Caenorhabditis elegans morphogenesis: The role of the cytoskeleton in elongation of the embryo

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Caenorhabditis elegans Morphogenesis: The Role of the Cytoskeleton in Elongation of the Embryo JAMES *MRC

Laboratory

R.

PRIESS*+’

AND

Biology, Hills Road, and Developmental Biology,

Cambridge University

of Molecular Cellular

Received

November

12, 1985; accepted

DAVID

I.

CB2 ZQH, of Colorado.

iw revised

HIRSH~~’ England, Boulder,

f&r-m

April

and TDepartment Colorado 80309

of Molecular,

2, 1986

During development Caenorhabditis elegans changes from an embryo that is relatively spherical in shape to a long thin worm. This paper provides evidence that the elongation of the body is caused by the outermost layer of embryonic cells, the hypodermis, squeezing the embryo circumferentially. The hypodermal cells surround the embryo and are linked together by cellular junctions. Numerous circumferentially oriented bundles of microfilaments are present at the outer surfaces of the hypodermal cells as the embryo elongates. Elongation is associated with an apparent pressure on the internal cells of the embryo, and cytochalasin D reversibly inhibits both elongation and the increase in pressure. Circumferentially oriented microtubules also are associated with the outer membranes of the hypodermal cells during elongation. Experiments with the microtubule inhibitors colcemid, griseofulvin, and nocodazole suggest that the microtubules function to distribute across the membrane stresses resulting from microfilament contraction, such that the embryo decreases in circumference uniformly during elongation. While the cytoskeletal organization of the hypodermal cells appears to determine the shape of the embryo during elongation, an extracellular cuticle appears to maintain the body shape after elongation. Q 1986 Academic press, IK INTRODUCTION

To understand the basis for morphogenesis in embryonic tissues it is desirable to know where cells divide, migrate, die, or change shape. The relatively few cells of the nematode Caenorhabditis elegans suggests that a detailed understanding of morphogenesis should be possible for this organism, and the complete cell lineage and anatomical studies on C. elegans embryos (Sulston et al, 1983) provide a useful background for such a study. In addition, several mutants with abnormal morphology have been isolated in C. elegans (Hirsh and Vanderslice, 1979; Wood et ah, 1980; Schierenberg et aL, 1980; H. Schnabel and R. Schnabel, unpublished results; J. Priess, unpublished results), providing an opportunity for identifying gene products that have controlling roles in morphogenesis. A particularly striking example of morphogenesis in the C. elegans embryo is the change that occurs in the shape of the body. The early development of the embryo involves rapid cell proliferation, but little change in shape from the approximately spherical fertilized egg. Midway through development the embryo begins to elongate rapidly along its anterior-posterior axis, increasing in length about fourfold. As the embryo elongates there is almost no division or migration or cells i To whom reprint ’ Current address: 0012-1606/86 Copyright All rights

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(Sulston et al., 1983). Instead cells throughout the embryo appear to change shape coordinately. Mechanisms of elongation in single cells have been studied for plant cells (see Lloyd and Barrow, 1982) and insect oocytes (Tucker and Meats, 1976; Went, 1978). In both systems cellular elongation appears to result from asymmetric growth. The plant cells are girdled by circumferentially oriented cellulose microfibrils, and the insect oocytes by follicular epithelial cells containing circumferentially oriented microtubules; these structures appear to limit radial expansion in an otherwise symmetrical growth pattern, and thus cause the cell to elongate. In contrast, growth does not appear to drive elongation in C. elegans embryos, which maintain a constant volume throughout embryonic development. This paper presents an analysis of elongation in the C. elegans embryo. The cells responsible for elongation are identified, and the cytoskeletal organization of these cells is analyzed. The distribution of actin is examined with fluorescently labeled phallotoxins, and antibodies against yeast or Drosophilia tubulins are used to examine the distribution of microtubules. The microfilament inhibitor cytochalasin D and the microtubule inhibitors colcemid, nocodazole, and griseofulvin are used to study the functions of actin and tubulin in elongation. A mutant that is able to elongate, but then shortens in length late in development, is examined to determine how the shape of the animal is maintained after elongation and hatching.

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Nematode strains and maintenance. Wild-type (NZ) Caenorhabditis elegans strain Bristol was cultured at 25°C on agar plates with Escherichia coli as food source (Brenner, 1974). sqt-3 (e2117) (kindly provided by Heinke Schnabel and Ralph Schnabel), sqt-3 (s&3), and sqt-3 (e24) (formerly dpy-15 (e2Q) were maintained at 15°C. Culture medium a.nd inhibitor solutions. The embryonic culture medium used in all experiments described here consisted of 3.7% sucrose and 0.1 MNaCl. The osmolality of the culture medium is critical, and is tested by determining whether the embryonic pseudocoelom (Sulston et al,, 1983) expands or contracts following laser permeabilization of the eggshell (see below). Stock solutions of inhibitors were prepared as follows: cytochalasin D (Sigma) at 2 mg/ml in 95% ethanol, colcemid (Sigma) at 1 mg/ml in H20, and griseofulvin (Sigma) at 10 mg/ml in dimethylformamide. Stock solutions were stored at 4°C and added to embryonic culture medium immediately before use. Laser permeabilixation of embryos. Embryos were cut from gravid hermaphrodites, then collected with a micropipette and clustered with an eyelash on a thin pad of 2% agarose following the general procedures of Sulston et al. (1983). A coverslip coated with 0.1% polylysine was placed on top of the embryos, then lifted gently from the pad with the attached embryos by flooding the pad with water. The coverslip was inverted and the embryos covered with a drop of FITC-polylysine prepared as follows: 1 mg of fluorescein isothiocyanate (Sigma) in 0.5 ml Hz0 was reacted with 10 mg of polylysine (Sigma) in 0.5 ml of coupling buffer (0.2 M NaHC03NaOH, pH 8.5), for 1 hr at room temperature. The solution was dialyzed overnight in the dark in phosphatebuffered saline (PBS, pH 7.4; 150 mM NaCl, 3 mM KCl, 8 mM Na2HP04, 1.5 mM KH2P04, 1 mM MgClJ. The embryos were incubated with the FITC-polylysine solution for about 1 min, then rinsed in culture medium and inverted over a depression microscope slide containing culture medium or inhibitor solutions. Embryos were examined with Nomarski differential interference contrast optics (Sulston and Horvitz, 1977) and a laser microbeam (Sulston and White, 1980) was used to irradiate eggshells of selected embryos. The eggshell of C. elegans consists of distinct inner and outer layers. Laser irradiation of the eggshell creates visible holes in the outer layer, which is coated directly with the FITCpolylysine. Complete permeabilization is achieved only when the inner layer adjacent to the site of irradiation breaks up into small bubbles. Scanning and transmission electron microscopy. Embryos were isolated by dissolving gravid adults with 1% hypochlorite, 0.5 M KOH for 5 min, collecting the eggs

