Development 111, 1-14 (1991)Printed in Great Britain © The Company of Biologists Limited 1991

Dynamic changes in the distribution of cytoplasmic myosin during

Drosophila embryogenesis

PAUL E. YOUNG, THOMAS C. PESACRETA and DANIEL P. KIEHART*

Department of Cellular and Developmental Biology, Harvard University, Biological Laboratories, 16 Divinity Avenue, Cambridge, MA02138, USA

* To whom all correspondence should be addressed

Summary

Dramatic changes in the localization of conventionalnon-muscle myosin characterize early embryogenesis inDrosophila melanogaster. During cellularization, myosinis concentrated around the furrow canals that form theleading margin of the plasma membrane as it plungesinward to package each somatic nucleus into a columnarepithelial cell. During gastrulation, there is specific anti-myosin staining at the apical ends of those cells thatchange shape in regions of imagination.

Both of these localizations appear to result from aredistribution of a cortical store of maternal myosin. Inthe preblastoderm embryo, myosin is localized to the eggcortex, sub-cortical arrays of inclusions, and, diffusely,the yolk-free periplasm. At the syncytial blastodermstage, myosin is found within cytoskeletal caps associ-ated with the somatic nuclei at the embryonic surface.Following the final syncytial division, these myosin caps

give rise to the myosin rings observed during cellulariz-ation.

These distributions are observed with both wholeimmune serum and affinity-purified antibodies directedagainst Drosophila non-muscle myosin heavy chain.They are not detected in embryos stained with anti-Drosophila muscle myosin antiserum or with preimmuneserum. Although immunolocalization can only suggestpossible function, these myosin localizations and thecoincident changes in cell morphology are consistentwith a key role for non-muscle myosin in poweringcellularization and gastrulation during embryogenesis.

Key words: cytoplasmic myosin, Drosophilaembryogenesis, cellularization, gastrulation, apicalconstriction, cell shape change.

Introduction

Non-muscle cytoplasmic myosins have been identifiedin eukaryotes throughout phytogeny. Presumably, theyparticipate in such diverse cellular motilities as intra-cellular vesicle movement, cytoplasmic streaming, cellsurface receptor capping and generation of corticaltension, cytokinesis, and cell shape changes (Pollard,1981; Warrick and Spudich, 1987; Spudich, 1989;Kiehart, 1990). Indeed, both antibody microinjectionstudies and molecular genetic approaches have demon-strated that myosin function is required for cytokinesis(Mabuchi and Okuno, 1977; Kiehart et al. 1982; deLozanne and Spudich, 1987; Knecht and Loomis, 1987).

We have previously identified a conventional non-muscle myosin isoform from Drosophila (Kiehart andFeghali, 1986), and have cloned and sequenced thegenes that encode its heavy and light chains (Kiehart etal. 1989; Ketchum et al. 1990; Chang et al. unpublisheddata). The heavy chain gene encodes a 6 kb transcriptthat is expressed in a developmentally regulatedfashion, with peaks of message accumulation at 4-12 h

of embryogenesis, early third instar larval and earlypupal stages. The gene is at polytene chromosome band60E9, and is distinct, by polytene location, by crosshybridization and by sequence analysis, from theDrosophila muscle myosin heavy chain gene at 36B(Bernstein et al. 1983; Rozek and Davidson, 1983;Kiehart et al. 1989; Ketchum et al. 1990).

The first insights into non-muscle myosin localizationduring Drosophila embryogenesis were made using anantibody against Sarcophaga flight muscle myosin(Warn et al. 1979, 1980). However, interpretation ofthese studies is limited by the use of a probe from aheterologous species, questions about myosin isoformspecificity, problems with background fluorescence,and a consideration of only those developmental stagesprior to gastrulation.

Here, we focus on the distribution of non-musclemyosin at cellularization and gastrulation during Dros-ophila embryogenesis. We also examine the myosindistribution at earlier stages, in order to address howthe pattern of myosin localization in these later stagesdevelops. We use antibodies specific for Drosophila

P. E. Young, T. C. Pesacreta and D. P. Kiehart

cytoplasmic myosin (Kiehart and Feghali, 1986) toestablish the localization of myosin in specimens fixedat various stages of embryogenesis. We review therelationship between the distribution of myosin andother cytoskeletal components in the embryo, particu-larly actin and spectrin, as determined by our owncolocalization studies and the staining patterns pub-lished by other investigators. Our observations suggestthat cytoplasmic myosin is in the right place at the righttime to contribute to force production in cellularization(cytokinesis) and various cell shape changes andmovements during gastrulation that are required forsuccessful embryogenesis. Our studies provide directevidence for a specific myosin localization in cellsundergoing an apical constriction coincident withchanges in cell sheet morphogenesis.

