A cleavage clock regulates features of lineage-specific differentiation in the development of a basal branching metazoan, the ctenophore Mnemiopsis leidyi
© Fischer et al.; licensee BioMed Central Ltd. 2013
Received: 20 August 2013
Accepted: 20 November 2013
Published: 31 January 2014
An important question in experimental embryology is to understand how the developmental potential responsible for the generation of distinct cell types is spatially segregated over developmental time. Classical embryological work showed that ctenophores, a group of gelatinous marine invertebrates that arose early in animal evolution, display a highly stereotyped pattern of early development and a precocious specification of blastomere fates. Here we investigate the role of autonomous cell specification and the developmental timing of two distinct ctenophore cell types (motile compound comb-plate-like cilia and light-emitting photocytes) in embryos of the lobate ctenophore, Mnemiopsis leidyi.
In Mnemiopsis, 9 h after fertilization, comb plate cilia differentiate into derivatives of the E lineage, while the bioluminescent capability begins in derivatives of the M lineage. Arresting cleavage with cytochalasin B at the 1-, 2- or 4-cell stage does not result in blastomere death; however, no visible differentiation of the comb-plate-like cilia or bioluminescence was observed. Cleavage arrest at the 8- or 16-cell stage, in contrast, results in the expression of both differentiation products. Fate-mapping experiments indicate that only the lineages of cells that normally express these markers in an autonomous fashion during normal development express these traits in cleavage-arrested 8- and 16-cell stage embryos. Lineages that form comb plates in a non-autonomous fashion (derivatives of the M lineage) do not. Timed actinomycin D and puromycin treatments show that transcription and translation are required for comb formation and suggest that the segregated material might be necessary for activation of the appropriate genes. Interestingly, even in the absence of cytokinesis, differentiation markers appear to be activated at the correct times. Treatments with a DNA synthesis inhibitor, aphidicolin, show that the number of nuclear divisions, and perhaps the DNA to cytoplasmic ratio, are critical for the appearance of lineage-specific differentiation.
Our work corroborates previous studies demonstrating that the cleavage program is causally involved in the spatial segregation and/or activation of factors that give rise to distinct cell types in ctenophore development. These factors are segregated independently to the appropriate lineage at the 8- and the 16-cell stages and have features of a clock, such that comb-plate-like cilia and light-emitting photoproteins appear at roughly the same developmental time in cleavage-arrested embryos as they do in untreated embryos. Nuclear division, which possibly affects DNA-cytoplasmic ratios, appears to be important in the timing of differentiation markers. Evidence suggests that the 60-cell stage, just prior to gastrulation, is the time of zygotic gene activation. Such cleavage-clock-regulated phenomena appear to be widespread amongst the Metazoa and these cellular and molecular developmental mechanisms probably evolved early in metazoan evolution.
KeywordsCtenophore Comb jelly Photocyte Comb cell Cytochalasin B Puromycin Actinomycin Cell cycle arrest Cell lineage
Many ctenophores, including the lobate ctenophore Mnemiopsis leidyi, generate distinct light-producing photocytes during development, which migrate to locations subjacent to each of the eight comb rows. Genome sequencing of Mnemiopsis has revealed two genomic clusters containing ten distinct copies of photoproteins closely resembling the luciferase-type photoproteins found at the base of the Metazoa. In situ hybridization studies have shown that at least subsets of these mRNAs are expressed in photocytes prior to when these embryos are bioluminescent. Thus, the presence or absence of bioluminescence is a strong indicator of the developmental fate of a differentiated photocyte.
Additional file 1: Normal embryonic development of Mnemiopsis leidyi. DIC microscopy time-lapse movie of developing Mnemiopsis embryos beginning with a zygote until 12 hpf. The aboral side is up. The animal shown is the same as in Figure 2A-P. (MOV 6 MB)
The stereotyped cleavage program in ctenophores allows each blastomere to be identified and its fate followed by the injection of intracellular lineage tracers[1, 4, 7] (summarized in Figure1C). For example, the mesoderm, including muscle, mesenchymal cells and photocytes, is generated by the micromeres born from endodermal precursors at the future oral pole (summarized in Figure1C). Early labeling experiments identified that the e1 micromeres give rise to the comb plate cilia[18, 19]; however, later fate-mapping experiments were able to detect the m1 micromere’s contributions to the formation of comb plates (summarized in Figure1C). Interestingly, when e1 micromeres are deleted, no comb plate cilia form, indicating that e1 micromeres are autonomously specified to give rise to comb plates while m1 micromeres require inductive signals[18, 20].
