Ribosome Heterogeneity in Zebrafish Germ and Soma provides insight into Gene Expression during Development and Disease by Arish N. Shah B. S., Bioengineering: Bioinformatics University of California, San Diego submitted to the Department of Biology in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY at the MASSACHUSETTS INSTITUTE OF TECHNOLOGY June 2024 © 2024 Arish N. Shah. All rights reserved. The author hereby grants to MIT a nonexclusive, worldwide, irrevocable, royalty-free license to exercise any and all rights under copyright, including to reproduce, preserve, distribute, and publicly display copies of the thesis, or release the thesis under an open-access license. Signature of Author ………………………………………………….…………………………… Arish N. Shah Department of Biology April 30, 2024 Certified by ..……………………………………………………………………………………… Eliezer Calo Associate Professor of Biology Department of Biology Thesis supervisor Accepted by ……...………………………………………………………………………………. Mary Gehring Professor of Biology Member, Whitehead Institute Director, Biology Graduate Committee 2 Ribosome Heterogeneity in Zebrafish Germ and Soma provides insight into Gene Expression during Development and Disease by Arish N. Shah Submitted to the Department of Biology on March 1, 2024 in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy ABSTRACT Ribosomes are the micromachines which produce the materials composing the molecular-cellular complexity of organisms. Regulation of gene expression by the translational machinery provides a layer of control over the timing, location, and amount of any given gene product and its associated functions. Protein synthesis during vertebrate development is driven by a common pool of ribosomes of two distinct origins: subunits synthesized by the mother during oogenesis and stored in the egg, and subunits produced after fertilization by the embryo. In most organisms, these two are the same. Recently, cell type-specific expression of two zebrafish ribosomal DNA (rDNA) genes was identified (Locati et al. 2017b). One rDNA variant located on chromosome 4 was found to be specifically expressed in eggs (aka maternal type), while expression of another rDNA variant located on chromosome 5 was specific to somatic cells (aka somatic type). Critically, these rRNAs vary in about 15% of their sequence. Ribosome structural heterogeneity is an appealing occurrence as it may uncover ribosome-specific functionality shaping translational control seen in development. Since variation observed between the zebrafish rRNA types is substantial, it has the potential to affect ribosome biogenesis, structure, and function at different levels. We use this system to investigate the possibility of germ cell-specific ribosome functionalization. This thesis contains research assessing two rDNA gene variants, the transcribed rRNAs, and the two sets of ribosome subunits they compose. We separately characterize maternal and somatic ribosomes using 6 - 120 hours post-fertilization (hpf) animals. Cryo- EM structure maps of each show compositionally different, yet structurally similar assemblies. Using transgenic labeling of maternal and somatic subunits, we confirm intersubunit compatibility forms cognate and hybrid monosomes. We show primordial germ cells transcribe the somatic rDNA gene upon genome activation, and, unexpectedly, shift transcription to the maternal rDNA gene at 72 hpf. Lastly, we demonstrate maternal ribosome-enriched translation of germ cell-specific mRNA in vivo. Zebrafish germ cells maintain a majority of maternal rDNA gene products at all measured times. Our findings solidify the chromosome 4 rDNA variant as a germ cell-specific rDNA gene. This work clarifies the structural, molecular, and cellular consequences of cell type- specific rRNA expression on ribosome heterogeneity. We indicate a germ cell-specific ribosome functionalization and frame the zebrafish dual rDNA gene variant system for future inquiry regarding ribosome biogenesis, translation control, and germ cell development. Thesis Supervisor: Eliezer Calo Title: Associate Professor of Biology 3 Acknowledgements Thank you to Dr. Eliezer Calo for taking me into the lab during my first year. No one could have expected the scientific trajectory that we took over the next 6 years, and this lack of expectation has led to successful scientific collaborations around the world. Thank you to the members of the Calo Lab for endless discussions about ribosome biology – especially to Dr. Nima Jaberi and Dr. Byron Lee. Thank you to my committee members, Dr. David Bartel and Dr. Peter Reddien for unending support during the trials and tribulations of this PhD. None of this work would have been done without the expert zebrafish husbandry by Dr. Adam Amsterdam. Thank you to many labs for sharing reagents, equipment, expertise, and mentoring – especially Dr. Iain Cheeseman, Dr. Mary Gehring, Dr. Chris Burge, Dr. Gene-Wei Li, and Dr. Adam Martin. A special thank you to Dr. Amy Keating for indispensable advice throughout the years. Thank you to various members of the MIT Biology community for many conversations about translation control, repetitive elements, developmental biology, and ribosome biogenesis – Dr. Jennifer Chu, Dr. Michael McGurk, Dr. David McWatters, Dr. Lidya Herzel, and Dr. Amelie Raz. Thank you to Dr. Trey Ideker, my undergraduate mentor, and Dr. Jean Lozach, my manager at Illumina, who showed me the power of bioinformatics. Thank you to Dr. Cecilia Moens for training me in zebrafish techniques and expanding my interests to developmental biology prior to arriving at MIT Biology. Thank you to Dr. Leonard Zon for stopping at my poster during a Society for Developmental Biology conference and recommending I work with members of his lab to profile zebrafish translation. Thank you to Dr. Seth Corey for chatting at the RiboClub conference, and establishing a fruitful collaboration with Dr. Usua Oyarbide. Thank you to Dr. Andrea Pauli for initiating a collaboration among Dr. Frieda Leesch and myself. The possibility of working on the same project as someone else can be terrifying and delightful, I’m grateful for the teamwork we were able to achieve. Thank you to my parents, Nitin and Pratima Shah, for supporting me in the best ways they know how, and to my aunts and uncles, Arvind and Sharda Shah, Rajni and Bindu Shah, and Hitendra and Bina Shah. As well as to the rest of my family: Shrey, Listya, Ian, Lucas, Ami, Nick, Ethan, Pinal, Jay, Biran, Cailin, Jiyana, Nitiksha, Harshil, Arav, Rushil, and Kanchi for the unending support along the way. Thank you to Dr. Daniel Ferrer, your scientific and personal reinforcement has been critical for the progress and completion of this thesis as well as my growth as a scientist. 4 Summary of Doctoral Thesis Work Research Publications 1. A dual ribosomal system in the zebrafish soma and germline Shah AN#, Leesch F#, Lorenzo-Orts L, Grundmann LE, Pribitzer C, Novatchkova M, Grishkovskaya I, Haselbach D, Calo E, Pauli A. In preparation. Manuscript is included as Chapter II. #, these authors contributed equally. 2. eIF6 Dosage Alleviates the TP53 Pathway in SBDS Deficient Cells Oyarbide U, Bezzerri V, Staton M, Bonni C, Shah AN, Cippolli M, Calo E, Corey SJ. In preparation. 3. Nucleolus activity-dependent recruitment and condensation by pH sensing Aryan F, Detrés D, Luo CC, Kim SX, Shah AN, Bartusel M, Flynn R, Calo E. Molecular Cell. 2023. 4. SBDSR126T rescues survival of sbds−/− zebrafish independently of Tp53 Oyarbide U, Shah AN, Staton M, Snyderman M, Sapra A, Calo E, Corey SJ. Life Science Alliance. 2023. 5. Loss of sbds in zebrafish leads to neutropenia and pancreas and liver atrophy Oyarbide U, Shah AN, Amaya-Mejia W, Snydermann M, Kell MJ, Allende DS, Calo E, Topczewski J, Corey SJ. Journal of Clinical Investigation Insight. 2020. 6. Calmodulin inhibitors improve erythropoiesis in Diamond-Blackfan anemia Taylor AM, Macari ER, Chan IT, Blair MC, Doulatov S, Vo LT, Raiser DM, Siva K, Basak A, Pirouz M, Shah AN, McGrath K, Humphries JM, Stillman E, Alter BP, Calo E, Gregory RI, Sankaran VG, Flygare J, Ebert BL, Zhou Y, Daley GQ, Zon LI. Science, Translational Medicine. 2020. Pedagogical Experiences o 2020-2024 Instructor and Founder – In Silico Tutoring o 2023 Content Contribution – Brilliant.org o 2020 Teaching Assistant, Cell Biology – MIT course 7.06 o 2018-2019 Teaching Assistant, TSR2 – MIT Office of Minority Education o 2017-2019 Activity Coordinator, Summer Dev Bio Outreach – MIT Biology o 2017 Teaching Assistant, Introductory Biology – MIT course 7.012 Awards o 2020 Teaching Award The Gene Brown – Merck Teaching Award, MIT o 2019 Best Poster Award, Research; 1 of 3 Department of Biology Retreat, MIT o 2019 Best Speaker Award, Research; First Place Northeast Regional Society for Developmental Biology o 2018 Best Poster Award, Research; 1 of 3 RiboClub Intl Conference on Ribosome Synthesis o 2018 Teaching Award Teresa Keng Graduate Teaching Award, MIT o 2018 Graduate Research Fellowship National Science Foundation (NSF GRFP) 5 Table of Contents ABSTRACT .................................................................................................................................................. 2 Acknowledgments ...................................................................................................................................... 3 Summary of Doctoral Thesis Work ........................................................................................................... 3 Table of Contents ....................................................................................................................................... 5 Preface ......................................................................................................................................................... 6 Specific nomenclature, abbreviations, and initialisms .............................................................................. 7 Chapter 0 ..................................................................................................................................................... 8 an abstracted elephant for inquiry ............................................................................................................ 9 Chapter I .................................................................................................................................................... 13 Chapter I preface ................................................................................................................................... 14 Ribosomes are required for gene expression ........................................................................................ 15 Ribosomes manufacture proteins by translating mRNA ........................................................................ 19 Ribosome structures support their functions .......................................................................................... 30 Ribosome biogenesis forms new subunits ............................................................................................. 37 Ribosome heterogeneity arises during biogenesis ................................................................................ 47 Ribosome specificity and the mRNA pool .............................................................................................. 51 Germ cell genomes are where evolution occurs .................................................................................... 56 Germ cell regulation of translation ......................................................................................................... 60 Danio rerio, a specific fish for inquiry ..................................................................................................... 64 References ............................................................................................................................................. 68 Chapter II ................................................................................................................................................... 89 Chapter II preface .................................................................................................................................. 90 Collaboration notes ................................................................................................................................ 91 A dual ribosomal system in the zebrafish soma and germline ............................................................... 93 Abstract .................................................................................................................................................. 94 Introduction ............................................................................................................................................ 95 Results ................................................................................................................................................... 97 Discussion ............................................................................................................................................ 109 Materials and Methods ......................................................................................................................... 111 References ........................................................................................................................................... 119 Supplementary Figures ........................................................................................................................ 124 Supplemental Tables ............................................................................................................................ 136 Acknowledgments ................................................................................................................................ 145 Author Contributions ............................................................................................................................. 146 Chapter III ................................................................................................................................................ 147 Chapter III preface ............................................................................................................................... 147 Summary and Model ............................................................................................................................ 148 Rephrasing the motivating questions ................................................................................................... 153 Hypotheses and Findings ..................................................................................................................... 155 Conclusions .......................................................................................................................................... 163 Note on nomenclature .......................................................................................................................... 164 References ........................................................................................................................................... 167 Note I ........................................................................................................................................................ 170 The blind and the elephant ................................................................................................................... 171 6 Preface Chapter 0 contains a short generalized version of the research question and illustrates, in abstract, the advantage of asking prying questions. Chapter I introduces the ribosome and its associated translational machinery necessary for gene expression in all lifeforms, along with concepts regarding ribosome heterogeneity, especially in germ cells. Chapter II provides a report of our work characterizing two types of zebrafish ribosomes, one specific to germ cells and another specific to somatic cells. Through biochemical, genetic, and molecular experimentation, we find the two are compositionally different, structurally distinct, and functionally compatible. Chapter III summarizes the motivations for researching two zebrafish cell type- specific ribosomes alongside our original hypotheses and our findings. Future research into this peculiar ribosome heterogeneity is likely to uncover rDNA gene dosage control, oocyte cell-specific ribosome biogenesis, and ribosome functional heterogeneity. Possible implications of this work on current research involving translation control and sex chromosomes are also discussed. 7 Specific nomenclature, abbreviations, and initialisms RNA ribonucleic acid rRNA ribosomal RNA mRNA messenger RNA tRNA transfer RNA ncRNA non-coding RNA bp base-pair DNA deoxyribonucleic acid rDNA ribosomal DNA (gene encoding rRNAs) NOR nucleolar organizing region ITS internal transcribed sequence ETS external transcribed sequence SSU small subunit of the ribosome, 40S LSU large subunit of the ribosome, 60S RP ribosome protein eIF eukaryotic initiation factor eEF eukaryotic elongation factor eRF eukaryotic release factor RPS/eS/uS ribosome protein of the small subunit RPL/eL/uL ribosome protein of the large subunit UTR untranslated region PGC primordial germ cell GFP green fluorescent protein IP immunoprecipitation Tg transgenic hpf hours post-fertilization dpf days post-fertilization Å angstrom, 0.1 nanometers S Svedberg unit 8 Chapter 0 Research is formalized curiosity. It is poking and prying with a purpose. It is a seeking that he who wishes may know the cosmic secrets of the world. – Zora Neale Hurston 1942 9 an abstracted elephant for inquiry The generalized question of my thesis research: Why are there two things in this system, when other systems only have one? Which perspectives can be used to query the system? An individual perspective of any system is an incomplete view of the whole. Each person uses their preferred approach to measure the system, and inherently “sees” limited information to answer the question. Each approach defines the which, how, when, and where methods can be used to measure the system; permuting these provides many perspectives of the process. Therefore, considering information from orthogonal measurements illustrates a better picture of the whole. Concepts regarding the benefits of multiple viewpoints and the conditional basis of knowledge are first described around 500 BCE, in the parable of the blind assessing an elephant; for more, see Note I. Here are four relevant perspectives and how each are used to approach this thesis question: A. Are the two different? à Measure contents of each B. Does the system use two? à Measure activity of each C. Does the system need two? à Measure the system after removing one D. If a system can have two, why not three? Who would assess the system? The person who asks A is capable of isolating each thing from the system for separate measurements. The person who asks B is capable of distinguishably measuring activity of each thing within the system. The person who asks C is willing to break the system and measure the ramifications of loss for an answer. The person who asks D is speculating on the system’s method of counting things to provide context to the discussion. Why this question for a Doctorate of Philosophy in Biology? While not as moving as the original French, my translation of Abraham Trembley’s memoires regarding the observations he made of the freshwater hydra provides one answer: She [Nature] must be explained by Nature; not by our own individual perspectives, each too narrow to grasp the totality of information contained within. … She provides us with the ideas that constitute her Infinite Wisdom; and therefore, concepts most worthy for cultivating the mind and heart. That is what we must propose in our Research (Trembley, 1744). Crossing paths with the Infinite Wisdom of Nature has been a demanding, edifying, and fulfilling experience. What benefit is it to probe this question in a biological system? This thesis draws heavy inspiration from one protocol: the developmental time course. For thousands of years, people have been building a model of how the commonplace phenomenon of development occurs by taking snapshots along the way. Separating developing embryos for measurement at successive times of an incubation period, aka developmental time course measurements (Fig. 0.1), were notably conducted by Aristotle with a chick egg over 3-weeks, in which he describes the formation of all major organs 10 from the otherwise inert biological materials of a fertilized egg. Over time our ability to answer complex questions about development has grown; matched by an increased sensitivity in the tools we use to measure the system from genomic, molecular, and cellular perspectives during each snapshot. As Aristotle remarked in 350 BCE, “Wonder remains the beginning of knowledge”, and, at least in my naïve evaluation, there is plenty of wonder yet to be revealed by examining the biological materials composing Life. Figure 0.1 – Experimental abstract Developmentally staged organisms and tissues composed of different cell types are used as input. Samples are lysed, homogenized, and processed to separate their molecular components and measure the degree of separation. Measurements inform models and generate hypotheses regarding molecular function. Which biological system did I use? For my Doctoral thesis work, the zebrafish (Danio rerio) model organism has been the system of choice. It has provided me with a molecularly robust and experimentally tractable means of querying the complex organismal system. This has made grasping at “the totality of information contained within” both possible and enjoyable. While the pursuit of biological knowledge is driven by the puzzling complexities seen in life, it also provides an opportunity to mechanistically understand molecular interactions in organismal health, and disease (Fig. 0.2). Collaborations with basic science researchers and translational clinician-scientists similarly using the zebrafish model for investigation, provide breadth and depth to this thesis work. Where and When can biological systems be assessed? At what scale? The organism is a unit of lifeform composed of cells, micrometer to millimeter scale lipid containers, that compartmentalize a subset of molecules; with each subset collectively functioning at the picometer to nanometer scale. Within in each cell, the genome (DNA) is expressed into RNA and translated into proteins by the ribosome. Together, nucleic acids, proteins, lipids, and sugars assemble the molecular programs materializing the cell in space (Fig. 0.2). In most cases, a multicellular lifeform, like us, begins as a single cell – the fertilized egg – which divides mitotically, over time, constructing the patterned composition of the embryo. The organism contains two populations of cells: the soma, non-reproductive cells which impart function to the body, and the germ, reproductive cells which transmit the genome to the next generation. Through the organism’s life cycle, cells of the germ and soma work together to ensure propagation of the genome. Dynamic manifestations of biologically-encoded functions arise from regulating gene expression programs in both time and space. Integration of small-scale molecular-cellular processes consequently produces the large-scale complexities exhibited by organisms: composition, metabolism, growth, adaptation, response, reproduction, and homeostasis. 11 Figure 0.2 – Building up and breaking down an organism. Building up (left to right): genes are expressed via the Central Dogma: DNAàRNAàProtein (Fig 1.1), where genomic information encoded in DNA is first transcribed into a messenger RNA (mRNA) that can be then translated into protein by the ribosome. These molecules interact to produce cellular programs and organismal functions. Distinct expression of genes (magenta, yellow, and green), shown as DNA sequences (boxes), RNA transcripts (lines), and proteins (filled circles) establish cell types (grey, blue, and orange squares) within the organism. Breaking down (right to left): an organism is a patterned composition of cells with different identities and counts. Organism-level functions like growth, homeostasis, and reproduction are realized from interactions among specific cellular programs over time and space. Cells enact diverse programs by modulating the composition of its gene products, aka gene expression. DNA, RNA, and Protein measurements can therefore assess gene expression in the organism, in specific tissues, or in single cells. What is the ribosome? Ribosomes are ribonucleoprotein machines found in all living systems (25-30 nanometer diameter in eukaryotes). The machinery, its biogenesis, and regulation noticeably appear in the set of genes shared across all known members of the tree of life. What is the ribosome’s function? The function of a ribosome is to assemble more ribosomes. To achieve this, ribosomes make ribosomal proteins, ribosomal biogenesis enzymes, ribosomal accessory factors, and RNA polymerases to transcribe ribosomal genes into ribosomal RNAs. Incidentally, ribosomes also function to assemble the molecular-cellular complexities seen in all known organisms. The ribosome simultaneously “reads” information encoded in ribonucleic acid polymers (RNA), “translates” the sequence, and assembles a protein by sequentially arranging amino acids in the order dictated by the message. Reading information in nucleic acid polymers and the unidirectional translation of the message into amino acid polymers is so fundamental to all known lifeforms, that biologists have termed this process “The Central Dogma” (Fig. 1.1). In the last 4 billion years, all biologically encoded proteins produced on Earth were fabricated by a ribosome. If cells need ribosomes to make ribosomes, where do cells get their first ribosomes? Like many things in life, the mother provides this necessary resource. A reduction of 50% of the translational output by ribosomes is enough to stop the cell’s ability to proliferate. Due to an essential role in gene expression, ribosome concentration and ribosome biogenesis are tightly regulated processes. An individual somatic cell can contain 1 x 106 (1,000,000) ribosomes. To maintain this ribosome concentration over many cellular divisions, the primary metabolism of a proliferative cell is ribosome biogenesis. A typical somatic cell grown in nutrient-rich conditions may require 24 hours to double its supply of ribosomes – thus allowing for one mother cell to duplicate into two daughter cells. 12 Unlike somatic cells, zebrafish eggs can contain nearly 3 x 1012 (3,000,000,000,000) ribosomes – a maternally-inherited stockpile supplying a newly fertilized single-cell zygote with ribosomes for embryogenesis. About 8 hours post-fertilization, the zygote will begin to add to the total number by generating its own ribosomes – thus allowing for thousands of cells to be born, patterned, and specified within 24 hours. The specific question of my thesis research: In zebrafish, ribosomes generated in the soma contain a different sequence and structure than maternally-deposited ribosomes found in the egg. Neither ongoing searches of metazoan literature regarding ribosomal gene variation, nor the inventory of known ribosomal gene sequences, nor publicly available ribosomal gene expression patterns include anything similar – possibly indicating this phenomenon as exclusive to zebrafish. Why are there two variant ribosomes in zebrafish, when other animals only have one? How did I pursue this research? I am incredibly grateful to have been given the opportunity to work with high quality collaborators, colleagues, and mentors at MIT and around the world to consider more perspectives regarding the molecular intricacies of this question. A. Are the two different? à Measure contents of each B. Does the system use two? à Measure activity of each C. Does the system need two? à Measure the system after removing one Research using approaches A and B is primarily concerned with measuring composition and translational activities of each ribosome. Our findings are reported in Chapter II. This work, now a collaboration, was independently initiated by myself in Dr. Eliezer Calo’s lab at MIT in Cambridge, and by Frieda Leesch in Dr. Andrea Pauli’s lab at the IMP in Vienna. Research using approach C was conducted collaboratively: with Dr. Alison Taylor in Dr. Leonard Zon’s lab at Boston Children’s Hospital, with Dr. Usua Oyarbide in Dr. Seth Corey’s lab at Cleveland Clinic, and with Fardin Aryan, Diego Detrés, and others within the Calo lab at MIT (see Summary of Doctoral Thesis Work). I worked with multiple researchers at multiple institutions in multiple countries who independently measured the same two zebrafish ribosome variants by employing multiple methods at multiple times in development. Progress was regularly fueled by the unique experiences and scientific insights I encountered while conducting collaborative research. 13 Chapter I Our real teacher has been and still is the embryo, who is, incidentally, the only teacher who is always right. – Viktor Hamburger 14 Chapter I preface Chapter I introduces the ribosome, a molecular machine required by all lifeforms. The chapter begins with a section about the machinery which allows for gene expression, followed by sections regarding mRNA translation, ribosome structures, and ribosome biogenesis. Ribosomes distinct to the zebrafish germ and soma are the subject of interrogation in this Thesis; therefore, the chapter contains sections about germline specification, germ cell translation, and the zebrafish model organism. A short section on a paralog investigation of a testis-specific ribosome heterogeneity found in the fruit fly is included for perspective – a progression of paralog protein publications provides a particularly appropriate case to consider the products of ribosomal genes. Ribosome biogenesis involves the coordinated activities of hundreds of proteins to form the ribosome. Structural and compositional differences in the construction of subunits may lead to molecularly distinct populations; some with added function, some with loss of function. The influence heterogeneous ribosomes may have on molecular-cellular systems is formidable. Two prominent models relating the effects of ribosome heterogeneity on translational control are described. In the literature, few pieces of evidence suggest ribosome functional heterogeneities alter the subset of mRNAs translated by a ribosome, and fewer support ribosome functionalization. The germline affords an additional alluring avenue for research. Ribosome heterogeneities are a source of variation potentially offering germ cells an advantage in natural selection at the cellular level. As embryonic germ cells establish the gametes, their population dynamics can greatly influence inheritance. An advantageous ribosome heterogeneity may provide germ cells with a competitive advantage; giving the genetically encoded heterogeneity a greater chance to be fixed in embryonic germ cell genomes for inheritance into the next generation. Our study, reported in Chapter II, works to characterize two heterogeneous ribosome populations – one is specific to somatic cells; another is specific to germ cells, and thus maternally deposited into the unfertilized egg. The conclusions are summarized in Chapter III and the implications of our findings on heterogeneous ribosomes in the germline are discussed. 15 Ribosomes are required for gene expression A major challenge in Biology is to understand how genes are expressed and regulated in both time and space in order to control cellular functions and organismal development. The sequential steps of gene expression are stated by the Central Dogma (Fig. 1.1), which describes a two-step reaction conducting the unidirectional flow of information experienced by all living systems: DNA à RNA à Protein (Crick 1970). First, gene information encoded in a DNA genome (gDNA), is transcribed into messenger RNA (mRNA), and second, the ribosome, with its associated machinery, simultaneously translates the mRNA into an order of amino acids and polymerizes the sequence to form a protein polypeptide. The specific order of polymerized amino acids composing a protein determine the specific functions of the protein, if made correctly. The ribosome’s catalytic role in gene expression is essential for the growth and survival of all lifeforms. Figure 1.1 – The Central Dogma Schematic cartoon of gene expression, as it follows the central dogma. The yellow gene is transcribed into an mRNA (shown with a yellow cap), and translated, by the ribosome (grey), into a protein (yellow circle). The translation machinery Ribosomes are the primary effectors concerning translation – the second step of gene expression. The enzyme at the core of each ribosome is a ribozyme which catalyzes the peptide bond reaction necessary for synthesizing a polypeptide (Cech 2000; Steitz and Moore 2003). Translation is an energy-intensive process involving a regulated sequence of three steps: initiation, elongation, and termination. The translation machinery contains three molecular components: ribosomes, transfer RNA (tRNA), and ribosome associated factors including translation factors (Archer et al. 2016; Shirokikh et al. 2017). Over various steps of translation, these components will interact to decode an mRNA and translate the resulting message into a nascent polypeptide. Functional impairment of any one of these components has the potential to alter gene expression programming (Mills and Green 2017). Structural impairment to any one of these components may force polypeptide failure, or generate a faulty protein (Tahmasebi et al. 2018), effectively inhibiting expression of the gene. For the sake of brevity and clarity, this Thesis chapter will draw from knowledge regarding the eukaryotic ribosome. 16 Ribosomes The discovery of the ribosome is credited to George Palade and Albert Claude for their work on the structural and functional organization of the cell in the 1950s. The newly discovered protein synthesis machine was initially referred to as the ‘microsome’ until a symposium held in 1958 at the Massachusetts Institute of Technology (MIT). Due to nomenclature difficulties in conversation among researchers, the name ‘ribosome’ was chosen to define the abundant ribonucleoprotein particle of the microsome fraction (Biophysical Society. et al. 1958). In 1974, the works of Palade, Claude, and Christian de Duve earned them a Nobel Prize in Physiology or Medicine. Later in 2009, another Nobel Prize in Chemistry was awarded to Venkatraman “Venki” Ramakrishnan, Thomas A. Steitz, and Ada E. Yonath for “studies of the structure and function of the ribosome”. Ribosomes are multicomponent macromolecular complexes composed of RNA and protein, weighing up to 4.3 megadaltons (in humans) (Fig. 1.2). When comparing ribosomes from various different organisms, a striking increase in molecular length is seen over evolutionary time (Bernier et al. 2018; Fox 2010; Gerbi 1986; Hariharan et al. 2023; Petrov et al. 2014). While structurally expanded on the exterior, the ribosome has maintained its functional core for the last 3.8 billion years. As a consequence of the ribosome’s ancient ancestry and ubiquitous usage, ribosomal DNA (rDNA) genes represent a chronicle of evolution across the domains of life (Petrov et al. 2014). Accordingly, ribosomal sequences can be used to create a phylogenetic tree relating all known organisms (Guo 2018). 60S LSU 40S SSU Figure 1.2 – An assembled human ribosome A surface representation of the 2.9 Å resolution structure of the human ribosome. The large subunit (LSU) rRNA components are shown in dark grey (top). Small subunit (SSU) rRNA components are shown in light grey (bottom). Ribosome proteins (RPs) are shown in white. Scale bar indicates 10 nm. (PDB: 6QZP) Ribosomes are made up of 2 subunits: the large subunit (LSU) and the small subunit (SSU) (Fig. 1.2). The subunits are themselves composed of roughly 80 ribosomal proteins, aka r-proteins (RPs), and 4 ribosomal RNAs (rRNAs) transcribed by RNA polymerase I (RNA Pol I). The SSU, aka 40S subunit, contains the decoding center where the ribosome ensures fidelity of codon-anticodon interactions during translation. The LSU, aka 60S subunit, contains the nascent peptide exit tunnel from where the newly polymerized peptide will emerge. The ribosome catalyzes peptide bonds between amino 17 acids via the enzyme peptidyl transferase; thus, forming the nascent polypeptide. It is also responsible for quality controlling the nascent chain on its exit out of the machine (Brandman and Hegde 2016; Dever and Green 2012; Merrick and Pavitt 2018). Transfer RNAs The translation machinery uses tRNAs to act as adapters between three nucleotide mRNA codons and one of twenty amino acids. tRNAs are transcribed by RNA polymerase III (RNA Pol III). Each tRNA is a single molecule of RNA folded with Watson-Crick interactions to form a cloverleaf structure (Holley et al. 1965). It consists of 4 arms: the D arm which acts to stabilize the tertiary structure of the tRNA, the amino acid acceptor arm with a 5’ CCA 3’ extension which is linked to, or “charged” with, an amino acid by specific aminoacyl tRNA synthetases, the TΨC arm which is involved in interactions with the ribosome, and critically, an anticodon arm that will interact with the codon present on an mRNA via base pairing. Lastly, there is a variable loop that is involved in aminoacyl tRNA synthetase recognition of specific tRNA (Giegé et al. 2012; Kim et al. 1974). Figure 1.3 – Structure of a tRNA Schematic representation of the primary and secondary structures of tRNA with the D, amino acid acceptor, TΨC, and anticodon arms labeled. Adapted from (Lorenz et al. 2017). Translation factors and ribosome associated factors Translation factors are RNA binding proteins (RBPs) classified into 3 categories: initiation, elongation, and release factors; each involved in initiation, elongation, and termination, respectively. Translation factors will either bind to ribosomal RNA or mRNA over various steps of translation. In eukaryotes, there are 18 different initiation factors (eIFs), 4 elongation factors (eEFs) and 2 release factors (eRFs) (Jackson et al. 2010; Safer 1989). Each translation factor plays a distinct role during translation. Proteins that transiently associate with the ribosome but are not specifically translation factors are known as ribosome-associated factors. These proteins compose the ribosome interactome (Reschke et al. 2013; Simsek et al. 2017). To identify ribosome accessory 18 factors, ribosomes can be isolated via immunoprecipitation or via centrifugation through a sucrose gradient or sucrose cushion. Methods for the improved detection of proteins by isobaric labeling and mass spectrometry (TMT-MS) have enabled the identification of many of these proteins and their stoichiometric relationship to the ribosome (Chen et al. 2021). Liquid chromatography tandem mass spectrometry (LC-MS/MS) can then be used to analyze the proteins associated with the translational machinery (Belin et al. 2010; Heiman et al. 2014; Reschke et al. 2013; Rivera et al. 2015; Simsek et al. 2017). Ribosome-associated factors only transiently associate with the ribosome and therefore only transiently alter functions of the translational machinery. Some factors influence the protein synthesis output of the ribosome, such as signal recognition particle (SRP). Associated factors, such as Sec61, may contribute to ribosome subcellular localization when in complex with the machinery (Voorhees et al. 2014), while others, such as poly(A) binding protein (PABP), stabilize interactions between an mRNA and the actively initiating ribosome translating it (Imami et al. 2018). Figure 1.4 – Layout of an mRNA Schematic of an mRNA molecule with its 5’ cap (yellow square), 5’ UTR, ORF (yellow rectangle), 3’ UTR, and poly(A) tail. The ORF begins at the +1 translation initiation codon, AUG, and terminates at a stop codon. mRNA transcripts Nascent pre-mRNAs are transcribed from the DNA genome by RNA polymerase II (RNA Pol II). Mature mRNAs contain 5’ and 3’ untranslated regions (UTRs), a translation initiation site defining an open reading frame (ORF) consisting of successive 3-nucleotide codons, and a termination site. Two more features often used for translation regulation include a 5’ cap and a 3’ poly(A) tail, are added to the mRNA; neither of which are encoded in DNA (Fig. 1.4). The poly(A) tail is added post-transcriptionally through polyadenylation of the 3’ end by poly(A) polymerase. This tail interacts with PABP to ensure the stability of mRNA and association with initiation factors (Sachs and Davis 1989). The same poly(A) tail is also required for the export of mRNA from the nucleus (Curinha et al. 2014). The cap prevents degradation of mRNAs by 5’-3’ exonuclease activity. The cap is an N7- methylated guanine (m7G, or m7Gppp) linked to the first mRNA nucleotide by a 5’-5’ triphosphate bond (Furuichi 2015). The inverted bond of a 5’ m7GpppGpNp 3’ is a characteristic feature of mRNA; translation factors use this handle to recognize an mRNA for the assembly of a translating ribosome. The rate-limiting step for protein synthesis is mRNA recognition by an available ribosome, known as initiation. 19 Ribosomes manufacture proteins by translating mRNA Translation is a sequential process by which a defined polypeptide chain (protein) is synthesized from mRNA. The translational machinery is composed of the small (40S) and large (60S) subunits of the ribosome, translation factors and ribosome-associated factors, as well as charged tRNAs. Ordered base-pairing between triplets of nucleotides in the mRNA sequence (codons) with complementary bases in the tRNAs (anticodons), establishes a sequential translation from the trinucleotide alphabet to the amino acid alphabet. Peptidyl transferase activity at each decoding step of the process ensures a polypeptide is synthesized in the N-terminal to C-terminal direction during translation. Decoding the mRNA message After being expressed from the genome, an mRNA must be processed and, in eukaryotes, exported from the nucleus to engage with various translation factors. Translation is performed by the ribosome, which, together with transfer RNAs (tRNAs), “reads” mRNA transcripts and translates the encoded open-reading frame into protein (Fig. 1.4). The sequence of nucleotide bases in an mRNA molecule dictates the sequence of amino acids, and thus, the identity, structure, and function of the resulting protein (Fig. 1.5). It is interesting to note that the essential translation activities are executed by rRNA and not RPs. Many ribosomal proteins which are constituent components of the machine are not essential for survival (Akanuma et al. 2012; Shoji et al. 2011). This suggests that the first ribosomes were likely composed entirely of RNA (Moore and Steitz 2002) and that the translation machinery could work before the evolution of coded ribosomal proteins (Fox 2010; Root-Bernstein and Root-Bernstein 2015). Figure 1.5 – Cartoon ribosome decoding an mRNA sequence into protein Ribosomes polymerize proteins using mRNA-encoded sequence information and tRNAs charged with specific amino acids. The 40S SSU (bottom) and 60S LSU (top) travel along an mRNA in the 5’ to 3’ direction (left to right) catalyzing the production of a coded protein by polymerizing amino acids in an ordered manner in the N to C direction (left to right). Three tRNAs are shown in the LSU decoding M (methionine), I (isoleucine), and T (threonine) from the codons AUG, AUU, and ACC respectively in the SSU (Table 1.1). 20 The genetic code As the genetic code contains 61 amino acid related codons and there are only 20 amino acids, the code is redundant (Crick et al. 1961). This means that there are multiple codons, frequently two to four, coding for the same amino acid. Because of this, it often does not matter what the 3rd base in a codon is, because all four codons will result in the same amino acid being used for addition to the nascent polypeptide. The decoding center in the ribosome strictly enforces Watson-Crick base pairing at the 1st and 2nd bases in the anticodon-codon interaction, and allows the 3rd base to experience a so-called wobble, such that nucleotides other than the original Watson-Crick base pairs can be used. Therefore, an organism does not need to produce tRNA genes for all 61 anticodons to still have effective translation. Most species encode about 45 (of the possible 61) tRNA genes for in their genome (Tang et al. 2009). single amino acid letter RNA codons code Isoleucine I AUU, AUC, AUA Leucine L CUU, CUC, CUA, CUG, UUA, UUG Valine V GUU, GUC, GUA, GUG Phenylalanine F UUU, UUC Methionine M AUG Cysteine C UGU, UGC Alanine A GCU, GCC, GCA, GCG Glycine G GGU, GGC, GGA, GGG Proline P CCU, CCC, CCA, CCG Threonine T ACU, ACC, ACA, ACG Serine S UCU, UCC, UCA, UCG, AGU, AGC Tyrosine Y UAU, UAC Tryptophan W UGG Glutamine Q CAA, CAG Asparagine N AAU, AAC Histidine H CAU, CAC Glutamic acid E GAA, GAG Aspartic acid D GAU, GAC Lysine K AAA, AAG Arginine R CGU, CGC, CGA, CGG, AGA, AGG Stop codons Stop UAA, UAG, UGA Table 1.1 – The genetic code The twenty amino acids found in biologically-encoded proteins are listed alongside the single-letter symbols used to represent them. All 64 possible 3-letter combinations of the RNA nucleotides adenine (A), uracil (U), cytosine (C), and guanine (G), so-called codons, are listed alongside the specific amino acid each codon encodes for (Fig. 1.5) or next to the stop codon that signals the end of a sequence (Fig. 1.4). While RNA messages can be expressed unambiguously into protein sequence it is impossible to predict an RNA sequence from its protein sequence due to redundancy. This property ensures the unidirectional flow of genetic information from nucleic acids to proteins (Fig. 1.1). Three steps for a ribosome to translate an mRNA Translation is often separated into three major steps: initiation, elongation, and termination. Over these steps of translation, the RNA and protein components of the translational machinery will interact to assemble an 80S ribosome, capable of decoding a specific mRNA and translating the resulting message into a nascent polypeptide. 21 Canonical cap-dependent initiation forms the machinery into an elongating 80S ribosome at the translation initiation site which also defines the ORF to be translated by the ribosome. The machinery forms into a peptide bond fabricator during elongation. Termination of translation involves the release of a nascent polypeptide and reforming the machinery back into its modular components. As initiation is considered to be the rate- limiting step of translation, control of 80S assembly on mRNAs is one of the most fundamental regulatory processes in gene expression. Many considerations must be taken to determine the probability that any specific RBP, translational machinery component, or otherwise, recognizes and binds to an mRNA. Importantly, the relative concentrations of each component and the relative affinities among each component are significant determinants. Many mRNA-specific features can also inform the probability of a given ORF to be recognized by the initiation complex, e.g., 5’UTR length, mRNA secondary structure, cap availability, and poly(A) tail length (Fig. 1.6). In cellular, and especially developmental, contexts, sequence-specific and non- specific RBP associations with mRNAs are robust translational control elements, known to regulate gene expression in a variety of ways (Harvey et al. 2018). The rate for a given mRNA (m) to successfully begin translation is the mRNA-specific initiation rate (ki). Figure 1.6 – Components of the translational machinery An mRNA (m) is bound by several RBPs, including translation factors, that change its availability and affinity to the 40S ribosomal SSU (light grey) and 60S ribosomal LSU (dark grey) of the ribosome. The twenty amino acids are covalently attached to specific tRNAs containing anticodons listed (Table 1.1). Translation factors and ribosome associated factors (green) coordinate the interactions among these modular components during each step of translation. Over various intermediate compositions of the translational machinery, an 80S ribosome eventually forms at the start codon of the ORF being translated into protein. Focusing on initiation Regulatory elements involved in translation initiation allow for regulated tuning of gene expression. Eukaryotic translation starts in the cytoplasm. To initiate translation, helicase eIF4A, cap-binding protein eIF4E, and scaffold protein eIF4G interact to form the eIF4F complex. This complex will bind to the 5’ cap of mRNAs. Initiation factor eIF4G interacts with PABP possibly aiding in circularization and re-initiation of translation after full cycle of initiation-elongation-termination (Ivanov et al. 2016). The initiator tRNA is complexed 22 to an eIF2-GTP, referred to as the eIF2 Ternary Complex, usually containing a methionine. The ternary complex binds a 40S subunit containing eIF1, eIF1a, eIF3, and eIF5, thus forming the 43S pre-initiation complex (PIC) (Fig. 1.7). Figure 1.7 – Cap-dependent translation initiation in eukaryotes The 43S pre-initiation complex (PIC) is composed of eIF1, eIF1a, eIF3, eIF5, and an eIF2-Met-tRNA ternary complex. An mRNA is bound by an eIF4F cap-recognition structure, composed of eIF4A, eIF4G, and eIF4E, and is thus capable of being recognized by an available 43S PIC. Upon binding, the 48S complex remains cap-tethered via eIF4E, traverses the 5’UTR by unwinding RNA via eIF4A, and maintains association with the mRNA via eIF4G. This 48S ribosome will “scan” 5’ to 3’ until a start codon is recognized. Then, eIF2 dissociation and temporary binding of eIF5B correctly position the 60S onto the 48S at the +1 codon. Adapted from (Komar and Merrick 2020). Recognition of any given mRNA by a PIC is the rate-limiting step of translation. The PIC binds an mRNA with an eIF4F complex and the newly formed 48S complex will slide along the 5’UTR of the mRNA scanning until the recognition of a start codon (AUG). Upon commitment to start codon-anticodon recognition, the 48S complex is stabilized, eIF2- GTP is hydrolyzed, thus dissociating eIF2, allowing for the binding by eIF5B, and subsequent fixation of the LSU assembly. This position in the mRNA is known as the translation initiation site, and defines the ORF that will be decoded for synthesis. When the LSU is fixed in place, eIF1, eIF1A, eIF3, eIF5, and eIF5B are ejected from the completed complex. The final product of initiation is a translation-competent 80S ribosome, bound to the mRNA, containing a methionine-tRNA in the P-site (Fig. 1.7). 