Materials and methods

AntibodiesPreparation of rabbit anti-Drosophila cytoplasmic myosin,muscle myosin and spectrin polyclonal sera has beendescribed previously (Kiehart and Feghali, 1986; Byers et al.1987). Anti-actin monoclonal antibody was a gift from DrJames Lessard and has been described elsewhere (Lessard,1988). Anti-human histone monoclonal antibody was pur-chased from Chemicon (Temecula, CA). Anti-tubulin mono-clonal antibody was a gift from Dr L. S. B. Goldstein.Affinity-purified rhodamine-labeled goat anti-mouse IgG,rhodamine-labeled goat anti-rabbit IgG, and fluorescein-labeled goat anti-rabbit IgG were purchased from HycloneLaboratories (Logan, Utah) and Tago Immunologicals(Burlingame, CA).

Affinity-purification of anti-cytoplasmic myosinpolyclonal serumRabbit anti-Drosophila cytoplasmic myosin antibody waspurified from polyclonal serum by elution from cytoplasmicmyosin absorbed to nitrocellulose using Pollard's (1984)modification of the method developed by Olmsted (1981), oron columns of Drosophila cytoplasmic myosin by modifi-cations of standard methods (Cuatrecasas and Anfinsen, 1971;Lutz and Kiehart, unpublished data). The concentration ofaffinity-purified antibody was estimated by a dye bindingassay (Smith et al. 1985) using bovine serum albumin as astandard.

Rhodamine-phalloidinA working solution was made by drying, under vacuum, 5jAof a 3.3 jiM stock of rhodamine phalloidin (Molecular Probes,Eugene, Oregon) in methanol and resuspending it in 500 [Aphosphate-buffered saline (PBS; prepared as described byKarr and Alberts, 1986).

Preparation of embryos for immunofluorescenceEmbryos from Drosophila melanogaster (Canton-S) flies werecollected at 25 °C on grape juice/agar plates at timed intervals(Elgin and Miller, 1978), rinsed off plates and dechorionatedby standard methods (KarT and Alberts, 1986). All sub-sequent fixation and immunostaining steps were performed atroom temperature by modification of a number of previouslydescribed methods (Zalokar and Erk, 1977; Mitchison andSedat, 1983; Karr and Alberts, 1986; Wieschaus and Nusslein-Volhard, 1986).

To verify patterns of contractile and cytoskeletal proteinlocalization, various fixatives, based on previously publishedmethods, were used. All gave similar results. They includedvarious formaldehyde concentrations (2.5, 3.7, 8% formal-dehyde; Karr and Alberts, 1986); a mixture of 4 ml 95%EtOH, 1 ml 50 % acetic acid, 1 ml formalin (from Zalokar andErk, 1977); and 90% methanol (Warn and Warn, 1986).

Embryos were fixed for 3-5 min in all cases. Exposure tofixatives, particularly formaldehyde-based fixatives, forlonger periods of time resulted in severely diminishedimmunofluorescent signal. The inclusion of 50 mM EGTA,pH7.5 in fixative solutions and in subsequent antibodyincubations gave improved immunofluorescent images.

Cryostat sectioning of embryosFixed embryos were prepared for sectioning as describedpreviously (Pesacreta et al. 1989). Sections were cut at athickness of 5 jim on a Minotome (IEC, Needham Heights,MA) cryostat. Staining solutions and times of incubation wereas detailed below for whole-mount embryos.

Antibody stainingEmbryos were blocked for 30min-lh in incubation solution(PBS+lOmgmn1 bovine serum albumin+0.4% TritonX-100). This concentration of Triton X improved staining andantibody penetration.

The solution was replaced with lml fresh incubationsolution containing an appropriate dilution of the primaryantiserum (1:1000), affinity-purified antibody (30figm\~l),rhodamine phalloidin (0.33 ^M), anti-tubulin monoclonalsupernatant (1:100), anti-histone monoclonal supernatant(1:1500) or Hoechst stain (ljUgml"1). Embryos were stainedwith each solution for lh.

Embryos were washed by resuspension in fresh incubationsolution for 1 h with three changes of fresh solution; incubatedfor 1 h in secondary antibody diluted 1:1000 with incubationsolution; and then washed again for at least lh with threechanges of fresh solution.

Observation of embryos and analysis of myosinlocalizationIncubation solution was replaced with a mounting solution of50 % glycerol in PBS. Embryos were allowed to equilibrate inthis solution at least 30 min prior to mounting. Embryos wereplaced on a microscope slide under a coverglass edged withsilicone grease to control the extent to which embryos wereflattened. Confocal (MRC-6000, Biorad, Cambridge, MA)imaging was performed on a Zeiss Axioplan microscope,using a 63 x lens (NA 1.40). Other images were obtainedthrough a Zeiss IM35 inverted microscope under NomarskiDIC optics with an epifluorescence attachment, using 16 x(NA 0.35), 40x (NA 0.75), and 63x (NA 1.4) lenses.Photobleaching of rhodamine-stained specimens was notsignificant for brief periods of observation. Photographs weretaken with an Olympus OM-2 camera mounted on the 35 mmport of the IM35 microscope using Kodak Tri-X filmprocessed with Diafine developer (Acufine, Inc., Chicago,IL). Prints of confocal images were obtained using a Sonyvideo printer (Park Ridge, New Jersey).