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through a 52 lrn Nitex Filter (Nitex Corp) then rinsing the eggs three times in M9 buffer (Brenner, 1974). Eggshells were digested from the embryos with chitinase following the procedure of Wolf et al., (1983). Only the outer layer of the eggshell is removed by chitinase treatment; the inner layer was disrupted mechanically by pippetting the chitinased embryos through a drawn Pasteur pipette. After removal of the eggshell the embryos were fixed in 3% glutaraldehyde in 0.1 M Na-cacodylate buffer, pH 7.4 for 1 hr, then rinsed and postfixed in 1% 0~0~ in 0.1 M Na-cacodylate, pH 7.2 for 15 min. For scanning electron microscopy the fixed embryos were rinsed with HzO, then pipetted onto coverslips coated with 0.1% polylysine, dehydrated through a graded acetone series, and critical point dried. Specimens were coated with gold for 4 min at 7 mA with a Technics Hummer sputter coater and viewed with an AMR model 1000 A scanning electron microscope. For transmission electron microscopy, the fixed embryos were rinsed, embedded, and sectioned according to the procedures of Ward et al. (1975). For analysis of the striated layer of the cuticle, an alternative procedure was followed: embryos were postfixed for 45 min in 1% 0~0~ and 0.8% K3Fe(CN)G (0.1 M sodium cacodylate buffer, pH 6.0). The embryos were rinsed in 0.05 M sodium cacodylate buffer, pH 7.0, then treated for 15 min with 0.2% tannic acid (Malinkrodt) in 0.05 M sodium cocodylate buffer, pH 7.0. Embryos then were rinsed in HZ0 and dehydrated and embedded. Fluorescence microscopy of actin jibers. Actin fibers were stained with tetramethylrhodaminyl-phalloidin (Rh-phalloidin) and fluorescein-isothiocyanate-conjugated phalloidin (FITC-phalloidin). These compounds were gifts from Dr. T. Wieland and have been described previously (Faulstich et al., 1983; Wulf et al., 1979). Patterns of actin fibers in the hypodermal cells of C elegans are difficult to resolve in the fluorescence microscope if the entire embryo is permeabilized and stained because of the large quantities of actin in the muscles and intestinal cells that underlie the hypodermis (see Fig. 1). We have had some success in visualizing the hypodermis specifically by fixing the embryos with formaldehyde and then adding dilute detergent to permeabilize cells, as described below. The hypodermal cells tend to permeabilize before the internal cells, and actin filaments in the hypodermal cells alone are visible for a brief period of time. The photographs presented in this paper are of embryos in advanced stages of elongation; actin fibers are visible in the fluorescence microscope at earlier stages, but the fibers appear to be closer together and to stain less intensely than in later stages and thus are difficult to resolve in photographs. Embryos were isolated from gravid adults and the eggshells removed as described above. The embryos were

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fixed in 1 ml of 3.0% formaldehyde (0.1 M sodium cacodylate buffer, pH 7.2) for 15 min. The embryos were rinsed in embryonic culture medium, then placed in 1 ml of 0.2% Triton X-100 in culture medium plus 5 ~1 of either Rh-phalloidin (0.4 mg/ml in methanol), FITCphalloidin (0.4 mg/ml in methanol), or unlabeled phalloidin (1 mg/ml in methanol). The embryos were placed on a microscope slide and immediately viewed with the fluorescence microscope. Immunojluorescence microscopy. In experiments with the MH27 antibody, embryos were prepared for immunofluorescence microscopy according to the proedure of Albertson (1984). The following modified version of this method was used to stain microtubules in the hypoderma1 cells. Embryos were washed from the agar petri plates used for nematode culture, treated for 3 min in 1 M KOH, then rinsed in Hz0 and pippetted onto a coverslip coated with 0.1% polylysine. A second coverslip also coated with 0.1% polylysine was laid gently across the embryos, and excess fluid removed until the top coverslip was next to the embryos. The embryos were frozen by placing the coverslips on dry ice for at least 10 min following the procedure of Albertson (1984). The coverslips were separated and immersed in methanol at -20°C for 4 min and acetone at -20°C for 4 min and then air dried. The embryo fragments were pre-incubated with Tris-buffered saline (TBS, 150 mM NaCl, 50 mM Tris-HCl, pH 7.4) containing 0.5% Tween 20 for 20 min, then incubated for 1 hr at room temperature with either a monoclonal antibody to tubulin YLl/ 2(Kilmartin et ah, 1982) or a monoclonal antibody to Drosophilia cr-tubulin (kindly provided by Dr. M. Fuller). Coverslips were rinsed in three changes of TBS-Tween 20, then incubated with FITC-anti-mouse, or anti-rat, IgG (Miles, Slough, England) for 30 min at 20°C and the coverslips rinsed again. Specimens were observed with a Zeiss RA microscope equipped for epifluorescence. RESULTS

Background

and Review

of Embryonic

Anatomy

Descriptions of the anatomy and the complete cell lineage of the C. elegans embryo have been published (Sulston et al, 1983). At the time of hatching, the fully developed embryo has an outer layer of epithelial cells, called the hypodermis, that surrounds internal cells such as muscles, intestinal cells, and neurons (Fig. 1). The hypodermis consists of five longitudinal rows of hypodermal cells: a dorsal row, left and right lateral rows, and left and right ventral rows. Each hypodermal cell has an outer, apical, surface and an inner, basal, surface associated with a basement membrane. For the purposes of this paper it is useful to describe the different surface margins of a hypodermal cell in terms of the axes of the

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apical

surface

belt desmosome

ventral FIG. 1. Schematic diagram of a mid-section of a C. elegans embryo at hatching. Five hypodermal cells (H) encircle the embryo. The hypodermal cells have an outer, apical, surface surrounded by a belt desmosome (wide black line) and an inner, basal, surface (white arrowhead). The small arrowheads on the apical surface of a lateral hypodermal cell point to the longitudinal margins of the belt desmosome surrounding the cell, and the small arrows point to the circumferential margins of the same belt desmosome. The internal cells shown are intestinal cells (I), neurons (N), and groups of muscles (M).

entire embryo. We define the longitudinal margins of an apical hypodermal surface as the margins that are parallel to the anterior-posterior axis of the embryo, and the circumferential margins as the margins that follow the circumference of the embryo (Fig. 1). The hypodermal cells are joined at their apical surfaces by specialized junctions which are analogous to belt desmosomes.Belt desmosomesare found commonly in vertebrate epithelial cells that are subject to mechanical stress, and are thought to play an important role in cell-cell adhesion (cf. Staehelin, 1974). Vertebrate belt desmosomesconsist of intracellular rings of microfilaments and associated proteins such as myosin and tropomyosin, and apparently are linked mechanically to adjacent cells through extracellular material. The belt desmosomes of C. elegans resemble vertebrate belt desmosomes ultrastructurally (see Fig. 7d), contain actin (see below), and stain positively with an antibody against human tropomyosin (J. Priess, unpublished results). A monoclonal antibody MH27 (kindly provided by R. Francis and R. Waterston) shows a spatial and temporal staining pattern in C. elegans embryos apparently identical to the pattern of belt desmosomesseen in the electron microscope, and is used in this paper to illustrate the shapes of the hypodermal cells during elongation (see Fig. 4). As the embryo elongates the relative posi-

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tions of hypodermal cells do not change, consistent with the idea that the hypodermal cells are coupled together mechanically (see Figs. 4b, d, f).

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FIG. 3. Elongation of C. elegans embryos at 25°C. The rate of elongation of the body is proportional to the rate of elongation of the head, which is measured more conveniently. The length of the head, defined here as the distance from the anterior tip of the embryo to the junction of the pharynx and intestine, was measured during embryogenesis for two animals (open and closed circles).