Here, we investigate the role of the cleavage program in the segregation and expression of the developmental potential of two distinct cell types (motile comb-plate-like cilia and light-emitting photocytes) during the development of the ctenophore Mnemiopsis. Using an inhibitor of filamentous actin polymerization (cytochalasin B) to block cytokinesis (but not nuclear division), we show that the formation of differentiation markers associated with each of these cell types appears only after they have segregated into their own distinct lineages and they are never co-expressed in the same cells. Furthermore, using inhibitors of protein synthesis, transcription and DNA synthesis, we provide evidence for the existence of a cleavage clock that is based on the number of rounds of DNA synthesis (and possibly controlled by the nuclear-to-cytoplasmic ratio), which determines the temporal appearance of differentiation products in cleavage-arrested embryos.
Mnemiopsis collection and embryos
Mnemiopsis leidyi adults were collected from the National Oceanic and Atmospheric Administration jetty and Eel Pond in Woods Hole, MA, during June and July and from 13660 Deering Bay Dr, Coral Gables, FL 33158, during December. Adults were brought into the lab and induced to spawn as described previously. Eggs were washed multiple times with 0.2 μm filtered seawater to remove any jelly or debris. Only batches in which a high percentage of embryos developed normally were used in the study. Fertilization in these animals occurs at spawning, so for developmental timing purposes this was designated as 0 hpf.
After the embryos were collected, they were transferred by pipet to 24-well culture dishes for drug treatments. The following drugs were used: cytochalasin B (Sigma, St. Louis, MO, USA, C6762), actinomycin D (Sigma, St. Louis, MO, USA, A1410), puromycin (Sigma, St. Louis, MO, USA, P7255) and aphidicolin (Sigma, St. Louis, MO, USA, A0781). Stock solutions of cytochalasin (1 mg/ml), actinomycin (1 mg/ml) and aphidicolin (1 mg/ml) were made up in dimethyl sulfoxide, with aliquots stored at -20°C. A puromycin stock solution (12.5 mg/ml) was made up in distilled water and stored at -20°C. Freshly thawed aliquots were used in each experiment. Working solutions were made by diluting stock solutions with filtered seawater. Approximately 1 ml of solution was added to each well containing embryos.
Hoechst staining and immunohistochemistry
The embryos were fixed for antibody staining in 4% paraformaldehyde and 0.02% glutaraldehyde, as previously described by Pang and Martindale. Following fixation, the embryos were removed from their membranes by gentle pipetting, washed with PBS plus 0.2% Triton (PBT), blocked in 5% goat serum for 1 h and then incubated in anti-tyrosine tubulin (Sigma, T9028) overnight at 4°C. The embryos were washed six times for 30 min in PBT and incubated with the secondary antibody, goat anti-mouse conjugated to Alexa-594 (Invitrogen, Molecular Probes, Carlsbad, CA, USA), Alexa-488 phalloidin (Invitrogen, Molecular Probes, Carlsbad, CA, USA) and Hoechst 33342 (Invitrogen, Molecular Probes, Carlsbad, CA, USA) overnight at 4°C. Afterwards the embryos were washed twice for 5 min in PBS and mounted for imaging.
After the embryos were collected, the vitelline membranes were mechanically removed using sharpened forceps. The de-membranated embryos were allowed to develop to the correct cleavage stage prior to injection with DiI, (Catalog No. D-282; Molecular Probes, Inc., Eugene, OR) a lipophilic membrane stain that diffuses laterally to stain the entire cell, as described in. Following DiI injection, the embryos were carefully transferred into individual wells of a Terasaki plate (Nunc, Roskilde, Denmark) so that each embryo remained separate. Each well of the plate contained 10 μl of filtered seawater (controls) or cytochalasin B solution (1 μg/ml). The Terasaki plate was then stored in a humidified chamber to minimize evaporation.