23 The start codon defines the translation initiation site It should be noted however, that translation can also be initiated from non-AUG start codons, such as near-cognate AUG codons in 5’ UTRs, or, during non-ATG (RAN) translation, from the multiplicity of repetitive RNA sequences. Intriguingly, this near- cognate initiation has been shown to especially occur in certain cell states, e.g., during cell stress and meiosis (Crosby et al. 2015; Ivanov et al. 2011). Some mRNAs do not require a cap structure to be translated. The presence of structured RNA cis-elements, like internal ribosome entry sites (IRESs) may allow for direct recruitment of translational machinery to the mRNA; effectively bypassing cap recognition and 5’UTR scanning altogether (Schuster and Hsieh 2019; Sonenberg and Hinnebusch 2009; Svitkin et al. 2005). IRESbase is a comprehensive literature database of experimentally validated functional minimal IRES elements currently containing 774 IRESs from 11 eukaryotes and 554 IRESs from 198 viruses. Likely due to a lack of reporter assays, 691 of the 774 known eukaryotic IRESs have been discerned from the human transcriptome while only 83 IRES sequences have been identified in other eukaryotes (Yang et al. 2021; Zhao et al. 2020). Start codon trinucleotides, referred to as upstream AUGs (uAUGs), and short open reading frames, termed upstream ORFs (uORFs) are often observed in 5′ UTRs of protein-coding genes (Morris and Geballe 2000). Sometimes, the scanning ribosome bypasses the initial AUG and begins translation at further downstream AUG start codons (Kozak 2002). The scanning-from-cap mechanism predicts that translation should initiate at the AUG codon most proximal to the 5′ end of the mRNA (Kozak 1978); however, the scanning 48S ribosome may encounter an “unfavorable nucleotide context” around the start codon and continue scanning (Herzog et al. 1995). This is why the sequence context surrounding the start codon is thought to be of particular importance to protein synthesis. It has been reported that ∼55% and ∼25% of mammalian genes have one or more uAUGs and uORFs, respectively (Crowe et al. 2006). These uAUGs and uORFs are conserved among species (Chew et al. 2016), have been shown to be involved in the down- regulation of translation (Meijer and Thomas 2002). In zebrafish, uORFs have been shown to be prevalent regulators of mRNA transcript levels, are actively translated through development, and act to regulate translation control of an mRNA (Johnstone et al. 2016). Specific translational machinery components may have altered affinities for the sequence surrounding the start codon and could change the mRNA-specific initiation rate for a given ORF. Mutating these gene sequences or occluding these mRNA nucleotides potentially affects the gene expression outcomes of particular transcripts. The consensus sequence surrounding the start codon was generated from 699 vertebrate genes and initially reported by Kozak as 5’GCC(A/G)CCAUG·GNN·NNN3’ in which the underlined AUG is the translation initiation site and therefore defines the 3-nucleotide spacing of the following ORF (Kozak 1987). The consensus motif calculated using over 10,000 human protein-coding genes is similarly 5’GCC(A/G)(C/A)CAUG·GCG·NNN3’ and comparable 24 to the motif calculated using 8,000 zebrafish genes 5’GCCAACAUG·GCG·NNN3’ (Nakagawa et al. 2008). It has been suggested that a short element adjacent to the start codon in a eukaryotic mRNA may directly base pair with an 18S rRNA to enhance translation initiation, in a similar manner to the interaction of the Shine-Dalgarno sequence with a 16S rRNA in a prokaryotic mRNA (Chappell et al. 2006; Dresios et al. 2006; Mauro and Edelman 2002, 2007). The data are insufficient to conclude whether sequence-specific mRNA-rRNA interactions occur near start codons to influence eukaryotic translation initiation rates. The possibility of mRNA-rRNA interactions influencing initiation, otherwise known as the ribosome filter hypothesis, is introduced later. Ribosomes elongate at codons and terminate at stop codons After initiation, three events are repeated for every codon: (1) recognition of the current codon with the help of a transfer RNA (tRNA); (2) peptidyl transfer allowing the next amino acid to be linked to the growing polypeptide; and (3) mRNA-tRNA translocation permitting the ribosome to move on to the next codon (Fig. 1.8). Each charged tRNA holds a unique combination of anticodon and amino acid, which allows for a precise translation of proteins based on the genetic code (Table 1.1). Many interactions ensure the correct positioning of mRNA in the SSU, the peptidyl transferase center (PTC) in the LSU, and the three correct mRNA codons to be displayed for tRNA interaction. Figure 1.8 – Translation elongation The 80S ribosome is shown with a Met-tRNA in the P-site. When the DC senses a correct anticodon-codon match from the eEF1A-bound tRNA in the A-site, in this case Val-tRNA, eEF1A hydrolyzes a GTP to place the amino acid in the PTC. As peptidyl transferase completes the addition of a single amino acid, the uncharged tRNA is moved to the E-site for exit, eEF2 hydrolyzes a GTP to sequentially translocate the ribosome to the next codon for another round. The nascent polypeptide is simultaneously pushed into the exit tunnel. Adapted from (Komar and Merrick 2020). Each ribosome possesses three tRNA binding sites: the A-site in which the aminoacyl- tRNA positions itself, the P-site where the peptidyl-tRNA is located, and the E-site in which the unloaded tRNA leaves the ribosome (Fig. 1.8). tRNAs are stabilized within the ribosome through numerous contacts with RPs and rRNAs. At the A-site, one tRNA is positioned to allow the correct contact between the mRNA codon and the anticodon in a zone of the SSU called the decoding center (DC). During one cycle of elongation, a ribosome recruits an aminoacyl-tRNA into the A-site, the DC checks for pairing between the mRNA codon and tRNA anticodon, if correct, the PTC forms the peptide bond, and the ribosome maneuvers into a conformation suitable for a new cycle of translation (Ben- 25 Shem et al. 2011; Melnikov et al. 2012). After forming a new peptide bond and displacing the uncharged tRNA from the E-site, the cycle can restart (Fig. 1.8). Translation termination occurs when ribosomes encounter a stop codon (UAA, UGA or UAG) in the A-site (Table 1.1, Fig. 1.4). This process is mainly assisted by translation factors eRF1 and eRF3, which form a complex with GTP. When eRF1 recognizes the stop codon, eRF3 hydrolyzes GTP, thus releasing the polypeptide chain, uncoupling the ribosome from the mRNA transcript, splitting the 40S and 60S subunits, and dissociating the machinery back into recycled modular components for continued rounds of translation (Dever and Green 2012; Hellen 2018) Regulating steps of translation Gene expression is a multistep process that involves transcription, translation, and turnover of mRNAs and proteins. In the past, analysis of gene expression regulation had been primarily focused on changes to the DNA that could alter mRNA levels, the transcriptional output. Recently, exemplary research points to the predominant role that translation rates control protein levels (Hentze et al. 2018) it is clear that each step of translation can be controlled by RBP- mediated regulatory events to obtain a specific cellular proteome (Fig. 1.9) . Translational control involves several signaling pathways which would allow Figure 1.9 – Expression regulation by RBPs the adjustment of the proteome Many RNA binding proteins (RBPs) will interact depending on the environmental sequence-specifically and non-specifically with conditions (nutrient availability, an mRNA. Binding can influence the mRNA’s ability to interact with cellular components hormones, stress, etc.), on the cell type, including the translational machinery. RBPs or even on the cell cycle phase (Ho et al. which alter turnover rates of mRNAs by 2020). Unsurprisingly, translational stabilization or destabilization are of particular control is critical for proper embryonic interest when assess gene expression (Harvey et development (Despic et al. 2017; al. 2018). Adapted from (Tran and Rao 2022). Tahmasebi et al. 2018). Regulators of translation are often described as having “global” effects on all mRNAs in the system or “specific” effects on subsets of mRNAs (Schwanhäusser et al. 2011). A portion of the cellular proteome codes for factors enabling regulation of global protein synthesis as well as translation control of specific. Regulation by RBPs can occur during mRNA transcription and processing, during transport, during decay, or during any of the three steps of translation (Perez-Perri et al. 2018). 26 Modeling protein synthesis, a polymerization reaction A reaction analysis of protein synthesis was first achieved by MacDonald and Gibbs, who derived the relevant rate equations (MacDonald and Gibbs 1969; MacDonald et al. 1968) and Lodish who determined the relevant assumptions necessary to derive initiation and elongation rates of translation in eukaryotic cells (Lodish 1974). The effective protein synthesis rate for a given mRNA in a cell is given by the equation (Fig. 1.10). Figure 1.10 – A model for polypeptide synthesis rate Q is the protein synthesis rate, m is the expression of the given mRNA, R is the ribosome concentration, and ki is the mRNA-specific initiation rate. Another term involves the mRNA’s specific length and ribosome density; including ke, the termination rate, and L, the number of codons occupied by one ribosome. The mRNA-specific initiation rate describes the combined steps for assembling an 80S monosome onto an mRNA: recognition of the mRNA 5’ cap, 43S scanning for a start codon, and 60S joining. By taking a range of R estimates, based on mass spectrometry (Kim et al. 2014), and a range of ki estimates (Noderer et al. 2014), based on reporter screens, simulated mRNA translation rates reveal the fundamentals of gene expression regulation (Fig. 1.11). Figure 1.11 – Protein output changes by ribosome concentration Matlab and pyplot were used for simulations of the model (Fig. 1.10). Similarly to (Mills and Green 2017), m, mRNA level, is set to 15, L, the number occupied codons, is set to 10, ki, represents a ranking of mRNA- specific initiation rates, set from 0.01 to 1, and R, the number of ribosomes, is set to range from 0.0004 to 1.2. Simulation plots are rotated 90º compared to those in (Mills and Green 2017). The colorimetric scale represents the row Z-score of simulated synthesis rates, with low synthesis in red and optimal synthesis in grey. Qt refers to the amount of protein synthesized over a specific period of time. Protein outputs of the magenta, yellow, and green mRNAs (below) are indicated by filled circles (right). 27 Figure 1.12 – Multiple initiations first increase, then lower synthesis Changing the concentration of 40S and 60S subunits (R) changes the density of ribosomes on a given mRNA (yellow). When the system is at an optimal ribosome concentration: the cycles of initiation, elongation, and release are timed to maximize ribosome density and minimize ribosome collisions, either an increase or a decrease lowers the protein synthesis rate. The circle fills refer to the amount of protein output (yellow) over a specific period of time (Qt). The amount which ΔR influences overall synthesis is dependent on the mRNA-specific initiation rate (ki). One of the key findings from this modeling, as well as others, is that the mRNA-specific initiation rate defines the influence of ribosome concentration (R) on protein synthesis rates (Q) (Fig. 1.12). Repeated initiations ribosomes on mRNAs can increase the protein synthesis rate by forming polysomes. However, a too-high ribosome concentration will eventually cause too many ribosomes to initiate, cause collisions, and lower the synthesis rate (MacDonald et al. 1968). The ribosome concentration regime between low density and optimal density is smaller for lowly initiated mRNAs (MacDonald and Gibbs 1969). Also, mRNAs with higher initiation rates are better able to buffer their synthesis rates when faced with changes to ribosome concentration. Under this model, limitations on translation initiation are predicted to differentially effect mRNAs depending on their individual initiation rates (Lodish 1974; Mills and Green 2017). Measuring translational components by separation Rate-zonal centrifugation is a widely used technique that separates particles based on sedimentation velocity, taking into consideration their size, shape, and density (Brakke 1951). Separating biomaterials was among the first usages of this technique (McQuillen et al. 1959). An approximate separation of non-translated and translated mRNAs by sedimentation rate is achieved using a density cushion – material below 60S contains free subunits and untranslated mRNAs, while material above 60S contains mRNAs with elongating ribosomes. The proportion of ribosome subunits committed into active translation can be evaluated by measuring RNA concentration in each of the two fractions. The proportion of an mRNA committed into active translation can be similarly evaluated. When taken as the fraction bound vs unbound, this measurement approximates binding affinity, and consequently serves as proxy for mRNA specific initiation rate (ki) (Fig. 1.6). Immobilizing ribosomes onto mRNA using translation inhibitors, such as cycloheximide (CHX), in the lysis buffer can help stabilize polysomes. This measurement is a snapshot assessing the translational status of the sample including all subunits and mRNAs: unbound, bound, and bound by multiple. 28 Figure 1.13 – Sucrose density gradient ultracentrifugation A) A schematic density gradient linearly ranging from 10% to 50% sucrose along the axis of the tube. A lysate (green) is loaded adjacent to the 10% sucrose. An ultracentrifuge accelerates the material with 210,000 xg for 2.5 hours. Molecules are fractionated along the gradient based on sedimentation velocities. Once finished, the gradient material can be collected in sequential fractions along the length of the tube (mm). B) Live UV A260 absorbance readings are shown (grey trace). The 40S SSU (light grey), and 60S LSU (dark grey) are illustrated above the peaks representing the position of free subunits. While 6-ribosome polysome peaks are detectable, only the monosome, 2-ribosome, and 4-ribosome polysome peaks are illustrated. For reference, objects on the surface of the Earth are subject to 1 xg, about 9.8 m/s2. Further separation of mRNA transcripts into several polysome fractions is achieved by sucrose density gradient ultracentrifugation and can be used to separate mRNA transcripts on the basis of the number of ribosomes bound to each of them (Arava et al. 2003; Hendrickson et al. 2009) (Fig. 1.13A). A homogenized suspension of the sample, the lysate, is pipetted into a thin layer on top of a higher density solution. The tube is subjected to high g-forces in a direction towards the bottom of the tube (Fig1.13A). Upon acceleration into the solution, all molecules in the lysate will move away from the starting position with a velocity determined by their specific size, shape, and density. At a specific time later, molecules will have transited into the solution, now spaced according to their relative velocities in the lysate. Therefore, the molecular qualities of size, shape, and density are nonlinearly mapped onto positions along the linear axis of the tube (Fig1.13B). The mapping relationship between sedimentation rate and a particle’s size is given by the Svedberg unit (S). The RNA concentration at various depths into the gradient, as measured by UV absorbance A260, corresponds to the fraction of ribosomes in the lysate existing as free subunits, as 80S monosomes, or in polysome assemblies (Fig. 1.13B, trace). Successive 29 polysomes form a graded series detectible in higher density solutions as each translating ribosome adds an additional 4 megadaltons to the molecule (Fig. 1.13B, peaks). Nuclease footprinting has long been recognized as a way to determine the positions of ribosomes on the mRNAs that they are actively translating (Steitz 1969; Wolin and Walter 1988). Cycloheximide-treated ribosomes become stalled during elongation and physically enclose 28–30 nucleotides of the mRNA transcript, consequently shielding this region from nuclease digestion. Polysome-bound mRNAs are typically digested down to monosomes using a ribonuclease (usually RNAseI or RNAseT1) to remove the mRNA regions between individual ribosomes. Sucrose density ultracentrifugation is then used to collect These protected mRNA fragments indicate the exact location of the cycloheximide- stalled ribosome (Ingolia et al. 2012; McGlincy and Ingolia 2017; Power 2022). 30 Ribosome structures support their functions As mentioned previously, protein synthesis is conducted by ribosomes, each composed of a large (LSU) and small (SSU) subunit. The ability to structurally resolve elements composing the ribosome has been entirely dependent on technical advances made in X- ray crystallography and, more recently, cryo-electron microscopy (cryo-EM). In the 1990’s electron microscopy was able to generate highly detailed images of molecules; however, the technology required optimization and was incapable of being applied to a complex as large as the 4 megadalton ribosome. Instead, a lower-than-atomic resolution structure (5.5 Å) was solved using X-ray crystallography; the work eventually earning a Nobel prize in 2009 (Clemons et al. 1999). An atomic-resolution (3 Å) structure of the eukaryotic (yeast) ribosome was first acquired using X-ray crystallography (Ben-Shem et al. 2011) with similarly high resolution structures of human, drosophila, and zebrafish ribosomes being resolved since then using cryo-EM (Hopes et al. 2022; Khatter et al. 2015; Leesch et al. 2023) (Fig. 1.14). These structures are publicly available at the Protein Data Bank (PDB, www.rscb.org) and are used for illustrative purposes in this Thesis. Human ribosomes, aka 80S ribosomes, contains 80 RPs and are scaffolded by 4 rRNA molecules. SSUs, aka 40S subunits, are composed of 33 RPs and an 18S rRNA. LSUs, aka 60S Figure 1.14 – An assembled fruit fly ribosome subunits, are composed of 47 RPs and An atomic model representation of the 3.0 Å the 5.8S, 28S, and 5S rRNAs (Anger et resolution structure of the fruit fly ribosome. The al. 2013; Ben-Shem et al. 2011; Khatter LSU rRNAs and RPs are in blue while the SSU rRNA and RPs are in yellow. (PDB: 6XU8) et al. 2015). The large subunit (LSU) The human LSU is scaffolded by 3 rRNAs – the 5.8S, 28S, and 5S – and contains 47 RPs distinct to the 60S subunit (Table 1.2 and Fig. 1.14). While previously known as RPL, these proteins are now known as eL and uL (e, referring to eukaryotic; u, referring to universal) (Ban et al. 2014). The LSU forms a variety of functional domains of the ribosome (Melnikov et al. 2012). Of particular relevance to this Thesis are the central protuberance (CP), the GTPase associated center (GAC), the sarcin-ricin loop (SRL), the P-stalk, the nascent polypeptide exit tunnel, and the peptidyl transferase center (PTC). The CP is made up of a three-member complex, known as the 5S RNP (composed of 5S rRNA, RPL11/uL5, and RPL5/uL18) and the 28S rRNA. Cryo-EM modeling demonstrated the CP participates in intersubunit bridge formation and coordinates translation by connecting the PTC with the decoding center in the SSU (Rhodin and Dinman 2011). 31 60S large subunit (LSU) 40S large subunit (SSU) universal human zebrafish universal human zebrafish uL1 L10A Rpl10a eS1 S3A Rps3a uL2 L8 Rpl8 uS2 SA Rpsa uL3 L3 Rpl3 uS3 S3 Rps3 uL4 L4 Rpl4 uS4 S9 Rps9 uL5 L11 Rpl11 eS4 S4 Rps4x uL6 L9 Rpl9 uS5 S2 Rps2 eL6 L6 Rpl6 eS6 S6 Rps6 eL8 L7A Rpl7a uS7 S5 Rps5, Rps5l uL10 P0 Rplp0 eS7 S7 Rps7 uL11 L12 Rpl12 uS8 S15A Rps15a uL13 L13A Rpl13a eS8 S8 Rps8a, Rps8b eL13 L13 Rpl13 uS9 S16 Rps16 uL14 L23 Rpl23 uS10 S20 Rps20 eL14 L14 Rpl14 eS10 S10 Rps10 uL15 L27A Rpl27a uS11 S14 Rps14 eL15 L15 Rpl15 uS12 S23 Rps23 uL16 L10 Rpl10 eS12 S12 Rps12 uL18 L5 Rpl5a, Rpl5b uS13 S18 Rps18 eL18 L18 Rpl18 uS14 S29 Rps29 eL19 L19 Rpl19 uS15 S13 Rps13 eL20 L18A Rpl18a uS17 S11 Rps11, Rps11l eL21 L21 Rpl21 eS17 S17 Rps17 uL22 L17 Rpl17 uS19 S15 Rps15 eL22 L22 Rpl22, Rpl22l1 eS19 S19 Rps19 uL23 L23A Rpl23a eS21 S21 Rps21 uL24 L26 Rpl26 eS24 S24 Rps24 eL24 L24 Rpl24 eS25 S25 Rps25 eL27 L27 Rpl27 eS26 S26 Rps26, Rps26l eL28 L28 Rpl28 eS27 S27 Rps27.1, Rps27.2, Rps27l uL29 L35 Rpl35 eS28 S28 Rps28 eL29 L29 Rpl29 eS30 S30 Faua, Faub uL30 L7 Rpl7, Rpl7l1 eS31 S27A Rps27a eL30 L30 Rpl30 eL31 L31 Rpl31 eL32 L32 Rpl32 eL33 L35A Rpl35a eL34 L34 Rpl34 eL36 L36 Rpl36 eL37 L37 Rpl37 eL38 L38 Rpl38 eL39 L39 Rpl39 eL40 L40 Uba52 eL41 L41 Si:dkey-151g10.6 eL42 L36A Rpl36a eL43 L37A --- P1 P1 Rplp1 P2 P2(αβ) Rplp2, Rplp2l Table 1.2 - Ribosome proteins with universal, human and zebrafish nomenclature The new standard nomenclature (Ban et al. 2014). Zebrafish ribosome proteins and their paralogs are listed alongside presumed universal and human orthologs. The GAC acts as an entry zone for translation-associated GTPases required during translation for GTP hydrolysis (Rodnina 2016). The GAC is composed of an interaction between the SRL and the P-stalk (Uchiumi and Kominami 1994). The flexible P-stalk is formed by acidic proteins RPLP0/uL10, RPLP1/P1, and RPLP2/P2 and is responsible for coordinating GTP hydrolysis with recruitment of elongation factors (Liljas and Sanyal 32 2018). Importantly, only RPLP0/uL10 forms the base of the P-stalk and has been shown to be necessary for ribosome function, hence its differing nomenclature (Ban et al. 2014). A B Figure 1.15 – Architecture of the nascent polypeptide exit tunnel A) A cross section cartoon of the exit tunnel illustrating its interior shape containing portions of RPs uL4, uL22, and eL39 (not shown). During elongation, the nascent chain (NC) is directed toward the solvent side, away from the PTC. The PE region contains portions of RPs uL23, uL24, and uL29. Adapted from (Bock et al. 2018). B) An illustrated model showing the 80S monosome with locations of aminoacyl (A), peptidyl (P), and exit (E) sites marked between the 40S and 60S subunits. The exit tunnel is drawn in dark grey with a dotted outline. The PE, peptide exit, region is marked. Exit Tunnel and Peptide Exit The newly synthesized amino acid is on the C terminus of the protein in the nascent polypeptide exit tunnel. This tunnel is formed by the 28S rRNA, RPL4/uL4, RPL17/uL22, and RPL39/eL39 (Fig. 1.15A). Note the placement of RPL17/uL22 as paralog expression of this RP is seen in the germ cells of various animals. The first RPs to interact with the nascent protein beyond the exit tunnel, on the solvent side, are RPL26/uL24, RPL35/uL29, and RPL23A/uL23 in the peptide exit (PE) region (Fig. 1.15A and 1.15B). RPL23A/uL23 is a known mTORC2 interactor during translation when mTOR kinase activity is thought to be positioned to phosphorylate nascent targets (Oh et al. 2010). Peptidyl transferase center This position contains the ribosome’s catalytic activity and is composed of 28S rRNA only (Klinge et al. 2012). The peptidyl transferase ribozyme catalyzes two chemical reactions – first, the amino acid addition to the nascent protein, formed by a peptide bond, and second, the release of the nascent polypeptide from the ribosome. The amino acid on the aminoacyl-tRNA of the A site forms a bond with the peptidyl-tRNA in the P site. The PTC and exit tunnel are interconnected (Fig. 1.15B). Critically, the presence of a polypeptide in the exit tunnel will induce ribosome stalling by inhibition of the PTC (Seidelt et al. 2009). 33 The small subunit and the decoding center The human SSU is scaffolded by the 18S rRNA and contains 33 RPs distinct to the 40S subunit (Table 1.2) While previously known as RPS, these proteins are now known as eS and uS (e, referring to eukaryotic; u, referring to universal) (Ban et al. 2014). The 18S rRNA forms an mRNA tunnel between the “head” and the “body” of the 40S. The SSU exhibits only contains one functional structure. The DC is located in the A-site position. It is composed of 5 helices (H18, H44, H34, H24, H31) of the 18S rRNA. It is surrounded by RPS3/uS3, RPS9/uS4, RPS2/uS5, RPS15/uS19, RPS23/uS12, and RPS30/eS30. The role of the DC during each elongation cycle is to ensure the recruitment of correct cognate tRNA, while preventing near-cognate anti-codons, differing by one mismatch. This structure is constantly monitoring the fidelity of mRNA-tRNA interaction. When an eEF1A-GTP-bound cognate tRNA binds to the mRNA codon in the A-site, a conformational change in eEF1A leads to GTP hydrolysis and eEF1A-GDP releases the tRNA into the A-site such that the amino acid is near the PTC (Poirot and Timsit 2016; Rodnina 2016; Timsit et al. 2021). At the end of translation, usually when a stop codon has reached the A-site, a hydrolysis reaction will free both the newly synthesized polypeptide and the final deacylated tRNA. The aminoacyl, peptidyl, and exit sites The 60S forms the A-, P-, and E- tRNA sites at the interface of the LSU and the SSU where the 40S displays the mRNA being decoded. The A-site, composed of RPL3/uL3 and the DC, will host aminoacyl-tRNAs for anticodon-codon recognition. The P-site contains a peptidyl-tRNA and is structured by RPL5/uL18, RPL10/uL16 and RPL36A/eL42 (Fig. 1.16). The E-site is for the deacylated tRNA after peptide bond formation. Critically, the E-site must be empty to move the uncharged tRNA from the P-site, else the ribosome will stall. Cycloheximide (CHX) works as a potent inhibitor of the elongation by Figure 1.16 – Intersubunit regions of interest A labeled schematic depicting various ribosome binding in the ribosomal E-site (Pestova features over a surface model of the 80S. The and Hellen 2003; Schneider-Poetsch et SSU (bottom) contains the decoding center and al. 2010). CHX is a natural fungicide its associated elongation factor binding sites. The produced by the bacterium Streptomyces LSU (top) subunit forms the aminoacyl, peptidyl, griseus. CHX was initially reported in and exit sites for tRNAs to sequentially position amino acids at the peptidyl transferase center 1946 by Alma Joslyn Whiffen-Barksdale. (PTC). The B1b/c intersubunit bridge connects CHX effectively kills fungal cultures at a 40S and 60S subunits between RPS18/uS13 and concentration of 0.2 µg/mL by halting RPL11/uL5 and is stabilized by 5S rRNA. translation. The CHX-bound ribosome Adapted from (Rhodin and Dinman 2011). will arrest on a translating mRNA at the specific codon its occupying. 34 Expansion segments When rDNA gene sequences from multiple organisms are compared, it is apparent that eukaryotic rRNAs are longer than prokaryotic rRNAs. Further, while rRNAs in yeast are longer than bacteria, the rRNAs of fishes, amphibians, birds, and mammals are longer still. This increase in length is due to nucleotide insertions of varying sizes at specific conserved positions on the rRNA. These species-specific regions are GC-rich and referred to as expansion segments (ESs) (Clark et al. 1984). Other than at these specific sites, rRNAs across all species exhibit a high degree of structural conservation regardless of primary sequence changes (Fig. 1.17). This is primarily facilitated by compensatory base-pair mutations which maintain RNA secondary structures over time. Figure 1.17 – Expansion segment complexification of the LSU rRNA The secondary structure maps of the A) 23S rRNA in E. coli, B) the 25S rRNA in S. cerevisiae, and C) the 28S rRNA in H. sapiens are shown. Distinct regions of the rRNA are color coded. Note the differences in ES27L (top, yellow) and ES7L (bottom, purple). The labeled map of human ES sites (right) has been modified with ESs in red and core regions in green. Adapted from (Bernier et al. 2018; Petrov et al. 2014). 35 ESs were originally thought to lack function relating to the ribosome and instead were thought to be tolerated insertion mutations in the rDNA gene, in rRNA folding, in ribosome biogenesis, in the mature ribosome, and in translation. This total lack of interference to ribosomal functionalities could explain their expansion (Clark et al. 1984). The possibility that neutral sequence accumulation could be the origination of sequence variation is addressed in the Chapter III. More recently, ribosome structure analyses have shown ESs to form into tentacle-like structures that protrude from the ribosome; not associated with the core RPs (Yusupova and Yusupov 2014) (Fig. 1.18). And, there has been growing interest suggesting potential roles rRNA ESs could be playing in translation – ranging from ribosome localization to translation fidelity (Fujii et al. 2018). The idea is that the structural core of the ribosome is common across prokaryotes and eukaryotes and the expanded areas could be platforms for additional functionalities (Fig. 1.17A and 1.17C). Figure 1.18 – Expansion segment tentacles of the LSU In line with the ribosome containing a common core and ESs existing outside of that core, an illustration of ES sequence tentacles found in the yeast LSU. Adapted from (Yusupova and Yusupov 2014). When regarding ribosome functional regions, large subunit ES4L, ES7L, ES27L (Fig. 1.17C), and small subunit ES3S, ES6S are a handful of expansion segment structures to note. ES4L in the 5.8S rRNA forms a cluster with other ESs – ES5L, ES19L, and ES31L – further stabilizing the 5.8S position along the B1b/c bridge and P-site-tRNA (Chandramouli et al. 2008) (Fig. 1.16) Notably, the majority of the length increases seen in 28S rRNAs of vertebrates come from just two ESs, ES7L and ES27L (Fig. 1.17C and Fig. 1.18). ES7L, an ES of the LSU, increases in length progressively from single-celled eukaryotes to mammals. Growth of ES7L occurs by iterative accretion of RNA fragments onto the more conserved basal 36 structure. For instance, the ES7L region ranges from around 20 nucleotides in bacteria, 80 nucleotides in archaea to about 210 nucleotides in non-vertebrates, and over 870 nucleotides in mammals (Petrov et al. 2014). ES3S and ES6S are in the small subunit, located where mRNA is unfolded before threading into the A-site. A tertiary interaction between ES3S and ES6S has been shown to promote unwinding and scanning in the 48S PIC (Alkemar and Nygård 2006; Díaz- López et al. 2019). It is thought that so-called “18S sticky regions” may provide rRNA complementary to mRNA 5’ UTRs during 48S scanning (Pánek et al. 2013). These conserved regions exist in ES3S, ES6S, ES7S, and ES12S. Intersubunit bridges 17 so-called intersubunit bridges stabilize the 80S structure by maintaining the positions of 40S and 60S subunits in proximity to each other (Fig. 1.19). The bridges consist of RNA-RNA interactions towards the core of the ribosome, RNA-protein, and protein- protein interactions towards the periphery. Stabilization of the structure is necessary during rotations occurring at translocation and tRNA release. RPL11/uL5, RPL19/eL19, RPL24/eL24 and RPL41/eL41 interact with the SSU while the expansion segments of the 60S subunit interact with RPS3A/eS1 and RPS8/eS8 of the 40S subunit (Klinge et al. 2012; Tamm et al. 2019). Notably, the position of 5S rRNA is stabilized by its binding partners RPL5/uL18 and RPL11/uL5 and forms the B1b/c bridge (Fig. 1.16). Figure 1.19 – Intersubunit bridges of the eukaryotic ribosome Top: an atomic model representation of the 3.0 Å resolution structure of the yeast ribosome shown in interface view, aka open book view, with LSU on the left and SSU on the right. 12 conserved bridges (B) are marked in blue, while 5 eukaryotic-specific bridges (eB) in red. Adapted from (Ben-Shem et al. 2011). Bottom: a cartoon representing 17 eukaryotic intersubunit bridges. The top row indicates the LSU portion of the bridge while the bottom row indicates the SSU portion of the bridge. For example, bridge B1a is formed by the interaction of LSU rRNA helix H38a and SSU RP Rps15/uS19. Note 11 contacts are formed by “I-bridges” between two components; while 6 are formed by “Y-bridges” among three components. 37 Ribosome biogenesis forms new subunits Ribosome biogenesis embodies a highly coordinated and controlled process by which ribosomes are synthesized and assembled. The processes occur primarily in the nucleolus with the final steps of subunit maturation occurring in the nucleoplasm and cytoplasm. Overall, ribosome biogenesis includes the activity of all three RNA polymerases, simultaneous transcription, processing, and assembly of 4 rRNAs, the synthesis, import, and incorporation of 80 RPs, and the coordination of more than 200 ribosome biogenesis factors (Brombin et al. 2015). This massive molecular manufacturing process consumes the majority of cellular energy (Warner 1999) and consequently requires tight regulation (Woolford and Baserga 2013). Figure 1.20 – Ribosome biogenesis overview Ribosome biogenesis occurs over 5 steps: synthesis of RPs, transcription of rDNA, pre-rRNA processing and modification, subunit assembly, and export to the cytoplasm for maturation. Adapted from (Li and Wang 2020). Despite the ubiquitous nature of this process, ribosome formation and protein translation need to be adapted according to the cell type and the cell environment. The proteome composition of a cell can be regulated during gene expression – either at the mRNA (transcription) or at the protein (translation) level. It has been suggested that proteome composition can differ between cells with identical translatomes (Buszczak et al. 2014). Therefore, ribosome biogenesis needs to be tunable to allow for such a finely tuned translational control. Research regarding eukaryotic ribosome biogenesis have primarily studied yeast. More recently, structures of human nucleolar and pre-60S assembly intermediates were 38 generated using cryo-electron microscopy (Vanden Broeck and Klinge 2023). A staggering 2.5 Å resolution perspective of ribosome biogenesis highlights the 60S biogenesis complex tethering pre-rRNA processing and RNA degradation to coordinate the conformational folding of, what will eventually constitute, the 28S rRNA. These findings underscore the fact that hundreds of molecules function to ensure ribosome assembly. The transcriptional outputs of RNA Polymerase III, II, and then I are described in the next sections. Each polymerase transcribes distinct regions of the genome. The transcripts they generate are used for a variety of inputs for the gene expression program. Relevant transcripts are described. Broadly, there are 5 steps in the eukaryotic ribosome biogenesis pathway: synthesis of RPs, transcription of rDNA, pre-rRNA processing and modification, subunit assembly, and export to the cytoplasm for maturation (Fig. 1.20). Ribosome biogenesis, by the numbers For perspective, in a rapidly growing culture of budding yeast, Saccharomyces cerevisiae, a single cell is capable of dividing into two daughter cells every 90 minutes. This requires doubling its ribosome content (SSUs and LSUs) from 300,000 to 600,000 molecules (Aitchison and Rout 2000). During these 90 minutes, a cell will import 150,000 ribosome proteins into the nucleus and simultaneously export 3,000 new ribosomes – per minute (von der Haar 2008). To accomplish this achievement of anabolism, RNA Pol I transcribes rDNA into the relevant rRNAs and will constitute 60% of the cell’s total transcription during this time (Warner 1999). Similarly, 50% of all RNA Pol II transcription and 90% of all mRNA splicing activity is allocated to the expression of RPs (Moss and Stefanovsky 2002). The cell will transcribe 3,000,000 tRNAs and 60,000 mRNAs to be used as materials for translation (Ares et al. 1999; Waldron and Lacroute 1975). While growing yeast are capable of producing a substantial quantity of components for translation, any single mRNA’s engagement with the ribosome is still rate-limited by the availability of free subunits (Shah et al. 2013). RNA Pol III transcribes 5S genes into 5S rRNA A mature eukaryotic ribosome including is constructed using 4 distinct rRNA molecules: 18S, 28S, 5.8S and 5S rRNAs. The 18S, 28S, and 5.8S rRNAs are synthesized in the nucleolus by transcription of the rDNA gene, sometimes known as the 47S rDNA. Transcription of the 5S rRNA occurs in the nucleoplasm (Fig. 1.20). Transcription initiation factors TIF-IIIA, TIF-IIIB, TIF-IIIC and RNA Pol III associate to a 5S rRNA promoter to express the 5S gene (White 2005). The 5S rRNA will then bind Rpl5/uL18 and Rpl11/uL5 to form the 5S RNP. This three-member complex will migrate to the nucleolus to be incorporated in the LSU. Presence of this 5S RNP outside the 60S is sensed as a proxy for cellular homeostasis; free 5S RNP indicates a perturbation of ribosome biogenesis (Pelava et al. 2016). RNA Pol II transcribes protein-coding genes into mRNA, mostly RP genes Genes encoding for ribosome proteins are scattered across the genome (Uechi et al. 2001). RNA Pol II transcribes all protein-coding genes; this includes ribosome protein genes encoding RPs for the large and small subunits. RP pre-mRNAs are then spliced, 39 capped, tailed, and finally exported from the nucleus as mature mRNA. Often, the removed intronic regions of RP mRNAs contain small nucleolar RNAs (snoRNAs). These snoRNAs are used in C/D and H/ACA snoRNP complexes that modify pre-rRNA. As mentioned above, the majority of metabolic flux flows through building and operating the translation machinery, accordingly, genes involved in ribosome biogenesis and protein synthesis are the primary targets for gene expression regulation when cells respond to stress conditions. The mRNAs encoding these genes often contain a 5’ terminal oligopyrimidine tract (5’ TOP) which links their mRNA-specific initiation rate to the mTOR Complex 1 nutrient sensor (mTORC1). Immediately following the 5’ m7G cap, these mRNAs encode a cytidine residue in the +1 position, followed by up to 13 consecutive pyrimidines (Levy et al. 1991). 5’ TOP motifs, e.g. 5’ m7GpppCUCUCUUUUUUC 3’) are found in all 80 human ribosome proteins, as well as other components of the translation machinery such as poly(A) binding protein (PABP) and subunits of eIF3, eIF4A, and eEF2 (Fonseca et al. 2015; Iadevaia et al. 2008). The 5’ TOP serves as a handle for translational control in a cell type-dependent manner (Avni et al. 1997). This form of regulation changes mRNA-specific initiation rates by altering recognition of the 5’ cap. mTORC1 has been shown to enhance 5′-TOP mRNA translation via two mechanisms; phosphorylation of 4EBP1 and/or LARP1, La Ribonucleoprotein 1, an RNA-binding protein (RBP). mTORC1-mediated phosphorylation of 4EBP1 leads to inactivation; thus favoring the interaction of eIF4E to the mRNA cap (Fonseca et al. 2015; Hong et al. 2017; Lahr et al. 2017; Tcherkezian et al. 2014; Thoreen et al. 2012). This translational switch has been shown to selectively enhance 5’ TOP mRNA translation (Hsieh et al. 2012; Thoreen et al. 2012). The hypophosphorylated state of LARP1 represses translation by similarly blocking eIF4E interaction to the cap, however, upon LARP1 phosphorylation by mTORC1, this repression is released acting to enhance translation (Hong et al. 2017). This is one of many signaling paths which uses RBP recognition of mRNA motifs to quickly modulate protein expression of a subset of genes in response to changes in cellular homeostasis. After RPs are synthesized in the cytoplasm, various chaperones and transporters are required for shuttling these gene products back to the nucleus, and more precisely to the nucleolus so that they can be incorporated into the pre-ribosome particle. To accomplish this, RPs contain nuclear localization signals (NLS) that are recognized by importin β-like nuclear transporters, i.e. Importin β, Transportin-1, RanBP5 and RanBP7 (Chou et al. 2010; Jäkel and Görlich 1998) (Fig. 1.20). The rDNA gene Ribosomal DNA (rDNA) genes in yeast are located in a single tandem array on chromosome XII, with lab strains containing 150-200 gene copies. Although most eukaryotes have similarly repetitive arrays of rDNA genes, referred to as nucleolar organizing regions (NORs) in the genome, the number of NORs and the number of rDNA repeats within a NOR is variable. Some species contain NORs with fewer than 100 rDNA copies while others contain NORs with more than 10,000 copies (Long and Dawid 1980). In the human genome, roughly 400 copies of rDNA genes are present in 10 NORs located 40 on the acrocentric chromosomes 13, 14, 15, 21, and 22 (McStay and Grummt 2008; Moss et al. 2007; O’Sullivan et al. 2002) (Fig. 1.21, top). In general, more than 50% of the rDNA genes are permanently silent in differentiated cells; however, the number of active rDNA gene copies varies widely depending on cell type and cell functions. Activation and repression of rDNA genes can result from environmental cues such as stress, nutrient availability, or growth factor signals (Sanij et al. 2008). Each rDNA repeat is approximately 43 kb, of which 30 kb correspond to the intergenic spacer region containing regulatory elements and 13 kb to the precursor rRNA (47S, pre-rRNA) (Fig. 1.21, bottom). 47S Figure 1.21 – The human rDNA gene and NOR architecture The human rDNA NOR is schematized at the top containing rDNA gene repeats. Arrows indicate polymerase promoter sites. A portion containing a single repeat is magnified and labeled below. The 13 kbp 47S pre-rRNA is transcribed between the promoter and the termination site. Coding regions for mature rRNAs are labeled in green. Adapted from (Potapova and Gerton 2019). Within any given genome, the rDNA gene is characterized by its multiplicity of copies and complex arrangement of copies, each containing other repetitive elements within them. Each rDNA gene contains the sequence of a polycistronic rRNA precursor (47S in human, 45S in mammals, and 35S in yeast), as well as non-transcribed sequences, intergenic spacers (IGSs) consisting of regulatory sequences (Gonzalez and Sylvester 2001) (Fig. 1.21). Many DNA regulatory motifs have been characterized near rDNA genes and in IGSs. Examples include: Runx2 (runt-related transcription factor 2) associated with repression of chromatin regions, acts here to prevent accessibility to rDNA genes in NORs (Young et al. 2007); c-MYC (v-myc avian myelocytomatosis viral oncogene homolog) which positively regulates rDNA transcription by favoring the recruitment of SL1, a transcription factor specific to RNA Pol I, resulting in an increased frequency of initiation and an increased rate of transcription (Arabi et al. 2005; Campbell and White 2014; Grandori et al. 2005); and possibly Tp53 (tumor protein Tp53) which has been proposed to negatively regulate rDNA transcription by binding TATA-box binding protein (TBP) and preventing the assembly RNA Pol I at the promoter (Budde and Grummt 1999). 41 RNA Pol I transcribes 47S pre-rRNA The 18S, 5.8S and 28S rRNAs are encoded in the rDNA gene located in the nucleolus. The 47S pre-rRNA transcript lacks a proper TATA-box and requires the assembly of a pre-initiation complex (PIC) at the rDNA promoter on active rDNA gene copies in order to initiate transcription (White 2005). RNA Pol I transcription is often equated with nucleolar activity. Several factors specifically interact with RNA Pol I, to form a pre-initiation complex (PIC) at the rDNA promoter. These specific interactions provide a framework for specific and early modulation of nucleolar activity (Fig. 1.20 and Fig. 1.22). Upstream binding transcription factor (UBF), selectivity factor 1 (SL1) and TIF-IA (also known as RRN3) recruit RNA Pol I to the rDNA gene promoter region to form the PIC and start transcription (Russell and Zomerdijk 2005). PIC formation is the rate-limiting step of ribosome biogenesis. The transcribed RNA is a single precursor called the 47S pre-rRNA. It comprises the 18S, 5.8S and 28S mature rRNAs, separated by two internal transcribed sequences (ITS1 and ITS2) and flanked by 5’ and 3’ external transcribed sequences (5’ETS and 3’ETS) (Fig. 1.22, top). Figure 1.22 – RNA Pol I specifically transcribes rDNA genes The mouse rDNA gene is schematized at the top containing intergenic spacers (IGS), an enhancer- promoter region, the sequence of the transcribed 47S pre-rRNA, RNA Pol I terminator sequences. ChIP enrichments for mammalian RNA Pol I, RRN3, TAF1B (as proxy for SL1), and UBF proteins are shown below schematically in relation to the top panel. Mouse cell ChIP data are sourced from (Daiß et al. 2022) Pol I is enriched at the spacer-promoter and binds over the whole transcribed region. RRN3 dissociates during promoter escape before the polymerase enters a productive elongation phase and is only slightly enriched at the stalled polymerases at the spacer- promoter. TAF1B as a component of SL1 clearly localizes to the spacer and gene promoters. UBF binds at the promoter and the whole gene body (Fig. 1.22, bottom). UBF is a nucleolar localized RNA Pol I specific transcription factor (Bell et al. 1988; Moss et al. 2007; Sanij et al. 2008). Upon binding to the rDNA gene enhancer and promoter region, UBF becomes transactivated via dimerization of its N-terminal domain; this allows 42 for binding by SL1 (Hempel et al. 1996; Moss and Stefanovsky 2002). UBF has been shown to bind different regions along the entire 43kb rDNA over the cell cycle (O’Sullivan et al. 2002; Roussel et al. 1993). The SL1 complex consists of TBP (TATA-box binding protein) together with several TAFs (TBP associated factors) (Comai et al. 1992; Eberhard et al. 1993; Heix et al. 1997). Notably, paralogous TAFs have been shown to have cell-specific expressions, potentially altering the specificity of SL1 (Hochheimer and Tjian 2003). SL1 directs species- specificity, hence its name (Heix et al. 1997; Murano et al. 2014; Russell and Zomerdijk 2005). For instance, while RNA Pol I complexes and UBF proteins are interchangeable between mice and human, SL1 is not; only promoting transcription of the rDNA gene from its original source organism (Bell et al. 1990; Comai et al. 1992). TIF-IA binds to the RNA Pol I subunit RPA43 (replication protein A43) and facilitates the interaction between the polymerase and the UBF-SL1 transcription factor complex bound to rDNA gene promoters (Cavanaugh et al. 2002; Yuan et al. 2002) (Fig. 1.22). The active site of RNA Pol I lies between subunits RPA1 and RPA2, aka A190 and A135, respectively. Modification and processing Modifications are made to the canonical A, U, C, and G nucleotides composing the pre- rRNA. These are added co-transcriptionally and post-transcriptionally. 2’O-methylations (2’OMe) are a common modification in human rRNA, found in 106 sites, followed by pseudouridine (Ψ), with 95 predicted sites (Krogh et al. 2016; Penzo and Montanaro 2018) (Fig. 1.23). 