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The translational regulator nanos is required for the survival and maintenance of primordial germ cells during embryogenesis. Three nanos homologs are present in the genome of the sea urchin Strongylocentrotus purpuratus, all of which are expressed with differential timing in the small micromere lineage. This lineage is set-aside during embryogenesis and contributes to constructing the adult rudiment. Small micromeres lacking Sp-nanos1 and Sp-nanos2 undergo an extra division and are not incorporated into the coelomic pouches. Further, these cells do not accumulate Vasa protein even though they retain vasa mRNA. Larvae that develop from Sp-nanos1 and 2 knockdown embryos initially appear normal, but do not develop adult rudiments; although they are capable of eating, over time they fail to grow and eventually die. We conclude that the acquisition and maintenance of multipotency in the small micromere lineage requires nanos, which may function in part by repressing the cell cycle and regulating other multipotency factors such as vasa. This work, in combination with other recent results in Ilyanassa and P. dumerilii, suggests the presence of a conserved molecular program underlying both primordial germ cell and multipotent cell specification and maintenance.
Multicellular animals are composed of many specialized cell types, but only the germ cells are capable of passing genetic information on to the next generation. To propagate the species, select cells are set aside during embryogenesis to form the future germline. A conserved set of genes is required to specify and/or maintain the germline during embryogenesis. Nanos, one such gene, is required for fertility in Drosophila, C. elegans, zebrafish, and mice (Kobayashi et al. 1996; Subramaniam and Seydoux 1999; Koprunner et al. 2001; Tsuda et al. 2003). Germline progenitor cells of the Drosophila embryo (pole cells) that lack nanos exhibit various defects, including an inability to migrate to the gonad, loss of transcriptional and mitotic quiescence, expression of somatic cell markers, and apoptosis; consequently nanos-null pole cells fail to develop into functional germ cells (Kobayashi et al. 1996; Asaoka-Taguchi et al. 1999; Deshpande et al. 1999; Hayashi et al. 2004). Nanos functions together with pumilio to repress the translation of the cell cycle regulator cyclin B and the pro-apoptotic gene head involution defect (hid) in pole cells, thus directly controlling the cell cycle and survival of these cells (Asaoka-Taguchi et al. 1999; Kadyrova et al. 2007). Similarly, loss of nanos function in C. elegans leads to premature primordial germ cell (PGC) proliferation and in mice its loss results in PGC apoptosis. Thus, nanos appears to have conserved functions in animal germlines (Subramaniam and Seydoux 1999; Suzuki et al. 2008).
In addition to its germline functions, nanos is expressed more broadly in multipotent cells. In Hydra polyps nanos is expressed in multipotent interstitial cells (I-cells), which give rise both to several somatic cell types and to germ cells (Mochizuki et al. 2000). In the polychaete annelid, Platynereis dumerilii, and the snail, Ilyanassa, both of which develop through a larval stage, multipotent cells of the embryo specifically express nanos (Rebscher et al. 2007; Rabinowitz et al. 2008). The nanos-positive multipotent cells of P. dumerilii give rise to both all of the trunk mesodermal cell types of the adult segments, and to germ cells (Rebscher et al. 2007). Nanos is also expressed in the 4d lineage of Ilyanassa; this lineage gives rise to adult mesodermal and endodermal tissues (Render 1997; Rabinowitz et al. 2008). Functional studies in Ilyanassa suggest that nanos is critical to maintain the fate of the 4d lineage, as the loss of nanos function in embryos results in the loss of all 4d-derived adult structures (Rabinowitz et al. 2008).
The majority of the species in the phylum Echinodermata are maximal indirect developers in which embryogenesis culminates with the formation of a free-swimming, feeding larva that supports its developing adult rudiment. At metamorphosis the rudiment will give rise to the juvenile. During embryogenesis in these maximally indirect developing organisms, groups of cells are set aside for use in adult rudiment construction. Unlike the cells that will give rise to the larval structures per se, these cells retain proliferative and developmental potential (Peterson et al. 1997). The small micromere lineage of the sea urchin embryo, which specifically accumulates nanos mRNA, is one such group of set-aside cells (Juliano et al. 2006). During embryogenesis, four small micromeres are formed at the 32-cell stage as a result of two unequal cleavage divisions: a vegetal unequal fourth cleavage division gives rise to a 16-cell embryo with 4 micromeres and a subsequent unequal division of the micromeres results in the formation of 4 small micromeres. In the blastula the small micromeres reside at the vegetal plate where they divide once before being transported through the blastocoel at the tip of the archenteron during gastrulation. The eight small micromere descendants are then partitioned into the left and right coelomic pouches, the site of adult rudiment formation in the pluteus larva. The coelomic pouches are derived from two sources: 60% from a subset of the macromere descendants and 40% from the small micromere descendants (Cameron et al. 1991). During larval formation, the small micromere descendants move into the coelomic pouches from their position at the tip of the archenteron in the gastrula (Pehrson and Cohen 1986; Tanaka and Dan 1990). It is clear that the small micromere lineage gives rise solely to adult tissues, but the identity of these tissues has not been experimentally determined (Pehrson and Cohen 1986; Tanaka and Dan 1990).