Elongation

FIG. 2. Scanning electron micrographs of the apical surfaces of hypodermal cells in embryos prior to elongation. (a) Dorsal view. The arrow points to a single dorsal hypodermal cell. The left and right rows of dorsal hypodermal cells have interdigitated and now form a single longitudinal row. (b) Lateral view. The longitudinal rows of dorsal, lateral, and ventral hypodermal cells are visible. The embryo has a small ventral bend (arrow). (c) Ventral view. The arrows point to individual ventral hypodermal cells. Right and left pairs of ventral hypodermal cells meet at the ventral midline of the embryo. In (b) and (c) the anterior (left) end of the embryo has not yet been enclosed by hypodermal cells, and neural precursors are visible (arrowheads). Calibration: 10 pm.

of C. elegans Embryos

The first 4 hr (25°C) of development in C. elegans is a period of rapid cell proliferation in which there is no appreciable change in the overall shape of the embryo. The hypodermal cells initially are clustered together on the dorsal surface of the embryo in six longitudinal rows: left and right dorsal, left and right lateral, and left and right ventral. At about 200 min the hypodermal cells spread across and enclose the embryo, and the two rows of dorsal cells interdigitate to form a single longitudinal row (Fig. 2a). As the hypodermal cells enclose the embryo a slight ventral bend develops in the body (Fig. 2b). This first change in body shape probably is not related to the process of elongation in older embryos, and can be ascribed instead to the dorsal side of the embryo becoming longer than the ventral side as the dorsal cells interdigitate. At about 250 min the embryo begins to elongate along its anterior-posterior axis. Elongation takes about 130 min, in which time the embryo decreases in circumference about threefold and increases in length approximately fourfold (Fig. 3, also see Figs. 4a, c, e). Because there is a consistent relationship between the age and shape of an embryo during elongation, developmental stages can be described in terms of the physical length of the embryo. For example, 2~ embryos at 25°C have

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FIG. 4. Changes in body and hypodermal cell shape during elongation. The left-right sets of pictures are scanning electron immunofluorescence micrographs, respectively, of comparable embryonic stages before (a, b), in the early stages (c, d), and in the (e, f) of elongation. (a) The embryonic sheath covers the embryo during elongation (see also c, d) such that individual hypodermal cell as seen in Fig. 2b, are no longer visible from the surface. In later stages of elongation (e), ridges are visible over the dorsal hypodermal surface (arrowheads, see also Fig. 7c) but not over the lateral hypodermal surfaces (arrow). (b) The boundaries of the cells are visible in embryos stained with the MH27 antibody and viewed in the fluorescence microscope (compare with Fig. 2b). of shape change in a single hypodermal cell during elongation is shown in the series b, d, f. The small white arrowheads point to the margins of a lateral hypodermal cell, and the short white arrows point to the circumferential margins. Note that the relative hypodermal cells do not appear to change during elongation (long white arrow in b, d, f). The dorsal hypodermal cells (large white in b, d) fuse in the early stages of elongation. Calibration: 10 pm.

and MH27final stages boundaries, and ventral hypodermal An example longitudinal positions of arrowheads

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increased their length twofold and are about 350 min in age. Hypodermal Cells Are Responsible for Elongation The hypodermal cells of the C. elegans embryo are necessary for elongation: embryos are unable to elongate normally if hypodermal cells are destroyed by laser microsurgery (Sulston et al., 1983; see also Fig. 10). In contrast, many of the internal cells of the embryo are not necessary for elongation: embryos elongate normally following removal of groups of neurons or body-wall muscles by laser microsurgery (J. Sulston, personal communication). Extending these results, we find that elongating embryos continue to elongate following destruction of both pharyngeal and intestinal cells (data not shown). of The Distance between the Longitudinal Margins Each Hypodermal Cell Decreases during Elongation

The hypodermal cells change shape markedly during elongation. Before elongation the distance between the two longitudinal margins of each hypodermal cell is much greater than the distance between the two circumferential margins of the same cell (Figs. 2b, 4b). As the embryo elongates the distance between longitudinal margins decreases until in the fully elongated animal the longitudinal margins are much closer than the circumferential margins (compare Figs. 4b and f). Because the hypodermal cells are responsible for elongation and appear to be coupled together mechanically, it is likely that the mechanism that causes the hypodermal cells to change shape is responsible for the embryo changing shape, i.e., elongating. The cytoskeletal organization of the hypodermal cells was examined to investigate how their shapes are determined. The Longitudinal Margins of Hypodermal Cells Appear to be Linked by Circumferentially Oriented Actin Fibers The distribution of actin-containing fibers (actin fibers) in the hypodermal cells was analyzed with Rhphalloidin and FITC-phalloidin, fluorescent reagents that bind specifically to filamentous actin (Wulf et al., 1979; Faulstich et al., 1983). Both reagents give identical staining patterns, and staining is reduced dramatically after preincubation with an excess of unlabeled phalloidin (see Materials and Methods). The following results are based on observations of over 100 stained embryos. Up to an hour before embryos begin to elongate, actin fibers are organized in complex networks associated with the membranes of all cells (Fig. 5). Immediately preced-

FIG. 5. Actin fibers in cells of early C. elegans shown is part of one cell in a two-cell embryo phalloidin. Calibration: 10 pm.

embryos. stained

The regions with FITC-

ing and throughout elongation, each hypodermal cell contains circumferentially oriented actin fibers that appear to connect the two longitudinal margins of the belt desmosome surrounding the cell (Fig. 6a, large arrow). The actin fibers in the lateral cells do not become circumferentially oriented until after the fibers in the dorsal and ventral cells. The actin fibers appear to shorten as the distance between the two longitudinal margins decreases during elongation, suggesting that the fibers may be contractile. In some of the embryos examined the belt desmosomes were disrupted, perhaps from either mechanical or detergent damage; in these embryos the circumferentially oriented actin fibers appeared to be continuous with, or to intermesh with, actin fibers in the belt desmosome (Fig. 6c, large arrows). The patterns of actin fibers in the two ventral hypodermal cells appear to be mirror images; individual actin fibers in the right ventral cells are aligned precisely with actin fibers in the left ventral cells. In contrast, actin fibers in the lateral cells are not aligned with fibers in either the dorsal or ventral cells, and the spacing of actin fibers in the dorsal and ventral cells consistently appears more regular than in lateral cells. By the time the embryo hatches all hypodermal cells appear to contain only unordered actin fibers (Fig. 6d), similar to cells in embryos before elongation. The distribution of actin fibers also was examined by electron microscopy. Bundles of microfilaments are found at the apical surfaces of dorsal and ventral hypodermal cells during elongation (Figs. 7a,b). The bundles are oriented perpendicular to longitudinal sections through embryos, and parallel to tangential and cross sections; they are thus circumferentially oriented. The

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FIG. 6. Actin fibers in the hypodermal cells. Embryos were stained with Rh- or FITC-phalloidin as described under Materials and Methods. (a) An elongating embryo showing the circumferentially oriented actin fibers (large white arrow) in the hypodermal cells, and actin in the belt desmosomes surrounding the cells. The nonstaining hypodermal cells (small white arrows) also were stained after continued incubation in detergent. The longitudinal (black arrowheads) and circumferential (black arrows) margins of the same lateral hypodermal cells shown in Figs. 4b, d, f are shown. (b) An embryo exposed to cytochalasin D for 25 min before staining with FITC-phalloidin. The embryo has retracted in length as described in the text, and shows little if any staining of actin fibers in the hypodermis (arrowhead). Longitudinally oriented actin fibers are visible in muscle cells that underlie the hypodermis (arrow). (c) A hypodermal cell in a broken embryo showing the circumferentially oriented actin fibers apparently connecting to remnants of the belt desmosome. The small arrow points to an aggregate of the circumferentiallyoriented fibers. (d) An embryo after elongation and near hatching showing the apparently random organization of actin fibers in the hypodermal cells. The small black arrow points to the belt desmosome between a dorsal and lateral hypodermal cell. Calibration: 10 pm.

