Correspondence to: Thomas Lecuit, Department of Molecular Biology, Princeton University, Washington Rd., Princeton, NJ 08544. Phone: (609) 258-5401. Fax:(609) 258-1547 E-mail:tlecuit{at}molbio.princeton.edu.
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Abstract |
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Cellularization of the Drosophila embryo is a specialized form of cytokinesis that results in the formation of a polarized epithelium. The mechanisms of membrane growth during cytokinesis are largely unknown. It is also unclear whether membrane growth and polarization represent distinct processes that occur simultaneously or whether growth of the membrane is involved in the emergence of polarity. Using a combination of surface labeling and particles tracking techniques, we monitored the dynamics of marked membrane regions during cellularization. We find that the major source of membrane is intracellular, rather than in the form of a plasma membrane reservoir. Depolymerization of microtubules inhibits the export of a newly synthesized transmembrane protein from the Golgi apparatus to the plasma membrane and simultaneously blocks membrane growth. Membrane insertion occurs in a defined sequence at specific sites, first apical, then apicallateral. Diffusion of the membrane appears insufficient to compete with the massive local insertion of new membrane. We thus identify a tightly regulated scheme of polarized membrane insertion during cellularization. We propose that such a mechanism could participate in the progressive emergence of apicalbasal polarity.
Key Words: cytokinesis, polarity, membrane growth, epithelial cells , cellularization
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Introduction |
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An important challenge in cell biology is to understand the mechanisms underlying the establishment and maintenance of cell polarity. Many different cell types show a polarized organization reflected in the asymmetric distribution of proteins and lipids in the plasma membrane and the asymmetric organization of the microtubule and actin cytoskeleton. This organization is key to their functional specialization. For example, it is essential during polar bud growth in the yeast Saccharomyces cerevisiae, polarized growth during axon formation, and the transport of nutrients across endothelial barriers. Some cases involve the imposition of polarity on previously existing membranes whereas others are associated with new membrane growth. In spite of obvious differences in the process, many components of the polarization pathway are shared across species and the mechanisms have been the subject of intense cell biological and biochemical investigations (for a review see
Cellularization of the Drosophila blastoderm is a remarkable process where the formation of a polarized epithelium is coupled to the process of cytokinesis. It offers an opportunity to understand the mechanisms of membrane growth during furrowing. It also allows one to ask whether membrane growth can contribute to the emergence of membrane polarity or whether these represent distinct processes. The first 13 nuclear divisions of the Drosophila embryo occur in a syncytium, resulting in 6,000 peripheral nuclei located beneath the plasma membrane. During cellularization, the membrane surface increases 25-fold, invaginates between the nuclei, and ultimately yields 6,000 epithelial cells 30 µm tall. Most studies so far have focussed on the cytoskeletal rearrangements that control cellularization (
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At the onset of gastrulation, the resultant epidermal cells have the hallmarks of polarized cells with adherens junctions separating the apical and basallateral domains. It is still unknown how this polarity is established. It may occur during cleavage of the embryo when the membrane grows and invaginates between the nuclei or, following the classical example of epithelial Madin-Darby canine kidney (MDCK) cells, it could occur after cellularization when contacts between cells trigger the formation of adherens and septate junctions. Nonpolarized MDCK cells use E-cadherinbased cellcell contacts as an external cue to position and to trigger the assembly of adherens junctions (
The work presented here uses Drosophila to address whether the polarization of epithelial cells can be governed by membrane growth under situations of rapid membrane mobilization. In other words, is polarity reflected in the regulated insertion of proteins and associated lipids at different locations as the membrane grows, or does polarity appear subsequently when nonpolarized cells interact in a manner akin to MDCK cells? To address these issues, we developed techniques that allow the visualization of the flux of membrane proteins in living embryos during cellularization. We compared these dynamic patterns to the distribution of a newly synthesized transmembrane protein in stage-fixed embryos. We conclude that growth of the plasma membrane stems from the remobilization of ER- and/or Golgi-derived membrane populations that insert at precise locations in a regulated manner. The localized membrane delivery we identify in our experiments is such that lectin-labeled membrane patches and microbeads bound to an intact membrane are consistently displaced away from the sites of insertion. This suggests a mechanism in which polarization of the plasma membrane might be inherently linked to the polarized pattern of membrane growth during embryonic cleavage.