Prior to cellularization, Hoechst or anti-histone stainingwas used to identify the developmental stage of each embryo,as well as to determine the mitotic phase and number of nucleiper unit area at the embryonic surface. The number of nucleiunambiguously defined the number of mitotic cycles that hadoccurred during development.

In all experiments, the pattern of specific antibody stainingwas compared directly to specimens stained with appropriate

Non-muscle myosin in Drosophila development 3

controls of preimmune or non-immune sera. To avoidrepetition, we only include the control data for the non-muscle myosin distribution during the cellular blastodermstage. Levels of preimmune and non-immune stainingremained at a comparably low level for all stages analyzed.

Quantitative immunoblotsThe quantitative immunoblot procedure used for the esti-mation of relative levels of cytoplasmic myosin present inembryos at different developmental stages was as describedpreviously (Pesacreta et al. 1989). Immunoblots were per-formed as described by Kiehart and Feghali (1986) butdilutions of sera were in Tween buffer (150 HIM NaCl, 20 mMTris-HCl, pH7.5, 0.1% Tween-20, and 0.01% thimerosal,with a final concentration of 5 % goat serum included forblocking purposes).

Nomenclature'Non-muscle' and 'cytoplasmic' are used interchangeablythroughout the text to describe the conventional (myosin-II),Drosophila non-muscle myosin isoform.

We assume the reader has a basic knowledge of earlyDrosophila development. For detailed descriptions of Dros-ophila embryogenesis, please consult Sonnenblick, 1950 andCampos-Ortega and Hartenstein, 1985.

Results

The distribution of non-muscle myosin changes dra-matically as the Drosophila embryo develops. Thechanges are most dramatic during cellularization andgastrulation.

Cytoplasmic myosin is at the furrow canals duringcellularizationDuring cellularization, cytoplasmic myosin is localizedat the leading edge of the furrow canals. Thisdistribution is consistent with a key role in generatingthe changes in cell shape that transform the syncytialblastoderm into the columnar epithelium of the cellularblastoderm.

As cellularization begins, myosin pervades the cortexof the embryo and is concentrated at high levels in aring that encircles each nucleus (Fig. 1). These ringsappear to interconnect and form a network thatencompasses the entire periphery of the Drosophilaembryo. Sagittal optical sections that bisect these ringsand the nuclei localize myosin to a dot on either side ofeach somatic nucleus (Fig. 1A, sagittal section, arrow-heads).

As cellularization proceeds, membrane extendsdeeper and deeper between adjacent nuclei as thefurrow canals are displaced towards the embryo interior(Fullilove and Jacobson, 1971). Fluorescent imagesconfirm that the polygonal rings of myosin are preciselycoincident with the location of the furrow canal(Fig. 1). Little or no myosin associates with themembrane that trails outward from the furrow canal tothe embryonic surface.

Once the furrow canals have reached the base of thenuclei, they become more triangular or wedge-shaped,as the dots of myosin staining are transformed into

teardrops (Fig. 1G, arrowheads). Their form suggeststhat the furrow canals on either side of a given nucleusare being drawn closer together and that the myosinrings have contracted to become smaller than whenthey surrounded each nucleus. In en face views, thepolygonal rings of myosin coincident with the furrowcanals have become distinct circles of myosin staining(Fig. 1H, I). Interestingly, the circles have begun todetach from one another, and in some cases no longercontact each other directly, but rather are intercon-nected by a meshwork of myosin (Fig. 1H, I). Weconsider it likely that the nearly isomorphic contractionof this network assists in drawing the furrow canalsdeeper and deeper into the embryo.

Once the furrow canals invaginate to their maximumextent (approx. 30/an), myosin is associated with thebasal and immediately adjacent lateral aspects of eachnewly forming cell (Fig. 1J, arrowheads). The myosincircles contract further within the surrounding myosinmeshwork (Fig. IK, L), and surround the bridge ofcytoplasm that connects each forming cell with the yolkmass. These small connections persist into earlygastrulation (Rickoll, 1976). Thus, during this finalphase of cellularization, non-muscle myosin is displacedboth in the direction of, and perpendicular to, theadvancement of the furrow canals.

Actin is localized in a similar fashion duringcellularization (Warn, 1986; Pesacreta et al. 1989;Young et al. unpublished results). Double-stainedpreparations show that the region of highest actinconcentration at the furrow canal is coincident with themost concentrated regioq of myosin. However, incontrast to myosin, actin is also localized along thelateral borders of the forming cells. Interestingly,spectrin also stains the lateral margins of the cells butfails to localize to the leading edge of the furrow canals(Pesacreta et al. 1989). Instead, spectrin lags slightlybehind the furrow canals.

