Abstract
A fertilized egg is initially transcriptionally silent and relies on maternally provided factors to initiate development. For embryonic development to proceed, the oocyte-inherited cytoplasm and the nuclear chromatin need to be reprogrammed to create a permissive environment for zygotic genome activation (ZGA). During this maternal-to-zygotic transition (MZT), which is conserved in metazoans, transient totipotency is induced and zygotic transcription is initiated to form the blueprint for future development. Recent technological advances have enhanced our understanding of MZT regulation, revealing common themes across species and leading to new fundamental insights about transcription, mRNA decay and translation.
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References
Gurdon, J. B. The developmental capacity of nuclei taken from intestinal epithelium cells of feeding tadpoles. Development 10, 622–640 (1962).
Takahashi, K. & Yamanaka, S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126, 663–676 (2006).
Vastenhouw, N. L., Cao, W. X. & Lipshitz, H. D. The maternal-to-zygotic transition revisited. Development 146, dev161471 (2019).
Despic, V. & Neugebauer, K. M. RNA tales—how embryos read and discard messages from mom. J. Cell Sci. 131, jcs201996 (2018).
Wu, Q. & Bazzini, A. A. Translation and mRNA stability control. Annu. Rev. Biochem. 92, 227–245 (2023).
Eckersley-Maslin, M. A., Alda-Catalinas, C. & Reik, W. Dynamics of the epigenetic landscape during the maternal-to-zygotic transition. Nat. Rev. Mol. Cell Biol. 19, 436–450 (2018).
Zhang, Y. & Xie, W. Building the genome architecture during the maternal to zygotic transition. Curr. Opin. Genet. Dev. 72, 91–100 (2022).
Ing-Simmons, E., Rigau, M. & Vaquerizas, J. M. Emerging mechanisms and dynamics of three-dimensional genome organisation at zygotic genome activation. Curr. Opin. Cell Biol. 74, 37–46 (2022).
Collart, C. et al. High-resolution analysis of gene activity during the Xenopus mid-blastula transition. Development 141, 1927–1939 (2014).
Heyn, P. et al. The earliest transcribed zygotic genes are short, newly evolved, and different across species. Cell Rep. 6, 285–292 (2014).
Owens, N. D. L. et al. Measuring absolute RNA copy numbers at high temporal resolution reveals transcriptome kinetics in development. Cell Rep. 14, 632–647 (2016).
Kwasnieski, J. C., Orr-Weaver, T. L. & Bartel, D. P. Early genome activation in Drosophila is extensive with an initial tendency for aborted transcripts and retained introns. Genome Res. 29, 1188–1197 (2019).
Aoki, F., Worrad, D. M. & Schultz, R. M. Regulation of transcriptional activity during the first and second cell cycles in the preimplantation mouse embryo. Dev. Biol. 181, 296–307 (1997).
Knowland, J. & Graham, C. RNA synthesis at the two-cell stage of mouse development. Development 27, 167–176 (1972).
Vassena, R. et al. Waves of early transcriptional activation and pluripotency program initiation during human preimplantation development. Development 138, 3699–3709 (2011).
Schulz, K. N. & Harrison, M. M. Mechanisms regulating zygotic genome activation. Nat. Rev. Genet. 10, 622 (2018).
Leesch, F. et al. A molecular network of conserved factors keeps ribosomes dormant in the egg. Nature 613, 712–720 (2023). This study uses mass spectrometry and cryo electron microscopy to establish that the low translational activity in oocytes and early embryos is caused by the action of four factors that maintain ribosomes in a ‘dormant’ state.
Kobayashi, W. et al. Nucleosome-bound NR5A2 structure reveals pioneer factor mechanism by DNA minor groove anchor competition. Nat. Struct. Mol. Biol. 31, 757–766 (2024).
Michael, A. K. et al. Mechanisms of OCT4-SOX2 motif readout on nucleosomes. Science 368, 1460–1465 (2020).
Echigoya, K. et al. Nucleosome binding by the pioneer transcription factor OCT4. Sci. Rep. 10, 11832 (2020).
Lerner, J., Katznelson, A., Zhang, J. & Zaret, K. S. Different chromatin-scanning modes lead to targeting of compacted chromatin by pioneer factors FOXA1 and SOX2. Cell Rep. 42, 112748 (2023).
Martinez-Sarmiento, J. A., Cosma, M. P. & Lakadamyali, M. Dissecting gene activation and chromatin remodeling dynamics in single human cells undergoing reprogramming. Cell Rep. 43, 114170 (2024).
Garcia, H. G., Tikhonov, M., Lin, A. & Gregor, T. Quantitative imaging of transcription in living Drosophila embryos links polymerase activity to patterning. Curr. Biol. 23, 2140–2145 (2013).
Lucas, T. et al. Live imaging of bicoid-dependent transcription in Drosophila embryos. Curr. Biol. 23, 2135–2139 (2013).
Hoppe, C. et al. Modulation of the promoter activation rate dictates the transcriptional response to graded BMP signaling levels in the Drosophila embryo. Dev. Cell 54, 727–741 (2020).
Pimmett, V. L. et al. Quantitative imaging of transcription in living Drosophila embryos reveals the impact of core promoter motifs on promoter state dynamics. Nat. Commun. 12, 4504 (2021).
Dufourt, J. et al. Imaging translation dynamics in live embryos reveals spatial heterogeneities. Science 372, 840–844 (2021).
Vinter, D. J., Hoppe, C., Minchington, T. G., Sutcliffe, C. & Ashe, H. L. Dynamics of hunchback translation in real-time and at single-mRNA resolution in the Drosophila embryo. Development 148, dev196121 (2021).
Pownall, M. E. et al. Chromatin expansion microscopy reveals nanoscale organization of transcription and chromatin. Science 381, 92–100 (2023). This study describes a new ChromExM method providing higher-resolution insights into nuclear organization and describes the ‘kiss-and-kick’ model of transcriptional activation, where transcriptional elongation kicks (displaces) enhancers away from the promoter.
Bhat, P. et al. SLAMseq resolves the kinetics of maternal and zygotic gene expression during early zebrafish embryogenesis. Cell Rep. 42, 112070 (2023).
Zhang, B. et al. Allelic reprogramming of the histone modification H3K4me3 in early mammalian development. Nature 537, 553–557 (2016).
Dahl, J. A. et al. Broad histone H3K4me3 domains in mouse oocytes modulate maternal-to-zygotic transition. Nature 537, 548–552 (2016).
Liu, X. et al. Distinct features of H3K4me3 and H3K27me3 chromatin domains in pre-implantation embryos. Nature 537, 558–562 (2016).
Zheng, H. et al. Resetting epigenetic memory by reprogramming of histone modifications in mammals. Mol. Cell 63, 1066–1079 (2016).
Wu, Y. et al. N6-Methyladenosine regulates maternal RNA maintenance in oocytes and timely RNA decay during mouse maternal-to-zygotic transition. Nat. Cell Biol. 24, 917–927 (2022).
Flyamer, I. M. et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 544, 110–114 (2017).
Ke, Y. et al. 3D chromatin structures of mature gametes and structural reprogramming during mammalian embryogenesis. Cell 170, 367–381.e20 (2017).
Du, Z. et al. Allelic reprogramming of 3D chromatin architecture during early mammalian development. Nature 547, 232–235 (2017).
Bazzini, A. A., Lee, M. T. & Giraldez, A. J. Ribosome profiling shows that miR-430 reduces translation before causing mRNA decay in zebrafish. Science 336, 233–237 (2012).
Subtelny, A. O., Eichhorn, S. W., Chen, G. R., Sive, H. & Bartel, D. P. Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature 508, 66–71 (2014).
Kronja, I. et al. Widespread changes in the posttranscriptional landscape at the Drosophila oocyte-to-embryo transition. Cell Rep. 7, 1495–1508 (2014).
Chen, L. et al. Global regulation of mRNA translation and stability in the early Drosophila embryo by the Smaug RNA-binding protein. Genome Biol. 15, R4 (2014).
Eichhorn, S. W. et al. mRNA poly(A)-tail changes specified by deadenylation broadly reshape translation in Drosophila oocytes and early embryos. eLife 5, e16955 (2016).
Xiang, K. & Bartel, D. P. The molecular basis of coupling between poly(A)-tail length and translational efficiency. eLife 10, e66493 (2021). This paper demonstrates that the limited levels of PABPC in early embryos provide a molecular explanation for the strong correlation between poly(A) tail length and translational efficiency during early embryonic development, as originally described by Subtelny et al. (2014).
Lee, M. T. et al. Nanog, Pou5f1 and SoxB1 activate zygotic gene expression during the maternal-to-zygotic transition. Nature 503, 360–364 (2013). This study, through ribosome footprinting, discovers three TFs that are highly translated in the early zebrafish embryo and demonstrates that collectively these factors activate a large subset of the first zygotic genes.
Zhang, C., Wang, M., Li, Y. & Zhang, Y. Profiling and functional characterization of maternal mRNA translation during mouse maternal-to-zygotic transition. Sci. Adv. 8, eabj3967 (2022).
Xiong, Z. et al. Ultrasensitive Ribo-seq reveals translational landscapes during mammalian oocyte-to-embryo transition and pre-implantation development. Nat. Cell Biol. 24, 968–980 (2022).
Zou, Z. et al. Translatome and transcriptome co-profiling reveals a role of TPRXs in human zygotic genome activation. Science 378, abo7923 (2022). This study employs low-input ribosome profiling to investigate the translatome in human oocytes and early embryos, identifying TRPX family TFs as key regulators of human ZGA.
Lorenzo-Orts, L. et al. eIF4E1b is a non-canonical eIF4E protecting maternal dormant mRNAs. EMBO Rep. 25, 404–427 (2023).
Shan, L.-Y. et al. LSM14B controls oocyte mRNA storage and stability to ensure female fertility. Cell. Mol. Life Sci. 80, 247 (2023).
Sun, J., Yan, L., Shen, W. & Meng, A. Maternal Ybx1 safeguards zebrafish oocyte maturation and maternal-to-zygotic transition by repressing global translation. Development 145, dev166587 (2018).
Bouvet, P. & Wolffe, A. P. A role for transcription and FRGY2 in masking maternal mRNA within Xenopus oocytes. Cell 77, 931–941 (1994).