Due to both their slow cell cycle and contribution to adult tissues, it has been suggested that the small micromeres are germline precursors (Tanaka and Dan 1990) . In support of this hypothesis, the conserved germline genes nanos, vasa, and piwi are specifically expressed in the small micromeres (Juliano et al. 2006; Voronina et al. 2008). However, the small micromeres proliferate shortly after they reach the coelomic pouches (Tanaka and Dan 1990). This would not be expected if the small micromeres were indeed equivalent to PGCs, which typically stay mitotically and/or transcriptionally quiescent until they reach the somatic gonad (Su et al. 1998; Subramaniam and Seydoux 1999; Seydoux and Braun 2006; Seki et al. 2007). An alternative hypothesis suggests that the small micromeres are instead multipotent and will give rise to various adult tissues (Ransick et al. 1996). These seemingly contradictory hypotheses can be reconciled when considering recent results pointing to a broader role for conserved germline genes in multipotent cells that give rise to both germ cells and to somatic cells (Mochizuki et al. 2000; Mochizuki et al. 2001; Reddien et al. 2005; Rebscher et al. 2007; Palakodeti et al. 2008; Pfister et al. 2008; Rabinowitz et al. 2008; Swartz et al. 2008). Thus, the small micromere lineage may give rise to both germ and somatic cell types of the sea urchin adult. Here we test the function of nanos in the small micromere lineage, a likely multipotent cell population, in order to further understand the potentially ancestral role of this gene in establishing and maintaining multipotential cell populations during embryogenesis.
Strongylocentrotus purpuratus were housed in aquaria with artificial seawater (ASW) at 16°C (Coral Life Scientific Grade Marine Salt; Carson, CA). Gametes were acquired by either 0.5M KCl injection or by shaking. Eggs were collected in ASW or filtered seawater and sperm was collected dry. To obtain embryos, fertilized eggs were cultured in ASW or filtered seawater at 16°C. When early stage embryos were required for labeling, fertilization was performed in the presence of 1mM 3-amino-triazol (3-AT) (Sigma; St. Louis, MO) to inhibit cross-linking of the fertilization envelopes. Before fixing, envelopes were removed by passing the embryos through 80μM and 64μM Nitex® mesh. Careful monitoring was required to ensure the integrity of the embryos.
Three Sp-nanos homologs were identified in the S. purpuratus genome (spbase.org) by BLAST analysis (Altschul et al. 1990). Full-length genes (complete ORF plus some UTR sequences) were amplified from 24 or 48-hour embryonic cDNA by PCR and cloned into pGEMT-EZ (Promega; Madison, WI) for sequencing. Amplification of Sp-nanos2 required only one round of PCR (Juliano et al., 2006), whereas Sp-nanos1 and Sp-nanos3 required 2 rounds of PCR with nested primer sets to amplify full-length sequences. Primers were as follows (listed 5′ to 3′): Sp-Nanos2-F1 -TTCTTGACTAGCTCTACGACGTACT; Sp-Nanos2-R1 - TCGAGACGAGTAGACCCTACA; Sp-Nanos1-F1 - TAGATCATTCAAGACAAGCTCT; Sp-Nanos1-F2 - GGAAGTACATCGCATTTTACAA; Sp-Nanos1-R1 - CTAGAAGATCTTAACGGTCG; Sp-Nanos1-R2 - TGGGGTTCGATACTGGGATC; Sp-nanos3-F1 - GTACACCCGTGTCCGTGAG; Sp-nanos3-F2 - CCAATACAACATTAATCTTCAAG; Sp-nanos3-R1 - TGTCAAAAACTTTGTGCCAGAA; Sp-nanos3-R2 - TACTTCCTACATAGGACGAC. The 3′ and 5′ UTR sequences of the Sp-nanos homologs were extended with RACE (Ambion; Austin, TX). Using PAUP, an unrooted neighbor-joining phylogram was made from full-length nanos coding sequences collected from NCBI (for accession numbers see Figure 1); bootstrap replicate values are from 1000 iterations (Swofford 2002).
Sequences used to make antisense WMISH probes for Sp-nanos1, Sp-nanos2, and Sp-nanos3 were amplified from 48-hour embryonic cDNA and cloned into pGEMT-EZ. The Sp-nanos3 probe template includes the entire ORF plus 650 bases of the 3′UTR; the primer sets used for nested PCR are described above. Two Sp-nanos2 probes were used: the first is previously described in Juliano et al. (2006) and the second includes 950 bases of the 3′UTR. The primer set used to amplify the Sp-nanos2 3′UTR is as follows (listed 5′ to 3′): Sp-Nanos2-F2 - TGTAGGGTCTACTCGTCTCGA; Sp-Nanos2-R2 - CACCCAGCAATCAGTACTTTC. The Sp-nanos1 probe template includes the entire ORF plus 130 bases of the 3′UTR; the nested primer sets used for amplification are as follows: forward primers are described above; Sp-nanos1-R3 - AGAATGGAGTACTTGCGTAC; Sp-nanos1-R4 - ATACACCCAGCAATCAGTAC. The pGEMT-EZ plasmids were linearized using either SalI (T7 transcription) or NcoI (SP6 transcription) (Promega; Madison, WI). Antisense DIG-labeled RNA probes were constructed using a DIG RNA labeling kit (Roche; Indianapolis, IN). WMISH experiments were performed as previously described (Minokawa et al. 2004) and the alkaline phosphatase reaction was carried out for 22 hours. The Sp-vasa probe is described in Juliano et al., 2006. A non-specific DIG-labeled RNA probe complimentary to pSPT 18 (Roche; Indianapolis, IN) was used as a negative control. For uninjected embryos, all steps were carried out in 2mL screw-top tubes (National Scientific Supply; Claremont, CA). For injected and manipulated embryos, all steps were carried out in 96-well round-bottom PVC plates (ThermoFisher Scientific; Rockford, IL). Samples were imaged on a Zeiss Axiovert 200M microscope equipped with a Zeiss color AxioCam MRc5 camera (Carl Zeiss, Inc.; Thornwood, NY).