Specificity of anti-cytoplasmie myosin stainingPreimmune serum fails to detect any antigens whenused to stain immunoblots of whole embryo homogen-ates (Fig. 2A, lane 1). In contrast, affinity-purifiedantibodies (data not shown) and anti-cytoplasmicmyosin antiserum (Fig. 2A, lane 2) both recognize asingle 2O5xlO3Mr band when used to stain similarimmunoblots. Quantitative analysis (Fig. 2B) indicatesthat there is a fairly uniform level of non-muscle myosinpresent in embryos through early gastrulation, at ornear the level observed in unfertilized eggs. About fourhours following fertilization, there is a significantincrease in the level of myosin, coincident with anincrease in the level of transcript accumulation from thenon-muscle myosin gene (Kiehart et al. 1989).

Similarly, preimmune serum produces no specificstaining patterns in immunofluorescence on fixedembryo specimens (Fig. 3A). Affinity-purified anti-bodies (Fig. 3B) and whole anti-cytoplasmic myosinantiserum (Fig. 3C) produce localization patterns thatare nearly identical to each other, although there is aslight increase in the level of diffuse staining observed

4 P. E. Young, T. C. Pesacreta and D. P. Kiehart

Of

Fig. 1. Myosin is localized at the leading edge of the furrow canals during cellularization. Confocal fluorescent micrographsof whole-mount embryos, fixed and stained with anti-cytoplasmic myosin antiserum at different times during cellularization,are shown. Panels A, D, G, and J represent optical sagittal sections of different embryos, fixed at progressively later timesduring cellularization (A D) Slow phase of cellularization; (G J) rapid phase Arrowheads in the photograh i t tout

p p g y , p g yduring cellularization. (A, D) Slow phase of cellularization; (G, J) rapid phase. Arrowheads in the photographs point ospecific alterations in the staining pattern: in A, 'dots' on either side of each nucleus; in G, 'teardrops'; in J, basal andimmediately adjacent lateral aspects of each forming cell. Panels B, E, H, and K are optical en face sections of theembryos depicted in panels A, D, G, and J, respectively, at a depth of focus coincident with the location of the furrowcanals. Through-focal series verify that these sections are at the peak of staining intensity. Panels C, F, I, and L are highermagnification images of panels B, E, H, and K. Scale bar in A is 10/an, and is for panels A, B, D, E, G, H, J, and K.Scale bar in C is 5/an and is for panels C, F, I, and L.

6 P. E. Young, T. C. Pesacreta and D. P. Kiehart

Fig. 4. Myosin concentrates in the pole cells and cell sheet furrows during gastrulation. Panels A, B, and C reveal myosinlocalization in whole-mount embryos, fixed and stained for anti-cytoplasmic myosin at progressively later times duringamnioproctodeal invagination. (A) Pole cells have begun to be carried onto the dorsal surface of the embryo. Pole cellsand the underlying plate of cells stain brightly for cytoplasmic myosin. Arrowhead indicates region of increasedfluorescence along pole cell membrane. (B) In a slightly older embryo, cells have begun to invaginate. Pole cells still stainbrightly, as does the region of invagination. There is a particularly bright line of staining at the anterior margin of thedepression. (C) The pole cells have disappeared into the embryo interior. Myosin concentrates in the anterior margins ofthe amnioproctodeal invagination, with decreased levels at posterior regions. Scale bar is for all panels and is 10 [an.

change. Thus, there is a tight correlation betweenmyosin redistribution and the specific cell shapechanges that have been proposed to initiate theinfolding of furrows and drive cell sheet shape changes(see Discussion).

Cytoplasmic myosin at earlier stagesWe have localized myosin at earlier developmentalstages in order to understand how myosin getspositioned to participate in cellularization and gastru-lation.

Preblastoderm embryosThe egg cortex of the preblastoderm embryo is enrichedin cytoplasmic myosin. There is a well-defined zone ofmyosin approximately 1-2 pan thick that extendscompletely around the egg periphery (Fig. 6A; arrow-head). Warn et al. (1979) observed a similar corticaldistribution of myosin in preblastoderm embryos withan antibody against Sarcophaga muscle myosin. Theantiserum that we raised against Drosophila musclemyosin fails to detect this cortical layer of myosin.

There is a more diffuse pool of cytoplasmic myosin inthe egg periplasm (Fig. 6A, bracket). The level ofstaining is clearly above the background. Immunologi-cal probes directed against other cytoskeletal com-ponents (e.g. actin, spectrin and tubulin) also give asimilar level of diffuse periplasmic labeling that is abovethat observed with preimmune or non-immune controlsera (see Warn et al. 1984; Karr and Alberts, 1986;Pesacreta et al. 1989; also Young et al. unpublished

results). The yolk mass is relatively devoid of cytoplas-mic myosin staining (Fig. 6A), but there is somestaining of the cytoplasm that surrounds the embryonicnuclei as they migrate towards the embryonic periphery(data not shown).