Medvedev, S., Pan, H. & Schultz, R. M. Absence of MSY2 in mouse oocytes perturbs oocyte growth and maturation, RNA stability, and the transcriptome1. Biol. Reprod. 85, 575–583 (2011).
Ivshina, M., Lasko, P. & Richter, J. D. Cytoplasmic polyadenylation element binding proteins in development, health, and disease. Annu. Rev. Cell Dev. Biol. 30, 1–23 (2014).
Voeltz, G. K., Ongkasuwan, J., Standart, N. & Steitz, J. A. A novel embryonic poly(A) binding protein, ePAB, regulates mRNA deadenylation in Xenopus egg extracts. Genes Dev. 15, 774–788 (2001).
Passmore, L. A. & Coller, J. Roles of mRNA poly(A) tails in regulation of eukaryotic gene expression. Nat. Rev. Mol. Cell Biol. 23, 93–106 (2022).
Voeltz, G. K. & Steitz, J. A. AUUUA sequences direct mRNA deadenylation uncoupled from decay during xenopus early development. Mol. Cell. Biol. 18, 7537–7545 (1998).
Lee, K., Cho, K., Morey, R. & Cook-Andersen, H. An extended wave of global mRNA deadenylation sets up a switch in translation regulation across the mammalian oocyte-to-embryo transition. Cell Rep. 43, 113710 (2024).
Gillian-Daniel, D. L., Gray, N. K., Åström, J., Barkoff, A. & Wickens, M. Modifications of the 5′ cap of mRNAs during Xenopus oocyte maturation: independence from changes in poly(A) length and impact on translation. Mol. Cell. Biol. 18, 6152–6163 (1998).
Zhang, S., Williams, C. J., Wormington, M., Stevens, A. & Peltz, S. W. Monitoring mRNA decapping activity. Methods 17, 46–51 (1999).
Yang, Y. et al. RNA 5-methylcytosine facilitates the maternal-to-zygotic transition by preventing maternal mRNA decay. Mol. Cell 75, 1188–1202 (2019).
Liu, J. et al. Developmental mRNA m5C landscape and regulatory innovations of massive m5C modification of maternal mRNAs in animals. Nat. Commun. 13, 2484 (2022).
Mendez, R. et al. Phosphorylation of CPE binding factor by Eg2 regulates translation of c-mos mRNA. Nature 404, 302–307 (2000).
Lim, J., Lee, M., Son, A., Chang, H. & Kim, V. N. mTAIL-seq reveals dynamic poly(A) tail regulation in oocyte-to-embryo development. Genes Dev. 30, 1671–1682 (2016).
Aanes, H. et al. Zebrafish mRNA sequencing deciphers novelties in transcriptome dynamics during maternal to zygotic transition. Genome Res. 21, 1328–1338 (2011).
Piqué, M., López, J. M., Foissac, S., Guigó, R. & Méndez, R. A combinatorial code for CPE-mediated translational control. Cell 132, 434–448 (2008).
Winata, C. L. et al. Cytoplasmic polyadenylation-mediated translational control of maternal mRNAs directs maternal-to-zygotic transition. Development 145, dev159566 (2017).
Sheets, M. D., Ogg, S. C. & Wickens, M. P. Point mutations in AAUAAA and the poly(A) addition site: effects on the accuracy and efficiency of cleavage and polyadenylation in vitro. Nucleic Acids Res. 18, 5799–5805 (1990).
Sheets, M. D., Fox, C. A., Hunt, T., Woude, G. V. & Wickens, M. The 3′-untranslated regions of c-mos and cyclin mRNAs stimulate translation by regulating cytoplasmic polyadenylation. Genes Dev. 8, 926–938 (1994).
Xiang, K., Ly, J. & Bartel, D. P. Control of poly(A)-tail length and translation in vertebrate oocytes and early embryos. Dev. Cell 59, 1058–1074 (2024).
Aoki, F., Hara, K. T. & Schultz, R. M. Acquisition of transcriptional competence in the 1‐cell mouse embryo: requirement for recruitment of maternal mRNAs. Mol. Reprod. Dev. 64, 270–274 (2003).
Liu, Y. et al. Remodeling of maternal mRNA through poly(A) tail orchestrates human oocyte-to-embryo transition. Nat. Struct. Mol. Biol. 30, 200–215 (2023).
Ulitsky, I. et al. Extensive alternative polyadenylation during zebrafish development. Genome Res. 22, 2054–2066 (2012).
Takada, Y. et al. Mature mRNA processing that deletes 3′ end sequences directs translational activation and embryonic development. Sci. Adv. 9, eadg6532 (2023).
Lim, J. et al. Mixed tailing by TENT4A and TENT4B shields mRNA from rapid deadenylation. Science 361, 701–704 (2018).
Wang, M. et al. ME31B globally represses maternal mRNAs by two distinct mechanisms during the Drosophila maternal-to-zygotic transition. eLife 6, e27891 (2017).
Hara, M. et al. Identification of PNG kinase substrates uncovers interactions with the translational repressor TRAL in the oocyte-to-embryo transition. eLife 7, e33150 (2018).
Lorenzo-Orts, L. & Pauli, A. The molecular mechanisms underpinning maternal mRNA dormancy. Biochem. Soc. Trans. 52, 861–871 (2024).
Bashirullah, A. et al. Joint action of two RNA degradation pathways controls the timing of maternal transcript elimination at the midblastula transition in Drosophila melanogaster. EMBO J. 18, 2610–2620 (1999).
Tadros, W. et al. SMAUG is a major regulator of maternal mRNA destabilization in Drosophila and its translation is activated by the PAN GU kinase. Dev. Cell 12, 143–155 (2007). This study shows that the SMAUG RBP facilitates the destabilization of maternal transcripts in D. melanogaster, demonstrating how a single RBP can broadly influence global mRNA levels during the MZT.
Giraldez, A. J. et al. Zebrafish miR-430 promotes deadenylation and clearance of maternal mRNAs. Science 312, 75–79 (2006).
Lund, E., Liu, M., Hartley, R. S., Sheets, M. D. & Dahlberg, J. E. Deadenylation of maternal mRNAs mediated by miR-427 in Xenopus laevis embryos. RNA 15, 2351–2363 (2009).
Baia Amaral, D., Egidy, R., Perera, A. & Bazzini, A. A. miR-430 regulates zygotic mRNA during zebrafish embryogenesis. Genome Biol. 25, 74 (2024).
Bartel, D. P. Metazoan microRNAs. Cell 173, 20–51 (2018).
Lytle, J. R., Yario, T. A. & Steitz, J. A. Target mRNAs are repressed as efficiently by microRNA-binding sites in the 5′ UTR as in the 3′ UTR. Proc. Natl Acad. Sci. USA 104, 9667–9672 (2007).
Moretti, F., Thermann, R. & Hentze, M. W. Mechanism of translational regulation by miR-2 from sites in the 5′ untranslated region or the open reading frame. RNA 16, 2493–2502 (2010).
Strayer, E. C. et al. NaP-TRAP, a novel massively parallel reporter assay to quantify translation control. Preprint at bioRxiv https://doi.org/10.1101/2023.11.09.566434 (2023).
Kloosterman, W. P., Wienholds, E., Ketting, R. F. & Plasterk, R. H. A. Substrate requirements for let-7 function in the developing zebrafish embryo. Nucleic Acids Res. 32, 6284–6291 (2004).
Bushati, N., Stark, A., Brennecke, J. & Cohen, S. M. Temporal reciprocity of miRNAs and their targets during the maternal-to-zygotic transition in Drosophila. Curr. Biol. 18, 501–506 (2008).
Hadzhiev, Y. et al. The miR-430 locus with extreme promoter density forms a transcription body during the minor wave of zygotic genome activation. Dev. Cell 58, 155–170 (2023).
Tani, S., Kusakabe, R., Naruse, K., Sakamoto, H. & Inoue, K. Genomic organization and embryonic expression of miR-430 in medaka (Oryzias latipes): insights into the post-transcriptional gene regulation in early development. Gene 449, 41–49 (2010).
Jiménez-Ruiz, C. A. et al. miR-430 microRNA family in fishes: molecular characterization and evolution. Animals 13, 2399 (2023).
Suh, N. et al. microRNA function is globally suppressed in mouse oocytes and early embryos. Curr. Biol. 20, 271–277 (2010).
Wu, S., Aksoy, M., Shi, J. & Houbaviy, H. B. Evolution of the miR-290–295/miR-371–373 cluster family seed repertoire. PLoS ONE 9, e108519 (2014).
Medeiros, L. A. et al. miR-290–295 deficiency in mice results in partially penetrant embryonic lethality and germ cell defects. Proc. Natl Acad. Sci. USA 108, 14163–14168 (2011).
Suh, M.-R. et al. Human embryonic stem cells express a unique set of microRNAs. Dev. Biol. 270, 488–498 (2004).
Houbaviy, H. B., Murray, M. F. & Sharp, P. A. Embryonic stem cell-specific microRNAs. Dev. Cell 5, 351–358 (2003).
Judson, R. L., Babiarz, J. E., Venere, M. & Blelloch, R. Embryonic stem cell–specific microRNAs promote induced pluripotency. Nat. Biotechnol. 27, 459–461 (2009).
Subramanyam, D. et al. Multiple targets of miR-302 and miR-372 promote reprogramming of human fibroblasts to induced pluripotent stem cells. Nat. Biotechnol. 29, 443–448 (2011).
Zhang, J.-M. et al. Argonaute 2 is a key regulator of maternal mRNA degradation in mouse early embryos. Cell Death Discov. 6, 133 (2020).
Stoeckius, M. et al. Global characterization of the oocyte‐to‐embryo transition in Caenorhabditis elegans uncovers a novel mRNA clearance mechanism. EMBO J. 33, 1751–1766 (2014).
Quarato, P. et al. Germline inherited small RNAs facilitate the clearance of untranslated maternal mRNAs in C. elegans embryos. Nat. Commun. 12, 1441 (2021).
Barckmann, B. et al. Aubergine iCLIP reveals piRNA-dependent decay of mRNAs involved in germ cell development in the early embryo. Cell Rep. 12, 1205–1216 (2015).