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Materials and Methods |
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Fly Strains
OregonR stocks were used as wild-type controls in all of our experiments. To generate embryos zygotically deficient for Neur from mothers with a wild-type Neur dosage, we crossed C(3)se females with males heterozygous for Df(3L)81k19.
Surface Label of Living Embryos with Wheat Germ Agglutinin
Embryos were collected at 25°C on agar plates after 30-min collections. Embryos were aged 90 min at 25°C, dechorionated in 30% bleach for 2 min, rinsed with water, and lined up on a strip of agar. Embryos were then transferred on a coverslip covered with a thin layer of glue. After dehydration, the embryos were covered with S700 halocarbon oil where they continued to develop normally.
At the desired stage, embryos were injected with lectin as follows. Wheat germ agglutinin (WGA) coupled to the Alexa488 fluorochrome (Molecular Probes, Inc.) was kept as a 1 mg/ml stock solution in PBS at -20°C. WGA was thawed, diluted 1:31:5 in PBS, and centrifuged at 14,000 rpm for 10 min. It was loaded in a fine capillary needle and injection was conducted in the perivitelline space at 50% egg length (further information available upon request from the authors at tlecuit@molbio.princeton.edu). The coverslip was mounted on an inverted Zeiss LSM510 confocal microscope where embryonic development proceeds normally. All images were processed using Adobe Photoshop® software.
Coating, Injection, and Tracking Fluorescent Microspheres in Living Embryos
0.5-µm YellowGreen fluorescent carboxylated microspheres (Polysciences, Inc.) were coated with WGAAlexa488 using the carbodiimide kit for covalent coupling (Polysciences, Inc.). WGA coated beads were diluted in PBS and injected in the vitelline space and followed under the confocal microscope. YellowGreen fluorescent beads were detected using a relatively large pinhole to be reliably tracked in 2-µm optical slices. The maximum area of contact between the beads and the membrane was calculated to be 0.25 x 0.25 x 3.14 = 0.19 µm2. The membrane surface at the onset of phase 3 was largely underestimated, with the approximation to a smooth cylinder 6.5 µm in diameter and 5 µm high: 2 x 3.25 x 3.14 x 5 = 102 µm. This gave a minimum estimate of the relative surface of 1/537.
Antibody Staining
Antibody staining against Neurotactin (Neur), Armadillo (Arm), and myosin required heat fixation, which was described in
In all cases, antibody staining was performed in BBT (PBS/0.1% BSA/0.1% Tween 20), except for an initial blocking step with 10% BSA in PBS and 0.1% Tween 20. Primary antibodies were incubated overnight at 4°C with the following dilutions: rabbit anti-Dlt 1:1,000 (
Colcemid Injections
Dechorionated embryos were lined up and dehydrated as described above. Colcemid (Sigma-Aldrich) was stored as a 5 mM stock solution at 4°, diluted in water to 1.25 mM. Injection with an Eppendorf transjector allowed a reproducible result with an estimated final concentration 1225 µM around the middle of the embryo. The effect was uniform around the circumference of the embryo at the injection site. Lower concentrations of colcemid showed variable effects and microtubules were not depolymerized reproducibly (data not shown). Injection was performed at 50% egg length at 1618°C.