2’OMe are methyl groups that are added on the oxygen of the C2 of the ribose sugar. Ψ involves the isomerization of uracil into 5-ribosyl uracil. In vertebrates, most small nucleolar ribonuclear RNAs (snoRNAs) guiding these interactions reside in introns of protein coding genes (Maxwell and Fournier 1995; Weinstein and Steitz 1999). Two classes of snoRNAs are involved in specifying the positions of two major rRNA modifications: C/D box snoRNAs (SNORDs) guide the methyltransferase Fibrillarin, resulting in the addition of methyl groups to ribose 2′-OH groups, while H/ACA box snoRNAs (SNORAs) guide Dyskerin, resulting in specific pseudouridylation (Sloan et al. 2017). Knockdown of Fibrillarin (Erales et al. 2017) and knockdown of Dyskerin (Jack et al. 2011) reduce global snoRNP activity for rRNA 2′OMe and Ψ levels, respectively. These studies allude to the importance of rRNA modifications in general. However, it seems that artificial dysregulation of these machineries by chemical or genetic means in cells does not represent a suitable approach for the identification of ribosome functional heterogeneity. Impairment of ribosome biogenesis, and the induction of associated cellular stress responses, preclude interrogation of the functions of 2′OMe or Ψ sites. Other studies allude to snoRNP complexes themselves being the functional agent; acting as sequence-specific RBPs to keep rRNA unfolded during biogenesis (Gulen et al. 2016). Under this model, modification could be an enzymatically necessary secondary effect of targeted Dyskerin and Fibrillarin binding to the rRNA. 43 Taken together, As the nascent pre-rRNA molecule exits RNA Pol I, RNA secondary structure folding starts to take place. The modification at specific rRNA positions is consistent with smooth interaction with the biogenesis machinery. In the context of subunit assembly, the presence of a modified nucleotide is thought to be a means of either delaying folding in a specific region or to prevent misfolding (Klinge and Woolford 2019). Figure 1.23 – Modification and processing of the human 28S rRNA The mods 2’-OMe, Ψ, m1A, m3U, m5C, m6A are shown schematically mapped onto the six color-coded domains (I-VI) of the human 28S rRNA. Root helices forming each domain are indicated as dark blocks. Nucleolar (states A, B, C, F) and nucleus (states I, L) forms of human LSU biogenesis (see Fig. 1.16) are shown to indicate modification and processing. Indicated chemical modifications are shown in grey or red. The positions key ribosome functional centers (PTC, GAC, L1 stalk) and rRNA expansion segments (ES7, ES15, and ES27) are boxed. Adapted from (Vanden Broeck and Klinge 2023). Pre-rRNA processing is concurrent with transcription and modification. rRNA processing consists of a series of cleavages in the presence of RPs and associated factors controlling the time and position of RNA cleavages over the course of biogenesis (Couté et al. 2006). The transcribed pre-rRNA is processed by programmed cleavages, into three rRNAs (18S, 5.8S, and 28S), effectively removing four internal and external sequences (5’ETS, ITS1, ITS2, and 3’ETS) (Fig. 1.24). In yeast, there are at least 170 distinct proteins acting as accessory factors in rRNA processing concurrently during and post ribosome biogenesis (Scull and Schneider 2019). Human ribosome assembly has been shown to include over 200 proteins functioning directly in ribosome subunit biogenesis and processing (Vanden Broeck and Klinge 2023) (Fig. 1.23). Assembly Simultaneously with pre-rRNA transcription from RNA Pol I, modification, cleavage, and interaction with biogenesis machineries, each nascent subunit must incorporate RPs. The assembly and export of the pre-40S ribosomal subunit precedes the assembly of the pre- 60S subunit (Peña et al. 2017). Ribosomal stoichiometry is determined by two mechanisms. One type of mechanism, exemplified by the 5S RNP, involves binding two ribosomal proteins to one assembly factor that simultaneously associates both RPs to 44 two target rRNA sequences. The RP components of the 5S RNP, uL18 and uL5, must first bind to Symportin1 to allow the RNP to integrate into the LSU. The other mechanism involves the formation of RP assemblies that are incorporated into a pre-ribosomal subunit (Peña et al. 2017). While much of ribosome assembly and maturation occurs in the nucleolus, in the nucleoplasm, pre-ribosomal particles are further remodeled and coupled to other proteins to become ready for cytoplasmic export (Brombin et al. 2015; Woolford and Baserga 2013) (Fig. 1.24). In the cytoplasm, pre-ribosomal subunits undergo the last modification steps, including release of 60S assembly factors, cleavage of 18S rRNA, and activation of the now mature 40S and 60S subunits (Warren 2018). A B Figure 1.24 – Assembly and processing of rRNA into ribosomes A) a schematic of the rDNA products mature rRNA, processed from the 47S pre-rRNA transcript. A characteristic Miller chromatin spread shows nascent ribosomes along the rRNA transcript. B) An 9800x electron microscope image of a Miller spread generated from the oocyte of a water beetle, Dytiscus marginalis. An rDNA ring with a contour length of 35 µm is shown with five actively transcribing repeats. Scale bar indicates 1 µm. Adapted from (Shaw 2015; Scheer 1987). Cytoplasmic maturation is a quality control step The SBDS protein is required for late cytoplasmic maturation of 60S ribosomal subunits and translational activation of ribosomes (Finch et al. 2011; Menne et al. 2007; Wong et al. 2011). In mammalian cells, SBDS and elongation factor like 1 (EFL1) catalyze the removal of the assembly factor eukaryotic initiation factor 6 (eIF6) from late cytoplasmic pre-60S ribosomal subunits (Finch et al. 2011; Warren 2018). Linkage analysis of families with SDS revealed a disease-associated interval at 7q11 (Goobie et al. 2001; Popovic et al. 2002). Causal mutations in the gene Shwachman- Bodian-Diamond syndrome (SBDS) were identified in a disease-linked region (Boocock et al. 2003). 90% of families with Shwachman-Diamond syndrome (SDS), a rare congenital disorder characterized by low white blood cells, poor growth, and non-specific skeletal abnormalities, have mutations in the SBDS gene. Loss of Sbds in mice leads to early embryonic lethality (Zhang et al. 2006) while knockdown of sbds in zebrafish recapitulates the human phenotype: exocrine pancreatic insufficiency, neutropenia, and variable skeletal defects (Provost et al. 2012; Oyarbide et al. 2020). 45 Localization of the membrane ribosome Ribosome accessory factors and cellular complexes can direct the localization of the ribosome to specific places within the cell. By binding to the translational machinery, such factors can temporarily change the composition of the complex and thus temporarily modify the function of the ribosome. For example, membrane-bound proteins, like those on the surface of cells must be produced by ribosomes physically adjacent to the endoplasmic reticulum (ER). For this physical translocation of the ribosome to occur, the translating mRNA/ribosome complex must first be localized to the ER. The signal recognition particle (SRP) is a ribonucleoprotein responsible for this function. The SRP recognizes the ER signals sequence near the N-terminus of an integral membrane protein or secretory protein that is actively being synthesized. Translation by the first ribosome on a mRNA molecule is stalled after the signal sequence is synthesized. Direct binding of the SRP to the signal sequence peptide and indirect binding to the ribosome stalls protein synthesis, after which the whole complex is guided to the ER. Binding to the SRP receptor in the ER membrane releases the SRP from the complex and translation resumes by synthesizing the protein through the ER membrane via a protein translocon (Akopian et al. 2013; Keenan et al. 2001; Lütcke 1995; Walter and Blobel 1980). The SRP consists of several proteins and a small non-coding RNA; SRP- RNA (also known as 7SL RNA), which is produced by RNA Pol III (Leung and Brown 2010; Walter and Blobel 1982). This SRP-RNA forms the backbone of the SRP, is about 300 nucleotides long, and has a conserved secondary structure with a small (Alu) domain and a large (S) domain (Siegel and Walter 1986). The Alu domain is responsible for translation arrest while the S domain recognizes the signal sequence and also binds to the SRP receptor (Lütcke 1995; Wild et al. 2019). The subcellular distributions of mRNA transcripts between cytosol and ER compartments are different (Reid and Nicchitta 2012, 2015; Schwartz and Blobel 2003; Voigt et al. 2017). Several ER membrane proteins, like SEC61β (a subunit of the Sec61 translocon) and LRRC59, are associated with ER-bound ribosomes. The mRNAs translated by SEC61β- or LRRC59-labeled ribosomes reveal commonly shared and some specifically enriched transcripts, indicating the contribution of these accessory factors to ribosome functional heterogeneity effecting gene expression (Hoffman et al. 2019). PKM2 was confirmed to be yet another ribosome accessory factor in the ribosome-interactome (Simsek et al. 2017) and is of particular interest to membrane translation. In mESCs, PKM2 is enriched on ER-associated ribosomes and is thought to directly binds ER-associated mRNAs to facilitate their translation. Taken together, these findings support the ability of ribosome associated factors to bind a localized subset of ribosomes or alter the subcellular location of ribosomes. The differently localized mRNAs in these compartments now interact with this specific subset of ribosomes. It is unclear how much the layer of spatial-specificity influences the potential layer of sequence-specificity thought to drive the preferential translation of mRNAs by specific subsets of ribosomes. 46 5S RNP and MDM2 regulate Tp53 A link between ribosome biogenesis and Tp53 was first established more than 20 years ago with immunoprecipitations revealing an interaction between RPL5/uL11, MDM2, and Tp53 (Marechal et al. 1994). Later, it became clear that RPL5/uL11 and RPL11/uL18 were both binding partners of MDM2, capable of suppressing MDM2’s E2 ligase activity and promoting the stabilization of Tp53 (Lohrum et al. 2003; Russo and Russo 2017; Zhang and Lu 2009). Further, it has been shown that the 5S rRNA also directly contacts MDM2 (Sloan et al. 2013). Interaction between more MDM2 became associated with 5S rRNA during ribosome biogenesis stress. More recently, the structures of the 5S RNP (Castillo Duque de Estrada et al. 2023; Pelava et al. 2016) have shown MDM2 binds RPL11/uL18 in the same region RPL11/uL18 uses to bind the 28S rRNA, containing an acidic domain and two zinc fingers. These data together demonstrate that all three components of the 5S RNP directly bind to MDM2 to coordinate Tp53 stabilization. Ribosome biogenesis and cell cycle progression RNA Pol I-mediated transcription levels oscillate during cell cycle progression. In particular, Cdk/cyclin complexes couple ribosome biogenesis regulation with cell cycle progression. Transcription rate reaches its maximum during S and G2 phases and decreases during M phase. During G1, rRNA transcription slowly recovers (Klein and Grummt 1999). rRNA transcription fluctuations during cell cycle progression are generated by Cdk/cyclin-dependent phosphorylation of both UBF and TIF-IB/SL1. In brief, during mitosis, RNA Pol I-dependent transcription silencing arises via Cdk1/cyclinB phosphorylation of TATA box binding protein associated factor (TAF), impairing the interaction of TIF-IB/SL1 with UBF (Heix et al. 1998; Kuhn et al. 1998). At the end of mitosis, dephosphorylation of TAF activates SL1 and relieves mitotic repression of rRNA transcription. On the other hand, the quality of ribosomes can limit cell cycle progression. It has been shown that the translation of cyclin E is specifically impaired upon ribosomal protein loss of function or rRNA haploinsufficiency. (Derenzini et al. 2005) Therefore, cells lacking a sufficient concentration of ribosomal components fail to express cyclin E, leading to a block in G1/S transition and stopping proliferation. Intriguingly, these proliferation-blocked cells do not stop growth (Volarevic et al. 2000). The rate of cell proliferation and growth has been shown to be directly proportional to the rate of ribosomal translation (Baxter and Stanners 1978; Johnson et al. 1975). In order to meet translational demands, proportional RNA Pol I and RNA Pol III transcription must be maintained for ribosome biogenesis, 5S rRNA transcription, and tRNA production. Various signaling pathways are involved in order to fine-tune RNA Pol I transcription in response to growth factor and cellular stress. The output of RNA Pol I transcription can be regulated in two ways: the copy number of active rDNA, set by epigenetic regulators of the rDNA, the rate of transcription across an rDNA, set by chromatin remodelers, polymerase co- factors, and rRNA folding dynamics (Grummt and Längst 2013; Russell and Zomerdijk 2005). 47 Ribosome heterogeneity arises during biogenesis Ribosome biogenesis must correctly incorporate and position all components of the ribosome structure – 80 RPs, 4 rRNAs, modification sites, processing cleavages, and maturation events. Fluctuations during biogenesis could lead to compositional and structural variation in the nascent ribosome. To achieve a steady-state homogenous population of ribosomes, the cell would need to execute perfect quality control over the lifetime of a ribosome complex, from biogenesis to degradation. This would mean degrading all incompletely assembled ribosomes such that they are non-detectable. In reality, the millions of ribosomes in cells and trillions of ribosomes in organisms are a structurally heterogeneous population. Ribosome compositional heterogeneities may arise from the probability that biogenesis incorporates one of many available paralogous components, or may arise via missing components which cannot be added later. Variation among rDNA genes may encode sequence divergence in 18S, 5.8S, or 28S rRNAs – intrinsically changing the ribosome scaffold. This form of variation is studied in Chapter II. RNA Pol I exclusively transcribes the rDNA gene. The cell coordinates ribosome biogenesis with RNA Pol I dynamic regulation of which and how many rDNA loci are actively transcribed. The relative expression of different rDNA gene variants may introduce differences in the population of rRNAs used for biogenesis; thus, increasing ribosome structural heterogeneity in the population. Structural heterogeneity may also arise from changes to the modification patterns (2′OMe or Ψ) at each nucleotide site. Figure 1.25 – Heterogeneous ribosomes Ribosome biogenesis generates ribosome structural heterogeneity (pink) and/or ribosome compositional heterogeneity (yellow) during incorporation. These, as well as ribosome interactions (blue) may impart ribosome functional heterogeneity to a subset of ribosomes. Specific structures offering advantageous interactions are potential sites for ribosome functionalization. Adapted from (Li and Wang 2020). 48 Another form of heterogeneity may arise from genes encoding ribosomal proteins (RPs) of the small and large subunit (Rps/eS and Rpl/eL). Just like other genes, RP gene mRNAs are transcribed in nucleus and translated into RPs in the cytoplasm. Upon import to the nucleolus, RPs will assemble with pre-rRNA co-transcriptionally. A form of ribosome compositional heterogeneity can arise from the stoichiometry or the substitution of a particular RP with a paralogous isoform. Alternative post-translational modification of Rps/eS and Rpl/eL proteins in the cytoplasm may lead to changes in cellular localization or distinct incorporation kinetics into the ribosome. Nomenclature for heterogeneous ribosomes An attempt was made to maintain these definitions of heterogeneous ribosomes: o Ribosome structural heterogeneity – to refer to primary sequence, secondary structure, and tertiary interactions among nucleotides and amino acids constructed into the structure of either subunit o Ribosome compositional heterogeneity – to refer to the number and identity of gene products in the ribosome: rRNA transcripts, RPs, and accessory factors o Ribosome functional heterogeneity – to refer to a measurable difference in ribosome activity due to a specific compositional or structural heterogeneity; often stability, likely localization, feasibly translational, otherwise unknown o Ribosome functionalization – to refer to the evolutionary consequence of a ribosome functional heterogeneity providing the organism (or cell) with an advantage against selective pressures; referred to as “under selection” Relevantly, the separation of these terms indicates that it is possible that heterogeneous ribosome populations might exist within a cell or within an organism, but may not necessarily exhibit distinguishable functions. Additionally, it is clear from the literature, that the evidence for ribosome structural and compositional heterogeneity is much stronger than the evidence for functional heterogeneity, with only a handful of cases supporting a physiologically advantageous role for such ribosome functionalization. Focusing on rDNA gene variation A genome contains hundreds of rDNA genes to allow for the massively parallel synthesis of rRNAs, with each repeat acting as a platform for ribosome biogenesis. Each instance of the rDNA gene in a NOR contains the sequence architecture necessary for stimulating recombination among other rDNA genes, leading to expansion or contraction of array. Regulation of rDNA gene expression increases the chances of stimulating double- stranded breaks in the DNA (DSBs) (Kobayashi et al. 1998). This feature is enriched in rDNA and leads to an important feature regarding the rDNA gene. Some DSBs of rDNA are repaired in the nucleolus by non-homologous end-joining (NHEJ). These repairs may be mutagenic at the repair site by introducing erroneous nucleotides in the rDNA gene sequence (Harding et al. 2015). Other DSBs are extruded from the nucleolus for homology-directed repair (HDR). These repairs are often non-mutagenic at the site of the 49 break and instead may generate recombinogenic errors altering the copy number of rDNA genes. Searching for a homologous sequence to use as a repair template, will identify many possible instances in the NOR and repair template choice may increase or decrease the copy number of rDNA genes (Sluis and McStay 2015). Variation of rDNA sequences within a genome (intragenomic variation) is understudied due to this distinctive volatility observed in rDNA genes (Bughio and Maggert 2019; Kobayashi 2014). This instability is so profound that the locus is often omitted from chromosome assemblies and is actively excluded from genomic research, like the human genome project (Salim and Gerton 2019), which involves intergenomic variation among organisms. The multiplicity, variability, and repetitive nature of rDNA genes obstructs detailed examination. The zebrafish rDNA gene variant studied in this Thesis was not present in the assembled zebrafish genome (through GRCz10) due to bioinformatic difficulties associated with assembling sequences near repetitive elements, like NORs and telomeres. The variant NOR was included in the 2017 version of the genome build (GRCz11). rRNA sequence variation in human genomes The human genome contains somewhere around 200 and 600 copies of rDNA (Gibbons et al. 2014; Parks et al. 2018). Sequence analysis of these regions is challenging due to the high number of repeats (Treangen and Salzberg 2011). Research into human genome sequences demonstrate the existence of rRNA variants (Tseng et al. 2008) and long-read sequencing has observed eleven new 47S rRNA sequences with 76 insertion-deletions (indels) and 25 single nucleotide variants (SNVs) (Kim et al. 2018). Sequence variations were found among the processed regions of ETS and ITS RNAs, as well as in 18S and 28S mature rRNAs. No variation has been detected in the human 5.8S rRNA. The majority of sequence variation contained in the 28S rRNA is located within ES27L, which coordinates communication between the mRNA tunnel and the peptide exit tunnel (Fujii et al. 2018). Expansion segments, like ES27L, are generally located on the surface of the ribosome and may have specific roles in translational control (Xue and Barna 2012). The human 28S variants display a tissue-specific expression, suggesting a precise function towards translation regulation. Other variants induce the formation of enlarged rRNA stem-loop structures resulting in novel interactions between RPs or among rRNA structures (Anger et al. 2013; Kim et al. 2018). Bioinformatic analysis of variants located at rRNA processing sites suggest variability in the processing efficiency of rRNA maturation (Kim et al. 2018). Sequence differences in rDNA sequence have been found among different units belonging to a single NOR (Caburet et al. 2005; Kuo et al. 1996), between different NORs (Kuo et al. 1996; Smirnov et al. 2006), among different cells (Gonzalez and Sylvester 2001), different tissues (Kuo et al. 1996; Parks et al. 2018; Smirnov et al. 2006) and among different human individuals in populations (Gibbons et al. 2014; Parks et al. 2018; Stults et al. 2008). It is valuable to note that no experiment has shown if these modified interactions impact the translation function of the ribosome. Recent work looking into the contribution of rRNA heterogeneity in mature ribosomes. Both intra- and interindividual nucleotide variations in 5S, 5.8S, 18S, and 28S rRNA 50 sequences were identified in humans and mice, as well as tissue-specific rRNA variant expression in mice (Parks et al. 2018). These rRNA nucleotide variants were found in actively translating ribosomes and mapped to RP binding sites and near the ribosome’s decoding center, suggesting that these rRNA variants could potentially have functional roles within the ribosome (Parks et al. 2018). rDNA sequence variation in other organisms In a handful of cases, rDNA variants have been shown to correlate expression with developmental stage of the organism. For example, during the life cycle of the malaria- causing parasite Plasmodium falciparum, specific rRNA variants are expressed between insect and mammalian hosts (Gunderson et al. 1987; Waters et al. 1997). The best studied system involving cell type-specific regulation of ribosomal gene expression is the African Clawed Frog, Xenopus laevis, which utilizes a dual 5S rRNA system. It has been well-documented that this amphibian expresses two variants of the 5S rRNA gene, aptly named oocyte-type and somatic-type (Brown and Gurdon 1977; Peterson et al. 1980; Wegnez et al. 1972). These two types differ at six nucleotide positions (Denis and Wegnez 1977), and as their names suggest, are express in different cell types during development. Roughly 400 copies of somatic-type 5S rRNA gene are expressed during oogenesis and again onwards after late embryogenesis. In contrast, the oocyte-type 5S rRNA gene has roughly 20,000 copies specifically transcribed in developing oocytes, scarcely during early embryogenesis, and then switched off completely in later stages (Denis and Wegnez 1977; Guinta et al. 1986; Wormington et al. 1983). The two 5S rRNA variants differentially bind to TFIIIA and ribosomal protein L5 (Rpl5/uL11). This difference between the two types is thought to be critical for proper oogenesis, during which, an excess of oocyte-type 5S rRNA will be stored in the cytoplasm for regulatory purposes while somatic-type 5S rRNA will bind Rpl5/uL11 and be rapidly integrated into the ribosome as the 5S RNP. Oocyte-type and somatic-type 5S, genes have also been observed a similar genomic organization in other species (Komiya et al. 1986). A literature search for indications of dual 5S rRNA systems reveals an enrichment for various species of fish, e.g. Mediterranean grey mullets, tilapia, leporinus, sturgeon, and loach (Ferreira et al. 2007; Gornung et al. 2007; Martins and Wasko 2006; Martins et al. 2002; Mashkova et al. 1981; Robles et al. 2005). 51 Ribosome specificity and the mRNA pool Ribosomes must be made in a manner that supports translation of mRNAs into proteins and capable of generating the phenotypic complexity seen in the molecular-cellular makeup of the organism. These two requirements for ribosomes are at the crux of many open hypotheses regarding molecular models of gene expression. Proteome makeup of a cell is a determinant of its identity and behavior, aka phenotype. This is dynamically regulated by translational controls of gene expression. The ribosome has often been hypothesized to exhibit specific translational functions on certain mRNAs. Ribosome structure and function in gene expression An early hypothesis involving the ribosome’s role in translation suggested a far less modular function than we know it be today. The hypothesis suggested that any given ribosome was built specifically for the expression of a single gene. Thus, during translation initiation, the mRNA would need to be recognized by a unique ribosome, wholly composed to be specific to one mRNA (Crick 1958). This hypothesis was refuted via experimentation with a bacteriophage infection of an E. coli culture where it was discovered that bacteriophage RNA was expressed into protein without any new ribosome synthesis (Brenner et al. 1961). Finally, Brenner and colleagues concluded that ribosomes were passive structures with no regulatory function that each have the inherent capacity of translating mRNAs. A summary of the history regarding ribosome structural heterogeneity is provided by Genuth and Barna: “the field vacillated from the most extreme view of ribosome specialization to the most extreme view of ribosome homogeneity” (Genuth and Barna 2018). Ribosomes were considered to be homogenous complexes which could not discriminate the identity of mRNAs prior to protein synthesis. In the last 30-40 years, detailed biochemical and genetic observations produced data which required modifying this somewhat dogmatic view of ribosome structural homogeneity. Studies conducted with mutated E. coli strains lacking ribosome proteins identified 17 viable RP-mutants (Dabbs 1986). This early study underscores that not all RPs are essential for cell survival and therefore ribosomes need not be homogenous structures to function in translation. At the same time, Gunderson et al. discovered structurally distinct rRNA variants in the Plasmodium berghei life cycle, finding that each variant was developmentally regulated during specific stages of the host mosquito’s life cycle (Gunderson et al. 1987). Similarly, developmental stage-specific RPs were characterized in the amoeba Dictyostelim discoideum, indicating 12 upregulated RPs and 18 downregulated RPs during the transition from unicellular to pluricellular phases (Ramagopal 1990). Since then, many studies in several organisms have indicated a varying degree of phenotypic penetrance induced by loss-of-function RP alleles (Polymenis 2020). Early research utilizing the Drosophila Minute mutants revealed developmental impairments, some of which are tissue specific, such as defective wing development or cardiomyopathy. The cell and/or tissue specific effects have been similarly seen in yeast (Komili et al. 2007; Parenteau et al. 2011), zebrafish (Amsterdam et al. 2004; Lai et al. 2009; Uechi et al. 2008), mouse (Barlow et al. 2010; Barna et al. 2008; Kondrashov et al. 52 2011; Perucho et al. 2014; Wilson-Edell et al. 2014), and human (Belin et al. 2009; Bolze et al. 2013; De Keersmaecker et al. 2013; Marcel et al. 2013; Rao et al. 2012). These conclusions also extend to other components of the translation machinery, particularly rRNA and ribosome associated factors; thus, challenging the assumptions of a standardized homogenous ribosome. Further research investigating ribosome- associated proteins will need to be undertaken before an appropriate mapping can be made among components, their functions, and the responses they elicit. Tissue specific phenotypes observed in diseases caused by the deregulation of ribosomes, known as ribosomopathies, offer a puzzling challenge to models of translation control (Farley- Barnes et al. 2019; Li and Wang 2020). Ribosome function and the expression loss of a specific mRNA Open hypotheses regarding ribosome heterogeneity involve whether, and if so, how, specific structures in translation machinery regulate translation and ultimately effect expression of cellular identity. In other words, how can perturbations to the general machinery lead to specific effects? There are currently two opposing theories. The first theory, referred to as the ribosome specialization model, considers the original (and now renewed) view in which the ribosome directly enacts a regulatory function on mRNAs during gene expression. That the mutation may abrogate a specific mRNA’s expression via the loss of translation output, imparted specifically to a subset of available ribosomes. This model relies on ribosome structural heterogeneity among cells as the variability causing the observed tissue-specific effects. The model predicts observed ribosome heterogeneities are a consequence of the selective advantage imparted by a ribosome functionalization; the subset of functionalized ribosomes adapted to initiating on and translating a subset of mRNAs (Fig. 1.26). Figure 1.26 – The ribosome filter hypothesis Interactions between an mRNA and specific ribosome components are schematized. The hypothesis predicts, subpopulations of ribosomes (denoted as A and B) displaying ribosome functional heterogeneities can influence translation in an mRNA-sequence specific manner. Thus, giving each population of ribosome the ability to “select” a subset of mRNA targets with which to initiate (denoted as kAi and kBi). 53 This may allow for ribosomes to be adapted to the cellular context, functionalized, to influence gene expression. There is growing support of the idea that ribosomes can regulate protein expression by selectively translating specific mRNAs (Ferretti and Karbstein 2019; Mauro and Matsuda 2016; Miller et al. 2023). According to the “ribosome filter hypothesis”, interactions between an mRNA and specific ribosome structural heterogeneities can influence initiation, giving the ribosome a functionalized filter which mRNA is translated in a specific cell or tissue (Gilbert 2011; Mauro and Edelman 2002). The ribosome-specific mRNA-specific initiation rates are different between subsets of ribosomes due to specific ribosome heterogeneities (Fig. 1.26). These differences can be classified into 3 categories: RP heterogeneity, rRNA heterogeneity, and ribosome associated factor heterogeneity. In the last decade, this model has gained momentum likely due to the vastly improved ability to biochemically measure these forms of heterogeneity (Ferretti and Karbstein 2019; Haag and Dinman 2019; Xue and Barna 2012). The kinds of ribosome structural heterogeneity required for this form of ribosome functionalization is now frequently measured (Gay et al. 2022; Genuth and Barna 2018; Kondrashov et al. 2011; Shi et al. 2017; Simsek et al. 2017). Intriguingly, bioinformatic analyses have shown conservation between 18S rRNA and UTRs of mRNAs supporting a complementarity-based mechanism of mRNA binding to rRNA (Mauro and Matsuda 2016; Pánek et al. 2013). Similar to RNA-binding proteins, interactions with 5’ UTRs of mRNAs may favor or inhibit translation at various steps: 43S initiation complex recognition of the mRNA, scanning of the 5’ UTR by the 48S, or 60S joining (Mauro and Edelman 2007). As mechanisms have yet to be elucidated, whether differences in ribosome-specific mRNA-specific protein synthesis involve complementary rRNA-mRNA sequence interaction is an open question. Importantly, while altered selectivity of ribosomes for mRNAs is a commonly considered functional difference among heterogeneous ribosomes, other translational activities of ribosomes could also be affected. A critical example of ribosome functional heterogeneity relies on the presence of Asc1/RACK1 in the small subunit (Ikeuchi and Inada 2016); responsible for no-go-decay, a ribosome-mediated mRNA quality control mechanism (Buskirk and Green 2017; Graille and Séraphin 2012; Joazeiro 2017; Simms et al. 2017). 54 Figure 1.27 – The ribosome concentration hypothesis Protein synthesis simulations schematizing how mRNAs with low initiation rates are nonlinearly affected by the inhibition of translation. Comparing the three example mRNAs, it is apparent that the magenta target will experience a greater loss in synthesis rate than mRNAs of higher initiation rates (yellow and green). Where wildtype animals (left) may exhibit a stable ribosome concentration for gene expression through development, mutant animals (right) may exhibit a loss of ribosome concentration differently among cells. The second theory, designated the ribosome concentration model, is built around one of the major findings from early measurements of the ribosome’s polymerization kinetics: losses in ribosome concentration preferentially effects protein synthesis of mRNAs with low initiation rates (Lodish 1974; MacDonald and Gibbs 1969). That the mutation may abrogate a specific mRNA’s expression via the loss of translational output, imparted generally by the population of available ribosomes. This model relies on the limited availability of initiating ribosome subunits among cells as the variability causing the observed tissue-specific effects (Kirby et al. 2015; Ludwig et al. 2014). The model predicts ribosome heterogeneities are a consequence of altered ribosome biogenesis; the accompanying loss of ribosome subunits preferentially abrogates protein synthesis from mRNAs with low initiation rates (Mills and Green 2017) (Fig. 1.27). In a consequential 2014 study on Diamond Blackfan anemia, a disease caused by altered ribosomes, it was demonstrated that RPS19/eS19 mutations lower the total level of ribosomes in a cell, suggesting that the observed decrease in GATA1 protein level is brought about by a reduced number of initiation complexes recognizing the highly structured 5’ UTR of Gata1, and not due to a specificity of RPS19-containing ribosomes to translate Gata1 mRNA (Ludwig et al. 2014). Similarly, in a pivotal 2017 review, Mills and Green suggest the broad variety of symptoms observed in RPL38/eL38 mutant mice, as well as many other ribosomopathy mutant phenotypes, could be due to a general decrease in ribosome number rather than a change in specific translation by specific ribosomes (Mills and Green 2017). Both hypotheses are schematized (Fig. 1.28) and are summarized here. The ribosome specialization hypothesis proposes that ribosomes “influence or filter the translation of various mRNAs” (Mauro and Edelman 2007). More specifically, this model proposes that heterogeneous ribosomes prefer to interact with specific mRNAs and that there are competitive interactions between mRNA sequences for binding to rRNA and/or ribosomal 55 proteins. Further, it suggests that this filter can be modulated by changing or blocking specific sites on the ribosome structure (Barna et al. 2022). By contrast, the ribosome concentration hypothesis posits that the overall number of functional ribosomes within a cell greatly influences the differential translation of specific mRNAs based on their mRNA initiation rates (Mills and Green 2017). Figure 1.28 – Two hypotheses regarding ribosome heterogeneity and translational control Left, ribosomes A and B are depicted to have distinct mRNA initiation rates (kAi and kBi). For the magenta gene, A initiates more than B. In wildtype (RWildtype), the combination of ribosomes translates adequate levels of protein (QWildtype), represented as filled circles. In mutants (RMutant, red), with lower levels of ribosome B, the B-specific lower initiation rate will cause a lower protein synthesis rate and result in deficient translation of the magenta protein (QMutant, red), represented as a lighter color. Right, in mutant or inhibited contexts, the number of subunits able to form ribosomes is lowered (RMutant, red). While mRNA initiation rates remain the same, the lower initiation rate of the magenta gene will cause a lower protein synthesis rate and result in deficient translation of the magenta protein (QMutant, red), represented as a lighter color. 56 Germ cell genomes are where evolution occurs The germline refers to the set of parental genetic information passed from one generation to the next. Only mutations in the germline genome can be transmitted to future generations and are the ultimate source of variation for all evolutionary processes. Germline stem cells (GSCs), labeled as primordial germ cells (PGCs) in an embryo, or spermatogonial and oogonial stem cells in juveniles, are precursors of subsequent gametes, egg, and sperm in adults. Like many other animals, fish, have two major cell lineages, namely the germline and soma. The germ-soma separation is one of the earliest events of embryonic development. These highly specialized cells originate at a species- specific area in the embryo and migrate to the developing gonadal ridges during embryonic development (Li et al. 2016; Linhartova et al. 2014, 2014; Presslauer et al. 2012; Saito et al. 2006). At this location, they undergo gametogenesis and eventual cellular differentiation into mature gametes, either eggs or sperm. Germ cell fate in metazoans can be either oocyte-inherited (predetermined) (Eddy 1975; Williamson and Lehmann 1996) or zygotically triggered (induced) (Lawson et al. 1999; Ying and Zhao 2001). In mammals, germ cells are generated during gastrulation in response to extracellular signals from the surrounding embryonic cells (Ying et al. 2001). On the other hand, many model organisms, such as C. elegans, D. melanogaster, X. laevis, and D. rerio, require maternal transmission of germ-cell-specific factors (germplasm) and their distribution into PGCs (Eddy 1975; Seydoux and Braun 2006). Sex-specific chromosomes and genomic regions with sex-specific function are enriched for gene duplications, mutagenic dsDNA breaks, and stage-dependent transcriptional silencing which contribute to the complexity of studying molecular functions of genes operating in the germline. (Hodson and Ross 2021; Pennell et al. 2023) Selection of germline variants allows for adaptation and engineering The germline genome consists of the genetic material present in germinal stem cells, set aside in early development. These genomes, used in meiosis, are passed to the next generation via eggs and sperm. The germline is a fascinating context for investigating the consequences of molecular heterogeneity on differentiation and cell fate. As embryonic germ cells establish the gametes, the specific cell type’s population dynamics and fate can greatly influence inheritance, and therefore the identity of the germline genome. While a variation in somatic function can provide the advantage for an organism to outcompete other organisms, a variation in germ cell functionality can provide the advantage to a cell and become the means of natural selection among germ cells. Optimization of gene expression may occur when translational control is vital for a cell’s competitive advantage and survival. Should this variation fall in the rRNA components of the ribosome a hypothetical sequence-specific regulation of translation may provide such an advantage. The fact that germ cells genomes are the only sequences that gets passed to the next generation is useful for genomic engineering technologies like CRISPR/Cas9 and transposable elements. Synthetic biology techniques allow for the manipulation of the genome to recombinantly express genetic tools useful for the biochemical, genetic, and 57 molecular research of interest. DNA transposons are genetic elements that move or transpose by a ‘cut and paste’ mechanism and have been extensively studied in plants and invertebrates. The use of transposons as a genetic tool in vertebrates started with the application of the Sleeping Beauty transposon system in which the transposase is provided in trans and the key DNA cargo is flanked by transposon end sequences (Ivics et al. 1997). This transposon paradigm remains the primary approach employed today with 10 transposons from four different superfamilies (TcI/Mariner, hAT, PIF/Harbinger, piggyBac) available for use in vertebrates (Ni et al. 2008). Tol2 and Sleeping Beauty transposon-based methods show efficient germline transmission in zebrafish (Davidson et al. 2003; Kawakami et al. 2000). Also, Tol2-mediated transgenes may not be subject to gene silencing effects since their expression persists through generations (Kawakami 2007). For this reason, as well as ease of cloning, Tol2-based plasmids are used to generate transgenic lines (Kwan et al. 2007). Germplasm and germ cell formation Germline determinant material, aka germplasm, is often described as dense particles, mitochondrial clusters, and “chromatoid” or electron-opaque substances found in positions where putative PGCs are formed. PGC development was initially identified by light and electron microscopy for several species of fish, including medaka, goldfish, guppy, trout, and zebrafish (Bruslé and Bruslé 1978, 1978; Eddy 1975; Johnston 1951; Satoh 1974) . PGCs are detectable during early and mid-gastrula stages within ectoderm and mesoderm. PGCs have large nuclei, possess prominent nucleoli, distinct nuclear membranes, and a relatively large volume of cytoplasm compared to other cell types (Johnston 1951). While characterization of zebrafish germplasm would not take place until much later, early microscopy data from research regarding yolk formation in oocytes includes these structures (Korfsmeier 1966). Along with mitochondria, maternally provided germplasm contains RNA and proteins necessary for PGC formation, migration, and development (Hashimoto et al. 2004; Herpin et al. 2007; Yoon et al. 1997). Germplasm molecular markers include genes known as vasa (vas), dead end (dnd), nanos (nos), bucky ball (buc), and dazl (daz). In zebrafish, the germplasm is formed from two components. The first is composed of mRNAs such as dnd, nanos3, and vasa, initially present in the animal pole of mature oocytes, and the second is composed of components including dazl RNA and Buc protein, vegetally localized during oogenesis and then recruited to the germ plasm during early cleavage (Theusch et al. 2006). Analyses of the localization patterns of dnd, nos1, vas, buc, and dazl RNAs during the first cell cycle have shown germplasm formation at the furrows of the first and second cleavage divisions where putative PGCs are formed. (Hashimoto et al. 2004; Herpin et al. 2007; Nagasawa et al. 2013; Presslauer et al. 2012; Theusch et al. 2006; Wang et al. 2015). The initial assembly of germplasm is actin dependent, whereas the later segregation and distal aggregation of germplasm requires myosin activity to remodel the so-called furrow microtubule array (Knaut et al. 2000; Theusch et al. 2006; Urven et al. 2006). Germplasm separates into four subcellular aggregates by the 32-cell stage, 58 eventually leading to four individual cells being specified as PGCs at the 1000-cell stage at 3 hours post-fertilization (Knaut et al. 2000) (Fig. 1.29). Figure 1.29 – Stages of germplasm localization in zebrafish Beginning from the top left and proceeding clockwise, early control of germplasm localization is entirely dependent on maternal products (pink) whereas activation of the zygotic genome is coincident with the now-specified four PGCs (yellow stripes). In the 1-cell stage embryo, germplasm becomes aggregated in the cleavage furrows between blastomeres. In high stage embryos, the germplasm is inherited into four cells. In shield stage embryos, PGCs are dividing and migrating in the gastrula. By 30 hours post- fertilization, PGCs have reached the prospective gonadal ridge and will reside here until sexual differentiation. In adult females, germplasm is generated and again becomes localized to the Balbiani body of stage I oocytes. By stage II of oogenesis, germplasm materials are transported to the vegetal cortex of the oocyte. In stages III to V, yolk formation grows the cell. Upon ovulation and egg activation, germplasm moves from vegetal regions to the animal pole. Adapted from (Kaufman and Marlow 2016). Germ cell specification, migration, and visualization In animal development zygotic transcription is activated in both PGCs and soma. Cells respond to the pre-patterned positions of molecules and migrate to particular regions to generate specific tissue types (Blaser et al. 2005; Kane and Kimmel 1993; Newport and Kirschner 1982; Urven et al. 2006). This period has been described as a transition from synchronous to asynchronous cell cycle events known as the midblastula transition (MBT) Zebrafish experience 8 synchronous cycles, after which the length of S phase increases, G1, and G2 phases intervene between M and S, each cell’s cycling rate is slowed, and synchrony among cells is lost during the next cycle (Kane and Kimmel 1993). PGCs have been shown migrating to the gonadal ridge using fixed and live methods. In situ hybridization (ISH) of germline-specific transcripts using complementary DNA/RNA hybridization of a labeled probe in a fixed embryo allows for imaging highly localized transcripts in PGCs (Saito et al. 2006). A green fluorescent protein (GFP) labeling technique is accomplished transgenically or by injecting synthetic mRNA in vitro transcribed with a GFP sequence fused to the nos3 3’ untranslated region (UTR) into 1- cell stage embryos (Köprunner et al. 2001). 59 Similar to nos3 mRNA, encoding RNA-binding zinc finger proteins, vasa mRNA, encoding a DEAD box RNA helicase, and dnd mRNA, encoding an RNA binding protein, are reliable markers for tracing PGCs during development (Köprunner et al. 2001). While all three of the listed mRNAs are essential for PGC specification, Dnd protein is required for pseudopod formation during the migration of PGCs (Liu et al. 2009; Nagasawa et al. 2013; Wang et al. 2015; Weidinger et al. 1999, 2003). As PGCs require the active translation of dnd mRNA into Dnd protein, injected translation-blocking morpholino-based gene knockdown methodology is commonly used to prevent PGC migration and survival (Ciruna et al. 2002). 60 Germ cell regulation of translation Genetic studies in many model organisms have established our current understanding of how RNA binding proteins (RBPs) and the regulated translation of individual mRNAs controls various aspects of germ cell development and early embryogenesis. Across many species, these proteins are conserved markers of germ cell identity including Nanos, Pumilio, Vasa, and Dazl (Lesch and Page 2012). These RBPs are pivotal for specification of germ cell identity, regulating germ cell differentiation, and preparing germ cells for entry into meiosis. Many excellent reviews describe germ cells, their specification, migration, and propagation in several model organisms (Huggins and Keiper 2020; Jamieson-Lucy and Mullins 2019; Lai and King 2013; Susor et al. 2016; Voronina et al. 2011). Oocytes must delay gene expression Transcription and translation are basic cellular processes that are commonly considered to be same for all cell types; oocytes are the notable exception. Oocytes must maintain their normal cellular metabolism, and at the same time, transcribe large quantities of specific mRNAs for developmentally timed usage later in embryogenesis. Translational control in the developing oocyte is especially complex (Richter and Lasko 2011; Winata and Korzh 2018). Many mRNAs must only be translationally silent during transport to their destinations (in space), while others must be silenced during cell type differentiation and development (in time). The strength of control elements being used by the oocyte to control mRNA-specific initiation rates is thought to be directly correlated with developmental outcomes of the embryo. All animal embryos pass through a stage during which developmental control is handed from maternally provided gene products to those synthesized from the zygotic genome. There are several mechanisms in play to dynamically use maternally deposited materials (Tadros and Lipshitz 2009) until this transition occurs. One of these mRNA silencing mechanisms revolves around the condition that mRNAs have no, or a short, poly(A) tail (Graindorge et al. 2006; Subtelny et al. 2014; Weill et al. 2012). During early embryogenesis polyadenylation and deadenylation of mRNAs occur, all of which contributes to the maternal-to-zygotic transition (MZT) where the maternal mRNAs are replaced by zygotic mRNAs. The MZT has been exceptionally reviewed multiple times (Vastenhouw et al. 2019). Translation heterogeneity in the germline Genetic analyses using loss-of-function alleles are fundamental for mapping an observed function to a gene. If cells require a specific gene for functionality, loss of gene function observed in mutants may force the cell to enter a disease state. Inhibiting genes involved in ribosome biogenesis has detrimental effects on the survivability of most cells. Several RPs have evolved paralogs that exhibit specificity to germline stem cells (GSCs). Ovary- specific or testis-specific RP expressions correlate with a potential mechanism for ribosome functional heterogeneity during gametogenesis (Li and Wang 2020). 