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FJG. 7. Transmission (a, b, d) and scanning (c) electron micrographs of hypodermal cells in 2.5~ embryos. All transmission micrographs are of longitudinal sections through the apical surfaces of dorsal or ventral hypodermal cells. (a, b) Apical region of a dorsal hypodermal cell at the convex surface (a) and of ventral hypodermal cell at a concave surface (b) of a curved body segment. Bundles of circumferentially oriented microfilaments (large black arrowheads) and circumferentially oriented microtubules (large white arrowheads) are associated with the apical membrane (small black arrow) of the cells. The embryonic sheath is the thin layer (small black arrowhead) overlying the cells (see Figs. 4a, c, e) (c) Surface view of folds (arrowhead) on concave surface of curved body segment. (d) Lower magnification showing the proximity of the embryonic sheath to the hypodermal apical membrane in regions overlying (small arrows) and adjacent to (small arrowheads) the microfilament bundles (large arrowheads). The white arrow points to one of the two circumferential margins of the belt desmosome surrounding the cell. Calibration: (a, b) 0.25 pm; (c) 5 wrn; (d) 0.5 pm.

microfilaments are about 7 nm in diameter, suggesting that they are likely to be composed of actin. The bundles are not observed in early embryos before elongation, or in late embryos after elongation. The bundles always are found immediately adjacent to the apical membranes of the dorsal and ventral hypodermal cells, suggesting that they may be attached to the membrane (Figs. 7a,b). While actin fibers are seen clearly in the lateral hypodermal cells after staining with Rh-phalloigin or FITCphalloidin, no microfilament bundles appeared to be associated directly with the surface membranes of the lateral cells in 7 embryos examined with the electron microscope. At present we do not know whether the failure to find microfilament bundles in the lateral cells was due to poor fixation, or represents a significant difference in the position or morphology of the microfilaments.

MicrojZament Function Is Required for Hypodermal Cells to Change Shape and
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min, but the fibers appeared to stain much less intensely than normal (data not shown). In 3 embryos exposed to the drug for 25 min, the hypodermal actin fibers were not visible (Fig. 6b). The effect of cytochalasin D on C. elegant embryos is reversible; all embryos treated with the drug were able to elongate, or to re-elongate, after the drug was removed (Figs. 8c, 9c). Most of the drug-treated embryos elon-

FIG. 8. Effect of cytochalasin D on embryos prior to elongation. (a) Two embryos at the same stage of development, About 1 min before photographing, the eggshell on the left embryo was permeabilized with the laser microbeam to expose the embryo to cytochalasin D in the medium (see Materials and Methods). The white arrowheads mark the diameter of the head. (b) After 45 min the embryo exposed to cytochalasin D has not elongated while the unexposed embryo on the right has more than doubled its length. (c) 60 min after washing out the cytochalasin D the left embryo has elongated about 2.5X and the head has decreased in diameter (white arrowheads). Calibration: 20 pm.

8a,b). No changes in the shapes of hypodermal cells were observed in 3 embryos processed for immunofluorescence with the MH27 antibody after ‘75 min of exposure to cytochalasin D (data not shown). If exposed to cytochalasin D during elongation, embryos retract within 15 min to almost their pre-elongation length and do not elongate further (Figs. 9a,b). To determine the effect of cytochalasin D on the hypodermal actin fibers, elongating embryos exposed to the drug for 15 and 25 min were stained with FITC-phalloidin and examined in the fluorescence microscope. Hypodermal actin fibers were visible in 4 embryos exposed to cytochalasin D for 15

FIG. 9. Effect of cytochalasin D on embryos during elongation. (a) The left embryo is beginning to elongate and is included here only to indicate the passage of time during the experiment. The right embryo is 2.5X, with the tail covering the head. About 1 min before photographing, the eggshell on the right embryo was permeabilized to expose the embryo to cytochalasin D in the medium. (b) After 15 min, the permeabilized embryo has retracted to nearly its original length before elongation, and its diameter (white arrowheads) has returned to nearly the size before elongation (compare with Fig. 8a). Note the position of the tip of the tail after retraction (white arrow). (c) 60 min after rinsing off the cytochalasin D, the right embryo has re-elongated to more than 2~ and has decreased once again in diameter. Calibration: 20 gm.

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gated 2.5X-3X, though normal embryos elongate approximately 4X. The body shape of fully elongated embryos near hatching (about 500 min) is not affected by treatment with cytochalasin D (data not shown). Because late embryos are covered by an extracellular cuticle that is impermeable to most compounds (Sulston et ab, 1983; Cox et al., 1982), the tips of the tails on the experimental animals were removed with a laser microbeam to ensure penetration of the drug (see Materials and Methods). Elongation of the Embryo Microjilament-Dependent

Is Associated with a Pressure on Internal

Cells

The results presented here suggest a model for elongation in which the microfilament bundles in the hypodermal cells contract, causing the hypodermal cells to constrict their apical surfaces circumferentially thus squeezing the internal cells of the embryo (see Discussion). This model predicts that internal cells should be subjected to pressure as the embryo elongates, and that the pressure should be sensitive to cytochalasin D. To test whether internal cells are subjected to pressure, a laser microbeam was used to create lesions in the hypodermal cells surrounding the embryo; if internal cells are under pressure they should burst out from such lesions. Internal cells do not burst from lesions created in hypodermal cells of embryos at any time before elongation (Figs. lOa,b). In contrast, internal cells burst within seconds from lesions created in hypodermal cells of embryos during elongation (Figs. lOa,b). Therefore an apparent pressure develops on internal cells during elongation. While internal cells do not burst from a hypodermal lesion in an embryo irradiated before elongation, some internal cells gradually emerge from the lesion as the embryo continues to develop (Fig. 10~). These embryos do not elongate; however, their heads and tails decrease in circumference (Fig. 10~). Because in this experiment the apparent pressure on internal cells develops without an intact cellular membrane surrounding the embryo, the pressure cannot be created osmotically. Therefore it seems likely that the internal cells are displaced by the mechanical force that causes the head and tail to decrease in circumference. To test whether microfilaments were involved in the generation of pressure on internal cells, hypodermal lesions were created in embryos that had been exposed previously to cytochalasin D. Internal cells never emerge from hypodermal lesions in embryos treated with cytochalasin D at any time before elongation (data not shown). Therefore cytochalasin D prevents the pressure from developing. In addition, internal cells do not emerge from lesions after elongating embryos are exposed to

FIG. 10. Development of internal pressure during elongation. (a) A 2.5X embryo (left) and an embryo prior to elongation (right). (b) Hypodermal cells in both embryos were ablated with a laser microbeam a few seconds before photographing. A large group of internal cells already have burst from the lesion (black arrowhead) in the left embryo, while no internal cells have emerged from the lesion (large white arrowhead) in the right embryo. Note the diameter (small white arrowheads) of the head of the right embryo. (c) 150 min later the left embryo has begun to lyse after extruding a large fraction of internal cells. While the embryo on the right has not elongated, the diameter of the head (small white arrowheads), and to a lesser degree that of the tail, have decreased as internal cells have emerged gradually from the hypodermal lesion (large white arrowhead). Calibration: 20 pm.