Kontur, C., Jeong, M., Cifuentes, D. & Giraldez, A. J. Ythdf m6A readers function redundantly during zebrafish development. Cell Rep. 33, 108598 (2020).
Zhao, B. S. et al. m6A-dependent maternal mRNA clearance facilitates zebrafish maternal-to-zygotic transition. Nature 542, 475–478 (2017).
Ivanova, I. et al. The RNA m6A reader YTHDF2 is essential for the post-transcriptional regulation of the maternal transcriptome and oocyte competence. Mol. Cell 67, 1059–1067 (2017).
Yao, H. et al. scm6A-seq reveals single-cell landscapes of the dynamic m6A during oocyte maturation and early embryonic development. Nat. Commun. 14, 315 (2023).
Wang, Y. et al. The RNA m6A landscape of mouse oocytes and preimplantation embryos. Nat. Struct. Mol. Biol. 30, 703–709 (2023).
Despic, V. et al. Dynamic RNA–protein interactions underlie the zebrafish maternal-to-zygotic transition. Genome Res. 27, 1184–1194 (2017).
Vejnar, C. E. et al. Genome wide analysis of 3′-UTR sequence elements and proteins regulating mRNA stability during maternal-to-zygotic transition in zebrafish. Genome Res. 29, 1100–1114 (2019).
Paillard, L. et al. EDEN and EDEN‐BP, a cis element and an associated factor that mediate sequence‐specific mRNA deadenylation in Xenopus embryos. EMBO J. 17, 278–287 (1998).
Semotok, J. L. et al. Smaug recruits the CCR4/POP2/NOT deadenylase complex to trigger maternal transcript localization in the early Drosophila embryo. Curr. Biol. 15, 284–294 (2005).
Laver, J. D. et al. Brain tumor is a sequence-specific RNA-binding protein that directs maternal mRNA clearance during the Drosophila maternal-to-zygotic transition. Genome Biol. 16, 94 (2015).
De Renzis, S., Elemento, O., Tavazoie, S. & Wieschaus, E. F. Unmasking activation of the zygotic genome using chromosomal deletions in the Drosophila embryo. PLoS Biol. 5, e117 (2007).
D’Agostino, I., Merritt, C., Chen, P.-L., Seydoux, G. & Subramaniam, K. Translational repression restricts expression of the C. elegans Nanos homolog NOS-2 to the embryonic germline. Dev. Biol. 292, 244–252 (2006).
Rabani, M., Pieper, L., Chew, G.-L. & Schier, A. F. A massively parallel reporter assay of 3′ UTR sequences identifies in vivo rules for mRNA degradation. Mol. Cell 68, 1083–1094 (2017).
Audic, Y., Omilli, F. & Osborne, H. B. Embryo deadenylation element-dependent deadenylation is enhanced by a cis element containing AUU repeats. Mol. Cell. Biol. 18, 6879–6884 (1998).
Ramos, S. B. V. et al. The CCCH tandem zinc-finger protein Zfp36l2 is crucial for female fertility and early embryonic development. Development 131, 4883–4893 (2004).
Detivaud, L., Pascreau, G., Karaïskou, A., Osborne, H. B. & Kubiak, J. Z. Regulation of EDEN-dependent deadenylation of Aurora A/Eg2-derived mRNA via phosphorylation and dephosphorylation in Xenopus laevis egg extracts. J. Cell Sci. 116, 2697–2705 (2003).
Haugen, R. J. et al. Regulation of the Drosophila transcriptome by Pumilio and the CCR4-NOT deadenylase complex. RNA 30, 866–890 (2024).
Liu, Y. et al. BTG4 is a key regulator for maternal mRNA clearance during mouse early embryogenesis. J. Mol. Cell Biol. 8, 366–368 (2016).
Yu, C. et al. BTG4 is a meiotic cell cycle-coupled maternal-zygotic-transition licensing factor in oocytes. Nat. Struct. Mol. Biol. 23, 387–394 (2016).
Zhao, L. et al. PABPN1L mediates cytoplasmic mRNA decay as a placeholder during the maternal‐to‐zygotic transition. EMBO Rep. 21, e49956 (2020).
Pinder, B. D. & Smibert, C. A. microRNA‐independent recruitment of Argonaute 1 to nanos mRNA through the Smaug RNA‐binding protein. EMBO Rep. 14, 80–86 (2013).
Nelson, M. R., Leidal, A. M. & Smibert, C. A. Drosophila Cup is an eIF4E‐binding protein that functions in Smaug‐mediated translational repression. EMBO J. 23, 150–159 (2004).
Zheng, W. et al. Homozygous mutations in BTG4 cause zygotic cleavage failure and female infertility. Am. J. Hum. Genet. 107, 24–33 (2020).
Reimão-Pinto, M. M., Castillo-Hair, S. M., Seelig, G. & Schier, A. F. The regulatory landscape of 5′ UTRs in translational control during zebrafish embryogenesis. Preprint at bioRxiv https://doi.org/10.1101/2023.11.23.568470 (2023).
Lim, J. et al. Uridylation by TUT4 and TUT7 marks mRNA for degradation. Cell 159, 1365–1376 (2014).
Chang, H. et al. Terminal uridylyltransferases execute programmed clearance of maternal transcriptome in vertebrate embryos. Mol. Cell 70, 72–82 (2018).
Liu, Y., Nie, H., Liu, H. & Lu, F. Poly(A) inclusive RNA isoform sequencing (PAIso-seq) reveals wide-spread non-adenosine residues within RNA poly(A) tails. Nat. Commun. 10, 5292 (2019).
Sha, Q.-Q. et al. Characterization of zygotic genome activation-dependent maternal mRNA clearance in mouse. Nucleic Acids Res. 48, 879–894 (2020).
Sha, Q.-Q. et al. Dynamics and clinical relevance of maternal mRNA clearance during the oocyte-to-embryo transition in humans. Nat. Commun. 11, 4917 (2020).
Morgan, M. et al. mRNA 3′ uridylation and poly(A) tail length sculpt the mammalian maternal transcriptome. Nature 548, 347–351 (2017).
Presnyak, V. et al. Codon optimality is a major determinant of mRNA stability. Cell 160, 1111–1124 (2015).
Bazzini, A. A. et al. Codon identity regulates mRNA stability and translation efficiency during the maternal‐to‐zygotic transition. EMBO J. 35, 2087–2103 (2016).
Mishima, Y. & Tomari, Y. Codon usage and 3′ UTR length determine maternal mRNA stability in zebrafish. Mol. Cell 61, 874–885 (2016). Together with Bazzini et al. (2016), this work identifies codon optimality as a key regulator of transcript stability during the MZT in zebrafish and many other species.
Buschauer, R. et al. The Ccr4-Not complex monitors the translating ribosome for codon optimality. Science 368, eaay6912 (2020).
Medina-Muñoz, S. G. et al. Crosstalk between codon optimality and cis-regulatory elements dictates mRNA stability. Genome Biol. 22, 14 (2021).
Rappol, T. et al. tRNA expression and modification landscapes, and their dynamics during zebrafish embryo development. Nucleic Acids Res 52, 10575–10594 (2024).
Reimão-Pinto, M. M. et al. The dynamics and functional impact of tRNA repertoires during early embryogenesis in zebrafish. EMBO J. https://doi.org/10.1038/s44318-024-00265-4 (2024).
Chen, K. Y., Park, H. & Subramaniam, A. R. Massively parallel identification of sequence motifs triggering ribosome-associated mRNA quality control. Nucleic Acids Res 52, 7171–7187 (2024).
Burke, P. C., Park, H. & Subramaniam, A. R. A nascent peptide code for translational control of mRNA stability in human cells. Nat. Commun. 13, 6829 (2022).
Calvo, S. E., Pagliarini, D. J. & Mootha, V. K. Upstream open reading frames cause widespread reduction of protein expression and are polymorphic among humans. Proc. Natl Acad. Sci. USA 106, 7507–7512 (2009).
Johnstone, T. G., Bazzini, A. A. & Giraldez, A. J. Upstream ORFs are prevalent translational repressors in vertebrates. EMBO J. 35, 706–723 (2016).
Hurt, J. A., Robertson, A. D. & Burge, C. B. Global analyses of UPF1 binding and function reveal expanded scope of nonsense-mediated mRNA decay. Genome Res. 23, 1636–1650 (2013).
Chan, L. Y., Mugler, C. F., Heinrich, S., Vallotton, P. & Weis, K. Non-invasive measurement of mRNA decay reveals translation initiation as the major determinant of mRNA stability. eLife 7, e32536 (2018).
May, G. E. et al. Unraveling the influences of sequence and position on yeast uORF activity using massively parallel reporter systems and machine learning. eLife 12, e69611 (2023).
Jia, L. et al. Decoding mRNA translatability and stability from the 5′ UTR. Nat. Struct. Mol. Biol. 27, 814–821 (2020).
Chew, G.-L., Pauli, A. & Schier, A. F. Conservation of uORF repressiveness and sequence features in mouse, human and zebrafish. Nat. Commun. 7, 11663 (2016).
Musaev, D. et al. UPF1 regulates mRNA stability by sensing poorly translated coding sequences. Cell Rep. 43, 114074 (2024).
Pickering, B. M., Mitchell, S. A., Spriggs, K. A., Stoneley, M. & Willis, A. E. Bag-1 internal ribosome entry segment activity is promoted by structural changes mediated by poly(rC) binding protein 1 and recruitment of polypyrimidine tract binding protein 1. Mol. Cell. Biol. 24, 5595–5605 (2004).
Kozak, M. Circumstances and mechanisms of inhibition of translation by secondary structure in eucaryotic mRNAs. Mol. Cell. Biol. 9, 5134–5142 (1989).
Beaudoin, J.-D. et al. Analyses of mRNA structure dynamics identify embryonic gene regulatory programs. Nat. Struct. Mol. Biol. 25, 677–686 (2018).
Shi, B. et al. RNA structural dynamics regulate early embryogenesis through controlling transcriptome fate and function. Genome Biol. 21, 120 (2020).
Jentoft, I. M. A. et al. Mammalian oocytes store proteins for the early embryo on cytoplasmic lattices. Cell 186, 5308–5327 (2023). This paper shows that cytoplasmic lattice structures in mouse oocytes are essential for stabilizing maternally deposited proteins, a process required for proper embryonic development.
