37/38, in the developing heart (Fig. 2C). This expression in the heart was limited to the atrium, reached a maximum by stage 43 (Fig. 2G-H) and then decreased progressively until stage 46 and was then undetectable. Starting at stage 40, some immunoreac-tivity could be detected in cells that line the endodermal mass. At stage 41, the endodermal mass is clearly delineated by a layer of α-SM actin-positive cells (Fig. 2D). At stage 43, the anterior region of the embryo presents a staining confined to the gut loops whereas more posterior regions of the embryo present a positive uniform layer of cells underlining the whole undifferentiated endodermal mass (Fig. 2F). As differentiation proceeds, the twists and turns of the intestine are all surrounded by a strongly positive layer of cells (Fig. 2I-J). From the beginning of vascular SM differentiation, aorta (Fig. 2K) and large vessels in the cephalic region (not shown) displayed an intense α-SM actin immunoreactivity, whereas small vessels in the caudal region were not found to express this marker. The only other tissues to be stained were the lungs (Fig. 2K), the bladder and the iris (not shown). All other tissues did not express α-SM actin at any time; in particular, skeletal muscles were negative from the onset of differentiation to the latest stages that were studied (premetamorphic, stage 50 embryo).

Fig. 2.

Developmental expression of α-SM actin expression. (A) Schematic representation of the developmental distribution of α-SM actin. From stage 37/38 to stage 46, α-SM actin is detected in the heart. From stage 40 and on, α-SM actin expression is predominently expressed in vascular and visceral SM cells. (B) At stage 23, no labeling is detected in the lateral plate (arrowheads). (C) By stage 37/38, a faint labeling is observed in the heart anlage (h), just beneath the pharyngial cavity (ph). (D) Stage-41 embryo displaying an α-SM actin immunoreactivity confined to the outer layer of the endoderm, where the first intestinal loop starts to differentiate. Two sections in the anterior (E) and posterior (F) regions of an embryo at stage 43 present a staining delineating the differentiating endoderm. (G) At the same stage heart is also labeled (arrowhead). (H) Enlargement of the heart area shown in (G); staining is restricted to the atrium (a) whereas ventricule (v) remains negative. (I,J) Stage-47 embryo where labeling is detected in the outer coat of the gut loops in anterior (I) and more posterior (J) regions. (K) Detail of a stage-47 embryo stained with the α-SM actin antibody, aorta (ao), lung (lu) and intestine (in) present a strong immunoreactivity whereas liver (li), notochord (n) and somites (s) do not. Magnification: B, D-G and I-J ×40; C, H and K ×80.

Fig. 2.

Developmental expression of α-SM actin expression. (A) Schematic representation of the developmental distribution of α-SM actin. From stage 37/38 to stage 46, α-SM actin is detected in the heart. From stage 40 and on, α-SM actin expression is predominently expressed in vascular and visceral SM cells. (B) At stage 23, no labeling is detected in the lateral plate (arrowheads). (C) By stage 37/38, a faint labeling is observed in the heart anlage (h), just beneath the pharyngial cavity (ph). (D) Stage-41 embryo displaying an α-SM actin immunoreactivity confined to the outer layer of the endoderm, where the first intestinal loop starts to differentiate. Two sections in the anterior (E) and posterior (F) regions of an embryo at stage 43 present a staining delineating the differentiating endoderm. (G) At the same stage heart is also labeled (arrowhead). (H) Enlargement of the heart area shown in (G); staining is restricted to the atrium (a) whereas ventricule (v) remains negative. (I,J) Stage-47 embryo where labeling is detected in the outer coat of the gut loops in anterior (I) and more posterior (J) regions. (K) Detail of a stage-47 embryo stained with the α-SM actin antibody, aorta (ao), lung (lu) and intestine (in) present a strong immunoreactivity whereas liver (li), notochord (n) and somites (s) do not. Magnification: B, D-G and I-J ×40; C, H and K ×80.