The embryos were scored for the presence or absence of comb cells using a Stereo Discovery (Zeiss, Inc) or Axio Scope (Zeiss, Inc) under transmitted light. Live comb plate cells were imaged using time-lapse microscopy with time intervals of five to ten images per second. Normal Mnemiopsis development was recorded at three images per minute using DIC (Differential interference contrast). Visualization of bioluminescence was performed on an Axio Scope, using the GFP filter (38 HE Green Fluorescent Prot. filter, excitation BP 470/40 nm, emission BP 525/50 nm) to stimulate photoprotein emission and with 0.5 to 1 sec exposure times with an AxioCam Mr to capture faint signals. Prior to visualization, an embryo was kept undisturbed in darkness for 10 to 20 min to prevent activation and depletion of the photoprotein.
Confocal imaging after immunohistochemical staining was performed using a Zeiss 710 confocal microscope. Images were processed using Zen software (Zeiss, Inc) and Volocity (Improvision, Inc) to create 3D image reconstructions of confocal sections.
Additional file 2: Regression of the cleavage furrow in a Mnemiopsis embryo after cytochalasin B treatment. Cytochalasin B was added during the first cleavage of a Mnemiopsis zygote. DIC microscopy time-lapse showing the ingression of the cleavage furrow. Once cytochalasin B has been added, cleavage stops and the cleavage furrow regresses. The animal shown is the same as in Figure 3A-D. (MOV 73 MB)
Additional file 3: Cleavage-arrested embryo with beating cilia at 2 comb cells. DIC microscopy movie showing the beating cilia on comb-cells in a cleavage-arrested embryo. The embryo shown is the same one as in figure 5A and 5A'. (MOV 6 MB)
Additional file 4: Close-up on one a comb cell with beating cilia in a cleavage-arrested embryo. DIC microscopy movie showing the beating cilia on comb-cells in a cleavage-arrested embryo. The embryo shown is the same one as in figure 5B and 5B'. (MOV 3 MB)
Additional file 5: Cleavage-arrested embryo with beating cilia at a comb cells. DIC microscopy movie showing the beating cilia on comb-cells in a cleavage-arrested embryo. The embryo shown is the same one as in figure 5C and 5C'. (MOV 2 MB)
Ctene cells in cleavage-arrested embryos are derived from e1 micromeres
Overview of the cytochalasin treatment experiments on Mnemiopsis leidyi embryos
Cytokinesis arrested with cytochalasin B
50% have combs
All combs that were formed were labeled
75% have combs
83% have combs
Cytokinesis arrested with cytochalasin B, labeling of all four E blastomeres
62% (8/13) have combs that are labeled
38% (5/13) did not form combs
Cytokinesis arrested with cytochalasin B at 16-cell stage, labeling of all four E blastomeres at the 8-cell stage
95/128 (74%) formed combs that are labeled
99% of embryos that formed combs were labeled, E cell lineage forms combs
32/128 (25%) did not form combs
1/128 (0.8%) formed combs that are not labeled
Cytokinesis arrested with cytochalasin B at the 16-cell stage, labeling of all e1 micromeres at the 16-cell stage
27/36 (75%) formed combs that are labeled
100% of embryos that formed combs were labeled, e1 cells form combs
9/36 (25%) did not form combs
Cytokinesis arrested with cytochalasin B at the 16-cell stage, labeling of all 1E macromeres at the 16-cell stage
No labeled combs
Only e1 cells at 16-cell stage form combs and E1 cells do not form combs
Cytokinesis arrested with cytochalasin B at the 8-cell stage, labeling of all M blastomeres
19/27 (70%) formed combs that are not labeled
95% of the embryos formed combs that were not labeled and combs do not arise from M lineage cells
7/27 (26%) did not form combs
1/27 (4%) formed combs that are labeled
8-cell stage and 16-cell stage
Cytokinesis arrested with cytochalasin B at the 8-cell stage or 16-cell stage, all E blastomeres were labeled
Unlabeled cells have photoprotein
M cells can make photoprotein even when they are cell-cycle arrested
To rule out the possibility that DiI labeling could induce the cell fate of the comb-plate-like cilia, we labeled all four M blastomeres at the 8-cell stage or all four 1M macromeres and m1 micromeres at the 16-cell stage and arrested the embryos with cytochalasin B. In only one case (which was likely to have been generated from a misinjected cell), were the cells producing the comb-plate-like cilia labeled (Figure 6D, G).
