The role of microfilaments in cranial neurulation in rat embryos: effects of short-term exposure to cytochalasin D

The presence of cytoplasmic microfilaments has been correlated with a wide variety of morphogenetic movements, suggesting that these are the contractile elements which generate cell movement and epithelial curvature (Wessels et al. 1971; Spooner, 1974). The importance of microfilament contraction in neurulation in amphibian, avian, and mammalian embryos is now widely accepted (Baker & Schroeder, 1967; Burnside, 1971; Freeman, 1972; M. Jacobson, 1978), but there is some disagreement as to whether they are actually responsible for generating curvature of the neural epithelium, or whether they simply bring about the observed shrinkage of the neural plate’s surface area which, in amphibian embryos at least, precedes the development of the neural folds (A. G. Jacobson, 1978).

The role and timing of microfilament-mediated contraction during cranial neurulation in mammalian embryos is not obvious. The cranial neural plate first forms convex neural folds which subsequently flatten and then become concave prior to neural tube closure (illustrated in accounts of human, pig, and rat development: Hamilton & Mossman, 1972; Patten, 1948; Morriss & Solursh, 1978). In the rat, and probably in other mammalian species also, these changes in neural fold shape are accompanied by changes in cellular shape and organization within the neural epithelium (Morriss-Kay, 1981). Dense apical microfilament bundles have been reported only at the concave stage (Morriss & New, 1979). At the 10-somite (early concave) stage in rodent and human (Hamilton & Mossman, 1972) embryos, a small point of neural fold apposition occurs at the forebrain-midbrain junction, dividing the anterior neuropore into two parts.

Cytochalasin B has been used in a variety of developing systems, where it has been found reversibly to inhibit microfilament structure and function (Wessels et al. 1971; Spooner, 1978). In a preliminary study on cultured rat embryos, the effect of cytochalasin B on the late stages of cranial neurulation suggested that these stages are microfilament-dependent (Morriss-Kay, 1981). In the present study we have used cytochalasin D which is more potent than cytochalasin B (Carter, 1967), having a higher affinity for the contractile-related binding site (Tannenbaum, 1978), and has an interpretative advantage in that it has little or no effect on hexose transport (Miranda, Godman, Deitch & Tanenbaum, 1974a; Tannenbaum, Tanenbaum & Godman, 1977). The aims were as follows: (a) to compare cytochalasin-D-induced effects with the normal shape and ultrastructure of the neural epithelium at three stages of neurulation (convex, early concave and late concave), in order to determine the relative importance of microfilament contraction at these stages; (b) to monitor recovery of neural fold shape and to discover whether neural tube closure could occur following short-term exposure to cytochalasin D at the same three stages; (c) to correlate changes in shape with ultrastructural changes in the microfilament-rich apical border of the concavestage neural epithelium.

Wistar-strain rat embryos were explanted in Tyrode’s saline at 9 –11 p.m. on day 9 of gestation (2-to 5-somite stage) or 9 –11 a.m. on day 10 (9-to 11-somite stage) and Reichert’s membrane was opened (day of positive vaginal smear = day 0). Day-10 embryos were examined under the dissecting microscope and divided into two groups: ‘preapposition-stage’ embryos (9-somite to early 10-somite stages, in which neural fold apposition at the forebrain-midbrain junction had not yet occurred, and ‘postapposition-stage’ embryos (11 somites) in which the open region of the cranial neural tube was divided into a midbrain/upper hindbrain spindle-shaped opening and the forebrain anterior neuropore (Morriss & New, 1979, fig. 3). 10-somite-stage embryos which had just initiated apposition were used as controls.