61 The ovary houses germline stem cells as they progress through maturation, directly supporting their growth into oocytes. Just as there exists heterogeneity in testis, interrogation into ovary tissues also reveals the potential for ribosome functionalization. Initially for Xenopus, and later on for other organisms, a maternal type 5S rRNA has been described in oocytes (Brown and Gurdon 1977; Guinta et al. 1986; Komiya et al. 1986; Wegnez et al. 1972). Tissue specific expression has been known to be correlated with particular genome organization (Peterson et al. 1980). Although these findings point towards the existence of heterogeneous ribosomes, it was not an intensely researched topic for many decades. Recently, the idea of ribosome functionalization by heterogenous ribosome composition has gained momentum due to the ability to biochemically characterize ribosomes (Xue and Barna 2012; Ferretti and Karbstein 2019; Haag and Dinman 2019). Germ cells also express specific paralogs of broadly used translation factors. For example, the C. elegans genome encodes at least five eIF4E-like genes, the function of which have recently been reviewed (Huggins and Keiper 2020). A number of these eIF4E isoforms play important roles in germline maintenance and development (Huggins et al. 2020). In particular, IFE-1 exhibits enriched expression in germ cells and the protein associates with P granules. Mutations in IFE-1 result in loss of fertility, including both reduced translation of specific maternally deposited mRNAs and defects in sperm development. Mutations in another eIF4E gene, IFE-3, result in defects in growth and germline sex determination. More specifically, the transition from spermatogenesis to oogenesis appears disrupted in IFE-3 mutant hermaphrodites. IFE-3 functions with its binding partner IFET-1 to regulate the translation of several germline sex determination factors, including the masculinizing gene, fem-3. The specificity of IFE-3’s effect on gene expression is mediated, in part, by association with IFET-1. This represents a sequence- specific RBP interaction that changes an mRNA-specific initiation rate. Similarly, the Drosophila genome contains eight eIF4E paralogs (Hernández et al. 2005), some of which exhibit specific enriched expression within gonads – with eIF4E-3 and eIF4G2 being essential in male fertility. The expansion of eIF4F complex members (eIF4E, eIF4A, and eIF4G) and the germline- specific expression of individual ribosome protein paralogs may provide a network of interactions for controlling germ cell-specific mRNA translation in space and time. RP heterogeneity in the drosophila germline; a case study on eL22-like In D. melanogaster RpL22-like/eL22-like has been identified as tissue restricted with the highest levels in the adult male germline (Mageeney et al. 2018) and, four paralogs are specifically expressed in the testis of mammals [RPS4Y2/eS4Y2, RPL22L1/eL22L1, RPL39L/eL39L and RPL10L/uL16L (Lopes et al. 2010; Nadano et al. 2002; Sugihara et al. 2010). The molecular origins of “why” such diverse animals would have evolved testis- specific paralogs of RpL22-like/eL22-like were, and still are, unclear. Of importance to potential ribosome functional heterogeneity, the paralog can replace RpL22/eL22 in the large subunit. RpL22/eL22 sits inside the exit tunnel, beyond the PTC (Fig. 1.15), just opposite the P-site, such that it is the first RP to contact newly polymerized C-termini of polypeptides (Anger et al. 2013). 62 From an evolutionary standpoint, ribosome heterogeneities due to RP gene duplications like RpL22/eL22 and RpL22-like/eL22-like, and ribosomal gene variants like the Xenopus oocyte-type and somatic-type 5S rRNAs, could have arisen by a neutral event. However, in the specific cellular context of the developing oocyte, where the demand for ribosomal gene products can be exceptionally high, this variant can quickly become subject to positive selection. During spermatogenesis, sex chromosomes become transcriptionally repressed and autosome-encoded paralogs are upregulated in a compensatory mechanism (Turner 2015). Given the placement of most testis-specific paralogs on autosomes and the presence of sex-linked-chromosome-encoded counterparts of these paralogs, this form of transcriptional regulation could explain testis-specific expression of RPs – arguing against ribosome functionalization for selective advantage. Looking from a different perspective, specific paralog expression driven by MSCI could provide an ideal context for neofunctionalization of the ribosome. Of note, both RPL22- like/eL22-like (Drosophila) and RPL22L1/eL22L1 (Human) are testis specific. Rescue experiments in genetic knockouts of RPL22 and RPL22-like do suggest that these two paralogs are functionally distinct (Mageeney et al. 2018). Recently, drosophila ribosomes containing either RPL22/eL22 or RPL22-like/eL22-like were shown to preferentially translate different sets of mRNAs (Mageeney and Ware 2019). RPL22/eL22-containing ribosomes were found to associate with mRNAs involved in organ development, while RPL22-like/eL22-like-containing ribosomes preferentially translated mRNAs involved in translation and protein transport across membranes. The mRNAs enriched into RPL22-ribosomes did show a preference for genes encoding proteins known to function in spermatogenesis. High resolution cryo-EM structures of poly-ribosome structures from the drosophila testis reveal no changes to functional regions of the ribosome due to RPL22/RPL22-like structural differences (Hopes et al. 2022) (Fig. 1.14). Whereas the evidence for ribosome structural heterogeneity is apparent, ribosome functionalization by specific RP-paralog incorporation is unclear. First, whether RPL22- like exerts its function from within the ribosome or via extra-ribosomal pathways is unknown. The evidence in favor of swapped RP-paralogs in ribosome composition leading to altered translation in spermatogenesis, or otherwise, requires further experimentation. In zebrafish, both RPL22 paralogs exhibit extra-ribosomal functions in the nucleus regarding specific intronic splicing that are critical to development (Zhang et al. 2013, 2017). Similarly, drosophila RPL22 paralogs may exhibit extra-ribosomal functions important for GSC-specific processes, like maintenance of a male germline niche via asymmetric GSC division or early fate specification of spermatogonia (Herrera and Bach 2018). 63 While RPL22-like mRNA and protein are specifically detected in sperm cell development where RPL22-like-containing ribosomes are specifically generated, it is still possible that ribosomes containing either of the RPL22 paralog-containing ribosomes do not exhibit specific functions advantageous to the organism. Advantageous functionality may arise between the paralog genes due to their genomic loci, the particular translational controls on their mRNAs, or RP interactions during ribosome biogenesis and quality control in GSCs. Modeling the effects of ribosome functional heterogeneity on gene expression can be difficult without knowing the biochemical nature of the distinction. The ribosome filter hypothesis proposes the biochemical nature of the distinction is ribosome-specific mRNA initiation rates. Heterogeneous ribosomes (depicted as yellow and blue) are used to model gene expression differences as a result of ribosome functional heterogeneity via specifc mRNA initiation rates (Fig. 1.30). Figure 1.30 – Ribosome functional heterogeneity via specific mRNA initiation rates The filter hypothesis predicts subpopulations of ribosomes (denoted as A and B in yellow and blue) can influence translation in an mRNA-sequence specific manner. Thus, giving each population of ribosome the ability to “select” a subset of mRNA targets with which to initiate (denoted as kAi and kBi), see (Fig. 1.11). For example, the circles at the bottom of the plot denote the initiation rates of multiple mRNA targets; note the swapped positions of green and magenta to indicate the ribosome-specific mRNA initiation rate. 64 Danio rerio, a specific fish for inquiry Zebrafish, Danio rerio, are a teleost of the cyprinid family in the Actinopterygii (ray-finned fish) class (Nüsslein-Volhard and Dahm 2002), named for their distinctive horizontal stripes. They are tropical freshwater fish that originated in a region at the base of the South-Eastern Himalayan Ridge (Talwar and Jhingran 1991). Relatively small, surprisingly resilient, and robustly fertile, Danio rerio as well as other Danio species were domesticated for ornamental purposes. In 1934, it was suggested by Charles W. Creaser that the pet store fish was favorable for embryological research (Creaser 1934). Later work from George Streisinger and colleagues led the groundwork for the vertebrate model organism that has allowed both the application of biochemical, genetic, and embryological methods (Streisinger et al. 1981). Today, the zebrafish is a well-established vertebrate system for researchers to ask genomic, molecular, cellular, and organismal questions; and to use as a platform for human disease modeling and clinical drug development (Baranasic et al. 2022; Lange et al. 2023; Patton et al. 2021). Development, in brief Upon fertilization of the egg by a sperm, the extra embryonic chorion separates from the zygote and cytoplasm moves to the animal pole to form the blastodisc. Zebrafish eggs are telolecithal, meaning that the majority of the cell is occupied by yolk, which contains nutritive elements. At first, cellular divisions are meroblastic cleavages i.e., occurring only at the animal pole of the embryo, such that all materials are still accessible to each division of the egg. 30 minutes after activation of the egg, a series of signaling events causes the cell to cleave every 15 minutes. These first cleavages of the animal occur synchronously. The newly fertilized single-cell stage zygote will form into a 512-cell blastula in only 2 hours 45 minutes, or 2.75 hours post-fertilization (hpf) (Fig. 1.31). At this stage, the midblastula transition (MBT) begins. This is characterized by activation of the zygotic genome (ZGA), loss of cell synchronicity, lengthening of the cell cycle, and emergence of cell motility. Epiboly follows, with the migration of cells around the yolk sac. Gastrulation begins, and includes specification of three cellular populations of precursor cells (primordia) which will give rise to all somatic tissue: the mesoderm, endoderm, and ectoderm. After 6 hpf involution occurs, by which cells of the future dorsal side migrate underneath surrounding cells, forming the embryonic shield. Bud stage at 10 hpf signifies the end of gastrulation. Next is segmentation and the formation of somites: mesodermal sections of tissues of the future spleen, muscle, and blood. The animal then subdivides the ectodermal neural plate into an axis-regionalized neural tube. By 24 hpf, segmentation completes, primary organs are visible, and most cells are lineage specified. In the next day, primitive hematopoiesis in the intermediate mesoderm generates blood cells and vertebrate-specific delamination of multipotent stem cells from the neural tube ectoderm will generate neural crest cells for further migration and specification. Between 48-72 hpf the embryo hatches into a “larva”. 65 Figure 1.31 – Developmental stages of the zebrafish embryo Images of the various developmental stages of the zebrafish from the 2-cell stage at 0.75 hpf up to prim- 25 stage at 36 hpf imaged using a light microscope. For each image, the accompanying developmental stage name is included on the bottom left and the time in hours post-fertilization (hpf) is included on the bottom right. Embryos were manually hatched for imaging. Adapted from (Kimmel et al. 1995). 66 The peculiar dual rDNA system of interest As previously mentioned, eukaryotic 18S, 5.8S and 28S rRNAs are processed from a single 47S transcript transcribed from the rDNA gene – normally existing in tandem arrays of over hundreds of copies in a genome. In 2017, Locati and Pagano and colleagues reported on two rDNA gene variants in the zebrafish, which transcribes rRNA from NOR clusters at two distinct genomic loci. As it appeared, a chromosome 4 rDNA variant was only expressed in eggs (to be maternally deposited), while the expression of a chromosome 5 rDNA variant was coupled with the ZGA; these are referred to as maternal type and somatic type rDNA genes respectively (Locati et al. 2017b). Zebrafish 5S rRNA genes also exhibit a similar maternal type and somatic type: chromosome 4 variants being expressed in eggs (to be maternally deposited), and expression of chromosome 18 variants are tied with the ZGA (Locati et al. 2017a). The primary sequences of each rRNA vary considerably; differences between maternal and somatic pairs of 5S, 18S, 5.8S, and 28S rRNA coding sequences indicate a ribosome structural heterogeneity and potential ribosome compositional heterogeneity. Both 5S rRNAs are 119 nucleotides showing 92.4% alignment (Locati et al. 2017a). The maternal 18S rRNA is 1939 nucleotides while the somatic 18S rRNA is 1889, with 91.3% alignment (ClustalW). The maternal 28S rRNA is 4270 nucleotides while the somatic 28S rRNA is 4106, these, only 87.0% similar (Locati et al. 2017b). These sequence differences imply that the divergent segments in rRNA display structural differences at different times of development. The rRNAs composing maternally deposited ribosomes are almost 100% transcribed from the maternal rDNA. Maternally deposited ribosomes are inherited by the zygote at fertilization (0 hpf). Detection of somatic type rRNA signifying zygotic rDNA transcription begins around 8-10 hpf at the end of gastrulation, several hours after the mRNA maternal- zygotic-transition. Separately, the Neugebauer lab showed this is precisely the time when embryonic cells generate a canonical nucleolus (Heyn et al. 2017). The somatic type rRNA comprises 0% of the 6 hpf embryo, 95% of the 120 hpf larva, and 100% of all adult somatic tissues. Sequence and secondary structure comparisons reveal somatic rRNAs are more similar to rRNA from other vertebrates (Locati et al. 2017b). This suggests that the maternal rDNA gene is more diverged, with a greater possibility of ribosome functional heterogeneity. 67 Nomenclature for zebrafish ribosomes For consistency with Locati and Pagano and colleagues, the first to report on the zebrafish dual ribosomal system (Locati et al. 2017a, 2017b, 2018): o The rDNA variant found near the q arm telomere of chromosome 4 at 77.5 Mbp (in GRCz11), its mature rRNA products, and the associated ribosome subunit products are referred to as “maternal” o The rDNA variant found near the p arm telomere of chromosome 5 at 00.8 Mbp (in GRCz11), its mature rRNA products, and the associated ribosome subunit products are referred to as “somatic” o The 5S rRNA variant found arrayed on chromosome 4 and its rRNA transcripts are referred to as “maternal” o The 5S rRNA variant found arrayed on chromosome 18 and its rRNA transcripts are referred to as “somatic” In light of our findings presented in Chapter II, this nomenclature ostensibly requires changing and is discussed in Chapter III. 68 References Aitchison JD, Rout MP. 2000. The Road to Ribosomes. J Cell Biol 151: 23–26. Akanuma G, Nanamiya H, Natori Y, Yano K, Suzuki S, Omata S, Ishizuka M, Sekine Y, Kawamura F. 2012. Inactivation of Ribosomal Protein Genes in Bacillus subtilis Reveals Importance of Each Ribosomal Protein for Cell Proliferation and Cell Differentiation. J Bacteriol 194: 6282–6291. Akopian D, Shen K, Zhang X, Shan S. 2013. Signal recognition particle: an essential protein-targeting machine. Annu Rev Biochem 82: 693–721. Alkemar G, Nygård O. 2006. Probing the Secondary Structure of Expansion Segment ES6 in 18S Ribosomal RNA. Biochemistry 45: 8067–8078. Amsterdam A, Sadler KC, Lai K, Farrington S, Bronson RT, Lees JA, Hopkins N. 2004. Many ribosomal protein genes are cancer genes in zebrafish. PLoS Biol 2: E139. Anger AM, Armache J-P, Berninghausen O, Habeck M, Subklewe M, Wilson DN, Beckmann R. 2013. Structures of the human and Drosophila 80S ribosome. Nature 497: 80–85. Arabi A, Wu S, Ridderstråle K, Bierhoff H, Shiue C, Fatyol K, Fahlén S, Hydbring P, Söderberg O, Grummt I, et al. 2005. c-Myc associates with ribosomal DNA and activates RNA polymerase I transcription. Nat Cell Biol 7: 303–310. Arava Y, Wang Y, Storey JD, Liu CL, Brown PO, Herschlag D. 2003. Genome-wide analysis of mRNA translation profiles in Saccharomyces cerevisiae. Proc Natl Acad Sci 100: 3889–3894. Archer SK, Shirokikh NE, Beilharz TH, Preiss T. 2016. Dynamics of ribosome scanning and recycling revealed by translation complex profiling. Nature 535: 570–574. Ares M, Grate L, Pauling MH. 1999. A handful of intron-containing genes produces the lion’s share of yeast mRNA. RNA N Y N 5: 1138–1139. Avni D, Biberman Y, Meyuhas O. 1997. The 5’ terminal oligopyrimidine tract confers translational control on TOP mRNAs in a cell type- and sequence context- dependent manner. Nucleic Acids Res 25: 995–1001. Ban N, Beckmann R, Cate JH, Dinman JD, Dragon F, Ellis SR, Lafontaine DL, Lindahl L, Liljas A, Lipton JM, et al. 2014. A new system for naming ribosomal proteins. Curr Opin Struct Biol 24: 165–169. Baranasic D, Hörtenhuber M, Balwierz PJ, Zehnder T, Mukarram AK, Nepal C, Várnai C, Hadzhiev Y, Jimenez-Gonzalez A, Li N, et al. 2022. Multiomic atlas with functional stratification and developmental dynamics of zebrafish cis-regulatory elements. Nat Genet 54: 1037–1050. Barlow JL, Drynan LF, Trim NL, Erber WN, Warren AJ, McKenzie ANJ. 2010. New insights into 5q- syndrome as a ribosomopathy. Cell Cycle Georget Tex 9: 4286– 4293. Barna M, Karbstein K, Tollervey D, Ruggero D, Brar G, Greer EL, Dinman JD. 2022. The promises and pitfalls of specialized ribosomes. Mol Cell 82: 2179–2184. Barna M, Pusic A, Zollo O, Costa M, Kondrashov N, Rego E, Rao PH, Ruggero D. 2008. Suppression of Myc oncogenic activity by ribosomal protein haploinsufficiency. Nature 456: 971–975. Baxter GC, Stanners CP. 1978. The effect of protein degradation on cellular growth characteristics. J Cell Physiol 96: 139–145. 69 Belin S, Beghin A, Solano-Gonzàlez E, Bezin L, Brunet-Manquat S, Textoris J, Prats A- C, Mertani HC, Dumontet C, Diaz J-J. 2009. Dysregulation of ribosome biogenesis and translational capacity is associated with tumor progression of human breast cancer cells. PloS One 4: e7147. Belin S, Hacot S, Daudignon L, Therizols G, Pourpe S, Mertani HC, Rosa-Calatrava M, Diaz J-J. 2010. Purification of Ribosomes from Human Cell Lines. Curr Protoc Cell Biol 49: 3.40.1-3.40.11. Bell SP, Jantzen HM, Tjian R. 1990. Assembly of alternative multiprotein complexes directs rRNA promoter selectivity. Genes Dev 4: 943–954. Bell SP, Learned RM, Jantzen H-M, Tjian R. 1988. Functional Cooperativity Between Transcription Factors UBF1 and SL1 Mediates Human Ribosomal RNA Synthesis. Science 241: 1192–1197. Ben-Shem A, Garreau de Loubresse N, Melnikov S, Jenner L, Yusupova G, Yusupov M. 2011. The structure of the eukaryotic ribosome at 3.0 Å resolution. Science 334: 1524–1529. Bernier CR, Petrov AS, Kovacs NA, Penev PI, Williams LD. 2018. Translation: The Universal Structural Core of Life. Mol Biol Evol 35: 2065–2076. Biophysical Society., Society B, Roberts RB. 1958. Microsomal particles and protein synthesis; papers presented at the First Symposium of the Biophysical Society, at the Massachusetts Institute of Technology, Cambridge, February 5, 6, and 8, 1958. Published on behalf of the Washington Academy of Sciences, Washington, D.C., by Pergamon Press, New York https://www.biodiversitylibrary.org/bibliography/6261. Blaser H, Eisenbeiss S, Neumann M, Reichman-Fried M, Thisse B, Thisse C, Raz E. 2005. Transition from non-motile behaviour to directed migration during early PGC development in zebrafish. J Cell Sci 118: 4027–4038. Bock LV, Kolář MH, Grubmüller H. 2018. Molecular simulations of the ribosome and associated translation factors. Curr Opin Struct Biol 49: 27–35. Bolze A, Mahlaoui N, Byun M, Turner B, Trede N, Ellis SR, Abhyankar A, Itan Y, Patin E, Brebner S, et al. 2013. Ribosomal protein SA haploinsufficiency in humans with isolated congenital asplenia. Science 340: 976–978. Boocock GRB, Morrison JA, Popovic M, Richards N, Ellis L, Durie PR, Rommens JM. 2003. Mutations in SBDS are associated with Shwachman-Diamond syndrome. Nat Genet 33: 97–101. Brakke MK. 1951. Density Gradient Centrifugation: A New Separation Technique 1. J Am Chem Soc 73: 1847–1848. Brandman O, Hegde RS. 2016. Ribosome-associated protein quality control. Nat Struct Mol Biol 23: 7–15. Brenner S, Jacob F, Meselson M. 1961. An Unstable Intermediate Carrying Information from Genes to Ribosomes for Protein Synthesis. Nature 190: 576–581. Brombin A, Joly J-S, Jamen F. 2015. New tricks for an old dog: ribosome biogenesis contributes to stem cell homeostasis. Curr Opin Genet Dev 34: 61–70. Brown DD, Gurdon JB. 1977. High-fidelity transcription of 5S DNA injected into Xenopus oocytes. Proc Natl Acad Sci U S A 74: 2064–2068. 70 Bruslé S, Bruslé J. 1978. An ultrastructural study of early germ cells in Mugil (Liza) auratus Risso, 1810 (Teleostei : Mugilidae). Ann Biol Anim Biochim Biophys 18: 1141– 1153. Budde A, Grummt I. 1999. p53 represses ribosomal gene transcription. Oncogene 18: 1119–1124. Bughio F, Maggert KA. 2019. The Peculiar Genetics of the Ribosomal DNA Blurs the Boundaries of Transgenerational Epigenetic Inheritance. Chromosome Res Int J Mol Supramol Evol Asp Chromosome Biol 27: 19–30. Buskirk AR, Green R. 2017. Ribosome pausing, arrest and rescue in bacteria and eukaryotes. Philos Trans R Soc Lond B Biol Sci 372: 20160183. Buszczak M, Signer RAJ, Morrison SJ. 2014. Cellular differences in protein synthesis regulate tissue homeostasis. Cell 159: 242–251. Caburet S, Conti C, Schurra C, Lebofsky R, Edelstein SJ, Bensimon A. 2005. Human ribosomal RNA gene arrays display a broad range of palindromic structures. Genome Res 15: 1079–1085. Campbell KJ, White RJ. 2014. MYC Regulation of Cell Growth through Control of Transcription by RNA Polymerases I and III. Cold Spring Harb Perspect Med 4: a018408. Castillo Duque de Estrada NM, Thoms M, Flemming D, Hammaren HM, Buschauer R, Ameismeier M, Baßler J, Beck M, Beckmann R, Hurt E. 2023. Structure of nascent 5S RNPs at the crossroad between ribosome assembly and MDM2-p53 pathways. Nat Struct Mol Biol 30: 1119–1131. Cavanaugh AH, Hirschler-Laszkiewicz I, Hu Q, Dundr M, Smink T, Misteli T, Rothblum LI. 2002. Rrn3 Phosphorylation Is a Regulatory Checkpoint for Ribosome Biogenesis *. J Biol Chem 277: 27423–27432. Cech TR. 2000. Structural biology. The ribosome is a ribozyme. Science 289: 878–879. Chandramouli P, Topf M, Ménétret J-F, Eswar N, Cannone JJ, Gutell RR, Sali A, Akey CW. 2008. Structure of the Mammalian 80S Ribosome at 8.7 Å Resolution. Structure 16: 535–548. Chappell SA, Dresios J, Edelman GM, Mauro VP. 2006. Ribosomal shunting mediated by a translational enhancer element that base pairs to 18S rRNA. Proc Natl Acad Sci U S A 103: 9488–9493. Chen X, Sun Y, Zhang T, Shu L, Roepstorff P, Yang F. 2021. Quantitative Proteomics Using Isobaric Labeling: A Practical Guide. Genomics Proteomics Bioinformatics 19: 689–706. Chew G-L, Pauli A, Schier AF. 2016. Conservation of uORF repressiveness and sequence features in mouse, human and zebrafish. Nat Commun 7: 11663. Chou C-W, Tai L-R, Kirby R, Lee I-F, Lin A. 2010. Importin β3 mediates the nuclear import of human ribosomal protein L7 through its interaction with the multifaceted basic clusters of L7. FEBS Lett 584: 4151–4156. Ciruna B, Weidinger G, Knaut H, Thisse B, Thisse C, Raz E, Schier AF. 2002. Production of maternal-zygotic mutant zebrafish by germ-line replacement. Proc Natl Acad Sci U S A 99: 14919–14924. Clark CG, Tague BW, Ware VC, Gerbi SA. 1984. Xenopus laevis 28S ribosomal RNA: a secondary structure model and its evolutionary and functional implications. Nucleic Acids Res 12: 6197–6220. 71 Clemons WM, May JL, Wimberly BT, McCutcheon JP, Capel MS, Ramakrishnan V. 1999. Structure of a bacterial 30S ribosomal subunit at 5.5 A resolution. Nature 400: 833– 840. Comai L, Tanese N, Tjian R. 1992. The TATA-binding protein and associated factors are integral components of the RNA polymerase I transcription factor, SL1. Cell 68: 965–976. Couté Y, Burgess JA, Diaz J-J, Chichester C, Lisacek F, Greco A, Sanchez J-C. 2006. Deciphering the human nucleolar proteome. Mass Spectrom Rev 25: 215–234. Creaser CW. 1934. The Technic of Handling the Zebra Fish (Brachydanio rerio) for the Production of Eggs Which Are Favorable for Embryological Research and Are Available at Any Specified Time Throughout the Year. Copeia 1934: 159–161. Crick F. 1970. Central Dogma of Molecular Biology. Nature 227: 561–563. Crick FH. 1958. On protein synthesis. Symp Soc Exp Biol 12: 138–163. Crick FHC, Barnett L, Brenner S, Watts-Tobin RJ. 1961. General Nature of the Genetic Code for Proteins. Nature 192: 1227–1232. Crosby MA, Gramates LS, dos Santos G, Matthews BB, St. Pierre SE, Zhou P, Schroeder AJ, Falls K, Emmert DB, Russo SM, et al. 2015. Gene Model Annotations for Drosophila melanogaster: The Rule-Benders. G3 GenesGenomesGenetics 5: 1737–1749. Crowe ML, Wang X-Q, Rothnagel JA. 2006. Evidence for conservation and selection of upstream open reading frames suggests probable encoding of bioactive peptides. BMC Genomics 7: 16. Curinha A, Oliveira Braz S, Pereira-Castro I, Cruz A, Moreira A. 2014. Implications of polyadenylation in health and disease. Nucl Austin Tex 5: 508–519. Dabbs ER. 1986. Mutant Studies on the Prokaryotic Ribosome. In Structure, Function, and Genetics of Ribosomes (eds. B. Hardesty and G. Kramer), Springer Series in Molecular Biology, pp. 733–748, Springer, New York, NY https://doi.org/10.1007/978-1-4612-4884-2_43 (Accessed January 19, 2024). Daiß JL, Pilsl M, Straub K, Bleckmann A, Höcherl M, Heiss FB, Abascal-Palacios G, Ramsay EP, Tlučková K, Mars J-C, et al. 2022. The human RNA polymerase I structure reveals an HMG-like docking domain specific to metazoans. Life Sci Alliance 5: e202201568. Davidson AE, Balciunas D, Mohn D, Shaffer J, Hermanson S, Sivasubbu S, Cliff MP, Hackett PB, Ekker SC. 2003. Efficient gene delivery and gene expression in zebrafish using the Sleeping Beauty transposon. Dev Biol 263: 191–202. De Keersmaecker K, Atak ZK, Li N, Vicente C, Patchett S, Girardi T, Gianfelici V, Geerdens E, Clappier E, Porcu M, et al. 2013. Exome sequencing identifies mutation in CNOT3 and ribosomal genes RPL5 and RPL10 in T-cell acute lymphoblastic leukemia. Nat Genet 45: 186–190. Denis H, Wegnez M. 1977. Biochemical research on oogenesis: oocytes of Xenopus laevis synthesize but do not accumulate 5S RNA of somatic type. Dev Biol 58: 212–217. Derenzini M, Montanaro L, Chillà A, Tosti E, Vici M, Barbieri S, Govoni M, Mazzini G, Treré D. 2005. Key role of the achievement of an appropriate ribosomal RNA complement for G1-S phase transition in H4-II-E-C3 rat hepatoma cells. J Cell Physiol 202: 483–491. 72 Despic V, Dejung M, Gu M, Krishnan J, Zhang J, Herzel L, Straube K, Gerstein MB, Butter F, Neugebauer KM. 2017. Dynamic RNA–protein interactions underlie the zebrafish maternal-to-zygotic transition. Genome Res. https://genome.cshlp.org/content/early/2017/05/13/gr.215954.116 (Accessed July 31, 2023). Dever TE, Green R. 2012. The elongation, termination, and recycling phases of translation in eukaryotes. Cold Spring Harb Perspect Biol 4: a013706. Díaz-López I, Toribio R, Berlanga JJ, Ventoso I. 2019. An mRNA-binding channel in the ES6S region of the translation 48S-PIC promotes RNA unwinding and scanning eds. N. Sonenberg, J.L. Manley, and N. Sonenberg. eLife 8: e48246. Dresios J, Chappell SA, Zhou W, Mauro VP. 2006. An mRNA-rRNA base-pairing mechanism for translation initiation in eukaryotes. Nat Struct Mol Biol 13: 30–34. Eberhard D, Tora L, Egly J-M, Grummt I. 1993. A TBP-containing multiprotein complex (TIF-IB) mediates transcription specificity of murine RNA polymerase I. Nucleic Acids Res 21: 4180–4186. Eddy EM. 1975. Germ plasm and the differentiation of the germ cell line. Int Rev Cytol 43: 229–280. Erales J, Marchand V, Panthu B, Gillot S, Belin S, Ghayad SE, Garcia M, Laforêts F, Marcel V, Baudin-Baillieu A, et al. 2017. Evidence for rRNA 2′-O-methylation plasticity: Control of intrinsic translational capabilities of human ribosomes. Proc Natl Acad Sci 114: 12934–12939. Farley-Barnes KI, Ogawa LM, Baserga SJ. 2019. Ribosomopathies: old concepts, new controversies. Trends Genet TIG 35: 754–767. Ferreira IA, Oliveira C, Venere PC, Galetti PM, Martins C. 2007. 5S rDNA variation and its phylogenetic inference in the genus Leporinus (Characiformes: Anostomidae). Genetica 129: 253–257. Ferretti MB, Karbstein K. 2019. Does functional specialization of ribosomes really exist? RNA N Y N 25: 521–538. Finch AJ, Hilcenko C, Basse N, Drynan LF, Goyenechea B, Menne TF, González Fernández Á, Simpson P, D’Santos CS, Arends MJ, et al. 2011. Uncoupling of GTP hydrolysis from eIF6 release on the ribosome causes Shwachman-Diamond syndrome. Genes Dev 25: 917–929. Fonseca BD, Zakaria C, Jia J-J, Graber TE, Svitkin Y, Tahmasebi S, Healy D, Hoang H- D, Jensen JM, Diao IT, et al. 2015. La-related Protein 1 (LARP1) Represses Terminal Oligopyrimidine (TOP) mRNA Translation Downstream of mTOR Complex 1 (mTORC1). J Biol Chem 290: 15996–16020. Fox GE. 2010. Origin and Evolution of the Ribosome. Cold Spring Harb Perspect Biol 2: a003483. Fujii K, Susanto TT, Saurabh S, Barna M. 2018. Decoding the Function of Expansion Segments in Ribosomes. Mol Cell 72: 1013-1020.e6. Furuichi Y. 2015. Discovery of m(7)G-cap in eukaryotic mRNAs. Proc Jpn Acad Ser B Phys Biol Sci 91: 394–409. Gay DM, Lund AH, Jansson MD. 2022. Translational control through ribosome heterogeneity and functional specialization. Trends Biochem Sci 47: 66–81. Genuth NR, Barna M. 2018. The Discovery of Ribosome Heterogeneity and Its Implications for Gene Regulation and Organismal Life. Mol Cell 71: 364–374. 73 Gerbi SA. 1986. The evolution of eukaryotic ribosomal DNA. Biosystems 19: 247–258. Gibbons JG, Branco AT, Yu S, Lemos B. 2014. Ribosomal DNA copy number is coupled with gene expression variation and mitochondrial abundance in humans. Nat Commun 5: 4850. Giegé R, Jühling F, Pütz J, Stadler P, Sauter C, Florentz C. 2012. Structure of transfer RNAs: similarity and variability. Wiley Interdiscip Rev RNA 3: 37–61. Gilbert WV. 2011. Functional Specialization of Ribosomes? Trends Biochem Sci 36: 127– 132. Gonzalez IL, Sylvester JE. 2001. Human rDNA: evolutionary patterns within the genes and tandem arrays derived from multiple chromosomes. Genomics 73: 255–263. Goobie S, Popovic M, Morrison J, Ellis L, Ginzberg H, Boocock GR, Ehtesham N, Bétard C, Brewer CG, Roslin NM, et al. 2001. Shwachman-Diamond syndrome with exocrine pancreatic dysfunction and bone marrow failure maps to the centromeric region of chromosome 7. Am J Hum Genet 68: 1048–1054. Gornung E, Colangelo P, Annesi F. 2007. 5S ribosomal RNA genes in six species of Mediterranean grey mullets: genomic organization and phylogenetic inference. Genome 50: 787–795. Graille M, Séraphin B. 2012. Surveillance pathways rescuing eukaryotic ribosomes lost in translation. Nat Rev Mol Cell Biol 13: 727–735. Graindorge A, Thuret R, Pollet N, Osborne HB, Audic Y. 2006. Identification of post- transcriptionally regulated Xenopus tropicalis maternal mRNAs by microarray. Nucleic Acids Res 34: 986–995. Grandori C, Gomez-Roman N, Felton-Edkins ZA, Ngouenet C, Galloway DA, Eisenman RN, White RJ. 2005. c-Myc binds to human ribosomal DNA and stimulates transcription of rRNA genes by RNA polymerase I. Nat Cell Biol 7: 311–318. Grummt I, Längst G. 2013. Epigenetic control of RNA polymerase I transcription in mammalian cells. Biochim Biophys Acta 1829: 393–404. Guinta DR, Tso JY, Narayanswami S, Hamkalo BA, Korn LJ. 1986. Early replication and expression of oocyte-type 5S RNA genes in a Xenopus somatic cell line carrying a translocation. Proc Natl Acad Sci U S A 83: 5150–5154. Gulen B, Petrov AS, Okafor CD, Vander Wood D, O’Neill EB, Hud NV, Williams LD. 2016. Ribosomal small subunit domains radiate from a central core. Sci Rep 6: 20885. Gunderson JH, Sogin ML, Wollett G, Hollingdale M, de la Cruz VF, Waters AP, McCutchan TF. 1987. Structurally distinct, stage-specific ribosomes occur in Plasmodium. Science 238: 933–937. Guo H. 2018. Specialized ribosomes and the control of translation. Biochem Soc Trans 46: 855–869. Haag ES, Dinman JD. 2019. Still Searching for Specialized Ribosomes. Dev Cell 48: 744– 746. Harding SM, Boiarsky JA, Greenberg RA. 2015. ATM Dependent Silencing Links Nucleolar Chromatin Reorganization to DNA Damage Recognition. Cell Rep 13: 251–259. Hariharan N, Ghosh S, Palakodeti D. 2023. The story of rRNA expansion segments: Finding functionality amidst diversity. WIREs RNA 14: e1732. 74 Harvey RF, Smith TS, Mulroney T, Queiroz RML, Pizzinga M, Dezi V, Villenueva E, Ramakrishna M, Lilley KS, Willis AE. 2018. Trans-acting translational regulatory RNA binding proteins. WIREs RNA 9: e1465. Hashimoto Y, Maegawa S, Nagai T, Yamaha E, Suzuki H, Yasuda K, Inoue K. 2004. Localized maternal factors are required for zebrafish germ cell formation. Dev Biol 268: 152–161. Heiman M, Kulicke R, Fenster RJ, Greengard P, Heintz N. 2014. Cell type–specific mRNA purification by translating ribosome affinity purification (TRAP). Nat Protoc 9: 1282–1291. Heix J, Vente A, Voit R, Budde A, Michaelidis TM, Grummt I. 1998. Mitotic silencing of human rRNA synthesis: inactivation of the promoter selectivity factor SL1 by cdc2/cyclin B-mediated phosphorylation. EMBO J 17: 7373–7381. Heix J, Zomerdijk JCBM, Ravanpay A, Tjian R, Grummt I. 1997. Cloning of murine RNA polymerase I-specific TAF factors: Conserved interactions between the subunits of the species-specific transcription initiation factor TIF-IB/SL1. Proc Natl Acad Sci 94: 1733–1738. Hellen CUT. 2018. Translation Termination and Ribosome Recycling in Eukaryotes. Cold Spring Harb Perspect Biol 10: a032656. Hempel WM, Cavanaugh AH, Hannan RD, Taylor L, Rothblum LI. 1996. The species- specific RNA polymerase I transcription factor SL-1 binds to upstream binding factor. Mol Cell Biol 16: 557–563. Hendrickson DG, Hogan DJ, McCullough HL, Myers JW, Herschlag D, Ferrell JE, Brown PO. 2009. Concordant Regulation of Translation and mRNA Abundance for Hundreds of Targets of a Human microRNA. PLOS Biol 7: e1000238. Hentze MW, Castello A, Schwarzl T, Preiss T. 2018. A brave new world of RNA-binding proteins. Nat Rev Mol Cell Biol 19: 327–341. Hernández G, Altmann M, Sierra JM, Urlaub H, Diez del Corral R, Schwartz P, Rivera- Pomar R. 2005. Functional analysis of seven genes encoding eight translation initiation factor 4E (eIF4E) isoforms in Drosophila. Mech Dev 122: 529–543. Herpin A, Rohr S, Riedel D, Kluever N, Raz E, Schartl M. 2007. Specification of primordial germ cells in medaka (Oryzias latipes). BMC Dev Biol 7: 3. Herrera SC, Bach EA. 2018. JNK signaling triggers spermatogonial dedifferentiation during chronic stress to maintain the germline stem cell pool in the Drosophila testis. eLife 7: e36095. Herzog E, Guilley H, Fritsch C. 1995. Translation of the second gene of peanut clump virus RNA 2 occurs by leaky scanning in vitro. Virology 208: 215–225. Heyn P, Salmonowicz H, Rodenfels J, Neugebauer KM. 2017. Activation of transcription enforces the formation of distinct nuclear bodies in zebrafish embryos. RNA Biol 14: 752–760. Ho JJD, Balukoff NC, Theodoridis PR, Wang M, Krieger JR, Schatz JH, Lee S. 2020. A network of RNA-binding proteins controls translation efficiency to activate anaerobic metabolism. Nat Commun 11: 2677. Hochheimer A, Tjian R. 2003. Diversified transcription initiation complexes expand promoter selectivity and tissue-specific gene expression. Genes Dev 17: 1309– 1320. 75 Hodson CN, Ross L. 2021. Evolutionary Perspectives on Germline-Restricted Chromosomes in Flies (Diptera). Genome Biol Evol 13: evab072. Hoffman AM, Chen Q, Zheng T, Nicchitta CV. 2019. Heterogeneous translational landscape of the endoplasmic reticulum revealed by ribosome proximity labeling and transcriptome analysis. J Biol Chem 294: 8942–8958. Holley RW, Apgar J, Everett GA, Madison JT, Marquisee M, Merrill SH, Penswick JR, Zamir A. 1965. STRUCTURE OF A RIBONUCLEIC ACID. Science 147: 1462– 1465. Hong S, Freeberg MA, Han T, Kamath A, Yao Y, Fukuda T, Suzuki T, Kim JK, Inoki K. 2017. LARP1 functions as a molecular switch for mTORC1-mediated translation of an essential class of mRNAs. eLife 6: e25237. Hopes T, Norris K, Agapiou M, McCarthy CGP, Lewis PA, O’Connell MJ, Fontana J, Aspden JL. 2022. Ribosome heterogeneity in Drosophila melanogaster gonads through paralog-switching. Nucleic Acids Res 50: 2240–2257. Hsieh AC, Liu Y, Edlind MP, Ingolia NT, Janes MR, Sher A, Shi EY, Stumpf CR, Christensen C, Bonham MJ, et al. 2012. The translational landscape of mTOR signalling steers cancer initiation and metastasis. Nature 485: 55–61. Huggins HP, Keiper BD. 2020. Regulation of Germ Cell mRNPs by eIF4E:4EIP Complexes: Multiple Mechanisms, One Goal. Front Cell Dev Biol 8: 562. Huggins HP, Subash JS, Stoffel H, Henderson MA, Hoffman JL, Buckner DS, Sengupta MS, Boag PR, Lee M-H, Keiper BD. 2020. Distinct roles of two eIF4E isoforms in the germline of Caenorhabditis elegans. J Cell Sci 133: jcs237990. Iadevaia V, Caldarola S, Tino E, Amaldi F, Loreni F. 2008. All translation elongation factors and the e, f, and h subunits of translation initiation factor 3 are encoded by 5′-terminal oligopyrimidine (TOP) mRNAs. RNA 14: 1730–1736. Ikeuchi K, Inada T. 2016. Ribosome-associated Asc1/RACK1 is required for endonucleolytic cleavage induced by stalled ribosome at the 3′ end of nonstop mRNA. Sci Rep 6: 28234. Imami K, Milek M, Bogdanow B, Yasuda T, Kastelic N, Zauber H, Ishihama Y, Landthaler M, Selbach M. 2018. Phosphorylation of the Ribosomal Protein RPL12/uL11 Affects Translation during Mitosis. Mol Cell 72: 84-98.e9. Ingolia NT, Brar GA, Rouskin S, McGeachy AM, Weissman JS. 2012. The ribosome profiling strategy for monitoring translation in vivo by deep sequencing of ribosome-protected mRNA fragments. Nat Protoc 7: 1534–1550. Ivanov A, Mikhailova T, Eliseev B, Yeramala L, Sokolova E, Susorov D, Shuvalov A, Schaffitzel C, Alkalaeva E. 2016. PABP enhances release factor recruitment and stop codon recognition during translation termination. Nucleic Acids Res 44: 7766– 7776. Ivanov IP, Firth AE, Michel AM, Atkins JF, Baranov PV. 2011. Identification of evolutionarily conserved non-AUG-initiated N-terminal extensions in human coding sequences. Nucleic Acids Res 39: 4220–4234. Ivics Z, Hackett PB, Plasterk RH, Izsvák Z. 1997. Molecular reconstruction of Sleeping Beauty, a Tc1-like transposon from fish, and its transposition in human cells. Cell 91: 501–510. Jack K, Bellodi C, Landry DM, Niederer RO, Meskauskas A, Musalgaonkar S, Kopmar N, Krasnykh O, Dean AM, Thompson SR, et al. 2011. rRNA Pseudouridylation 76 Defects Affect Ribosomal Ligand Binding and Translational Fidelity from Yeast to Human Cells. Mol Cell 44: 660–666. Jackson RJ, Hellen CUT, Pestova TV. 2010. The mechanism of eukaryotic translation initiation and principles of its regulation. Nat Rev Mol Cell Biol 11: 113–127. Jäkel S, Görlich D. 1998. Importin beta, transportin, RanBP5 and RanBP7 mediate nuclear import of ribosomal proteins in mammalian cells. EMBO J 17: 4491–4502. Jamieson-Lucy A, Mullins MC. 2019. The vertebrate Balbiani body, germ plasm, and oocyte polarity. Curr Top Dev Biol 135: 1–34. Joazeiro CAP. 2017. Ribosomal Stalling During Translation: Providing Substrates for Ribosome-Associated Protein Quality Control. Annu Rev Cell Dev Biol 33: 343– 368. Johnson LF, Penman S, Green H. 1975. Increasing content of poly A(+) mRNA of serum- stimulated cells in the absence of ribosome synthesis. J Cell Physiol 87: 141–146. Johnston PM. 1951. The embryonic history of the germ cells of the largemouth black bass, Micropterus salmoides salmoides (Lacépède). J Morphol 88: 471–542. Johnstone TG, Bazzini AA, Giraldez AJ. 2016. Upstream ORFs are prevalent translational repressors in vertebrates. EMBO J 35: 706–723. Kane DA, Kimmel CB. 1993. The zebrafish midblastula transition. Dev Camb Engl 119: 447–456. Kaufman OH, Marlow FL. 2016. Methods to study maternal regulation of germ cell specification in zebrafish. In Methods in Cell Biology, Vol. 134 of, pp. 1–32, Elsevier https://linkinghub.elsevier.com/retrieve/pii/S0091679X16000133 (Accessed November 29, 2023). Kawakami K. 2007. Tol2: a versatile gene transfer vector in vertebrates. Genome Biol 8 Suppl 1: S7. Kawakami K, Shima A, Kawakami N. 2000. Identification of a functional transposase of the Tol2 element, an Ac-like element from the Japanese medaka fish, and its transposition in the zebrafish germ lineage. Proc Natl Acad Sci U S A 97: 11403– 11408. Keenan RJ, Freymann DM, Stroud RM, Walter P. 2001. The Signal Recognition Particle. Annu Rev Biochem 70: 755–775. Khatter H, Myasnikov AG, Natchiar SK, Klaholz BP. 2015. Structure of the human 80S ribosome. Nature 520: 640–645. Kim J-H, Dilthey AT, Nagaraja R, Lee H-S, Koren S, Dudekula D, Wood Iii WH, Piao Y, Ogurtsov AY, Utani K, et al. 2018. Variation in human chromosome 21 ribosomal RNA genes characterized by TAR cloning and long-read sequencing. Nucleic Acids Res 46: 6712–6725. Kim M-S, Pinto SM, Getnet D, Nirujogi RS, Manda SS, Chaerkady R, Madugundu AK, Kelkar DS, Isserlin R, Jain S, et al. 2014. A draft map of the human proteome. Nature 509: 575–581. Kim SH, Suddath FL, Quigley GJ, McPherson A, Sussman JL, Wang AH, Seeman NC, Rich A. 1974. Three-dimensional tertiary structure of yeast phenylalanine transfer RNA. Science 185: 435–440. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. 1995. Stages of embryonic development of the zebrafish. Dev Dyn Off Publ Am Assoc Anat 203: 253–310. 77 Kirby TJ, Lee JD, England JH, Chaillou T, Esser KA, McCarthy JJ. 2015. Blunted hypertrophic response in aged skeletal muscle is associated with decreased ribosome biogenesis. J Appl Physiol Bethesda Md 1985 119: 321–327. Klein J, Grummt I. 1999. Cell cycle-dependent regulation of RNA polymerase I transcription: the nucleolar transcription factor UBF is inactive in mitosis and early G1. Proc Natl Acad Sci U S A 96: 6096–6101. Klinge S, Voigts-Hoffmann F, Leibundgut M, Ban N. 2012. Atomic structures of the eukaryotic ribosome. Trends Biochem Sci 37: 189–198. Klinge S, Woolford JL. 2019. Ribosome assembly coming into focus. Nat Rev Mol Cell Biol 20: 116–131. Knaut H, Pelegri F, Bohmann K, Schwarz H, Nüsslein-Volhard C. 2000. Zebrafish vasa RNA but not its protein is a component of the germ plasm and segregates asymmetrically before germline specification. J Cell Biol 149: 875–888. Kobayashi T. 2014. Ribosomal RNA gene repeats, their stability and cellular senescence. Proc Jpn Acad Ser B Phys Biol Sci 90: 119–129. Kobayashi T, Heck DJ, Nomura M, Horiuchi T. 1998. Expansion and contraction of ribosomal DNA repeats in Saccharomyces cerevisiae: requirement of replication fork blocking (Fob1) protein and the role of RNA polymerase I. Genes Dev 12: 3821–3830. Komar AA, Merrick WC. 2020. A Retrospective on eIF2A-and Not the Alpha Subunit of eIF2. Int J Mol Sci 21: 2054. Komili S, Farny NG, Roth FP, Silver PA. 2007. Functional specificity among ribosomal proteins regulates gene expression. Cell 131: 557–571. Komiya H, HASEGAWA M, TAKEMURA S. 1986. Differentiation of Oocyte- and Somatic- Type 5S rRNAs in Animals 1. J Biochem (Tokyo) 100: 369–374. Kondrashov N, Pusic A, Stumpf CR, Shimizu K, Hsieh AC, Ishijima J, Shiroishi T, Barna M. 2011. Ribosome-mediated specificity in Hox mRNA translation and vertebrate tissue patterning. Cell 145: 383–397. Köprunner M, Thisse C, Thisse B, Raz E. 2001. A zebrafish nanos-related gene is essential for the development of primordial germ cells. Genes Dev 15: 2877–2885. Korfsmeier KH. 1966. Zur Genese des Dottersystems in der Oocyte von Brachydanio rerio. Z Für Zellforsch Mikrosk Anat 71: 283–296. Kozak M. 1987. An analysis of 5’-noncoding sequences from 699 vertebrate messenger RNAs. Nucleic Acids Res 15: 8125–8148. Kozak M. 1978. How do eucaryotic ribosomes select initiation regions in messenger RNA? Cell 15: 1109–1123. Kozak M. 2002. Pushing the limits of the scanning mechanism for initiation of translation. Gene 299: 1–34. Krogh N, Jansson MD, Häfner SJ, Tehler D, Birkedal U, Christensen-Dalsgaard M, Lund AH, Nielsen H. 2016. Profiling of 2’-O-Me in human rRNA reveals a subset of fractionally modified positions and provides evidence for ribosome heterogeneity. Nucleic Acids Res 44: 7884–7895. Kuhn A, Vente A, Dorée M, Grummt I. 1998. Mitotic phosphorylation of the TBP- containing factor SL1 represses ribosomal gene transcription. J Mol Biol 284: 1–5. 78 Kuo BA, Gonzalez IL, Gillespie DA, Sylvester JE. 1996. Human ribosomal RNA variants from a single individual and their expression in different tissues. Nucleic Acids Res 24: 4817–4824. Kwan KM, Fujimoto E, Grabher C, Mangum BD, Hardy ME, Campbell DS, Parant JM, Yost HJ, Kanki JP, Chien C-B. 2007. The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn Off Publ Am Assoc Anat 236: 3088–3099. Lahr RM, Fonseca BD, Ciotti GE, Al-Ashtal HA, Jia J-J, Niklaus MR, Blagden SP, Alain T, Berman AJ. 2017. La-related protein 1 (LARP1) binds the mRNA cap, blocking eIF4F assembly on TOP mRNAs. eLife 6: e24146. Lai F, King ML. 2013. Repressive translational control in germ cells. Mol Reprod Dev 80: 665–676. Lai K, Amsterdam A, Farrington S, Bronson RT, Hopkins N, Lees JA. 2009. Many ribosomal protein mutations are associated with growth impairment and tumor predisposition in zebrafish. Dev Dyn Off Publ Am Assoc Anat 238: 76–85. Lange M, Granados A, VijayKumar S, Bragantini J, Ancheta S, Santhosh S, Borja M, Kobayashi H, McGeever E, Solak AC, et al. 2023. Zebrahub – Multimodal Zebrafish Developmental Atlas Reveals the State-Transition Dynamics of Late- Vertebrate Pluripotent Axial Progenitors. 