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Results |
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Four Phases in the Formation of a Polarized Epithelium
Cleavage of the syncytial blastoderm begins on entry into interphase of the 14th nuclear cycle and lasts 70 min at 20°C. Living embryos can be followed with differential interferance contrast (DIC) microscopy as shown in Fig 1. Immediately after mitosis 13 nuclei, reappear as spheres 5 µm in diameter beneath the plasma membrane (Fig 1 A, arrow). The cell surface above each nucleus protrudes in a dome shape known as a somatic bud, a feature of the membrane during interphase 914 (
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The initial stages of cellularization can be followed with greater accuracy using a membrane-attached PDZ-containing protein, Dlt (
The Furrow Canal Is a Distinct Membrane Region Formed in Phase 1
To understand how the polarization of the membrane arises and how to characterize the sites of new membrane insertion, we developed a labeling technique in living embryos. We labeled the plasma membrane by injecting the fluorescent lectin WGA in the perivitelline space of a living embryo. Under physiological conditions, WGAAlexa488 is a heterodimer that selectively binds to N-acetylglucosamine and N-acetylneuraminic acid (sialic acid) residues found on numerous membrane glycoproteins. When WGAAlexa488 is injected, only a very small area of plasma membrane is labeled and occurs within seconds after injection, effectively generating a localized pulse of labeled membrane. No WGA is detected in a free unbound form in the vitelline space. This contrasts with other fluorescent lectins, such as soybean agglutinin, which poorly binds to the membrane, diffuses around the entire circumference of the embryo, and remains unbound in the vitelline space (data not shown). We then used this labeling technique to characterize membrane synthesis, the rationale being that if new intracellular membrane is inserted, it will be seen as unlabeled membrane that either dilutes or displaces "old" labeled membrane. We first injected WGAAlexa488 during phase 1. Initially, WGA is detected over the entire surface of the somatic buds (Fig 2 A, top), reflecting the numerous villous projections observed in electron microscopy (1 µm within 2 min (+33' and +35'). The label is still detectable late in phase 4 and moves much faster (+56' and +58').
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We conclude from these observations that two adjacent membrane regions form in phase 1 that show distinct behaviors (Fig 2 D): the surface above the nuclei shows a rapid flux or turnover, whereas the FC defines a more stable membrane region that persists throughout the course of cellularization. Our data do not distinguish whether the persistent high levels of staining in the FC in phase 1 reflect only low turnover rates in that structure compared with the apical membrane or the active relocalization of the label from the apical membrane to the FCs. With the caveat that WGA may alter the normal behavior of the plasma membrane, these data suggest that the FC forms as a separate membrane region. Consistent with this view, note that Dlt becomes restricted to and remains in the FC throughout cellularization (Fig 1) and that a basal junction with high levels of Arm forms at the boundary between the FC and the adjacent membrane of the somatic bud (Fig 1). This junction may be required to isolate the FC from the rest of the plasma membrane.
Apical Insertion of New Membrane and Lateral Transfer of Old Membrane
The FC can actually only be labeled during phase 1. WGAAlexa488 injected during phase 2 appears to be excluded from the canal (Fig 3B and Fig B', arrowhead), although it binds uniformly to the remainder of the embryo's surface (Fig 3 A, purple). As membrane invaginates at a slow speed during phase 3, the label is gradually depleted from the apical membrane while the lateral label persists and the size of the labeled domain even expands (Fig 3 A, blue; compare double arrow at time 0 and +18'). The parallel nature of these processes suggests that membrane initially on the apical surface may be transferred laterally as cellularization proceeds, continuing with what happens during phase 1.
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The edge of the lateral labeled patches maintain their association with the invaginating FC, and the labeled regions behave as though they simply accompany the inward growth of the membrane until the end of cellularization. In fact, the original most intensely labeled membrane regions move basally at the same speed as the FCs through phases 3 and 4. This suggests that there is no obvious incorporation of unlabeled membrane at the invagination front or that if it occurred it would not contribute to the growth of the plasma membrane. Instead, new membrane incorporation appears to occur predominantly apically and accounts for the bulk of the membrane growth during phase 3.