To determine whether e1 cells specifically gave rise to the comb plates, we labeled all four e1 micromeres at the 16-cell stage and immediately afterwards treated the embryos with cytochalasin B. All of the resulting embryos formed cells producing comb-plate-like cilia that were labeled with DiI (Figure 6E, Table 1, Additional file7). We also labeled all four 1E macromeres after the division of the E blastomeres into e1 and 1E at the 16-cell stage and cleavage arrested the embryos with cytochalasin B. None of the cells producing comb-plate-like cilia that formed were labeled (Table 1). These data demonstrate that only E lineage blastomeres at the 8-cell stage and e1 micromeres at the 16-cell stage give rise to comb-bearing cells. 1E macromeres, but not M lineage descendants, appear to be able to contribute to the formation of comb-plate-like cilia.
Photocytes are specified by the M lineage in cleavage-arrested embryos
Additional file 8: Bioluminescence in photocytes of a control embryos. Light microscopy movie showing light emission from the photocytes that was stimulated by a brief illumination with fluorescent light. The photocytes are well visible on either side of the embryo. The animal shown is the same one as in Figure 7B. (MOV 1 MB)
Additional file 9: Bioluminescence in photocytes of a cleavage arrested embryos. Light microscopy movie showing light emission from the photocytes that was stimulated by a brief illumination with fluorescent light. Those cells that contain the bioluminescent protein are photocytes. The embryo shown is the same one as in Figure 7C. (MOV 4 MB)
In embryos that were treated with cytochalasin B at the 8- or 16-cell stage, we observed green bioluminescence in the cleavage-arrested blastomeres. This began at the same time as in control embryos at around 9 hpf (Additional file9). Because DiI labeling of M blastomeres masked the weak florescence of the photoproteins in injected embryos, we labeled the E blastomeres at the 8-cell stage with DiI. Observation of DiI-injected E lineage cleavage-arrested embryos at the 8- or 16-cell stage revealed that the photoprotein was located in the unlabeled M (8-cell) and 1M macromeres (precursors to the 2M lineage) at the 16-cell stage (Figure 7C, D, E, F).
Transcription and translation are required for the formation of comb-plate-like cilia in cleavage-arrested embryos
The differentiation of comb-plate-like cilia in E lineage cells and photoproteins in M lineage descendants in cleavage-arrested embryos suggests that developmental potential is faithfully segregated to the correct spatial lineages by the early cleavage program. However, it is unclear what key factors may be segregated into the lineages. To gain insight into the molecular nature of the determinants for the formation of comb-plate-like cilia, we used inhibitors of mRNA and protein synthesis to see if these classes of molecules could be responsible for the expression of these structures.
Overview of the aphidicolin, actinomycin D and puromycin treatment experiments on Mnemiopsis leidyi embryos
Arrest of DNA synthesis for 2 h with aphidicolin, 2 μg/ml
No combs formed (0/30)
DNA synthesis is essential for comb formation.
27/30 (90%) formed combs, comb formation was delayed by 2 h
29/30 (97%) formed combs, comb formation was delayed by 2 h
29/30 (97%) formed combs, comb formation was delayed by 2 h
60-cell stage and 4 hpf
Arrest of DNA synthesis and cytokinesis with aphidicolin, 2 µg/ml, plus cytochalasin B, 1 to 2 μg/ml
Immediately arrested, single nuclei observed, neither combs nor photoprotein were formed in any embryo (0/30)
DNA synthesis is essential for comb formation even in cleavage-arrested embryos.
1- to 2-cell stage
Blocking of transcription with actinomycin D, 10 μg/ml
0/30 gastrulated, development arrests at 32- to 60-cell stage
Zygotic transcription begins around the 32- to 60-cell stage and the specification of the combs requires more than only maternal RNAs.
4- to 8-cell stage
0/30 gastrulated, development arrests at 32- to 60-cell stage
16- to 32-cell stage
10/30 (33%) gastrulate, 20/30 (67%) arrest at 32- to 60-cell stage
28/30 (93%) gastrulate but no pharynx is formed, 2/30 (7%) do not gastrulate
5 hpf (post-gastrulation)
No pharynx is formed
Arrest of translation with puromycin, 125 μg/ml, 25 μg/ml
0/30 formed combs
Comb protein is translated prior to 6 hpf, comb formation requires protein that is translated in the embryo and not maternally loaded.