Cytoplasmic myosin is also concentrated in arrays ofinclusions confined to the cortex and subjacent eggperiplasm (Fig. 6B, C; arrows). These arrays arelongitudinal distributions of particles that radiateoutward from either or both of the embryonic poles,and span the entire length of the embryo. Each arraycontains two types of stained inclusions: points ofstaining 1-2/xm across and spheres of staining approx.4/im in diameter. These arrays of inclusions disappearfrom the region of the somatic nuclei during thesyncytial blastoderm stage. However, they are stillobserved at the posterior embryonic pole in the regionof pole bud formation, at times when they are no longervisible across the rest of the embryo. Antisera againstother cytoskeletal components fail to detect these samearrays. For example, punctate actin staining occursuniformly across the entire embryonic surface and is notorganized in the longitudinal arrays seen with antimyo-sin (Karr and Alberts, 1986; Warn, 1986; Pesacretaetal. 1989).

Syncytial blastodermCytoplasmic myosin redistributes as the nuclei ap-proach the embryonic surface between the end ofnuclear cycle 9 and interphase of nuclear cycle 10. Thelongitudinal arrays of inclusions begin to disappear,

8 P. E. Young, T. C. Pesacreta and D. P. Kiehart

membrane. Our observations of living embryos (Kie-hart, Young and Inoue", unpublished observations andKiehart et al. 1990) indicate that these staining patternsare coincident with a period of increased movement andpole cell surface activity.

During pole cell formation, there is an increase in thelevel of cytoplasmic myosin around the base of eachpole bud, as it pinches off from the bulk of the embryo(Fig. 9A, arrowhead). Warn and colleagues (1985)document a comparable localization of actin duringpole cell formation. The co-localization of actin andmyosin in this region strongly suggests that theseproteins play a role in the cytokinetic event that

Fig. 6. Cytoplasmic myosin is concentrated in the cortexand in subcortical arrays in preblastoderm embryos.(A) Confocal sagittal section of fixed embryo stained forcytoplasmic myosin. Myosin is localized to a narrow bandimmediately apposed to the plasma membrane in the eggcortex (arrowhead). Myosin concentrates diffusely inadjacent periplasmic regions (bracket), with relatively littlestaining of the egg interior. Stained aggregates visible inoptical section (arrow) are seen more clearly in B and C.Abbreviations: p, periplasmic staining; c, cortical staining.(B and C) Whole-mount face views of fixed preblastodermembryos, stained for cytoplasmic myosin. Linear assembliesof anti-myosin-stained aggregates radiate outwards fromthe embryonic poles (B) and follow the length of theembryo (C). Such arrays contain both 'points' and 'spheres'of fluorescent staining. The spacing between adjacent linearassemblies appears fairly constant. There is a highconcentration of such aggregates at both embryonic poles(anterior and posterior). Scale bar is for all panels and is10/im.

separates the pole cells from the bulk of the syncytialblastoderm.

Myosin at later stagesWe have not investigated the myosin distribution indetail at stages later than early gastrulation. A diffusecytoplasmic distribution with some additional concen-tration in the cortex appears to characterize virtually allcells. Interestingly, appreciable amounts of myosinaccumulate in the developing nervous system (Fig. 10)to levels above those seen for surrounding tissues.

Discussion

Our data demonstrate that non-muscle myosin is in theright place at the right time to contribute to cell shapechanges that drive cellularization and gastrulation ofthe Drosophila embryo. We speculate that conven-tional non-muscle myosin is a key motor for thesemovements. The data establish the distribution ofcytoplasmic myosin in developing Drosophila embryosfrom egg deposition to early gastrulation. We obtainsimilar patterns of localization in whole-mount andsectioned embryos and in specimens prepared by avariety of different protocols with both formaldehyde-and alcohol-based fixatives (see Materials and methodsfor details). Immunoblot analysis (Fig. 2B) and devel-opmental Northerns (Kiehart et al. 1989) suggest thatearly changes in localization involve a reorganization ofa maternally derived myosin pool. After early gastru-lation (approx. 4h), zygotic transcription and trans-lation contribute new myosin to the existing pool.

The distributions that we document through thecellular blastoderm stage are consistent with prelimi-nary studies by Warn and his colleagues (Warn et al.1979, 1980). They used high concentrations of anantiserum against a muscle myosin isoform from aheterologous species. Anti-Drosophila muscle myosinantibodies that strongly label the developing muscu-lature of fixed Drosophila embryos fail to duplicate thepatterns that we have documented using our antibodies

Non-muscle myosin in Drosophila development

Fig. 7. Myosin is reorganized in the cortex when nucleiarrive during cycle 10. (A) Fixed whole-mount nuclearcycle 10 embryo stained with affinity-purified anti-cytoplasmic myosin antibody, face view. Myosin caps haveformed within the cortex and overlie each somatic nucleus.(B) Optical, sagittal section of fixed whole-mount embryoat nuclear cycle 10, stained with anti-cytoplasmic myosinantiserum. There is an accumulation of mydsin in thecortex above each nucleus (arrowheads), as verified in Cwhere the embryo is double-labeled with Hoechst toidentify nuclear position. Scale bar is for all panels and is20 urn.

against Drosophila cytoplasmic myosin (data notshown). We interpret the staining patterns seen byWarn et al. (1979, 1980) as resulting from a low level ofantibodies directed against one or more of the smallnumber of epitopes shared by muscle and cytoplasmicisoforms of Drosophila myosin (for cross-reactivity ofthe anti-cytoplasmic and anti-muscle myosin sera usedin this study, see Fig. 7 of Kiehart and Feghali, 1986; forsequence similarities between cytoplasmic and musclemyosin isoforms, see George et al. 1989 and Ketchum etal. 1990).