The embryos were cultured at 38°C in 60ml cylindrical bottles rotating at 30r.p.m. The culture medium consisted of 2 ·5 ml immediately centrifuged, heat-inactivated rat serum (Steele & New, 1974) and 2 ·5 ml Tyrode’s saline containing 50 μgml^-1 streptomycin and penicillin and either cytochalasin D or DMSO. The gas phase was 5 % O₂/5 % CO₂/90 % N₂ (New, Coppola & Cockroft, 1976a,b). After 1 h, some embryos were removed, washed in Tyrode’s saline and fixed for in 2 ·5% cacodylate-buffered glutaraldehyde (0 ·1 M, pH7 ·2). They were then transferred to buffer and viewed with the dissecting microscope. Yolk sac and amnion were removed from all day-10 embryos and from day-9 embryos which were to be prepared for scanning electron microscopy (SEM). Neural fold shape and somite number were assessed before further processing for SEM or for light microscopy (LM) and transmission electron microscopy (TEM).

The remaining embryos were transferred to fresh medium prepared as described above but without the addition of cytochalasin D or DMSO, regassed with the same gas mixture, and continued in culture. Each culture bottle contained a maximum of eleven embryos for the first hour, and a maximum of six embryos thereafter. Eighteen day-9 embryos and thirteen day-10 embryos (eight preapposition and five postapposition) were removed and fixed after a further 12 h, and eleven day-10 embryos (six preapposition and five postapposition) were removed and fixed after 5 h. All remaining embryos were cultured until the morning of day 11 (36 or 24 h in the addition-free medium), being regassed on late day 10 with 5 % CO₂ in air (New et al. 1976a, b). They were then washed, fixed, and transferred to buffer, and the membranes removed. Somite number, general morphology, and neural tube morphology was recorded for all embryos; some were then prepared for SEM or LM and TEM.

Cytochalasin D (Aldrich Chemical Co.) was prepared as a 10% aqueous stock solution of lmg cytochalasin D in 1ml dimethylsulphoxide (DMSO) (Sigma) and stored frozen. 10% aqueous DMSO was used for controls. In a series of preliminary cultures, 2 ·5 to 25 μl of the cytochalasin D solution was added to 5 ml serum to produce a range of concentrations of 0 ·05 to 0 ·5 μgml^-1, in each of which four or five day-10 embryos were cultured for 1 –3 h. Cranial neural fold shape, which was partly or wholly concave in profile at the start, was examined in the living state and by SEM, LM, and TEM, and compared with that of controls. Some embryos were transferred to addition-free medium after exposure to cytochalasin D or DMSO, cultured for a further 24h, and examined as at 1-3 h. On the basis of these experiments, a cytochalasin D concentration of 0 ·15 μgml^-1 (3 ×10^–6M; 7 ·5 μl stock solution) and an exposure period of lh were chosen for all subsequent experiments, being the minimum concentration and time which produced complete collapse of the neural folds. 7 ·5 μl of aqueous DMSO was added to 5 ml medium for control embryos.

Exposure of day-10 embryos to 0 ·15 μgml^-1 cytochalasin D had similar morphological effects to that of 0 ·5 μgml^-1 cytochalasin B as used previously (Morriss-Kay, 1981), confirming the greater potency of cytochalasin D.

All embryos not used for LM and TEM were prepared for SEM. They were dehydrated in graded acetones, critical-point dried, mounted on aluminium stubs with double-sided Sellotape, coated with gold in a sputter coater, and viewed in a JEOL JSM-T20 scanning electron microscope.

Embryos were photographed whole for reference during subsequent sectioning. They were then postfixed in cacodylate-buffered osmium tetroxide, washed, dehydrated, and embedded individually in Spun resin at an orientation appropriate for cutting transverse sections. 1 gm sections were mounted on glass slides and stained with 0 ·5 % methylene blue/0 ·5 % azure II in 1 % borax for light microscopy. Adjacent sections were cut ultrathin for TEM, and stained with uranyl acetate and lead citrate. Two to four embryos of each of the initial neural fold stages and from each culture period were prepared for LM and TEM.

The development and appearance with LM, TEM and SEM of control-cultured embryos used in this study was not detectably different from that of embryos cultured in addition-free medium in other studies in this laboratory, or from embryos freshly explanted at equivalent stages. We therefore conclude that the DMSO added to the culture medium of both control and cytochalasin-treated embryos had no detectable effect on development.