0/30 formed combs
0/30 formed combs
6 hpf (post-gastrulation)
32/30 (97%) formed combs
It is interesting to note that embryos that were treated at the 1-, 2-, 4-, 8- and 16-cell stages continued to divide normally and arrested only prior to gastrulation at about the 60-cell stage. Embryos that were treated with actinomycin D at the 32- or 60-cell stage divided only a few more times and then became arrested (Table 2). This suggests that if actinomycin D is inhibiting the bulk of mRNA synthesis, then zygotic transcription appears to begin at about the 32- or 60-cell stage and there are sufficient stores of maternal mRNAs to allow the embryo to reach these stages of development.
To block protein synthesis, we treated embryos at various stages with puromycin (25 to 125 μg/ml). Comb formation was only observed in embryos that were treated after 6 hpf (at 25 μg/ml puromycin), suggesting that the key determinant is not solely that a protein is required for comb formation, but that proteins must be translated between 5 and 6 hpf (Table 2).
The differentiation of combs is regulated by the number of nuclear divisions
Cells producing ctene-like cilia and photocytes are formed at the same time in control embryos and in embryos treated with cytochalasin B at the 8-cell stage or later, suggesting that the embryo has a molecular timing mechanism that operates in the absence of cytokinesis. Because nuclear divisions appear to proceed normally in embryos treated with cytochalasin B (Figure 4), the number of nuclear divisions may be an important component of the timing mechanism. To test whether the timing of the formation of the comb-plate-like cilia is dependent on the number of nuclear divisions, we treated embryos with the DNA synthesis inhibitor aphidicolin (2 μg/ml) at various cleavage stages up through 5 hpf (Table 2). When treated during early cleavage, up to the 32-cell stage, we did not observe the formation of any comb-plate-like cilia. When treated at the 60-cell stage, 4 hpf, and 5 hpf, we did observe comb formation, although it was delayed by approximately 2 to 3 h (12 hpf as opposed to 9 hpf). Embryos that were treated with aphidicolin at 5 hpf for 2 h and then removed from the drug, formed comb-plate-like cilia with a delay corresponding approximately to the length of the drug treatment. These results are consistent with the fact that the aphidicolin treatment is reversible, that DNA synthesis is able to resume following the rescue (for example, the drug is not lethal) and that the correct total lineage-specific DNA content is important for the timing of the formation of comb-plate-like cilia. We did not see any comb formation in embryos that were treated with both cytochalasin B and aphidicolin, even after 6 hpf. The double treatment with cytochalasin B and aphidicolin is not reversible and embryos that were treated for 2, 3 or 4 hpf did not recover and did not form comb cells. These results showed that that the combined effects of blocking DNA synthesis and cytokinesis are more detrimental than either drug alone and indicate that a critical number of nuclear divisions is essential.
We used cytochalasin B to inhibit cytokinesis in early ctenophore embryos to examine how developmental potential is partitioned into two distinct cell lineages: those giving rise to comb plates and those forming the photocytes. When cytokinesis is arrested in early Mnemiopsis leidyi embryos, treated embryos do not undergo apoptosis, but rather nuclear divisions continue and give rise to multinucleated cells. Cytochalasin B was used to study the segregation of developmental potential in Mnemiopsis by Freeman. Freeman inhibited certain divisions including the first, second or third cleavage or the second and third cleavage by short treatments with cytochalasin B. When only the first cleavage was inhibited by a brief exposure, most embryos developed like the control embryos but were delayed by one cleavage cycle. However, when the second, third or second and third cleavages were reversibly blocked, the inhibited cleavages were skipped and the embryos continued the cleavage program with a reduced number of cells.
Surprisingly, as we demonstrate here, if cytokinesis is arrested at the 8-cell stage or later and is permitted to remain arrested for the following hours of development, comb-plate-like cilia and the photoprotein-mediated bioluminescence characteristic of photocytes form at the correct time in development compared with control embryos. Lineage-tracing experiments reveal that comb-plate-like cells are derived from E cells at the 8-cell stage and from the e1 micromeres at the 16-cell state. These are the same cell lineages that autonomously generate comb-plate-like cilia when isolated from the rest of the embryo. Previous intracellular fate-mapping experiments showed that m1 micromere descendants also have the capacity to make comb-plate-like cilia but M lineage descendants were not found to give rise to comb-plate-like cilia in cleavage-arrested embryos. Likewise, the bioluminescence characteristic of M-lineage-derived photocytes was only seen in M-lineage-derived cells in cleavage-arrested embryos. Earlier experiments showed that the m1 micromeres require inductive signals first from the e11 or e12 micromeres and later from endomesodermal cells derived from E and M macromeres before they form combs[14, 20]. Our results support the finding that m1 micromeres require additional inductive signals from endomesodermal cells for comb formation, since no combs were formed by m1 micromeres in cleavage-arrested embryos. This supports the finding that the presence of e1 micromeres is not sufficient to induce comb formation in m1 micromeres[14, 20].