Sp-vasa FISH on mock-injected and Sp-nanos1 and 2 knockdown embryos was performed as previously described above through the blocking steps. Subsequently, samples were incubated overnight with anti-DIG-POD (Roche; Indianapolis, IN) diluted 1:1500 in blocking buffer II (Minokawa et. al., 2004) at room temperature with rotation. Samples were washed 6 times for 1 hour in TBST (10mM Tris-HCl, pH7.4; 0.15M NaCl; 0.1% Tween-20). Signal was detected using the Tyramide Signal Amplification (TSA) kit (Perkin Elmer; Waltham, MA). Samples were washed once with 1X amplification solution and then incubated with cyanine 3 TSA working solution for 15 minutes. Samples were washed 6 times with TBST, the 5th wash contained a 1:1000 dilution of a 10mg/mL Hoechst stock solution (Molecular Probes; Carlsbad, CA) for DNA labeling. Z-stacks were acquired for 5 mock-injected and 14 Sp-nanos1/2 MASO-injected embryos on an LSM 510 laser scanning confocal microscope (Carl Zeiss, Inc.; Thornwood, NY) and Sp-vasa-positive cells were counted.
RNA was extracted from 75 mock-injected and 75 Sp-nanos1/2 MASO-injected embryos at each time point (1 day, 3 days, 5 days, 7 days) using the RNeasy Micro Kit (Qiagen; Valencia, CA). cDNA was prepared using the TaqMan ® Reverse Transcription Reagents kit (Applied Biosystems; Foster City, CA). QPCR was performed on the 7300 Real-Time PCR system (Applied Biosystems; Foster City, CA) with the SYBER Green PCR Master Mix Kit (Applied Biosystems; Foster City, CA). Sp-nanos1 and 2 and Sp-vasa primer sets are described in Juliano et al. (2006). Experiments were run in triplicate and the data were normalized to ubiquitin RNA levels.
Full-length Sp-nanos2 was cloned downstream of the 6X-Histidine tag found in the pTAT vector (Nagahara et al. 1998). Recombinant protein was expressed in BL21 bacteria, purified on a ProBond nickel column (Invitrogen; Carlsbad, CA), and used to raise antiserum in rabbits as previously described (Wong and Wessel 2004). For affinity purification, recombinant Sp-Nanos2 protein was immobilized using the Pierce AminoLink Plus Immobilization Kit (ThermoFisher Scientific; Rockford, IL) as per the manufacturer’s instructions. Heat-inactivated antiserum was passed over the antigen-immobilized column and bound antibodies were eluted with 1mL 100mM glycine (pH 2.5) into 50uL 1M Tris (pH 9.5).
For immunoblot analysis of Sp-Nanos2, S. purpuratus uninjected and Sp-nanos1/2 MASO-injected embryos were cultured to the gastrula stage and were used to make embryo extracts as follows. Two hundred embryos were pelleted, resuspended in heated SDS sample loading buffer, boiled for 5 minutes, and DTT (Roche; Indianapolis, IN) was added at a final concentration of 5mM. Samples were incubated at 100°C for 5 minutes, spun at 14K RPM for 2 minutes, then loaded onto Tris-glycine, 4–20% gradient gels (Invitrogen; Carlsbad, CA). After transfer to nitrocellulose (Pall Corporation; Pensacola, FL), blots were probed with affinity-purified Sp-nanos2 antibody at 1μg/mL in Blotto (3% dry milk, 170mM NaCl, 50mM Tris, 0.05% Tween20). For visualization, blots were probed with an anti-rabbit-HRP secondary antibody diluted 1:5000 in Blotto (Jackson ImmunoResearch Laboratories; West Grove, PA), and visualized by standard ECL detection. Blots were stripped by two 30-minute incubations at 80°C in 200mM glycine-HCl, 0.05% Tween20 (pH 2.5) and then re-probed with α-YP30 serum diluted 1:30,000 in Blotto (Wessel et al. 2000).
Embryos were cultured as described above and samples were collected at indicated stages of development for whole mount antibody labeling. Embryos were fixed in 4% paraformaldehyde (Electron Microscopy Sciences; Hatfield, PA)/ASW for 10 minutes at room temperature, extracted in 100% MeOH (−20°C) for 1 minute, washed 3 times with PBS-Tween, and stored at 4°C. Antibody labeling was preceded by a blocking step of at least 30 minutes in 4% sheep serum (Sigma; St. Louis, MO) /PBS-Tween. For labeling, embryos were incubated overnight at 4°C with affinity-purified Sp-nanos2 antibody diluted in blocking buffer to a concentration of 10μg/mL. The embryos were washed 3 times with PBS-Tween and then incubated with anti-rabbit Cy3 conjugated antibody (Jackson ImmunoResearch; West Grove, PA) diluted 1:300 in blocking buffer for one hour at room temperature. The embryos were then washed 3 times with PBS-Tween. Labeled embryos were imaged on a Zeiss AxioPlan microscope (Carl Zeiss Inc.; Thornwood, NY) with an Orca-ER CCD camera (Hammamatsu Corporation; Bridgewater, NJ). For injected and manipulated samples the immunolabeling procedure was the same with the following exceptions: Samples were fixed in 24-well BSA-coated polystyrene plates (Corning; Corning, NY) or 9-well glass plates. Advanced larvae were fixed with 90% MeOH for 1 hour at −20°C. All subsequent labeling steps were done in BSA-coated 60-well mini trays (Nunc; Rochester, NY). For Sp-Vasa immunolabeling, Sp-vasa antibody was diluted in blocking buffer to a concentration of 10μg/mL (Voronina et al., 2008). The secondary antibody used was anti-rabbit Alexa Flour® 488 conjugated antibody (Invitrogen; Carlsbad, CA) diluted 1:500 in blocking buffer. Embryos and larvae were incubated with 1:1000 dilution of a 10mg/mL Hoechst stock solution for 10 minutes in PBS-Tween for DNA labeling (Molecular Probes; Carlsbad, CA) for DNA labeling. Labeled embryos and larvae were imaged on an LSM 510 laser scanning confocal microscope (Carl Zeiss, Inc.; Thornwood, NY).