Lateral Insertion of New Membrane Becomes Predominant in Phase 4
The rate of membrane invagination greatly increases in phase 4. We find that the pattern of membrane insertion is very different from that characterized for the preceding stages. When WGA label is injected at the end of phase 3, fluorescence is initially detected both apically and laterally (Fig 3 C, red and middle at time 0). In contrast to the earlier stages, the apical membrane maintains its label, though intensity slightly decreases, as cellularization proceeds. A gap of unlabeled membrane emerges in the apical portion of the lateral region (arrowhead in Fig 3 C, 12 min after time 0). This gap grows in size and the lateral membrane previously labeled at time 0 is displaced basally and maintains its association with the FC (not shown). We conclude that the point of new unlabeled membrane insertion changes during phase 4, shifting from predominantly apical to predominantly lateral. Given the mild reduction of the apical membrane label, we conclude that apical insertion still persists, but our data show that the major site of insertion is lateral.
The changes in distribution of the labeled lectin at the membrane surface do not reflect a change in distribution of the WGA receptors under either normal or experimental conditions. For instance, if lateral transfer of WGA during phase 3 stemmed from increasing levels of WGA receptors laterally as a result of local changes in membrane composition, injection of WGA at the end of phase 3 would predominantly label the lateral membrane. However, our data show that the ability of WGA to bind the apical membrane appears constant at any time during cellularization (compare Fig 3A and Fig C, at time 0). Clearance of fluorescent WGA from apical regions and lateral transfer could also be due to a redistribution of the WGA receptors induced by the binding of the WGA ligand. To control that the apical membrane still contains the WGA receptors, we sequentially labeled the membrane with WGA conjugated with different fluorochromes. First, we injected WGAAlexa488 during phase 3, leading to the lateral transfer of that label as described in Fig 3 A. At the onset of phase 4, WGArhodamine was injected in the same region of the embryo. As shown in Fig 3 D, this second label (red) is predominantly apical and tapers off along the lateral membrane, while the first label (green) is mostly in the basal lateral membrane. These experiments argue that the dynamic changes in WGA are unlikely due to the natural or induced redistribution of the WGA receptors.
In conclusion, throughout the course of cellularization, unlabeled membrane is constantly incorporated into labeled plasma membrane. This incorporation occurs in a defined sequence and a specific pattern, first apical, then apicallateral. Note that the membrane miscibility appears limited under conditions where WGA is bound to the plasma membrane.
Displacement of Microspheres on the Membrane Surface
To further test this idea, we probed the dynamic properties of the plasma membrane in a situation where it is unaffected. We followed the movement of 0.5-µm microspheres bound to an intact plasma membrane. These beads were coated with WGA so they bind the membrane very rapidly upon injection in the vitelline space. The maximum area of contact between the bead and the membrane (0.19 µm2) is much smaller than the total plasma membrane surface area at the onset of phase 3 (at least 100 µm2; see Materials and Methods). Therefore, the beads can be considered as small passive patches representing 1/550th of the total membrane when invagination begins and even less later on. Such beads are thus predicted to follow the global movement of membrane populations they are imbedded in. In particular, if membrane mixing is important and creates a constant homogeneous state in the membrane, the locally delivered new membrane will effectively distribute evenly around the bead and, over time, the movement of the bead will be random. However, if membrane mixing is limited, new inserted membrane should consistently displace both the recipient membrane and the beads away from the sites of insertion, thus creating anisotropic growth of the membrane.
We injected WGA-coated fluorescent 0.5-µm beads in the vitelline space of living embryos at different stages. The dilution was such that, on average, less than one bead bound the membrane of a forming cell, to avoid any possible interaction between beads. During phases 1 or 2, beads are seen in the FC a few minutes after injection and remain at the invaginating front through phases 3 and 4. Fig 4 A shows a bead at time 0 in red and at the end of phase 4 in green (the arrowhead points to the FC). This behavior is observed in 96% of the cases (n = 55) when beads are injected at early stages (Fig 4 D, top, blue). The remaining 4% represent beads located just above the FC which also accompany the movement of the FC. When injected at the beginning of phase 3, the beads rapidly bind the incipient lateral membrane. Fig 4B and Fig B', shows two representative examples where the left panel shows the DIC image at the initial time point: the arrowhead and the small arrows point to the initial positions of the FC and the beads, respectively. The right panel shows the positions of the beads at different time points: the blue bead indicates the initial location (arrows), and subsequent locations are in red and green. The beads are displaced away from the apical sites of membrane insertion and move in register with the advancing FC (Fig 4B and Fig B'). 89% of the beads (n = 122) have such a behavior and are eventually located along the basal portion of the lateral membrane (Fig 4 D, green shaded area). The remaining 11% are found in the medial portion of the lateral membrane (Fig 4 D, white area). During phase 4, when insertion becomes predominantly apicallateral, a different situation is observed. Beads injected at the onset of phase 4 remain in the apical region of the membrane, above the insertion site, in 88% of cases (n = 56) (Fig 4 D, red shaded area) and don't accompany the rapid movement of the invaginating membrane. Fig 4 C shows the position of three beads at the end of cellularization; the two arrowheads show the extent of membrane growth during that time window.