Interestingly, neither comb-plate-like cilia nor bioluminescence was detected in cleavage-arrested zygotes or 2- or 4-cell stage embryos, suggesting that the segregation of distinct E and M lineages is crucial for the expression of the lineage-specific differentiation products. These results suggest that factors that are required for comb-plate and photocyte determination are already present and localized at the 8- and 16-cell stages and may have mutually exclusive activity.
The segregation of developmental determinants
One of the fundamental processes of development is the localization of factors that are required to establish cell polarity, break symmetry and drive the asymmetric specification of cell fates. Experimental embryologists have provided evidence for the segregation of factors that determine specific cell fates in a diverse variety of different organisms (examples are summarized in[32–34]). In some embryos, segregated factors are asymmetrically localized maternally to the cell cortex such as the developmental potential required for gastrulation in echinoderm embryos[35–39] and for the establishment of dorso-ventral polarity in many embryos that undergo spiral cleavage[40–43], or the maternal gradients in insect eggs[44–46]. Other cytoplasmic factors are segregated actively at the time of fertilization as in ascidians[47–49], amphibians[50, 51] and soil nematodes[52, 53].
In ctenophore embryos, all evidence suggests that the asymmetrical localization of developmental potential is actively segregated by the cleavage process itself. Each of the early cell divisions leads to a definite asymmetric cell fate. The site of first cleavage gives rise to the oral-aboral axis. Interestingly, a similar correlation between the site of the first cleavage and the site of gastrulation, and thus the formation of the oral-aboral axis, has been observed in the hydrozoan Clytia hemisphaerica (previously called Phialidium gregarium). Freeman showed that the site of the first cleavage can be altered from the side of the polar body formation experimentally in ctenophores as well as in hydrozoans[10, 54]. In contrast, the eggs of several sea urchin species are already polarized along the animal-vegetal axis at the time of fertilization (summarized in). Starfish eggs and even ascidian eggs show a similar polarization[56, 57]. Based on currently available data, it could be assumed that an irrevocable polarization of the unfertilized egg was an evolutionary novelty of bilaterians whereas in basally branching taxa, such as ctenophores and cnidarians, the egg is polarized by the position of the female pronucleus, which determines the site of the first cleavage, after fertilization. However, a broader taxon sampling is desired to further support this hypothesis.
In ctenophores, the second division gives rise to the anal axis[4, 15, 19]. The third division separates the E and M lineages[4, 18], and the fourth division segregates fates between micromere lineages (for example, comb rows) and macromere lineages (for example, photocyte cells)[4, 18]. Consequently, when cytokinesis is inhibited, the factors remain associated with the proper lineages that would have generated the descendants during normal development. Interestingly, photoprotein formation and comb-plate formation never appeared to occur in the same cell. It is possible that the developmental determinants, which are required for comb-plate and photoprotein differentiation, inhibit each other and only after these factors have been spatially segregated can they be activated.
Although we have little information on the cell biological nature of the segregation process in ctenophores, centrosomes have been shown to be a causal factor in the segregation of factors that specify cell fates in other systems. In Caenorhabditis elegans, the sperm-derived centrosome, a complex of several proteins that acts as the microtubule-organizing center, breaks the symmetry of the oocyte and sets off a series of events that relies on maternally deposited proteins and eventually leads to the establishment of the anterior-posterior, dorsal-ventral and left-right body axes[58, 59]. A surprisingly large number of mRNAs are localized to specific cell lineages in the snail Ilyanassa. These mRNAs are associated with the centrosomes, and as the cells divide the mRNAs are subsequently distributed in an asymmetric manner to specific daughter cells. Experimental evidence shows that these factors play an active role in the unique development of these cells[60, 61]. It would be interesting to investigate whether a centrosome-dependent mechanism is also involved in the segregation of developmental factors in Mnemiopsis, where distinct developmental fates are decided at each of the early divisions.