2023.03.06.531398. https://www.biorxiv.org/content/10.1101/2023.03.06.531398v2 (Accessed January 30, 2024). Lawson KA, Dunn NR, Roelen BA, Zeinstra LM, Davis AM, Wright CV, Korving JP, Hogan BL. 1999. Bmp4 is required for the generation of primordial germ cells in the mouse embryo. Genes Dev 13: 424–436. Leesch F, Lorenzo-Orts L, Pribitzer C, Grishkovskaya I, Roehsner J, Chugunova A, Matzinger M, Roitinger E, Belačić K, Kandolf S, et al. 2023. A molecular network of conserved factors keeps ribosomes dormant in the egg. Nature 613: 712–720. Lesch BJ, Page DC. 2012. Genetics of germ cell development. Nat Rev Genet 13: 781– 794. Leung E, Brown JD. 2010. Biogenesis of the signal recognition particle. Biochem Soc Trans 38: 1093–1098. Levy S, Avni D, Hariharan N, Perry RP, Meyuhas O. 1991. Oligopyrimidine tract at the 5’ end of mammalian ribosomal protein mRNAs is required for their translational control. Proc Natl Acad Sci U S A 88: 3319–3323. Li D, Wang J. 2020. Ribosome heterogeneity in stem cells and development. J Cell Biol 219: e202001108. Li M, Hong N, Xu H, Song J, Hong Y. 2016. Germline replacement by blastula cell transplantation in the fish medaka. Sci Rep 6: 29658. Liljas A, Sanyal S. 2018. The enigmatic ribosomal stalk. Q Rev Biophys 51: e12. Linhartova Z, Saito T, Psenicka M. 2014. Embryogenesis, visualization and migration of primordial germ cells in tench (Tinca tinca). J Appl Ichthyol 30: 29–39. Liu Y, Bourgeois CF, Pang S, Kudla M, Dreumont N, Kister L, Sun Y-H, Stevenin J, Elliott DJ. 2009. The Germ Cell Nuclear Proteins hnRNP G-T and RBMY Activate a Testis-Specific Exon. PLOS Genet 5: e1000707. Locati MD, Pagano JFB, Abdullah F, Ensink WA, van Olst M, van Leeuwen S, Nehrdich U, Spaink HP, Rauwerda H, Jonker MJ, et al. 2018. Identifying small RNAs derived 79 from maternal- and somatic-type rRNAs in zebrafish development. Genome 61: 371–378. Locati MD, Pagano JFB, Ensink WA, van Olst M, van Leeuwen S, Nehrdich U, Zhu K, Spaink HP, Girard G, Rauwerda H, et al. 2017a. Linking maternal and somatic 5S rRNA types with different sequence-specific non-LTR retrotransposons. RNA N Y N 23: 446–456. Locati MD, Pagano JFB, Girard G, Ensink WA, van Olst M, van Leeuwen S, Nehrdich U, Spaink HP, Rauwerda H, Jonker MJ, et al. 2017b. Expression of distinct maternal and somatic 5.8S, 18S, and 28S rRNA types during zebrafish development. RNA N Y N 23: 1188–1199. Lodish HF. 1974. Model for the regulation of mRNA translation applied to haemoglobin synthesis. Nature 251: 385–388. Lohrum MAE, Ludwig RL, Kubbutat MHG, Hanlon M, Vousden KH. 2003. Regulation of HDM2 activity by the ribosomal protein L11. Cancer Cell 3: 577–587. Long EO, Dawid IB. 1980. Repeated genes in eukaryotes. Annu Rev Biochem 49: 727– 764. Lopes AM, Miguel RN, Sargent CA, Ellis PJ, Amorim A, Affara NA. 2010. The human RPS4 paralogue on Yq11.223 encodes a structurally conserved ribosomal protein and is preferentially expressed during spermatogenesis. BMC Mol Biol 11: 33. Lorenz C, Lünse CE, Mörl M. 2017. tRNA Modifications: Impact on Structure and Thermal Adaptation. Biomolecules 7: 35. Ludwig LS, Gazda HT, Eng JC, Eichhorn SW, Thiru P, Ghazvinian R, George TI, Gotlib JR, Beggs AH, Sieff CA, et al. 2014. Altered translation of GATA1 in Diamond- Blackfan anemia. Nat Med 20: 748–753. Lütcke H. 1995. Signal recognition particle (SRP), a ubiquitous initiator of protein translocation. Eur J Biochem 228: 531–550. MacDonald CT, Gibbs JH. 1969. Concerning the kinetics of polypeptide synthesis on polyribosomes. Biopolymers 7: 707–725. MacDonald CT, Gibbs JH, Pipkin AC. 1968. Kinetics of biopolymerization on nucleic acid templates. Biopolymers 6: 1–5. Mageeney CM, Kearse MG, Gershman BW, Pritchard CE, Colquhoun JM, Ware VC. 2018. Functional interplay between ribosomal protein paralogues in the eRpL22 family in Drosophila melanogaster. Fly (Austin) 12: 143–163. Mageeney CM, Ware VC. 2019. Specialized eRpL22 paralogue-specific ribosomes regulate specific mRNA translation in spermatogenesis in Drosophila melanogaster. Mol Biol Cell 30: 2240–2253. Marcel V, Ghayad SE, Belin S, Therizols G, Morel A-P, Solano-Gonzàlez E, Vendrell JA, Hacot S, Mertani HC, Albaret MA, et al. 2013. p53 acts as a safeguard of translational control by regulating fibrillarin and rRNA methylation in cancer. Cancer Cell 24: 318–330. Marechal V, Elenbaas B, Piette J, Nicolas JC, Levine AJ. 1994. The ribosomal L5 protein is associated with mdm-2 and mdm-2-p53 complexes. Mol Cell Biol 14: 7414– 7420. Martins C, Wasko AP. 2006. ORGANIZATION AND EVOLUTION OF 5S RIBOSOMAL DNA IN THE FISH GENOME. https://www.semanticscholar.org/paper/ORGANIZATION-AND-EVOLUTION-OF- 80 5S-RIBOSOMAL-DNA-IN-Martins- Wasko/fca5fd69ba11f8c3f4112fb166850a4c4a18372f (Accessed January 19, 2024). Martins C, Wasko AP, Oliveira C, Porto-Foresti F, Parise-Maltempi PP, Wright JM, Foresti F. 2002. Dynamics of 5S rDNA in the tilapia (Oreochromis niloticus) genome: repeat units, inverted sequences, pseudogenes and chromosome loci. Cytogenet Genome Res 98: 78–85. Mashkova TD, Serenkova TI, Mazo AM, Avdonina TA, Timofeyeva MYa null, Kisselev LL. 1981. The primary structure of oocyte and somatic 5S rRNAs from the loach Misgurnus fossilis. Nucleic Acids Res 9: 2141–2151. Mauro VP, Edelman GM. 2002. The ribosome filter hypothesis. Proc Natl Acad Sci U S A 99: 12031–12036. Mauro VP, Edelman GM. 2007. The Ribosome Filter Redux. Cell Cycle Georget Tex 6: 2246–2251. Mauro VP, Matsuda D. 2016. Translation regulation by ribosomes: Increased complexity and expanded scope. RNA Biol 13: 748–755. Maxwell ES, Fournier MJ. 1995. The small nucleolar RNAs. Annu Rev Biochem 64: 897– 934. McGlincy NJ, Ingolia NT. 2017. Transcriptome-wide measurement of translation by ribosome profiling. Methods 126: 112–129. McQuillen K, Roberts RB, Britten RJ. 1959. SYNTHESIS OF NASCENT PROTEIN BY RIBOSOMES IN ESCHERICHIA COLI. Proc Natl Acad Sci U S A 45: 1437–1447. McStay B, Grummt I. 2008. The epigenetics of rRNA genes: from molecular to chromosome biology. Annu Rev Cell Dev Biol 24: 131–157. Meijer HA, Thomas AAM. 2002. Control of eukaryotic protein synthesis by upstream open reading frames in the 5’-untranslated region of an mRNA. Biochem J 367: 1–11. Melnikov S, Ben-Shem A, Garreau de Loubresse N, Jenner L, Yusupova G, Yusupov M. 2012. One core, two shells: bacterial and eukaryotic ribosomes. Nat Struct Mol Biol 19: 560–567. Menne TF, Goyenechea B, Sánchez-Puig N, Wong CC, Tonkin LM, Ancliff PJ, Brost RL, Costanzo M, Boone C, Warren AJ. 2007. The Shwachman-Bodian-Diamond syndrome protein mediates translational activation of ribosomes in yeast. Nat Genet 39: 486–495. Merrick WC, Pavitt GD. 2018. Protein Synthesis Initiation in Eukaryotic Cells. Cold Spring Harb Perspect Biol 10: a033092. Miller SC, MacDonald CC, Kellogg MK, Karamysheva ZN, Karamyshev AL. 2023. Specialized Ribosomes in Health and Disease. Int J Mol Sci 24: 6334. Mills EW, Green R. 2017. Ribosomopathies: There’s strength in numbers. Science 358: eaan2755. Moore PB, Steitz TA. 2002. The involvement of RNA in ribosome function. Nature 418: 229–235. Morris DR, Geballe AP. 2000. Upstream Open Reading Frames as Regulators of mRNA Translation. Mol Cell Biol 20: 8635–8642. Moss T, Langlois F, Gagnon-Kugler T, Stefanovsky V. 2007. A housekeeper with power of attorney: the rRNA genes in ribosome biogenesis. Cell Mol Life Sci CMLS 64: 29–49. 81 Moss T, Stefanovsky VY. 2002. At the Center of Eukaryotic Life. Cell 109: 545–548. Murano K, Okuwaki M, Momose F, Kumakura M, Ueshima S, Newbold RF, Nagata K. 2014. Reconstitution of human rRNA gene transcription in mouse cells by a complete SL1 complex. J Cell Sci 127: 3309–3319. Nadano D, Notsu T, Matsuda T, Sato T-A. 2002. A human gene encoding a protein homologous to ribosomal protein L39 is normally expressed in the testis and derepressed in multiple cancer cells. Biochim Biophys Acta BBA - Gene Struct Expr 1577: 430–436. Nagasawa K, Fernandes JM, Yoshizaki G, Miwa M, Babiak I. 2013. Identification and Migration of Primordial Germ Cells in Atlantic Salmon, Salmo salar: Characterization of Vasa, Dead End, and Lymphocyte Antigen 75 Genes. Mol Reprod Dev 80: 118–131. Nakagawa S, Niimura Y, Gojobori T, Tanaka H, Miura K. 2008. Diversity of preferred nucleotide sequences around the translation initiation codon in eukaryote genomes. Nucleic Acids Res 36: 861–871. Newport J, Kirschner M. 1982. A major developmental transition in early Xenopus embryos: II. Control of the onset of transcription. Cell 30: 687–696. Ni J, Clark KJ, Fahrenkrug SC, Ekker SC. 2008. Transposon tools hopping in vertebrates. Brief Funct Genomic Proteomic 7: 444–453. Noderer WL, Flockhart RJ, Bhaduri A, Diaz de Arce AJ, Zhang J, Khavari PA, Wang CL. 2014. Quantitative analysis of mammalian translation initiation sites by FACS-seq. Mol Syst Biol 10: 748. Nüsslein-Volhard C, Dahm R, eds. 2002. Zebrafish: a practical approach. 1st ed. Oxford University Press, Oxford. Oh WJ, Wu C, Kim SJ, Facchinetti V, Julien L-A, Finlan M, Roux PP, Su B, Jacinto E. 2010. mTORC2 can associate with ribosomes to promote cotranslational phosphorylation and stability of nascent Akt polypeptide. EMBO J 29: 3939–3951. O’Sullivan AC, Sullivan GJ, McStay B. 2002. UBF binding in vivo is not restricted to regulatory sequences within the vertebrate ribosomal DNA repeat. Mol Cell Biol 22: 657–668. Oyarbide U, Shah AN, Amaya-Mejia W, Snyderman M, Kell MJ, Allende DS, Calo E, Topczewski J, Corey SJ. 2020. Loss of Sbds in zebrafish leads to neutropenia and pancreas and liver atrophy. JCI Insight 5: e134309, 134309. Pánek J, Kolář M, Vohradský J, Shivaya Valášek L. 2013. An evolutionary conserved pattern of 18S rRNA sequence complementarity to mRNA 5′ UTRs and its implications for eukaryotic gene translation regulation. Nucleic Acids Res 41: 7625–7634. Parenteau J, Durand M, Morin G, Gagnon J, Lucier J-F, Wellinger RJ, Chabot B, Elela SA. 2011. Introns within ribosomal protein genes regulate the production and function of yeast ribosomes. Cell 147: 320–331. Parks MM, Kurylo CM, Dass RA, Bojmar L, Lyden D, Vincent CT, Blanchard SC. 2018. Variant ribosomal RNA alleles are conserved and exhibit tissue-specific expression. Sci Adv 4: eaao0665. Patton EE, Zon LI, Langenau DM. 2021. Zebrafish disease models in drug discovery: from preclinical modelling to clinical trials. Nat Rev Drug Discov 20: 611–628. 82 Pelava A, Schneider C, Watkins NJ. 2016. The importance of ribosome production, and the 5S RNP–MDM2 pathway, in health and disease. Biochem Soc Trans 44: 1086– 1090. Peña C, Hurt E, Panse VG. 2017. Eukaryotic ribosome assembly, transport and quality control. Nat Struct Mol Biol 24: 689–699. Pennell M, Rodriguez OL, Watson CT, Greiff V. 2023. The evolutionary and functional significance of germline immunoglobulin gene variation. Trends Immunol 44: 7– 21. Penzo M, Montanaro L. 2018. Turning Uridines around: Role of rRNA Pseudouridylation in Ribosome Biogenesis and Ribosomal Function. Biomolecules 8: 38. Perez-Perri JI, Rogell B, Schwarzl T, Stein F, Zhou Y, Rettel M, Brosig A, Hentze MW. 2018. Discovery of RNA-binding proteins and characterization of their dynamic responses by enhanced RNA interactome capture. Nat Commun 9: 4408. Perucho L, Artero-Castro A, Guerrero S, Ramón y Cajal S, LLeonart ME, Wang Z-Q. 2014. RPLP1, a crucial ribosomal protein for embryonic development of the nervous system. PloS One 9: e99956. Pestova TV, Hellen CUT. 2003. Translation elongation after assembly of ribosomes on the Cricket paralysis virus internal ribosomal entry site without initiation factors or initiator tRNA. Genes Dev 17: 181–186. Peterson RC, Doering JL, Brown DD. 1980. Characterization of two xenopus somatic 5S DNAs and one minor oocyte-specific 5S DNA. Cell 20: 131–141. Petrov AS, Bernier CR, Hsiao C, Norris AM, Kovacs NA, Waterbury CC, Stepanov VG, Harvey SC, Fox GE, Wartell RM, et al. 2014. Evolution of the ribosome at atomic resolution. Proc Natl Acad Sci U S A 111: 10251–10256. Poirot O, Timsit Y. 2016. Neuron-Like Networks Between Ribosomal Proteins Within the Ribosome. Sci Rep 6: 26485. Polymenis M. 2020. Ribosomal proteins: mutant phenotypes by the numbers and associated gene expression changes. Open Biol 10: 200114. Popovic M, Goobie S, Morrison J, Ellis L, Ehtesham N, Richards N, Boocock G, Durie PR, Rommens JM. 2002. Fine mapping of the locus for Shwachman-Diamond syndrome at 7q11, identification of shared disease haplotypes, and exclusion of TPST1 as a candidate gene. Eur J Hum Genet EJHG 10: 250–258. Potapova TA, Gerton JL. 2019. Ribosomal DNA and the nucleolus in the context of genome organization. Chromosome Res 27: 109–127. Power L. 2022. Beginners guide to ribosome profiling. The Biochemist 44: 30–34. Presslauer C, Nagasawa K, Fernandes JMO, Babiak I. 2012. Expression of vasa and nanos3 during primordial germ cell formation and migration in Atlantic cod (Gadus morhua L.). Theriogenology 78: 1262–1277. Provost E, Wehner KA, Zhong X, Ashar F, Nguyen E, Green R, Parsons MJ, Leach SD. 2012. Ribosomal biogenesis genes play an essential and p53-independent role in zebrafish pancreas development. Dev Camb Engl 139: 3232–3241. Ramagopal S. 1990. Induction of cell-specific ribosomal proteins in aggregation- competent nonmorphogenetic Dictyostelium discoideum. Biochem Cell Biol Biochim Biol Cell 68: 1281–1287. 83 Rao S, Lee S-Y, Gutierrez A, Perrigoue J, Thapa RJ, Tu Z, Jeffers JR, Rhodes M, Anderson S, Oravecz T, et al. 2012. Inactivation of ribosomal protein L22 promotes transformation by induction of the stemness factor, Lin28B. Blood 120: 3764–3773. Reid DW, Nicchitta CV. 2015. Diversity and selectivity in mRNA translation on the endoplasmic reticulum. Nat Rev Mol Cell Biol 16: 221–231. Reid DW, Nicchitta CV. 2012. Primary role for endoplasmic reticulum-bound ribosomes in cellular translation identified by ribosome profiling. J Biol Chem 287: 5518–5527. Reschke M, Clohessy JG, Seitzer N, Goldstein DP, Breitkopf SB, Schmolze DB, Ala U, Asara JM, Beck AH, Pandolfi PP. 2013. Characterization and analysis of the composition and dynamics of the mammalian riboproteome. Cell Rep 4: 1276– 1287. Rhodin MHJ, Dinman JD. 2011. An Extensive Network of Information Flow through the B1b/c Intersubunit Bridge of the Yeast Ribosome. PLOS ONE 6: e20048. Richter JD, Lasko P. 2011. Translational Control in Oocyte Development. Cold Spring Harb Perspect Biol 3: a002758. Rivera MC, Maguire B, Lake JA. 2015. Isolation of ribosomes and polysomes. Cold Spring Harb Protoc 2015: 293–299. Robles F, de la Herrán R, Ludwig A, Rejón CR, Rejón MR, Garrido-Ramos MA. 2005. Genomic organization and evolution of the 5S ribosomal DNA in the ancient fish sturgeon. Genome 48: 18–28. Rodnina MV. 2016. The ribosome in action: Tuning of translational efficiency and protein folding. Protein Sci 25: 1390–1406. Root-Bernstein M, Root-Bernstein R. 2015. The ribosome as a missing link in the evolution of life. J Theor Biol 367: 130–158. Roussel P, André C, Masson C, Géraud G, Hernandez-Verdun D. 1993. Localization of the RNA polymerase I transcription factor hUBF during the cell cycle. J Cell Sci 104 ( Pt 2): 327–37. Russell J, Zomerdijk JCBM. 2005. RNA-polymerase-I-directed rDNA transcription, life and works. Trends Biochem Sci 30: 87–96. Russo A, Russo G. 2017. Ribosomal Proteins Control or Bypass p53 during Nucleolar Stress. Int J Mol Sci 18: 140. Sachs AB, Davis RW. 1989. The poly(A) binding protein is required for poly(A) shortening and 60S ribosomal subunit-dependent translation initiation. Cell 58: 857–867. Safer B. 1989. Nomenclature of initiation, elongation and termination factors for translation in eukaryotes. Eur J Biochem 186: 1–3. Saito T, Fujimoto T, Maegawa S, Inoue K, Tanaka M, Arai K, Yamaha E. 2006. Visualization of primordial germ cells in vivo using GFP-nos1 3’UTR mRNA. Int J Dev Biol 50: 691–699. Salim D, Gerton JL. 2019. Ribosomal DNA instability and genome adaptability. Chromosome Res Int J Mol Supramol Evol Asp Chromosome Biol 27: 73–87. Sanij E, Poortinga G, Sharkey K, Hung S, Holloway TP, Quin J, Robb E, Wong LH, Thomas WG, Stefanovsky V, et al. 2008. UBF levels determine the number of active ribosomal RNA genes in mammals. J Cell Biol 183: 1259–1274. Satoh N. 1974. An ultrastructural study of sex differentiation in the teleost Oryzias latipes. Development 32: 195–215. 84 Scheer U. 1987. Contributions of Electron Microscopic Spreading Preparations (“Miller Spreads”) to the Analysis of Chromosome Structure. In Structure and Function of Eukaryotic Chromosomes (ed. W. Hennig), Results and Problems in Cell Differentiation, pp. 147–171, Springer, Berlin, Heidelberg https://doi.org/10.1007/978-3-540-47783-9_10 (Accessed February 10, 2024). Schneider-Poetsch T, Ju J, Eyler DE, Dang Y, Bhat S, Merrick WC, Green R, Shen B, Liu JO. 2010. Inhibition of Eukaryotic Translation Elongation by Cycloheximide and Lactimidomycin. Nat Chem Biol 6: 209–217. Schuster SL, Hsieh AC. 2019. The Untranslated Regions of mRNAs in Cancer. Trends Cancer 5: 245–262. Schwanhäusser B, Busse D, Li N, Dittmar G, Schuchhardt J, Wolf J, Chen W, Selbach M. 2011. Global quantification of mammalian gene expression control. Nature 473: 337–342. Schwartz T, Blobel G. 2003. Structural Basis for the Function of the β Subunit of the Eukaryotic Signal Recognition Particle Receptor. Cell 112: 793–803. Scull CE, Schneider DA. 2019. Coordinated Control of rRNA Processing by RNA Polymerase I. Trends Genet TIG 35: 724–733. Seidelt B, Innis CA, Wilson DN, Gartmann M, Armache J-P, Villa E, Trabuco LG, Becker T, Mielke T, Schulten K, et al. 2009. Structural insight into nascent polypeptide chain-mediated translational stalling. Science 326: 1412–1415. Seydoux G, Braun RE. 2006. Pathway to totipotency: lessons from germ cells. Cell 127: 891–904. Shah P, Ding Y, Niemczyk M, Kudla G, Plotkin JB. 2013. Rate-Limiting Steps in Yeast Protein Translation. Cell 153: 1589–1601. Shaw PJ. 2015. Nucleolus. In Encyclopedia of Life Sciences, pp. 1–11, Wiley https://onlinelibrary.wiley.com/doi/10.1002/9780470015902.a0001352.pub4 (Accessed February 10, 2024). Shi Z, Fujii K, Kovary KM, Genuth NR, Röst HL, Teruel MN, Barna M. 2017. Heterogeneous Ribosomes Preferentially Translate Distinct Subpools of mRNAs Genome-wide. Mol Cell 67: 71-83.e7. Shirokikh NE, Archer SK, Beilharz TH, Powell D, Preiss T. 2017. Translation complex profile sequencing to study the in vivo dynamics of mRNA-ribosome interactions during translation initiation, elongation and termination. Nat Protoc 12: 697–731. Shoji S, Dambacher CM, Shajani Z, Williamson JR, Schultz PG. 2011. Systematic chromosomal deletion of bacterial ribosomal protein genes. J Mol Biol 413: 751– 761. Siegel V, Walter P. 1986. Removal of the Alu structural domain from signal recognition particle leaves its protein translocation activity intact. Nature 320: 81–84. Simms CL, Yan LL, Zaher HS. 2017. Ribosome Collision Is Critical for Quality Control during No-Go Decay. Mol Cell 68: 361-373.e5. Simsek D, Tiu GC, Flynn RA, Byeon GW, Leppek K, Xu AF, Chang HY, Barna M. 2017. The Mammalian Ribo-interactome Reveals Ribosome Functional Diversity and Heterogeneity. Cell 169: 1051-1065.e18. Sloan KE, Bohnsack MT, Watkins NJ. 2013. The 5S RNP couples p53 homeostasis to ribosome biogenesis and nucleolar stress. Cell Rep 5: 237–247. 85 Sloan KE, Warda AS, Sharma S, Entian K-D, Lafontaine DLJ, Bohnsack MT. 2017. Tuning the ribosome: The influence of rRNA modification on eukaryotic ribosome biogenesis and function. RNA Biol 14: 1138–1152. Sluis M van, McStay B. 2015. A localized nucleolar DNA damage response facilitates recruitment of the homology-directed repair machinery independent of cell cycle stage. Genes Dev 29: 1151–1163. Smirnov E, KALMÁROVÁ M, KOBERNA K, ZEMANOVÁ Z, MALÍNSKÝ J, MAŠATA M, CVAČKOVÁ Z, MICHALOVÁ K, RAŠKA I. 2006. NORs and Their Transcription Competence during the Cell Cycle. Folia Biol (Praha) 52: 59–70. Sonenberg N, Hinnebusch AG. 2009. Regulation of Translation Initiation in Eukaryotes: Mechanisms and Biological Targets. Cell 136: 731–745. Steitz JA. 1969. Polypeptide Chain Initiation: Nucleotide Sequences of the Three Ribosomal Binding Sites in Bacteriophage R17 RNA. Nature 224: 957–964. Steitz TA, Moore PB. 2003. RNA, the first macromolecular catalyst: the ribosome is a ribozyme. Trends Biochem Sci 28: 411–418. Streisinger G, Walker C, Dower N, Knauber D, Singer F. 1981. Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature 291: 293–296. Stults DM, Killen MW, Pierce HH, Pierce AJ. 2008. Genomic architecture and inheritance of human ribosomal RNA gene clusters. Genome Res 18: 13–18. Subtelny AO, Eichhorn SW, Chen GR, Sive H, Bartel DP. 2014. Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature 508: 66–71. Sugihara Y, Honda H, Iida T, Morinaga T, Hino S, Okajima T, Matsuda T, Nadano D. 2010. Proteomic analysis of rodent ribosomes revealed heterogeneity including ribosomal proteins L10-like, L22-like 1, and L39-like. J Proteome Res 9: 1351– 1366. Susor A, Jansova D, Anger M, Kubelka M. 2016. Translation in the mammalian oocyte in space and time. Cell Tissue Res 363: 69–84. Svitkin YV, Herdy B, Costa-Mattioli M, Gingras A-C, Raught B, Sonenberg N. 2005. Eukaryotic Translation Initiation Factor 4E Availability Controls the Switch between Cap-Dependent andInternal Ribosomal Entry Site-Mediated Translation. Mol Cell Biol 25: 10556–10565. Tadros W, Lipshitz HD. 2009. The maternal-to-zygotic transition: a play in two acts. Dev Camb Engl 136: 3033–3042. Tahmasebi S, Khoutorsky A, Mathews MB, Sonenberg N. 2018. Translation deregulation in human disease. Nat Rev Mol Cell Biol 19: 791–807. Talwar PK, Jhingran AG. 1991. Inland fishes of India and adjacent countries. Balkema, Rotterdam. Tamm T, Kisly I, Remme J. 2019. Functional Interactions of Ribosomal Intersubunit Bridges in Saccharomyces cerevisiae. Genetics 213: 1329–1339. Tang DT, Glazov EA, McWilliam SM, Barris WC, Dalrymple BP. 2009. Analysis of the complement and molecular evolution of tRNA genes in cow. BMC Genomics 10: 188. Tcherkezian J, Cargnello M, Romeo Y, Huttlin EL, Lavoie G, Gygi SP, Roux PP. 2014. Proteomic analysis of cap-dependent translation identifies LARP1 as a key regulator of 5’TOP mRNA translation. Genes Dev 28: 357–371. 86 Theusch EV, Brown KJ, Pelegri F. 2006. Separate pathways of RNA recruitment lead to the compartmentalization of the zebrafish germ plasm. Dev Biol 292: 129–141. Thoreen CC, Chantranupong L, Keys HR, Wang T, Gray NS, Sabatini DM. 2012. A unifying model for mTORC1-mediated regulation of mRNA translation. Nature 485: 109–113. Timsit Y, Sergeant-Perthuis G, Bennequin D. 2021. Evolution of ribosomal protein network architectures. Sci Rep 11: 625. Tran TM, Rao DS. 2022. RNA binding proteins in MLL-rearranged leukemia. Exp Hematol Oncol 11: 80. Treangen TJ, Salzberg SL. 2011. Repetitive DNA and next-generation sequencing: computational challenges and solutions. Nat Rev Genet 13: 36–46. Tseng H, Chou W, Wang J, Zhang X, Zhang S, Schultz RM. 2008. Mouse ribosomal RNA genes contain multiple differentially regulated variants. PloS One 3: e1843. Turner JMA. 2015. Meiotic Silencing in Mammals. Annu Rev Genet 49: 395–412. Uchiumi T, Kominami R. 1994. A functional site of the GTPase-associated center within 28S ribosomal RNA probed with an anti-RNA autoantibody. EMBO J 13: 3389– 3394. Uechi T, Nakajima Y, Chakraborty A, Torihara H, Higa S, Kenmochi N. 2008. Deficiency of ribosomal protein S19 during early embryogenesis leads to reduction of erythrocytes in a zebrafish model of Diamond-Blackfan anemia. Hum Mol Genet 17: 3204–3211. Uechi T, Tanaka T, Kenmochi N. 2001. A complete map of the human ribosomal protein genes: assignment of 80 genes to the cytogenetic map and implications for human disorders. Genomics 72: 223–230. Urven LE, Yabe T, Pelegri F. 2006. A role for non-muscle myosin II function in furrow maturation in the early zebrafish embryo. J Cell Sci 119: 4342–4352. Vanden Broeck A, Klinge S. 2023. Principles of human pre-60S biogenesis. Science 381: eadh3892. Vastenhouw NL, Cao WX, Lipshitz HD. 2019. The maternal-to-zygotic transition revisited. Dev Camb Engl 146: dev161471. Voigt F, Zhang H, Cui XA, Triebold D, Liu AX, Eglinger J, Lee ES, Chao JA, Palazzo AF. 2017. Single-Molecule Quantification of Translation-Dependent Association of mRNAs with the Endoplasmic Reticulum. Cell Rep 21: 3740–3753. Volarevic S, Stewart MJ, Ledermann B, Zilberman F, Terracciano L, Montini E, Grompe M, Kozma SC, Thomas G. 2000. Proliferation, but not growth, blocked by conditional deletion of 40S ribosomal protein S6. Science 288: 2045–2047. von der Haar T. 2008. A quantitative estimation of the global translational activity in logarithmically growing yeast cells. BMC Syst Biol 2: 87. Voorhees RM, Fernández IS, Scheres SHW, Hegde RS. 2014. Structure of the mammalian ribosome-Sec61 complex to 3.4 Å resolution. Cell 157: 1632–1643. Voronina E, Seydoux G, Sassone-Corsi P, Nagamori I. 2011. RNA granules in germ cells. Cold Spring Harb Perspect Biol 3: a002774. Waldron C, Lacroute F. 1975. Effect of growth rate on the amounts of ribosomal and transfer ribonucleic acids in yeast. J Bacteriol 122: 855–865. 87 Walter P, Blobel G. 1980. Purification of a membrane-associated protein complex required for protein translocation across the endoplasmic reticulum. Proc Natl Acad Sci U S A 77: 7112–7116. Walter P, Blobel G. 1982. Signal recognition particle contains a 7S RNA essential for protein translocation across the endoplasmic reticulum. Nature 299: 691–698. Wang Y, Zhu T, Li Q, Liu C, Han F, Chen M, Zhang L, Cui X, Qin Y, Bao S, et al. 2015. Prmt5 is required for germ cell survival during spermatogenesis in mice. Sci Rep 5: 11031. Warner JR. 1999. The economics of ribosome biosynthesis in yeast. Trends Biochem Sci 24: 437–440. Warren AJ. 2018. Molecular basis of the human ribosomopathy Shwachman-Diamond syndrome. Adv Biol Regul 67: 109–127. Waters AP, van Spaendonk RM, Ramesar J, Vervenne RA, Dirks RW, Thompson J, Janse CJ. 1997. Species-specific regulation and switching of transcription between stage-specific ribosomal RNA genes in Plasmodium berghei. J Biol Chem 272: 3583–3589. Wegnez M, Monier R, Denis H. 1972. Sequence heterogeneity of 5 S RNA in Xenopus laevis. FEBS Lett 25: 13–20. Weidinger G, Stebler J, Slanchev K, Dumstrei K, Wise C, Lovell-Badge R, Thisse C, Thisse B, Raz E. 2003. dead end, a novel vertebrate germ plasm component, is required for zebrafish primordial germ cell migration and survival. Curr Biol CB 13: 1429–1434. Weidinger G, Wolke U, Köprunner M, Klinger M, Raz E. 1999. Identification of tissues and patterning events required for distinct steps in early migration of zebrafish primordial germ cells. Development 126: 5295–5307. Weill L, Belloc E, Bava F-A, Méndez R. 2012. Translational control by changes in poly(A) tail length: recycling mRNAs. Nat Struct Mol Biol 19: 577–585. Weinstein LB, Steitz JA. 1999. Guided tours: from precursor snoRNA to functional snoRNP. Curr Opin Cell Biol 11: 378–384. White RJ. 2005. RNA polymerases I and III, growth control and cancer. Nat Rev Mol Cell Biol 6: 69–78. Wild K, Juaire KD, Soni K, Shanmuganathan V, Hendricks A, Segnitz B, Beckmann R, Sinning I. 2019. Reconstitution of the human SRP system and quantitative and systematic analysis of its ribosome interactions. Nucleic Acids Res 47: 3184–3196. Williamson A, Lehmann R. 1996. Germ cell development in Drosophila. Annu Rev Cell Dev Biol 12: 365–391. Wilson-Edell KA, Kehasse A, Scott GK, Yau C, Rothschild DE, Schilling B, Gabriel BS, Yevtushenko MA, Hanson IM, Held JM, et al. 2014. RPL24: a potential therapeutic target whose depletion or acetylation inhibits polysome assembly and cancer cell growth. Oncotarget 5: 5165–5176. Winata CL, Korzh V. 2018. The translational regulation of maternal mRNAs in time and space. Febs Lett 592: 3007–3023. Wolin SL, Walter P. 1988. Ribosome pausing and stacking during translation of a eukaryotic mRNA. EMBO J 7: 3559–3569. Wong CC, Traynor D, Basse N, Kay RR, Warren AJ. 2011. Defective ribosome assembly in Shwachman-Diamond syndrome. Blood 118: 4305–4312. 88 Woolford JL, Baserga SJ. 2013. Ribosome Biogenesis in the Yeast Saccharomyces cerevisiae. Genetics 195: 643–681. Wormington WM, Schlissel M, Brown DD. 1983. Developmental regulation of Xenopus 5S RNA genes. Cold Spring Harb Symp Quant Biol 47 Pt 2: 879–884. Xue S, Barna M. 2012. Specialized ribosomes: a new frontier in gene regulation and organismal biology. Nat Rev Mol Cell Biol 13: 355–369. Yang T-H, Wang C-Y, Tsai H-C, Liu C-T. 2021. Human IRES Atlas: an integrative platform for studying IRES-driven translational regulation in humans. Database 2021: baab025. Ying Y, Qi X, Zhao GQ. 2001. Induction of primordial germ cells from murine epiblasts by synergistic action of BMP4 and BMP8B signaling pathways. Proc Natl Acad Sci U S A 98: 7858–7862. Ying Y, Zhao GQ. 2001. Cooperation of endoderm-derived BMP2 and extraembryonic ectoderm-derived BMP4 in primordial germ cell generation in the mouse. Dev Biol 232: 484–492. Yoon C, Kawakami K, Hopkins N. 1997. Zebrafish vasa homologue RNA is localized to the cleavage planes of 2- and 4-cell-stage embryos and is expressed in the primordial germ cells. Dev Camb Engl 124: 3157–3165. Young DW, Hassan MQ, Pratap J, Galindo M, Zaidi SK, Lee S, Yang X, Xie R, Javed A, Underwood JM, et al. 2007. Mitotic occupancy and lineage-specific transcriptional control of rRNA genes by Runx2. Nature 445: 442–446. Yuan X, Zhao J, Zentgraf H, Hoffmann-Rohrer U, Grummt I. 2002. Multiple interactions between RNA polymerase I, TIF-IA and TAFI subunits regulate preinitiation complex assembly at the ribosomal gene promoter. EMBO Rep 3: 1082–1087. Yusupova G, Yusupov M. 2014. High-Resolution Structure of the Eukaryotic 80S Ribosome. Annu Rev Biochem 83: 467–486. Zhang S, Shi M, Hui C-C, Rommens JM. 2006. Loss of the mouse ortholog of the shwachman-diamond syndrome gene (Sbds) results in early embryonic lethality. Mol Cell Biol 26: 6656–6663. Zhang Y, Duc A-CE, Rao S, Sun X-L, Bilbee AN, Rhodes M, Li Q, Kappes DJ, Rhodes J, Wiest DL. 2013. Control of hematopoietic stem cell emergence by antagonistic functions of ribosomal protein paralogs. Dev Cell 24: 411–425. Zhang Y, Lu H. 2009. Signaling to p53: ribosomal proteins find their way. Cancer Cell 16: 369–377. Zhang Y, O’Leary MN, Peri S, Wang M, Zha J, Melov S, Kappes DJ, Feng Q, Rhodes J, Amieux PS, et al. 2017. Ribosomal Proteins Rpl22 and Rpl22l1 Control Morphogenesis by Regulating Pre-mRNA Splicing. Cell Rep 18: 545–556. Zhao J, Li Y, Wang C, Zhang H, Zhang H, Jiang B, Guo X, Song X. 2020. IRESbase: A Comprehensive Database of Experimentally Validated Internal Ribosome Entry Sites. Genomics Proteomics Bioinformatics 18: 129–139. 89 Chapter II Oogenesis has proved the most difficult problem that I had hitherto attacked, and at one time I despaired of ever unravelling the intricate story of the origin and nature of the complicated granulations of the oocyte. – James Brontë Gatenby 1922 90 Chapter II preface Only a highly specialized oogonial cell, aka egg, has the competence to form an embryonic body and, consequently, the responsibility of transmitting hereditary information to the next generation. Gene expression regulatory mechanisms in egg cells are of particular importance to the propagation of a species. Therefore, these processes are thought to be under heightened selective pressure – favoring adaptation and possibly optimization of actively expressed genes in the egg. As explained in Chapter I, in all animals, this includes rDNA genes – the substrates for ribosome biogenesis and templates for ribosome structure. In 2017, Locati and Pagano and colleagues reported on a previously unknown rDNA gene in zebrafish; the rRNA expressed in somatic cells differs from rRNAs maternally deposited in eggs. The heightened selective pressure mentioned above may act specifically on the maternal rDNA gene. Adaptation may act on variation in the rDNA gene template, leading to compositional or structural heterogeneity during biogenesis, and ultimately to ribosome-specific functional heterogeneity. Two prominent hypotheses regarding the effects of ribosome heterogeneity on gene expression are presented in Chapter I. Notwithstanding, my rationale for comparing maternal and somatic ribosomes stems from observing two unique rDNA loci, expressed differentially within the zebrafish across time and space. Chapter II contains our report on the recently discovered cell-type restricted rDNA gene variant in zebrafish. The timing and location of rRNA expression in germ cells and somatic tissues is further characterized. RP composition and sub- stoichiometric ribosome associated factors are given for maternal and somatic ribosomes. Through this investigation high resolution atomic structures of ribosomes at these times of development were generated, and are provided. The two ribosomes only coincide in early development; where we demonstrate the capacity of our maternal and somatic ribosome-specific transgenic tagging method to address cognate and hybrid subunit compatibility. Lastly, we show maternal ribosomes do translate different sets of mRNAs in the developing embryo; preferentially enriched with cell type-specific transcripts. We conclude zebrafish ribosomes in germ cells and somatic cells are compositionally different, structurally distinct, and functionally compatible. In this chapter of my Thesis, I present the major findings of my doctoral research. My motivation was to develop methodologies and protocols for evaluating maternal and somatic ribosomes in zebrafish development both in germ and somatic cells. In this chapter we address the following: A. Are the two different? physically à We measure compositions and structures of each functionally à We measure a specific target for translation activity by each B. Does the system use two? individually à We measure each for translation activity in the system compatibly à We measure each for subunit rearrangement of transgenic labels 91 Collaboration notes In 2019, just before the onset of a global health crisis, we learned that Friederike “Frieda” Leesch, a doctoral candidate in Andrea Pauli's lab at the IMP in Vienna, was considering the same question in the same system by using similar probing methods. The “competitive” nature of our fields and the contemplation of potentially being outpaced in our research during a lockdown was initially discouraging. However, in our first call with each other, the two of us immediately recognized the benefits of collaborating with someone who had been reading the same literature and investigating the same system. After several in-depth discussions defining our distinct perspectives of the system, over quarantines and through 2023, Frieda and I recognized that merging our findings revealed an intriguing narrative. By considering measurements from both of our perspectives, we could clarify the role ribosome heterogeneity might be playing in embryonic development. Although the draft report included as the contents of this chapter is yet to be published and some data are still pending inclusion, we chose to integrate this narrative into our respective Theses. Our data collectively redefine the rDNA variant on chromosome 4 of the domesticated zebrafish, Danio rerio, as a germ cell-specific rDNA gene, confirm the usage of the variant in germ cell-specific ribosome biogenesis, and demonstrate the resulting ribosomal subunit products, maintain their germ cell-specificity. I consider our findings and conclusions to epitomize the parable of the blind and the elephant (see Note I). It underscores the advantages secured by the open exchange of ideas and data. Two PhD students on opposite sides of the world, both asking similar questions about the contribution of maternal material into eggs, each apply unique perspectives to the same system at the same times of development, plan the same assays (even design the same primers!) to interrogate the same two ribosomes, end up replicating each other’s findings, combining data from various methods, and working together to tell a more complete story about the complexities of a germ cell-specific ribosome in zebrafish. This scientific engagement has proved to be indispensable for progressing research into this biological system as well as for our respective trainings as doctoral candidates. 92 Collaboration statement on general contributions We each independently assessed ribosome compositions by MS, translational activity by polysome profiling, and rRNA expressions of both maternal and somatic ribosome types by assaying PCR fragment length polymorphisms in various tissues and across developmental time. While Frieda and colleagues solved the structures of either ribosome, unresolved regions involving ribosome intersubunit bridges still prevented the confirmation of a hybrid subunit arrangement. I transgenically engineered two zebrafish lines to maternally label or somatically label 60S subunits, and used each to experimentally show interchangeability among subunits. I labeled and sorted primordial germ cells for measuring rRNA expression and analyzed associated short read NGS data. I performed the RIP measuring germ cell-specific mRNA enrichments of each ribosome. Collaboration statement on specific contributions The order of findings and specific claims presented in this manuscript were initially drafted by Frieda. We asynchronously wrote the text. I generated all schematics, diagrams, plots, and cartoons as well as arranged all figures, tables, and supplementary information seen in Chapter II. Frieda provided the images seen in Fig. 2.3, Fig. 2.S6C, Fig. 2.S6D, and Fig. 2.S7 as well as primary data seen in other panels. Data contributed by either author are separated by the panel each are presented in: Data generated at MIT. Data generated at IMP. Contributed by A.N.S. Contributed by F.L. Fig. 2.1A, 2.1D Fig. 2.1B, 2.1C Fig. 2.4 Fig. 2.2 Fig. 2.5 Fig. 2.3 Fig. 2.6 Fig. 2.S2A, 2.S2B Fig. 2.S1 Fig. 2.S5 Fig. 2.S2C Fig. 2.S6 Fig. 2.S3 Fig. 2.S7 Fig. 2.S4 Fig. 2.S8 Fig. 2.S9 Fig. 2.S10 Fig. 2.S11 Fig. 2.S12 93 A dual ribosomal system in the zebrafish soma and germline Arish N. Shah3,#, Friederike Leesch1,2,#, Laura Lorenzo-Orts1, Lorenz E. Grundmann1,2, Carina Pribitzer1, Maria Novatchkova1, Irina Grishkovskaya1, David Haselbach1, Eliezer Calo3,4,*, and Andrea Pauli1,* 1 Research Institute of Molecular Pathology, Vienna BioCenter, Vienna, Austria. 2 Vienna BioCenter PhD Program, Doctoral School of the University of Vienna and Medical University of Vienna, Vienna, Austria. 3 Department of Biology, Massachusetts Institute of Technology, Cambridge, United States. 4 David H. Koch Institute for Integrative Cancer Research, Massachusetts Institute of Technology, Cambridge, United States. # These authors contributed equally * Lead authors 94 Abstract Protein synthesis during vertebrate embryogenesis is driven by ribosomes of two distinct origins: maternal ribosomes synthesized during oogenesis and stored in the egg, and somatic ribosomes, produced by the developing embryo after zygotic genome activation (ZGA). In zebrafish, these two ribosome types are expressed from different genomic loci and also differ in their ribosomal RNA (rRNA) sequence. To characterize this dual ribosome system further, we examined the expression patterns of maternal and somatic rRNAs during embryogenesis and in adult tissues. We found that maternal rRNAs are not only expressed during oogenesis but are continuously produced in the zebrafish germline. Proteomic analyses of maternal and somatic ribosomes unveiled differences in core ribosomal protein composition. Cryo-EM structures of maternal and somatic ribosomes revealed no remarkable differences between the core of these two ribosome types, despite the nucleotide differences observed in the structures. Our structural and in vivo data demonstrate that both maternal and somatic ribosomes can be translationally active in the embryo. Using transgenically tagged maternal or somatic ribosome subunits, we experimentally confirm the presence of hybrid 80S ribosomes composed of 40S and 60S subunits from both origins and demonstrate the preferential in vivo association of maternal ribosomes with germline-specific transcripts. Our study identifies a distinct type of ribosomes in the zebrafish germline and thus presents a foundation for future explorations into possible regulatory mechanisms and functional roles of heterogeneous ribosomes. 95 Introduction Ribosomes are macromolecular complexes composed of ribosomal RNAs (rRNAs) and proteins that are responsible for protein synthesis in the cell. Ribosomes of two different origins are present during embryogenesis. Large numbers of maternal ribosomes are synthesized during oogenesis and stored in the egg. Upon fertilization, maternal ribosomes are solely responsible for all translational activity during early embryogenesis (Bazzini et al. 2016; Hensey and Gautier 1997). Maternal ribosomes are progressively degraded during embryogenesis and replaced by newly synthesized ribosomes (i.e. somatic ribosomes) that are produced after zygotic genome activation (ZGA) in the embryo (Heyn et al. 2017; Schramm and Bavister 1999). In addition to their roles in translation (Bazzini et al. 2016; Cenik et al. 2019; Hensey and Gautier 1997; Noack Watt et al. 2016), maternal ribosomes have been proposed to act as a source of pyrimidines required for the synthesis of new ribosomes (Liu et al. 2018b). In the absence of transcription, the synthesis of new proteins in the early vertebrate embryo needs to be regulated at the post-transcriptional level. Several mechanisms have been described to regulate translation during early embryogenesis, including mRNA localization (Kloc and Etkin 2005; Lorenzo-Orts et al. 2023; Maegawa et al. 1999; Medioni et al. 2012), binding of proteins or miRNAs to mRNAs (Giraldez et al. 2006), and regulation of polyA tail length (Eichhorn et al. 2016; Lim et al. 2016; Liu et al. 2023; Subtelny et al. 2014). While much effort has been devoted to understanding maternal mRNA regulation in the past, we have recently discovered that maternal ribosomes associate with a specific set of factors that bind to key sites on the ribosome and contribute to its repression and stability (Leesch et al. 2023). Thus, control of maternal ribosome function also plays an important role in translational regulation during embryogenesis. While ribosomes have traditionally been considered homogeneous macromolecular complexes with no regulatory role in translation, several studies proposed that ribosomes are heterogeneous in their composition (Ferretti et al. 2017; Komili et al. 2007; Mageeney and Ware 2019; O’Leary et al. 2013; Shi et al. 2017; Simsek et al. 2017; Xue et al. 2015). Ribosome compositional heterogeneity is thought to be caused by variations in ribosome accessory factors, ribosome proteins, and ribosomal RNA, as well as by modifications of nucleotides and amino acids. For instance, distinct rRNAs have been reported in Plasmodium berghei ribosomes isolated from different life cycle stages (Gunderson et al. 1987; Waters et al. 1997), and in ribosomes from Xenopus oocytes (Peterson et al. 1980). Moreover, in Drosophila, ribosomes isolated from the germline and soma showed a different protein composition (Hopes et al. 2022). Although heterogeneous ribosomes exist in various developmental and cellular contexts, the functional implications of ribosome heterogeneity remain controversial (Barna et al. 2022; Ferretti and Karbstein 2019). In zebrafish, maternal and somatic ribosomes differ in their composition (Locati et al. 2017a, 2017b), which allows studying these specific ribosome types in an experimentally well-established vertebrate model organism. Maternal ribosomes differ in all four rRNA 96 sequences (28S, 18S, 5.8S, and 5S) and originate from a different ribosomal DNA (rDNA) locus compared to somatic ribosomes (Locati et al. 2017a, 2017b). Maternal rRNAs are transcribed during oogenesis from a single rDNA cluster located on chromosome 4 (Locati et al. 2017b), which is methylated and thus transcriptionally silenced in the soma (Ortega-Recalde et al. 2019). Maternal rRNAs differ from somatic rRNAs in up to 14% of their sequence. However, the consequences of these differences on ribosome functions are not yet known. Furthermore, it remains unclear whether maternal and somatic ribosomes differ in protein composition and structure. Here we analyze the expression, protein composition, and structure of maternal and somatic ribosomes in zebrafish. Our results show that despite the differences in rRNA sequences, maternal and somatic ribosomes are found in polysomes indicating their translational activity, have similar core structures, and can form hybrid ribosomes containing subunits of both origins. Further, we identify germ cells as a cell type that continues to express maternal-type ribosomes throughout development, suggesting a potential functional importance in germ cell-specific translation. 97 Results Expression of rRNA variants across development and cell types in zebrafish To investigate the expression of maternal and somatic rRNA variants during zebrafish development, we analyzed total RNA during the first five days of embryogenesis. Consistent with previous findings (Locati et al. 2017a, 2017b), we observed that maternal rRNAs originating from the maternal rDNA locus on chromosome 4 (GRCz11; 4: 77,564,140 – 77,555,053) were the sole rRNA variants present in the egg and during early stages of embryogenesis before zygotic genome activation (ZGA). Following ZGA, somatic rRNAs started to be transcribed from the somatic rDNA locus on chromosome– 5 (GRCz11; 5: 827,807 – 819,029). The levels of somatic rRNAs increased progressively and reached the same level as maternal rRNAs between 20 to 30 hours post-fertilization (hpf). By 5 days post-fertilization (dpf), more than 98% of the detected rRNAs were of somatic origin, reflecting the transition from maternal to somatic ribosomes (Fig. 2.1A and 2.1B). Although most ribosomes are of somatic origin by day 5 of development, maternal rRNA variants may be expressed in specific cell types or tissues beyond oocytes. To investigate this, we employed a PCR-based assay that could discriminate between maternal and somatic rRNA variants based on the amplification of highly variable Expansion Segments (ESs) in 18S and 28S RNAs (Supplementary Fig. 2.S1A). To confirm the applicability of the PCR-based assay, we recapitulated the transition from maternal to somatic 18S rRNA (Supplementary Fig. 2.S1B and 2.S1C). While in most adult tissues a single DNA band corresponding to somatic 18S and 28S rRNAs was detected, testis samples showed evidence for the presence of both somatic and maternal rRNAs (Supplementary Fig. 2.S2). Next-generation sequencing (NGS) experiments using total RNA from testes dissected from five different adult male zebrafish revealed variable levels of maternal and somatic rRNAs, with maternal rRNA variants accounting for 2% to 27% of the total rRNAs (Fig. 2.1C). Together, these results suggest that both maternal and somatic ribosomes are present in the male germline. Recent interrogation of human cancer biopsies in The Cancer Genome Atlas (TCGA) (Liu et al. 2018a) has similarly shown differential expression of rRNA variants (Rothschild et al. 2023). To test whether zebrafish maternal ribosomes might also be re-expressed in immortalized cells or during tumorigenesis, we analyzed cDNAs from several cultured zebrafish cell lines and excised adult tumors (Berghmans et al. 2005; Driever and Rangini 1993; Heilmann et al. 2015; Patton et al. 2005; Paw and Zon 1999; Perez et al. 2018). 18S rRNA compositional analysis in the assayed ex vivo or neoplastic contexts showed no evidence of reactivated maternal rRNA (Supplementary Fig. 2.S3A and 2.S3B). Similarly testing regenerating fin blastema through 5 days post amputation indicates dedifferentiated cells losing morphology and re-entering the cell cycle (Pfefferli and Jaźwińska, 2015) do not express maternal rRNA (Supplementary Fig. 2.S3C). To gain deeper insights into the expression of maternal rRNAs in the germline, we re- analyzed total RNA sequencing data from whole embryos and larvae, as well as 98 published total RNA sequencing data from FACS-sorted primordial germ cells (PGCs) during larval development up to 10 dpf (Redl et al. 2021). PGCs, which are the precursors of the adult germline, showed persistently high levels of maternal rRNAs compared to somatic larval tissues throughout 10 dpf (Fig. 2.1D). Consistent with a potential role for maternal ribosomes in the germline, expression analysis of maternal ITS1 (internal transcribed spacers, Fig. 1A), which are removed from the nascent 47S rRNA during ribosome biogenesis, indicated active transcription of maternal rRNAs in PGCs, but not in the soma, from 3 dpf (Supplementary Fig. 2.S4), long before the production of early- stage oocytes between 10-25 dpf in juveniles (Dranow et al. 2016; Takahashi 1977). Figure 2.1: Expression of maternal and somatic rRNA variants in zebrafish A) Organization of maternal (yellow) and somatic (blue) rDNA genes in the zebrafish genome. The 5S rRNA genes are encoded separately, while 18S, 5.8S, and 28S are derived from a single 47S pre-rRNA transcript containing additional spacer sequences (ITS1 and ITS2 (internal transcribed spacers)) that are removed in several processing steps. Expansion segments (ES3S and ES31L) are indicated by lighter-colored boxes and PCR primer icons (see Supplementary Fig. 2.S1A). B) Relative expression levels of the four rRNAs (28S, 5.8S, 18S, and 5S), comparing maternal (yellow) and somatic (blue) variants. C) Fraction of maternal versus somatic rRNAs, obtained by quantifying reads with variant-specific SNPs (single nucleotide polymorphisms) during the first five days of development, in adult somatic organs, and in testes of 5 different males. D) Expression of maternal and somatic rRNA variants in whole embryo lysate and FACS-sorted PGCs. ES, expansion segment. hpf, hours post-fertilization. FACS, fluorescence activated cell sorting. 99 Despite our evidence for persistent expression of the rDNA locus on chromosome 4 in the zebrafish germline, for simplicity and consistency with previous publications, we will continue to refer to the rRNAs, the subunits they compose, and the ribosomes generated from this locus as “maternal” throughout this manuscript. Maternal and somatic ribosomes differ in protein composition Maternal and somatic ribosomes have so far only been shown to differ in their rRNAs. To investigate potential differences in protein composition, we performed proteomic analyses of ribosomes isolated at different times during embryogenesis and compared the expression of core ribosomal proteins (RPs). We chose time points containing either only maternal ribosomes (egg, 1 hpf, 3 hpf and 6 hpf), similar amounts of maternal and somatic ribosomes (24 hpf) and only somatic ribosomes (120 hpf) (Fig. 2.1B and Fig. 2.2A, bottom). We detected 8 RPs with more than one paralog associated to maternal or somatic ribosomes during embryogenesis, namely Rps27 (eS27), Rps8 (eS8), Rps26 (eS26), Rps17 (eS17), Rplp2 (P2), Rpl7 (uL30), Rpl5 (uL18) and Rpl22 (eL22) (Fig. 2.2A, top). Moreover, alternative isoforms were evident for Rps5 (uS7), Rps18 (uS13), Rps19 (eS19) and Rpl9 (uL6) (Fig. 2.2A). To gain further insights into the expression patterns of these alternative RPs, we analyzed previously published RNA-seq datasets covering different stages of oogenesis, embryogenesis, and adult tissues (Cabrera-Quio et al. 2021; Noda et al. 2022; Pauli et al. 2014) (Fig. 2.2B and Supplementary Fig. 2.S5). While only one protein variant was predominantly present for all RPs (Fig. 2.2A), mRNAs encoding for Rps27.1/Rps27.2, Rplp2/Rplp2l, Rpl5a/Rpl5b, and Rps17 were expressed at similar levels during development and in adult tissues (Fig. 2.2B). In some cases, one RP paralog was associated with purified ribosomes at all stages while the other was ribosome-bound only at specific times. Examples include Rpl22 and Rpl5b, which were associated with ribosomes throughout embryogenesis, whereas their paralogs Rpl22l1 and Rpl5a, respectively, only showed increased ribosome incorporation at 5 dpf, correlating with the expression of somatic rRNA variants (Fig. 2.2B and Fig. 2.2C). This proteomic analysis of maternal and somatic ribosomes during embryogenesis provided evidence for RP composition differences in assembled ribosomes at different times of development, but did not reveal a single RP paralog pair that mirrored the switch between maternal and somatic rRNAs. 