Mitchell, L. E. Maternal effect genes: update and review of evidence for a link with birth defects. Hum. Genet. Genom. Adv. 3, 100067 (2021).
Zhang, H. et al. Stable maternal proteins underlie distinct transcriptome, translatome, and proteome reprogramming during mouse oocyte-to-embryo transition. Genome Biol. 24, 166 (2023).
Peshkin, L. et al. On the relationship of protein and mRNA dynamics in vertebrate embryonic development. Dev. Cell 35, 383–394 (2015).
Cao, W. X. et al. Precise temporal regulation of post-transcriptional repressors is required for an orderly Drosophila maternal-to-zygotic transition. Cell Rep. 31, 107783 (2020).
Nguyen, T. et al. Differential nuclear import sets the timing of protein access to the embryonic genome. Nat. Commun. 13, 5887 (2022).
Ryu, S., Holzschuh, J., Erhardt, S., Ettl, A.-K. & Driever, W. Depletion of minichromosome maintenance protein 5 in the zebrafish retina causes cell-cycle defect and apoptosis. Proc. Natl Acad. Sci. USA 102, 18467–18472 (2005).
Zavortink, M. et al. The E2 Marie Kondo and the CTLH E3 ligase clear deposited RNA binding proteins during the maternal-to-zygotic transition. eLife 9, e53889 (2020).
Yang, Y. et al. The E3 ubiquitin ligase RNF114 and TAB1 degradation are required for maternal‐to‐zygotic transition. EMBO Rep. 18, 205–216 (2017).
Shen, W. et al. Comprehensive maturity of nuclear pore complexes regulates zygotic genome activation. Cell 185, 4954–4970 (2022). Together with Nguyen et al. (2022) (X. laevis), this work (zebrafish) shows that the nuclear import of maternally deposited factors plays a pivotal role in initiating ZGA.
Chen, K. et al. A global change in RNA polymerase II pausing during the Drosophila midblastula transition. eLife 2, e00861 (2013).
Abe, K. et al. The first murine zygotic transcription is promiscuous and uncoupled from splicing and 3′ processing. EMBO J. 34, 1523–1537 (2015).
Cvetesic, N. et al. Global regulatory transitions at core promoters demarcate the mammalian germline cycle. Preprint at bioRxiv https://doi.org/10.1101/2020.10.30.361865 (2020).
Zaret, K. S. Pioneer transcription factors initiating gene network changes. Annu. Rev. Genet. 54, 1–19 (2020).
Liang, H.-L. et al. The zinc-finger protein Zelda is a key activator of the early zygotic genome in Drosophila. Nature 456, 400 403 (2008). This paper is the first to identify a sequence-specific TF regulating ZGA in any organism, identifying Zelda as a key ZGA regulator in D. melanogaster.
Nien, C.-Y. et al. Temporal coordination of gene networks by Zelda in the early Drosophila embryo. PLoS Genet. 7, e1002339 (2011).
Harrison, M. M., Li, X.-Y., Kaplan, T., Botchan, M. R. & Eisen, M. B. Zelda binding in the early Drosophila melanogaster embryo marks regions subsequently activated at the maternal-to-zygotic transition. PLoS Genet. 7, e1002266 (2011).
Leichsenring, M., Maes, J., Mössner, R., Driever, W. & Onichtchouk, D. Pou5f1 transcription factor controls zygotic gene activation in vertebrates. Science 341, 1005–1009 (2013).
Miao, L. et al. The landscape of pioneer factor activity reveals the mechanisms of chromatin reprogramming and genome activation. Mol. Cell 82, 986–1002 (2022). This study demonstrates how the PFs Nanog, Pou5f3 and Sox19b collaborate in zebrafish to make chromatin accessible for ZGA, while also showing that histone acetylation can bypass the need for these factors in initiating transcription.
Riesle, A. J. et al. Activator-blocker model of transcriptional regulation by pioneer-like factors. Nat. Commun. 14, 5677 (2023).
Gao, M. et al. Pluripotency factors determine gene expression repertoire at zygotic genome activation. Nat. Commun. 13, 788 (2022).
Gentsch, G. E., Spruce, T., Owens, N. D. L. & Smith, J. C. Maternal pluripotency factors initiate extensive chromatin remodelling to predefine first response to inductive signals. Nat. Commun. 10, 4269 (2019).
Charney, R. M. et al. Foxh1 occupies cis-regulatory modules prior to dynamic transcription factor interactions controlling the mesendoderm gene program. Dev. Cell 40, 595–607 (2017).
Paraiso, K. D. et al. Endodermal maternal transcription factors establish super-enhancers during zygotic genome activation. Cell Rep. 27, 2962–2977 (2019).
Gaskill, M. M., Gibson, T. J., Larson, E. D. & Harrison, M. M. GAF is essential for zygotic genome activation and chromatin accessibility in the early Drosophila embryo. eLife 10, e66668 (2021).
Duan, J. et al. CLAMP and Zelda function together to promote Drosophila zygotic genome activation. eLife 10, e69937 (2021).
Colonnetta, M. M., Abrahante, J. E., Schedl, P., Gohl, D. M. & Deshpande, G. CLAMP regulates zygotic genome activation in Drosophila embryos. Genetics 219, iyab107 (2021).
Soluri, I. V., Zumerling, L. M., Parra, O. A. P., Clark, E. G. & Blythe, S. A. Zygotic pioneer factor activity of Odd-paired/Zic is necessary for late function of the Drosophila segmentation network. eLife 9, e53916 (2020).
Koromila, T. et al. Odd-paired is a pioneer-like factor that coordinates with Zelda to control gene expression in embryos. eLife 9, e59610 (2020).
Ji, S. et al. OBOX regulates mouse zygotic genome activation and early development. Nature 620, 1047–1053 (2023).
Golbus, M. S., Calarco, P. G. & Epstein, C. J. The effects of inhibitors of RNA synthesis (α-amanitin and actinomycin D) on preimplantation mouse embryogenesis. J. Exp. Zool. 186, 207–216 (1973).
Maeso, I. et al. Evolutionary origin and functional divergence of totipotent cell homeobox genes in eutherian mammals. BMC Biol. 14, 45 (2016).
Gassler, J. et al. Zygotic genome activation by the totipotency pioneer factor Nr5a2. Science 378, 1305–1315 (2022).
Festuccia, N. et al. Nr5a2 is dispensable for zygotic genome activation but essential for morula development. Science 386, eadg7325 (2024).
Wu, J. et al. The landscape of accessible chromatin in mammalian preimplantation embryos. Nature 534, 652–657 (2016).
Lai, F. et al. NR5A2 connects zygotic genome activation to the first lineage segregation in totipotent embryos. Cell Res. 33, 952–966 (2023).
Zhao, Y. et al. Nr5a2 ensures inner cell mass formation in mouse blastocyst. Cell Rep. 43, 113840 (2024).
Li, L. et al. Lineage regulators TFAP2C and NR5A2 function as bipotency activators in totipotent embryos. Nat. Struct. Mol. Biol. 31, 950–963 (2024).
Lu, F. et al. Establishing chromatin regulatory landscape during mouse preimplantation development. Cell 165, 1375–1388 (2016).
Choi, S. H. et al. DUX4 recruits p300/CBP through its C-terminus and induces global H3K27 acetylation changes. Nucleic Acids Res. 44, 5161–5173 (2016).
Hendrickson, P. G. et al. Conserved roles of mouse DUX and human DUX4 in activating cleavage-stage genes and MERVL/HERVL retrotransposons. Nat. Genet. 49, 925–934 (2017).
De Iaco, A. et al. DUX-family transcription factors regulate zygotic genome activation in placental mammals. Nat. Genet. 49, 941–945 (2017).
Whiddon, J. L., Langford, A. T., Wong, C.-J., Zhong, J. W. & Tapscott, S. J. Conservation and innovation in the DUX4-family gene network. Nat. Genet. 49, 935–940 (2017). Together with Ji et al. (2023), Gassler et al. (2022), Lu et al. (2016), Hendrickson et al. (2017) and De Iaco et al. (2017), this work identifies important sequence-specific TFs contributing to chromatin accessibility and activation of early expressed genes in mouse embryos; however, these and subsequent studies suggest redundancy among some of these TFs in activating the earliest ZGA genes.
Chen, Z. & Zhang, Y. Loss of DUX causes minor defects in zygotic genome activation and is compatible with mouse development. Nat. Genet. 51, 947–951 (2019).
De Iaco, A., Verp, S., Offner, S., Grun, D. & Trono, D. DUX is a non-essential synchronizer of zygotic genome activation. Development 147, dev177725 (2019).
Guo, Y. et al. Obox4 promotes zygotic genome activation upon loss of Dux. eLife 13, e95856 (2024).
Frederick, M. A. et al. A pioneer factor locally opens compacted chromatin to enable targeted ATP-dependent nucleosome remodeling. Nat. Struct. Mol. Biol. 30, 31–37 (2023).
Kubinyecz, O. N. et al. Maternal SMARCA5 is required for major ZGA in mouse embryos. Preprint at bioRxiv https://doi.org/10.1101/2023.12.05.570276 (2023).
Bultman, S. J. et al. Maternal BRG1 regulates zygotic genome activation in the mouse. Genes Dev. 20, 1744–1754 (2006).
Li, X.-Y., Harrison, M. M., Villalta, J. E., Kaplan, T. & Eisen, M. B. Establishment of regions of genomic activity during the Drosophila maternal to zygotic transition. eLife 3, e03737 (2014).
Cho, C.-Y. & O’Farrell, P. H. Stepwise modifications of transcriptional hubs link pioneer factor activity to a burst of transcription. Nat. Commun. 14, 4848 (2023).
Schulz, K. N. et al. Zelda is differentially required for chromatin accessibility, transcription factor binding, and gene expression in the early Drosophila embryo. Genome Res. 25, 1715–1726 (2015).
Sun, Y. et al. Zelda overcomes the high intrinsic nucleosome barrier at enhancers during Drosophila zygotic genome activation. Genome Res. 25, 1703–1714 (2015). Together with Schulz et al. (2015), this work reports studies in D. melanogaster which established that the ZGA-initiating TF Zelda can open regions of closed nucleosomal chromatin, a hallmark of pioneer factors, paving the way for subsequent research on genome activation.