Morpholino antisense oligos (MASO) directed against the Sp-nanos1 and/or Sp-nanos2 5′UTR were synthesized by Gene Tools (Philomath, OR). Microinjections of zygotes were performed as previously described (Cheers and Ettensohn 2004). In brief, eggs were de-jellied with acidic sea water (pH 5.0), washed with ASW or filtered sea water three times, rowed with a mouth pipette onto protamine sulfate-coated 60×15mm petri dishes, fertilized in the presence of 1mM 3-AT, and injected using the Femto Jet ® injection system (Eppendorf; Hamburg, Germany). To make the injection needles, 1×90 mm glass capillaries with filaments (Narishige; Tokyo, Japan) were pulled on a vertical needle puller (Narishige; Tokyo, Japan). Morpholino injection solutions were made as follows: 500μM MASO, 20% glycerol, 1mM 10,000 MW Dextran conjugated to Texas Red ® (Invitrogen; Carlsbad, CA). For rescue experiments, Sp-nanos1 mRNA with Xenopus β-globin UTRs was added to the morpholino injection solution to a final concentration of approximately 500ng/ L. Sp-nanos1 mRNA was made using an SP6 Message Machine kit as per the manufacture’s instructions (Ambion; Austin, TX). Mock injection solutions were identical, but without morpholino or RNA. Injected embryos were removed from the injection plate by mouth pipette and cultured at 16°C in noble agar-coated 24-well plates for up to 3 days. If larval stages were required, embryos were moved to pre-rinsed 15 or 50mL conical tubes (BD Falcon; Franklin Lakes, NJ) and rotated end over end at 16°C. Larvae were washed and fed every 3–4 days with the algal species Chaetoceros gracilis (UTEX; Austin, TX). To induce metamorphosis, between 6–8 weeks after fertilization, larvae were moved to 60×15mm petri dishes with biofilm and incubated at 16°C.
Zygotes were injected, as described above, with either Sp-nanos1/2 MASO or mock injection solutions. Both injected (donor) and uninjected (recipient) embryos were moved by mouth pipette from the injection plate to BSA-coated 60×15mm petri dishes. The process of removing the embryos from the injection plate removes the fertilization envelope. Embryos were cultured to the 16-cell stage and then micromere transplants were performed as previously described (Yajima 2007). However, instead of rhodamine, donor micromeres were labeled with Texas Red ® conjugated to dextran from the injection solution. After incorporation of the donor micromeres into the recipient embryo, transplanted embryos were moved to single wells of a polystyrene 96-well plate (Corning; Corning, NY) and cultured at 16°C. After 3 days, transplanted embryos were moved to individual 15mL conical tubes and cultured to late larval stages as described above. Images of transplanted embryos and larvae were taken on a Zeiss AxioPlan microscope (Carl Zeiss Inc.; Thornwood, NY) with an Orca-ER CCD camera (Hammamatsu Corporation; Bridgewater, NJ).
A computational search, using basic local alignment search tool (BLAST), of the S. purpuratus genome (spbase.org) revealed the presence of 3 nanos homologs (Altschul et al. 1990), which were designated as Sp-nanos1, Sp-nanos2, and Sp-nanos3 (Figure 1A; Juliano et al., 2006). The open reading frames (ORFs) of Sp-nanos1 and Sp-nanos2 have approximately 90% identical nucleotide sequences and nearly identical amino acid sequences. The most significant difference between the ORFs of Sp-nanos1 and Sp-nanos2 is an additional 12 amino acids (104–115) present in Sp-nanos2 (Figure 1A). However, Sp-nanos1 and Sp-nanos2 have different flanking sequences in the genome and thus appear to be two distinct genes rather than different alleles of the same gene. Furthermore, 3′ RACE analysis demonstrates that the Sp-nanos2 3′UTR is approximately 1.2 Kb. The Sp-nanos1 3′UTR is only 0.2Kb long and shares 60% sequence identity with the first 0.2 Kb of the Sp-nanos2 3′ UTR (data not shown). By contrast, Sp-nanos3 shares approximately 20% amino acid sequence identity over the entire ORF with Sp-nanos1 and Sp-nanos2 (Figure 1A). A phylogenetic analysis demonstrates that the three S. purpuratus nanos homologs cluster together on an unrooted neighbor joining tree, suggesting that they arose from recent duplication events (Figure 1B).