100 Figure 2.2: Expression of alternative ribosomal core proteins in zebrafish development A) Heatmap of protein expression (normalized IBAQ values) for variant ribosomal protein pairs (paralogs, top; alternative isoforms, bottom) associated with zebrafish ribosomes at different developmental stages. Below, the presence of maternal (yellow) or somatic (blue) rRNA variants at these developmental stages is indicated separately. B) Heatmap of RNA expression of paralogs and alternative isoforms for ribosomal 101 protein variants during embryogenesis. See also Supplementary Fig. 2.S5. The relative presence of maternal (yellow) or somatic (blue) rRNA variants at these developmental stages is indicated in the scheme below the heat map. C) Volcano plot based on mass spectrometry data showing fold enrichment of proteins in the ribosome fraction of 5 dpf larvae versus 6 hpf embryos (n = 3 for each time point). Pairs of ribosomal protein variants with significantly different associations are shown in different shades. Non-significant but detected variants are shown in green. Permutation-based false discovery rates (FDRs) are shown as dotted (FDR < 0.01) and dashed (FDR < 0.05) lines. All significantly enriched or depleted proteins are listed in Supplementary Table 2.S1. hpf, hours post fertilization. dpf, days post fertilization. TPM, transcripts per million. iBAQ, intensity Based Absolute Quantification. Cryo-EM reveals similar core structures of maternal and somatic ribosomes rRNAs are known to not only provide the structural framework of the ribosome but also form many of its core functional sites. Since maternal and somatic ribosomes contain different rRNAs and RPs, we used Cryo-EM to investigate how these differences affect the ribosome structure. We isolated somatic ribosomes from zebrafish larvae at 5 dpf and obtained a 3.5 Å structure (Supplementary Fig. 2.S6, 2.S7, and Supplementary Table 2.S2), which we then compared to a previously published structure of the maternal zebrafish ribosome at 6 hpf (Fig. 2.3A). We specifically chose to compare ribosomes from these two time points because the majority of ribosomes contained rRNAs from one genomic location (maternal versus somatic rRNAs) and were in a similar translational state (Leesch et al. 2023), thus allowing potential structural differences to be attributed to differences in rRNAs. In spite of the differences observed in rRNA and protein composition, maternal and somatic ribosomes did not show major structural differences (Fig. 2.3A). This is likely due to the fact that the majority of sequence differences between maternal and somatic rRNAs are located in expansion segments (ESs), which are highly flexible RNA helices on the surface of the ribosome that cannot be resolved by cryo-EM (Fig. 2.3B). While we were able to see 27% of the total differences between maternal and somatic rRNA in our structures, most of these differences did not alter the structure (Supplementary Table 2.S3). Some nucleotide differences were present in the two parallel sides of a helix, thus preserving the interaction between two nucleotides. For instance, both strands of helix 12 (H12) of the 28S rRNA differ in maternal and somatic ribosomes, with the somatic variant containing a U (U68) and the maternal sequence containing a C (C68). In the opposite strand, the maternal variant encodes a compensatory G73, whereas the somatic variant contains A73, thereby maintaining the Watson-Crick interaction (U-A in the somatic rRNA, C-G in the maternal rRNA). We also found differences in nucleotides that were not base paired in maternal and somatic ribosomes, such as 4 nucleotides located at the tip of H11 in the 18S rRNA. Although these nucleotides interact with Rps8a, we did not observe any differences in Rps8a binding. To investigate whether the presence of maternal rRNAs in the ribosome may affect ribosomal function, we first examined structural differences in functionally important sites of the ribosome, such as the sarcin-rich domain (SRD, or sarcin-rich loop, SRL) (Anger et al. 2013; Khatter et al. 2015). The SRL contains two nucleotide differences (Usom3716/Cmat3864 and Asom3738/Gmat3886) located at the helix that forms the base of 102 the SRL stem-loop (Fig. 2.3B), which is important for anchoring elongation factors during tRNA translocation (Shi et al. 2012; Szewczak et al. 1993). We did not observe any structural differences in the SRLs of maternal and somatic ribosomes, as Watson-Crick base pairing was maintained in the RNA helix. Other functionally important regions like the GTPase association center (GAC), which acts as a binding platform for elongation factors and ribosome rescue factors (Uchiumi and Kominami 1994), were not resolved in our cryo-EM datasets in the region containing three nucleotide differences between maternal and somatic 28S RNAs (Asom1558/Cmat1573, Usom1560/Cmat1575 and Usom1565/Cmat1580). In a second line of investigation, we analyzed whether the differences between maternal and somatic rRNAs corresponded to evolutionarily variable nucleotides because functionally important nucleotides tend to be conserved throughout evolution. We found that the Shannon entropy index (Schmitt and Herzel 1997) was higher than 0.8 for most nucleotides, indicating a high degree of variability at the single nucleotide level across evolution (Supplementary Table 2.S3). Together, our cryo-EM analyses of maternal and somatic ribosomes show that despite the substantial amount of nucleotide sequence differences, the core ribosomal structures are conserved. 103 Figure 2.3: Location of sequence and structural differences in rRNA variants A) Maps of zebrafish ribosomes isolated from 6 hpf embryos (left, maternal ribosome (grey)) and 5 dpf larvae (right, somatic ribosome (green)). B) Sequence differences in maternal and somatic rRNA variants mapped to the rRNA secondary structure prediction. Sequence differences modeled in cryo-EM structures obtained from 6 hpf zebrafish embryos and 5-day-old larvae (circles indicate sequence differences). Regions not modeled due to poor density are shown in grey. Sequence differences in modeled regions were classified based on interactions with other nucleotides and proteins and highlighted in different colors (see inset legend). C) A single region with structural changes involving an rRNA sequence difference was observed. The region is indicated in (B) with a pink circle and dashed line. hpf, hours post fertilization. dpf, days post fertilization. 104 Formation of hybrid ribosomes comprising maternal and somatic subunits To investigate whether maternal and somatic ribosomal subunits may interact and form 80S “hybrid” ribosomes, we focused our structural analyses on rRNA sequence differences and alternative ribosomal proteins in the contact regions between 40S and 60S subunits, termed intersubunit bridges (see Table 2.1). 40S and 60S subunits interact through 17 bridges (B1a-eB14) (Anger et al. 2013; Ben-Shem et al. 2011; Melnikov et al. 2012). Most of these bridges are formed by RNA-RNA or RNA-protein contacts, with the exception of B1b/c, which is formed by the contact of two ribosomal proteins, Rps18 (uS13) and Rpl11 (uL5). Five bridges are specific for eukaryotic ribosomes (eB8-eB14). On the rRNA level, sequence differences in the intersubunit bridges B3-B6 and eB13 did not result in structural changes, while differences in ES31L, ES41L, and ES6S (Fig. 2.3B), which form the eukaryotic-specific intersubunit bridges eB8, eB11, eB12, and eB14, were not resolved in our ribosome structures and could therefore not be assessed (Table 2.1). In addition to rRNA sequence differences, one ribosomal protein, Rps8 (eS8), known to form an intersubunit bridge (Tamm et al. 2019) has a paralogous RP and both are expressed in early zebrafish embryogenesis (Fig. 2.2A). However, in our data, Rps8a was more abundant at the protein and mRNA level in both maternal and somatic ribosomes, suggesting no difference in RP paralog usage at the intersubunit bridge between maternal and somatic ribosomes (Fig. 2.2A and Fig. 2.2B). Therefore, our analysis provided no evidence for structural differences that could prevent the formation of hybrid ribosomes. Table 2.1: Structural differences of intersubunit bridge regions in maternal and somatic ribosomes Intersubunit bridges (listed in the first column) are connections between 40S SSU (composed of the 18S rRNA) and 60S LSU (composed of the 5S, 5.8S, and 28S rRNAs). Bridges composed of rRNA sequences and r-proteins with differences between maternal and somatic types are marked. Bridges composed of poorly modeled rRNA regions are marked in grey. H, helix. ES, expansion segment. 105 To further investigate the possibility of hybrid ribosome formation in vivo, 24 hpf zebrafish larvae were used since maternal and somatic rRNA variants are present at equal levels at this developmental stage (Fig. 2.1B). To assess whether maternal ribosomes can be found in polysomes at this developmental stage, we performed polysome gradients of total lysates, which showed peaks for the 40S and 60S ribosomal subunits, a large 80S (monosome) peak and several polysome peaks (Fig. 2.4A). Besides detecting maternal and somatic rRNA types in the fraction corresponding to 40S, 60S and 80S subunits, we identified both rRNA types in translationally active polysomes (Fig. 2.4B), suggesting that both maternal and somatic ribosomes support translation at this stage of embryogenesis. Figure 2.4: Maternal and somatic ribosomal subunits are translationally active in 24 hpf embryos A) Representative polysome profile with continuous UV absorbance (A260) reading from sucrose density gradients containing lysate from 24 hours post-fertilization (hpf) zebrafish embryos. Position in the gradient and collected fractions are indicated. B) Gel electrophoretic analysis of PCR-based detection of maternal and somatic small (18S) and large (28S) rRNA variants in the individual factions indicated in (A). See Supplementary Table 2.S4 for lengths of PCR products from maternal and somatic rRNAs. hpf (hours post-fertilization), bp (base pair), H (helix), ES (expansion segment). 106 To achieve selective labeling of maternal and somatic ribosomal subunits, FLAG-tagged Rpl10a was specifically expressed during oogenesis via the oogenesis-specific promoter sycp (synaptonemal complex protein) (Tg(Mat-RiboFLAG), ensuring the restricted incorporation into maternal ribosomes. In contrast, a male-contributed transgene using a ubiquitously expressed promoter ubi (ubiquitin b) (Tg(Som-RiboFLAG)) enabled specific labeling of somatic ribosomes (Supplementary Fig. 2.S8, 2.S10, 2.S11). To validate our experimental setup (Supplementary Fig. 2.S9), we performed FLAG- immunoprecipitation (FLAG-IP) experiments using maternal and somatic FLAG-tagged Rpl10a embryo lysates treated with EDTA to dissociate 80S ribosomes into individual subunits. By analyzing the relative abundance of maternal and somatic 28S rRNA in the input and IP fractions using RT-qPCR, we confirmed the specificity of our tagging, precipitation, and detection approaches (Fig. 2.5A and 2.5B). Figure 2.5: In vivo evidence for hybrid ribosome formation in 1 day post-fertilization embryos A) Scheme summarizing the generation of embryos with tagged ribosomes by crossing Tg(Mat-RiboFLAG) females to wildtype males and Tg(Som-RiboFLAG) males to wildtype females. B) RT-qPCR analysis of the relative amounts of maternal and somatic 28S rRNAs detected in input or eluate (IP) fractions of a FLAG- IP experiment, which used EDTA-treated lysates containing either tagged maternal (Tg(Mat-RiboFLAG)) or somatic (Tg(Som-RiboFLAG)) 60S subunits as input. C) RT-qPCR analysis of the relative amounts of maternal and somatic 28S and 18S rRNAs detected in input or eluate (IP) fractions of a FLAG-IP experiment. Cycloheximide (CHX) was added during lysis of embryos containing either tagged maternal (Tg(Mat-RiboFLAG)) or somatic (Tg(Som-RiboFLAG)) 60S subunits. In (B) and (C), significance was calculated by t-test (if not indicated, p-value > 0.05). 107 To test whether hybrid ribosomes form in vivo, we performed a second FLAG-IP experiment in the presence of cycloheximide (CHX), a well-established translation inhibitor known to stabilize translating ribosomes. This allowed us to assess the presence of maternal or somatic small subunits (SSUs) in the IP fraction of the tagged maternal or somatic LSUs (Fig. 2.5C). Intriguingly, we observed similar levels of maternal and somatic SSUs in the IP fractions independently of whether maternal or somatic tagged LSUs had served as baits. These findings provide direct experimental evidence supporting the in vivo formation of hybrid ribosomes. Co-enrichment of maternal ribosomes with PGCs and PGC-specific mRNAs Building on our observation that maternal rRNAs are continuously expressed in PGCs (Fig. 2.1D and Supplementary Fig. 2.4B) and that maternal ribosomes are actively translating in 1-day-old zebrafish embryos, we reasoned that maternal ribosomes should be present in PGCs and may be translating PGC-specific transcripts. To test these hypotheses, we injected in vitro transcribed eGFP mRNA fused to the nanos3-3'UTR into the 1-cell stage embryo. The nanos3-3’UTR stabilizes the transcript only in PGCs, enabling PGC-specific eGFP translation at 1 dpf (Fig. 2.6A, 2.6B). To confirm that PGCs maintain a higher ratio of maternal versus somatic ribosomes at 1 dpf (Fig. 2.1D), we used RT-qPCR to assess the ratio of maternal and somatic rRNA variants in FACS-isolated eGFP-positive PGCs and non-eGFP expressing somatic cells. As expected, a higher abundance of maternal 18S and 28S rRNA variants were detected in the isolated PGCs compared to somatic cells, confirming the persistence of maternal ribosomes at higher levels in germline cells (Fig. 2.6C). To directly test whether maternal ribosomes associate with PGC-specific transcripts and thus may contribute to germ cell- specific translation, we performed a RNA immunoprecipitation (RIP) assay using biotinylated eGFP-nanos3-3'UTR mRNAs extracted from 1-day-old cycloheximide- treated larvae. This approach allowed us to analyze the pool of ribosomes bound to this specific PGC-localized mRNA. Notably, we observed a significant enrichment of the maternal ribosomes compared to somatic ribosomes associated with biotin-labeled PGC- localized mRNAs (Fig. 2.6D). However, this level of enrichment was similar to the level of enrichment of maternal ribosomes in isolated PGCs (Fig. 2.6C), providing no evidence for selectivity of maternal ribosomes in associating with germ cell-specific transcripts. Nevertheless, our results reveal a co-occurrence between maternal ribosomes and germline-specific transcripts in PGCs. 108 Figure 2.6: Enrichment of PGC-localized mRNAs with maternal ribosomes at 1 day post-fertilization A) Schematic of the experimental strategy. eGFP-nanos3’UTR containing mRNA was injected into 1-cell stage zebrafish embryos from which two different experimental approaches were used to investigate the ratio of maternal and somatic ribosomes in PGCs and to test which ribosomes are bound to PGC-localized mRNAs. B) Brightfield and eGFP images of 1 day post-fertilization larvae, previously injected with no mRNA, eGFP-nanos3’UTR mRNA, or biotinylated eGFP-nanos3-3’UTR mRNAs. Scale bars indicate 500 μm. C) Ratio of maternal (yellow) versus somatic (blue) 18S and 28S rRNA in FACS-isolated eGFP-positive (PGCs) and eGFP-negative (somatic) cells analyzed by RT-qPCR. D) Ratio of maternal (yellow) versus somatic (blue) 18S and 28S variants associated with biotinylated eGFP-nanos3-3’UTR mRNAs isolated by RIP (RNA immunoprecipitation). Non-biotinylated eGFP-nanos3-3’UTR mRNA served as control. In (C) and (D), significance was calculated by t-test (if not indicated, p-value > 0.05). 109 Discussion Previous studies have revealed that zebrafish ribosomes of maternal and somatic origin are composed of two distinct sets of rRNAs (Locati et al. 2017a, 2017b). While one set of rRNAs was thought to be exclusively present in maternal ribosomes, our PCR-based and RNA-seq analyses indicate that these rRNAs are not only transcribed during oogenesis, but are also present in PGCs and the male germ line. Translation in PGCs is tightly regulated and differs from that in the soma, as these cells are specified before the onset of zygotic genome activation (ZGA) and contain specific maternal mRNAs (Dranow et al. 2016; Johnson et al. 2011; Yu et al. 2016). Through at least 10 dpf, both inherited and zygotically generated maternal ribosomes constitute the major ribosome type in PGCs. Therefore, maternal ribosomes may comprise a distinct set of ribosomes that drive translation in the germline - a finding that may have broad implications for understanding germline determination and development in zebrafish. Heterogeneous rRNA sequences have been described in various organisms, including fungi, invertebrates (Dimarco et al. 2012), Xenopus, and humans (Pasolini et al. 2006; Vierna et al. 2013). However, whether ribosome heterogeneity translates into specialized ribosomal functions is still a matter of scientific debate. Apart from Plasmodium species (Gunderson et al. 1987; Li et al. 1997; Waters et al. 1997) and the 5S rRNA in Xenopus (Peterson et al. 1980), zebrafish embryos are the only known system that contains two distinct ribosomes composed of different sets of rRNAs. Differential expression of rRNA variants between tissues may indicate the ability of rRNA variant ribosomes to have distinct functions. Therefore, zebrafish embryogenesis, especially germline versus soma studies, provides a powerful system to study ribosome-mRNA interactions in the context of the functional consequences of ribosome heterogeneity. However, the cell-to-cell variability in their abundance requires future techniques to address these questions at the single cell level. Importantly, we observed that at 1 dpf, when maternal and somatic ribosomes are present at similar levels, both types of ribosomes associate with polysomes and are thus likely involved in translation. Consistent with the role of maternal ribosomes in supporting early development, translation by maternally deposited ribosomes is essential and increases during embryogenesis before the onset of ribosome biogenesis (Bazzini et al. 2016; Hensey and Gautier 1997; Leesch et al. 2023). Notably, in nematode worms lacking zygotic production of ribosomes (Cenik et al., 2016) and in zebrafish mutants with impaired ribosome biogenesis (Bielczyk-Maczyńska et al. 2015; Noack Watt et al. 2016), maternal ribosomes have been shown to be sufficient to sustain embryonic development. If both maternal and somatic ribosomes are capable of translation, an obvious question is why a dual ribosomal system has evolved in zebrafish. One hypothesis is that maternal ribosomes may favor the translation of certain transcripts. Fully addressing this question would require swapping the maternal and somatic rRNA locus, a challenging experiment in zebrafish. However, we did not observe major structural differences in the core of maternal and somatic ribosomes, suggesting that transcript specificity (if any) may be caused by the different expansion segments or solvent-exposed surfaces of maternal and 110 somatic ribosomes. In agreement with this, we observed 80S “hybrid” monosomes composed of 40S and 60S subunits of different origins. An alternative and non-exclusive hypothesis is that the maternal rDNA locus may be involved in PGC fate and sex determination in zebrafish. Zebrafish lack sex chromosomes and, unlike mammals, rely on the transfer of specific maternal factors for germ cell development. Several reports suggest a role for the maternal rDNA locus in sex determination, with specific changes (such as demethylation and amplification on extrachromosomal circles) occurring during oocyte expansion and being associated with feminization (Breit et al. 2020; Dranow et al. 2013; Ortega-Recalde et al. 2019; Tao et al. 2020). Another possibility is that the distinct set of rRNAs affects ribosome stability, as it has been suggested that maternal ribosomes are degraded more rapidly than their somatic counterparts (Locati et al. 2017b). A recent study proposed that ribosome ubiquitination plays a role in targeting and degrading maternal ribosomes during zebrafish embryogenesis (Ugajin et al., 2023). Additionally, one of the expansion segments, ES4L, is located on the ribosome surface and is formed by the 3' end of the 5.8S rRNA and the 5' end of the 28S rRNA. This 3' half of the 5.8S rRNA has previously been suggested to play a role in ribosome degradation (Locati et al., 2018), and therefore this region could potentially serve as a target for degradation of maternal ribosomes. Taken together, our study describes a distinct set of ribosomes present in the zebrafish germline. The presence of two distinct ribosomes during zebrafish embryogenesis provides an experimentally accessible system for future studies of ribosome specialization and defines a novel potential mechanism of translational regulation in the zebrafish germline. 111 Materials and Methods Zebrafish lines and husbandry Zebrafish (D. rerio) experiments in the Pauli lab were conducted according to Austrian and European guidelines for animal research and approved by local Austrian authorities (animal protocols for work with zebrafish: GZ 342445/2016/12 and MA 58-221180-2021- 16), and zebrafish lines in the Calo lab were housed in AAALAC-approved facilities and maintained according to protocols approved by the Massachusetts Institute of Technology Committee on Animal Care (CAC). Zebrafish transgenic lines containing maternally or somatically tagged ribosomes were generated as part of this study and are described below. In vivo samples were allocated randomly to the experiment and treated equally. TLAB fish, generated by crossing zebrafish AB with the natural variant TL (Tupfel Longfin), served as wild-type zebrafish for experiments in the Pauli lab (Fig. 2.1, 2.2, 2.3, 2.S2A, 2.S2B, 2.S5, 2.S6). All experiments in the Calo Lab were performed in the AB/Tübingen (TAB5/14) zebrafish genetic background (Fig. 2.4, 2.5, 2.6, 2.S1, 2.S2C, 2.S3, 2.S4, 2.S8, 2.S10, 2.S11). Fish were raised according to standard protocols (28 °C; 14:10 h light:dark cycle). Zebrafish embryo and tissue collection Zebrafish embryos were collected at the indicated times post-fertilization, and staged according to (Kimmel et al. 1995). Zebrafish AB9 fibroblasts, originally derived from an adult caudal fin (ATCC CRL-2298), were cultured on cell culture-treated 10 cm plates (Genesee Scientific, 25-500), collected using trypsin (Gibco, 25200072) and standard cell culture procedures. AB9 cells were grown at 28 °C wth 5% CO2 in Dulbecco’s Modified Eagle Medium (DMEM, Genesee Scientific, 25–500) supplemented with 10% fetal bovine serum (FBS, Gemini Bio- products) and 1% Penicillin/Streptomycin (Gibco, 10378–016). Zebrafish ZMEL melanoma cells, originally derived from tp53M214K/M214K tumors expressing human oncogenic BRAFV600E were cultured and collected using the same protocols as AB9 cells. ZMEL cells were a kind gift from the laboratory of Dr. Leonard Zon (Boston Children’s Hospital). Zebrafish ZF4 fibroblasts, originally derived from a 1 dpf embryo (ATCC CRL-2050), were cultured on cell culture-treated 10 cm plates (Genesee Scientific, 25-400), collected using trypsin (Gibco, 25200072) and standard cell culture procedures. ZF4 cells were grown at 28 °C with 5% CO2 in a 1:1 mixture of Dulbecco’s Modified Eagle Medium and Ham’s F- 12 media (DMEM/F12, Genesee Scientific, 25-503) supplemented with 10% fetal bovine serum (FBS, Gemini Bio-products) and 1% Penicillin/Streptomycin (Gibco, 10378–016). For all cell lines, cells from a 50-70% confluent 10 cm plate were collected and either flash frozen or directly lysed in TRIzol. 112 Zebrafish tumors used in this study were derived from the genotypes Tg(mitfa: GNAQQ209L); tp53M214K/M214K (referred to as GNAQ melanoma), Tg(BRAFV600E); tp53M214K/M214K (referred to as BRAF melanoma), and tp53M214K/M214K (referred to as MPNST, malignant peripheral nerve sheath tumor) and excised using standard protocols approved by the Committee on Animal Care at MIT. Adult zebrafish were monitored and euthanized before tumors (3-4 mm in diameter) were excised with a clean scalpel and placed into a 35 mm dish. Tumors were incubated with 2 mL of Dissection Media [Dulbecco’s Modified Eagle Medium (DMEM, Genesee Scientific, 25–500), 1% Penicillin/Streptomycin (Gibco, 10378–016), and 75 μg/mL of Liberase (Roche, 5401020001)], manually disaggregated with a clean razor blade, and let sit at room temperature until a cellular slurry formed. Tumors were further broken down by trituration with an additional 5 mL of Wash Solution [1x PBS (Genesee Scientific, 25– 508), 1% Penicillin/Streptomycin (Gibco, 10378–016), and 10% fetal bovine serum (FBS, Gemini Bio-products)]. Cells were passed through a 40 μm strainer (Falcon, 352340), pelleted for 2 in at 500 xg, washed twice with 1x PBS, pelleted again, and lysed in TRIzol. Generation of transgenic fish lines Plasmid construction was based on the Tol2/Gateway zebrafish kit (Kwan et al. 2007). The p5e-ubi entry clone was a generous gift from the laboratory of Dr. Leonard Zon (Boston Children’s Hospital). The p5e-sycp plasmid was cloned by amplifying the sycp promoter from genomic DNA before using Gateway BP Clonase II to insert the gel- selected product into pDONRP4-P1R (#219) using previously described primers (Gautier et al. 2013). The pMe-3xFLAG-eGFP-Rpl10a plasmid was cloned by stitching together PCR products containing 3xFLAG, eGFP, and Rpl10a coding sequence (amplified from 1 dpf cDNA) via Gibson Assembly (Gibson et al. 2009), then amplified using gateway primers before using Gateway BP Clonase II (ThermoFisher Scientific, 11789020) to clone the insert product. The pMe-3xFLAG-eGFP-Rpl10a-430-3’UTR plasmid was similarly cloned. The p3e-polyA entry clone (#302) and pDestTol2 (#393) were used from the Tol2Kit in conjunction with the above plasmids and LR Clonase II (ThermoFisher Scientific, 11791020) to generate ubi:eGFP-3xFLAG-Rpl10a:polyA and sycp:eGFP- 3xFLAG-Rpl10a-430-3’UTR:polyA and are referred to as Tg(Som-RiboFLAG) and Tg(Mat-RiboFLAG) respectively. To generate mosaic transgenic fish lines, a 1 nL mixture of plasmid DNA (40 ng/μL) and capped-mRNA encoding Tol2 Transposase (20 ng/μL) was loaded into needles pulled from glass capillary tubing (Warner, 64–0766) and injected into one-cell-stage embryos using a pico-liter injector (Warner Instruments, PLI90A). Mosaic founders were crossed with wild-type animals to generate a stable line. F2 and F3 embryos were used for images and experiments. eGFP fluorescence and brightfield images were acquired on a Leica M205 FCA fluorescence stereo microscope. Images were taken with the same settings for all embryos. 113 FACS isolation of primordial germ cells (PGCs) One-cell-stage zebrafish embryos were injected with 1-2 nL eGFP-nanos3’UTR mRNA using a pico-liter injector (Warner Instruments, PLI90A) and needles pulled from glass capillary tubing (Warner, 64–0766). At 24 hpf, embryos were sorted for eGFP expression, dechorionated using 1 mg/mL pronase (Pronase from Streptomyces griseus, Cat# 000000010165921001, Sigma-Aldrich), and washed with 1x PBS (Genesee Scientific, 25–508). 80 eGFP-positive embryos were incubated in a 5 mL solution of TrypLE Express (ThermoFisher Scientific, 12605036) and 0.003% Tricaine (ethyl-3-aminobenzoate- methanesulfonate) for 60 min at 31 °C in a Thermomixer (Eppendorf, Thermomixer R) set to 100 rpm, followed by trituration using a glass Pasteur pipette until tissues were digested into single cells. Cells were passed through a 40 μm strainer (Falcon, 352340), pelleted for 5 min at 500 xg at room temperature, and washed with 1x PBS. Cell pellets were resuspended in 1x PBS supplemented with 5% fetal bovine serum (FBS, Gemini Bio-products). Using a BD FACSAria cell sorter, approximately 1,200 eGFP-expressing cells (primordial germ cells) and 1,200 non-eGFP-expressing cells (somatic cells) were obtained from 80 embryos and sorted directly into TRIzol. Additions to polysome gradient protocols To obtain a better resolution of the low molecular weight fractions, 24 hour post- fertilization embryos were batch deyolked according to the previously published protocol (Link et al. 2006). While 200 to 250 embryos were used per sample for non-deyolked samples, 500 embryos were used per gradient for deyolked embryos. The deyolking buffer was supplemented with 1x protease inhibitors (EDTA-free, Roche) and 0.1 μg/mL cycloheximide. To avoid DNA contamination in the higher molecular weight factions, obtained larval lysates were treated with 1:125 Turbo DNase (QIAGEN, 79254) according to the manufactures protocol and incubated for 10 min at 28 °C. To isolate RNA after fractionation, 400 μL from each collected fraction were transferred into a Safe-lock Eppendorf tube, 1000 μL TRIzol (ThermoFisher Scientific, 15596-018) reagent were added and samples were vortexed for 10 s immediately. The following steps were performed according to the standard TRIzol based RNA isolation protocol. Western blotting of polysome fractions 400 μL of each fraction was first mixed with 700 μL water to dilute sucrose. Sodium deoxycholate and TCA were added to a final concentration of 0.08% and 20% respectively. Samples were vortexed and incubated overnight at -20 °C and centrifuged at 20,000 xg at 4 °C for 30 min. Protein pellets were washed twice with 2 volumes of cold (-20 °C) 100% acetone, then pelleted again. After acetone was removed, protein pellets were air dried, resuspended in 100 μL Laemmli buffer, sonicated for 5 min (30 s on, 30s off), and boiled for 5 min, before running a portion on a 4-20% Tris-glycine polyacrylamide gel (Invitrogen, XV04205). Transfer to a PVDF membrane previously activated with methanol, was run in cold Tris-glycine transfer buffer (1x Tris-glycine, 20% methanol) at 80 V for 80 min. After transfer, membranes were blocked in 5% milk in PBST (1x PBS with 0.1% Tween-20) for 1 h and blotted with primary antibodies (1: 2000 anti-FLAG, 114 Millipore Sigma, F3165, and 1:500 anti-Rpl7, ThermoFisher Scientific, A300-741A) overnight at 4 °C. Membranes were washed with PBST and blotted with secondary antibodies (1:10,000 anti-mouse, Invitrogen, 32230, or 1:10,000 anti-rabbit, Invitrogen, 32260) for 1 h at room temperature. All blots were sequentially probed SuperSignal West Femto (ThermoFisher Scientific, 34096) was used to develop the blot and a Bioanalytical Imaging System model c500 (Azure Biosciences) was used for imaging. See Supplementary Fig. S12 for uncropped original images. Isolation of zebrafish embryos for total RNA time course Embryos were snap-frozen in liquid nitrogen and stored at −80 °C. To maintain a uniform genetic background, all embryos were collected from the same clutch of a single fish stock (for the time course experiment). Detection of maternal and somatic rRNA variants by PCR Due to the high G/C content of rRNAs and therefore of the corresponding cDNA, standard PCR conditions were adapted to these more complex templates. RNA was isolated as previously described (Leesch et al. 2023). The PCR reaction differed from the manufacturer’s protocol in the addition of 1.3x Q5® Hot Start High-Fidelity (from a 2x MasterMix, New England Biolabs M0494) and additionally 1 M betaine (Sigma-Aldrich, B0300). The PCR conditions were modified according to the standard protocol by extending of the elongation time up to 90s in each PCR cycle and a final elongation step of 15 min. Primer sequences and annealing temperatures are listed in Supplementary Table 2.S4. Isolation of zebrafish ribosomes for CryoEM 5-day-old zebrafish larvae were obtained from TLAB wild-type zebrafish crosses according to the animal care guidelines and protocols described in (Leesch et al. 2023). To collect 5-day-old larvae, 0.01% (w/v) tricaine (25× stock solution in dH2O, buffered to pH 7-7.5 with 1 M Tris pH 9.0) was added to blue water consisting of fish water, 0.025% (v/v) Instant Ocean salts (Aquarium Systems, 218035), and 0.0001% (v/v) methylene blue (Sigma-Aldrich, M9140), pH 7. Embryos and larvae were lysed in freshly prepared lysis buffer containing 20 mM HEPES- KOH (pH7.4), 150 mM KCl, 10 mM MgCl2, 0.5 mM DTT, rRNasin (Promega), 1 X cOmplete-EDTA-free protease inhibitor (Roche) using a precooled 2 ml Dounce homogenizer. The lysis buffer was adapted from a previous publication about the isolation ribosomes from zebrafish larvae (Trinh et al. 2017). The protocol for isolation of crude ribosomes was adapted from previously published CryoEM protocols (Khatter et al. 2014, 2015). The following steps were identical to the isolation protocol for zebrafish ribosomes described (Leesch et al. 2023), and are summarized in Fig. 2.S6A and Table 2.S2. The established isolation protocol was designed to provide highly homogeneous samples for MS and CryoEM. While the use of translation inhibitors or starvation in this system was not possible to obtain more homogeneous samples and ultimately higher resolution. Therefore, the protocol included high salt washes and centrifugation steps that would exclude higher molecular weight polysomes as well as individual subunits. The purity and 115 quality of the samples was assessed by the band pattern on an SDS page and the RNA concentration in each fraction (Fig. 2.S6B and 2.S6C). After sucrose was removed from the fractions used for CryoEM, sample quality and ribosome concentration were assessed by negative staining electron microscopy (EM) (Fig. 2.S6D). Processing CryoEM data Processing of obtained CryoEM datasets was performed using the same program settings, protocols and processing sets as described (Leesch et al. 2023). In the following section, only the processing steps of the 5 dpf ribosome are discussed, as the other datasets are published in (Leesch et al. 2023). All details used in the generation of these different datasets are listed in Table 2.S2. Cryosparc version 2.16.0 was primarily used to process the somatic ribosome dataset (Punjani et al. 2017). Micrographs were motion-corrected and dose-weighted using Patch Motion Correction, and Patch CTF was used to determine contrast transfer function (CTF) parameters. Approximately 1000 particles were manually selected and 2D classified to create templates. These templates were then used for automated particle selection and were further manually inspected. The remaining particles were extracted and subjected to 2D classification. Classes with clear ribosomal densities were selected for heterogeneous ab initio model generation, while the remaining particles were refined in 3D to generate a consensus model. RNA extraction from TRIzol Homogenized lysate was mixed with 0.2 mL chloroform per mL of TRIzol, vortexed, and centrifuged for 15 minutes at 16,000 xg at 4 °C. The upper aqueous phase containing RNA was mixed with an equal volume of 100% ethanol, vortexed, processed with a Zymo column (RNA Clean and Concentrator, Zymo, R1014) including on-column DNase treatment, and eluted in 13 μL RNAse-free water. cDNA synthesis and RT-qPCR cDNA was synthesized using SuperScript IV VILO Master Mix (ThermoFisher Scientific, 11766050). For each sample, an 8 μL reaction composed of 6.4 μL RNA, 0.8 μL 10x ezDNase Buffer, and 0.8 μL ezDNAse enzyme was incubated at 37 °C for 5 min before centrifuging and placing on ice. Then, 3 μL of SuperScript IV VILO Master Mix or No RT Control and 4 μL of RNase-free water was added to each reaction, followed by incubation at 25 °C for 10 min, 60 °C for 10 min, and 85 °C for 5 min. The synthesized cDNA was diluted 5-1000 fold with RNase-free water, depending on the input material, and used in a 10 μL RT-qPCR reaction composed of 5 μL 2x KAPA SYBR FAST qPCR Master Mix (Roche, KK4600), 2.5 μL cDNA, and 0.25 μM of each primer. Reactions were arrayed into a 384-well reaction plate or a 96-well reaction and measured with a Light Cycler 480 II Real-time PCR Machine: 95 °C for 30 s, 35 cycles of 95 °C for 5 s, 60 °C for 15 s and 68 °C for 15 s. Three biological replicates were run in technical triplicate for each measurement. No reverse-transcriptase samples were included as negative controls. Relative fold differences of specific targets were calculated using the 116 2ΔCt method. Statistical analyses were performed using Student’s unpaired t-test where p < 0.05 was considered significant. In vitro transcription of mRNA and biotin labeling To generate the RNA for microinjections, 2 μg of a construct encoding eGFP-nanos3’UTR was digested using NotI-HF (New England BioLabs, R3189S) for 1 h. The linearized product was purified according to manufacturer specifications via gel selection using NucleoSpin (Macherey-Nagel, 740588.50). 500 ng of DNA was used in an overnight IVT reaction using mMESSAGE mMACHINE T7 ULTRA Transcription Kit (Invitrogen, AM1345). The product was treated with TURBO DNase for 30 min. RNA was poly-A- tailed using the provided tailing reaction and samples were run on agarose gels to confirm sizes. RNA was purified using Zymo RNA Clean and Concentrator columns (Zymo Research, R1016) and concentration was measured using a Nanodrop ND-1000 Spectrophotometer (ThermoFisher Scientific). End labeling of RNA was achieved using previously described methods (Wang et al. 2007). Briefly, 20 pmol of in vitro transcribed mRNA (approximately 10 ug) was subjected to labeling with 1 nmol of pCp-Biotin (Jena Bioscience, NU-1706-BIO) in a 20 μL reaction comprising 5 μL DMSO, 2 μL 10 mM ATP, 1 μL T4 RNA Ligase 1 (NEB, M0437M), 1 μL SuperAseIn (Invitrogen AM2694), and 2 μL T4 RNA Ligase Buffer at 25 °C for 2 h. A no- labeling reaction was similarly incubated with the reaction components lacking pCp- Biotin. All mRNAs were purified using Zymo RNA columns and adjusted to 200 ng/μL. Isolation of zebrafish ribosome subunits by immunoprecipitation Embryos were washed twice with 500 μL of ice-cold Subunit Buffer (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM EDTA, and 1 mM DTT. As much buffer as possible was removed before samples were either immediately processed or flash frozen and stored at -80 °C. 400 μL of ice-cold Subunit Lysis Buffer (Subunit Buffer with 1% Triton X-100, 1x cOmplete-EDTA-free protease inhibitor (Sigma, 11873580001), and 20 mM EDTA) was added to either fresh or frozen tissue. Keeping tubes on ice, a pre-cooled 1 mL syringe attached to a 26-gauge needle was used to shear samples by slowly triturating material for 20-30 strokes until no visible tissue remained. Lysates were incubated on ice for 5 min, then cleared by centrifugation at 10,000 xg for 10 min at 4 °C. Cleared S10 fractions were adjusted to 500 μL with Subunit Lysis Buffer and overlaid onto 2 mL of a 1 M sucrose cushion prepared in Subunit Buffer in a 15x51 mm 3 mL thickwall polycarbonate ultracentrifuge tube (Beckman Coulter, 349622) on ice. Ribosomal subunits were pelleted using a TLA100.3 rotor (Beckman Coulter, 349490) at ~300,000 xg at 4 °C for 2 h. Supernatants were carefully poured off and pellets were resuspended by horizontal rotation on ice for at least 30 min in 235 μL of Subunit Buffer. Resuspended subunits were incubated with 2 μL of anti-FLAG antibody (Millipore Sigma, F3165-.2MG) for 1 h at 4 °C. Then, 14 μL of equilibrated Dynabeads Protein G (ThermoFisher Scientific, 10003D) were added and incubated at 4 °C for at least 4 h. Beads were subsequently collected using a magnetic rack, washed with Subunit Buffer three times, and resuspended in TRIzol-LS reagent (Invitrogen, 10296028). 117 Isolation of zebrafish monosomes by immunoprecipitation Embryos were washed twice with 500 μL of ice-cold Ribosome Buffer [20 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT]. As much buffer as possible was removed before samples were either immediately processed or flash frozen and stored at -80 °C. 200 μL of ice-cold Ribosome Lysis Buffer [Ribosome Buffer containing 1% Triton X-100, 1x cOmplete-EDTA-free protease inhibitor (Sigma, 11873580001), and 100 μg/mL cycloheximide] was added to either fresh or frozen tissue. Keeping tubes on ice, a pre- cooled 1 mL syringe attached to a 26-gauge needle was used to shear samples by slowly triturating material for 20-30 strokes until no visible tissue remained. Lysates were incubated on ice for 5 min, cleared by centrifugation at 10,000 xg for 10 min at 4 °C, then treated with 1 unit per embryo of RNAse T1 (ThermoFisher, EN0541). After 90 min of rotating at room temperature, 2 units per embryo of SuperAseIn (Invitrogen, AM2694) were added to stop the reaction. Cleared and digested S10 fractions were adjusted to 250 μL with Ribosome Lysis Buffer and overlaid onto a continuous 10-50% (w/v) sucrose gradient prepared in Ribosome Buffer containing 100 μg/mL cycloheximide. Gradients were generated using a BioComp Gradient Master device and 14x89 mm ultracentrifuge tubes (Seton Scientific, 7030). Gradients were centrifuged in a SW41 Ti rotor (Beckman Coulter, 331362) at 35,000 rpm (rmax of 210,000 xg) for 150 min at 4 °C followed by analysis and fractionation using a BioComp Gradient Station coupled to a Model Triax™ Flow Cell detector (FC-2). Approximately 1500 μL from the monosome fraction were manually collected. The resulting monosomes were incubated with 2 μL of anti-FLAG antibody (Millipore Sigma, F3165-.2MG) for 1 h at 4 °C. Then, 14 μL of equilibrated Dynabeads Protein G (ThermoFisher Scientific, 10003D) were added and incubated at 4 °C for at least 4 h. Beads were subsequently collected using a magnetic rack, washed three times with Ribosome Buffer containing 100 μg/mL cycloheximide, and resuspended in TRIzol-LS reagent (Invitrogen, 10296028). RIP of biotinylated mRNAs One-cell-stage zebrafish embryos were injected with 1-2 nL of either biotinylated or non- biotinylated eGFP-nanos3’UTR mRNA. At 24 hpf, embryos were dechorionated using pronase (Pronase from Streptomyces griseus, Cat# 10165921001, Sigma-Aldrich) and checked for eGFP expression. Approximately 400 eGFP-positive embryos were washed twice with 500 μL of ice-cold Ribosome Buffer [20 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT]. As much buffer as possible was removed before samples were immediately processed. 400 μL of ice-cold Ribosome Lysis Buffer [Ribosome Buffer containing 1% Triton X-100, 1x cOmplete-EDTA-free protease inhibitor (Sigma, 11873580001), and 100 μg/mL cycloheximide] was added to fresh tissue. Keeping tubes on ice, a pre-cooled 1 mL syringe attached to a 26-gauge needle was used to shear samples by slowly triturating 118 material for 20-30 strokes until no visible tissue remained. Lysates were incubated on ice for 5 min, then cleared by centrifugation at 10,000 xg for 10 min at 4 °C. Cleared S10 fractions were adjusted to 500 μL with Ribosome Lysis Buffer and overlaid onto 2 mL of a 1 M sucrose cushion prepared in Ribosome Buffer containing 100 μg/mL cycloheximide in a 15x51 mm 3 mL thickwall polycarbonate ultracentrifuge tube (Beckman Coulter, 349622) on ice. Ribosomes were pelleted using a TLA100.3 rotor (Beckman Coulter, 349490) at 80,000 rpm (rmax of 346,000 xg) for 2 h at 4 °C. Supernatants were carefully poured off and pellets were resuspended by horizontal rotation on ice for at least 30 min in 235 μL of Ribosome Buffer with 100 μg/mL cycloheximide and 5 μL SuperAseIn (Invitrogen AM2694). Streptavidin magnetic beads (Invitrogen, 65001) were washed twice with Ribosome Buffer, then added to resuspended ribosomes, and incubated with rotation overnight at 4 °C. Beads were magnetically collected, washed three times with Ribosome Buffer containing 100 μg/mL cycloheximide and resuspended in TRIzol LS reagent (Invitrogen, 10296028). 119 References Anger AM, Armache J-P, Berninghausen O, Habeck M, Subklewe M, Wilson DN, Beckmann R. 2013. Structures of the human and Drosophila 80S ribosome. Nature 497: 80–85. Barna M, Karbstein K, Tollervey D, Ruggero D, Brar G, Greer EL, Dinman JD. 2022. The promises and pitfalls of specialized ribosomes. Mol Cell 82: 2179–2184. Bazzini AA, Del Viso F, Moreno-Mateos MA, Johnstone TG, Vejnar CE, Qin Y, Yao J, Khokha MK, Giraldez AJ. 2016. Codon identity regulates mRNA stability and translation efficiency during the maternal-to-zygotic transition. EMBO J 35: 2087– 2103. Ben-Shem A, Garreau de Loubresse N, Melnikov S, Jenner L, Yusupova G, Yusupov M. 2011. The structure of the eukaryotic ribosome at 3.0 Å resolution. Science 334: 1524–1529. Berghmans S, Murphey RD, Wienholds E, Neuberg D, Kutok JL, Fletcher CDM, Morris JP, Liu TX, Schulte-Merker S, Kanki JP, et al. 2005. tp53 mutant zebrafish develop malignant peripheral nerve sheath tumors. Proc Natl Acad Sci U S A 102: 407– 412. Bielczyk-Maczyńska E, Lam Hung L, Ferreira L, Fleischmann T, Weis F, Fernández- Pevida A, Harvey SA, Wali N, Warren AJ, Barroso I, et al. 2015. The Ribosome Biogenesis Protein Nol9 Is Essential for Definitive Hematopoiesis and Pancreas Morphogenesis in Zebrafish. PLoS Genet 11: e1005677. Breit TM, Rauwerda H, Pagano JFB, Ensink WA, Nehrdich U, Spaink HP, Dekker RJ. 2020. Immunoglobulin switch-like recombination regions implicated in the formation of extrachromosomal circular 45S rDNA involved in the maternal-specific translation system of zebrafish. Developmental Biology http://biorxiv.org/lookup/doi/10.1101/2020.01.31.928739 (Accessed July 31, 2023). Cabrera-Quio LE, Schleiffer A, Mechtler K, Pauli A. 2021. Zebrafish Ski7 tunes RNA levels during the oocyte-to-embryo transition. PLoS Genet 17: e1009390. Cenik ES, Meng X, Tang NH, Hall RN, Arribere JA, Cenik C, Jin Y, Fire A. 2019. Maternal Ribosomes Are Sufficient for Tissue Diversification during Embryonic Development in C. elegans. Dev Cell 48: 811-826.e6. Dimarco E, Cascone E, Bellavia D, Caradonna F. 2012. Functional variants of 5S rRNA in the ribosomes of common sea urchin Paracentrotus lividus. Gene 508: 21–25. Dranow DB, Hu K, Bird AM, Lawry ST, Adams MT, Sanchez A, Amatruda JF, Draper BW. 2016. Bmp15 Is an Oocyte-Produced Signal Required for Maintenance of the Adult Female Sexual Phenotype in Zebrafish ed. M.C. Mullins. PLOS Genet 12: e1006323. Dranow DB, Tucker RP, Draper BW. 2013. Germ cells are required to maintain a stable sexual phenotype in adult zebrafish. Dev Biol 376: 43–50. Driever W, Rangini Z. 1993. Characterization of a cell line derived from zebrafish (Brachydanio rerio) embryos. In Vitro Cell Dev Biol Anim 29A: 749–754. Eichhorn SW, Subtelny AO, Kronja I, Kwasnieski JC, Orr-Weaver TL, Bartel DP. 2016. mRNA poly(A)-tail changes specified by deadenylation broadly reshape translation in Drosophila oocytes and early embryos. eLife 5: e16955. 120 Ferretti MB, Ghalei H, Ward EA, Potts EL, Karbstein K. 2017. Rps26 directs mRNA- specific translation by recognition of Kozak sequence elements. Nat Struct Mol Biol 24: 700–707. Ferretti MB, Karbstein K. 2019. Does functional specialization of ribosomes really exist? RNA N Y N 25: 521–538. Gautier A, Goupil A-S, Le Gac F, Lareyre J-J. 2013. A Promoter Fragment of the sycp1 Gene Is Sufficient to Drive Transgene Expression in Male and Female Meiotic Germ Cells in Zebrafish1. Biol Reprod 89: 89, 1–14. Gibson DG, Young L, Chuang R-Y, Venter JC, Hutchison CA, Smith HO. 2009. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6: 343– 345. Giraldez AJ, Mishima Y, Rihel J, Grocock RJ, Van Dongen S, Inoue K, Enright AJ, Schier AF. 2006. Zebrafish MiR-430 promotes deadenylation and clearance of maternal mRNAs. Science 312: 75–79. Gunderson JH, Sogin ML, Wollett G, Hollingdale M, de la Cruz VF, Waters AP, McCutchan TF. 1987. Structurally distinct, stage-specific ribosomes occur in Plasmodium. Science 238: 933–937. Heilmann S, Ratnakumar K, Langdon E, Kansler E, Kim I, Campbell NR, Perry E, McMahon A, Kaufman C, van Rooijen E, et al. 2015. A quantitative system for studying metastasis using transparent zebrafish. Cancer Res 75: 4272–4282. Hensey C, Gautier J. 1997. A developmental timer that regulates apoptosis at the onset of gastrulation. Mech Dev 69: 183–195. Heyn P, Salmonowicz H, Rodenfels J, Neugebauer KM. 2017. Activation of transcription enforces the formation of distinct nuclear bodies in zebrafish embryos. RNA Biol 14: 752–760. Hopes T, Norris K, Agapiou M, McCarthy CGP, Lewis PA, O’Connell MJ, Fontana J, Aspden JL. 2022. Ribosome heterogeneity in Drosophila melanogaster gonads through paralog-switching. Nucleic Acids Res 50: 2240–2257. Johnson AD, Richardson E, Bachvarova RF, Crother BI. 2011. Evolution of the germ line- soma relationship in vertebrate embryos. Reprod Camb Engl 141: 291–300. Khatter H, Myasnikov AG, Mastio L, Billas IML, Birck C, Stella S, Klaholz BP. 2014. Purification, characterization and crystallization of the human 80S ribosome. Nucleic Acids Res 42: e49. Khatter H, Myasnikov AG, Natchiar SK, Klaholz BP. 2015. Structure of the human 80S ribosome. Nature 520: 640–645. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. 1995. Stages of embryonic development of the zebrafish. Dev Dyn Off Publ Am Assoc Anat 203: 253–310. Kloc M, Etkin LD. 2005. RNA localization mechanisms in oocytes. J Cell Sci 118: 269– 282. Komili S, Farny NG, Roth FP, Silver PA. 2007. Functional specificity among ribosomal proteins regulates gene expression. Cell 131: 557–571. Kwan KM, Fujimoto E, Grabher C, Mangum BD, Hardy ME, Campbell DS, Parant JM, Yost HJ, Kanki JP, Chien C-B. 2007. The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn Off Publ Am Assoc Anat 236: 3088–3099. 121 Leesch F, Lorenzo-Orts L, Pribitzer C, Grishkovskaya I, Roehsner J, Chugunova A, Matzinger M, Roitinger E, Belačić K, Kandolf S, et al. 2023. A molecular network of conserved factors keeps ribosomes dormant in the egg. Nature 613: 712–720. Li J, Gutell RR, Damberger SH, Wirtz RA, Kissinger JC, Rogers MJ, Sattabongkot J, McCutchan TF. 1997. Regulation and trafficking of three distinct 18 S ribosomal RNAs during development of the malaria parasite. J Mol Biol 269: 203–213. Lim J, Lee M, Son A, Chang H, Kim VN. 2016. mTAIL-seq reveals dynamic poly(A) tail regulation in oocyte-to-embryo development. Genes Dev 30: 1671–1682. Link V, Shevchenko A, Heisenberg C-P. 2006. Proteomics of early zebrafish embryos. BMC Dev Biol 6: 1. Liu J, Lichtenberg T, Hoadley KA, Poisson LM, Lazar AJ, Cherniack AD, Kovatich AJ, Benz CC, Levine DA, Lee AV, et al. 2018a. An Integrated TCGA Pan-Cancer Clinical Data Resource to Drive High-Quality Survival Outcome Analytics. Cell 173: 400-416.e11. Liu Y, Zhao H, Shao F, Zhang Y, Nie H, Zhang J, Li C, Hou Z, Chen Z-J, Wang J, et al. 2023. Remodeling of maternal mRNA through poly(A) tail orchestrates human oocyte-to-embryo transition. Nat Struct Mol Biol 30: 200–215. Liu Y, Zou W, Yang P, Wang L, Ma Y, Zhang H, Wang X. 2018b. Autophagy-dependent ribosomal RNA degradation is essential for maintaining nucleotide homeostasis during C. elegans development. eLife 7: e36588. Locati MD, Pagano JFB, Ensink WA, van Olst M, van Leeuwen S, Nehrdich U, Zhu K, Spaink HP, Girard G, Rauwerda H, et al. 2017a. Linking maternal and somatic 5S rRNA types with different sequence-specific non-LTR retrotransposons. RNA N Y N 23: 446–456. Locati MD, Pagano JFB, Girard G, Ensink WA, van Olst M, van Leeuwen S, Nehrdich U, Spaink HP, Rauwerda H, Jonker MJ, et al. 2017b. Expression of distinct maternal and somatic 5.8S, 18S, and 28S rRNA types during zebrafish development. RNA N Y N 23: 1188–1199. Lorenzo-Orts L, Strobl M, Steinmetz B, Leesch F, Pribitzer C, Schutzbier M, Dürnberger G, Pauli A. 2023. eIF4E1b is a non-canonical eIF4E required for maternal mRNA dormancy. Developmental Biology http://biorxiv.org/lookup/doi/10.1101/2023.06.10.544440 (Accessed July 31, 2023). Maegawa S, Yasuda K, Inoue K. 1999. Maternal mRNA localization of zebrafish DAZ-like gene. Mech Dev 81: 223–226. Mageeney CM, Ware VC. 2019. Specialized eRpL22 paralogue-specific ribosomes regulate specific mRNA translation in spermatogenesis in Drosophila melanogaster. Mol Biol Cell 30: 2240–2253. Medioni C, Mowry K, Besse F. 2012. Principles and roles of mRNA localization in animal development. Dev Camb Engl 139: 3263–3276. Melnikov S, Ben-Shem A, Garreau de Loubresse N, Jenner L, Yusupova G, Yusupov M. 2012. One core, two shells: bacterial and eukaryotic ribosomes. Nat Struct Mol Biol 19: 560–567. Noack Watt KE, Achilleos A, Neben CL, Merrill AE, Trainor PA. 2016. The Roles of RNA Polymerase I and III Subunits Polr1c and Polr1d in Craniofacial Development and in Zebrafish Models of Treacher Collins Syndrome. PLoS Genet 12: e1006187. 122 Noda T, Blaha A, Fujihara Y, Gert KR, Emori C, Deneke VE, Oura S, Panser K, Lu Y, Berent S, et al. 2022. Sperm membrane proteins DCST1 and DCST2 are required for sperm-egg interaction in mice and fish. Commun Biol 5: 332. O’Leary MN, Schreiber KH, Zhang Y, Duc A-CE, Rao S, Hale JS, Academia EC, Shah SR, Morton JF, Holstein CA, et al. 2013. The ribosomal protein Rpl22 controls ribosome composition by directly repressing expression of its own paralog, Rpl22l1. PLoS Genet 9: e1003708. Ortega-Recalde O, Day RC, Gemmell NJ, Hore TA. 2019. Zebrafish preserve global germline DNA methylation while sex-linked rDNA is amplified and demethylated during feminisation. Nat Commun 10: 3053. Pasolini P, Costagliola D, Rocco L, Tinti F. 2006. Molecular organization of 5S rDNAs in Rajidae (Chondrichthyes): Structural features and evolution of piscine 5S rRNA genes and nontranscribed intergenic spacers. J Mol Evol 62: 564–574. Patton EE, Widlund HR, Kutok JL, Kopani KR, Amatruda JF, Murphey RD, Berghmans S, Mayhall EA, Traver D, Fletcher CDM, et al. 2005. BRAF mutations are sufficient to promote nevi formation and cooperate with p53 in the genesis of melanoma. Curr Biol CB 15: 249–254. Pauli A, Norris ML, Valen E, Chew G-L, Gagnon JA, Zimmerman S, Mitchell A, Ma J, Dubrulle J, Reyon D, et al. 2014. Toddler: an embryonic signal that promotes cell movement via Apelin receptors. Science 343: 1248636. Paw BH, Zon LI. 1999. Primary fibroblast cell culture. Methods Cell Biol 59: 39–43. Perez DE, Henle AM, Amsterdam A, Hagen HR, Lees JA. 2018. Uveal melanoma driver mutations in GNAQ/11 yield numerous changes in melanocyte biology. Pigment Cell Melanoma Res 31: 604–613. Peterson RC, Doering JL, Brown DD. 1980. Characterization of two xenopus somatic 5S DNAs and one minor oocyte-specific 5S DNA. Cell 20: 131–141. Pfefferli C, Jaźwińska A. 2015. The art of fin regeneration in zebrafish. Regeneration (Oxf) 2: 72–83. Punjani A, Rubinstein JL, Fleet DJ, Brubaker MA. 2017. cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat Methods 14: 290–296. Redl S, de Jesus Domingues AM, Caspani E, Möckel S, Salvenmoser W, Mendez-Lago M, Ketting RF. 2021. Extensive nuclear gyration and pervasive non-genic transcription during primordial germ cell development in zebrafish. Dev Camb Engl 148: dev193060. Rothschild D, Susanto TT, Spence JP, Genuth NR, Sinnott-Armstrong N, Pritchard JK, Barna M. 2023. A comprehensive rRNA variation atlas in health and disease. 2023.01.30.526360. https://www.biorxiv.org/content/10.1101/2023.01.30.526360v1 (Accessed August 29, 2023). Schmitt AO, Herzel H. 1997. Estimating the entropy of DNA sequences. J Theor Biol 188: 369–377. Schramm RD, Bavister BD. 1999. Onset of nucleolar and extranucleolar transcription and expression of fibrillarin in macaque embryos developing in vitro. Biol Reprod 60: 721–728. 123 Shi X, Khade PK, Sanbonmatsu KY, Joseph S. 2012. Functional role of the sarcin-ricin loop of the 23S rRNA in the elongation cycle of protein synthesis. J Mol Biol 419: 125–138. Shi Z, Fujii K, Kovary KM, Genuth NR, Röst HL, Teruel MN, Barna M. 2017. Heterogeneous Ribosomes Preferentially Translate Distinct Subpools of mRNAs Genome-wide. Mol Cell 67: 71-83.e7. Simsek D, Tiu GC, Flynn RA, Byeon GW, Leppek K, Xu AF, Chang HY, Barna M. 2017. The Mammalian Ribo-interactome Reveals Ribosome Functional Diversity and Heterogeneity. Cell 169: 1051-1065.e18. Subtelny AO, Eichhorn SW, Chen GR, Sive H, Bartel DP. 2014. Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature 508: 66–71. Szewczak AA, Moore PB, Chang YL, Wool IG. 1993. The conformation of the sarcin/ricin loop from 28S ribosomal RNA. Proc Natl Acad Sci U S A 90: 9581–9585. Takahashi H. 1977. Juvenile Hermaphroditism in the Zebrafish, Brachydanio rerio. 北海 道大學水産學部研究彙報 28: 57–65. Tamm T, Kisly I, Remme J. 2019. Functional Interactions of Ribosomal Intersubunit Bridges in Saccharomyces cerevisiae. Genetics 213: 1329–1339. Tao B, Lo LJ, Peng J, He J. 2020. rDNA subtypes and their transcriptional expression in zebrafish at different developmental stages. Biochem Biophys Res Commun 529: 819–825. Trinh LA, Chong-Morrison V, Gavriouchkina D, Hochgreb-Hägele T, Senanayake U, Fraser SE, Sauka-Spengler T. 2017. Biotagging of Specific Cell Populations in Zebrafish Reveals Gene Regulatory Logic Encoded in the Nuclear Transcriptome. Cell Rep 19: 425–440. Uchiumi T, Kominami R. 1994. A functional site of the GTPase-associated center within 28S ribosomal RNA probed with an anti-RNA autoantibody. EMBO J 13: 3389– 3394. Varshney GK, Pei W, LaFave MC, Idol J, Xu L, Gallardo V, Carrington B, Bishop K, Jones M, Li M, et al. 2015. High-throughput gene targeting and phenotyping in zebrafish using CRISPR/Cas9. Genome Res 25: 1030–1042. Vierna J, Wehner S, Höner zu Siederdissen C, Martínez-Lage A, Marz M. 2013. Systematic analysis and evolution of 5S ribosomal DNA in metazoans. Heredity 111: 410–421. Wang H, Ach RA, Curry B. 2007. Direct and sensitive miRNA profiling from low-input total RNA. RNA 13: 151–159. Waters AP, van Spaendonk RM, Ramesar J, Vervenne RA, Dirks RW, Thompson J, Janse CJ. 1997. Species-specific regulation and switching of transcription between stage-specific ribosomal RNA genes in Plasmodium berghei. J Biol Chem 272: 3583–3589. Xue S, Tian S, Fujii K, Kladwang W, Das R, Barna M. 2015. RNA regulons in Hox 5’ UTRs confer ribosome specificity to gene regulation. Nature 517: 33–38. Yu J, Lan X, Chen X, Yu C, Xu Y, Liu Y, Xu L, Fan H-Y, Tong C. 2016. Protein synthesis and degradation are essential to regulate germline stem cell homeostasis in Drosophila testes. Dev Camb Engl 143: 2930–2945. 124 Supplementary Figures Supplementary Figure 2.S1: Detection of maternal and somatic rRNAs in embryonic development A) Schematic representation of the rRNA expansion segments (ES) depicted in Fig. 2.1A which are targeted for a PCR-based fragment length polymorphism (FLP) assay. Either ES3S (helical regions a and b) in the 18S rRNA or ES31L (helical regions a and b) in the 28S rRNA are PCR amplified using a single primer pair (indicated by primer icons) detecting both M-type and S-type rRNA variants in the same reaction. Resulting PCR products differ in size due to the different lengths of M-type and S-type ES regions. Supplementary Table 2.S4 contains the relevant PCR conditions and primer sequences. B) Gel electrophoretic analysis of PCR-based detection of maternal and somatic rRNA variants at the indicated developmental times. For each developmental time assayed, three individual embryos, each from an independent natural cross comprising one wildtype female and one wildtype male, were used for RNA extraction and cDNA synthesis. C) Averaged (n=3) line profiles of densitometry scans (ImageJ) from each lane of the gel shown in (B). The regions representing signal from M-type and S-type 18S rRNA PCR FLP are indicated and used for relative quantification in Fig. 2.1B. H, helix. ES, expansion segment. bp, base pair. hpf, hours post-fertilization. NRT, no RT control. NTC, no template control. M, 100 bp marker. 125 Supplementary Figure 2.S2: Analysis of rRNA variant expression in different tissues Gel electrophoretic analysis of PCR-based detection of maternal and somatic A) 18S rRNA and B) 28S rRNA variants in different adult zebrafish tissues dissected from males (n=3, each). Positive control reactions using 2 dpf and egg cDNAs are included for reference. C) Gel electrophoretic analysis of PCR- based detected of maternal and somatic 18S rRNA in different adult zebrafish tissues dissected from females and males (n=1, each). Positive control reactions using AB9 cell and 2-cell stage cDNAs are included for reference. See Supplementary Fig. 2.S1 and Supplementary Table 2.S4 for indicated lengths of PCR products from maternal and somatic rRNAs. H, helix. dpf, days post-fertilization. bp, base pair. M, 100 bp marker. 126 Supplementary Figure 2.S3: Detecting rRNA variants in cell lines, tumors, and regenerating fins A) Gel electrophoretic analysis of PCR-based fragment length polymorphism (FLP) 18S rRNA variant detection from the indicated cell lines. Diagnostic PCR lengths for maternal and somatic 18S rRNAs are indicated. Positive control reactions using 3 hpf, 24 hpf, and adult fin clip cDNAs are included to compare maternal-only, maternal and somatic, or somatic-only band patterns, respectively. B) Gel electrophoretic analysis of 18S rRNA variant detection from the indicated excised tumors. See Materials and Methods for a complete description of each cell line and tumor. hpf, hours post-fertilization. NTC, no template control. AB9 (fibroblast cells), ZMEL (melanoma cells), ZF4 (fibroblast cells), BRAF (melanoma tumor), GNAQ (melanoma tumor), MPNST (malignant peripheral nerve sheath tumor). C) Gel electrophoretic analysis of 18S rRNA variants from regenerating fin blastema excised at the indicated times post initial amputation. 127 Supplementary Figure 2.S4: Transcription of rRNA variants in zebrafish and primordial germ cells (PGCs) from 1 through 10 days post fertilization Expression of maternal (yellow) and somatic (blue) ITS1 sequences, indicative of de novo rRNA transcription, in A) total embryos/larvae and B) FACS-sorted PGCs. Public dataset PRJNA597223 (Redl et al. 2021) was obtained from the Sequence Read Archive at NCBI and analyzed as described. 128 Supplementary Figure 2.S5: Expression of alternative ribosomal core proteins in zebrafish oogenesis and adult tissue Heatmap of RNA expression of paralogs and alternative isoforms encoding ribosomal protein variants assayed A) over progressive stages of oogenesis and B) across adult tissues and organs. The relative presence of maternal (yellow) or somatic (blue) rRNA variants at these developmental stages is indicated in the scheme below the heat map. 129 Supplementary Figure 2.S6: Overview of ribosome isolation from zebrafish embryos and larvae A) Scheme summarizing the main steps of the isolation protocol. B) RNA concentration measurement from an example ribosome isolation (6 hpf) after gradient fractionation. C) SDS-PAGE of the gradient fractions stained with InstantBlue. The fraction indicated in yellow in (B) (fraction #14) was used for mass spectrometry (MS) and Cryo-EM analyses. D) Representative negative staining of fraction #14 to determine ribosome concentration and amount to use for the preparation of Cryo-EM grids. 130 Supplementary Figure 2.S7: Processing pipeline for the 5 dpf zebrafish ribosome Schematic overview of the methodologies used for capturing, particle picking, 2D classification, and 3D refinement of the structure. All steps were done in Cryosparc v3.2.0. Maps are shown in gray, masks in blue. The orientation distribution plot for all particles contributing to Map1 and the Gold-Standard Fourier Shell Correlation (GSFSC) of the respective map is shown on the bottom-right. Local resolution maps were calculated for Map1, Map2 and Map3. Using 9,456 micrographs as input, 569,541 particles were obtained to generate the final 2.8Å resolution map. dpf, days post-fertilization 131 Supplementary Figure 2.S8: Generation of Tg(Mat-RiboFLAG) and Tg(Som-RiboFLAG) lines A) Cartoon depicting the generation of two transgenic lines containing either FLAG-tagged maternal or FLAG-tagged somatic ribosomes (see Materials and Methods for details). Mosaic F0 were raised to adulthood and transmission of either transgene via the germline was screened for in B) 48 hours post- fertilization (hpf) embryos using a green heart marker (cmlc2 promoter driving eGFP) also encoded on the plasmid (Kwan et al. 2007) as indicated by the white arrow. As expected, only half of the progeny derived from a Tg(Som-RiboFLAG) male and a wildtype female express the transgene. Scale bars indicate 500 μm. hpf, hours post-fertilization. eGFP, enhanced green fluorescent protein. Tg, transgene. 132 Supplementary Figure 2.S9: Experimental strategies with transgenic lines Schematic illustration of the methodologies used with Tg(Mat-RiboFLAG) and Tg(Som-RiboFLAG). Transgenic 24 hpf larvae express either maternal (yellow) or somatic (blue) 3xFLAG-Rpl10a-tagged ribosomes (see Supplementary Fig. 2.S8, 2.S10, and 2.S11). All experiments are conducted with F2 and F3 adults. Biological replicates are derived from independent crosses. CHX treatment maintains the polysome architecture. EDTA treatment disassembles monosomes into 40S and 60S subunits. CHX with RNAse treatment generates 80S monosomes with ribosome-protected fragments of mRNA. hpf, hours post-fertilization. CHX, cycloheximide. EDTA, ethylenediaminetetraacetic acid. 133 Supplementary Figure 2.S10: Transgenic expression 3xFLAG-eGFP-Rpl10a Scheme summarizing the generation of embryos with tagged ribosomes. Brightfield and fluorescent images of developing zebrafish embryos from either A) maternally-provided Tg(Mat-RiboFLAG), B) paternally- provided Tg(Som-RiboFLAG), or C) paternally-provided Tg(Mat-RiboFLAG) parents. Embryos depicted in (A) contain maternally deposited 3xFLAG-eGFP-Rpl10a at 0 hpf which dilutes over time while embryos depicted in (B) begin expression of 3xFLAG-eGFP-Rpl10a which accumulates after 6 hpf. See Supplementary Fig. 2.S8 for expression at 48 hpf. As expected, progeny derived from paternally-provided Tg(Mat-RiboFLAG) show no transgene expression. Note 22 hpf embryos do not yet express the included green heart marker (cmlc2 promoter driving eGFP). Scale bars indicate 500 μm. hpf, hours post-fertilization. 134 Supplementary Figure 2.S11: Incorporation of Tg(3xFLAG-Rpl10a) into translating ribosomes A) Representative polysome profiles with continuous A260 reading from embryos at 24 hours post- fertilization (hpf). Position in the gradient and collected fraction numbers are indicated. B) Western blot of polysome gradient fractions containing lysate from 24 hpf embryos with maternally-provided Tg(Mat- RiboFLAG) detecting expression of transgenic FLAG-Rpl10a in the fraction of translating ribosomes. Negative control Western blot of polysome gradient fractions containing lysate from 24 hpf embryos with paternally-provided Tg(Mat-RiboFLAG) lacking expression of transgenic FLAG-Rpl10a. hpf, hours post- fertilization. kDa, kilodalton. 135 Supplementary Figure 2.S12: Polysome Western blots Uncropped versions of blots shown in Supplementary Fig. 2.S11B. Note the anti-Rpl7 detection of non- specific signal near 50 kDa is distinct from the 56 kDa signal generated by anti-FLAG detection of FLAG- eGFP-Rpl10a. kDa (kilodaltons). 136 Supplemental Tables Supplementary Table 2.S1: Proteins differentially associated with maternal and somatic zebrafish ribosomes. Proteins significantly enriched or depleted in ribosomes isolated from 6 hpf embryos versus 5 dpf larvae (permutation-based FDR <0.05). Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] si:dkey-7j14.6 7.43E+04 5.32E+04 4.29E+04 2.37E+08 1.06E+08 3.63E+08 -11.88 1.46E-06 rnasel3 7.43E+04 5.32E+04 4.29E+04 3.66E+08 5.93E+07 1.46E+08 -11.37 7.21E-06 retsatl 7.43E+04 5.32E+04 4.29E+04 4.22E+07 9.07E+07 4.58E+07 -9.98 1.45E-06 gyg1a 7.43E+04 3.96E+05 4.29E+04 2.22E+08 2.06E+08 1.13E+06 -8.43 1.08E-02 slc43a1a 7.43E+04 5.32E+04 1.02E+05 2.07E+07 1.16E+07 2.69E+07 -7.98 6.32E-06 vldlr 7.43E+04 5.32E+04 4.29E+04 1.21E+07 3.78E+06 1.51E+07 -7.32 3.96E-05 si:ch73-189n23.1 7.43E+04 9.52E+06 2.94E+07 1.86E+08 1.41E+09 2.95E+08 -7.28 2.25E-02 rnasel2 1.85E+06 1.51E+06 6.88E+06 3.80E+08 2.40E+08 1.45E+08 -6.46 1.67E-04 rps17 3.26E+06 6.99E+07 9.77E+07 2.20E+09 2.49E+09 2.19E+09 -6.34 3.69E-03 lamp2 7.43E+04 5.32E+04 4.29E+04 1.62E+07 3.77E+06 1.13E+06 -6.21 9.72E-04 lamp1b 7.43E+04 5.32E+04 4.29E+04 8.55E+06 5.57E+05 1.12E+07 -6.09 2.79E-03 neb 4.81E+05 1.06E+04 3.15E+04 5.07E+06 6.98E+06 1.13E+06 -5.97 9.35E-03 ELAPOR2 7.43E+04 5.32E+04 4.29E+04 7.46E+06 1.57E+06 3.51E+06 -5.96 1.43E-04 hsp90b1 7.43E+04 3.79E+04 4.29E+04 2.32E+06 9.23E+06 1.13E+06 -5.87 5.60E-04 mast1a 7.43E+04 1.76E+06 3.58E+04 9.98E+06 9.81E+06 9.29E+06 -5.86 8.13E-03 tspan36 4.91E+05 4.61E+05 4.29E+04 1.69E+07 1.43E+07 5.96E+06 -5.72 2.24E-03 si:ch211-226h8.14 5.63E+05 5.32E+04 9.73E+05 1.50E+07 2.17E+08 1.13E+06 -5.65 4.04E-02 pdia4 7.43E+04 3.03E+05 4.29E+04 5.34E+06 1.11E+07 1.13E+06 -5.36 3.47E-03 tspan3a 3.37E+06 1.84E+06 4.29E+04 2.37E+07 1.61E+07 3.48E+07 -5.20 2.33E-02 itm2ba 7.43E+04 5.32E+04 4.29E+04 5.75E+06 3.17E+05 2.06E+06 -4.81 5.09E-03 tpte 7.43E+04 5.32E+04 4.29E+04 2.73E+06 5.57E+05 2.29E+06 -4.78 6.96E-04 zgc:171717 4.88E+06 1.05E+07 4.67E+07 2.83E+08 7.10E+08 2.09E+08 -4.70 3.31E-03 tspan7b 7.43E+04 6.47E+05 2.56E+05 5.98E+06 3.24E+06 9.73E+06 -4.63 2.60E-03 naalad2 1.37E+05 5.32E+04 4.29E+04 2.18E+06 5.54E+05 3.80E+06 -4.61 2.24E-03 sptbn1 1.52E+05 3.46E+04 3.64E+04 4.23E+05 5.57E+05 5.81E+06 -4.27 1.32E-02 myh9b 1.41E+05 4.38E+03 7.62E+04 4.23E+05 5.57E+05 1.13E+06 -4.15 2.48E-02 myom1a 7.43E+04 9.96E+04 7.12E+03 4.23E+05 5.57E+05 1.13E+06 -4.10 1.13E-02 znf1157 5.46E+04 2.68E+04 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -4.02 4.35E-04 pon3.2 7.43E+04 3.08E+05 4.29E+04 4.60E+05 5.76E+06 1.13E+06 -3.86 1.85E-02 ryr3 3.42E+04 6.01E+04 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -3.85 4.37E-04 prdx4 7.43E+04 7.37E+04 4.29E+04 3.96E+05 1.52E+06 1.13E+06 -3.84 1.14E-03 krt15 7.43E+04 2.88E+04 4.40E+04 4.23E+05 5.57E+05 1.13E+06 -3.82 8.02E-04 jupa 3.59E+04 3.13E+05 4.29E+04 4.23E+05 2.57E+06 1.13E+06 -3.77 1.49E-02 tm9sf2 7.43E+04 5.32E+04 4.29E+04 1.09E+06 6.76E+05 4.87E+05 -3.68 3.42E-04 zgc:174164 7.43E+04 5.32E+04 4.29E+04 5.04E+05 5.09E+05 1.13E+06 -3.58 5.18E-04 mrpl44 7.43E+04 3.04E+04 7.41E+04 4.23E+05 5.57E+05 1.13E+06 -3.55 1.34E-03 myha 7.43E+04 8.94E+04 3.29E+04 4.23E+05 5.57E+05 1.13E+06 -3.42 1.70E-03 calr 7.43E+04 1.94E+05 4.29E+04 4.32E+05 1.44E+06 1.13E+06 -3.39 5.18E-03 bms1 3.62E+04 1.61E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -3.35 4.79E-03 zgc:113102 2.95E+05 2.38E+04 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -3.26 2.13E-02 ddx27 7.43E+04 1.91E+05 2.40E+04 4.23E+05 5.57E+05 1.13E+06 -3.20 1.14E-02 actn3b 7.43E+04 7.60E+04 6.19E+04 4.23E+05 5.57E+05 1.13E+06 -3.19 8.48E-04 emg1 7.43E+04 1.27E+05 4.39E+04 4.23E+05 5.57E+05 1.13E+06 -3.11 2.70E-03 nol11 7.43E+04 2.83E+05 2.39E+04 4.23E+05 5.57E+05 1.13E+06 -3.02 2.40E-02 exosc10 5.28E+04 1.98E+05 4.85E+04 4.23E+05 5.57E+05 1.13E+06 -3.01 7.15E-03 137 Supplementary Table 2.S1 continued Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] vtg4 1.74E+06 5.83E+06 1.12E+07 7.33E+07 5.51E+07 1.39E+07 -2.98 2.26E-02 BX548076.3 7.43E+04 7.93E+05 1.15E+06 4.23E+05 6.71E+07 1.13E+06 -2.96 2.21E-01 rrp12 8.01E+04 2.70E+05 2.59E+04 4.23E+05 5.57E+05 1.13E+06 -2.96 2.21E-02 srpk1a 7.43E+04 3.18E+04 2.41E+05 4.23E+05 5.57E+05 1.13E+06 -2.96 1.49E-02 si:dkey-159f12.2 1.01E+06 5.32E+04 1.06E+06 4.23E+05 5.46E+07 1.13E+06 -2.95 2.25E-01 rpl37 4.89E+07 3.34E+08 2.49E+07 5.39E+08 7.02E+08 4.74E+08 -2.93 2.85E-02 eif4a1a 7.43E+04 4.24E+05 2.20E+04 4.23E+05 5.57E+05 1.13E+06 -2.86 4.82E-02 mtdha 7.43E+04 4.65E+04 2.02E+05 4.23E+05 5.57E+05 1.13E+06 -2.86 8.03E-03 utp4 7.43E+04 1.64E+05 5.75E+04 4.23E+05 5.57E+05 1.13E+06 -2.86 4.23E-03 si:dkey-65b12.6 7.43E+04 1.40E+05 6.76E+04 4.23E+05 5.57E+05 1.13E+06 -2.86 2.71E-03 aqr 7.43E+04 2.61E+05 3.70E+04 4.23E+05 5.57E+05 1.13E+06 -2.85 1.63E-02 ywhabb 7.43E+04 2.53E+05 4.11E+04 4.23E+05 5.57E+05 1.13E+06 -2.81 1.45E-02 ddx54 3.62E+04 1.26E+05 1.72E+05 4.23E+05 5.57E+05 1.13E+06 -2.80 1.10E-02 lsg1 6.40E+04 9.69E+04 1.31E+05 4.23E+05 5.57E+05 1.13E+06 -2.79 2.76E-03 utp20 1.15E+05 1.80E+05 3.96E+04 4.23E+05 5.57E+05 1.13E+06 -2.78 9.79E-03 ddx52 7.43E+04 2.55E+05 5.54E+04 4.23E+05 5.57E+05 1.13E+06 -2.66 1.30E-02 mybphb 7.43E+04 9.33E+04 1.71E+05 4.23E+05 5.57E+05 1.13E+06 -2.60 4.63E-03 cct8 7.43E+04 2.54E+05 7.50E+04 4.23E+05 5.57E+05 1.13E+06 -2.52 1.22E-02 prrc2c 7.43E+04 8.47E+04 2.26E+05 4.23E+05 5.57E+05 1.13E+06 -2.52 9.04E-03 dhx9 7.43E+04 1.24E+05 1.54E+05 4.23E+05 5.57E+05 1.13E+06 -2.52 4.63E-03 tbl3 7.13E+04 4.69E+05 4.32E+04 4.23E+05 5.57E+05 1.13E+06 -2.51 4.83E-02 rbfox2 1.65E+05 2.12E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -2.49 1.95E-02 adar 7.43E+04 1.33E+05 1.66E+05 4.23E+05 5.57E+05 1.13E+06 -2.45 5.85E-03 vtg1 1.70E+07 7.30E+07 7.78E+07 3.72E+08 3.36E+08 1.26E+08 -2.45 2.42E-02 kpnb3 7.43E+04 1.48E+05 1.61E+05 4.23E+05 5.57E+05 1.13E+06 -2.41 6.40E-03 heatr1 6.26E+04 2.70E+05 1.05E+05 4.23E+05 5.57E+05 1.13E+06 -2.41 1.60E-02 ywhae1 7.43E+04 4.43E+05 5.75E+04 4.23E+05 5.57E+05 1.13E+06 -2.38 4.27E-02 wdr75 7.43E+04 1.98E+05 1.30E+05 4.23E+05 5.57E+05 1.13E+06 -2.37 8.36E-03 eef2l2 7.43E+04 6.93E+04 3.78E+05 4.23E+05 5.57E+05 1.13E+06 -2.37 3.07E-02 ddx18 7.43E+04 2.52E+05 1.10E+05 4.23E+05 5.57E+05 1.13E+06 -2.34 1.29E-02 elavl1b 3.16E+05 1.56E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -2.33 3.69E-02 wdr12 7.43E+04 5.67E+05 5.18E+04 4.23E+05 5.57E+05 1.13E+06 -2.31 6.68E-02 mrpl15 1.68E+05 3.04E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -2.31 3.71E-02 utp6 7.43E+04 2.04E+05 1.62E+05 4.23E+05 5.57E+05 1.13E+06 -2.25 1.16E-02 ufl1 7.43E+04 3.85E+05 9.04E+04 4.23E+05 5.57E+05 1.13E+06 -2.23 3.29E-02 crnkl1 7.43E+04 4.22E+05 8.31E+04 4.23E+05 5.57E+05 1.13E+06 -2.23 3.94E-02 bysl 7.43E+04 7.29E+04 4.92E+05 4.23E+05 5.57E+05 1.13E+06 -2.22 5.25E-02 luc7l 1.43E+06 4.52E+04 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -2.20 1.90E-01 ywhaz 1.86E+05 3.60E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -2.18 5.44E-02 rpl29 4.08E+08 1.51E+08 2.27E+08 1.02E+09 9.74E+08 1.27E+09 -2.17 5.55E-03 hspb1 7.43E+04 6.11E+05 6.61E+04 4.23E+05 5.57E+05 1.13E+06 -2.16 7.64E-02 eif2s1a 7.43E+04 1.41E+05 2.92E+05 4.23E+05 5.57E+05 1.13E+06 -2.15 2.17E-02 ktn1 1.14E+05 2.20E+05 1.31E+05 4.23E+05 5.57E+05 1.13E+06 -2.12 9.36E-03 RPL37A 8.70E+08 2.05E+09 3.99E+09 7.94E+09 8.42E+09 8.06E+09 -2.08 1.81E-02 hdlbpa 7.43E+04 1.94E+05 2.47E+05 4.23E+05 5.57E+05 1.13E+06 -2.08 2.17E-02 rcl1 7.43E+04 4.08E+05 1.21E+05 4.23E+05 5.57E+05 1.13E+06 -2.06 4.10E-02 pkhd1l1 7.43E+04 3.29E+05 1.52E+05 4.23E+05 5.57E+05 1.13E+06 -2.05 3.01E-02 cdk5rap3 7.04E+04 3.35E+05 1.76E+05 4.23E+05 5.57E+05 1.13E+06 -2.00 3.63E-02 plrg1 7.43E+04 4.39E+05 1.27E+05 4.23E+05 5.57E+05 1.13E+06 -2.00 4.89E-02 metap1 7.43E+04 1.87E+05 3.14E+05 4.23E+05 5.57E+05 1.13E+06 -1.98 3.32E-02 eif3m 7.43E+04 1.88E+05 3.22E+05 4.23E+05 5.57E+05 1.13E+06 -1.96 3.53E-02 138 Supplementary Table 2.S1 continued Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] flot2a 7.43E+04 4.26E+05 1.44E+05 4.23E+05 5.57E+05 1.13E+06 -1.96 4.94E-02 nme2b.2 7.43E+04 1.42E+06 4.38E+04 4.23E+05 5.57E+05 1.13E+06 -1.95 2.14E-01 ttn.2 1.77E+05 2.33E+05 1.15E+05 4.23E+05 5.57E+05 1.13E+06 -1.94 1.40E-02 abce1 7.43E+04 1.85E+05 3.61E+05 4.23E+05 5.57E+05 1.13E+06 -1.92 4.34E-02 si:ch211-198a12.6 1.05E+05 1.11E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -1.91 1.85E-01 atp2a2a 1.11E+05 1.11E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -1.89 1.88E-01 rps27a 2.13E+08 3.20E+08 6.31E+08 1.34E+09 1.31E+09 1.19E+09 -1.87 1.22E-02 krt17 7.43E+04 3.83E+05 2.16E+05 4.23E+05 5.57E+05 1.13E+06 -1.81 5.69E-02 dnaja2b 7.43E+04 6.20E+05 1.59E+05 4.23E+05 5.57E+05 1.13E+06 -1.73 1.04E-01 ltv1 7.43E+04 2.89E+05 3.45E+05 4.23E+05 5.57E+05 1.13E+06 -1.72 6.75E-02 plecb 7.95E+05 2.69E+05 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -1.62 2.07E-01 tuba8l3 7.43E+04 1.65E+06 9.26E+04 4.23E+05 5.57E+05 1.13E+06 -1.52 2.88E-01 rplp1 2.75E+08 3.16E+08 3.36E+08 7.28E+08 9.78E+08 9.66E+08 -1.52 7.44E-03 ldhba 7.43E+04 1.17E+06 1.99E+05 4.23E+05 5.57E+05 1.13E+06 -1.32 2.74E-01 pbdc1 7.43E+04 1.12E+06 2.09E+05 4.23E+05 5.57E+05 1.13E+06 -1.32 2.68E-01 ufm1 7.43E+04 1.87E+06 1.26E+05 4.23E+05 5.57E+05 1.13E+06 -1.31 3.54E-01 fbxl6 3.30E+05 4.94E+05 1.17E+05 4.23E+05 5.57E+05 1.13E+06 -1.27 1.29E-01 arglu1a 7.43E+04 5.24E+05 6.13E+05 4.23E+05 5.57E+05 1.13E+06 -1.16 2.72E-01 rps27l 3.62E+08 2.08E+08 2.38E+08 5.59E+08 5.92E+08 5.90E+08 -1.15 3.11E-02 cyt1l 7.43E+04 3.44E+05 1.03E+06 4.23E+05 5.57E+05 1.13E+06 -1.12 3.27E-01 naa40 7.43E+04 2.37E+05 1.52E+06 4.23E+05 5.57E+05 1.13E+06 -1.11 3.81E-01 exosc4 7.43E+04 1.06E+06 3.78E+05 4.23E+05 5.57E+05 1.13E+06 -1.06 3.57E-01 npm1a 7.52E+05 4.64E+04 9.01E+05 4.23E+05 5.57E+05 1.13E+06 -1.03 4.46E-01 rps12 3.46E+09 3.58E+09 4.58E+09 7.69E+09 7.87E+09 7.44E+09 -1.00 3.64E-02 si:ch211-106j21.4 8.00E+05 6.38E+04 7.12E+05 4.23E+05 5.57E+05 1.13E+06 -0.96 4.22E-01 sf3b4 7.43E+04 1.41E+06 3.58E+05 4.23E+05 5.57E+05 1.13E+06 -0.94 4.39E-01 h2ax 3.69E+06 8.92E+06 4.29E+04 4.23E+05 1.81E+07 1.13E+06 -0.87 7.32E-01 hnrnpul1 1.14E+06 1.02E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -0.81 5.83E-01 cryba2a 9.85E+05 1.24E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -0.78 5.97E-01 srp9 7.43E+04 7.24E+05 1.01E+06 4.23E+05 5.57E+05 1.13E+06 -0.76 5.16E-01 si:ch73-366l1.5 8.74E+05 1.71E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -0.69 6.54E-01 cryba1b 1.69E+05 1.33E+06 3.04E+05 4.23E+05 5.57E+05 1.13E+06 -0.65 4.92E-01 mdka 7.43E+04 1.61E+06 6.19E+05 4.23E+05 5.57E+05 1.13E+06 -0.62 6.26E-01 imp3 7.43E+04 1.56E+06 6.82E+05 4.23E+05 5.57E+05 1.13E+06 -0.58 6.45E-01 cirbpb 7.43E+04 6.81E+05 1.60E+06 4.23E+05 5.57E+05 1.13E+06 -0.57 6.53E-01 srsf6a 7.43E+04 3.69E+06 4.06E+05 4.23E+05 5.57E+05 1.13E+06 -0.42 7.81E-01 si:dkey-108k21.21 7.43E+04 1.88E+06 9.79E+05 4.23E+05 5.57E+05 1.13E+06 -0.32 8.11E-01 snrpd3 1.54E+06 2.12E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 -0.31 8.52E-01 si:ch211-76m11.8 2.28E+05 7.47E+05 9.70E+05 4.23E+05 5.57E+05 1.13E+06 -0.23 7.63E-01 rbfox1 7.43E+04 1.06E+06 2.22E+06 4.23E+05 5.57E+05 1.13E+06 -0.20 8.85E-01 rpl9 7.43E+04 2.09E+06 1.22E+06 4.23E+05 5.57E+05 1.13E+06 -0.16 9.07E-01 chtopa 7.43E+04 3.64E+06 7.17E+05 4.23E+05 5.57E+05 1.13E+06 -0.15 9.19E-01 tnnt3b 1.94E+06 4.87E+05 2.22E+05 4.23E+05 5.57E+05 1.13E+06 -0.12 9.02E-01 cirbpa 7.43E+04 1.69E+06 1.81E+06 4.23E+05 5.57E+05 1.13E+06 -0.08 9.57E-01 srp19 7.43E+04 1.78E+06 2.01E+06 4.23E+05 5.57E+05 1.13E+06 0.00 9.98E-01 zgc:136908 1.59E+05 1.66E+06 1.19E+06 4.23E+05 5.57E+05 1.13E+06 0.08 9.40E-01 nol12 4.66E+05 1.42E+06 5.11E+05 4.23E+05 5.57E+05 1.13E+06 0.11 8.68E-01 mylpfb 3.39E+06 2.99E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 0.23 9.01E-01 ppiab 8.49E+05 2.15E+06 3.65E+05 4.23E+05 5.57E+05 1.13E+06 0.44 6.00E-01 mrpl41 1.18E+06 2.32E+06 2.71E+05 4.23E+05 5.57E+05 1.13E+06 0.49 6.11E-01 flna 2.25E+06 6.77E+05 5.48E+05 4.23E+05 5.57E+05 1.13E+06 0.55 4.80E-01 139 Supplementary Table 2.S1 continued Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] hnrnpa0l 7.43E+04 4.10E+06 2.80E+06 4.23E+05 5.57E+05 1.13E+06 0.56 7.42E-01 ccdc59 2.48E+06 5.74E+05 6.20E+05 4.23E+05 5.57E+05 1.13E+06 0.57 4.78E-01 purbb 1.27E+07 4.51E+05 1.77E+05 4.23E+05 5.57E+05 1.13E+06 0.64 7.09E-01 snrpg 7.43E+04 2.56E+06 6.22E+06 4.23E+05 5.57E+05 1.13E+06 0.72 6.89E-01 pfdn4 7.43E+04 4.59E+06 4.83E+06 4.23E+05 5.57E+05 1.13E+06 0.87 6.33E-01 eif1axa 1.94E+06 8.85E+05 9.97E+05 4.23E+05 5.57E+05 1.13E+06 0.89 1.75E-01 tmem67 7.54E+08 4.00E+08 3.92E+08 2.51E+08 2.54E+08 2.71E+08 0.93 8.02E-02 srsf3a 5.93E+05 1.61E+06 3.12E+06 4.23E+05 5.57E+05 1.13E+06 1.16 1.82E-01 snrpe 7.43E+04 8.61E+06 5.06E+06 4.23E+05 5.57E+05 1.13E+06 1.20 5.47E-01 tp53inp2 7.43E+04 1.89E+07 3.09E+06 4.23E+05 5.57E+05 1.13E+06 1.34 5.33E-01 rps27.1, rps27.2 2.60E+09 1.44E+09 1.28E+09 6.71E+08 6.13E+08 6.54E+08 1.39 2.10E-02 skiv2l 1.54E+06 1.77E+06 2.19E+06 4.23E+05 5.57E+05 1.13E+06 1.49 2.81E-02 zgc:153867 3.25E+06 1.15E+06 1.70E+06 4.23E+05 5.57E+05 1.13E+06 1.52 5.01E-02 gadd45gip1 4.20E+07 3.56E+06 4.29E+04 4.23E+05 5.57E+05 1.13E+06 1.53 5.60E-01 hmgb2b 1.26E+06 3.13E+06 1.66E+06 4.23E+05 5.57E+05 1.13E+06 1.54 4.25E-02 eef1a1l1 2.00E+06 4.97E+06 2.51E+06 1.37E+06 5.57E+05 1.13E+06 1.62 3.35E-02 eif3i 1.09E+06 1.97E+06 3.78E+06 4.23E+05 5.57E+05 1.13E+06 1.64 4.82E-02 crygn2 1.60E+06 3.20E+06 1.60E+06 4.23E+05 5.57E+05 1.13E+06 1.65 2.90E-02 rpl10 1.19E+08 8.41E+07 7.63E+07 2.18E+07 3.97E+07 2.61E+07 1.69 9.34E-03 hsp90ab1 1.06E+06 3.82E+06 2.74E+06 4.23E+05 5.57E+05 1.13E+06 1.79 4.01E-02 eif3ba 1.49E+06 2.66E+06 2.94E+06 4.23E+05 5.57E+05 1.13E+06 1.82 1.85E-02 srpk1b 1.76E+06 3.30E+06 2.13E+06 4.23E+05 5.57E+05 1.13E+06 1.84 1.59E-02 hmgb3a 1.18E+06 3.61E+06 3.59E+06 4.23E+05 5.57E+05 1.13E+06 1.95 2.85E-02 rtn2a 1.27E+06 9.98E+06 1.35E+06 4.23E+05 5.57E+05 1.13E+06 2.00 8.32E-02 ilf3b 1.85E+06 1.76E+06 5.30E+06 4.23E+05 5.57E+05 1.13E+06 2.00 2.41E-02 mrpl12 2.13E+06 6.13E+06 1.47E+06 4.23E+05 5.57E+05 1.13E+06 2.06 2.97E-02 syncrip 3.65E+06 3.41E+06 1.74E+06 4.23E+05 5.57E+05 1.13E+06 2.12 1.09E-02 eif3ha 3.32E+06 2.85E+06 2.38E+06 4.23E+05 5.57E+05 1.13E+06 2.13 6.40E-03 mvp 7.31E+06 1.38E+07 4.58E+06 3.51E+06 1.12E+06 1.32E+06 2.16 1.97E-02 larp1b 3.47E+06 3.03E+06 2.27E+06 4.23E+05 5.57E+05 1.13E+06 2.16 6.55E-03 ppan 1.36E+06 5.12E+06 3.63E+06 4.23E+05 5.57E+05 1.13E+06 2.19 2.02E-02 eif5a 2.99E+06 3.54E+06 2.54E+06 4.23E+05 5.57E+05 1.13E+06 2.22 5.35E-03 snd1 4.27E+06 5.84E+06 2.72E+06 9.19E+05 5.57E+05 1.13E+06 2.29 4.54E-03 eif3s10 2.94E+06 3.89E+06 2.76E+06 4.23E+05 5.57E+05 1.13E+06 2.30 4.69E-03 rsl1d1 1.72E+06 3.85E+06 4.80E+06 4.23E+05 5.57E+05 1.13E+06 2.30 1.10E-02 eif4a3 2.22E+06 6.50E+06 2.54E+06 4.23E+05 5.57E+05 1.13E+06 2.37 1.10E-02 mvp 1.07E+09 1.10E+09 5.20E+08 3.32E+08 1.06E+08 1.21E+08 2.39 9.88E-03 naa15a 3.16E+06 2.46E+06 5.07E+06 4.23E+05 5.57E+05 1.13E+06 2.40 5.63E-03 nop2 1.39E+06 5.13E+06 5.60E+06 4.23E+05 5.57E+05 1.13E+06 2.41 1.81E-02 rnps1 3.83E+06 4.98E+06 2.28E+06 4.23E+05 5.57E+05 1.13E+06 2.45 5.59E-03 tubb5 3.86E+06 2.98E+06 3.79E+06 4.23E+05 5.57E+05 1.13E+06 2.45 3.27E-03 ckmb 3.26E+06 7.08E+06 1.96E+06 4.23E+05 5.57E+05 1.13E+06 2.47 1.11E-02 akap8l 2.57E+06 4.65E+06 3.91E+06 4.23E+05 5.57E+05 1.13E+06 2.48 4.13E-03 pfdn6 2.12E+06 5.27E+06 4.56E+06 4.23E+05 5.57E+05 1.13E+06 2.52 6.31E-03 rpl7l1 3.00E+06 6.24E+06 2.74E+06 4.23E+05 5.57E+05 1.13E+06 2.53 5.59E-03 eif3c 2.40E+06 3.46E+06 6.21E+06 4.23E+05 5.57E+05 1.13E+06 2.53 6.05E-03 si:dkey-183i3.5 1.52E+06 1.48E+07 2.33E+06 4.23E+05 5.57E+05 1.13E+06 2.54 4.23E-02 eif3s6ip 5.80E+06 4.18E+06 2.87E+06 4.23E+05 5.57E+05 1.13E+06 2.68 3.27E-03 tuba8l4 1.86E+06 7.58E+06 5.34E+06 4.23E+05 5.57E+05 1.13E+06 2.71 9.51E-03 srsf1a 4.21E+06 2.65E+06 6.80E+06 4.23E+05 5.57E+05 1.13E+06 2.72 4.26E-03 hnrnpaba 1.77E+06 6.29E+06 7.52E+06 4.23E+05 5.57E+05 1.13E+06 2.77 1.04E-02 140 Supplementary Table 2.S1 continued Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] tuba8l 3.00E+06 9.41E+06 2.98E+06 4.23E+05 5.57E+05 1.13E+06 2.77 7.05E-03 krt91 1.45E+06 9.71E+06 6.71E+06 4.23E+05 5.57E+05 1.13E+06 2.82 1.78E-02 ebna1bp2 2.64E+07 1.16E+07 5.08E+06 6.91E+06 5.57E+05 1.13E+06 2.83 4.73E-02 zgc:163061 6.71E+06 1.15E+07 1.29E+06 4.23E+05 5.57E+05 1.13E+06 2.85 2.37E-02 zgc:56095 7.56E+07 5.00E+07 9.13E+07 8.43E+06 1.53E+07 6.60E+06 2.89 1.47E-03 fbl 4.96E+06 6.09E+06 3.63E+06 4.23E+05 5.57E+05 1.13E+06 2.89 1.79E-03 znf622 9.58E+05 1.13E+07 1.03E+07 4.23E+05 5.57E+05 1.13E+06 2.90 3.90E-02 eif3ea 8.18E+06 2.48E+06 5.81E+06 4.23E+05 5.57E+05 1.13E+06 2.93 4.71E-03 ftsj3 5.23E+06 5.80E+06 3.89E+06 4.23E+05 5.57E+05 1.13E+06 2.93 1.51E-03 bxdc2 3.81E+06 1.00E+07 3.09E+06 4.23E+05 5.57E+05 1.13E+06 2.93 4.90E-03 btf3 6.56E+08 6.95E+08 1.24E+09 6.39E+07 4.12E+08 4.53E+07 2.96 2.00E-02 rrs1 2.26E+06 6.45E+06 8.90E+06 4.23E+05 5.57E+05 1.13E+06 2.97 6.07E-03 srsf7a 7.12E+06 3.45E+06 5.97E+06 4.23E+05 5.57E+05 1.13E+06 3.03 1.90E-03 icn2 7.84E+06 1.04E+07 2.36E+06 4.23E+05 5.57E+05 1.13E+06 3.16 5.73E-03 rpf2 3.91E+06 4.91E+06 1.02E+07 4.23E+05 5.57E+05 1.13E+06 3.17 2.23E-03 rsl24d1 3.69E+06 6.55E+06 8.26E+06 4.23E+05 5.57E+05 1.13E+06 3.18 1.68E-03 nsa2 4.99E+06 7.18E+06 6.52E+06 4.23E+05 5.57E+05 1.13E+06 3.26 8.52E-04 hbae1.3 7.18E+06 1.32E+07 2.67E+06 4.23E+05 5.57E+05 1.13E+06 3.30 5.05E-03 hnrnpa1a 1.82E+06 7.60E+06 1.92E+07 4.23E+05 5.57E+05 1.13E+06 3.32 1.46E-02 zc3h15 3.30E+06 8.98E+06 9.27E+06 4.23E+05 5.57E+05 1.13E+06 3.34 2.31E-03 calm3a 7.34E+07 4.58E+07 2.63E+07 1.47E+07 5.13E+06 1.13E+06 3.34 1.83E-02 eif6 4.39E+06 1.75E+07 2.63E+07 4.17E+05 4.08E+06 1.13E+06 3.34 2.28E-02 apoa2 4.59E+06 1.49E+07 4.20E+06 4.23E+05 5.57E+05 1.13E+06 3.36 3.35E-03 si:dkey-12j5.1 1.23E+07 7.43E+06 3.68E+06 4.23E+05 5.57E+05 1.13E+06 3.43 2.14E-03 gnl3 5.30E+06 9.15E+06 7.23E+06 4.23E+05 5.57E+05 1.13E+06 3.45 7.55E-04 ifrd1 1.92E+07 5.08E+06 3.62E+06 4.23E+05 5.57E+05 1.13E+06 3.46 5.12E-03 naa15b 6.17E+06 5.43E+06 1.35E+07 4.23E+05 5.57E+05 1.13E+06 3.57 1.19E-03 igf2bp1 4.20E+06 1.17E+07 9.92E+06 4.23E+05 5.57E+05 1.13E+06 3.61 1.38E-03 ncl 2.24E+07 9.76E+06 2.45E+06 4.23E+05 5.57E+05 1.13E+06 3.66 8.01E-03 krt5 3.07E+06 1.49E+07 1.34E+07 4.23E+05 5.57E+05 1.13E+06 3.72 3.66E-03 tnnc2 1.96E+07 5.15E+06 6.31E+06 4.23E+05 5.57E+05 1.13E+06 3.74 2.07E-03 tnnt3b 2.41E+07 5.14E+06 5.28E+06 4.23E+05 5.57E+05 1.13E+06 3.75 3.53E-03 actb1 1.78E+06 1.43E+07 1.19E+07 1.73E+05 5.57E+05 1.13E+06 3.81 1.41E-02 srsf4 1.50E+07 8.32E+06 6.47E+06 4.23E+05 5.57E+05 1.13E+06 3.85 6.66E-04 lyar 2.51E+06 1.86E+07 2.15E+07 4.23E+05 5.57E+05 1.13E+06 3.96 7.09E-03 khdrbs1a 6.62E+06 1.86E+07 8.87E+06 4.23E+05 5.57E+05 1.13E+06 4.00 7.73E-04 snrpd2 1.79E+07 1.24E+07 5.15E+06 4.23E+05 5.57E+05 1.13E+06 4.02 1.09E-03 naa10 4.78E+06 1.31E+07 2.07E+07 4.23E+05 5.57E+05 1.13E+06 4.08 1.49E-03 apoa1a 4.09E+06 2.42E+07 1.44E+07 4.23E+05 5.57E+05 1.13E+06 4.13 2.47E-03 snrpd3l 9.46E+06 1.58E+07 9.71E+06 4.23E+05 5.57E+05 1.13E+06 4.14 3.04E-04 si:dkey-164f24.2 2.97E+07 1.21E+07 4.58E+06 4.23E+05 5.57E+05 1.13E+06 4.20 2.43E-03 si:ch211-113a14.11 1.87E+07 4.02E+07 2.20E+07 3.98E+06 5.57E+05 1.13E+06 4.23 2.43E-03 srsf3b 5.66E+06 1.33E+07 2.86E+07 4.23E+05 5.57E+05 1.13E+06 4.32 1.38E-03 si:dkey-108k21.10 2.27E+07 1.97E+07 5.67E+06 4.23E+05 5.57E+05 1.13E+06 4.40 1.06E-03 srsf5b 1.05E+07 1.75E+07 1.54E+07 4.23E+05 5.57E+05 1.13E+06 4.46 1.91E-04 myhz1.1 2.14E+07 1.59E+07 9.16E+06 4.23E+05 5.57E+05 1.13E+06 4.50 2.95E-04 si:ch73-368j24.11 1.25E+07 2.23E+07 1.14E+07 4.23E+05 5.57E+05 1.13E+06 4.51 2.38E-04 serbp1a 1.10E+09 6.13E+08 9.22E+08 8.03E+07 2.78E+07 2.27E+07 4.53 4.17E-04 gtpbp4 1.52E+07 1.83E+07 1.18E+07 4.23E+05 5.57E+05 1.13E+06 4.53 1.59E-04 atp5f1b 1.71E+07 4.54E+07 4.33E+06 4.23E+05 5.57E+05 1.13E+06 4.54 3.59E-03 LO017835.1 4.14E+06 2.27E+07 3.70E+07 4.23E+05 5.57E+05 1.13E+06 4.56 3.21E-03 141 Supplementary Table 2.S1 continued Signal Area (5 dpf) Signal Area (6 hpf) log2fc p-value gene name rep I rep II rep III rep I rep II rep III [5d/6h] [5d/6h] myl1 7.60E+06 2.84E+07 1.80E+07 4.23E+05 5.57E+05 1.13E+06 4.61 6.03E-04 fkbp1aa 1.42E+07 1.88E+07 1.88E+07 4.23E+05 5.57E+05 1.13E+06 4.73 1.13E-04 si:ch211-113a14.12 2.07E+07 2.41E+07 1.69E+07 4.23E+05 5.57E+05 1.13E+06 4.98 8.75E-05 mylz3 2.20E+07 1.81E+07 2.82E+07 4.23E+05 5.57E+05 1.13E+06 5.12 8.20E-05 mrto4 1.39E+07 3.08E+07 3.52E+07 4.23E+05 5.57E+05 1.13E+06 5.26 1.63E-04 drg1 5.99E+07 2.39E+07 1.10E+07 4.23E+05 5.57E+05 1.13E+06 5.28 5.66E-04 srsf6b 1.22E+07 3.77E+07 4.34E+07 4.23E+05 5.57E+05 1.13E+06 5.40 2.90E-04 srsf5a 1.92E+07 3.47E+07 3.14E+07 4.23E+05 5.57E+05 1.13E+06 5.42 7.64E-05 actc1b 2.90E+08 1.16E+08 1.05E+08 4.60E+06 7.00E+06 1.13E+06 5.52 7.17E-04 si:ch211-133n4.4 4.97E+06 6.19E+07 1.05E+08 4.23E+05 5.57E+05 1.13E+06 5.63 4.30E-03 tpp2 3.02E+07 4.63E+07 2.50E+07 4.23E+05 5.57E+05 1.13E+06 5.67 5.93E-05 mylpfa 3.79E+07 3.06E+07 3.12E+07 4.23E+05 5.57E+05 1.13E+06 5.68 3.86E-05 cyt1 1.84E+07 6.86E+07 5.18E+07 4.23E+05 5.57E+05 1.13E+06 5.97 1.67E-04 pvalb2 3.41E+07 1.18E+08 2.05E+07 4.23E+05 5.57E+05 1.13E+06 6.08 3.27E-04 gapdh 5.44E+06 4.43E+08 4.18E+08 4.23E+05 6.72E+06 1.13E+06 6.09 2.55E-02 fthl27 9.02E+07 3.00E+07 4.00E+07 4.23E+05 5.57E+05 1.13E+06 6.21 8.49E-05 tuba1a 3.93E+07 8.84E+07 3.37E+07 4.23E+05 5.57E+05 1.13E+06 6.25 6.78E-05 hspa8 5.78E+07 9.34E+07 7.87E+07 4.23E+05 1.86E+06 1.13E+06 6.29 9.15E-05 zgc:55461 3.07E+07 1.03E+08 4.07E+07 4.23E+05 5.57E+05 1.13E+06 6.29 9.95E-05 tpma 5.27E+08 3.43E+07 3.61E+07 4.23E+05 5.57E+05 1.13E+06 7.07 1.19E-03 llph 8.31E+07 9.11E+07 9.01E+07 4.23E+05 5.57E+05 1.13E+06 7.10 1.05E-05 serbp1b 1.66E+08 7.71E+07 1.23E+08 4.23E+05 5.57E+05 1.13E+06 7.50 1.56E-05 atp2a1 2.08E+08 2.56E+08 5.44E+07 4.23E+05 5.57E+05 1.13E+06 7.79 6.82E-05 CABZ01093502.2 1.20E+09 8.84E+07 8.44E+07 4.23E+05 5.57E+05 1.13E+06 8.33 4.53E-04 rpl22l1 4.75E+08 9.10E+08 6.66E+08 4.23E+05 1.26E+07 1.13E+06 8.50 6.68E-04 zdhhc20b 4.69E+08 2.37E+08 2.48E+08 4.23E+05 5.57E+05 1.13E+06 8.87 5.94E-06 rpl5a 9.11E+08 7.62E+08 6.04E+08 4.23E+05 5.57E+05 1.13E+06 10.18 1.68E-06 142 Supplementary Table 2.S2: Summary of cryoEM data collection, image processing, model building, refinement and statistics. 1 hpf zebrafish 6 hpf zebrafish 5 dpf zebrafish PDB ID 7OYA 7OYB unpublished Data collection Microscope Thermo Fisher Titan Krios Acceleration Voltage (kV) 300 Detector Falcon 3 EC Falcon 3 EC Falcon 3 EC Magnification (nominal) 81 81 81 Defocus range (µm) 0.5-4 0.5 -3 Calibrates pixel size (Å/px) 1.1 1.04 1.061 Electron exposure (e-/Å2) 43 48 40 Exposure rate (e-/Å2/s) 43 48 40 Number of frames per movie 39 39 Collection software Cryosparc 3.2.0 Cryosparc 3.2.0 Cryosparc 3.2.0 Number of micrographs 17,040 11,860 9,456 Initial particle number 1,961,364 1,285,670 1,049,941 Final particle number 535,633 775,288 569,541 Map resolution (Å, FSC=0.143) 3.2 2.6 2.8 Refinement Software Phenix 1.17.1 Phenix 1.17.1 Phenix 1.17.1 Initial model(s) 4UG0 4UG0 4UG0 6MTE 5DAT Correlation coefficient (CCmask) 0.83 0.84 0 .53 Map sharpening factor (Å2) -108.5 -89.2 Model composition (chains) 82 77 Non-hydrogen atoms 199,914 194,870 Protein residues 11,179 10,795 Nucleotides 5,130 5,035 Ligands MG: 202 MG: 203 ZN: 7 ZN: 7 B factors (Å2, min/max/mean) Protein 4.4 / 281.9 / 53.3 12.8 / 350.6 / 51.7 19.5 / 223.05/ 72.8 RNA 13.9 / 234.1 / 53.1 13.9 / 205.7 / 56.6 19.5 / 223.05/ 72.8 Ligands 7.8 / 102.4 / 28.4 7.8 / 102.4 / 27.0 57.6 / 131.99/ 94.8 RMS deviations Bond length (Å) (#>4 sigma) 0.003 0.003 0.002 Bond angle (°) (#>4 sigma) 0 .62 0 .614 0 .448 Validation MolProbity score 1.59 1.72 1.88 Clashscore 6.04 8.01 10.30 Poor Rotamers (%) 1.3 1.32 1.27 Cß deviations (%) 0.01 0 0 CABLAM outliers (%) 2.35 2.36 175 Favored (%) 97.03 96.86 96 Allowed (%) 2.97 3.14 4 Disallowed (%) 0 0 0 143 Supplementary Table 2.S3: Structural comparison of sequence differences between maternal and somatic rRNA variants in zebrafish maternal (6 hpf) and somatic (5 dpf) ribosome models. 144 Supplementary Table 2.S4: Primer sequences and PCR conditions used to detect maternal and somatic rRNA variants by PCR-based fragment length polymorphism (FLP) and RT-qPCR. Primer Sequence PCR Products Source 18S_ES3S_FLP_F CTAATACATGCCAACGAGCGCC NEB Taq 200 bp from Maternal 18S MIT 18S_ES3S_FLP_R CCGAGGTTATCTAGAGTCACCAAAGC 62 °C 170 bp from Somatic 18S IMP 18S_Mat_F CGTGGGCGGTGGAGAGG KAPA SYBR MIT 18S_Mat_R CCATGGTAGGCGACGGACC 68 °C 169 bp from Maternal 18S 18S_Som_F GCGGCGTGGGCTCCCCCTTCG KAPA SYBR MIT 18S_Som_R CCATGGTAGGCGCCTAAAG 68 °C 158 bp from Somatic 18S 28S_ES31L_FLP_F1 [M13]GGTGAAATACCACTACTCTTATCG NEBNext Taq 254* bp from Maternal 28S MIT 28S_ES31L_FLP_R1 [PIG]GTTACCGTTTGACAGGTGTAC 68 °C 210* bp from Somatic 28S 28S_ES31L_FLP_F2 CTCTTATCGTTTCCTCACTTACCCG Q5 + 1M betaine 196 bp from Maternal 28S IMP 28S_ES31L_FLP_R2 TACCGCCCCAGTCAAACTCC 70 °C 152 bp from Somatic 28S 28S_Mat_F AGGTGCCCTGACCCCCGTTCCCGAGCCG KAPA SYBR MIT 28S_Mat_R CGCCCGACCCCCGCGGACGGGGAAAG 68 °C 155 bp from Maternal 28S 28S_Som_F CGGCGCCCCCTCTCGTTCCCGTCTCC KAPA SYBR MIT 28S_Som_R CCTCCCCCACCCGAAGGCGGGGGC 68 °C 110 bp from Somatic 28S The 18S_ES3S_FLP primer pair was identically and independently designed by both A.N.S and F.L. *The 28S_ES31L_FLP_F1 primer contains a 5’ M13F tail (19-nucleotides) and the 28S_ES31L_FLP_R1 primer contains a 5’ PIG tail (6-nucleotides) adding 25 bp to the final product length. These were included to allow a third primer (5’ FAM-labelled M13F) to generate fluorescent PCR products for quantification using formamide denaturing capillary electrophoresis (Varshney et al. 2015). Note the 28S PCR FLP in Fig. 2.4B uses pair 1 (generating 254 bp and 210 bp products) while the 28S PCR FLP in Supplementary Fig. 2.S2B uses pair 2 (generating 196 bp and 152 bp products). 145 Acknowledgments We would like to thank Anton Meinhart (IMP) for his support in Cryo-EM analyses; the Mass Spectrometry Facility at the Vienna BioCenter Core Facilities (VBCF) headed by Elisabeth Roitinger and Karl Mechtler, in particular Richard Imre, for the processing and analysis of proteomics data; the VBCF Electron Microscopy Facility for the support and maintenance of the EM facility; the IMP aquatics facility personnel, in particular F. Ecker, K. Rattner, J. König and D. Sunjic, for their excellent care of fish, and the Koch Institute Frontier Research Program, the Casey and Family Foundation Cancer Research Fund, the Michael (1957) and Inara Erdei Fund, the Swanson Biotechnology Center Microscopy Core, Grace Phelps from Dr. Lee’s lab at MIT for providing tumor-inducing zebrafish lines; the Koch Institute Zebrafish Core Facility, headed by Adam Amsterdam, and MIT’s BioMicro Center for their support and the VBC RNA Salon and the RNA-Deco-SFB community for providing useful feedback and suggestions. We would also like to thank the entire Pauli and Calo groups for valuable discussions on the project, and for feedback on the manuscript. Funding: Work in the Pauli lab was supported by the Institute of Molecular Pathology (IMP), which receives institutional funding from Boehringer Ingelheim and the Austrian Research Promotion Agency (Headquarter grant FFG-852936), and by the European Research Council (ERC) consolidator grant (‘GaMe’, 101044495 to A.P.), the FWF START program (Y 1031-B28 to A.P.), the Human Frontier Science Program (HFSP) Career Development Award (CDA00066/2015 to A.P.), the SFB RNA-Deco (project number F 80 to A.P.) and a HFSP Young Investigator Grant (RGY0079/2020 to A.P.). L.L.-O. was supported by an SNF Early Postdoc Mobility fellowship (P2GEP3_191204), an EMBO long-term fellowship (ALTF 1165-2019) and an MSCA-IF-EF-SE (890218). We acknowledge the cryo-EM facility CEITEC MU of CIISB, Instruct-CZ Centre, supported by MEYS CR (LM2018127), and Diamond Light Source for granting us access and support at the cryo-EM facilities at the UK’s National Electron Bio-imaging Centre (eBIC) under proposal EM BI25222, funded by the Wellcome Trust, MRC and BBSRC. Work in the Calo lab was supported by the National Institute of General Medical Sciences (R35GM142634), the Pew Charitable Trusts, and the National Cancer Institute (P30- CA14051 to E.C.). A.N.S was supported by the National Science Foundation Graduate Research Fellowship under Grant Number 174530. For the purpose of Open Access, the authors have applied a CC BY public copyright license to any Author Accepted Manuscript version arising from this submission. 146 Author Contributions A.N.S. conceptualization, data curation, formal analysis, investigation, methodology (in addition to general parts, hybrid ribosome and germline experiments), writing F.L. conceptualization, data curation, formal analysis, investigation, methodology (in addition to general parts, structural sample preparation and analysis), writing L.L.-O. data curation, formal analysis, investigation, methodology (model building of the 5 dpf ribosome), writing L.E.G. formal analysis, investigation, methodology, validation (CryoEM analysis) C.P. methodology M.N. investigation, methodology (RNA-Seq analysis) I.G. methodology (CryoEM data acquisition) D.H. data curation, formal analysis, investigation, methodology, supervision, validation, writing E.C. conceptualization, project administration, supervision, funding acquisition, writing A.P. conceptualization, project administration, supervision, funding acquisition, writing 147 Chapter III There is still the question [of] whether the nucleoli of egg cells and of somatic cells should be considered homologous. – Thomas H. Montgomery Jr. 1898 148 Chapter III preface In Chapter 0, a question inspiring this Thesis work is given. The ribosome is a molecular machine required by all organisms to execute the gene expression programming required for cellular life. Expression of cellular identities across time and space serve as the basis for complex organismal functions: composition, metabolism, growth, adaptation, response, reproduction, and homeostasis. In Chapter I, the ribosome is introduced as the abundant multi-component complex found in all living systems. The Central Dogma and the structures performing the regulated steps of protein synthesis are described. Focus is given to ribosome heterogeneity and the potential impact it could have on the molecular- cellular complexity of cells and organisms. While all cells inherit copies of the genome, the germline is a specialized cell lineage that houses the genome to be inherited by the next generation. In Chapter II, the domesticated zebrafish, Danio rerio, commonly found in pet stores, is the system used for inquiry. Laboratory strains of the model organism were used for measurement and experimentation involving two variations of ribosome: one specific to germ cells and the other specific to somatic cells. In Chapter III, the perplexing question regarding “why” zebrafish have two developmentally regulated ribosomes is used as a framework to summarize findings made during examination of the system, to discuss implications of our conclusions on current research, and to provide future directions for continued investigation. The rDNA gene, ribosome biogenesis, 40S and 60S maturation, and translation control in germ cells are the ribosomal gene products of interest. 