cytochalasin D and allowed to retract in length (data not shown). Because internal cells in elongating embryos normally are subjected to pressure, this result indicates that the pressure is abolished by treatment with cytochalasin D. The effect of cytochalasin D is reversible; a few internal cells gradually emerge from hypodermal lesions after removal of the drug (data not shown). In contrast to the apparent pressure that exists on internal cells during elongation of the embryo, the turgor pressure of postembryonic animals does not appear to be mediated by microfilaments. Nematode larvae in

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general are characterized by a high turgor pressure that is thought to play a role in locomotion (Harris and Crofton, 1957). Internal cells burst from lesions in the body and surrounding cuticle of newly hatched C. elegans larvae, as is the case for elongating embryos. However, pretreating larvae with cytochalasin D, after removing the tips of their tails with the laser microbeam to ensure permeabilization, does not prevent internal cells from erupting at surface lesions. Hypodermal Microtubules In addition to the bundles of microfilaments, two other structures are associated with the apical membranes of hypodermal cells during elongation that may play a role in elongation. The first is an array of circumferentially oriented microtubules described in this section, and the second is an extracellular layer, the embryonic sheath, described in the following section. The distribution of microtubules in the hypodermal

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cells was examined by indirect immunofluorescence with antibodies against either yeast tubulin (Kilmartin et al., 1982) or Drosophila cy-tubulin (M. Fuller, personal communication), which gave identical results. Patterns of microtubules in the hypodermis are difficult to resolve in whole-mount preparations of embryos because of the large amounts of tubulin in internal cells (see Fig. llc), so a procedure was developed for separating fragments of the hypodermis from internal cells for analysis (see Materials and Methods). The patterns of microtubules in the hypodermal fragments appear identical to the patterns visible in whole-mount preparations analyzed carefully with the fluorescence microscope. The following results are based on observations of between 50 and 100 embryos. In the period of embryogenesis before elongation, microtubules are found in complex cytoplasmic networks in interphase cells, or associated with the spindle apparatus in mitotic cells (Albertson, 1984). The microtubules in the lateral hypodermal cells remain as a net-

FIG. 11. Anti-tubulin staining of hypodermal fragments. All large arrowheads point to either dorsal or ventral hypodermal cells, and all large arrows point to lateral hypodermal cells. (a) Dorsal and lateral hypodermal cells in an embryo beginning to elongate. Essentially all microtubules in the dorsal hypodermis appear circumferentially oriented, in contrast to the lateral hypodermal cells where they appear to be oriented randomly. (b) Dorsal, lateral, and ventral hypodermal cells of an embryo during elongation. (c) Ventral or dorsal and lateral hypodermal cells of an embryo in a later stage of elongation. Some randomly oriented microtubules (small arrow) appear in the dorsal or ventral hypodermal cells. The white area on the right of the embryo is a region where internal cell did not separate from the hypodermis, and illustrates the difficulty of resolving microtubules in whole-mount preparations. (d) Dorsal, lateral, and ventral hypodermal cells of an embryo after hatching. Several circumferentially oriented microtubules remain in the dorsal and ventral cells, but many randomly oriented microtubules now are present. Calibration: 10 pm.

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work throughout embryogenesis (Fig. 11, large white arrows). However the microtubules in the dorsal and ventral hypodermal cells undergo a remarkable transition in pattern after the hypodermal cells have enclosed the embryo but before elongation begins; essentially all the microtubules in the dorsal and ventral hypodermal cells become circumferentially oriented (Fig. 11, large white arrowheads). Embryos in early stages of elongation appear to have the greatest density of circumferentially oriented microtubules, and embryos during and after elongation have a lower density and probably fewer total (compare Figs. lla and c). In the late stages of elongation and following elongation, cytoplasmic microtubules with various orientations also appear in the hypodermal cells (Figs. llc,d, small arrows). In elongating embryos the circumferentially oriented microtubules in the dorsal and ventral hypodermal cells all appear at the same focal plane in the fluorescence microscope, suggesting that the microtubules are associated with one of the hypodermal membranes. The microtubules in the lateral hypodermal cells are seen in several focal planes, and appear to be distributed in the cytoplasm. In electron micrographs of these embryos, numerous circumferentially oriented microtubules are found adjacent to the apical surface membranes of dorsal and ventral, but not lateral, hypodermal cells (Figs. 7a,b, white arrowheads). Cytoplasmic granules are excluded from a small region surrounding each microtubule, possibly representing space occupied by microtubule-associated proteins (Dentler et al., 1975). To test the function of the microtubules, embryos before, during, or after elongation were treated with the microtubule inhibitors colcemid, nocodazole, or griseofulvin. These compounds are unrelated chemically and appear to prevent microtubule assembly by binding to tubulin dimers (Wilson and Bryan, 1974; Wehland et al., 1977). The concentrations of colcemid and griseofulvin used in these experiments prevent mitosis in C elegans embryos (50 pg/ml colcemid, 20 pg/ml griseofulvin; Strome and Wood; 1983). While the microtubule inhibitors immediately block mitosis, presumably by preventing spindle microtubules from forming, studies in other systems indicate that they also can disrupt cytoplasmic microtubules (see Dustin, 1978). Between 10 and 20 C. elegans embryos were exposed to each of the microtubule inhibitors at various times before, during, and after elongation. Embryos treated with any of the microtubule inhibitors before or at any time during elongation continue to elongate, but do not elongate fully and instead develop a variety of surface abnormalities ranging from sharp constrictions to broad depressions (Fig. 12a). Surface constrictions and depressions do not form when embryos are pretreated

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with cytochalasin D, suggesting that they are caused by the microfilament bundles in the hypodermal cells. The surface morphology of fully elongated embryos near or after hatching is not affected by the microtubule inhibitors. To determine the effect of the microtubule inhibitors on the hypodermal microtubules, 9 elongating embryos were exposed to nocodazole (1 pg/ml) 1 hr and then processed for electron microscopy (see Materials and Methods). Almost no profiles of microtubules were visible in longitudinal sections through the hypodermals cells of any of the embryos examined, indicating that the microtubules had either broken down or were no longer circumferentially oriented (data not shown). While the pattern of hypodermal microtubules was altered significantly, the distribution of microfilament bundles appeared normal in these embryos. To examine further whether the microtubule inhibitors altered the pattern of actin fibers in the hypodermis, 15 embryos that had been exposed to nocodazole (1 pg/ml) for 2 hours were stained with FITC-phalloidin and examined with the fluorescence microscope. The pattern of actin fibers in these embryos appeared normal; there was no apparent aggregation of fibers in regions of the hypodermis where constrictions had developed (data not shown). The Embryonic

Sheath

The embryonic sheath is a distinct extracellular layer that is secreted over the surface of the embryo prior to elongation (Fig. 4a). The embryonic sheath appears to be attached to the hypodermal surface directly above the microfilament bundles: As the embryo elongates and moves within the eggshell, various regions of the body transiently bend and twist (see Fig. 4e). At the concave surface of any bend, the dorsal and ventral (but not lateral) hypodermal surfaces fold into multiple circumferentially oriented ridges (Fig. 7~). In sections through the surface, the troughs (Fig. 7d, black arrows) between the ridges are seen to correspond without exception to the location of microfilament bundles (Fig. 7d, large black arrowheads). The embryonic sheath invariably contacts the hypodermal membrane directly above each microfilament bundle, but not at the membrane on either side of a bundle (Fig. 7d, small black arrowheads). To determine what role, if any, the embryonic sheath performed in elongation, it would be desirable to test whether elongation is normal after removal of the sheath. We find that elongating embryos treated with the proteolytic enzyme trypsin at 5 pg/ml for 15 min develop lesions in their sheaths (see Fig. 12c, black arrowhead). Pre-elongation stages incubated with trypsin for up to 40 min show no evidence of cell dissociation or permeabilization (as assayed by FITC-phalloidin stain-