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Our data show that the microspheres have a reproducible nonrandom mobility and are displaced away from insertion sites with their final distribution depending on the time when they bind the membrane: the most basally located beads were injected earlier than more apically located beads. Given the relatively small size of the beads with respect to the surrounding membrane surface, the movement of the beads most likely reveals the behavior of the supporting membrane as a whole. We therefore infer that membrane growth appears anisotropic, mainly occurring at insertion sites. This situation might reveal that diffusion in the plane of the membrane, although effective, is not enough to cancel the effect of massive membrane insertion in a defined site.
Insertion of Neurotactin, a Newly Synthesized Transmembrane Protein
The labeling experiments suggest that membrane growth is mostly accounted for by a reservoir of intracellular membrane originating from the secretory pathway. To assess the contribution of ER and Golgi-based membranes to cellularization, we used antibodies to Neur, a transmembrane protein whose synthesis during cellularization requires de novo RNA synthesis in late syncytial embryos (Fig 5 A). We followed the localization of Neur along the biosynthetic pathway. Neur is first detected in early phase 2 during nuclear elongation after the basal junction and FCs have formed (Fig 5 B). At this point, the membrane has not started to grow and most of Neur is localized in punctate structures in the apical and basal cytoplasm (Fig 5 B, arrowheads). The number and intensity of punctate Neur structures increase in the basal cytoplasm during phase 3 (Fig 5 C, bracket), suggesting an increased level of synthesis. We compared the cytoplasmic localization of Neur to that of ER and Golgi markers. The ER resident chaperone BiP is concentrated in an apical perinuclear region (Fig 5 F, arrowhead), and extends at lower levels in an intricate web into the basal cytoplasm (Fig 5 F, bracket). In contrast to mammalian cells, the Golgi apparatus of insects is dispersed (
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During phase 3, Neur also becomes visible along the invaginating lateral membrane (Fig 5 C, arrowhead). Towards the end of phase 3, Neur levels increase in the lateral membrane, at the expense of a reduced cytoplasmic localization below the nuclei (compare Fig 5C and Fig D). It is also detected on the apical surface, but is absent from the FC (marked using an antibody to myosin in Fig 5 D, inset, red). This latter point is in agreement with our finding that the FC is a membrane region set aside very early before Neur can be detected in the embryo. At the end of cellularization, membrane staining of Neur is even stronger, especially in the apical regions of the lateral membrane (Fig 5 E, arrowhead). The distribution of Neur is eventually asymmetric, with higher levels coinciding with the zone of late apicallateral membrane insertion (region 3) observed in WGA labeling experiments. Fig 5 J shows an embryo labeled with WGA in late phase 3 and viewed at the end of cellularization to illustrate this parallel: the arrowhead points to the zone of unlabeled membrane insertion, which appears to coincide with the zone of increased Neur staining (compare Fig 5E and Fig J, arrowheads).
In conclusion, the transmembrane protein Neur is induced during cellularization and localized in the secretory pathway in proximity to cis-Golgi membranes. Neur is continuously inserted to the growing plasma membrane. These observations further support the view that new membrane originating from the secretory pathway is constantly used as raw material for membrane growth.