Previously published whole-mount RNA in situ hybridization (WMISH) results revealed that Sp-nanos2 is expressed in the small micromere lineage of the S. purpuratus embryo (Juliano et al. 2006). Here we demonstrate that Sp-nanos2 is first detected in the small micromeres at the 60-cell stage (Figure 2C). Probes directed against the Sp-nanos1 ORF and the Sp-nanos2 3′UTR also indicate that these transcripts accumulate in the small micromere lineage (Supp. Figure 1). However, it is not possible to make a specific Sp-nanos1 probe due to its short 3′UTR, and it is likely that WMISH probes directed against the ORFs of these two homologs significantly cross-hybridize. Sp-nanos3 transcripts accumulate specifically in the small micromere lineage of the gastrula and then are associated with the left coelomic pouch of the pluteus (Figure 2G-I). We conclude that all three Sp-nanos homologs are dynamically and specifically expressed in the small micromere lineage.
A polyclonal antibody generated against the entire Sp-Nanos2 ORF recognizes an approximately 24kDa band on an immunoblot, consistent with the 24.7kDa predicted molecular weight (MW) of Sp-nanos2 (Figure 3E and data not shown). The specificity of the antibody was tested by pre-incubation of the antibody with recombinant Sp-Nanos2 protein and reactivity was lost on an immunoblot when it was pre-incubated with recombinant Sp-Nanos2, thus confirming the specificity of the antibody (data not shown). However, given the nearly identical amino acid sequences of Sp-nanos1 and Sp-nanos2, this antibody likely recognizes both proteins. Since the predicted MW of Sp-Nanos1 (24.2 kD) is only 500Da less than Sp-Nanos2, we are not able to resolve these two homologs using SDS-PAGE. Immunofluorescence analysis demonstrates that Sp-Nanos1 and 2 proteins accumulate specifically in the small micromere lineage, similar to the Sp-nanos1and 2 transcripts (Figure 2J, K).
To determine the function of Sp-nanos1 and Sp-nanos2 homologs during sea urchin embryogenesis, we employed an antisense morpholino oligonucleotide (MASO) knock down strategy. The immediate sequences upstream of the start codon in the 5′UTR of both Sp-nanos1 and Sp-nanos2 are identical, thus allowing us to design one morpholino to simultaneously knockdown both Sp-Nanos1 and Sp-Nanos2 protein expression. This morpholino will be referred to as Sp-nanos1/2 MASO. Sp-nanos1/2 MASO was injected into newly fertilized zygotes, which were subsequently cultured to the gastrula stage and analyzed for Sp-Nanos1 and 2 protein expression. Sp-Nanos1 and 2 proteins were no longer detected at gastrula stages by immunoblot analysis or by immunofluorescence in Sp-nanos1/2 MASO-injected embryos as compared to uninjected controls (Figure 3). Therefore, this morpholino effectively knocks down Sp-Nanos1 and 2 proteins.
To examine the effect of Sp-nanos1 and 2 knockdown on small micromere development during embryogenesis we tested its effect on the expression of Sp-vasa, a well-characterized small micromere enriched gene (Voronina et al. 2008). Sp-vasa mRNA levels do not significantly change over the first 7 days of development after Sp-nanos1 and 2 knockdown as indicated by real-time quantitative PCR (QPCR) (data not shown). As shown by WMISH, the Sp-vasa mRNA accumulation pattern is not disrupted by Sp-nanos1 and 2 knockdown; Sp-vasa mRNA is detected at the vegetal plate of the blastula, in the small micromeres of the gastrula, and in both coelomic pouches of the pluteus in both Sp-nanos1 and 2 knockdown embryos and controls (Figure 4A-H). By contrast, when Sp-nanos1 and 2 are knocked down, Sp-Vasa protein accumulation in the small micromere descendants is reduced at the gastrula stage as compared to mock-injected controls (Figure 4K, L; Voronina et al. 2008).
Four small micromeres arise from an asymmetric division of the micromeres as the embryo develops from the 16-cell to 32-cell stage. Subsequently, the small micromeres divide once when in the vegetal plate of the blastula and then remain quiescent for the rest of embryogenesis. Therefore, 8 small micromere descendants travel at the tip of the archenteron and then are incorporated into the coelomic pouches of the pluteus. However, by WMISH, Sp-nanos1 and 2 knockdown gastrulae appear to have an increased number of cells at the tip of the archenteron that accumulate Sp-vasa and Sp-nanos1 and 2 transcripts. (Figure 4F, Figure 8G). To quantitate their number, fluorescent in situ hybridization (FISH) was used in combination with Hoechst labeling to count the number of Sp-vasa RNA-positive cells (Figure 4I, J). Sp-nanos1 and 2 knockdown gastrulae contain approximately twice the number of Sp-vasa RNA-positive cells at the tip of the archenteron. This may indicate that neighboring cells have acquired Sp-vasa mRNA expression. However, given that the number of Sp-vasa mRNA-positive cells is approximately double, it is more likely that the small micromere descendants have undergone one extra cell division. Therefore, Sp-nanos1 and 2 may be required to maintain the mitotically quiescent state of the small micromeres during embryogenesis, an important feature of set-aside cells.
Aside from the changes in the small micromere lineage described above, Sp-nanos1 and 2 knockdown embryos proceed through embryogenesis normally, leading to formation of a pluteus larva, similar to mock-injected controls (Figure 5A, D). However, when larvae that develop from Sp-nanos1 and 2 knockdown embryos are fed and cultured, their coelomic pouches fail to grow and the rudiment does not form (Figure 5E, F, J, O). With prolonged culture, Sp-nanos1 and 2 knockdown larvae die after 4–6 weeks; these larvae are smaller than controls, but this lack of growth does not appear to be due to an inability to feed because the gut contracts and algae are present in the stomach. By contrast, larvae that develop from mock-injected embryos under the same culture conditions construct adult rudiments and can be induced to metamorphose (Figure 5B, C, G, M, N).