These results demonstrate that the overall expression of this SM marker is restricted to the ventral aspects of the differentiating embryo.

α-SM actin expression is activated following mesoderm induction

By the stage-8 blastula, isolated animal and vegetal parts do not form mesoderm, but when combined in a conjugate, vegetal tissues induce the animal cap cells to form mesoderm (Nieuwkoop, 1969). We used this experimental procedure to determine if the α-SM actin gene was activated by this inductive interaction. Animal and vegetal parts were isolated at stage 8 and cultured in isolation or as conjugates. Explants were fixed and sectioned for immunostain-ing after 5 days of culture (equivalent stage 47) to ascertain that α-SM actin expression did not reflect its transient expression in the heart. Neither animal cap (Fig. 3A,B) nor vegetal tissue (Fig. 3C,D) expressed α-SM actin when cultured in isolation. When both tissues were combined, α-SM actin expression was observed in animal cap derivatives (Fig. 3E,F). α-SM actin was expressed in a very small group of cells that were widely scattered in the different sections of conjugates and always located in the vicinity of the endodermal mass (Fig. 3E,F). A time course of induction of α-SM actin expression in conjugates was carried out to determine the earliest expression of this marker. It appears that α-SM actin can be detected after 3 days of culture in approximately 53% of the conjugates analyzed (n= 15).

Fig. 3.

Expression of α-SM actin in stage-8 animal cap, vegetal pole and conjugates. (A) Blastula-isolated animal caps do not present any immunoreactivity with the α-SM actin antibody. (B) Corresponding phase contrast. (C) Vegetal pole cells are also negative for this antibody. (D) Corresponding phase contrast. (E,F) When both tissues are combined as conjugates, an α-SM actin immunoreactivity can be observed in the animal cap derivatives (arrowheads). Magnification: A-I, ×50.

Fig. 3.

Expression of α-SM actin in stage-8 animal cap, vegetal pole and conjugates. (A) Blastula-isolated animal caps do not present any immunoreactivity with the α-SM actin antibody. (B) Corresponding phase contrast. (C) Vegetal pole cells are also negative for this antibody. (D) Corresponding phase contrast. (E,F) When both tissues are combined as conjugates, an α-SM actin immunoreactivity can be observed in the animal cap derivatives (arrowheads). Magnification: A-I, ×50.

Thus, α-SM actin presents all the characteristics of a marker activated by mesoderm induction. In conjugate experiments, it can be detected as soon as 3 days of culture, i.e. at an equivalent stage 40/41, in perfect correlation with its first expression in the gut in vivo.

α-SM actin expression is regionalized to ventrolateral marginal zone explants

Regional differences in the inductive specificity of the vegetal hemisphere (prospective endoderm) are known to be responsible for the regionalization of mesoderm in the early embryo (Dale and Slack, 1987). In order to determine if SM precursor cells are already regionalized after mesoderm induction, explants of different areas of stage-11 (gastrula) embryos were isolated, cultured in vitro until control embryos reached stage 47 and analyzed for α-SM actin immunoreactivity. Stage-11 embryos were chosen since this is a stage at which the dorsal side of the embryo can be identified without ambiguity by the appearance of the dorsal blastopore lip. The results are summarized in Table 1. The animal pole region, referred to as a disc of tissue in the centre of the pigmented hemisphere, revealed no immunoreactivity to the anti-α-SM actin antibody (Fig. 4A,B) in all cases examined (n=10). Similarly, endodermal explants (n=7) were never found to express the SM marker (Fig. 4C,D). Dorsal mesoderm explants in 22 out of 24 cases were also found to be negative for this SM marker (Fig. 4E,F). As in previous reports (Dale et al., 1985, 1987), these explants gave rise, among other tissues, to notochord easily recognizable by its vacuolar structure (Fig. 4F). α-SM actin expression was only observed (cf. Table 1) in explants stemming from the ventral (Fig. 4G,H) and lateral (Fig. 4I-L) marginal zone, the latter with a greater efficiency (77%). α-SM actin-positive cells detected in ventral and lateral marginal zone explants were either associated with the epidermal layer and delimiting the cavities formed within the explants (Fig. 4L), or as patches of positive cells (Fig. 4G,I,J).