Microtubules Mediate the Mobilization of Secretory Membranes to the Growing Plasma Membrane
Neur must be transported over a large distance of 20 µm from the Golgi apparatus to the sites of insertion in the plasma membrane. Blocking this transport should immediately stall cellularization if the flux of membrane from the Golgi apparatus is the primary source of new membrane for cellularization. As shown in Fig 6, a good candidate for the mechanism underlying an apical movement of Golgi membranes is microtubular transport. Microtubule asters are located apical to the nuclei beneath the plasma membrane (Fig 6 A, arrowhead) and very long microtubules extend basally up to 30 µm deep inside the embryo (Fig 6A and Fig B) well within the area where the bulk of the cis-Golgi membranes are detected (Fig 6 A) and where Neur accumulates in punctate structures (Fig 5 C).
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Depolymerization of microtubules by injection of colchicine or colcemid at the onset of cellularization blocks membrane invagination (
Although we cannot exclude additional roles for microtubules, these data establish a link between plasma membrane growth and export of Neur from Golgi-associated membranes. This requirement further establishes that the membrane needed for cellularization originates from intracellular organelles.
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Discussion |
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Using novel techniques to follow the dynamics of the plasma membrane in living embryos, we explored the modalities of membrane growth during cellularization.
We find that cellularization involves the remobilization of an intracellular membrane reservoir and its regulated insertion at well-defined sites in a precise sequence, first apical, then apicallateral. This unexpected control of membrane delivery during cleavage of the embryo may contribute to the initiation and slow emergence of apicalbasal polarity as the epithelium forms.
Cellularization Involves the Remobilization of an Intracellular Membrane Reservoir
The origin of the membrane required for cytokinesis and for cellularization, in particular, has been a long-standing unresolved issue. We conclude that the major reservoir of membrane is intracellular, based on the following observations. First, at any time during cellularization, the previously labeled plasma membrane incorporates membrane that is unlabeled and therefore must originate from inside the cell. Second, Neur, a transmembrane protein that we showed is induced de novo during cellularization, accumulates in Golgi-associated membranes and is continuously transferred to the growing plasma membrane. When microtubules are depolymerized, cellularization immediately stalls and Neur accumulates in abnormally abundant punctate structures of the basal cytoplasm reminiscent of those seen under normal conditions. The exclusion of BiP staining from this region and the presence of ßCOP punctate structures in close proximity or colocalizing with Neur argue that Neur is properly exported from the ER to the Golgi apparatus and accumulates in Golgi-associated membranes when microtubules are depolymerized. This is in contrast to mammalian cells, where ER to Golgi export is a microtubule-dependent process and may identify a special feature of the ER and Golgi apparatus organization in early Drosophila embryos. Taken together, our data argue that ER and Golgi membranes constitute the major raw material for plasma membrane growth during cellularization. This is at odds with the idea that membrane grows through the simple redistribution of apical membrane and that the villous projections constitute the major reservoir for invagination (
Membrane is normally made in the ER and passes through the Golgi apparatus constitutively on its way to the plasma membrane. Given the relatively constant amount of membranes stained with either BiP or ßCOP antibodies from cycle 914 embryos (not shown), it is probable that the intracellular membrane reservoir is made long before cellularization begins and that cellularization involves the regulated mobilization of this membrane pool to the plasma membrane. It will be interesting to see which steps of the secretory pathway are regulated to accommodate the need for rapid membrane export. Moreover, the shift from a slow rate to a fast rate of invagination suggests that during cellularization regulation of membrane export changes. The existence of a mutant that results in a specific defect in membrane growth during phases 13, but where phase 4 (fast phase) proceeds normally (
Insertion of New Membrane as an Active Mechanism during Cytokinesis?
A surprising result is that the majority of the membrane accounting for growth and invagination is not targeted to the front of the invaginating membrane, which is in contradiction to a report from
A Link between Membrane Growth and Polarization during Cytokinesis?