Two-week-old control and Sp-nanos1 and 2 knockdown larvae were assessed for Sp-Vasa protein accumulation by immunofluorescence (Figure 5H, I, K, L). In control larvae Sp-Vasa protein is detected in the developing adult rudiment as well as the amniotic invagination (Figure 5H, I). Of note here is that the amniotic invagination is of ectodermal origin, and thus the expression of Sp-Vasa protein in this region must be new (Pearse and Cameron 1991). This is in contrast to the Vasa-positive cells in the developing rudiment, many of which are likely small micromere descendants. In larvae that develop from Sp-nanos1 and 2 knockdown embryos, only a small number of Vasa-positive cells are detected in the poorly developing coelomic pouch and the amniotic invagination does not form (Figure 5K, L).
To determine the relative contributions of Sp-nanos1 and Sp-nanos2 to coelomic pouch formation, two additional morpholinos were designed to specifically target each nanos homolog. This was possible because approximately 40 bases upstream of the starting methionine the 5′UTR sequences of Sp-nanos1 and Sp-nanos2 diverge. Sp-nanos1 knockdown larvae lacked significant coelomic pouches at one week and did not recover them, similar to Sp-nanos1 and 2 knockdown larvae (Figure 6B). Sp-nanos2 knockdown larvae initially displayed a delay in coelomic pouch growth, but recovered and developed a normal adult rudiment (Figure 6C, data not shown). When Sp-nanos1 and Sp-nanos2 are knocked down together by co-injection of both specific morpholinos, the phenotype is more severe than Sp-nanos1 knockdown alone (Figure 6D). Therefore, both Sp-nanos1 and Sp-nanos2 play an important role in coelomic pouch development, but Sp-nanos1 is relatively more important as larvae cannot form an adult rudiment when this gene is knocked down by itself.
To test if the aberrant phenotype induced by the Sp-nanos1 morpholino is due entirely to the loss of Sp-nanos1 expression, a rescue experiment was performed (Figure 6E, F). When Sp-nanos1 mRNA with a morpholino-insensitive 5′UTR was co-injected with the Sp-nanos1 morpholino, larvae were able to recover both their proper shape and size, and coelomic pouches developed normally (Figure 6F). Furthermore, the Vasa-positive amniotic invagination was observed, which is not of coelomic pouch origin (Figure 6F’). Thus, the dramatic larval phenotypes observed in Sp-nanos1 morpholino-injected larvae are due entirely to the loss of Sp-nanos1 expression.
We next tested the effect of Sp-nanos1 and 2 knockdown specifically in the micromere lineage of an otherwise wild type embryo. To accomplish this, micromeres were transplanted from an Sp-nanos1 and 2 knockdown (Sp-nanos1/2 MASO plus red dextran) or mock-injected (red dextran only) 16-cell embryo to an uninjected recipient 16-cell embryo whose micromeres were previously removed (Figure 7A). This experiment allows us to follow the fate of Sp-nanos1 and 2 knockdown small micromeres during development. Furthermore, it tests if the requirement for Sp-nanos1 and 2 in coelomic pouch development is specific to the micromere lineage. This lineage gives rise both to the small micromere lineage and to the large micromere lineage, which contributes solely to the larval skeleton.
After each transplant experiment, in both Sp-nanos1 and 2 knockdown embryos and controls, labeled small micromere descendants were observed at the tip of the archenteron (data not shown). Furthermore in 5/5 control transplant embryos, labeled cells were observed in both coelomic pouches of an early pluteus (Figure 7B, C). However, in 7/8 Sp-nanos1 and 2 knockdown transplants, labeled cells were not observed in coelomic pouches indicating that the small micromere descendants were not incorporated there (Figure 7 E, F). In the one case where small micromeres were observed in the coelomic pouches, the intensity of the fluorescence was notably lower likely indicating a lower amount of morpholino injected in the donor embryo. The presence of red, large micromere-derived, skeletal cells in each case confirms a successful micromere transplant (Figure 7D, G).
Larvae that develop from embryos with transplanted Sp-nanos1 and 2 knockdown micromeres are able to develop adult rudiments to varying degrees (Figure 7H-M). This is in contrast to larvae that develop from Sp-nanos1/2 MASO-injected zygotes, which are not able to make an adult rudiment (Figure 5O). This result may indicate that in a wildtype embryo Sp-nanos1 and/or Sp-nanos2 are required outside of the small micromere lineage for proper coelomic pouch formation and adult rudiment development. Neither gene product is detected by WMISH outside of this lineage in wildtype embryos or larvae through one week of development, but this does not rule out low levels of expression in additional cell lineages. Alternatively, this result may indicate that when the descendants of Sp-nanos1 and 2 knockdown small micromeres are not incorporated into the coelomic pouches, new Sp-nanos1 and/or 2 expression in non-small micromere descendants rescues the adult rudiment. This rescue would not be possible when the Sp-nanos1/2 MASO is ubiquitously present because newly transcribed Sp-nanos1 and 2 mRNA would not be translated. In support of this latter hypothesis, Sp-nanos1 and 2 knockdown embryos exhibit a 2-fold increase in Sp-nanos1 and 2 mRNA accumulation, suggesting that the embryo compensates for the loss of Sp-Nanos1 and 2 protein by upregulating Sp-nanos1 and/or 2 transcription (Figure 8A). Furthermore, after Sp-nanos1 and 2 knockdown, Sp-nanos1 and 2 mRNA is detectable in the left coelomic pouch several days into larval development by WMISH (Figure 8F-I). By contrast, in control larvae, Sp-nanos1 and 2 mRNA is no longer detected by WMISH shortly after pluteus formation (Figure 8B-E). Given that the Sp-nanos1 and 2 knockdown small micromeres are not incorporated into the coelomic pouches (see Figure 7), this result suggests that an alternative cell population is accumulating Sp-nanos1 and/or 2 mRNA in the coelomic pouch of Sp-nanos1 and 2 knockdown larvae. Furthermore, this new population of Sp-nanos1 and/or 2-positive cells may be responsible for the recovery of the adult rudiment observed in our transplant experiments (see Figure 7).