Veil, M., Yampolsky, L., Gruening, B. & Onichtchouk, D. Pou5f3, SoxB1, and Nanog remodel chromatin on high nucleosome affinity regions at zygotic genome activation. Genome Res. 29, 383–395 (2019).
Pálfy, M., Schulze, G., Valen, E. & Vastenhouw, N. L. Chromatin accessibility established by Pou5f3, Sox19b and Nanog primes genes for activity during zebrafish genome activation. PLoS Genet. 16, e1008546 (2020).
Xu, Z. et al. Impacts of the ubiquitous factor Zelda on Bicoid-dependent DNA binding and transcription in Drosophila. Genes Dev. 28, 608–621 (2014).
Brennan, K. J. et al. Chromatin accessibility in the Drosophila embryo is determined by transcription factor pioneering and enhancer activation. Dev. Cell 58, 1898–1916 (2023).
Yamada, S. et al. The Drosophila pioneer factor Zelda modulates the nuclear microenvironment of a dorsal target enhancer to potentiate transcriptional output. Curr. Biol. 29, 1387–1393 (2019).
Hansen, J. L., Loell, K. J. & Cohen, B. A. A test of the pioneer factor hypothesis using ectopic liver gene activation. eLife 11, e73358 (2022).
Li, L. et al. Multifaceted SOX2–chromatin interaction underpins pluripotency progression in early embryos. Science 382, eadi5516 (2023).
Gibson, T. J., Larson, E. D. & Harrison, M. M. Protein-intrinsic properties and context-dependent effects regulate pioneer factor binding and function. Nat. Struct. Mol. Biol. 31, 548–558 (2024).
Pluta, R. et al. Molecular basis for DNA recognition by the maternal pioneer transcription factor FoxH1. Nat. Commun. 13, 7279 (2022).
Soufi, A. et al. Pioneer transcription factors target partial DNA motifs on nucleosomes to initiate reprogramming. Cell 161, 555–568 (2015).
Zhu, F. et al. The interaction landscape between transcription factors and the nucleosome. Nature 562, 76–81 (2018).
Fernandez Garcia, M. et al. Structural features of transcription factors associating with nucleosome binding. Mol. Cell 75, 921–932.e6 (2019).
Sönmezer, C. et al. Molecular co-occupancy identifies transcription factor binding cooperativity in vivo. Mol. Cell 81, 255–267.e6 (2021).
Larson, E. D., Marsh, A. J. & Harrison, M. M. Pioneering the developmental frontier. Mol. Cell 81, 1640–1650 (2021).
Soufi, A., Donahue, G. & Zaret, K. S. Facilitators and impediments of the pluripotency reprogramming factors’ initial engagement with the genome. Cell 151, 994–1004 (2012).
Sinha, K. K., Bilokapic, S., Du, Y., Malik, D. & Halic, M. Histone modifications regulate pioneer transcription factor cooperativity. Nature 619, 378–384 (2023).
Chronis, C. et al. Cooperative binding of transcription factors orchestrates reprogramming. Cell 168, 442–459 (2017).
Li, S., Zheng, E. B., Zhao, L. & Liu, S. Nonreciprocal and conditional cooperativity directs the pioneer activity of pluripotency transcription factors. Cell Rep. 28, 2689–2703 (2019).
Li, D. et al. Chromatin accessibility dynamics during iPSC reprogramming. Cell Stem Cell 21, 819–833 (2017).
Gaskill, M. M. et al. Localization of the Drosophila pioneer factor GAF to subnuclear foci is driven by DNA binding and required to silence satellite repeat expression. Dev. Cell 58, 1610–1624 (2023).
Zou, Z., Wang, Q., Wu, X., Schultz, R. M. & Xie, W. Kick-starting the zygotic genome: licensors, specifiers, and beyond. EMBO Rep. 25, 4113–4130 (2024).
Akdogan-Ozdilek, B., Duval, K. L. & Goll, M. G. Chromatin dynamics at the maternal to zygotic transition: recent advances from the zebrafish model. F1000Res. 9, 299 (2020).
Veenstra, G. J. C. Dynamics of chromatin remodeling during embryonic evelopment. In Xenopus: From Basic Biology to Disease Models in the Genomic Era (eds Fainsod, A. & Moody, S. A.) 173–184 (CRC, 2022).
Wilkinson, A. L., Zorzan, I. & Rugg-Gunn, P. J. Epigenetic regulation of early human embryo development. Cell Stem Cell 30, 1569–1584 (2023).
Harrison, M. M., Marsh, A. J. & Rushlow, C. A. Setting the stage for development: the maternal-to-zygotic transition in Drosophila. Genetics 225, iyad142 (2023).
Potok, M. E., Nix, D. A., Parnell, T. J. & Cairns, B. R. Reprogramming the maternal zebrafish genome after fertilization to match the paternal methylation pattern. Cell 153, 759–772 (2013).
Jiang, L. et al. Sperm, but not oocyte, DNA methylome is inherited by zebrafish early embryos. Cell 153, 773–784 (2013).
Veenstra, G. J. C. & Wolffe, A. P. Constitutive genomic methylation during embryonic development of Xenopus. Biochim. Biophys. Acta. 1521, 39–44 (2001).
Dimitrov, S., Almouzni, G., Dasso, M. & Wolffe, A. P. Chromatin transitions during early xenopus embryogenesis: changes in histone H4 acetylation and in linker histone type. Dev. Biol. 160, 214–227 (1993).
Dworkin-Rastl, E., Kandolf, H. & Smith, R. C. The maternal histone H1 variant, H1M (B4 Protein), is the predominant H1 histone in Xenopus pregastrula embryos. Dev. Biol. 161, 425–439 (1994).
Freedman, B. S. & Heald, R. Functional comparison of H1 histones in xenopus reveals isoform-specific regulation by Cdk1 and RanGTP. Curr. Biol. 20, 1048–1052 (2010).
Pérez-Montero, S., Carbonell, A., Morán, T., Vaquero, A. & Azorín, F. The embryonic linker histone H1 variant of Drosophila, dBigH1, regulates zygotic genome activation. Dev. Cell 26, 578–590 (2013).
Hergeth, S. P. & Schneider, R. The H1 linker histones: multifunctional proteins beyond the nucleosomal core particle. EMBO Rep. 16, 1439–1453 (2015).
Funaya, S., Ooga, M., Suzuki, M. G. & Aoki, F. Linker histone H1FOO regulates the chromatin structure in mouse zygotes. FEBS Lett. 592, 2414–2424 (2018).
Henn, L. et al. Alternative linker histone permits fast paced nuclear divisions in early Drosophila embryo. Nucleic Acids Res. 48, 9007–9018 (2020).
Hammoud, S. S. et al. Distinctive chromatin in human sperm packages genes for embryo development. Nature 460, 473–478 (2009).
Brykczynska, U. et al. Repressive and active histone methylation mark distinct promoters in human and mouse spermatozoa. Nat. Struct. Mol. Biol. 17, 679–687 (2010).
Loppin, B. et al. The histone H3.3 chaperone HIRA is essential for chromatin assembly in the male pronucleus. Nature 437, 1386–1390 (2005).
Lin, C.-J., Koh, F. M., Wong, P., Conti, M. & Ramalho-Santos, M. Hira-mediated H3.3 incorporation is required for DNA replication and ribosomal RNA transcription in the mouse zygote. Dev. Cell 30, 268–279 (2014).
Inoue, A. & Zhang, Y. Nucleosome assembly is required for nuclear pore complex assembly in mouse zygotes. Nat. Struct. Mol. Biol. 21, 609–616 (2014).
Szenker, E., Lacoste, N. & Almouzni, G. A developmental requirement for HIRA-dependent H3.3 deposition revealed at gastrulation in Xenopus. Cell Rep. 1, 730–740 (2012).
Ishiuchi, T. et al. Reprogramming of the histone H3.3 landscape in the early mouse embryo. Nat. Struct. Mol. Biol. 28, 38–49 (2021).
Wen, D. et al. Histone variant H3.3 is an essential maternal factor for oocyte reprogramming. Proc. Natl Acad. Sci. USA 111, 7325–7330 (2014).
Cheloufi, S. et al. The histone chaperone CAF-1 safeguards somatic cell identity. Nature 528, 218–224 (2015).
Ibarra-Morales, D. et al. Histone variant H2A.Z regulates zygotic genome activation. Nat. Commun. 12, 7002 (2021).
Murphy, P. J., Wu, S. F., James, C. R., Wike, C. L. & Cairns, B. R. Placeholder nucleosomes underlie germline-to-embryo DNA methylation reprogramming. Cell 172, 993–1006 (2018).
Hurton, M. D., Miller, J. M. & Lee, M. T. H3K4me2 distinguishes a distinct class of enhancers during the maternal-to-zygotic transition. Preprint at bioRxiv https://doi.org/10.1101/2024.08.26.609713 (2024).
Liu, X. et al. Hierarchical accumulation of histone variant H2A.Z regulates transcriptional states and histone modifications in early mammalian embryos. Adv. Sci. 9, 2200057 (2022).
Zhang, B. et al. Widespread enhancer dememorization and promoter priming during parental-to-zygotic transition. Mol. Cell 72, 673–686 (2018).
Akkers, R. C. et al. A hierarchy of H3K4me3 and H3K27me3 acquisition in spatial gene regulation in xenopus embryos. Dev. Cell 17, 425–434 (2009).
Zhu, W., Xu, X., Wang, X. & Liu, J. Reprogramming histone modification patterns to coordinate gene expression in early zebrafish embryos. BMC Genomics 20, 248 (2019).
Hörmanseder, E. et al. H3K4 methylation-dependent memory of somatic cell identity inhibits reprogramming and development of nuclear transfer embryos. Cell Stem Cell 21, 135–143 (2017).
Mazzetto, M., Gonzalez, L. E., Sanchez, N. & Reinke, V. Characterization of the distribution and dynamics of chromatin states in the C. elegans germline reveals substantial H3K4me3 remodeling during oogenesis. Genome Res. 34, 57–69 (2024).
Vastenhouw, N. L. et al. Chromatin signature of embryonic pluripotency is established during genome activation. Nature 464, 922–926 (2010).
Lindeman, L. C. et al. Prepatterning of developmental gene expression by modified histones before zygotic genome activation. Dev. Cell 21, 993–1004 (2011).