In the following description, not all of the features described for control embryos are illustrated. Illustrations of many of the features referred to here may be found in Morriss & Solursh (1978), Morriss & New (1979) and Morriss-Kay (1981).

At the start of culture, embryos were at the 2-to 5-somite stage, with convex cranial neural folds. LM of preculture embryos, and control embryos after 1 hour’s culture, showed a 50 μm-deep neural groove consisting of approximately five supranotochordal cells and close apical surface apposition of the adjacent ten cells or so on each side (Fig. 1A).

After 1 hour’s culture in medium containing cytochalasin D the shape appeared unchanged by dissecting microscope (live embryos) and SEM (Fig. 2A,B) observations. LM revealed that the neural groove had opened out to form a V-shape in the region which was closely apposed in the midline in control embryos (Fig. 1B). Otherwise, the LM appearance of neuroepithelial cell shape and organization was similar to that of control embryos, having a columnar or slightly pseudostratified form. TEM (not illustrated) showed that the cells were approximately the same breadth from base to apex, lacking the narrow necks and apical surface bulges seen at later stages. At their apical border they were joined by short desmosome-like junctions. Filamentous material was attached to these junctions, extending for only a few nm in the plane of section, in both control and cytochalasin D-treated embryos.

After a further 12 h culture in addition-free medium (Fig. 2C,D), both control and cytochalasin D-treated embryos had gained six to seven somite pairs, so that they ranged in somite number from 8 to 11. All of the treated embryos showed neural tube/fold development (form of cranial neural folds, spinal neural tube, and posterior neuropore) appropriate to the previous somite stage, e.g. embryos with nine pairs of somites had neural tube/fold development resembling that of 8-somite-stage controls. Only one treated embryo (with eleven pairs of somites) showed any concavity of the cranial neural epithelium; this epithelium had the ultrastructure as well as the shape of that of a 10-somite control embryo, with apical microfilament bundles and junctions oriented parallel to the apical surface but not forming the near-continuous line normally seen at the 11-somite stage onwards. Heart rate, yolk-sac blood island development, and yolk-sac and amnion expansion were similar in control and treated embryos; ‘turning’ was slightly retarded relative to somite number in the treated embryos.

Of the eighteen cytochalasin D-treated embryos cultured from day 9 to day 11 (Table 1), twelve formed neural tubes which were completely closed except for the small posterior neuropore. These differed from control embryos in being slightly smaller, and in having a less well developed cranial flexure (Fig. 2E,F). LM and TEM (not illustrated) showed a normally organized neural epithelium but with many pyknotic cells and less well-expanded brain vesicles and neural canal than those of controls.

The six embryos with open neural tube defects (Fig. 2G) were all unturned; the embryonic axis formed a tight U-shape, with the amnion closely applied to the surface of the neural folds. There was very little amniotic fluid present, no yolk-sac circulation, and the heartbeat was weak. However, it was striking that even in these embryos the otic pit had sunk beneath the surface to form a closed otocyst. (Four of these embryos were from one culture bottle which contained no other embryos, and two were from a bottle which also contained four embryos whose brain tubes closed.) Since twelve of these embryos succeeded in forming closed brain tubes, we conclude that failure to do so was not due to a primary effect on neurulation, and this result will not be discussed further. One control embryo was also unturned, and compressed into a tight U-shape within a fluid-deficient amniotic cavity. There were neural tube defects in the midbrain and spinal regions.

This embryo was normal by all visible criteria when the medium was changed 17 h after the start of culture in DMSO-free medium, and is therefore assumed to have been damaged by handling at this stage.

Monitoring during culture showed that all cytochalasin-D-treated embryos developed a yolk-sac circulation later than controls; six of the twelve embryos with closed neural tubes had not achieved this at termination.