Sp-nanos1 and 2 are required for the incorporation of the small micromere descendants into the larval coelomic pouches and are therefore critical to maintain small micromere fate. Furthermore, in the absence of Sp-nanos1 and 2, coelomic pouches do not develop and the adult rudiment does not form, thus the functions of these genes are required for juvenile development (Figure 9). Given that Sp-nanos1 and 2 are specifically expressed in the small micromere lineage, this suggests that these cells are multipotent and give rise to multiple juvenile tissues. Alternatively, the small micromere descendants could have a critical signaling role in directing proliferation and fate determination in other cells that contribute to coelomic pouch and adult rudiment formation. These two possibilities are not mutually exclusive and currently we cannot distinguish between their relative contributions to adult rudiment development. The small micromere descendants proliferate rapidly upon their arrival at the coelomic pouch, which supports the hypothesis that these cells contribute substantially to the adult rudiment, and are therefore multipotent (Tanaka and Dan 1990). The complete lack of adult rudiment structures in the Sp-nanos1 and 2 knockdowns also implies that other cells that contribute to juvenile development require the presence of the small micromere lineage to adopt their fates. For example, the cells of the amniotic invagination do not invaginate in Sp-nanos1 and 2 knockdown larvae. Thus, the small micromere lineage may contribute to adult rudiment development both by contributing directly to the tissues and in an instructive manner.
Although the sea urchin embryo is patterned very early by maternal inputs and zygotic gene expression, some of the cells retain plasticity, thus allowing for remarkable regulative capacity. For example, the large micromere descendants give rise to the primary mesenchyme cells (PMCs), which ingress into the blastocoel before gastrulation and eventually give rise to the entire larval skeleton. However, if the PMCs are surgically removed from the early gastrula then a population of non-skeletal mesenchyme cells transfate into skeletal mesenchyme cells and a larval skeleton is built (Ettensohn and McClay 1988; Ettensohn et al. 2007). If the micromeres are removed at the 16-cell stage then both the large and small micromere lineages are lost. However, the resulting larvae have a skeleton and develop an adult rudiment that produces normal, fertile adults (Ransick et al. 1996). Given the significant contribution of the small micromeres to the coelomic pouches, this result suggests that micromere-deleted embryos successfully recover multipotent cells for adult rudiment formation.
Recent work demonstrated that the regulation of Sp-vasa expression might play an important role in the recovery of multipotent small micromere fates (Voronina et al. 2008). When the Vasa-positive micromeres are removed, Vasa protein expression is upregulated in the entire embryo, followed by a restriction to a subset of cells, which may then take on the fate of the former small micromere lineage (Voronina et al. 2008). Here we demonstrate that Sp-nanos1 and 2 knockdown embryos develop into larvae that cannot produce an adult rudiment. Interestingly, when Sp-nanos1 and 2 knockdown is restricted to the micromere lineage, the resulting larvae are capable of producing an adult rudiment, despite the fact that the small micromeres are not incorporated into the coelomic pouches. This result implies that the embryo is able to recover the lost population of multipotent cells early in larval development. New expression of Sp-nanos1 and/or 2 must be required for the specification of new multipotent cells because adult rudiments cannot form when the Sp-nanos1/2 MASO is ubiquitously present. In Sp-nanos1 and 2 knockdown embryos, accumulation of Sp-nanos1 and 2 RNA is increased and persists in the coelomic pouch further into larval development. Given that macromere descendants are the only other lineage to populate the coelomic pouches, these cells may take on small micromere fates when the small micromere descendants are lost.
Nanos is required to maintain germline fate throughout the Drosophila life cycle and appears to have a similar function in maintaining germline fate in other organisms. In C. elegans embryos and larvae, nanos is required to maintain repressive chromatin marks and cell cycle arrest (Subramaniam and Seydoux 1999; Schaner et al. 2003). In the mouse, nanos3−/− PGCs undergo apoptosis (Tsuda et al. 2003; Suzuki et al. 2008). In adult germlines nanos is required to maintain oocyte production in zebrafish and spermatogonia populations in mice (Draper et al. 2007; Lolicato et al. 2008). In the Xenopus oocyte, cell cycle arrest requires nanos-directed repression of cyclinB (Nakahata et al. 2001; Nakahata et al. 2003). Therefore, nanos has a conserved function in preserving animal germlines by maintaining a quiescent state.
The function of nanos in the small micromeres appears to be analogous to its conserved germline functions. Small micromere descendants in Sp-nanos1 and 2 knockdown embryos may undergo an extra division and are not incorporated into the coelomic pouches. We do not know what happens to Sp-nanos1 and 2 knockdown small micromeres, but two possibilities exist. First, they may undergo apoptosis, similar to nanos−/− pole cells in Drosophila and nanos3−/− PGCs in mice (Tsuda et al. 2003; Hayashi et al. 2004; Sato et al. 2007; Suzuki et al. 2008). Alternatively, nanos-depleted small micromere descendants may take on differentiated cell fates and thus become incorporated into other larval structures. This outcome would be analogous to Drosophila pole cells taking on somatic cell fates when the apoptosis pathway is suppressed (Hayashi et al. 2004). However, no evidence of small micromere incorporation into alternative larval structures was detected in our transplant experiments, thus we favor the hypothesis that nanos-depleted small micromeres undergo apoptosis.