Table 1.

α-SM actin expression in stage 11 isolated explants

α-SM actin expression in stage 11 isolated explants
α-SM actin expression in stage 11 isolated explants
Fig. 4.

Immunodetection of α-SM actin in explants isolated at stage 11 and cultured for 5 days. Animal caps (A) and endodermal tissuesm (C) do not present any immunoreactivity to the anti-α-SM actin antibody; (B,D) corresponding phase contrast. (E) Explants derived from the dorsal marginal zone, in which notochord (n) differentiate, are also negative to this antibody; (F) corresponding phase contrast. (G) Ventral marginal zone-derived explants express α-SM actin in patches of cells in the centre of the explant; (H) corresponding phase contrast. (I-L) Lateral marginal zone-derived explants present also an immunoreactivity to the α-SM actin antibody, either as a cluster of cells within the explant (I,J) or as a layer of cells associated with the differentiated epidermis (L). (K) Phase contrast of (J). Magnification: A-K, × 50; L, × 100.

Fig. 4.

Immunodetection of α-SM actin in explants isolated at stage 11 and cultured for 5 days. Animal caps (A) and endodermal tissuesm (C) do not present any immunoreactivity to the anti-α-SM actin antibody; (B,D) corresponding phase contrast. (E) Explants derived from the dorsal marginal zone, in which notochord (n) differentiate, are also negative to this antibody; (F) corresponding phase contrast. (G) Ventral marginal zone-derived explants express α-SM actin in patches of cells in the centre of the explant; (H) corresponding phase contrast. (I-L) Lateral marginal zone-derived explants present also an immunoreactivity to the α-SM actin antibody, either as a cluster of cells within the explant (I,J) or as a layer of cells associated with the differentiated epidermis (L). (K) Phase contrast of (J). Magnification: A-K, × 50; L, × 100.

It appears then that mesoderm at the early gastrula stage is already regionalized in its ability to give rise to α-SM actin-expressing cells; only ventral and lateral explants of the marginal zone possess this potential.

ρ-SM actin expression is specifically induced by PGFs

Using the animal cap assay, we determined which types of PGFs were able to induce α-SM actin expression. We expected that PGFs of the FGF family at low or high concentrations and of the TGF-β family at low concentration, which induce ventrolateral-type mesoderm (Slack et al., 1987; Green et al., 1990), could induce this actin isoform, whereas PGFs of the TGF-β family at high concentration, which induce dorsal-type mesoderm (Green et al., 1990), would not. We made use in this study of recombinant bFGF at 50 ng ml−1 and 200 ng ml−1, and of conditioned medium of a cell clone derived from XTC cell line: XTC-GTX-11, which is known to produce activin-like activity (van den Eijnden-Van Raaij et al., 1990). The results are summarized in Table 2. Untreated stage-8 animal cap (n=26) did not express α-SM actin (Fig. 5A,B). In the presence of undiluted XTC-GTX-11-conditioned medium, animal cap explant presented classical extension (Jones and Woodland, 1987) and, in the 21 cases examined, 20 did not present any α-SM actin immunoreactivity (Fig. 5C,D). Notochord was frequently observed (Fig. 5D) together with skeletal muscle. When XTC-GTX-11-conditioned medium was used at a 1:3 dilution, among the 18 explants analyzed, 16 expressed strong α-SM actin immunoreactivity (Fig. 5E,F). Those explants displayed the typical morphology of ventrally induced animal cap with a large vesicle delimited by epidermis, α-SM actin-expressing cells constituting a layer commonly designated as mesothelium. bFGF-treated animal caps, at both concentrations (50 and 200 ng ml−1), and in all cases examined (n=10 and n=19 respectively), exhibited a strong expression of α-SM actin, located in the vicinity of the epidermal layer (Fig. 5G,H). bFGF-treated caps presented also the characteristic morphology of ventrally induced explants.