Myosin in cellularizing embryosThe dramatic reorganization of myosin during cellular-

Fig. 8. Myosin is associated with the plasma membrane ofsurface caps during syncytial blastoderm stages (nuclearcycles 11-13). (A) Confocal sagittal section of earlysyncytial stage embryo, fixed and double-labeled with anti-cytoplasmic myosin antiserum and anti-histone antibody.Myosin caps (arrowheads) are immediately apposed to theplasma membrane and directly overlie each interphasenucleus (arrows). One of the nuclei is out of the plane offocus. (B) Confocal en face image of an embryo fixed andstained for cytoplasmic myosin at a later syncytialblastoderm stage, at a depth of focus just interior to theoutermost surface of the surface caps. Myosin forms a'ring' around each nucleus (nuclei are not stained in thispanel), observed both in caps that do, and do not, contactone another. Regions of the embryonic surface betweencaps are relatively devoid of staining. Scale bar is for bothpanels and is 10/tm.

ization is consistent with an active role for thischemomechanical force producer in the shape changesthat transform the syncytial blastoderm into the cellularblastoderm. The localization of myosin around theinvaginating furrow canals, the coexistence of actin atthe furrow canals that we have observed and that hasbeen described by others (Warn et al. 1984; Warn, 1986;Pesacreta etal. 1989; Warn etal. 1990), and experimentsthat show anti-myosin injection blocks cellularization(Lutz and Kiehart, 1987; and unpublished data), areconsistent with a role for myosin in force production forcellularization. The changes in distribution parallel therecruitment of myosin to the contractile ring asevidenced by antibody localization studies in dividingcells over a wide range of phyla from slime molds(Dictyostelium; Yumura and Fukui, 1985) to mammals(Fujiwara and Pollard, 1976; Nunnally et al. 1980;reviewed in Conrad and Schroeder, 1990). In culturedcells, such studies have been confirmed by elegant,fluorescent myosin light chain microinjection studiesthat allow the changing distribution of myosin to befollowed in living cells (Mittal et al. 1987).

It is generally accepted that myosin plays a keyfunctional role in powering cytokinesis (Mabuchi andOkuno, 1977; Kiehart et al. 1982; de Lozanne and

10 P. E. Young, T. C. Pesacreta and D. P. Kiehart

Fig. 9. Elevated levels of myosin are present in the polecells and region of pole cell formation. (A) Cryostatsection at posterior pole of fixed embryo at nuclear cycle12, stained for cytoplasmic myosin. There is appreciablemyosin in the pole cell cytoplasm (arrow), at levels higherthan that observed within the surface cap cytoplasm. Anaccumulation of myosin is also detected at the base of thispole cell (arrowhead), where it is pinching off from therest of the embryo. (B) Face view at posterior pole of fixedwhole-mount embryo, nuclear cycle 12, stained forcytoplasmic myosin. Cytoplasmic myosin accumulates atthe margins of some pole cells, particularly in regions ofcontact (arrowhead). Scale bar is for both panels and is

Spudich, 1987; Knecht and Loomis, 1987) and thatmyosin, with actin, is localized in a band of thick andthin filaments known as the contractile ring. Contractilering function in animal cell cleavage is believed to bethe consequence of the sliding of actin filaments,anchored to the membrane or cortex and mediated bybipolar myosin filaments. Thus, the shortening contrac-tile ring behaves as a purse-string, serving to constrictthe cell at the furrow in order to partition the cytoplasmfollowing mitosis. However, the ultrastructural organiz-ation, assembly, regulation, function and disassemblyof the contractile ring are not understood in detail.Therefore, a complete understanding of the mechanismof myosin function in force production during cellulariz-ation, specifically, and cytokinesis, in general, remainselusive.

Cytokinesis of the cellular blastoderm is topologicallycomplex. However, all the elements of contractilityappear to be present and it is likely that the overallmechanism is comparable to cytokinesis in other cells.Indeed in Drosophila, a network of some 6000contractile rings are organized so that they form acontinuous ring around the entire embryo. The radial

Fig. 10. Myosin accumulates in the ventral nervous system.Confocal sagittal section of fixed embryo stained forcytoplasmic myosin. Myosin concentrates in the fiber tractsand the commissures. Scale bar is 20 fun.

constrictive force of each ring, being opposed byadjacent rings, results in movement directed toward theembryo interior. Thus it is formally possible that asimilar, purse string-like mechanism, that works inconcert with a contracting actomyosin mesh work,serves to mediate cytokinesis during cellularization. Wespeculate that this complex arrangement of actomyosin,in rings connected by a meshwork, is required for thesimultaneous cytokinesis of the 6000 cells of the cellularblastoderm. As in other systems, it is likely that therings and meshwork form numerous attachments to thecell surface so that localized contractile function caneffect cellularization in part of the embryo, despitedisruption of the contractile machinery in other regions.