149 Summary and Model Figure 3.1 – Gene expression and the regulated biogenesis of molecular-cellular complexity A schematic of a eukaryotic cell and the topics discussed in Chapter I (see Fig 1.6 for explanations of each of the illustrations. The gDNA (left) is depicting as containing multiple types of genes: rDNA (grey), protein coding (yellow), and ncRNA (multi). These are transcribed by RNA polymerases I, II, and III, respectively. The ribosomal DNA (rDNA) gene holds information on how to build a ribosome and exists in many multiple repetitive copies in NORs. RNA Polymerase I (Pol I) transcribes rDNA into rRNA (Fig. 3.1, top). During ribosome biogenesis in the nucleolus, nascent rRNAs, 80 RPs, the 5S RNP, and hundreds of components of the biogenesis machinery are involved in the maturation of two ribosomal subunits. Concentration and availability of 40S and 60S subunits, the fundamental units of translation, can influence gene expression via translational control of specific mRNAs. From elsewhere in the genome (gDNA), RNA Polymerase II (Pol II) (Fig. 3.1, middle) transcribes protein-coding genes into mRNAs which require recognition by an initiating ribosome to synthesize the encoded protein according to the Central Dogma of gene expression. A variety of non-coding RNA genes (ncRNAs) are transcribed by RNA Polymerase III (Fig. 3.1, bottom) and used by the cell to maintain gene expression: 5S RNP, structuring the P-site of the 60S and the B1b/c intersubunit bridge, is formed by 5S rRNA, transfer RNAs (tRNAs) used for translating mRNA codons during protein synthesis, and 7SL RNA forms the signal recognition particle (SRP) to halt and localize the elongating ribosome. 150 The probability of any given mRNA gaining access to the ribosome, aka the mRNA- specific initiation rate, is set by the particular combination of its primary nucleotide sequence and the mixture of RNA-binding proteins (RBPs), translation associated factors, ribosome associated factors, ribosomal subunits, and tRNAs. Tuning the quantity and availability of these components can control translational output generally for all mRNAs or specifically for some mRNAs in the system. Cell type-specific programming utilizes both transcriptional and translational controls to regulate gene expression across time and space. These system-level controls allow for dynamic regulations seen in tissues constituting an organism. This includes the egg – a totipotent cell – assembled with the reproductive capability of generating a new organism. Ribosome heterogeneity involves the variation among constructions during ribosome biogenesis. The multicomponent nature of the ribosome allows for structural and compositional heterogeneity in its makeup. Like many other structure/function relationships, identifying how the arrangement of system components (structure) gives rise to a process (function) is critical for mechanistic understanding. Ribosome functional heterogeneity may arise given variation in these structures. Further, selective pressures on gametes could lay the foundation for germ cell-specific ribosome functionalization. Locati and Pagano and colleagues recently discovered and reported on a dual ribosomal system in zebrafish (Locati et al., 2017a; Locati et al., 2017a). The rRNA composing ribosomes maternally-deposited into eggs are exclusively transcribed from a specific rDNA gene variant on chromosome 4, while ribosomes in somatic cells are exclusively transcribed from a specific rDNA gene variant on chromosome 5. The dynamic replacement of maternal ribosomes for somatic ribosomes starts at 8-10 hours post- fertilization (hpf) and by 120 hpf, over 95% of embryonic ribosomes contain somatic type rRNA (Fig. 3.2). Sequence divergences between the two rDNA genes are largely observed in expansion segment regions of the rRNA – possibly aiding interaction with ribosome accessory factors. Figure 3.2 – Developmentally regulated maternal and somatic rRNAs in zebrafish A colorimetric representation of rRNA composition in embryonic and larval stage zebrafish at the indicated hours post-fertilization, illustrated below. M-type (yellow) and S-type (blue) refer to maternal and somatic rDNA variants respectively. At 24 hpf, similar amounts of each rRNA are detected in whole embryo lysates. 151 In what would otherwise be a simple case of heterogeneous ribosomes, this system exhibits all three components thought to potentially drive ribosome functionalization: intragenomic rDNA variant genes, variation in ribosome biogenesis leading to ribosome heterogeneity (structural and compositional changes with possible functional outcomes), and germline-based selective pressures which can be overcome by optimizations in gene expression. The temporal concurrence of maternal-rRNA-containing ribosomes with oocyte and embryonic mRNA, and the dynamic shift to somatic-rRNA containing ribosomes in embryonic and larval stages may be grounds for ribosome functionalization and potential ribosome-specific mRNA-specific translation (Fig. 3.3). Figure 3.3 – Model of the duplicated ribosomal gene products in zebrafish The zebrafish dual ribosomal system is schematized. Maternal (yellow) and somatic (blue) rDNA genes are transcribed by Pol I during ribosome biogenesis to form 40S and 60S subunits. Each set of subunits may exhibit differences in gene expression. Nuclei of a stage Ib oocyte and an epithelial cell are shown as trace outlines with grey fills for nucleoli. The somatic cell nucleus (bottom) is enlarged 8x for illustrative effect. Taking from the findings presented in this Thesis, the available and memorable literature, and scientific discussions with colleagues, I organized Chapter III to discuss the functional implications of ribosome heterogeneity using a perspective framed by separating each ribosome gene products. The following products allow for interactions at multiple levels of biological organization (Fig. 0.2) and are discussed as player of functionality involving the ribosomal gene: o Molecular – protein synthesis is the process enabled by the rDNA gene. It is made possible by the enzymatic functions of the ribosome and dynamic interactions with components of the associated translational machinery. This is measured by the copy number of synthesized proteins, aka translational output (Qt). Is ribosome heterogeneity acting as a layer of regulation on gene expression? Does this putative regulatory scheme mechanistically occur at initiation, elongation, or termination? Do the ribosomes translate different sets of mRNAs? Or is ribosome functionalization via ribosome-specific translation control an idealization of an ancient molecule with multiple interactions? o Molecular – The 40S and 60S ribosomal subunits are the RNP products of the rDNA gene. Maturation, quality control, and storage of subunits are a few of the molecular 152 interactions involving subunit structures outside of translation. The ribosome subunit copy number (R) is measured by UV absorbance (A260), often in gradient conditions. How different are the ribosomal subunits in structure and composition? How do maturation, localization, and quality control processes generate heterogeneous ribosomes with altered functions? o Cellular – The nucleolus is the organellar product of the rDNA gene. Active transcription of ribosomal RNAs during ribosome biogenesis nucleates a structural arrangement of the membrane-less organelle. The nucleolus, its size, copy number, and configuration are detected in cells by imaging. Posed over 125 years ago by Thomas Montgomery – there is still the question of whether the nucleoli of egg cells and of somatic cells should be considered homologous. How different are the nucleoli? Are the components of ribosome biogenesis different among somatic cell types? Between germ and soma? Do nucleolar configurations or outputs vary as well? Is germ cell ribosome biogenesis mechanistically distinct? Possibly to aid in ribosome storage? Are developmental ribosomopathy nucleoli similar to low fertility germ cells? o Genomic – Chromatin regulation by the ~11.6 kbp rDNA gene nucleotide sequence is yet another compelling product. As explained earlier, rDNA genes are notorious for gene instability. The copy number and the specific locus of rDNA genes influences heterochromatinization outcomes of each rDNA gene and its genomic neighborhood. While the cell-specific regulatory status of any given DNA region can be directly measured using chromatin accessibility, DNA marks, or histone modifications, the transcriptional output (RNA) from the region is often a proxy. The NOR is a hot-spot for genomic recombination events as homology directed repair machinery will identify many possible repair templates for dsDNA breaks in rDNA. What is the genomic context of each rDNA gene? Does activity of each NOR regulate nearby chromatin in the same way? Do germ cells specifically express variants of other translational machinery components? How much does chromosomal activity influence NOR usage? What events led to the stabilization of a specific rDNA copy on chromosome 4 in germ cells? Are there other germ cell-specific functions associated with the region? Finally, I provide a speculation on the ability of combined Genomic-Molecular-Cellular interactions of the rDNA gene, specifically in germ cells, to influence sex determination in zebrafish (Aharon and Marlow, 2022; Brunet and Doolittle, 2015; Doolittle, 2022; Dranow et al., 2013; Pfeiffer et al., 2018). 153 Rephrasing the motivating questions The straightforward answer to the question: “why are there two variant ribosomes in zebrafish?” is that there exist two sets of subunits forming non-identical, but similar structures. Subunits are generated by two discrete ribosome biogenesis events taking place in different cell populations at different times. Each cell type distinctly uses one of two variant rDNA genes which encode for the observed differences. Ribosome functional heterogeneity may produce ribosome-specific forms of translational control (Fig. 3.3). How ribosome heterogeneity impacts development and other aspects of the biology of multicellular organisms is under active investigation. Clearly, the idea of functionalization amongst subsets of ribosomes is appealing because it represents another layer of potential regulation that may be influencing cell proliferation rates, gene expression programs, and cellular fate decisions. In an effort to assess possible ribosome functional heterogeneity in the zebrafish, a significant portion of the work presented in Chapter II of this Thesis represent attempts to assess ribosome structure and composition. While most “why” questions in Biology cannot be answered completely, we instead provide in-depth characterizations of two distinct ribosome structures with potential regulatory mechanisms in germ cell-specific gene expression. We better define the zebrafish as an experimentally accessible system for future studies involving developmentally regulated ribosome types. From considering these perspectives, we learned that two different ribosomes are built from two variant ribosomal genes expressed at distinct times and places. The following sections use the question as a means to summarize data on the variant rDNA in question. If two sets of subunits arise from two distinct ribosome biogenesis events each using different rDNA genes, the question ultimately boils down to: Why are there two variant rDNA genes used by zebrafish? The first interpretation of “why” deals with historical origination of the duplicated rDNA sequence. Investigation involving the origins of this rDNA gene variant system in zebrafish remains open for future inquiry. Another interpretation of “why” deals with a teleological description of the evolutionary fitness events occurring between the duplication and the current variants. Evolutionary fitness depends on the reliable transmittance of genetic information by germ cells (egg and sperm). This requirement is thought to act as a potent selective agent on molecular- cellular variation in germ cell programming – possibly in favor of optimizations contributing to germ cell proliferation and consequent propagation of the species. In this manner, heterogeneity involving the rDNA gene, ribosome biogenesis, subunit maturation, and translation control in germ cells may be the substrate for adaptation – leading to fixation of genetic elements. Adaptation requires that the trait be heritable, serves the function of germ cell proliferation, and increases the fitness of organisms that carry it. 154 1 – make a copy Gene duplication is one of the central avenues of biological innovation. The evolutionary potential of duplication was put into a coherent framework by Ohno in his tellingly entitled 1970 book ‘Evolution by Gene Duplication’ (Ohno, 1970). Ohno posited that, after a duplication, one of the two identical copies of a gene becomes free of selective constraints and prone to accumulating mutations that would have been wiped out by purifying selection before the duplication. DNA polymerase slipping, unequal meiotic crossing over, and transposable elements are methods of gene duplication. Notably, the rDNA gene is capable of regulating the rate of nearby recombination events, thus influencing the copy number of rDNA genes in the genome (Arnau et al., 2022; Nomura et al., 2013). 2 – escape elimination The most common fate of a duplicated gene copy will be mutational inactivation, pseudogenization, and eventual elimination. These events are underreported as the only genetic material that can be assessed are those found by sequencing modern genomes. 3 – achieve fixation Duplicates can become fixed in the genome “accidentally” by chance (aka genetic drift) or by virtue of mutation leading to new or altered function providing advantage against a selective pressure (adaptation). Functional differences between gene products must have arisen by either of these means. This requirement is at the crux of many (all?) inquiries asking “why” specific genes exist in genomes. The events by which sequences achieve fixation in the genome are hotly debated One of the most intriguing aspects of rDNA genes that has not yet been discussed in this Thesis is intragenomic concerted evolution. Sequence variation is actively removed within a genome – effectively homogenizing rDNA sequences found within or between individuals of a species (Ganley and Kobayashi, 2007; Goffová and Fajkus, 2021; Kobayashi et al., 1998; Lunerová et al., 2017). Variation in rDNA is only seen in a handful of species. Either a subset of rDNA gene variants have escaped concerted evolution or the organism is under selective pressure to evolve multiple/new rDNA sequences (Carranza et al., 1999; Eickbush and Eickbush, 2007; Keller et al., 2006). The failure of zebrafish to maintain concerted evolution of rDNA genes is most perplexing. All aspects of cellular growth are intimidatingly interconnected with the list of rDNA gene products: Protein Synthesis, Ribosome Subunits, Ribosome Biogenesis, or Chromatin Regulation. The rest of Chapter III discusses findings, implications, and future directions regarding each of the four ribosomal gene products discussed above: Protein Synthesis, Ribosome Subunits, Ribosome Biogenesis, and Gene Instability. 155 Hypotheses and Findings In this Thesis work, a variety of approaches were used to interrogate the dual ribosome system in zebrafish. Key findings from each inquiry are fully arranged in Chapter II and summarized here. A discussion of a few of these findings in relation to the zebrafish dual ribosome system and to ribosomal gene products are discussed later in Chapter III. Approaches to assay Protein Synthesis, Ribosome Structure, Ribosome Biogenesis, and Genome Regulation included posing the following queries: Does the system use two ribosomes for translation? Yes. We use sucrose density ultracentrifugation to show that the embryo utilizes both maternal and somatic rRNA-containing ribosomal subunits for translation. We originally hypothesized that maternal and somatic ribosomes must participate in embryonic translation and that any differences between the two may arise in their relative contributions to monosome vs polysome configurations. Recently, it was shown that approximately half of maternally deposited ribosomes are locked by a 3-protein dormancy complex (Leesch et al., 2023), incapable of translating mRNA. Our data support an early embryonic release of maternal ribosomes from dormancy for later participation in translation and polysome formation (Fig. 3.4). Interestingly, both labs independently detect maternal 28S rRNA tightly aligning with fractions containing polysome peaks from 24 hpf embryos; more so than somatic 28S rRNA (Fig. 2.4). Figure 3.4 – Embryonic translation simultaneously uses both ribosome subunit variants Top, polysome profile data from a sucrose density gradient centrifugation is plotted with schematized and labeled ribosomal subunits, monosomes, and polysomes. Below, maternal (yellow) and somatic (blue) intersubunit bridges are shown (see Fig. 1.19). 7 bridge structures (dark) are identical in sequence. 5 bridges (dark, ≈) contain divergent sequences, but form identical structures in our cryo-EM maps. The remaining 5 bridges (light) contain divergent sequences (in indicated sites) but are not resolved in our data. 156 Future directions involving the translational outputs of each ribosome type will involve subcellular localization analysis of each ribosome, ribosome-specific footprint analysis, and defining any ribosome-specific mRNA-specific initiation rates. Can each of the two ribosomes be pulled out of the system for measurement? Yes. We genetically engineer transgenic zebrafish and utilize a directional cross approach to epitope labeling either maternal or somatic 60S subunits (LSU). Immunoprecipitation from a lysate of subunits confirms the specificity of our methodology. Our transgenic lines (Mat-RiboFLAG and Som-RiboFLAG (Fig. 2.S8) are among the first developed tools to separately measure zebrafish maternal and somatic subunits for biochemical inquiry. Are the two sets of subunits translationally compatible with each other? Yes. We use our engineered transgenic methodology (Fig. 2.S9) to show that embryonic subunits form cognate monosomes (maternal-40S with maternal-60S and somatic-40S with somatic-60S) and hybrid monosomes (somatic-40S with maternal-60S and maternal- 40S with somatic-60S) (Fig. 3.4). Our cryo-EM maps of cognate maternal and cognate somatic ribosomes reveal 12 of 17 intersubunit bridge structures as being identical between LSUs and SSUs, potentially supporting the interchangeability of subunits and the formation of hybrid monosomes (Fig. 3.5). Prior to mapping sequence variation to known intersubunit bridges (Ben-Shem et al., 2011; Tamm et al., 2019), we originally hypothesized that rDNA sequence variation may lead to incompatible ribosome subunits as the lack of interchangeability could explain the pressure to maintain two unique developmentally regulated rDNA genes. We pull tagged maternal-60S-containing or tagged somatic-60S-containing monosomes when both ribosomes co-exist at similar levels in the 24 hpf embryo (Fig. 3.2). Accordingly, our measurements of rRNA composing immunoprecipitated RNase-treated ribosomes provide direct in vivo evidence of the formation of both cognate and hybrid monosomes (Fig. 2.5). Are the two ribosomes compositionally different? Yes. Beyond their rRNA sequence differences, each ribosome is molecularly distinct. We use TMT-MS and LC-MS/MS proteomics to measure protein compositions of both the ribosome and ribosome accessory factors, respectively, using material derived from 6 hpf and 120 hpf animals (Fig. 3.2). First, our analysis reveals a striking sameness in the RP composition of both ribosomes and highlights a handful of RP paralogs which may impart function to the ribosome structure. The differences in RPs composing each ribosome correlate with mRNA expression differences among the corresponding RP gene paralogs over developmental time (Fig. 2.2). Do the two ribosomes structurally look different? Yes. Structural discrepancy between the two is apparent, even if comprised of relatively minor differences in rRNA secondary structure. 157 We use cryo-electron microscopy (cryo-EM) to take thousands of images of ribosomes purified from early stage embryos (containing maternal type ribosomes) and late stage embryos (containing somatic type ribosomes). We originally hypothesized the ribosomes to be more dissimilar due to their heterogeneous rRNA sequences and cell types of origin. In fact, our models show the majority of electron density in each ribosome core are identical (Fig. 3.5). These data point to ribosome accessory factors and rRNA expansion segments as the major remaining unresolved regions of potential structural heterogeneity. Regions of interest for divergent ribosome structure and function are indicated (Fig. 3.6). Figure 3.5 – An assembled zebrafish ribosome Atomic model representations of our 3.2 Å resolution structure of the zebrafish maternal (6 hpf) ribosome (PDB: 7OYB). All RP atoms are shown in ball-and-stick form in grey. All rRNA atoms are shown in sphere form. Colors for maternal 5S, 18S, 5.8S (not visible), and 28S rRNAs are consistent with Fig. 2.1. Do the two rRNAs provide distinct scaffolds for two ribosomes? Yes. While most sequence discrepancies found in rRNA helical structures exhibit compensatory mutations maintaining base pairing and RNA secondary structure, many unpaired nucleotides are unique unpaired structures between each ribosome. We use our high resolution (3.2 Å and 2.8 Å) structures of each ribosome to measure base pairing of nucleotides in ribosomes composed of either maternal or somatic rRNA variants (Fig. 3.5). As much of the sequence divergence falls in expansion segments, we hypothesized most rRNA structures to be preserved between the two. Our assessment of individual nucleotides reveals the majority of sequence differences between variants do not form altered scaffolds, rather, our data support compensatory base-pair mutations 158 that maintains rRNA helical structures (Fig. 2.2). Flexible rRNA regions, including expansion segments, that cannot be structurally resolved would benefit from further study to determine the full extent of altered scaffolding abilities by either ribosome. Does ribosome heterogeneity support ribosome functional heterogeneity? Possibly. While the core of each ribosome is largely identical, several unresolved areas have been shown to participate in ribosome functional heterogeneity in other systems. We use our high resolution structures and proteomics data of each ribosome to assess the level of heterogeneity at specific positions around the ribosome (Fig. 3.6). Divergent sequences near these sites may be platforms for the ribosome to interact with RBPs or ribosome accessory factors. Investigation involving functional differences between the two ribosomes remains open for further inquiry (Anger et al., 2013; Brimacombe, 1981; Hopes et al., 2022; Kraushar et al., 2021; Wells et al., 2020; Wolin and Walter, 1988). Figure 3.6 – Variant usage in ribosome biogenesis produces heterogeneous structures Cartoon representations of the ribosome (dark grey 60S and light grey 40S) and various heterogeneous structures thought to be involved in ribosome functional heterogeneity and translational control. In the center, a ribosome is shown with E- P- and A- sites labeled. i) The peptide exit (PE) region where nascent proteins are extruded is in close proximity to divergent rRNA regions E27L, H59, H53, and H24. ii) The GTPase association center (GAC) is a binding platform for elongation factors and three divergent rRNA nucleotides were not resolved. Rplp2/P2α and Rplp2l/P2β associated with both maternal and somatic ribosomes. iii) The decoding center (DC) is where A- and P-site tRNAs (orange and red, respectively) will 159 be assessed for codon-anticodon pairing and is influenced by divergent rRNA regions ES3S and ES6S. iv) The 3’ end of the 5.8S rRNA and the 5’ end of the 28S rRNA contain divergent sequences and have been thought to play a role in disome collision-based marking of ribosomes for degradation. v) Sequence divergence in ES7L, ES15L, and ES27L regions are exemplified by varying lengths of GGC tandem- repeats. Changing the number of helices in ESs is thought to regulate RBP interactions with the ribosome. Future directions involving the differences in translational structures by each ribosome type will utilize sucrose gradient material to assess inter-ribosomal interactions (Hopes et al., 2022; Norris et al., 2021; Petrov et al., 2014) in monosomes, disomes, and polysomes. More precise fractionation of cellular components prior to grid formation for structural analysis will allow for a better look at variation of ribosome subunit structures in the cell. Can each of the two rRNAs be distinguished for measurement? Yes. We use a PCR-based fragment length polymorphism (FLP) method and an NGS- based method to measure the relative levels of either rRNA in many tissues across the zebrafish life cycle (Fig. 2.S1). Does somatic rRNA transcription trigger the degradation of maternally-deposited rRNA? Unknown. Both labs independently recapitulated findings showing the coincident increase of somatic rRNAs with a decrease of maternal rRNAs (Fig. 3.7) (Locati et al., 2017b). Figure 3.7 – The maternal-to-zygotic rRNA transition in zebrafish Each data point represents a relative measurement of maternal-type rRNAs transcribed from the rDNA gene on chromosome 4 (yellow) and somatic type rRNAs transcribed from the rDNA gene on chromosome 5 (blue). 18S rRNAs are shown in light colors and 28S rRNAs are shown in dark colors. Zebrafish were grown for the indicated hours post-fertilization at 28.5 ºC before total RNA was extracted from whole animal lysates for detection. Data generated at the Massachusetts Institute of Technology (MIT) in Cambridge are shown as filled triangles (p). Data generated at the Institute of Molecular Pathology (IMP) in Vienna are shown as empty squares (o). See Chapter II for full methods. Note the x-axis spacing change after 24 hrs. Our data confirm maternal rRNA transcription is stopped in the mature oocyte (Fig. 2.S4) (Newport and Kirschner, 1982). Therefore, the replacement of maternal rRNA must be achieved by a combination of somatic rRNA transcription and maternal rRNA degradation. However, this would require the maternal ribosome turnover rate to be considerably faster than the reported ribosome half-life of 4-6 d in rat liver (Stoykova et al., 1983) or 9-30 d in frog eggs (Brown and Gurdon, 1964). We hypothesized somatic ribosome biogenesis 160 would require the nucleotides and amino acids recovered from the biased degradation of maternal ribosomes. However, ribosome-specific degradation is difficult to distinguish during development. Our data only point to a coincident timing of these events (Fig. 3.9I). Another hypothesis involving unbiased degradation remains a likely and untested scenario. Somatic ribosome biogenesis increases the pool of translating subunits after 8 hpf, which leads to ribosome collisions and ubiquitination (Ugajin et al., 2023). As the majority of ribosomes, post-gastrulation, are maternal type, an equivalent degradation rate of all ribosomes will affect more maternal ribosomes. As disome collisions are co- translational, the loss of any given ribosome could be related to its translation efficiency. Does ribosome biogenesis of maternal and somatic ribosomes differ? Possibly. As mentioned earlier, a multi-nucleolated organization of membrane-adjacent nucleoli has long been appreciated as a phenomenon unique to oocytes (Gatenby, 1922; Montgomery Jr., 1898). Cell type-specificity of biogenesis components (rDNA transcription, rRNA processing, ribosome assembly, etc) remains open for study. Components of the Pol I PIC are duplicated in the zebrafish and some exhibit oocyte cell type-specific expression (Girbig et al., 2022; Scull and Schneider, 2019; White, 2005). We hypothesize that apart from the identity of rDNA NOR being used for ribosome biogenesis, there may be other cell type-specific molecular differences occurring in the nucleoli of oocytes. Difference between nucleoli of zebrafish germ and somatic cells are easily seen using light microscopy (Fig. 3.8). It is a coincidence that germ cells use the chromosome 4 rDNA locus for ribosome biogenesis and also achieve a multi-nucleolated architecture. The most striking difference among these cells is their size – the cross sectional area of an epithelial cell is roughly 1/70th of a stage Ib oocyte and 1/2200th of a stage III oocyte (see scale bars and insets in Fig. 3.8). In fact, many individual nucleoli in stage 1b oocytes are larger than the entire nuclei of somatic cells. Figure 3.8 – Molecularly-cellularly distinct ribosome biogenesis events occur in zebrafish Hematoxylin and eosin staining of a histological slice from a 2 year-old female zebrafish emphasizing an A) epithelial cell, a B) stage Ib oocyte, and a C) stage III oocyte. Cell dimensions are given above each image and x-y bounds are indicated as white bars. Scale bars are below each image. Insets in B and C include traced outlines of cells in A and B, drawn to scale. 161 Future investigations of ribosome biogenesis events in the zebrafish will utilize a combination of imaging and structural approaches. Cryo-EM analyses of purified nucleolar material and Cryo-FIB-SEM integration with cryo-EM, immunofluorescence, and RNA FISH will identify the structural differences being generated during each biogenesis step. Is the maternal rDNA transcribed in somatic cells? No. Consistent with others, we confirm somatic tissues to be absent of maternal type rDNA expression during normal development (Fig. 3.9IV). We also fail to measure activation of the maternal rDNA in the context of cellular dedifferentiation – specifically in mesenchymal cells during fin clip regeneration and in neoplastic melanoma experiencing neural crest stem cell-like states (Fig. 3.9V). Figure 3.9 – Differential expression of ribosome gene variants in the zebrafish life cycle The fill in each box represents the rRNA composition at the indicated times in development. I) All maternally deposited ribosomes in the egg and through 6 hpf are maternal type until 8-10 hpf when somatic type rRNA is transcribed. Germ cells likely also transcribe somatic type rRNA around this time. II) By 24 hpf half of all embryonic ribosomes are somatic type and will continue to replace maternal ribosomes in the soma by 5 dpf. PGCs will maintain 70% of maternal ribosomes by 24 hpf. Maternal type rDNA is utilized at 3 dpf and will replace somatic type ribosomes in the germline by 10 dpf. III) Females continue to utilize maternal type rDNA and maintain oocytes while males fail to continue to utilize maternal type rDNA and undergo an apoptotic event leading to male identity. IV) During growth and adult maintenance phases, the germ cell- specific chromosome 4 rDNA is specific to the gonads. V) We have yet to detect maternal rDNA transcription outside of germ cells even in disease and neoplastic conditions. Is maternal rDNA transcribed by the zygote? Or somatic rDNA transcribed in germ cells? Yes. Our data clearly indicates the misnomer “maternal” chromosome 4 rDNA undergoing active transcription in larval germ cells as early as 3 dpf (Fig. 3.9II). Our data also show primordial germ cell transcription of the somatic chromosome 5 rDNA at the same time 162 before shifting entirely to the maternal locus. We measured the majority presence of maternal ribosomes in germ cells at all measured times. We separately confirm the in vivo association of maternal ribosomes with germ cell specific transcripts at 1 day post- fertilization. While primarily restricted to the ovary, we also sporadically detect mature rRNA transcribed from the chromosome 4 rDNA variant in adult testes (Fig. 3.9IV). Our germ cell expression data distinguishes the sub-telomeric rDNA variant on chromosome 4 as a cell type-specific rDNA gene. Is maternal rDNA functioning to shape the genome? Zebrafish lack sex chromosomes and, unlike mammals, rely on the transfer of specific maternal factors for germ cell development. At least two reports directly suggest a role for the maternal rDNA locus in sex determination. A 1 Mbp sequence at the end of chromosome 4 and beginning of chromosome 5 is provided in Fig. 3.10 and Fig. 3.11. Specifically, authors suggest demethylation and amplification on extrachromosomal circles is associated with feminization (Breit et al., 2020a; Breit et al., 2020b; Tao et al., 2020). A focus on characterizing DNA methylation within the zebrafish germline, uncovered oocyte specific amplification of a 11.6 kb region that contains the maternal rDNA gene (Ortega-Recalde et al., 2019). Interestingly, the demethylation and amplification of this locus correlates with the expansion of Ib oocytes (Fig. 3.8). At this stage, oocytes contain multiple nucleoli and provide signals that drive the feminization of the gonad (Wilson et al., 2024). These results suggest modification of rDNA chromatin is linked with sex determination. Consequently, ribosome biogenesis, ribosome subunit structure and translational control are also potentially linked. 163 Conclusions In Chapter I, four properties of the rDNA gene are introduced: Protein Synthesis, Ribosome Biogenesis, Ribosome Subunits, and Gene Instability. Chapter II of this Thesis evaluates the zebrafish and its peculiar dual ribosomal system. Chapter III of this Thesis attempts to approach the questions of why two rDNA genes exist in zebrafish, let alone, how a well-studied model organism had an uncovered variation for so long. While two established perspectives in the field exist: one speculated from specific effects arising from ribosome structural heterogeneity and another constructed around general changes in ribosome concentration giving rise to specific effects; both perspectives assume the observed trait – either specific or general translational control – is selectively advantageous. By considering the findings we made via these approaches, and from several other nearby observation points, we suggest a third perspective on the origins of the system, discussed here, aimed at rDNA gene instability, specifically involving the germ cell- specific rDNA gene on chromosome 4. This perspective removes the assumption of adaptation and suggests constructive neutral evolution may provide an answer. Further research considerations should aim to incorporate observations of these four perspectives – possibly revealing molecular complexities still concealed by the origins of the zebrafish dual ribosome system. 164 Note on nomenclature For the sake of clarity and consistency with Locati and colleagues (Locati et al., 2017b; Locati et al., 2018), the chromosome 4 rDNA gene, its mature rRNA products, and the ribosomal subunits products are referred to as “maternal”, and the chromosome 5 rDNA gene, its mature rRNA products, and the ribosome subunit products are referred to as “somatic”. Prior to our reported findings in Chapter II of this Thesis, there had been an absence of direct evidence that the somatic type arises exclusively from somatic cell lineages. No investigations had measured somatic type rRNA expression in germ cell lineages. In addition, the was an absence of evidence showing maternal type arises exclusively during oogenesis for maternal inheritance. No investigations had measured maternal type rRNA expression later in development. Our work supports the naming of “somatic” type rDNA – its rRNA and subunits are restricted to the soma. Certainty of t restriction will require differentially measuring nascent transcription of either rRNA – especially in the nucleoli somatic cells of the ovary and testis. Our work does not support the naming of “maternal” type rDNA – its rRNA and subunits are primarily generated in stage I and II oocytes, minorly present in testes, and zygotically transcribed in the larval stage germline. Names like “germ cell rDNA” may make more sense from a cell biology perspective; however, gene names including a cell type name of the gene’s cell type-specific expression is cumbersome when dealing with ex vivo, in vitro, and recombinant contexts. Names like “fem-rDNA” may make sense from an organismal function perspective, as it is thought that the maternal rDNA locus is tied to feminizing signals in the ovary (Ortega- Recalde et al., 2019). Similarly, names like “early” and “late” (Ramachandran et al., 2020) are not specific enough outside the context of the developing embryo. Making matters more complicated, at least three more rDNA gene variants have been shown to be reliably expressed at low levels (Locati et al., 2017b; Tao et al., 2020) in the zebrafish. I propose yet another possible nomenclature for rDNA variant genes in zebrafish: rDNA477 the rDNA gene chr4: 77,555,053 – 77,564,140 (GRCz11) rDNA500 the rDNA gene chr5: 00,819,029 – 00,827,807 (GRCz11) And offer examples of its usage in reference to rDNA gene products: r477_ES27L the ES27L sequence found in the 28S of rDNA477 r500_28S the 28S rRNA transcribed from rDNA500 r477_SSU the SSU built using rDNA477 maternal ribosome ribosomes in the egg; mostly r477_SSU and r477_LSU zygotic r477_28S zygotically transcribed 28S rRNA from rDNA477 (in PGCs) 165 166 Figure 3.10 and Figure 3.11 – 1 Mbp regions surrounding the zebrafish rDNA A) The repetitive elements contained in each rDNA surrounding transcribed regions are schematized to scale with the 18S, 5.8S, and 28S rRNAs. B) DNA nucleotide characters A, T, C, and G are shown as blue, green, red, and yellow pixels. Each of the 10 columns represents sequential 100 kbp regions of the genome. Each column is similar to a FASTA file – read left to right, with 100 characters on each line; 1000 lines down. The repetitive composition of the sub-telomeric regions is readily apparent in this arrangement. 167 References Aharon, D. and Marlow, F. L. (2022). Sexual determination in zebrafish. Cell. Mol. Life Sci. 79, 1–19. Anger, A. M., Armache, J.-P., Berninghausen, O., Habeck, M., Subklewe, M., Wilson, D. N. and Beckmann, R. (2013). Structures of the human and Drosophila 80S ribosome. Nature 497, 80–85. Arnau, V., Barba-Aliaga, M., Singh, G., Ferri, J., García-Martínez, J. and Pérez-Ortín, J. E. (2022). A feedback mechanism controls rDNA copy number evolution in yeast independently of natural selection. PloS One 17, e0272878. Ben-Shem, A., Garreau de Loubresse, N., Melnikov, S., Jenner, L., Yusupova, G. and Yusupov, M. (2011). The structure of the eukaryotic ribosome at 3.0 Å resolution. Science 334, 1524–1529. Breit, T. M., Rauwerda, H., Pagano, J. F. B., Ensink, W. A., Nehrdich, U., Spaink, H. P. and Dekker, R. J. (2020a). Immunoglobulin switch-like recombination regions implicated in the formation of extrachromosomal circular 45S rDNA involved in the maternal-specific translation system of zebrafish. Developmental Biology. Breit, T. M., Pagano, J. F. B., Jagt, P. L. van der, Mittring, E., Ensink, W. A., Olst, M. van, Leeuwen, S. van, Leeuw, W. de, Nehrdich, U., Spaink, H. P., et al. (2020b). New observations on non-coding RNAs involved in the dual translation system in zebrafish development. 869651. Brimacombe, R. (1981). Secondary structure and evolution of ribosomal RNA. Nature 294, 209–210. Brown, D. D. and Gurdon, J. B. (1964). Absence of ribosomal rna synthesis in the anucleolate mutant of xenopus laevis. Proc. Natl. Acad. Sci. 51, 139–146. Brunet, T. D. P. and Doolittle, W. F. (2015). Multilevel Selection Theory and the Evolutionary Functions of Transposable Elements. Genome Biol. Evol. 7, 2445– 2457. Carranza, S., Baguñà, J. and Riutort, M. (1999). Origin and evolution of paralogous rRNA gene clusters within the flatworm family Dugesiidae (Platyhelminthes, Tricladida). J. Mol. Evol. 49, 250–259. Doolittle, W. F. (2022). All about levels: transposable elements as selfish DNAs and drivers of evolution. Biol. Philos. 37, 24. Dranow, D. B., Tucker, R. P. and Draper, B. W. (2013). Germ cells are required to maintain a stable sexual phenotype in adult zebrafish. Dev. Biol. 376, 43–50. Eickbush, T. H. and Eickbush, D. G. (2007). Finely Orchestrated Movements: Evolution of the Ribosomal RNA Genes. Genetics 175, 477–485. Ganley, A. R. D. and Kobayashi, T. (2007). Highly efficient concerted evolution in the ribosomal DNA repeats: Total rDNA repeat variation revealed by whole-genome shotgun sequence data. Genome Res. 17, 184–191. Gatenby, J. B. (1922). The Cytoplasmic Inclusions of the Germ-Cells: Part X. The Gametogenesis of Saccocirrus1. J. Cell Sci. s2-66, 1–48. Girbig, M., Misiaszek, A. D. and Müller, C. W. (2022). Structural insights into nuclear transcription by eukaryotic DNA-dependent RNA polymerases. Nat. Rev. Mol. Cell Biol. 23, 603–622. 168 Goffová, I. and Fajkus, J. (2021). The rDNA Loci—Intersections of Replication, Transcription, and Repair Pathways. Int. J. Mol. Sci. 22, 1302. Hopes, T., Norris, K., Agapiou, M., McCarthy, C. G. P., Lewis, P. A., O’Connell, M. J., Fontana, J. and Aspden, J. L. (2022). Ribosome heterogeneity in Drosophila melanogaster gonads through paralog-switching. Nucleic Acids Res. 50, 2240– 2257. Keller, I., Chintauan-Marquier, I. C., Veltsos, P. and Nichols, R. A. (2006). Ribosomal DNA in the grasshopper Podisma pedestris: escape from concerted evolution. Genetics 174, 863–874. Kobayashi, T., Heck, D. J., Nomura, M. and Horiuchi, T. (1998). Expansion and contraction of ribosomal DNA repeats in Saccharomyces cerevisiae: requirement of replication fork blocking (Fob1) protein and the role of RNA polymerase I. Genes Dev. 12, 3821–3830. Kraushar, M. L., Krupp, F., Harnett, D., Turko, P., Ambrozkiewicz, M. C., Sprink, T., Imami, K., Günnigmann, M., Zinnall, U., Vieira-Vieira, C. H., et al. (2021). Protein Synthesis in the Developing Neocortex at Near-Atomic Resolution Reveals Ebp1- Mediated Neuronal Proteostasis at the 60S Tunnel Exit. Mol. Cell 81, 304-322.e16. Leesch, F., Lorenzo-Orts, L., Pribitzer, C., Grishkovskaya, I., Roehsner, J., Chugunova, A., Matzinger, M., Roitinger, E., Belačić, K., Kandolf, S., et al. (2023). A molecular network of conserved factors keeps ribosomes dormant in the egg. Nature 613, 712–720. Locati, M. D., Pagano, J. F. B., Ensink, W. A., van Olst, M., van Leeuwen, S., Nehrdich, U., Zhu, K., Spaink, H. P., Girard, G., Rauwerda, H., et al. (2017a). Linking maternal and somatic 5S rRNA types with different sequence-specific non-LTR retrotransposons. RNA N. Y. N 23, 446–456. Locati, M. D., Pagano, J. F. B., Girard, G., Ensink, W. A., van Olst, M., van Leeuwen, S., Nehrdich, U., Spaink, H. P., Rauwerda, H., Jonker, M. J., et al. (2017b). Expression of distinct maternal and somatic 5.8S, 18S, and 28S rRNA types during zebrafish development. RNA N. Y. N 23, 1188–1199. Locati, M. D., Pagano, J. F. B., Abdullah, F., Ensink, W. A., van Olst, M., van Leeuwen, S., Nehrdich, U., Spaink, H. P., Rauwerda, H., Jonker, M. J., et al. (2018). Identifying small RNAs derived from maternal- and somatic-type rRNAs in zebrafish development. Genome 61, 371–378. Lunerová, J., Renny-Byfield, S., Matyâšek, R., Leitch, A. and Kovařík, A. (2017). Concerted evolution rapidly eliminates sequence variation in rDNA coding regions but not in intergenic spacers in Nicotiana tabacum allotetraploid. Plant Syst. Evol. 303, 1043–1060. Montgomery Jr., T. S. H. (1898). Comparative cytological studies, with especial regard to the morphology of the nucleolus. J. Morphol. 15, 265–582. Newport, J. and Kirschner, M. (1982). A major developmental transition in early Xenopus embryos: II. Control of the onset of transcription. Cell 30, 687–696. Nomura, M., Nogi, Y. and Oakes, M. (2013). Transcription of rDNA in the Yeast Saccharomyces cerevisiae. In Madame Curie Bioscience Database [Internet], p. Landes Bioscience. Norris, K., Hopes, T. and Aspden, J. L. (2021). Ribosome heterogeneity and specialization in development. WIREs RNA 12, e1644. 169 Ohno, S. (1970). Evolution by Gene Duplication. Springer Science & Business Media. Ortega-Recalde, O., Day, R. C., Gemmell, N. J. and Hore, T. A. (2019). Zebrafish preserve global germline DNA methylation while sex-linked rDNA is amplified and demethylated during feminisation. Nat. Commun. 10, 3053. Petrov, A. S., Bernier, C. R., Hsiao, C., Norris, A. M., Kovacs, N. A., Waterbury, C. C., Stepanov, V. G., Harvey, S. C., Fox, G. E., Wartell, R. M., et al. (2014). Evolution of the ribosome at atomic resolution. Proc. Natl. Acad. Sci. U. S. A. 111, 10251– 10256. Pfeiffer, J., Tarbashevich, K., Bandemer, J., Palm, T. and Raz, E. (2018). Rapid progression through the cell cycle ensures efficient migration of primordial germ cells - The role of Hsp90. Dev. Biol. 436, 84–93. Ramachandran, S., Krogh, N., Jørgensen, T. E., Johansen, S. D., Nielsen, H. and Babiak, I. (2020). The shift from early to late types of ribosomes in zebrafish development involves changes at a subset of rRNA 2′-O-Me sites. RNA 26, 1919–1934. Scull, C. E. and Schneider, D. A. (2019). Coordinated Control of rRNA Processing by RNA Polymerase I. Trends Genet. TIG 35, 724–733. Stoykova, A. S., Dudov, K. P., Dabeva, M. D. and Hadjiolov, A. A. (1983). Different Rates of Synthesis and Turnover of Ribosomal RNA in Rat Brain and Liver. J. Neurochem. 41, 942–949. Tamm, T., Kisly, I. and Remme, J. (2019). Functional Interactions of Ribosomal Intersubunit Bridges in Saccharomyces cerevisiae. Genetics 213, 1329–1339. Tao, B., Lo, L. J., Peng, J. and He, J. (2020). rDNA subtypes and their transcriptional expression in zebrafish at different developmental stages. Biochem. Biophys. Res. Commun. 529, 819–825. Ugajin, N., Imami, K., Takada, H., Ishihama, Y., Chiba, S. and Mishima, Y. (2023). Znf598- mediated Rps10/eS10 ubiquitination contributes to the ribosome ubiquitination dynamics during zebrafish development. Molecular Biology. Wells, J. N., Buschauer, R., Mackens-Kiani, T., Best, K., Kratzat, H., Berninghausen, O., Becker, T., Gilbert, W., Cheng, J. and Beckmann, R. (2020). Structure and function of yeast Lso2 and human CCDC124 bound to hibernating ribosomes. PLOS Biol. 18, e3000780. White, R. J. (2005). RNA polymerases I and III, growth control and cancer. Nat. Rev. Mol. Cell Biol. 6, 69–78. Wilson, M. L., Romano, S. N., Khatri, N., Aharon, D., Liu, Y., Kaufman, O. H., Draper, B. W. and Marlow, F. L. (2024). Rbpms2 promotes female fate upstream of the nutrient sensing Gator2 complex component, Mios. Developmental Biology. Wolin, S. L. and Walter, P. (1988). Ribosome pausing and stacking during translation of a eukaryotic mRNA. EMBO J. 7, 3559–3569. 170 Note I We are not in the publishing business; we are in the business of revealing truth. – Angelika Amon 2016 171 The blind and the elephant – Andhgajanyāyah The story This parable is an account of a group of blind people who have never before experienced an elephant, and use touch to probe the unfamiliar animal for information about its identity. Each person separately feels a distinct part of the elephant's body, such as the ear, the leg, or the tusk. After independent sightless evaluation, they share their respective interpretations of the animal. Every portrayal is different; often with conflicting assessments. Only upon humble consideration of multiple viewpoints do they realize each of their ideas about the animal are both incomplete and valid. The discrepancies point to a more complex answer – consistent with their knowledge and possibly consistent with unknown qualities still yet to be assessed by others using different methods. The origin Andhgajanyāyah, originates around 500 BCE, teaching the dialectical concepts of Anekäntaväda and Syädväda, which best translate to “many-sidedness” and “conditioned perspective”, respectively. Reality is described as being multifaceted, composed of many dimensions; with any one individual’s conditional perspective limiting their understanding to only the few principal components apparent to them. The story teaches that there may be truth to what someone perceives, regardless of if their interpretations contrast with your own. The provided axiom holds that all knowledge claims are only ever tentative and should be considered in the form “X may be Y”, even if expressed as “X is Y”. The interpretation Sometimes, our personally privileged perspective permits us to perceive truth. Sometimes we cannot. This unknown probability should encourage skepticism of even our most well-supported models. Another individual may be advantaged to witness the system from a different perspective we do not have access to. This unknown probability should encourage collaboration to acquire complementary support to our conclusions. Only by incorporating multiple orthogonally taken measurements from multiple conditionally positioned perspectives can we make an attempt to comprehend the unknown.