Table 2.

α-SM actin expression in stage-8 animal caps treated with peptide growth factors

α-SM actin expression in stage-8 animal caps treated with peptide growth factors
α-SM actin expression in stage-8 animal caps treated with peptide growth factors
Fig. 5.

Immunodetection of α-SM actin in stage-8 animal caps treated with peptide growth factors. (A) Untreated animal caps are negative to the anti-α-SM actin antibody; (B) corresponding phase contrast. (C) Animal caps treated with undiluted XTC-GTX-11-conditioned medium display induction of axial structures such as notochord (n) but do not express α-SM actin; (D) corresponding phase contrast. (E) Animal caps treated with XTC-GTX-11 conditioned medium at a 1:3 dilution present strong α-SM actin immunoreactivity; staining is associated with a layer of cells commonly designated as mesothelium in ventrally induced explants; (F) corresponding phase contrast. (G) Animal caps treated with bFGF at 200 ng ml-1 are intensely labeled with the anti-α-SM actin antibody; (H) corresponding phase contrast. Magnification: A-H, × 60.

Fig. 5.

Immunodetection of α-SM actin in stage-8 animal caps treated with peptide growth factors. (A) Untreated animal caps are negative to the anti-α-SM actin antibody; (B) corresponding phase contrast. (C) Animal caps treated with undiluted XTC-GTX-11-conditioned medium display induction of axial structures such as notochord (n) but do not express α-SM actin; (D) corresponding phase contrast. (E) Animal caps treated with XTC-GTX-11 conditioned medium at a 1:3 dilution present strong α-SM actin immunoreactivity; staining is associated with a layer of cells commonly designated as mesothelium in ventrally induced explants; (F) corresponding phase contrast. (G) Animal caps treated with bFGF at 200 ng ml-1 are intensely labeled with the anti-α-SM actin antibody; (H) corresponding phase contrast. Magnification: A-H, × 60.

α-SM actin expression in this animal cap assay validates the prediction initially stated; there is an excellent correla-tion between the expression of this SM marker and the potential mesodermal fate specified by the PGFs that were tested.

It is commonly accepted that mesoderm can be divided into four parts with regard to its destiny: (i) the notochord origin of the backbone, (ii) the somites, which give rise to axial muscles (myotomes), skeleton (sclerotome) and connective tissues of the skin (dermatome), (iii) the intermediate mesoderm, which forms part of the urogenital system, and (iv) the lateral plate mesoderm, which gives rise, among other tissues, to blood islands, heart and smooth muscles (Carl-son, 1981).

Attempts to understand the molecular basis of regional patterning of the mesoderm in the frog embryo was so far almost exclusively focused on the formation of dorsal structures for which cell identity (Zanetti et al., 1985; Gurdon et al., 1985; Hopwood et al., 1989; Smith et al., 1991; Kint-ner and Brockes, 1984) and positional markers (Ruiz i Altaba and Melton, 1989; de Robertis et al., 1989) are well defined. Because there have been no molecular markers to document the differentiation of ventrolateral mesoderm derivatives, the ventrolateral mesoderm patterning has not been so extensively studied. The objective of this work was to analyze ventrolateral mesoderm specification in the Xenopus embryo using a new tool to document ventrolateral mesoderm derivatives. Such a regional marker of mesoderm should fulfil four criteria: (i) it should be expressed in ventrolateral mesoderm derivatives, (ii) it should be activated following mesoderm induction, (iii) it should be regionalized within the early embryo, and (iv) it should be induced by PGFs which are potentially involved in the spec-ification of this area of the mesoderm. Since SM cells are thought to arise from the splanchnic layer of the lateral plate mesoderm, we investigated whether an α-actin isoform specific for the SM could serve as a reliable marker of ventrolateral mesoderm.