A major feature of cellularization that distinguishes itfrom cytokinesis in simpler cells is that the netdisplacement of membrane occurs in two directions,orthogonal to one another. First, the furrow canalsmust move some 30-40fim into the embryo, therebylaterally segregating each nucleus from its adjacentneighbors. Then, towards the end of this movement,the contractile ring at the base of each forming cell mustconstrict in a direction orthogonal to its originalmovement and parallel to the surface of the embryo, soas to form the basal margins of each columnar cell. Ourdata suggest that this transition in the direction ofmovement begins at the end of the slow phase ofcellularization (Mahowald, 1963), when the furrowcanals have reached the base of the nuclei. En faceimages of embryos, at a depth of focus coincident withthe furrow canals, indicate a transition in the myosinnetwork that occurs roughly when the furrow canalsreach the base of the nuclei (see Fig. 1). The

Non-muscle myosin in Drosophila development 11

interlocking rings of myosin that encircle each nucleusno longer make direct contact with each other, butrather appear separated from one another, heldtogether in a meshwork of more diffuse myosinstaining. During the fast phase of cellularization thatfollows, these rings decrease in diameter, suggesting aconstriction that could serve to pinch off the newlyforming cells.

The striking change in the speed of progress of thefurrow canals at the slow phase-fast phase transition isconsistent with a change in the direction or generationof force within the embryo at this time. We believe thatthe coincident rearrangement of the actomyosinnetwork supports its direct involvement in cellulariz-ation; however, the interactions between the variouselements of the contractile apparatus that allow thesecomplicated changes to occur synchronously and inprecise spatial register remain a mystery (see Merrill etal. 1988 and Wieschaus and Sweeton, 1988 formutagenesis studies that begin to address how cytoskel-etal and cellular elements may interact during cellular-ization).

Myosin in gastrulating embryosWe observe an accumulation of myosin at the apicalends of cells within furrows in cell sheets during thegastrulation of the Drosophila embryo. We speculatethat myosin is recruited to the apical region of thesecells and contracts in a purse-string fashion to initiatecell shape changes that lead to the displacement of thecell sheets. Such changes are similar to those observedduring amphibian gastrulation, in which it is thoughtthat apical constriction results in the bending ofepithelial sheets (e.g. Hardin and Keller, 1988), andsupport computer-generated models (e.g. Odell et al.1981) that demonstrate that an apical constriction candrive the infolding of cell sheets. Treatment of embryosof another Dipteran species with microfilament-destabi-lizing drugs results in a complete arrest of allgastrulation movements (Kaiser and Went, 1987).

Our localization of myosin to the apical ends of cellsduring early gastrulation is novel. The coexistence ofactin at the cell apices in these regions in Drosophila(Callaini, 1989) is consistent with the involvement of anactomyosin network in producing an apical constrictionin regions of invagination. Other studies have localizedmyosin (e.g. Lee et al. 1983) and other cytoskeletalproteins (actin, e.g. Sadler et al. 1982; o--actinin, e.g.Lash et al. 1985) to the apical ends of cells duringvertebrate neurulation. Although actin often shows afar more uniform accumulation to the apical ends ofcells across the embryo, myosin is fairly restricted to theapical ends of cells in regions undergoing an apicalconstriction, and subsequent cell sheet infolding.However, the data from these studies do not permitprecise correlation of myosin localization with indi-vidual cell shape changes in the region of infolding. Ourresults unambiguously demonstrate that an apicalmyosin localization is restricted to those cells that areundergoing or have just undergone apical constriction.Indeed, the tight temporal correlation between cell

shape change and myosin localization fuels our specu-lation that non-muscle myosin plays an active role inforce production for these movements. We attribute theclarity of our results to our use of high-affinityantibodies against Drosophila cytoplasmic myosin, ourfixation conditions, and the powerful optical sectioningcapabilities of the confocal microscope.

Origins and regulation of spatial heterogeneity ofmyosin during cellularization and gastrulationChanges in myosin localization during cellularizationand gastrulation likely reflect reorganization of myosinwithin the cells participating in a given movement. It islikely that complex regulatory cascades dictate theprecise changes in myosin localization and functionwithin regions of invagination (for review of myosinregulation, see Sellers and Adelstein, 1987). Forexample, the solubility of non-muscle myosin in vitro,and potentially the subcellular localization in vivo, isinfluenced by the phosphorylation state of myosinregulatory light chain. At least three kinases canphosphorylate the light chains and influence theirfunction. The activity of these kinases is in turnregulated by a variety of soluble factors, includingcAMP, free Ca2+ concentration and the stage of the cellcycle. Thus, such post-translational regulation ofmyosin function is likely to be quite complex. Ofcourse, we cannot rule out additional regulation ofmyosin distribution due to changes in myosin transcrip-tion or translation in specific regions of the embryo.