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FIG. 12. Effect of microtubule inhibitors (a) or trypsin (b, c) on elongation. (a) The embryo shown was permeabilized at the start of elongation, and has been in colcemid for 120 min (see Materials and Methods). While all embryos exposed to the microtubule inhibitors colcemid, nacodazole or griseofulvin developed constrictions, there was variability in both the type of constriction and the degree of elongation of the embryo. The embryo shown here is an example of an animal with broad surface depressions; other experimental embryos had acute constrictions similar to the embryo shown in (b) which was exposed instead to trypsin. (b) An embryo treated with trypsin for 15 min, rinsed with trypsin inhibitor, and allowed to develop for 120 min. Prominent furrows are seen on the dorsal surface of the animal (white arrows). (c) Electron micrograph of longitudinal section taken through the embryo in (b) showing a furrow (black arrow) within a hypodermal cell. The furrow almost reaches the basement membrane (white arrows) separating the hypodermal cell from underlying muscles. The embryonic sheath (black arrowhead) appeared disrupted in several regions. Calibration: (a, b) 20 Frn; (c) 0.5 pm.

ing of intracellular actin fibers, see Materials and Methods), though some cells are permeabilized in embryos incubated for 50 min and longer. Embryos before or during elongation were treated with trypsin for 15 min, then the medium was replaced with fresh culture medium containing excess trypsin inhibitor (see Materials and Methods). The treated embryos did not elongate, or continue to elongate, properly and instead developed prominent constrictions in the hypodermis (Fig. 12b), similar to constrictions observed in some embryos treated with microtubule inhibitors. Embryos processed for electron microscopy showed several hypodermal cells with one or more deep furrows extending through almost the entire cell body (Fig. 12c, black arrow). Hypodermal constrictions do not develop if the embryos are exposed to cytochalasin D and trypsin simultaneously (data not shown), suggesting that they are caused by contraction of the microfilament bundles.

The Cuticle Appears to Maintain Embryo following Elongation

the Shape

of the

During elongation, microfilaments maintain (and probably determine) the shape of the embryo because embryos rapidly change shape when exposed to cytochalasin D (seeFig. 9). However, fully elongated embryos near and after hatching do not change shape when exposed to the drug. Indeed, the hypodermal microfilaments, which seem the most likely candidates to maintain (as well as determine) the shape of the body during elongation, become disorganized sometime before the embryo hatches (see Fig. 6d). How then is the shape of the body maintained after elongation? After elongating, the embryo secretes an extracellular cuticle (Sulston et aL, 1983), and it is reasonable to expect that the cuticle performs an important role in maintaining the body shape at least after hatching; cuticles

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Kramer, J. Priess unpublished results). Of three sqt-3 alleles examined, sqt-3 (e2117) has the most severe phenotype, with embryos elongating normally and then retracting to nearly their original length before elongation (Fig. 13). sqt-3 (ezll?‘) is temperature-sensitive; at 15°C the embryos maintain the normal elongated shape, while at 25°C the embryos retract after elongation. No obvious ultrastructural differences in body cells or basement membranes are observed between sqt-3’ (e2117) embryos raised at 15 and 25°C. However, there is a significant difference in cuticle morphology; at 15”C, sqt-3 (e2117) animals appear to develop a normal cuticle (Fig. 14a), while at 25°C a major structural layer of the cuticle, the striated layer (Cox et al., 1982), appears to be missing in the mutant embryos that retract (Fig. 14b). In wildtype embryos a rudimentary striated layer is visible at about 430 min but not at 380 min (J. Priess, unpublished results), suggesting that the striated layer normally is formed near the time elongation is completed (see Fig. 3). DISCUSSION

Elongation of the Embryo by Circumferential Constriction of the Apical Hypodermal Surfaces The results presented in this paper indicate that the hypodermal cells of C. elegans contain numerous circumferentially oriented bundles of microfilaments associated with their apical membranes, that the microfilament bundles shorten as the hypodermal cells change shape as the embryo elongates, and that microfilament

FIG. 13. Development of sqt-3 (e.2117) embryos at 25°C. Large white arrowheads mark the diameter of the head, a small white arrowhead marks the anterior limit of the head, and a black arrowhead marks the posterior limit of the head at the junction with the intestine. (a) A sqt-3 embryo beginning to elongate. (b) A fully elongated sot-9 embryo. Only the head is visible in this focal plane; the tail lies on a lower focal plane and curls around the egg. Note how the head (measured from the small white arrowhead to the large black arrowhead) extends almost around the egg. (c) The same sqt-3 embryo after retraction (about 600 min in development). Now both the head and the tip of the tail are visible in the same focal plane. The length of the head is reduced almost by half of the length in (b), and the diameter of the head has increased appreciably. Calibration: 20 gm.

separated from body tissues still retain the general contour of the living animal (Cox et aZ., 1982). Analysis of sqt-S mutants in C, elegans supports the hypothesis that the cuticle is necessary to maintain the body shape after elongation. sqt-3 mutant embryos elongate normally, but subsequently retract in length (J.

FIG. 14. Electron microscopy of cuticles of sqt-3 (&?117) embryos raised at 15°C (a) or 25°C (b). The cuticle is the extracellular structure above the hypodermal cell membranes (white arrows). (a) The striated layer (black arrowhead) in the cuticle of 15’C embryo appears similar to wild-type. (b) At 25°C sqt-3 (e.2117) embryos retract after elongation (see Fig. 13) and the striated layer of the cuticle appears to be missing (black arrowhead). Calibration: 0.5 km.

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function is required for the hypodermal cells to change shape. Therefore it seems likely that the microfilament bundles in the hypodermal cells are contractile, though we have not determined whether proteins characteristic of contractile systems, such as myosin, are associated with the microfilaments. Contractile bundles of microfilaments have been found in nonmuscle cells in other organisms, for example the stress fibers of vertebrate fibroblasts (Isenberg et al., 1976; Kreis and Birchmeier, 1980). The circumferentially oriented actin fibers in C. elegans, which correspond presumably to the microfilament bundles, appear to attach to the belt desmosome surrounding each hypodermal cell, as cytoplasmic microfilaments have been shown to attach to desmosomes in other systems (Burnside 1971,1973; Bernstein and Wollman, 1976). If the belt desmosomes in C. elegans provide mechanical coupling between adjacent hypodermal cells, as seems likely, contraction of the actin fibers in individual hypodermal cells would force the embryo to decrease in circumference. The decrease in circumference should create pressure on the internal cells of the embryo, and this paper reports evidence that a microfilament-dependent pressure does exist during elongation. Internal cells would be squeezed radially as the circumference decreased, and the embryo would elongate as cytoplasm was displaced along the anterior-posterior axis. The Role of the Hypodermal Embryonic Sheath