Hontelez, S. et al. Embryonic transcription is controlled by maternally defined chromatin state. Nat. Commun. 6, 10148 (2015).
Haberle, V. et al. Two independent transcription initiation codes overlap on vertebrate core promoters. Nature 507, 381 (2014). This study shows that maternal and zygotic transcripts in zebrafish utilize distinct TSSs, highlighting the differences in promoter grammar between the maternal and zygotic states.
Clouaire, T. et al. Cfp1 integrates both CpG content and gene activity for accurate H3K4me3 deposition in embryonic stem cells. Genes Dev. 26, 1714–1728 (2012).
van Heeringen, S. J. et al. Principles of nucleation of H3K27 methylation during embryonic development. Genome Res. 24, 401–410 (2014).
Zenk, F. et al. Germ line-inherited H3K27me3 restricts enhancer function during maternal-to-zygotic transition. Science 357, 212–216 (2017).
Xia, W. et al. Resetting histone modifications during human parental-to-zygotic transition. Science 365, 353–360 (2019).
Inoue, A., Jiang, L., Lu, F., Suzuki, T. & Zhang, Y. Maternal H3K27me3 controls DNA methylation-independent imprinting. Nature 547, 419–424 (2017).
Chen, Z., Djekidel, M. N. & Zhang, Y. Distinct dynamics and functions of H2AK119ub1 and H3K27me3 in mouse preimplantation embryos. Nat. Genet. 53, 551–563 (2021).
Samata, M. et al. Intergenerationally maintained histone H4 lysine 16 acetylation is instructive for future gene activation. Cell 182, 127–144 (2020).
Chan, S. H. et al. Brd4 and P300 confer transcriptional competency during zygotic genome activation. Dev. Cell 49, 867–881 (2019).
Gupta, R., Wills, A., Ucar, D. & Baker, J. Developmental enhancers are marked independently of zygotic Nodal signals in Xenopus. Dev. Biol. 395, 38–49 (2014).
Sato, Y. et al. Histone H3K27 acetylation precedes active transcription during zebrafish zygotic genome activation as revealed by live-cell analysis. Development 146, dev179127 (2019).
Wang, M., Chen, Z. & Zhang, Y. CBP/p300 and HDAC activities regulate H3K27 acetylation dynamics and zygotic genome activation in mouse preimplantation embryos. EMBO J. 41, e112012 (2022).
Sakamoto, M. et al. Detection of newly synthesized RNA reveals transcriptional reprogramming during ZGA and a role of Obox3 in totipotency acquisition. Cell Rep. 43, 114118 (2024).
Ciabrelli, F. et al. CBP and Gcn5 drive zygotic genome activation independently of their catalytic activity. Sci. Adv. 9, eadf2687 (2023).
Zhou, J. J. et al. Histone deacetylase 1 maintains lineage integrity through histone acetylome refinement during early embryogenesis. eLife 12, e79380 (2023).
Theis, A. & Harrison, M. M. Reprogramming of three-dimensional chromatin organization in the early embryo. Curr. Opin. Struct. Biol. 81, 102613 (2023).
Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).
Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).
Zenk, F. et al. HP1 drives de novo 3D genome reorganization in early Drosophila embryos. Nature 593, 289–293 (2021).
Strom, A. R. et al. Phase separation drives heterochromatin domain formation. Nature 547, 241–245 (2017).
Hug, C. B., Grimaldi, A. G., Kruse, K. & Vaquerizas, J. M. Chromatin architecture emerges during zygotic genome activation independent of transcription. Cell 169, 216–228 (2017). This study is the first to examine 3D genome architecture throughout embryo development in any organism, revealing that chromatin architecture formation in D. melanogaster coincides with ZGA, although it is not dependent on zygotic transcription.
Ogiyama, Y., Schuettengruber, B., Papadopoulos, G. L., Chang, J.-M. & Cavalli, G. Polycomb-dependent chromatin looping contributes to gene silencing during Drosophila development. Mol. Cell 71, 73–88 (2018).
Niu, L. et al. Three-dimensional folding dynamics of the Xenopus tropicalis genome. Nat. Genet. 53, 1075–1087 (2021).
Chen, X. et al. Key role for CTCF in establishing chromatin structure in human embryos. Nature 576, 306–310 (2019).
Wike, C. L. et al. Chromatin architecture transitions from zebrafish sperm through early embryogenesis. Genome Res. 31, 981–994 (2021).
Kaaij, L. J. T., van der Weide, R. H., Ketting, R. F. & de Wit, E. Systemic loss and gain of chromatin architecture throughout zebrafish development. Cell Rep. 24, 1–10 (2018).
Laue, K., Rajshekar, S., Courtney, A. J., Lewis, Z. A. & Goll, M. G. The maternal to zygotic transition regulates genome-wide heterochromatin establishment in the zebrafish embryo. Nat. Commun. 10, 1551 (2019).
Acemel, R. D., Maeso, I. & Gómez‐Skarmeta, J. L. Topologically associated domains: a successful scaffold for the evolution of gene regulation in animals. WIREs Dev. Biol. 6, e265 (2017).
Merkenschlager, M. & Nora, E. P. CTCF and cohesin in genome folding and transcriptional gene regulation. Annu. Rev. Genom. Hum. Genet. 17, 1–27 (2015).
Nègre, N. et al. A comprehensive map of insulator elements for the Drosophila fenome. PLoS Genet. 6, e1000814 (2010).
Lupiáñez, D. G., Spielmann, M. & Mundlos, S. Breaking TADs: how alterations of chromatin domains result in disease. Trends Genet. 32, 225–237 (2016).
Zhang, K. et al. Analysis of genome architecture during SCNT reveals a role of cohesin in impeding minor ZGA. Mol. Cell 79, 234–250 (2020).
Olbrich, T. et al. CTCF is a barrier for 2C-like reprogramming. Nat. Commun. 12, 4856 (2021).
Gao, T. et al. Nuclear reprogramming: the strategy used in normal development is also used in somatic cell nuclear transfer and parthenogenesis. Cell Res. 17, 135–150 (2007).
Sun, F. et al. Nuclear reprogramming: the zygotic transcription program is established through an “erase-and-rebuild” strategy. Cell Res. 17, 117–134 (2007).
Zhu, Y. et al. Relaxed 3D genome conformation facilitates the pluripotent to totipotent-like state transition in embryonic stem cells. Nucleic Acids Res. 49, 12167–12177 (2021).
Espinola, S. M. et al. cis-Regulatory chromatin loops arise before TADs and gene activation, and are independent of cell fate during early Drosophila development. Nat. Genet. 53, 477–486 (2021).
Batut, P. J. et al. Genome organization controls transcriptional dynamics during development. Science 375, 566–570 (2022).
Ing-Simmons, E. et al. Independence of chromatin conformation and gene regulation during Drosophila dorsoventral patterning. Nat. Genet. 53, 487–499 (2021).
Ghavi-Helm, Y. et al. Highly rearranged chromosomes reveal uncoupling between genome topology and gene expression. Nat. Genet. 51, 1272–1282 (2019).
Despang, A. et al. Functional dissection of the Sox9–Kcnj2 locus identifies nonessential and instructive roles of TAD architecture. Nat. Genet. 51, 1263–1271 (2019).
Ou, H. D. et al. ChromEMT: visualizing 3D chromatin structure and compaction in interphase and mitotic cells. Science 357, eaag0025 (2017).
Mazzocca, M., Fillot, T., Loffreda, A., Gnani, D. & Mazza, D. The needle and the haystack: single molecule tracking to probe the transcription factor search in eukaryotes. Biochem. Soc. Trans. 49, 1121–1132 (2021).
Kuznetsova, K. et al. Nanog organizes transcription bodies. Curr. Biol. 33, 164–173.e5 (2023).
Sabari, B. R. et al. Coactivator condensation at super-enhancers links phase separation and gene control. Science 361, eaar3958 (2018).
Nair, S. J. et al. Phase separation of ligand-activated enhancers licenses cooperative chromosomal enhancer assembly. Nat. Struct. Mol. Biol. 26, 193–203 (2019).
Chong, S. et al. Imaging dynamic and selective low-complexity domain interactions that control gene transcription. Science 361, eaar2555 (2018).
Boija, A. et al. Transcription factors activate genes through the phase-separation capacity of their activation domains. Cell 175, 1842–1855 (2018).
Mir, M. et al. Dynamic multifactor hubs interact transiently with sites of active transcription in Drosophila embryos. eLife 7, e40497 (2018).
Dufourt, J. et al. Temporal control of gene expression by the pioneer factor Zelda through transient interactions in hubs. Nat. Commun. 9, 5194 (2018).
Gibson, B. A. et al. Organization of chromatin by intrinsic and regulated phase separation. Cell 179, 470–484 (2019).
Stasevich, T. J. et al. Regulation of RNA polymerase II activation by histone acetylation in single living cells. Nature 516, 272–275 (2014).
Narita, T. et al. Enhancers are activated by p300/CBP activity-dependent PIC assembly, RNAPII recruitment, and pause release. Mol. Cell 81, 2166–2182 (2021).
Schoenfelder, S. & Fraser, P. Long-range enhancer–promoter contacts in gene expression control. Nat. Rev. Genet. 20, 437–455 (2019).
Bartman, C. R., Hsu, S. C., Hsiung, C. C.-S., Raj, A. & Blobel, G. A. Enhancer regulation of transcriptional bursting parameters revealed by forced chromatin looping. Mol. Cell 62, 237–247 (2016).
Benabdallah, N. S. et al. Decreased enhancer–promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484 (2019).
Chen, H. et al. Dynamic interplay between enhancer–promoter topology and gene activity. Nat. Genet. 50, 1296–1303 (2018).
Alexander, J. M. et al. Live-cell imaging reveals enhancer-dependent Sox2 transcription in the absence of enhancer proximity. eLife 8, e41769 (2019).
Cho, W.-K. et al. Mediator and RNA polymerase II clusters associate in transcription-dependent condensates. Science 361, 412–415 (2018).
Hilbert, L. et al. Transcription organizes euchromatin via microphase separation. Nat. Commun. 12, 1360 (2021). This study demonstrates that nascent mRNA in early zebrafish embryos undergoing ZGA can displace chromatin, revealing how transcription influences chromatin organization.