In vertebrates, actins constitute a multigene family expressed in a tissue-specific manner. One can identify, on the basis of the amino acid sequences, (i) two cytoskeletal actins, termed β and γ, which are expressed in virtually all cell types, and (ii) two sarcomeric actins, one restricted to the heart (α-cardiac), one specific for skeletal muscle (α-skeletal) and two SM actins, α and γ, which are expressed in the vascular and the visceral SM (Vandekerckhove and Weber, 1984). Xenopus laevis possesses three cytoskeletal actin genes (Cross et al., 1988) that differ from the β- and γ-types (Vandekerckhove et al., 1981), and one of them is specifically expressed in the muscle (Mohun and Garrett, 1987). The α-cardiac and α-skeletal actin genes (Mohun et al., 1984; Gurdon et al., 1985) are found to be expressed in a tissue-specific manner in the adult (Mohun et al., 1984). However, unlike other vertebrates, there is only one SM actin gene of α type that is detected in this anuran (Van-dekerckhove and Weber, 1984).

Since the Xenopus actin gene family appears to differ from those of other vertebrates, it was important to determine the specificity of the α-SM actin antibody used in this study (Skalli et al., 1986). In adult Xenopus tissue extracts, this antibody recognized a single band in SM-containing tissues. The expression of α-SM actin is in perfect correlation with what was previously reported in other species (Vandekerckhove and Weber, 1979, 1981). Therefore, the anti-α-SM actin antibody presented all the characteristics of a specific marker of SM cells in Xenopus laevis and could thus be used to analyze mesodermal patterning in this species.

During development, the distribution of the α-SM actin seems to present a greater tissue specificity than the two α-sarcomeric actins previously described in Xenopus (Mohun et al., 1984); both α-cardiac and α-skeletal actin genes are transcribed simultaneously in striated muscles of the somites and in the heart at the early neurula stage, even if later in development they will both present a tissue-specific pattern of expression (Mohun et al., 1984). α-SM actin expression is rapidly restricted to visceral and vascular SM cells, and is only transiently expressed in the developing heart from stage 37/38 to stage 46. A similar pattern of expression was also observed in other vertebrate species. Ruzicka and Schwartz (1988) have shown that the initial expression of SM actin transcripts take place in the epimy-ocardium of the avian embryo between stage 8 and 10; and, in the rat, by gestational day 10 and onward, α-SM actin is detected in cardiomyocytes (Sawtell and Lessard, 1989; Woodcock-Mitchell et al., 1988). There is no clear correlation between this transient expression of α-SM actin in heart anlage and the stage at which heart begins to beat (stage 33/34 NF). Neither the functional significance of this transient and “ectopic” expression of α-SM actin in the heart nor the mechanisms involved in the down-regulation of its expression by stage 46 are known.

The major sites of α-SM actin expression are SM cells of the presumptive gut, where it can be detected as early as stage 40/41 and the vascular SM cells of large vessels, which are mainly located in the anterior part of the embryo. Throughout development, α-SM actin is expressed in a layer of cells that is initially associated to the endodermal mass and later constitutes the outer coating of the intestinal loops. Since Xenopus possesses only one SM actin isoform (Vandekerckhove and Weber, 1984), it is not surprising to find that intestinal SM cells express the α-actin isoform, which is more specific for the vascular SM cells in other vertebrates.

Since the expression of α-SM actin can be induced by vegetal pole cells in animal cap isolated at the blastula stage, this gene behaves as others that are activated during mesoderm induction, like α-cardiac actin (Gurdon et al., 1985), MyoD (Hopwood et al., 1989), Mix.1 (Rosa, 1989), Xwnt-8 (Christian et al., 1991) and Brachyury T (Smith et al., 1991) genes. In conjugate experiments, this SM differentiation marker can be detected after 3 days of culture (equivalent stage 40/41), which is a much later stage than the other markers.