During the 9-to 11-somite stages, the midbrain/hindbrain cranial neural epithelium of normal rat embryos begins to develop a concave surface, apposes and fuses at the forebrain-midbrain junction to form the spindle-shaped midbrain-upper hindbrain neuropore, which then narrows as the lateral edges move towards each other (illustrated in Morriss & New, 1979). At the same time, the two sides of the forebrain (anterior neuropore) move together and become apposed.

TEM of control embryos of these stages showed that where the neural epithelium has a flat or only slightly concave apical surface, the cells have narrow necks with apical bulges, with the subapical junctions predominantly parallel with the long axis of the cells (i.e. vertical). Microfilaments could be seen attached to the subapical junctions but only rarely formed a continuous bundle stretching between two junctions within the plane of section (Figs 3A, 4A). Where the epithelial surface was more concave, many of the subapical junctions were orientated parallel to the surface (i.e. horizontal); together with the microfilament bundles they formed an almost unbroken subapical line, with rounded areas of cell above it. This pattern was also characteristic of the concave neural epithelium of later stage embryos (Figs 3E, 4F).

After a lh exposure to cytochalasin D, embryos which had not formed the forebrain-midbrain apposition point at the start of culture (late 9- or early 10-somite stage, Fig. 5A) showed a convex midbrain-hindbrain neural epithelium, i.e. neural folds which had begun to form concave surfaces had flopped laterally (Fig. 5B). The lateral forebrain neural epithelium was also slightly everted, but the deep optic sulci were maintained. LM and TEM showed the neuroepithelial cells to have much broader surfaces and neck regions than those of equivalent regions in control embryos (Fig. 4B). Microfilament-like material could be seen attached to the subapical junctions or free within the cytoplasm; this was either fuzzy and indefinite in structure (Fig. 4B) or in the form of broken filaments (Fig. 4C).

Embryos which had formed the forebrain-midbrain apposition point at the start of culture (11-somite-stage embryos) showed a widely gaping midbrain-hindbrain neuropore, but retained a concavely curved neuroepithelial surface in this region. The TEM appearance of the apical region was similar to that of unapposed embryos, with few intact microfilament bundles, horizontal junctions or apical surface bulges. Maintenance of the concave shape therefore appeared to be due to the limitation on lateral movement of the neural folds imposed by the spindle-shaped neuropore rather than to the structural organization of the epithelium itself.

After culture in addition-free medium, recovery of neural fold shape had progressed well in both preapposition- and postapposition-stage embryos, but had not yet achieved the degree of elevation seen at the start of culture (Fig. 5C,F). TEM showed some narrowing of the neck region and apical surface of the neuroepithelial cells, some reappearance of apical microfilament bundles, and slight bulging of the apical cell surfaces. However, no horizontal junctions were seen, and the few microfilament bundles observed were less clearly organized and closer to the apical surface than those of control embryos (Fig. 4D).

After 12 h culture in addition-free medium, control embryos (Fig. 5G) were all normal and had gained five or six pairs of somites and a yolk-sac circulation. Cytochalasin-D-treated embryos had gained five to seven somite pairs except for one 11-somite-stage embryo which increased to nineteen somite pairs. They had a slightly weaker heartbeat than the controls, and blood islands but no circulation in the yolk sac. Of the five embryos which were at the postapposition stage at the start of culture, two had completed cranial neurulation, and the other three had small inverted-teardrop-shaped midbrain-upper hindbrain openings (Fig. 51,J), one with a small anterior neuropore also.

None of the eight embryos which were at the preapposition stage at the start of culture had achieved forebrain-midbrain apposition, even though the widely collapsed cytochalasin-D-affected neural folds had regained their concave curvature and in three embryos (with sixteen to seventeen somite pairs) the lateral edges were very close to each other in the midline (Fig. 5H).

The ultrastructure of the apical region of the cranial neural epithelium which had regained its concave curvature was similar in all embryos examined, whether pre- or postapposition at the time of cytochalasin exposure. Microfilament bundles and horizontal junctions were present, but did not form such a clear subapical line as that seen in late-preclosure-stage control embryos, though apical surface bulges were well developed (Figs 3D,E and 4E,F).