Henninger, J. E. et al. RNA-mediated feedback control of transcriptional condensates. Cell 184, 207–225 (2021).
Edgar, B. A. & Datar, S. A. Zygotic degradation of two maternal Cdc25 mRNAs terminates Drosophila’s early cell cycle program. Genes Dev. 10, 1966–1977 (1996).
Shimuta, K. et al. Chk1 is activated transiently and targets Cdc25A for degradation at the Xenopus midblastula transition. EMBO J. 21, 3694–3703 (2002).
Dalle Nogare, D. E., Pauerstein, P. T. & Lane, M. E. G2 acquisition by transcription-independent mechanism at the zebrafish midblastula transition. Dev. Biol. 326, 131–142 (2009).
Farrell, J. A., Shermoen, A. W., Yuan, K. & O’Farrell, P. H. Embryonic onset of late replication requires Cdc25 down-regulation. Genes Dev. 26, 714–725 (2012).
Collart, C., Smith, J. C. & Zegerman, P. Chk1 inhibition of the replication factor Drf1 guarantees cell-cycle elongation at the Xenopus laevis mid-blastula transition. Dev. Cell 42, 82–96 (2017).
Zhang, M., Kothari, P., Mullins, M. & Lampson, M. A. Regulation of zygotic genome activation and DNA damage checkpoint acquisition at the mid-blastula transition. Cell Cycle 13, 3828–3838 (2014).
Farrell, J. A. & O’Farrell, P. H. Mechanism and regulation of Cdc25/twine protein destruction in embryonic cell-cycle remodeling. Curr. Biol. 23, 118–126 (2013).
Blythe, S. A. & Wieschaus, E. F. Zygotic genome activation triggers the DNA replication checkpoint at the midblastula transition. Cell 160, 1169–1181 (2015).
Collart, C., Allen, G. E., Bradshaw, C. R., Smith, J. C. & Zegerman, P. Titration of four replication factors is essential for the Xenopus laevis midblastula transition. Science 341, 893–896 (2013).
Murphy, C. M. & Michael, W. M. Control of DNA replication by the nucleus/cytoplasm ratio in Xenopus. J. Biol. Chem. 288, 29382–29393 (2013).
Joseph, S. R. et al. Competition between histone and transcription factor binding regulates the onset of transcription in zebrafish embryos. eLife 6, 1328 (2017).
Amodeo, A. A., Jukam, D., Straight, A. F. & Skotheim, J. M. Histone titration against the genome sets the DNA-to-cytoplasm threshold for the Xenopus midblastula transition. Proc. Natl Acad. Sci. USA 112, E1086–E1095 (2015).
Chari, S., Wilky, H., Govindan, J. & Amodeo, A. A. Histone concentration regulates the cell cycle and transcription in early development. Development 146, dev177402 (2019).
Shindo, Y. & Amodeo, A. A. Excess histone H3 is a competitive Chk1 inhibitor that controls cell-cycle remodeling in the early Drosophila embryo. Curr. Biol. 31, 2633–2642 (2021).
Almouzni, G. & Wolffe, A. P. Constraints on transcriptional activator function contribute to transcriptional quiescence during early Xenopus embryogenesis. EMBO J. 14, 1752–1765 (1995).
Jevtić, P. & Levy, D. L. Both nuclear size and DNA amount contribute to midblastula transition timing in Xenopus laevis. Sci. Rep. 7, 7908 (2017).
Chen, H., Einstein, L. C., Little, S. C. & Good, M. C. Spatiotemporal patterning of zygotic genome activation in a model vertebrate embryo. Dev. Cell 49, 852–866 (2019).
Jukam, D., Kapoor, R. R., Straight, A. F. & Skotheim, J. M. The DNA-to-cytoplasm ratio broadly activates zygotic gene expression in Xenopus. Curr. Biol. 31, 4269–4281 (2021).
Syed, S., Wilky, H., Raimundo, J., Lim, B. & Amodeo, A. A. The nuclear to cytoplasmic ratio directly regulates zygotic transcription in Drosophila through multiple modalities. Proc. Natl Acad. Sci. USA 118, e2010210118 (2021).
Newport, J. & Kirschner, M. A major developmental transition in early Xenopus embryos: I. characterization and timing of cellular changes at the midblastula stage. Cell 30, 675–686 (1982).
Lu, X., Li, J. M., Elemento, O., Tavazoie, S. & Wieschaus, E. F. Coupling of zygotic transcription to mitotic control at the Drosophila mid-blastula transition. Development 136, 2101–2110 (2009).
Balachandra, S., Sarkar, S. & Amodeo, A. A. The nuclear-to-cytoplasmic ratio: coupling DNA content to cell size, cell cycle, and biosynthetic capacity. Annu. Rev. Genet. 56, 165–185 (2022).
Edgar, B. A., Kiehle, C. P. & Schubiger, G. Cell cycle control by the nucleo-cytoplasmic ratio in early Drosophila development. Cell 44, 365–372 (1986).
Chen, H. & Good, M. C. Nascent transcriptome reveals orchestration of zygotic genome activation in early embryogenesis. Curr. Biol. 32, 4314–4324 (2022).
Edgar, B. A. & Schubiger, G. Parameters controlling transcriptional activation during early Drosophila development. Cell 44, 871–877 (1986).
Strong, I. J. T., Lei, X., Chen, F., Yuan, K. & O’Farrell, P. H. Interphase-arrested Drosophila embryos activate zygotic gene expression and initiate mid-blastula transition events at a low nuclear–cytoplasmic ratio. PLoS Biol. 18, e3000891 (2020).
Blythe, S. A. & Wieschaus, E. F. Establishment and maintenance of heritable chromatin structure during early Drosophila embryogenesis. eLife 5, e20148 (2016).
Veenstra, G. J. C., Destrée, O. H. J. & Wolffe, A. P. Translation of maternal TATA-binding protein mRNA potentiates basal but not activated transcription in Xenopus embryos at the midblastula transition. Mol. Cell. Biol. 19, 7972–7982 (1999).
Larson, E. D. et al. Premature translation of the Drosophila zygotic genome activator Zelda is not sufficient to precociously activate gene expression. G3 12, jkac159 (2022).
Rother, F. et al. Importin α7 is essential for zygotic genome activation and early mouse development. PLoS ONE 6, e18310 (2011).
Forbes Beadle, L. et al. Combined modelling of mRNA decay dynamics and single-molecule imaging in the Drosophila embryo uncovers a role for P-bodies in 5′ to 3′ degradation. PLOS Biol. 21, e3001956 (2023).
Riemondy, K., Henriksen, J. C. & Rissland, O. S. Intron dynamics reveal principles of gene regulation during the maternal-to-zygotic transition. RNA 29, rna.079168.122 (2023).
Gentsch, G. E., Owens, N. D. L. & Smith, J. C. The spatiotemporal control of zygotic genome activation. iScience 16, 485–498 (2019).
Holler, K. et al. Spatio-temporal mRNA tracking in the early zebrafish embryo. Nat. Commun. 12, 3358 (2021).
Boettiger, A. N. & Levine, M. Synchronous and stochastic patterns of gene activation in the Drosophila embryo. Science 325, 471–473 (2009).
Stapel, L. C., Zechner, C. & Vastenhouw, N. L. Uniform gene expression in embryos is achieved by temporal averaging of transcription noise. Genes Dev. 31, 1635–1640 (2017).
Little, S. C., Tikhonov, M. & Gregor, T. Precise developmental gene expression arises from globally stochastic transcriptional activity. Cell 154, 789–800 (2013).
Artieri, C. G. & Fraser, H. B. Transcript length mediates developmental timing of gene expression across Drosophila. Mol. Biol. Evol. 31, 2879–2889 (2014).
Falco, G. et al. Zscan4: a novel gene expressed exclusively in late 2-cell embryos and embryonic stem cells. Dev. Biol. 307, 539–550 (2007).
Srinivasan, R. et al. Zscan4 binds nucleosomal microsatellite DNA and protects mouse two-cell embryos from DNA damage. Sci. Adv. 6, eaaz9115 (2020).
Lécuyer, E. et al. Global analysis of mRNA localization reveals a prominent role in organizing cellular architecture and function. Cell 131, 174–187 (2007).
Kigami, D., Minami, N., Takayama, H. & Imai, H. MuERV-L is one of the earliest transcribed genes in mouse one-cell embryos1. Biol. Reprod. 68, 651–654 (2003).
Jachowicz, J. W. et al. LINE-1 activation after fertilization regulates global chromatin accessibility in the early mouse embryo. Nat. Genet. 49, 1502–1510 (2017).
Percharde, M. et al. A LINE1–nucleolin partnership regulates early development and ESC identity. Cell 174, 391–405 (2018).
Li, X. et al. LINE-1 transcription activates long-range gene expression. Nat. Genet. 56, 1494–1502 (2024).
Sakashita, A. et al. Transcription of MERVL retrotransposons is required for preimplantation embryo development. Nat. Genet. 55, 484–495 (2023).
Ge, S. X. Exploratory bioinformatics investigation reveals importance of “junk” DNA in early embryo development. BMC Genom. 18, 200 (2017).
Yang, J., Cook, L. & Chen, Z. Systematic evaluation of retroviral LTRs as cis-regulatory elements in mouse embryos. Cell Rep. 43, 113775 (2024).
Macfarlan, T. S. et al. Endogenous retroviruses and neighboring genes are coordinately repressed by LSD1/KDM1A. Genes Dev. 25, 594–607 (2011).
Vega-Sendino, M. et al. The homeobox transcription factor DUXBL controls exit from totipotency. Nat. Genet. 56, 697–709 (2024).
Asimi, V. et al. Hijacking of transcriptional condensates by endogenous retroviruses. Nat. Genet. 54, 1238–1247 (2022).
Meng, F. W., Murphy, K. E., Makowski, C. E., Delatte, B. & Murphy, P. J. Competition for H2A.Z underlies the developmental impacts of repetitive element de-repression. Development 150, dev202338 (2023).
Ugolini, M. et al. Transcription bodies regulate gene expression by sequestering CDK9. Nat. Cell Biol. 26, 604–612 (2024).
Sugie, K. et al. Expression of Dux family genes in early preimplantation embryos. Sci. Rep. 10, 19396 (2020).