Microtubules

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wrapped with contractile “bands.” The plates then might function to distribute evenly constrictive stresses resulting from contraction of the bands. If the circumferentially oriented microtubules are to resist microfilament contraction, the stress of contraction must be transmitted longitudinally from the microfilament bundles to the adjacent microtubules because almost all of the microtubules are between, rather than beneath, microfilament bundles (See Fig. 7a). While in electron micrographs no longitudinally oriented fibers appear to be associated with the hypodermal membrane, it is possible that such fibers exist but do not stain, or that the membrane itself is sufficient to transmit stress longitudinally. A further possibility is that the extracellular embryonic sheath functions to transmit stress longitudinally. The sheath is secreted before elongation, and appears to be attached to the hypodermal surface above each of the microfilament bundles, where stress is likely to be generated. The sheath could be compared to a tarpaulin draped across a scaffold of microtubules, as shown in Fig. 15. Constrictions develop across the hypodermal

and

Elongation conceivably could require only a closely spaced array of circumferentially oriented contractile fibers covering the surface of the embryo. If the rate of fiber contraction was coordinated properly in all regions of the body, the embryo should decrease uniformly in circumference. However, the observation that embryos do not elongate fully when exposed to microtubule inhibitors, and instead develop surface constrictions, suggests that microtubules also are involved in elongation. Microtubules are considered to be relatively stiff components of the cytoskeleton in many types of cells (cf. Tucker, 1977). Therefore it is possible that the numerous circumferentially oriented microtubules associated with the dorsal and ventral (but not lateral) hypodermal surfaces of the C. elegans embryo serve a structural role in strengthening these surfaces. The circumferential margins of the dorsal and ventral cells do not decrease in length as much as the circumferential margins of the lateral cells, consistent with the idea that the dorsal and ventral surfaces are relatively stiff. Mechanically, the embryo might be modeled as a cluster of cells sandwiched between two relatively rigid “plates” that are

FIG. 15. Schematic diagram illustrating some of the possible components involved in the generation and distribution of force at the surface of a dorsal or ventral hypodermal cell. A “scaffold” of microtubules (MT) is shown draped by a “tarpulin,” which could be the embryonic sheath (S) or hypodermal membrane, with attached “drawstrings” consisting of microfilament bundles (MF). Contraction of the microfilament bundles, which are attached to the belt desmosomes (not shown), pulls the sheath against the microtubules, thus distributing the force of contraction across the entire surface.

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cells if the embryonic sheath is disrupted by brief digestion with trypsin, consistent with the hypothesis that the embryonic sheath normally functions to transmit stress. However, the possibility remains that trypsin additionally affects the distribution of microtubules in the hypodermal cells, which also might result in hypodermal constrictions. Further resolution of the roles of the microfilaments, microtubules, and the embryonic sheath in elongation may be possible through analysis of mutants in C. elegant with abnormal morphology (H. Schnabel, R. Schnabel, and J. Priess, unpublished results). Mutants have been found that either fail to elongate or do not elongate fully as embryos, and thus may be defective in some aspect of the contractile machinery. Other mutants develop surface constrictions similar to wild-type embryos treated with either microtubule inhibitors or trypsin, and thus may be defective in microtubular organization or the embryonic sheath. The Genetic Control of Mwphogenesis The results presented in this paper suggest that the genetic program that determines body shape in C. elegans embryos involves gene products that specify cytoskeletal organization. A hierarchy of control seems likely: Actin-bundling proteins are involved presumably in assembling microfilaments into bundles in the hypodermal cells as in other systems (see Schliwa, 1981), and other a&in-binding proteins and microtubule-associated proteins may be necessary for the microfilament bundles and microtubules to associate with the hypodermal membranes. Additional gene products may control the pattern of association with the membrane such that the microfilament bundles are spaced regularly and both microfilaments and microtubules are circumferentially oriented. As described in this paper, there is variation in the cytoskeletal organization of the different hypodermal cell groups; the distribution of microtubules in the lateral hypodermal cells differs significantly from the distribution of microtubules in adjacent dorsal and ventral hypodermal cells, and these cell groups also may differ in microfilament pattern. Thus the gene products that establish a particular cytoskeletal pattern possibly are controlled at higher levels by genes involved in selecting which hypodermal cells adopt that pattern. While the basis for the distinctive cytoskeletal patterns in the groups of dorsal, lateral, and ventral hypodermal cells is not known, the availability of the complete embryonic lineage of C. elegant (Sulston et al., 1983) allows the origins of cells within each group to be compared. The lineage shows the lateral, dorsal, and ventral hypodermal cells are not generated as separate clones

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of cells. Instead some precursors generate both dorsal and lateral, or ventral and lateral, hypodermal cells at terminal divisions. Therefore it seems possible that the position, rather than ancestry, of a hypodermal cell may influence the pattern of cytoskeletal organization it adopts. Geometry

and Mwrphogenesis

Elongation in C. elegant embryos involves generating a force that can alter coordinately the shapes of a relatively large number of cells. As shown in this paper, the force appears to be generated at the outer surface membranes of hypodermal cells, so the total force acting on the body is proportional to the outer surface area of the embryo. However, the amount of cellular material that is deformed during elongation is proportional to the volume of the embryo. Because the ratio of surface area to volume must decrease with increased body size, it is possible that there is a mechanical limit to the size of an embryo that can elongate like C. elegant. The adult forms of different nematode species vary greatly in size, ranging from 1 to about 350 mm in length with corresponding variation in the number of body cells (Barnes, 1980). Yet in contrast to most organisms, all nematode

FIG. 16. Schematic diagrams comparing elongation of a small, closed cylinder of cells with folding by a plane of cells. The plane of cells also could represent a small section from a large cylinder composed of many cells. Both the small cylinder and the plane are composed of cells that contract their apical surfaces anisotropically (top). This apical contraction forces the small cylinder to elongate, while the planar array begins to fold.

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species produce eggs and embryos that are remarkably similar in size, between 50 and 90 pm in length (Bird, 1976). Perhaps in the evolutionary history of nematodes mutations that increased the size of eggs resulted in larger embryos that were unable to elongate into worms. While elongation might be precluded by increased size, increasing the number of hypodermal cells in a large embryo could create the potential for new morphogenetic patterns. With more cells, changes in hypodermal cell shape could lead to changes in the shape of the hypodermal layer itself, without deforming the entire embryo as in C. etegans. For example, part of the hypodermal layer might fold into a tube if each hypodermal cell contracted its apical surface anisometrically, as shown in Fig. 16. The ability of a layer of epithelial cells to change shape is a major feature in the morphogenesis of a wide range of organisms, for example in the development of the neural tube in vertebrates and insects. The formation of the newt neural tube and the subsequent expansions of the brain cavities all occur without growth and instead involve changes in the shape of a sheet of epithelial cells, the neural ectoderm, that is only one cell in thickness (Jacobson, 1978). Both experimental and theoretical studies have suggested that an important feature in the folding of the neural ectoderm is the ability of the individual cells to contract their apical surfaces (Glaser, 1914; Schroeder, 1970; Jacobson and Gordon, 1976; Ode11 et al., 1981). Thus the strikingly different morphogenetic processes of a spherical embryo elongating into a long thin worm, and a sheet of embryonic cells folding into a tube may have a similar cytomechanical basis. We thank K. Breitweiser, J. Cox, A. Fire, J. Hodgkin, J. Kramer, A. Staehelin, and especially J. S&ton and J. White for many interesting and helpful discussions in the course of this work, and A. Fire, J. Hodgkin, J. Sulston, B. Waterston, and J. White for a critical reading of the manuscript. We also are indebted to N. Thomson and N. Wolf for expert assistance with transmission electron microscopy. This work was supported in part by a National Science Foundation scholarship and a National Institutes of Health Fellowship, GM 09751, to J. P. and by U.S. Public Health Service Grant GM 19851. REFERENCES ALBERTSON, D. G. (1984). Formation of the first cleavage spindle in nematode embryos. Dev. BioL 101,61-72. BARNES, R. D. (1980). The Asehelminths. In “Invertebrate Zoology,” pp. 288-307. Holt, Rinehart & Winston, Japan. BIRD, A. F. (1976). The development and organization of skeletal structures in nematodes. In “The Organization of Nematodes” (N. A. Croll, ed.), pp. 107-137. Academic Press, New York. BRENNER, S. (1974). The genetics of Cuenorhabditti elegans. Genetics 77,71-94. BRENNER, S. L., and KORN, E. D. (1979). Substoichiometric concentrations of cytochalasin D inhibit actin polymerization. J. Biol. Chem. 254,9982-9985.