After 24 h in addition-free medium (Fig. 6A –C and Table 1) the cranial neural tube of all embryos of the postapposition group was closed. However, like cytochalasin-treated day-9 embryos cultured to day 11, the cranial flexure was less acute than in controls, the brain vesicles less well expanded, and there was much pyknosis in the neural epithelium and elsewhere. All had turned.

None of the ‘preapposition’ embryos had succeeded in forming the apposition point, and the cranial neural tube was wide open from the forebrain to the metencephalon, i.e. the whole area which had been open at the start of culture. Five had not completed turning.

The otic pit was wide open in all embryos exposed to cytochalasin D on day 10, whereas in all of the controls a closed otocyst had formed. Treated embryos were smaller than controls, the heart rate was slower, and the yolk-sac circulation was poor or absent. Spinal neural tube closure progressed normally except for the slight retardation described above.

During the 24 h period from late day 9 of gestation, rat embryos undergo a complex integrated sequence of morphogenetic changes. All of these events involve cell movement, exocytosis, or endocytosis, and may therefore be assumed to be dependent to some degree on microfilaments. The possibility of an adverse effect on some or all of them, and secondary effects through them, must be born in mind when interpreting the results presented here. Cytochalasin D must have affected all microfilament-dependent functions, even though its effect on the thick subapical line of microfilament bundles in the curving neural epithelium is the only one which was immediately obvious after a lh exposure period. For instance, during the longer recovery periods, cytochalasin-D-treated embryos were observed to develop a yolk-sac circulation more slowly than controls, and to have a slower heart beat. Consequently a less efficient supply of oxygen and nutrients may have contributed to the pyknosis seen in the neural epithelium with LM and TEM in these embryos, and to their smaller size.

Yolk-sac-mediated nutrition involves phagocytosis, and embryos of the stages used here are dependent on this process for their survival, growth, and normal development (Beck & Lloyd, 1966). Mimura & Asano (1976) observed a 30% inhibition of phagocytosis in peritoneal macrophages in medium containing 0 ·5gml^-1 cytochalasin D but no inhibition at 0 ·1 μg ml^-1. Although the concentration of 0 ·15 μg ml^-1 used in the present study is low in relation to the macrophage study, the slightly smaller size of the treated embryos when compared with controls at day 11 suggests that there may have been some effect on nutrient uptake. But while an effect on phagocytosis may have had some effect on growth, it could not have been responsible for the initial morphogenetic alterations brought about by cytochalasin D treatment. Transfer of the breakdown products of nutrients phagocytosed by yolk-sac endoderm cells at this stage is a relatively long-term process (Beck et al. 1967), whereas we observed neural fold collapse within one hour.

The effects of cytochalasin D on day-10 embryos showed a clear correlation between loss of neuroepithelial shape and loss of microfilament bundles. During subsequent culture in addition-free medium, the pretreatment cranial neural fold shape was regained in more than 5h but less than 12 h; ultrastructurally this was correlated with regeneration of subapical microfilament bundles, suggesting a causal relationship between the two events.

Studies on the mechanism of action of cytochalasins suggest that they have up to four effects on microfilament structure, function and organization, all of which are reversible. Different effects have been observed using different cell types and different experimental protocols. MacLean-Fletcher & Pollard (1980) observed an effect on filament-filament interactions, whereas Schliwa (1982) found that side-to-end junctions of actin filaments were intact after cytochalasin treatment. Cytochalasins can also prevent filament elongation, binding to the fast assembly end of the microfilament and thereby preventing further polymerization except from the slow assembly end (Flanagan & Lin, 1980; Lin, Tobin, Grumet & Lin, 1980; Brown & Spudich, 1981). They can also induce rapid depolymerization of filamentous actin (Casella, Flanagan & Lin, 1981, using platelets in medium containing 0 ·5 μgml^-1 cytochalasin D), although this effect was not observed in various established cell lines such as HEp-2 and HeLa (Morris & Tannenbaum, 1980; Miranda et al. 1974a; Miranda, Godman & Tanenbaum, 1974b). Disruption of filaments with the release of filament fragments was the chief effect observed by Schliwa (1982).