Different lines of evidence also suggest that polarization of the plasma membrane might be inherently linked to the process of membrane growth during cleavage of the Drosophila blastoderm. Membrane insertion does not occur at random and uniformly over the entire surface of the plasma membrane. Rather, new membrane is delivered at well-defined sites that change from apical to apicallateral at different phases of cellularization. This change coincides with a sudden twofold increase in the rate of membrane invagination from phase 3 to 4. Therefore, during cleavage of the Drosophila embryo, membrane trafficking is regulated in a way that allows polarized membrane delivery. Genetic analysis of syntaxin1, a target membrane protein termed t-SNARE, in Drosophila cellularizing embryos (
Two independent sets of experiments argue that membrane miscibility might be insufficient to compete with the effect of massive local membrane delivery, thus lending support to our model. At any time during cellularization, WGA-labeled membrane regions located basal to the sites of membrane delivery move in register with the advancing front of invagination and segregate from the newly inserted membrane as if displaced away from the sites of insertion. We do not think this is a simple effect of WGA binding to the membrane, as a similar conclusion was drawn when we tracked WGA-coated microspheres bound to an otherwise intact plasma membrane. The microbeads serve as landmarks on the plasma membrane and reveal its dynamic properties. Isolated beads move away from the sites of membrane insertion and follow the inward movement of the FC. This situation would not occur if diffusion in the plane of the membrane was enough to cancel the effect of local membrane delivery. Together, these experiments argue that the history of membrane insertion is partly recapitulated in the spatial arrangements of distinct membrane regions, the most basal being also the oldest, generally speaking. This would provide a powerful mechanism to polarize the forming epithelium if the composition of the newly inserted membrane from the ER and Golgi apparatus changes as a function of time through the progressive induction of new transmembrane proteins. Each set of transmembrane proteins might in turn recruit specific groups of membrane-bound proteins and further define each region as a compositionally unique membrane domain. We studied the transmembrane protein Neur and found that its localization in phase 4 reflects the scheme of apicolateral membrane insertion in agreement with our model. Neur is absent from the FC, a membrane region that is formed before induction of Neur. Neur is also localized at higher levels in the apicallateral sites of membrane insertion.
Such a model could explain how a prepolarity initiated during cellularization would subsequently participate in recruitment of proteins required for the formation of a mature apical adherens junction. The apical adherens junction is only effective in tightly connecting adjacent cells and serving as a diffusion barrier after cellularization is completed and when gastrulation begins. At this stage, Arm/ß-catenin is concentrated in a tight apical belt (
To conclude, our data show that cellularization, a specialized form of cytokinesis, is characterized by a precise scheme of membrane growth and insertion that accompanies the emergence of cell polarity. Cellularization may use a novel strategy to polarize the membrane, which we suggest occurs through the sequential insertion of new membrane at different sites and is supported by the fact that membrane mixing between the recipient and newly inserted membrane populations may not be enough to compete with the overwhelming insertion of new membrane. It will be interesting to see whether this pertains to other situations of massive membrane growth such as axon formation. Our findings also open the interesting possibility that membrane insertion might actively participate in furrowing during cytokinesis.
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Footnotes |
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1 Abbreviations used in this paper: Arm, Armadillo; DIC, differential interference contrast; Dlg, Discs-large; Dlt, Discs-lost; FC, furrow canal; MDCK, Madin-Darby canine kidney; Neur, Neurotactin; WGA, wheat germ agglutinin.
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Acknowledgements |
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We are very grateful to G. Waters, T. Vogt, Y. Ahmed, and R. Hoang for insightful and useful comments on the manuscript; and to M. Bhat and H. Bellen for the antibody to Dlt, C. Fields for the antibody to myosin, and V. Ripoche and V. Malhotra for the antibody to ßCOP. We also thank J. Goodhouse for help with confocal microscopy.
E. Wieschaus is an investigator of the Howard Hughes Medical Institute. T. Lecuit was supported by the Howard Hughes Medical Institute, the Philippe Foundation, and a long-term fellowship from the Human Frontier Science Program Organization.
Submitted: 12 June 2000
Accepted: 26 June 2000
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References |
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