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BURNSIDE, B. (1971). Microtubules and microfilaments in newt neurulation. Dev. Biol 15,432-450. BURNSIDE, B. (1973). Microtubules and microfilaments in amphibian neurulation. Am. 2001. 13, 989-1006. Cox, G. N., KUSCH, M., and EDGAR, R. S. (1981). The cuticle of Caenorhabditis elegans: Its isolation and partial characterization. J. Cell Biol. SO, 7-17. DENTLER, W. L., GRANETT, S., and ROSENBAUM, J. (1975). Ultrastructural localization of the high molecular weight proteins associated with in vitro-assembled brain microtubules. J. Cell Biol. 65,237-241. DUSTIN, P. (1978). Microtubule poisons. In “Microtubules,” pp. 167225. Springer-Verlag, New York. FLANAGAN, M. D., and LIN, S. (1980). Cytochalasins block actin filament elongation by binding to high affinity sites associated with F-a&in. J Biol. Chem. 225, 835-838. FAULSTICH, H., TRISCHMANN, H., and MAYER, D. (1983). Preparation of tetramethylrhodaminyl-phalloidin and uptake of the toxin into short-term cultured hepatocytes by endocytosis. E~J. Cell Res. 144, 73-82. GLASER, 0. C. (1914). On the mechanism of morphological differentiation in the nervous system. Anat. Rec. 8, 525-551. HARRIS, J. E. and CROFTON, H. D. (1957). Structure and function in the nematode-Internal pressure and cuticular structure in Ascaris. J. Exp. Biol. 34, 116-130. HIRSH, D., and VANDERSLICE, R. (1976). Temperature-sensitive developmental mutants of Cuenorhabditis elegans. Dev. Biol. 46,220-235. ISENBERG, G., RATHKE, P. C., HULSMAN, N. FRANKE, W. W., and WOHLFARTH-BOTTERMAN, K. E. (1976). Cell Tissue Res. 166,427-431. JACOBSON, A. G., and GORDON, R. (1976). Changes in the shape of the developing vertebrate nervous system analyzed experimentally, mathematically and by computer simulation, J. Exp. 2001. 197,191246. JACOBSON, A. G. (1978). Some forces that shape the nervous system. In “Formshaping Movements in Neurogenesis” (Carl-Olof Jacobson and Ted Ebendal, eds.), pp. 13-22. Almqvist and Wiksell International, Stockholm. KILMARTIN, J. V., WRIGHT, B., and MILSTEIN, C. (1982). Rat monoclonal antitubulin antibodies derived by using a new nonsecreting rat cell line. J. Cell Biol. 93, 576-582. KREIS, T. E., and BIRCHMEIER, W. (1980). Stress fiber sarcomeres of fibroblasts are contractile. Cell 22.555-561. LLOYD, C. W., and BARLOW, P. W. (1982). The co-ordination of cell division and elongation-The role of the cytoskeleton. In “The Cytoskeleton in Plant Growth and Development” (C. W. Lloyd, ed.) pp. 203-228. Academic Press. London. ODELL, G. M., OSTER, G., ALBERCH, P., and BURNSIDE, B. (1981). The mechanical basis of morphogenesis. Dev. Biol. 85, 446-462. SCHIERENBERG, E., MIWA, J., and VON EHRENSTEIN, G. (1980). Cell lineages and developmental defects of temperature-sensitive embryonic arrest mutants in Cuenorhabditis elegoks. Dev. Biol. 76,141-159. SCHLIWA, M. (1981). Proteins associated with cytoplasmic actin. Cell 31,587-590 SCHROEDER, T. E. (1970). Neurulation in Xenopus laevti. An analysis and model based upon light and electron microscopy. J. Embryol. Exp. Morphol. 23,427-462. STAEHELIN, L. A. (1974). Structure and function of intercellular junctions Int. Rev. Cytol. 39, 191-283. STROME, S., and WOOD, W. B. (1983). Generation of asymmetry and segregation of germ-line granules in early C. elegans embryos. Cell 35.15-25. SULSTON, J. E., and HORVITZ, H. R. (1977). Postembryonic cell lineages of the nematode Caenorhabditis elegans. Dev. Biol. 56,110-156. SULSTON, J. E., SCHIERENBERG, E., WHITE, J. G. and THOMSON, J. N.

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(1983). The embryonic cell lineage of the nematode Cuenorhabditis elegans. Dev. BioL 100, 64-119. SULSTON, J. E., and WHITE, J. G. (1980). Regulation and cell autonomy during postembryonic development of Cuenorhabditis elegans. Dev. Biol. 78, 577-597. TANENBAUM, S. W. (1978). “Cytochalasins: Biochemical and Cell Biological Aspects.” Elsevier-North-Holland, Amsterdam. TUCKER, J. B., and MEATS, M. (1976). Microtubules and control of insect egg shape. J. Cell Biol. 71, 207-217. TUCKER, J. B. (1977). Spatial organization of microtubules. In “Microtubules” (K. Roberts and J. S. Hyams, eds.), pp. 315-359. Academic Press, London/New York. WARD, S., THOMSON, N., WHITE, J. G., and BRENNER, S. (1975). Electron microscopical reconstruction of the anterior sensory anatomy of the nematode Caenorhabditis elegans. .I Camp. Neural. 160,313-338. WEHLAND. J., HERZOG, W., and WEBER, K. (1977). Interaction of gri-

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seofulvin with microtubules, microtubule protein and tubulin. J. MoL BioL 111, 329-342. WENT, D. F. (1978). Oocyte maturation without follicular epithelium alters egg shape in a dipteran insect. J. Exp. 2001 205,149-X5. WILSON, L., and BRYAN, J. (1974). Biochemical and pharmacological properties of microtubules. Adv. Cell Mol. Biol. 3,22-72. WOLF, N., PRIESS, J., and HIRSH, D. (1983). Segregation of germline granules in early embryos of Cuenorhabditis elegans: An electron microscopic analysis. J. EmbryoL Exp. Morph,01 73,297-306. WOOD, W. B., HECHT, R., CARR, S., VANDERSLICE, R., WOLF, N., and HIRSH, D. (1980). Parental effects and phenotypic characterization of mutations that affect early development in Caenorhabditti elegans. Dev. Biol. 74, 446-469. WULF, E., DEBOBEN, A., BAUTZ, F., FAULSTICH, H., and WIELAND, T. (1979). Fluorescent phallotoxin, a tool for the visualization of cellular actin. Proc. Natl. Acad. Sci. USA 76, 4498-4502.