What is the nature of segregated developmental determinants?
Ctenophore development is rapid, with cleavages occurring every 15 to 20 min. The asymmetrical localization of proteins or mRNAs into distinct lineages during each division could be a simple way to distribute components quickly so as to distinguish different cell fates. Freeman showed that the developmental determinants that are required for comb formation begin to be localized to the aboral region already during the 2-cell stage. Although crude and not gene specific, our experiments that inhibit transcription and translation suggest that the distributed determinants are not simply represented by the full set of comb-plate protein or comb-plate mRNA. Instead subsequent transcription and translation are required for comb formation. The nature of the components that are asymmetrically localized during the early cleavage stages in Mnemiopsis remains unknown.
In other systems, RNAs, proteins, protein complexes or a mix of these are the developmental determinants that are distributed unevenly during development and which break the symmetries. A well-known example is the localization of the protein Dishevelled – a key player of the Wnt/ β-catenin pathway – to one side of an unfertilized egg. This is one of the first symmetry-breaking events in many bilaterians[17, 62, 63]. Other well-known examples of proteins with asymmetric localizations are Miranda, Prospero and Staufen during Drosophila neuroblast division and the Par proteins during C. elegans development.
Besides the localization of proteins or protein complexes, RNAs are often asymmetrically stored and thus establish cell polarity and determine the different daughter cell fates. One of the most famous examples of RNA localization is the localization of the Oskar mRNA, which is one of the key components used to establish polarity in the Drosophila oocyte during Drosophila oogenesis[66–69]. Other examples of RNA localization are ASH1 in budding yeast, bicoid in Drosophila embryos, Vg1 in Xenopus oocytes and CamKIIa in distal dendrites in mammalian neurons (summarized in).
Future experiments will be needed to reveal the nature of the developmental determinants that are segregated during early ctenophore development.
Temporal regulation of comb-plate formation: counting cell divisions
The regulation of growth and differentiation are key to developmental processes. Moreover, the timing of developmental events is a particularly important aspect of development. We showed that the differentiation of comb cells in Mnemiopsis embryos, which are cleavage arrested at the 8-cell stage or later, subsequently occurs at the same time as in untreated control embryos. Furthermore, our results show that comb-plate formation is blocked if DNA replication is inhibited by aphidicolin in addition to the inhibition of cytokinesis by cytochalasin B. This indicates that the number of nuclear division cycles or the amount of DNA (the nuclear-to-cytoplasmic ratio) appear to be important factors for comb-cell differentiation and not simply the amount of time that has elapsed since the third cleavage. As previously shown, after the second and/or third cleavage is blocked, Mnemiopsis embryos skip the respective cleavages and continue in a timely manner with their subsequent cleavage program, suggesting that a timing system determines the orientation of each division. Once this process is activated, the cleavage plane is determined by this timing mechanism and cleavages do not follow a set order with respect to each prior cleavage division. Since the inhibition of the first cleavage only causes a delay of the program, which otherwise occurs normally, Freeman concluded that the timing mechanism is initiated with the completion of the first cleavage.
Embryos that undergo exact numbers of cell divisions are known from a broad range of developmental systems; however, little is understood about the underlying mechanisms[71–75]. A stunning example is the highly stereotypic cell division pattern of the ventral germ band in malacostracan crustaceans. In Cherax destructor, for example, all ectoteloblasts undergo exactly 15 rounds of asymmetric cell division and give birth to ectodermal blast cells, which undergo two distinct divisions each.
Fundamental experiments by Whittaker showed that ascidian embryos developed muscle-specific acetylcholinesterase and brain pigment cell tyrosinase in specific blastomeres even in cleavage-arrested embryos and concluded that specific positional information is differentially segregated during early development. Since then several studies have employed cytochalasin B to investigate the cell fate specification and cell lineage of ascidian embryos (for example,[77–80]). Satoh and Ikegami performed a series of experiments combining cytochalasin B and aphidicolin in ascidian embryos, showing that future muscle cells must undergo eight cell-division cycles before they start expressing the muscle cell lineage marker acetylcholinesterase. Nevertheless, the question of how cells ‘count’ the number of cell cycles remains unanswered.