Myosin in earlier embryosWe speculate that the subcortical inclusions that stainwith anti-myosin in preblastoderm embryos represent astorage form of myosin that is recruited during myosincap formation at nuclear cycle 10. In Drosophila andother organisms, similar inclusions have been observedthat contain other cytoskeletal proteins (Strome, 1986;Warn, 1986; Weisenberg et al. 1987) and may alsorepresent a storage form of these proteins. An alternateinterpretation of these staining patterns is that theyrepresent an association of myosin with vesicularinclusions, as data point towards association of bothconventional and mini-myosins to membranes andvesicular inclusions in other systems (Adams andPollard, 1986; Berrios and Fisher, 1986; Grolig et al.1988; Adams and Pollard, 1989; Tang et al. 1989).

The cortex of the early Drosophila embryo isenriched in cytoplasmic myosin, as it is for a number ofcytoskeletal components including actin and spectrin(Warn et al. 1984; Karr and Alberts, 1986: Pesacreta etal. 1989). However, in contrast to actin, myosin doesnot localize to discrete foci believed to be surfacemicrovilli, a distribution consistent with the absence ofconventional, non-muscle myosins from microvilli inother systems (reviewed by Mooseker, 1985).

We speculate that the incorporation of myosin intothe cytoskeletal caps associated with each somaticnucleus of the syncytial blastoderm represents recruit-ment of myosin already present in the egg cortex,periplasm and, potentially, the subcortical arrays of

12 P. E. Young, T. C. Pesacreta and D. P. Kiehart

inclusions. The steady state levels of myosin areconstant at this time, and, because little or no myosinheavy chain message is present before 4-12 h ofdevelopment, we think it is unlikely that new proteinsynthesis contributes to the pattern of localization. Suchrecruitment parallels the redistribution of actin andspectrin. Signals that presumably originate in thecentrosomes (Rappaport, 1986; Raff and Glover, 1989)appear to play a key role in organizing the actinnetwork, but at this time remain a mystery.

The localization of myosin in the cortex of theDrosophila embryo prior to the onset of cellularizationcontrasts the lack of myosin in the cortex of otherinterphase cells where myosin is apparently recruitedjust seconds before its function in cytokinesis (e.g.Fujiwara and Pollard, 1976; Mittal et al. 1987;Schroeder, 1987; Schroeder and Otto, 1988). Perhapsmyosin plays additional roles in maintaining corticalstructure and organization in the Drosophila embryothat it does not play in other species. For example, thiscortical myosin may contribute to the establishment andmaintenance of nuclear positioning at the embryonicsurface. Experiments designed to disrupt myosinfunction or to disrupt microfilaments in Drosophilaembryos during syncytial blastoderm (by the micro-injection of antibodies or microfilament-destabilizingdrugs) result in the loss of nuclear positioning at theembryo surface (Zalokar and Erk, 1976; Foe andAlberts, 1983; Edgar etal. 1987; Lutz and Kiehart, 1987and unpublished data).

Myosin at later stages of developmentThis paper does not address in detail the distribution ofnon-muscle myosin in embryos following early gastru-lation. However, if myosin indeed plays an active rolein the cell shape changes that drive gastrulation, webelieve that later stages of development will becharacterized by specific myosin distributions associ-ated with other cell sheet rearrangements. Forexample, the strong localization of non-muscle myosinto the nervous system, and similar localizations ofmyosin and actin in the growth cones of neurons ofvarious organisms in primary culture (Kuczmarski andRosenbaum, 1979; Bridgman and Dailey, 1989;Forscher and Smith, 1988), are consistent with a role foran actomyosin contractile network in neurite growthand extension.

We thank F. Kuebler for help with antibody characteriz-ation and affinity-purification; M. Shea, A. Ketchum, X.-j.Chang, T.-l. Chen, K. Edwards, A. Richman, G. Thomas, C.Schmidt, K. Guthrie, S. Bukata, J. Glazeroff, S. Mones, R.Warn, D. Branton, L. Goldstein, F.C. Kafatos, and D. Beggfor their stimulating discussions and helpful suggestions; K.Guthrie for assistance with embryo preparation and staining;J. Minden for suggesting anti-histone staining for visualizationof nuclei on the confocal microscope; the reviewers for helpfulsuggestions and comments; and D. Begg for critically readingan early version of the manuscript. Preliminary accounts ofthis work appear in abstract form (Young et al. 1987) and arereferred to in a review on Drosophila cytoplasmic myosin(Kiehart et al. 1990). Funding: National Institutes of Health

Award No. GM33830 to DPK and an NSF PredoctoralFellowship to PEY.

This work is to be submitted in partial fulfillment of therequirements of Harvard University for the degree of Doctorof Philosophy for Paul Young.

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(Accepted 10 October 1990)