The small micromere lineage specifically expresses vasa, nanos, and piwi, all of which have conserved roles in germline determination and maintenance, yet the small micromeres do not appear to give rise exclusively to germ cells (Juliano et al. 2006). We propose that the sea urchin uses a two-step germline determination mechanism. The first step is the formation of an embryonic multipotent precursor under the control of a conserved molecular program, which includes specific expression of nanos, vasa, and, piwi. In the second step of this mechanism the multipotent precursors give rise to both somatic cells and to germ cells later in development. In the sea urchin, this likely occurs as the adult rudiment forms during larval development. This mechanism is in contrast to animals that only exhibit specific expression of vasa, nanos, and piwi in cells fated to be exclusively germline, which we refer to here as one-step germline determination. The two-step germline determination mechanism was also proposed for P. dumerilii, a lophotrochozoan that segregates its germline late in larval development from a multipotent precursor established during embryogenesis. This lineage of multipotent cells is termed the mesodermal growth zone (MPGZ) and like the small micromere lineage of the sea urchin, cells of the MPGZ specifically express vasa, nanos, and piwi (Rebscher et al. 2007). Vasa and nanos also are expressed specifically in the 4d lineage of Ilyanassa, another lophotrochozoan and loss of nanos leads to significant loss of adult structures derived from this lineage (Rabinowitz et al. 2008; Swartz et al. 2008). This same program also may operate in the multipotent I-cells of the adult Hydra, a cnidarian. Vasa and nanos are specifically expressed in Hydra I-cells, which give rise both to somatic cells and to germ cells (Mochizuki et al. 2000; Mochizuki et al. 2001). Conservation of the gene expression profiles and of nanos function in PGCs and various multipotent cell lineages suggests the presence of a conserved molecular program shared in these cell types. What is the evolutionary relationship between cells that contain this program among metazoans? We propose the following two mutually exclusive hypotheses to explain how the small micromeres obtained this program:
Maximal indirect development is prevalent in both the lophotrochozoans and in echinoderms, although substantial evidence suggests that these branches independently acquired this form of development (Raff 2008). Therefore, a correlative link exists between maximal indirect development and the two-step germline determination mechanism. If the two-step germline determination is ancestral (Figure 10A) then perhaps it was the retention of this mechanism that facilitated the independent acquisition of maximal indirect development. In such a case, pre-existing multipotent precursors could easily be set aside for use in the adult, thus protecting the germline during the transition. Furthermore, the acquisition of maximal indirect development may have locked many echinoderms and lophotrochozoans into an ancestral two-step germline determination mechanism. By contrast, if two-step germline determination is derived (Figure 10B), then it may have independently evolved in these two groups with maximal indirect development. It has been hypothesized that the larvae of maximal indirect developers co-opted GRNs from the adult for use in building its larval structures (Sly et al. 2003). For example, the GRN used to build the sea urchin larval skeleton was likely co-opted from the adult skeletogenic GRN (Gao and Davidson 2008). Therefore, if two-step germline determination is derived, it is possible that the small micromere lineage co-opted its molecular program from adult PGCs.
These models are based on data from a limited number of animal species. To discern the evolutionary origins of the small micromere molecular program it is critical to decorate the tree with data from a wide variety of organisms. However, based on this study and those reported in Hydra, planarians, Ilyanassa, and P. dumerilii it is clear that genes traditionally classified as “germline genes” have a broader role in maintaining multipotency (Shibata et al. 1999; Mochizuki et al. 2000; Mochizuki et al. 2001; Reddien et al. 2005; Salvetti et al. 2005; Guo et al. 2006; Juliano et al. 2006; Rebscher et al. 2007; Palakodeti et al. 2008; Rabinowitz et al. 2008; Swartz et al. 2008; Voronina et al. 2008). Specification and maintenance of potency in all of the cell types that express genes such as vasa, nanos, and piwi likely share a common underlying mechanism of regulatory control. Thus, comparing these mechanisms in diverse animals that use varied developmental strategies will allow us to uncover the critical and ancestral portions of this program.
Supplemental Figure 1. Sp-nanos1 and Sp-nanos2 mRNA accumulates in the small micromere lineage. WMISH was performed on the indicated embryonic stages of S. purpuratus. Sp-nanos1 and Sp-nanos2 share approximately 90% identity in their ORFs. The Sp-nanos2 probe is specific because it targets the divergent 3′UTR. The Sp-nanos1 probe targets both the ORF and the significantly shorter 3′UTR, thus some cross-reactivity to Sp-nanos2 may occur with this probe. Sp-nanos1 is first detected in the small micromeres at the 60-cell-stage, remains associated with this lineage in the blastula, and can no longer be detected in the gastrula. Sp-nanos2 is first detected in the small micromere lineage at the vegetal plate of the blastula, remains associated with this lineage in the gastrula, and is no longer detected in the pluteus. The negative control is a non-specific probe directed against pSPT 18. Arrows indicate the Sp-nanos mRNA accumulation in the small micromere lineage.
We thank all current and past members of PRIMO for helpful discussions. Specifically we thank Zak Swartz and Adrian Reich for help in editing this manuscript. We thank two anonymous reviewers for helpful and critical feedback on this manuscript. This work was supported by grants from the NIH and the NSF.
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