A number of counting mechanisms have been suggested for this phenomenon. Based on the observation that genomic DNA is highly methylated in zygotes and gradually demethylated during development, which tends to de-repress transcriptional activity in mammals, Kataoka et al. suggested that changes in DNA methylation could be involved in keeping count of the number cell cycles in ascidians. However, their own results do not support this hypothesis as they were unable to detect any changes in DNA methylation. Another possible explanation is that a specific ratio of DNA to cytoplasm is required. This ratio is crucial during early Xenopus development, where zygotic transcription is initiated after 11 to 12 rounds of cell division. The required ratio of DNA-to-cytoplasmic volume can also be acquired when cytokinesis is blocked with cytochalasin B but DNA replication is allowed to continue further. In contrast, in C. elegans, the timing of gut marker gene expression only depends on an early period of DNA synthesis until the 8-cell stage when the gut is clonally established. Subsequent rounds of cell divisions, which usually occur after the 8-cell stage, can be inhibited without preventing the expression of the marker gene at a later stage, thus this is independent of the DNA-to-cytoplasmic ratio or the number of DNA synthesis rounds.
Future studies are required to unravel the mechanism that regulates the timing of cell differentiation in ctenophores, whether this is the ratio of DNA to cytoplasm, DNA methylation, protein degradation, telomeric alterations, a combination of different factors or an entirely different mechanism that is currently unknown.
The onset of zygotic transcription
Early developmental events are regulated by gene products, which may be provided maternally. The activation of the zygotic genome marks the maternal-to-zygotic transition (MZT), which was first described for Xenopus and has since been studied in many model organisms. It is estimated that around 40% (in mice) to 75% (in sea urchins) of all protein coding genes are represented as maternal mRNAs and are degraded throughout early development[85, 86].
Other than bilaterians, the onset of zygotic transcription has only been described in cnidarians. In Clytia hemisphaerica, zygotic transcription starts at the late blastula to early gastrula stage[63, 87]. The sea anemone Nematostella vectensis exhibits an upregulation of gene expression around 10 hpf, at a time in development when cell division is asynchronous.
We used actinomycin D to inhibit transcription and we showed that treatment prior to the 60-cell stage did not visibly affect development until the animal reached the 60-cell stage, suggesting that maternally loaded transcripts are sufficient to support the early cleavage program up to this stage but not subsequent development or differentiation. These results suggest that the onset of zygotic transcription starts at about the 60-cell stage, just prior to gastrulation. The rapid early development of ctenophores also suggests that there may be little time for the transcription of new messages until the cell cycle began to slow down. Future experiments using labeled nucleotides will help to determine the precise onset of zygotic transcription in Mnemiopsis.
The transition from maternal to zygotic transcription is most likely an ancient feature in animal evolution and might be tightly linked with the evolution of multicellularity and sexual reproduction. If ctenophores are the sister group of all metazoans, as suggested by recent studies[90, 91], one could conclude that the inclusion of the MZT is part of the metazoan ground pattern. However, so far data about the MZT in sponges are not available and the phylogenetic position of ctenophores is still very much under debate. There are currently several contradictory hypotheses, including: (a) ctenophores and cnidarians are sister groups[93–95] and (b) ctenophores are the sister group to all bilaterians. Recent developmental studies support the idea that ctenophores lack several characters that are shared between cnidarians and bilaterians[13, 97–101] and thus they support the idea that (c) ctenophores are the sister group to all remaining eumetazoans or even, as mentioned above, that (d) ctenophores are the sister group to all metazoans[90, 91].
A phylogenetic comparison of maternally loaded proteins and the molecular regulation of the maternal-to-zygotic transition in bilaterian and non-bilaterian taxa might provide further insight into the evolution of sexual reproduction and life history.
hours post fertilization
PBS plus 0.2% Triton
Differential interference contrast
We are exceptionally thankful to Prof William Browne (University of Miami, FA) for allowing us to perform some of the experiments in his laboratory and providing animals. We also want to thank the community of the Marine Biological Laboratory for facilitating these studies, in particular the 2009 Embryology class, the faculty and the course directors, Lee Niswander and Nipam Patel, where this project was initiated.
This work is supported by a Boehringer Ingelheim Travel grant and an MBL Financial Aid Award to AHLF to attend the Embryology course 2009. It is further supported by National Science Foundation grants awarded to JQH (No. 1121268) and KP (No. 1158629). MQM was supported by the National Science Foundation and NASA.
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