Our ultrastructural observations suggest that the effect of cytochalasin D on neurulation in rat embryos is mediated through microfilament disruption, involving both depolymerization and fragmentation, and that loss of cross linking between parallel actin filaments within microfilament bundles may also occur.

Cytochalasin B was first used to study neurulation by Linville & Shepard (1972). They exposed chick embryos in culture to 2 ·5, 5, and 10 μgml^-1 throughout the period of neurulation. At the two higher concentrations many of the embryos had open neural tube defects, but at 2 ·5 μgml^-1 only 15 of 22 viable embryos were affected (though all were retarded in the timing of closure). These results are surprising in view of our earlier observation of neural fold collapse in rat embryos exposed to only 0 ·5 μgml^-1 cytochalasin B (Morriss-Kay, 1981). The difference may be due to methodology or to a species difference; ultrastructural observations were not made.

Lofberg (1974) observed only retardation of neurulation and an elongated neural plate in axolotl embryos treated with 1 μgml^-1 cytochalasin B, whereas 2 ·5 and 5 μgml^-1 resulted in disaggregation of the neuroepithelial cells and consequent neurulation failure. In this case the differences from our results are more likely to involve a species difference, since even the disaggregating neural tissue formed elevating neural folds, whereas in our rat embryos neural fold collapse occurred at cytochalasin B or D concentrations much lower than those which induced even minor signs of disaggregation. Our results also suggest that in rat embryos cranial neurulation is more vulnerable than spinal neurulation to very low concentrations of cytochalasins.

We saw no indication of any effect on premitotic nuclear migration or cytokinesis, perhaps due to the short exposure time as much as to the low concentration used. Webster & Langman (1978) observed these effects, together with some disaggregation, in the neural epithelium of day-11 mouse embryos exposed to 10 μgml^-1 cytochalasin B for 2h in culture. This concentration is twenty times that used in our cytochalasin B study.

Studies on the effects of cytochalasins on neurulation in mammalian embryos in vivo show that neural tube defects can be brought about by ingestion of these drugs (Shepard & Greenaway, 1977; Wiley, 1980; Austin, Wind & Brown, 1982). However, the cellular effects associated with exencephaly and encephalocoele did not involve microfilaments at the lower dose levels, suggesting a maternal-mediated effect, to which a direct effect on embryonic microfilaments was added at the higher dose levels (Wiley, 1980).

Our results suggest that although there may have been minor effects on phagocytosis and (less likely) exocytosis, direct interference with the role of microfilaments in epithelial morphogenesis was the major effect of cytochalasin D on day-9 and day-10 embryos. The relative importance of the effects on the mechanisms of neurulation, otocyst formation, cranial flexure, and turning depended on the stage of development at the time of exposure.

Some conclusions may be drawn for neurulation. In late day-9 embryos, microfilaments are mainly involved in maintaining close apposition of the apical neuroepithelial cell surfaces adjacent to the midline, and do not play a major role in maintaining shape of the convex neural folds. In day-10 embryos, the subapical line of microfilament bundles is essential for generation and maintenance of the V-shaped or concave form of the neural folds. Following cytochalasin-induced collapse of the elevated neural folds, re-elevation can take place as the microfilament bundles regenerate. But if the forebrain-midbrain midline fusion (apposition) point has not been achieved prior to cytochalasin exposure, it will not form after re-elevation even if the neural folds are closely apposed. This developmental event, involving cell-cell adhesion, appears to be finely timed and of crucial importance in brain tube formation.

We wish to thank Martin Barker, Beth Crutch and Janet Kilcoyne for technical and photographic assistance, and the MRC for financial support.