Interestingly, explants of mesoderm isolated from different areas of the marginal zone of gastrula stage embryos do not present the same potential to express this SM marker. Explants of the ventral and lateral marginal zone do pos-sess this capability, while prospective dorsal mesoderm does not. Thus, α-SM actin appears to be regionalized in its expression. At the gastrula stage, lateral and ventral mesoderm are committed to forming α-SM actin-expressing cells. It is likely that α-SM actin expression might represent the last step of a cascade of molecular regulation initiated by mesodermal-inducing PGFs. A number of recent findings support the idea that bFGF could be a natural mesoderm inducer in Xenopus (Kimelman and Kirschner, 1988; Slack and Isaacs, 1989; Shiurba et al., 1991; Amaya et al., 1991). In contrast, the role of the members of the TGF-β family, such as activin, remains unclear since their spatial and temporal expression has not been found to be correlated with the time at which mesoderm induction occurs in the embryo (Thomsen et al., 1991). Recently, it was shown that a member of the Wnt oncogene-related proteins family, Xwnt-8 (Christian et al., 1991), could play a role in the specification of mesoderm by changing the character of mesoderm induced by bFGF (Christian et al., 1992). So far, there is no direct evidence for the involvement of these different PGFs in mesoderm induction, but their study may serve as a paradigm to address the mechanisms underlying this inductive event. By the use of the animal cap assay, we have analyzed whether PGFs of the FGF and TGF-β families, potentially involved in the specification of the different mesoderm areas, could also be involved in the induction of α-SM actin expression. It appears that a specificity in the induction of α-SM actin expression exists. bFGF and a low concentration of XTC-GTX-11-conditioned medium have the potential to specify the SM fate in this assay. Conditioned medium from the XTC-GTX-11 cell line at a high concentration failed to induce α-SM actin expression in blastula animal cap. This is in good agreement with the prediction that members of the FGF family and a low level of activin can induce ventral mesoderm (Slack et al., 1987; Amaya et al., 1991; Green et al., 1990) and that a high concentration of activin induces dorsal mesoderm (Green et al., 1990; Thomsen et al., 1990).

Therefore, α-SM actin fulfils the four criteria of a ventrolateral marker of mesoderm, stated at the beginning of the discussion: it is expressed in ventrolateral aspects of the embryo, it is activated following mesoderm induction, it is regionalized to the ventrolateral region of the marginal zone and it is specifically induced by bFGF and by low levels of activin, which are the putative PGFs involved in the determination of this area of mesoderm. The expression of this novel marker, together with previous data on the differ entiation of notochord, skeletal muscle and blood cells in isolated marginal zone explants (Dale and Slack, 1987), indicate that SM is the cell type more frequently observed in derivatives of the lateral marginal zone, whereas blood cells and notochord are respectively more representative of ventral and dorsal marginal zone derivatives. Although α-SM actin can only be detected by the tadpole stage, it represents an excellent marker to identify, at a molecular level, ventrolateral mesoderm.

We thank Drs van den Eijden-Van Raaij and Prats for provid-ing us respectively with the XTC-GTX-11-conditioned medium and recombinant bFGF. We are thankful to Dr Michael Matthay for reviewing the English manuscript. Dominique Morineau is acknowledged for his expert photographic assistance.

This work was supported in part by CNRS and by a grant of the Fondation pour la Recherche sur les Myopathies (Contract AFM 1990-1991). The financial support of Telethon (Italy) to the project “Molecular modulation of cell adhesion and cytoskeletal molecules during neuromuscular development and regeneration” is gratefully acknowledged. V. K. is a directeur de recherche at INSERM. J.-P. S.-J. was supported by a fellowship of the Association pour la Recherche sur le Cancer (Contract 1990-1991) and of the Association Française contre les Myopathies (Contract 1991-1992).

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