Li, F. et al. mRNA isoform switches during mouse zygotic genome activation. Cell Prolif. 57, e13655 (2024).
Nepal, C. et al. Dynamic regulation of the transcription initiation landscape at single nucleotide resolution during vertebrate embryogenesis. Genome Res. 23, 1938–1950 (2013).
Atallah, J. & Lott, S. E. Evolution of maternal and zygotic mRNA complements in the early Drosophila embryo. PLoS Genet. 14, e1007838 (2018).
Kim, H. H.-S. & Lakadamyali, M. Microscopy methods to visualize nuclear organization in biomechanical studies. Curr. Opin. Biomed. Eng. 30, 100528 (2024).
Treen, N., Heist, T., Wang, W. & Levine, M. Depletion of maternal cyclin B3 contributes to zygotic genome activation in the ciona embryo. Curr. Biol. 28, 1150–1156 (2018).
Calvo, L., Birgaoanu, M., Pettini, T., Ronshaugen, M. & Griffiths-Jones, S. The embryonic transcriptome of Parhyale hawaiensis reveals different dynamics of microRNAs and mRNAs during the maternal–zygotic transition. Sci. Rep. 12, 174 (2022).
Fukushima, H. S., Takeda, H. & Nakamura, R. Incomplete erasure of histone marks during epigenetic reprogramming in medaka early development. Genome Res. 33, 572–586 (2023).
Wei, J. et al. Temporospatial hierarchy and allele-specific expression of zygotic genome activation revealed by distant interspecific urochordate hybrids. Nat. Commun. 15, 2395 (2024).
Halstead, M. M., Ma, X., Zhou, C., Schultz, R. M. & Ross, P. J. Chromatin remodeling in bovine embryos indicates species-specific regulation of genome activation. Nat. Commun. 11, 4654 (2020).
Zhou, C., Halstead, M. M., Bonnet‐Garnier, A., Schultz, R. M. & Ross, P. J. Histone remodeling reflects conserved mechanisms of bovine and human preimplantation development. EMBO Rep. 24, e55726 (2023).
Phelps, W. A. et al. Hybridization led to a rewired pluripotency network in the allotetraploid Xenopus laevis. eLife 12, e83952 (2023).
Avilés-Pagán, E. E. & Orr-Weaver, T. L. Activating embryonic development in Drosophila. Semin. Cell Dev. Biol. 84, 100–110 (2018).
Schvartzman, J. M., Thompson, C. B. & Finley, L. W. S. Metabolic regulation of chromatin modifications and gene expression. J. Cell Biol. 217, 2247–2259 (2018).
Nagaraj, R. et al. Nuclear localization of mitochondrial TCA cycle enzymes as a critical step in mammalian zygotic genome activation. Cell 168, 210–223.e11 (2017).
Li, W. et al. Nuclear localization of mitochondrial TCA cycle enzymes modulates pluripotency via histone acetylation. Nat. Commun. 13, 7414 (2022).
Li, J. et al. Lactate regulates major zygotic genome activation by H3K18 lactylation in mammals. Natl Sci. Rev. 11, nwad295 (2023).
Gerber, A. P., Luschnig, S., Krasnow, M. A., Brown, P. O. & Herschlag, D. Genome-wide identification of mRNAs associated with the translational regulator PUMILIO in Drosophila melanogaster. Proc. Natl Acad. Sci. USA 103, 4487–4492 (2006).
Zhao, L.-W. et al. Nuclear poly(A) binding protein 1 (PABPN1) mediates zygotic genome activation-dependent maternal mRNA clearance during mouse early embryonic development. Nucleic Acids Res. 50, 458–472 (2021).
Zhang, J. et al. The role of maternal VegT in establishing the primary germ layers in Xenopus embryos. Cell 94, 515–524 (1998).
Pritchard, D. K. & Schubiger, G. Activation of transcription in Drosophila embryos is a gradual process mediated by the nucleocytoplasmic ratio. Genes Dev. 10, 1131–1142 (1996).
Ali-Murthy, Z., Lott, S. E., Eisen, M. B. & Kornberg, T. B. An essential role for zygotic expression in the pre-cellular Drosophila embryo. PLoS Genet. 9, e1003428 (2013).
Asami, M. et al. Human embryonic genome activation initiates at the one-cell stage. Cell Stem Cell 29, 209–216 (2022).
Kane, D. A. & Kimmel, C. B. The zebrafish midblastula transition. Development 119, 447–456 (1993).
Braude, P., Bolton, V. & Moore, S. Human gene expression first occurs between the four- and eight-cell stages of preimplantation development. Nature 332, 459–461 (1988).
Jukam, D., Shariati, S. A. M. & Skotheim, J. M. Zygotic genome activation in vertebrates. Dev. Cell 42, 316–332 (2017).
O’Farrell, P. H. Growing an embryo from a single cell: a hurdle in animal life. Cold Spring Harb. Perspect. Biol. 7, a019042 (2015).
Foe, V. E. & Alberts, B. M. Studies of nuclear and cytoplasmic behaviour during the five mitotic cycles that precede gastrulation in Drosophila embryogenesis. J. Cell Sci. 61, 31–70 (1983).
Niakan, K. K., Han, J., Pedersen, R. A., Simon, C. & Pera, R. A. R. Human pre-implantation embryo development. Development 139, 829–841 (2012).
Aiken, C. E. M., Swoboda, P. P. L., Skepper, J. N. & Johnson, M. H. The direct measurement of embryogenic volume and nucleo-cytoplasmic ratio during mouse pre-implantation development. Reproduction 128, 527–535 (2004).
Seydoux, G. et al. Repression of gene expression in the embryonic germ lineage of C. elegans. Nature 382, 713–716 (1996).
Mishima, Y. et al. Differential regulation of germline mRNAs in soma and germ cells by zebrafish miR-430. Curr. Biol. 16, 2135–2142 (2006).
Kedde, M. et al. RNA-binding protein dnd1 inhibits microRNA access to target mRNA. Cell 131, 1273–1286 (2007).
Siddiqui, N. U. et al. Genome-wide analysis of the maternal-to-zygotic transition in Drosophila primordial germ cells. Genome Biol. 13, R11 (2012).
Kane, D. A. et al. The zebrafish epiboly mutants. Development 123, 47–55 (1996).
Acknowledgements
The authors thank members of the Giraldez laboratory for critical feedback, particularly D. Musaev, S. Krishna, F. Sievers, H. Lee, L. Miao, E. Strayer and G. Jaschek. This work was funded by the Jane Coffin Childs Foundation postdoctoral fellowship #61-1730 (to M.L.K.), the Human Frontiers postdoctoral fellowship LT0073/2022-L and EMBO long-term postdoctoral fellowship ALTF #794-2021 (to C.H.), and National Institutes of Health (NIH) grants R01 HD100035 and R35 GM122580 (to A.J.G.).
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M.L.K. and C.H. researched the literature. M.L.K., C.H. and A.J.G. contributed substantially to discussion of the content, wrote the article and reviewed and/or edited the manuscript before submission.
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Glossary
- CCR4–NOT complex
-
During maternal-to-zygotic transition (MZT), this multiprotein complex plays a critical role in regulating gene expression by controlling mRNA deadenylation.
- Cleavage cycles
-
The series of rapid mitotic cell divisions that occur in the early embryo following fertilization, essential for increasing cell numbers in the embryo while maintaining a constant overall size, except for Drosophila spp. where cleavage cycles occur in a syncytium resulting in a growing number of nuclei in a shared cytoplasm.
- Deadenylation
-
The process by which the poly(A) tail of an mRNA molecule is shortened or removed by deadenylating enzymes, which regulates the stability and lifespan of the mRNA molecule.
- Erase and rewrite
-
A developmental strategy that involves the removal (erase) of maternal signatures to a naive state, followed by the establishment (rewrite) of zygotic signatures.
- Histone acetylation
-
Acetyl groups on histone tails that modify the functional properties of DNA, added by histone acetyltransferases (HATs) and removed by histone deacetylases (HDACs).
- Maternal decay
-
(M-decay). Refers to the degradation of maternally deposited mRNAs before zygotic genome activation (ZGA) or independent of zygotically produced factors.
- Maternal-to-zygotic transition
-
(MZT). The transition period during embryogenesis when control of embryonic development transitions from maternal factors to zygotic factors.
- Mid-blastula transition
-
(MBT). A transition phase in embryonic development, characterized by lengthening cell cycles, acquisition of cell motility and, in Drosophila spp., cellularization.
- Nuclear-to-cytoplasmic volume ratio
-
(N:C ratio). The relative nuclear-to-cytoplasmic ratio within a cell. During embryogenesis (zygote to gastrulation) the size of the embryo does not change; cell sizes are halved with every cleavage cycle.
- ORF-mediated decay
-
A decay pathway driven by the protein Upf1, whereby the translation status of the main open reading frame (ORF), affected by upstream ORFs and ORF length, influences decay dynamics.
- Pioneer transcription factors
-
(PFs). Specialized transcription factors (TFs) with the unique ability to bind to condensed or inaccessible regions of chromatin, promoting chromatin opening and making these regions accessible for other regulatory proteins.
- Protamines
-
Small proteins that replace histones in sperm (except in zebrafish) and help to compact the sperm genome.
- Re-adenylation
-
(Cytoplasmic polyadenylation). Lengthening of poly(A) tails by specialized poly(A) polymerases in the cytoplasm, which leads to translational upregulation.
- Totipotent
-
The ability of a cell to give rise to all cell types in an organism, including both embryonic and extra-embryonic tissues.
- Zygote
-
Describes a fertilized egg and the earliest developmental stage of a multicellular organism.
- Zygotic decay
-
(Z-decay). Refers to the clearance of maternally deposited mRNAs dependent on zygotically produced factors.
- Zygotic genome activation
-
(ZGA). The process during embryogenesis where the zygotic genome becomes transcriptionally active.
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Kojima, M.L., Hoppe, C. & Giraldez, A.J. The maternal-to-zygotic transition: reprogramming of the cytoplasm and nucleus. Nat Rev Genet 26, 245–267 (2025). https://doi.org/10.1038/s41576-024-00792-0
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DOI: https://doi.org/10.1038/s41576-024-00792-0
